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From: Colin.Veitch-at-csiro.au
Date: Tue, 1 May 2007 00:56:33 -0500
Subject: [Microscopy] imaging plant (carrot) cell walls

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

G'day,

We have been asked to look at the cell wall structure of some carrots
and have essentially no experience in this area at all. At this stage
we are interested in the cellulose structure to start with.

We have access to a Hitachi VP FESEM and JEOL TEM as well as a range of
optical techniques. What would be the best technique for sample
preparation (CPD etc) and imaging?

Thank you very much in advance for your help with this.

Colin Veitch

Electron Microscopist
CSIRO Textile and Fibre Technology
PO Box 21, BELMONT, Vic. 3216. Australia.
colin.veitch-at-csiro.au
http://www.tft.csiro.au

+61 (0) 3 5246 4000
0438 538 475
+61 (0) 3 5246 4811

The information contained in this e-mail message may be privileged or
confidential information. If you are not an intended recipient, you may
not copy, distribute or take any action in reliance on it. If you have
received this message in error, please telephone CSIRO Textile and Fibre
Technology on +61 3 5246 4000.



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From: keith.morris-at-ucl.ac.uk
Date: Tue, 1 May 2007 03:50:03 -0500
Subject: [Microscopy] AskAMicroscopist: Building Criteria for Optical

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Vibration control is important and our confocal/time-lapse/laser dissection
microscopes have at least an in-house made heavy cheap damping plate (140kg)
with anti-vibrational pads (which prevent banging on the support worktop
wobbling the image on the VDU screen) or at best a professional air-table
with the confocals and TIRF. Our facility is on the second floor in busy
London and the building has no special anti-vibration features - the
opposite really as shutting an outside corridor door can cause a judder on
the screen without some sort of isolation system. Stout, well fitted thick
worktops with extra support under the microscope, also helps (particularly
with 140+kg on it). Very basic lab microscopes (~£5,000) used at 10x to 40x
seem to get by with minimal vibration protection, probably as their optics
are so poor that is the least of their problems being optimised for under
200x magnification. Our Leica SP2 AOBS came with a natty Leica granite and
squash ball number and the PALM micro-dissection system with the heavy
plate/rubber pads approach, both similar to our cheap in-house approach, but
both work OK up to the full 1,000x mag - i.e. you don't seem to need much
more vibrational protection for optical microscopes. This is important as
invariably throughout their life the microscopes move slowly about the
building from room to room as the wind of departmental politics change.

The main problem we get is people leaning the anti-vibrational tables to
write, and things like focus and manual stage drift caused by air
temperature variations in the air-conditioned room (although this is
minimised by using a large incubator enclosing the stage and objectives for
live cell work, and the use of motorised stages), see below.

At UKAEA Harwell we did have a Hitachi low voltage hi-res SEM that was
moved, during business 'restructuring', from it's purpose built
anti-vibrational room with faraday cage shielding and window blinds, and
ended up in the ground floor of our building in a normal lab. The windows
were covered with tin foil, and, particularly every time a large vehicle
drove by, the image at high mag was unusable (the hi-mag image always
wobbled whatever). The old purpose built suite of microscope rooms, at
another site, were converted to offices. This was 15 years ago, so I don't
know if such SEM's have better vibration protection now, but in this case
the very expensive purpose built room was certainly better.

In general vibration is the least of our problems, although all the rooms
(even those being built for microscopes) were designed with little thought
having tacky thin wobbly worktop fittings. More importantly things like
putting the cold feed out vent above the microscope is common to all the
rooms and temperature control is only to office standard with variations of
typically 4 to 5 oC (and dropping to 16oC overnight with the microscopes
off, making the Zeiss/Leica anti-fluorescent immersion go cloudy - heat to
40oC to clear it). A temperature change of 15oC, as the live cell incubator
warms up, moves focus out by about 35um, so ambient temperature changes are
a very serious problem. This plays havoc with focus and any manual XY stage
(both drift). Ideally keep the room fairly warm (22oC, or higher if you use
incubators, is ideal) and shield any air-conditioning out vents from the
microscope area. In all cases we have had to pay £1,000+ to improve the
'balanced' air condition temperature control system - the art of the
Victorian concept of a thermostat, set to 22oC, seems to have been lost. So
make sure you get this right. Even a large stage incubator can get into
trouble fighting against a poorly setup up air conditioning system. Plus
ensure that the exhaust heat from the argon laser of a confocal is suitably
ducted out of the building (they give out 3 to 5 kW). Heat generated from
these confocals with also test the quality of any room's air conditioning
system.

We also get mains interference effects on transmission confocal images with
some systems occasionally (according to the manufacturers engineers, but I
suspect a dodgey internal connection though). All of our systems have some
of rudimentary mains spike/surge filtration and UPS PC protection, and it
would be worth thinking of putting quality mains spike protection into the
wiring system of a microscope room when it's being built (with UPS being a
stand alone box by the PC).

Keith

---------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology
Institute of Ophthalmology
UCL, London EC1V 9EL





-----Original Message-----
X-from: ph2-at-sprynet.com [mailto:ph2-at-sprynet.com]
Sent: 01 May 2007 03:17
To: keith.morris-at-ucl.ac.uk


1. A recent study was published in Sound and Vibration on Microscope
Acceptability of vibration for optical ones. (I'm at home and don't have a
copy with me)

- Vibration Sensitivity of Laboratory Bench Microscopes
Hal Amick and Matthew Stead, Colin Gordon & Associates

Benchtop optical microscopes are among the most common tools found in R&D
facilities. The importance of vibration isolation to maximize image quality
is discussed. Vibration criteria are presented.

2. I talked to the author last week (I have always been interested in
this since 3D vibration analysis work at GA Tech years ago) about some other
things that could be used as criteria for assessment and asked him to submit
an abstract for the upcoming Inter-Micro in Chicago, IL in July. Hopefully
he'll make it. In the mean time if you need a copy of the article contact
me or the S&V journal (It's not on their website yet
http://www.sandv.com/home.htm ).

3. This article does not cover the building aspects, but a good
vibration engineer could help there in conjunction with a structural
engineer, particularly in combination with the frequency response data from
this article.

Tony

.....................................................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
pH2, LLC
5250 E US 36, Suite 830
Avon, IN 46123
(317) 718-7020 off
(317) 718-7038 fax
(317) 409-3238 cell

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-----Original Message-----
X-from: sandra.filippi-at-montgomerycollege.edu
[mailto:sandra.filippi-at-montgomerycollege.edu]
Sent: Monday, April 30, 2007 9:21 PM
To: ph2-at-sprynet.com

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (sandra.filippi-at-montgomerycollege.edu) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday,
April 30, 2007 at 20:12:50
---------------------------------------------------------------------------

Email: sandra.filippi-at-montgomerycollege.edu
Name: Sandra Lee Filippi, Campus Planner

Organization: montgomery college

Education: Graduate College

Location: Rockville MD

Question:


Dear Sir or Madam Microscopist:

Montgomery College is in design development for a new academic teaching
science center. The lab planner for the design team is citing ASHRAE TC2.6
Guideline Criteria for use in Planning New Facilities and insists that we
design to the VC-B level because the biology department uses compound
microscopes with total magnification of 1,000x (oil immersion). I managed
academic science labs for a two-year college for more than 20 years. We
routinely used light microscopes at total magnification of 1,000x (oil
immersion) in a facility built in 1965 without special vibration control.
When we opened our new science center in 1995 it did not have special
vibration control either. I have studied at Georgetown University and did
student research at The NIH. Neither of these facilities required special
vibration control for a light microscope.
I have consulted with the Westover Scientific Customer Service/Product
Specialist ñ Microscopy who states, ìTo be honest with you I have never
heard of such a thing, neither have my colleagues. ÝThere are tables that
are manufactured to reduce vibrations, but Iíve never heard of a building
being designed around a microscope. ÝWe have customers who use microscopes
in old buildings all over the world. Westover recommends that a microscope
be used on a vibration free surface which typically means not locating your
microscope on a table with other equipment that can cause vibrations such as
fans and other lab equipment. Microscopes have been used for years and
years on standard benchtops and other tables without issue. It's not our
position to recommend additional support when building a new structure. In
my entire career I have never heard of this requirement.î Westover
manufactures microscopes for Fisher Scientific.
The lab planner is going to cost us dearly unless I can refute his
recommendation. What is your professional opinion in this matter? Thank
you very much for your time.
Sandra

Sandra Lee Filippi, Campus Planner
Montgomery College
Suite 200
40 West Gude Drive Room 223
Rockville MD 20850-1166
301.251.7362 vox
301.251.7379 fax
240.882.6672 cell


---------------------------------------------------------------------------


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From: Sandra.Filippi-at-montgomerycollege.edu
Date: Tue, 1 May 2007 06:41:21 -0500
Subject: [Microscopy] AskAMicroscopist: Building Criteria for Optical

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Tony:

May I please have a copy of the article? I am able to receive scanned (PDF) attachments to email and/or fax to the numbers in my signature. THANK YOU.

Sandra

Sandra Lee Filippi, Campus Planner

Suite 200
40 West Gude Drive Room 223
Rockville MD 20850-1166
301.251.7362 vox
301.251.7379 fax
240.882.6672 cell
sandra.filippi-at-montgomerycollege.edu

-----Original Message-----
X-from: Tony Havics [mailto:ph2-at-sprynet.com]
Sent: Monday, April 30, 2007 10:13 PM
To: Filippi, Sandra; Micrscopy Listserve

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (sandra.filippi-at-montgomerycollege.edu) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday,
April 30, 2007 at 20:12:50
---------------------------------------------------------------------------

Email: sandra.filippi-at-montgomerycollege.edu
Name: Sandra Lee Filippi, Campus Planner

Organization: montgomery college

Education: Graduate College

Location: Rockville MD

Question:


Dear Sir or Madam Microscopist:

Montgomery College is in design development for a new academic teaching
science center. The lab planner for the design team is citing ASHRAE TC2.6
Guideline Criteria for use in Planning New Facilities and insists that we
design to the VC-B level because the biology department uses compound
microscopes with total magnification of 1,000x (oil immersion). I managed
academic science labs for a two-year college for more than 20 years. We
routinely used light microscopes at total magnification of 1,000x (oil
immersion) in a facility built in 1965 without special vibration control.
When we opened our new science center in 1995 it did not have special
vibration control either. I have studied at Georgetown University and did
student research at The NIH. Neither of these facilities required special
vibration control for a light microscope.
I have consulted with the Westover Scientific Customer Service/Product
Specialist ñ Microscopy who states, ìTo be honest with you I have never
heard of such a thing, neither have my colleagues. ÝThere are tables that
are manufactured to reduce vibrations, but Iíve never heard of a building
being designed around a microscope. ÝWe have customers who use microscopes
in old buildings all over the world. Westover recommends that a microscope
be used on a vibration free surface which typically means not locating your
microscope on a table with other equipment that can cause vibrations such as
fans and other lab equipment. Microscopes have been used for years and
years on standard benchtops and other tables without issue. It's not our
position to recommend additional support when building a new structure. In
my entire career I have never heard of this requirement.î Westover
manufactures microscopes for Fisher Scientific.
The lab planner is going to cost us dearly unless I can refute his
recommendation. What is your professional opinion in this matter? Thank
you very much for your time.
Sandra

Sandra Lee Filippi, Campus Planner
Montgomery College
Suite 200
40 West Gude Drive Room 223
Rockville MD 20850-1166
301.251.7362 vox
301.251.7379 fax
240.882.6672 cell


---------------------------------------------------------------------------


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12, 13 -- From: sandra.filippi-at-montgomerycollege.edu (by way of
Ask-A-Microscopist)
12, 13 -- Subject: AskAMicroscopist: Building Criteria for Optical
Microscope Lab
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From: baskin-at-bio.umass.edu
Date: Tue, 1 May 2007 06:48:34 -0500
Subject: [Microscopy] Re: imaging plant (carrot) cell walls

Contents Retrieved from Microscopy Listserver Archives
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G'day Colin,
I will assume that when you wrote "some carrots" you mean
carrot roots, the big bright orange things we eat. I can give you a
couple of suggestions although clearly without knowing more about the
goals I am a little in the dark.

1. Good old polarized light microscopy. As I am sure you know from
fibers (i.e., fibres), cellulose is birefringent. If you embed the
roots in your favorite resin, cut semi thin sections (don't stain),
you can then image in polarized light. This can give you net
orientation (mean extinction angle) and information about the extent
of orientation (retardance). Note that while mature xylem
fibers/vessels in the carrot root will have a retardance on the order
of cotton, that of the rank and file cells in the root will be much
less, so you need a reasonably sensitive polarized light set up.

2. SEM. A nice trick to image the inner most layer of the cell wall
with SEM is to make fresh sections. By this I mean cut the roots
before you fix them. The turgor pressure ejects most/all of the cell
contents, including the plasma membrane. Cut the sections in somthing
simple like PBS and rinse for 10 min. I have never tried this with
carrot roots but on a variety of other plant tissues, this works.
What you do next depends on your instrument. In my lab, we fix,
dehydrate in ethanol, do CPD, coat with Pt, and image with high vac
FESEM. This shows cell wall structure nicely. Microfibrillar
structures revealed in this way are on on order of 10 nm. There has
been no extraction so they contain more than cellulose. With a VP
instrument, you might be able to skip all that and image directly,
right after rinsing. You may or may not be able to image these
reliably in VP mode.

Hope this helps. Feel free to contact me with questions.

Tobias


}
}
} G'day,
}
} We have been asked to look at the cell wall structure of some carrots
} and have essentially no experience in this area at all. At this stage
} we are interested in the cellulose structure to start with.
}
} We have access to a Hitachi VP FESEM and JEOL TEM as well as a range of
} optical techniques. What would be the best technique for sample
} preparation (CPD etc) and imaging?
}
} Thank you very much in advance for your help with this.
}
} Colin Veitch
}
} Electron Microscopist
} CSIRO Textile and Fibre Technology
} PO Box 21, BELMONT, Vic. 3216. Australia.
} colin.veitch-at-csiro.au
} http://www.tft.csiro.au
}
} +61 (0) 3 5246 4000
} 0438 538 475
} +61 (0) 3 5246 4811
}
} The information contained in this e-mail message may be privileged or
} confidential information. If you are not an intended recipient, you may
} not copy, distribute or take any action in reliance on it. If you have
} received this message in error, please telephone CSIRO Textile and Fibre
} Technology on +61 3 5246 4000.
}
}
}
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--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Biology Department
/_ / __ /__ \ \ \__ 611 N. Pleasant St.
/ / / \ \ \ University of Massachusetts
/ / / \ \ \ Amherst, MA, 01003
/ / ___ / \ \__/ \ ____
http://www.bio.umass.edu/biology/baskin/
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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From: bliss5-at-llnl.gov
Date: Tue, 1 May 2007 11:42:05 -0500
Subject: [Microscopy] Re: viaWWW: desktop microscopist

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Maria:

I fear that Desktop Microscopist is no more. I have tried, with no
luck, to contact the source; Lacuna Labs. The authors did not respond
to my email, after I searched them out, several months ago, when the
computer my application ran on died.

I have used that particular application for years in my work with
Laue x-ray backscatter diffraction. The version I have is 2.1.x from
1998. It will not work on an operating system beyond 9.x and my
onsite computer geeks don't support that old stuff anymore.

I now use another application called ScanOrient to back up my DM
information. But it is not quite the same.
--

+++++++++++++++++++++++++++++

R. Ann Bliss
Technologist, Chemistry Materials and Life Science
Materials Science and Technology Division
Lawrence Livermore National Laboratory

_____________________________

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From: tbargar-at-unmc.edu
Date: Tue, 1 May 2007 15:39:40 -0500
Subject: [Microscopy] Help with enhancing contrast of Mitochondrial cristae

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Dear listers,

Anyone out there have any advice on how to enhance the contrast and
definition of the membranes of the cristae of mitochondria? The samples
brought to me are monolayer cell cultures of cancer cells grown on
Thermanox coverslips. This is how I'm currently processing the samples:
Primary fixation is 2% glutaraldehyde, 2% paraformaldehyde, 0.5% acrolein
in 0.1M Sorensen's phosphate buffer pH 7.2. Post-fixation in 1%Osmium
Tetroxide in 0.1M Sorensen's phosphate buffer. Dehydration in 50, 70, 90,
95, 100%X3 ethanol solutions. embedding in Araldite. Sections are stained
with 2% uranyl acetate aqueous 15 minutes and Reynold's lead citrate 10
minutes. The density of the cytoplasm and mitochondrial matrix are
similar with the result that the contrast of the mitochondria is similar to
the cytoplasm. The mitochondria and it's membranes (outer and that of the
cristae) don't really "stand out". The researchers involved want to see
really contrasty mitochondrial cristae.

The next thing I'm going to try is post-fixation with a mix of osmium
tetroxide and potassium ferrocyanide.

Anyone have any other ideas? Thanks in advance.


Tom Bargar
University of Nebraska Medical Center
Core Electron Microscopy Research Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu


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From: lcgould-at-med.cornell.edu
Date: Tue, 1 May 2007 16:01:22 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial cristae

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Tom-
You anticipated one of my responses by stating that you are going to
use K-Ferrocyanide with the osmium.
Another thing you can try (along with the Os/K-fer) is to use the
following as your primary fix:
4% pfa, 2.5% glut, 0.002% picric acid in No-Cacod. buffer (original
ref: It & Karnovsky J Cell Biol Vol 89 Abstract #418, 1968).

I was first told about this fix by a group who studied outer rod
segments of the eye....LOTS of membranes!

Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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From: Michal.Jarnik-at-fccc.edu
Date: Tue, 1 May 2007 16:31:26 -0500
Subject: [Microscopy] SYTO Orange stains

Contents Retrieved from Microscopy Listserver Archives
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I need to stain nucleic acids in the 540 nm excitation range on semi
thin cryosections. I selected SYTO 83 from Molecular Probes(exc. 543,
emission 559), but so far was not very successful - the cytoplasm
actually shows more staining than the nucleus. They are not very
specific about the protocol (except that one should not use phosphate
buffers). Does anybody there in cyberspace have any experience with
these dyes?

Thanks,

--
Michal Jarnik


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From: Walter.Bobrowski-at-pfizer.com
Date: Wed, 2 May 2007 07:33:20 -0500
Subject: [Microscopy] Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Tom, try an ethanolic-based UA stain:

Mix in the following proportions:
0.2g UA
3ml 50% ethanol

Let mix at least several hours on rocker/shaker.
Next day, try a series of UA staining times and examine (say, every 5
minutes). I've found 10-15 minutes optimal, as longer times appeared to
also darken surrounding plastic matrix.
Pick your optimal time, stain, rinse, and counterstain with Reynold's
LC, ~5-10 minutes, rinse.

Post-fixing in reduced osmium as you state is my other suggestion, but
see if ethanolic UA gives you what you need before re-embedding more
samples. Also see if the two techniques combined (reduced osmium,
ethanolic UA stain) give you even better results!

Be sure to post your findings.
Best regards,

Walter F. Bobrowski
Sr. Scientist
Pfizer Global R&D
2800 Plymouth Rd.
Ann Arbor, MI 48105

TEL: 734-622-7814
FAX: 734-622-5718

"Like the strength of a steel rod, the true character of a person can
only be known under extreme stress." - Leslie Fieger




-----Original Message-----
X-from: tbargar-at-unmc.edu [mailto:tbargar-at-unmc.edu]
Sent: Tuesday, May 01, 2007 4:46 PM
To: Bobrowski, Walter


Dear listers,

Anyone out there have any advice on how to enhance the contrast and
definition of the membranes of the cristae of mitochondria? The samples
brought to me are monolayer cell cultures of cancer cells grown on
Thermanox coverslips. This is how I'm currently processing the samples:
Primary fixation is 2% glutaraldehyde, 2% paraformaldehyde, 0.5%
acrolein
in 0.1M Sorensen's phosphate buffer pH 7.2. Post-fixation in 1%Osmium
Tetroxide in 0.1M Sorensen's phosphate buffer. Dehydration in 50, 70,
90,
95, 100%X3 ethanol solutions. embedding in Araldite. Sections are
stained
with 2% uranyl acetate aqueous 15 minutes and Reynold's lead citrate 10
minutes. The density of the cytoplasm and mitochondrial matrix are
similar with the result that the contrast of the mitochondria is similar
to
the cytoplasm. The mitochondria and it's membranes (outer and that of
the
cristae) don't really "stand out". The researchers involved want to see
really contrasty mitochondrial cristae.

The next thing I'm going to try is post-fixation with a mix of osmium
tetroxide and potassium ferrocyanide.

Anyone have any other ideas? Thanks in advance.


Tom Bargar
University of Nebraska Medical Center
Core Electron Microscopy Research Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu


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25, 31 -- From Walter.Bobrowski-at-pfizer.com Wed May 2 07:33:14 2007
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From: dsoren-at-umich.edu
Date: Wed, 2 May 2007 10:52:50 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial cristae

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Tom,

It is possible that you are losing your cellular membranes during the
dehydration steps. First, en bloc stain with saturated uranyl
acetate in water and then go through the dehydration steps quickly,
about 2 minutes for each step. Begin with 50 % EtOH and end with
only one change of 100 % EtOH. If you are infiltrating with EtOH:
araldite, keep the steps with high EtOH content short, too, about a
30 minute maximum.

I hope that helps!

Dotty Sorenson

On May 1, 2007, at 4:44 PM, tbargar-at-unmc.edu wrote:

}
}
}
} ----------------------------------------------------------------------
} ------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/
} MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------
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}
}
} Dear listers,
}
} Anyone out there have any advice on how to enhance the contrast and
} definition of the membranes of the cristae of mitochondria? The
} samples
} brought to me are monolayer cell cultures of cancer cells grown on
} Thermanox coverslips. This is how I'm currently processing the
} samples:
} Primary fixation is 2% glutaraldehyde, 2% paraformaldehyde, 0.5%
} acrolein
} in 0.1M Sorensen's phosphate buffer pH 7.2. Post-fixation in 1%Osmium
} Tetroxide in 0.1M Sorensen's phosphate buffer. Dehydration in 50,
} 70, 90,
} 95, 100%X3 ethanol solutions. embedding in Araldite. Sections are
} stained
} with 2% uranyl acetate aqueous 15 minutes and Reynold's lead
} citrate 10
} minutes. The density of the cytoplasm and mitochondrial matrix are
} similar with the result that the contrast of the mitochondria is
} similar to
} the cytoplasm. The mitochondria and it's membranes (outer and that
} of the
} cristae) don't really "stand out". The researchers involved want
} to see
} really contrasty mitochondrial cristae.
}
} The next thing I'm going to try is post-fixation with a mix of osmium
} tetroxide and potassium ferrocyanide.
}
} Anyone have any other ideas? Thanks in advance.
}
}
} Tom Bargar
} University of Nebraska Medical Center
} Core Electron Microscopy Research Facility
} 986395 Nebraska Medical Center
} Omaha, NE 68198-6395
} 402-559-7347
} tbargar-at-unmc.edu
}
}
} ==============================Original
} Headers==============================
} 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39 2007
} 8, 20 -- Received: from zixvpm02.unmc.edu (zixvpm02.unmc.edu
} [192.198.54.127])
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} 15:39:38 -0500 (CDT)
} 8, 20 -- Subject: Help with enhancing contrast of Mitochondrial
} cristae
} 8, 20 -- To: Microscopy-at-MSA.Microscopy.Com
} 8, 20 -- X-Mailer: Lotus Notes Release 7.0.1 January 17, 2006
} 8, 20 -- Message-ID: {OFDC8670F1.FEB83264-ON862572CE.
} 006F5771-862572CE.00717D6E-at-unmc.edu}
} 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu}
} 8, 20 -- Date: Tue, 1 May 2007 15:39:27 -0500
} 8, 20 -- X-MIMETrack: Serialize by Router on UNMCNOTES01.UNMC.EDU/
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}
}

Dorothy Sorenson
Microscopy and Image-analysis Laboratory
Department of Cell and Developmental Biology
University Of Michigan Medical School
A807 BSRB
109 Zina Pitcher Place
Ann Arbor, MI 48109-2200
(734)763-1170
FAX (734)763-1166



==============================Original Headers==============================
9, 19 -- From dsoren-at-umich.edu Wed May 2 10:52:50 2007
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9, 19 -- Subject: Re: [Microscopy] Help with enhancing contrast of Mitochondrial cristae
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==============================End of - Headers==============================




From: TindallR-at-missouri.edu
Date: Wed, 2 May 2007 11:46:18 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Along this same line of thought, if you have access to a
variable-wattage laboratory microwave suitable for histo- or EM
processing, you can drastically shorten your dehydration solvent
exposure times to seconds, rather than half an hour or more.

Processing times for all steps are greatly reduced by using microwaves.
It is possible to go from fresh sample to polymerized blocks in 4-5
hours, or sometimes less, with often superior results compared to
conventional processing. Extraction of sample components is minimized
by the short exposures to the various reagents, especially in
dehydration steps.

You can find our "generic" microwave protocol at
http://www.emc.missouri.edu/Pdfs/General%202-ME%20Microwave%20Processing
%20Protocol.pdf. Like most everything else in EM work, it gets modified
all the time, but it's a good starting point.

Good luck,
Randy

Randy Tindall
Senior EM Specialist
Electron Microscopy Core Facility---We Do Small Well!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu
On-line calendar:
http://biotech.rnet.missouri.edu/cgi-bin/Calcium310.pl?Op=Splash&Amount=
Week&NavType=Both&Type=TimePlan


-----Original Message-----
X-from: dsoren-at-umich.edu [mailto:dsoren-at-umich.edu]
Sent: Wednesday, May 02, 2007 10:55 AM
To: Tindall, Randy D.

Dear Tom,

It is possible that you are losing your cellular membranes during the
dehydration steps. First, en bloc stain with saturated uranyl acetate
in water and then go through the dehydration steps quickly, about 2
minutes for each step. Begin with 50 % EtOH and end with only one
change of 100 % EtOH. If you are infiltrating with EtOH:
araldite, keep the steps with high EtOH content short, too, about a 30
minute maximum.

I hope that helps!

Dotty Sorenson

On May 1, 2007, at 4:44 PM, tbargar-at-unmc.edu wrote:

}
}
}
} ----------------------------------------------------------------------
} ------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America To Subscribe/Unsubscribe -- http://www.microscopy.com/
} MicroscopyListserver On-Line Help
} http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------
} ------
}
}
} Dear listers,
}
} Anyone out there have any advice on how to enhance the contrast and
} definition of the membranes of the cristae of mitochondria? The
} samples brought to me are monolayer cell cultures of cancer cells
} grown on Thermanox coverslips. This is how I'm currently processing
} the
} samples:
} Primary fixation is 2% glutaraldehyde, 2% paraformaldehyde, 0.5%
} acrolein in 0.1M Sorensen's phosphate buffer pH 7.2. Post-fixation in

} 1%Osmium Tetroxide in 0.1M Sorensen's phosphate buffer. Dehydration
} in 50, 70, 90, 95, 100%X3 ethanol solutions. embedding in Araldite.
} Sections are stained with 2% uranyl acetate aqueous 15 minutes and
} Reynold's lead citrate 10
} minutes. The density of the cytoplasm and mitochondrial matrix are
} similar with the result that the contrast of the mitochondria is
} similar to the cytoplasm. The mitochondria and it's membranes (outer
} and that of the
} cristae) don't really "stand out". The researchers involved want to
} see really contrasty mitochondrial cristae.
}
} The next thing I'm going to try is post-fixation with a mix of osmium
} tetroxide and potassium ferrocyanide.
}
} Anyone have any other ideas? Thanks in advance.
}
}
} Tom Bargar
} University of Nebraska Medical Center
} Core Electron Microscopy Research Facility
} 986395 Nebraska Medical Center
} Omaha, NE 68198-6395
} 402-559-7347
} tbargar-at-unmc.edu
}
}
} ==============================Original
} Headers==============================
} 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39 2007 8, 20 --
} Received: from zixvpm02.unmc.edu (zixvpm02.unmc.edu
} [192.198.54.127])
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ESMTP
} id l41KddQH023357
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} 15:39:38 -0500 (CDT)
} 8, 20 -- Subject: Help with enhancing contrast of Mitochondrial
} cristae 8, 20 -- To: Microscopy-at-MSA.Microscopy.Com 8, 20 -- X-Mailer:
} Lotus Notes Release 7.0.1 January 17, 2006 8, 20 -- Message-ID:
} {OFDC8670F1.FEB83264-ON862572CE.
} 006F5771-862572CE.00717D6E-at-unmc.edu}
} 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu} 8, 20 -- Date: Tue, 1
} May 2007 15:39:27 -0500 8, 20 -- X-MIMETrack: Serialize by Router on
} UNMCNOTES01.UNMC.EDU/ Servers/UNEBR at 05/01/2007 03:39:38 8, 20 --
} PM 8, 20 -- MIME-Version: 1.0 8, 20 -- Content-type: text/plain;
} charset=US-ASCII ==============================End of -
} Headers==============================
}
}

Dorothy Sorenson
Microscopy and Image-analysis Laboratory Department of Cell and
Developmental Biology University Of Michigan Medical School
A807 BSRB
109 Zina Pitcher Place
Ann Arbor, MI 48109-2200
(734)763-1170
FAX (734)763-1166



==============================Original
Headers==============================
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==============================Original Headers==============================
21, 25 -- From TindallR-at-missouri.edu Wed May 2 11:46:18 2007
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From: nairvinods-at-gmail.com
Date: Wed, 2 May 2007 13:52:26 -0500
Subject: [Microscopy] "Rockefeller University seeks Manager for Electron Microscopy unit"

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello,
This is a posting for one of my friends, so please do not respond to me.
Thanks,
Vinod



The Rockefeller University, a world-renowned center for research and
graduate education in the biomedical sciences, seeks a Manager of
Electron Microscopy to join the Bio-Imaging Resource Center: see
http://www.rockefeller.edu/bioimaging/. The position will involve the
day-to-day management and running of the EM service and the promotion
of new technologies offered by the center. The successful applicant
will work alongside the Director of the BIRC, who oversees the optical
microscopy center directly and provides general oversight of the EM
service.

We have recently expanded the range of equipment and capabilities of
the EM service to include modern, low-temperature technologies.
Current equipment includes:
• An FEI Tecnai G2 Spirit BioTwin TEM with Gatan 4K x 4K digital camera;
• A LEO 1550 SEM with field-emission electron gun;
• Two JEM 100x TEMs (conventional film);
• Leica UC6b/EM FC6 cryoultramicrotome and Reichert Ultra Cut E ultramicrotomes;
• Leica EMPACT2 high pressure freezing system with rapid-transfer
loading device;
• Leica AFS freeze-substitution system.

We are seeking a dynamic candidate with excellent communication
skills, an enthusiastic approach towards new techniques, motivation to
maintain a broad knowledge of state-of-the-art EM technology and the
flexibility to interact with a diverse group of researchers.
Candidates should have significant research experience using EM
techniques with a PhD (preferred) or Master's degree in Biology or a
related field. Familiarity with immunolabeling techniques would be
highly advantageous.

Responsibilities will include:
• Applying EM techniques to research questions on a wide range of
biological specimens, including viruses, bacteria, cultured cells, and
animal and plant tissues;
• Day-to-day management and financial oversight of the EM service;
• The selection, installation and maintenance of new high-end
equipment to ensure that the University is constantly at the forefront
of imaging technology;
• Education, outreach and instruction of users;
• Attending national and international meetings to keep up-to-date
with current technologies;
• Striving for maximum efficiency and user-friendliness of the EM service.

The Rockefeller University is located on a beautiful campus on
Manhattan's Upper East Side and offers an excellent benefits package,
tuition reimbursement and a competitive salary.

Please click the URL below to apply for this job:
http://www.rockefeller.edu/hr/jobs.php?url=https://recruit.rockefeller.edu/OA_HTML/OA.jsp?OAFunc=IRC_VIS_VAC_DISPLAY%26p_svid=2844%26p_spid=3068%26p_site_id=21

The Rockefeller University is an Affirmative Action/VEVRAA/Equal
Opportunity Employer.

Please feel free to contact Dr. Alison North, the Director of the
BIRC, at northa-at-rockefeller.edu for further information or with
specific questions.


==============================Original Headers==============================
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From: larry.ackerman-at-ucsf.edu
Date: Wed, 2 May 2007 14:24:09 -0500
Subject: [Microscopy] Re: Labware washers

Contents Retrieved from Microscopy Listserver Archives
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Randy,
I had good luck with a Hotpack, model # H-1125 in another lab. Very
reliable and good manufacturer support if needed. Measure your space
very carefully! Some other units were 3/4" too tall to fit under the lab
top.
Larry

TindallR-at-missouri.edu wrote:
} ----------------------------------------------------------------------------
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} Dear Listers,
}
} Does anyone have a recommendation for a laboratory glassware washer that
} will fit under a counter (about like a home dishwasher size), handle
} light duty of two or three loads a week, wash labware to a level
} acceptable for EM use, and, ahem, not break the bank? Does something
} like this exist for $5000 or thereabouts? Am I delusional? (Don't
} answer that......)
}
} We currently have users privileges on a really nice huge heavy duty lab
} dishwasher, but the day is coming when this will end. Also, we don't
} always have access exactly when we most need it. We want our own!
}
} Thanks in advance. Vendor replies are most welcome.
}
} Cheers,
} Randy
}
} Randy Tindall
} Senior EM Specialist
} Electron Microscopy Core Facility---We Do Small Well!
} W125 Veterinary Medicine
} University of Missouri
} Columbia, MO 65211
} Tel: (573) 882-8304
} Fax: (573) 884-2227
} Email: tindallr-at-missouri.edu
} Web: http://www.emc.missouri.edu
} On-line calendar:
} http://biotech.rnet.missouri.edu/cgi-bin/Calcium310.pl?Op=Splash&Amount=
} Week&NavType=Both&Type=TimePlan
}
}
} ==============================Original Headers==============================
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}

--
Larry Ackerman, Associate Specialist
UCSF, Dept. of Anatomy, Rm S1347
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu


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From: Robert.Zonis-at-sanford.com
Date: Wed, 2 May 2007 15:14:46 -0500
Subject: [Microscopy] Optical Microscopy - Objectives for Nikon Eclipse ME600L

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear listers,

Sorry if this is a double post. I am looking for 2X and 4X CFI Plan Achromat Objectives for a Nikon Eclipse ME600. 1X Achromat and/or 2X or 4X Apochromats would be great if they were affordable (under $1,000 each) but I won't hold my breath waiting for that. Does anyone have or know of these objectives, new or used, in stock and for sale anywhere? Nikon USA does not have them in stock, neither does our Nikon dealer, and we need to get at least one of these objectives fairly soon - within a week if possible.

The low resolution is so that I can increase the field of view for digital image analysis. If anyone knows of another way to accomplish this, or knows of third-party suppliers that make objectives that fit the Nikon, I'd love to hear about it.

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


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From: Konrad.Jarausch-at-hitachi-hta.com
Date: Wed, 2 May 2007 15:47:49 -0500
Subject: [Microscopy] Gas reaction chemistry in a "large" chamber SEM?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

RE: Gas reaction chemistry in a large-chamber SEM?

I am looking for people who have found creative ways to do
gas-reaction chemistry in a "large" chamber SEM, ideally with a heating
stage, gas injection and better control of the vacuum/cleanliness
(near-UHV). Ideas about chamber within a chamber designs, better
cleaning/pumping for the large chamber and/or experience with
gas-injection in an environmental SEM would be most welcome! I am
familiar with the designs for TEM/STEM, but am looking for ideas based
on SEMs. Please feel free to respond either directly or include the
distribution. Thank you in advance for your suggestions!

konrad


Disclaimer: i work for the Hitachi electron microscope division so
please do NOT disclose any confidential or proprietary information, only
open domain discussion!!



==============================Original Headers==============================
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From: kraftpiano-at-gmail.com
Date: Wed, 2 May 2007 18:50:48 -0500
Subject: [Microscopy] SEM Sample prep advice.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I've started getting information from the kids about what kind of
samples they want to look at using our SEM when it is up and running,
and one student gave me a bit of a baffler. I'm generally familiar
with various sample prep techniques for different samples of fibers,
biologicals, non-organic materials, etc., but this one stumped me.

The student read somewhere that spider webs achieve their elasticity
by small bundles of fibers inside the sticky "glue" on the web. They
want to see this first hand. Any suggestions on how to prepare a
spider web? There is no shortage of them around here.

--Justin A. Kraft
Atlantis Academy
West Palm Beach, FL

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From: jgsheridanmicroscopy-at-gmail.com
Date: Wed, 2 May 2007 18:58:47 -0500
Subject: [Microscopy] viaWWW: ES vision software

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This Question/Comment was submitted to the Microscopy Listserver
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Email: jgsheridanmicroscopy-at-gmail.com
Name: John Sheridan

Organization: TCD

Title-Subject: [Filtered] EDX: ES vision software

Question: Hi all!

Does anyone know who makes "ES vision" , it's a piece of software for EDX analysis. Google has not helped me so far.

Many thanks,

John


---------------------------------------------------------------------------

==============================Original Headers==============================
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From: rra-at-stowers-institute.org
Date: Wed, 2 May 2007 18:59:15 -0500
Subject: [Microscopy] viaWWW: Mitochondrial Staining

Contents Retrieved from Microscopy Listserver Archives
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Email: rra-at-stowers-institute.org
Name: Rhonda Allen

Organization: Stowers Institute

Title-Subject: [Filtered] Mitochondrial Staining

Question: I am sure someone else can elaborate more than I, but Hyatt state that Phosphotungstic Acid after glutaraldehyde fixation in an aqueous acidic medium intensely stains mitochoindrial matix, cisternae of er, and others.

This is from "Principles and Techniques of Electron Microscopy", 4th edition, page 313.


Rhonda Allen

---------------------------------------------------------------------------

==============================Original Headers==============================
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From: tchallman-at-case4n6.com
Date: Wed, 2 May 2007 18:59:49 -0500
Subject: [Microscopy] viaWWW: Hitachi SEM 2460N Service Contract

Contents Retrieved from Microscopy Listserver Archives
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Email: tchallman-at-case4n6.com
Name: Tracy Challman

Organization: CASE Forensics Corp

Title-Subject: [Filtered] Hitachi SEM 2460N Service Contract

Question: Once again, we are looking for qualified and experienced people to provide an alternative to Hitachi's SEM Service Contract - annual preventative maintenance and occassional mishaps in the electronics. Do you recommend anyone, particularly in the Pacific Northwest?

All suggestions are welcome.

Thank you,
Tracy

---------------------------------------------------------------------------

==============================Original Headers==============================
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8, 11 -- From: tchallman-at-case4n6.com (by way of MicroscopyListserver)
8, 11 -- Subject: viaWWW: Hitachi SEM 2460N Service Contract
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From: robert.zonis-at-sanford.com
Date: Wed, 2 May 2007 19:00:15 -0500
Subject: [Microscopy] viaWWW: Help with optical objectives

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Email: robert.zonis-at-sanford.com
Name: Robert Zonis

Organization: Sanford

Title-Subject: [Filtered] Help with optical objectives

Question: Does anybody know where I can get Nikon CFI Plan Achromat objectives in UW 2x or 4x relatively quickly? I'm doing some image analysis and I need a wider field of view.

Alternatively, does anyone know if there's a third-party supplier of objectves for Nikons?

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. ñ A Newell Rubbermaid Company
Liquid Manufacturing and Technology Center
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613


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From: cgarber-at-2spi.com
Date: Wed, 2 May 2007 19:52:25 -0500
Subject: [Microscopy] ES vision?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Email: jgsheridanmicroscopy-at-gmail.com
John Sheridan wrote:
==================================================
Title-Subject: [Filtered] EDX: ES vision software

Question: Hi all!

Does anyone know who makes "ES vision" , it's a piece of software for EDX analysis. Google has not helped me so far.
=================================================
Are you thinking about the Electron Flight Simulator (EFS) software used in conjuction with EDS, see URL
http://www.2spi.com/catalog/software/efs.html

Chuck


===================================================
Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
President
SPI SUPPLIES FAX: 1-(610)-436-5755
PO BOX 656 e-mail: cgarber-at-2spi.com
West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com


Look for us!
############################
WWW: http://www.2spi.com
############################
==================================================

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From: zaluzec-at-microscopy.com
Date: Wed, 2 May 2007 20:32:23 -0500
Subject: [Microscopy] ES Vision

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


John

ESVision was originally written by a company called EMISpec.

EMISpec was acquired by FEI in 2003.


Nestor
Your Friendly Neighborhood SysOp

==============================Original Headers==============================
6, 11 -- From zaluzec-at-microscopy.com Wed May 2 20:32:22 2007
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6, 11 -- From: "Nestor J. Zaluzec" {zaluzec-at-microscopy.com}
6, 11 -- Subject: ES Vision
6, 11 -- Content-Type: text/plain; charset="us-ascii"
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From: protrain-at-emcourses.com
Date: Thu, 3 May 2007 06:06:37 -0500
Subject: [Microscopy] Optical Microscopy

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

All the fuss over microprobes was fun but if you want to get your teeth into
names used by scientists (?) what about "optical microscopy"?

With my 43 years in the business I have always taught that whilst light
microscopes have "light optics" electron microscopes have "electron optics",
the principles are the same. So when I see a piece about optical
microscopes I always look to see if it should be of interest to me?

What is the general thought?

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


==============================Original Headers==============================
6, 26 -- From protrain-at-emcourses.com Thu May 3 06:06:37 2007
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From: Mike.Bode-at-olympus-sis.com
Date: Thu, 3 May 2007 06:21:28 -0500
Subject: [Microscopy] ES Vision

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Unless you meant ESIVision (with an "I") for EELS imaging, in which case
you can contact me.

mike

Michael Bode

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS CORP.
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-Mail: Mike.Bode-at-olympus-sis.net
www.olympus-sis.net

-----Original Message-----
X-from: zaluzec-at-microscopy.com [mailto:zaluzec-at-microscopy.com]
Sent: Wednesday, May 02, 2007 19:37
To: Mike Bode


John

ESVision was originally written by a company called EMISpec.

EMISpec was acquired by FEI in 2003.


Nestor
Your Friendly Neighborhood SysOp

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15, 25 -- From Mike.Bode-at-olympus-sis.com Thu May 3 06:21:27 2007
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From: frank.karl-at-degussa.com
Date: Thu, 3 May 2007 07:00:42 -0500
Subject: [Microscopy] Re: Optical Microscopy

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Guilty!

Having worn glasses since eight grade, I have come to associate optical
with glass the photons.

Phototonic microscopy? sounds like a title of a paper in search of a
little primping
Non-modified bright field? - sounds like a title of a paper in search of a
little primping from a major university
Light microscopy? - sort of suggest their might be a heavy microscopy
Refractory microscopy - sound like microscopy in hell
Specular microscopy - the study of the stock market in great detail..

Tongue firmly implanted in cheek: Let's call it Bruce. I got the idea
from a microscopy society bog (
http://www.msneo.org/2006/06/blog-mail.html)
who knows..........

Stay safe..............Frank




protrain-at-emcourse
s.com To: frank.karl-at-degussa.com
cc:
05/03/2007 07:09 Subject: [Microscopy] Optical Microscopy
AM
Please respond to
protrain








----------------------------------------------------------------------------

The Microscopy ListServer -- CoSponsor: The Microscopy Society of America


Hi

All the fuss over microprobes was fun but if you want to get your teeth
into
names used by scientists (?) what about "optical microscopy"?

With my 43 years in the business I have always taught that whilst light
microscopes have "light optics" electron microscopes have "electron
optics",
the principles are the same. So when I see a piece about optical
microscopes I always look to see if it should be of interest to me?

What is the general thought?

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


==============================Original
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From: David.Patton-at-uwe.ac.uk
Date: Thu, 3 May 2007 07:33:53 -0500
Subject: [Microscopy] Re: Optical Microscopy

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Light Microscopy? I once got an assignment where the student thought
this was a special tool to study the activity of chlorophyll!

Dave

-----Original Message-----
X-from: frank.karl-at-degussa.com [mailto:frank.karl-at-degussa.com]
Sent: 03 May 2007 13:05
To: David Patton

Guilty!

Having worn glasses since eight grade, I have come to associate optical
with glass the photons.

Phototonic microscopy? sounds like a title of a paper in search of a
little primping
Non-modified bright field? - sounds like a title of a paper in search
of a
little primping from a major university
Light microscopy? - sort of suggest their might be a heavy microscopy
Refractory microscopy - sound like microscopy in hell
Specular microscopy - the study of the stock market in great detail..

Tongue firmly implanted in cheek: Let's call it Bruce. I got the idea
from a microscopy society bog (
http://www.msneo.org/2006/06/blog-mail.html)
who knows..........

Stay safe..............Frank





protrain-at-emcourse

s.com To:
frank.karl-at-degussa.com

cc:

05/03/2007 07:09 Subject: [Microscopy]
Optical Microscopy
AM

Please respond to

protrain










------------------------------------------------------------------------
----

The Microscopy ListServer -- CoSponsor: The Microscopy Society of
America


Hi

All the fuss over microprobes was fun but if you want to get your teeth
into
names used by scientists (?) what about "optical microscopy"?

With my 43 years in the business I have always taught that whilst light
microscopes have "light optics" electron microscopes have "electron
optics",
the principles are the same. So when I see a piece about optical
microscopes I always look to see if it should be of interest to me?

What is the general thought?

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


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==============================Original Headers==============================
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From: gas19-at-daimlerchrysler.com
Date: Thu, 3 May 2007 07:38:02 -0500
Subject: [Microscopy] viaWWW: RE: Help with optical objectives

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Email: gas19-at-daimlerchrysler.com
Name: Gerald Shulke

Organization: DaimlerChrysler

Title-Subject: [Filtered] Re: [Microscopy] viaWWW: Help with optical objectives

Question: We have bought two low magnfiication objectives for one of our Nikon microscopes. A 1.5x CF Plan BF EPI Achromat na 0.045 wd 3.6mm for roughly $3k, and a 2.5x CF BF Plan EPI Achromat na 0.075 wd 8.8 mm for roughly $1k. Both have Nikon Japan on the lens. You may try some of the Nikon distributors. Our local distributor is Mager Scientific in Dexter, MI. www.magersci.com. (734) 426-3885.

Gerald Shulke
DaimlerChrysler
Material Characterization Labs

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From: baskin-at-bio.umass.edu
Date: Thu, 3 May 2007 08:21:03 -0500
Subject: [Microscopy] Re: Optical Microscopy

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Steve,
Is it ever? I mean of interest to you? More specifically,
about electrons rather than photons? My feeling is that despite folks
talking about 'electron optics' that the term optical microscopy
always means light. Probably the irrationality of names but one could
suggest that optics means something different than optical. Optics
originally referred to photons but once it was discovered that
electrons (and refrigerators) also have wave properties, the term
optics was expanded. Wheras optical seems to have retained its photon
orientation.

My second order guess.

Tobias


}
} Hi
}
} All the fuss over microprobes was fun but if you want to get your teeth into
} names used by scientists (?) what about "optical microscopy"?
}
} With my 43 years in the business I have always taught that whilst light
} microscopes have "light optics" electron microscopes have "electron optics",
} the principles are the same. So when I see a piece about optical
} microscopes I always look to see if it should be of interest to me?
}
} What is the general thought?
}
} Steve Chapman
} Senior Consultant Protrain
} For electron microscopy consultancy and training world wide
} Tel +44 1280 816512 Fax +44 1280 814007
} Mobile +44 7711 606967 Web www.emcourses.com
}

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Biology Department
/_ / __ /__ \ \ \__ 611 N. Pleasant St.
/ / / \ \ \ University of Massachusetts
/ / / \ \ \ Amherst, MA, 01003
/ / ___ / \ \__/ \ ____
http://www.bio.umass.edu/biology/baskin/
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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From: ph2-at-sprynet.com
Date: Thu, 3 May 2007 10:36:28 -0500
Subject: [Microscopy] Optical Microscopy

Contents Retrieved from Microscopy Listserver Archives
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Agreed.

For kicks, here's the definition in

Glossary of Microscopical Terms and Definitions, 2nd Ed., NY Microscopical
Society, 1989.


Optical microscope. Very ambiguous term since all microscopes involve
optics; better to specify light, acoustic, x-ray or electron microscope,
etc.


Also, for Frank, from the foreword:

"Words, like eyeglasses, blur everything that they do not make clear."
- Joseph Joubert (1754-1824)



Tony

Ps Microprobe is not in the glossary

Pps Glasses since 4th grade.


......................................................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
pH2, LLC
5250 E US 36, Suite 830
Avon, IN 46123
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-----Original Message-----
X-from: protrain-at-emcourses.com [mailto:protrain-at-emcourses.com]
Sent: Thursday, May 03, 2007 7:13 AM
To: ph2-at-sprynet.com

Hi

All the fuss over microprobes was fun but if you want to get your teeth into

names used by scientists (?) what about "optical microscopy"?

With my 43 years in the business I have always taught that whilst light
microscopes have "light optics" electron microscopes have "electron optics",

the principles are the same. So when I see a piece about optical
microscopes I always look to see if it should be of interest to me?

What is the general thought?

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


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From: donald-at-uwm.edu
Date: Thu, 3 May 2007 10:45:22 -0500
Subject: [Microscopy] Re: viaWWW: ES vision software

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Good Morning John,

You may be referring to ES Vision developed by Emispec several years
ago at the State University of Arizona, and now part of the FEI
organisation.

Try: ESVisionSupport-at-FEICO.com

This is from a communication I received from them last year.

Good Luck,

Donald Robertson

/////////////////////////////////////////////////
From: ESVisionSupport-at-FEICO.com
Subject: RE: Support service for ES Vision
Date: July 28, 2006 1:06:43 PM CDT
To: donald-at-uwm.edu

Hi Donald,

The support email address you used for this message is the best way to
obtain support for your ES Vision system. Please forward your questions
or problems to this address and we will respond as soon as possible.

Best Regards,

ES Vision Support

/////////////////////////////////////

On May 2, 2007, at 7:01 PM, jgsheridanmicroscopy-at-gmail.com wrote:

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}
} Email: jgsheridanmicroscopy-at-gmail.com
} Name: John Sheridan
}
} Organization: TCD
}
} Title-Subject: [Filtered] EDX: ES vision software
}
} Question: Hi all!
}
} Does anyone know who makes "ES vision" , it's a piece of software
} for EDX analysis. Google has not helped me so far.
}
} Many thanks,
}
} John

Donald Robertson
Sr. Instrumentation Specialist
HRTEM Lab
Department of Physics
College of Letters and Science
University of Wisconsin - Milwaukee
tel: (414) 229 2753




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From: jquinn-at-www.matscieng.sunysb.edu
Date: Thu, 3 May 2007 11:21:13 -0500
Subject: [Microscopy] re: optical microscopy

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Folks

ASTM standard E175(2005)
Standard Terminology of Microscopy

} re: Optical microscope.
} Very ambiguous term since all microscopes involve
} optics; better to specify light, acoustic, x-ray or
} electron microscope, etc.
}
}
}

regards,

JQuinn



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From: bozzola-at-siu.edu
Date: Thu, 3 May 2007 15:18:38 -0500
Subject: [Microscopy] TEM: Hitachi H7650 question

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I have some questions for users of the new Hitachi H7650 TEM.

If you could contact me offline, I would greatly appreciate it.

Thank you.

JB
--
##############################################################
John J. Bozzola, Ph.D., Director
I.M.A.G.E. (Integrated Microscopy & Graphics Expertise)
750 Communications Drive - MC 4402
Southern Illinois University
Carbondale, IL 62901 U.S.A.
Phone: 618-453-3730
Email: bozzola-at-siu.edu
##############################################################

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From: tomw-at-uidaho.edu
Date: Thu, 3 May 2007 18:40:16 -0500
Subject: [Microscopy] viaWWW: Simulated EDS Spectrum

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Email: tomw-at-uidaho.edu
Name: Tom Williams

Organization: Univ. of Idaho

Title-Subject: [Filtered] Simulated EDS Spectrum

Question: One of our Geology Faculty asked me if I had software that could simulate EDS spectrums for minerals (such as a feldspars or quartz) he could use to generate a number of patterns for educational purposes (without having to actually run each sample on the SEM/EDS-itís a time-constraint issue). I have never seen software that can do this, and told him it is far from trivial. As far as I can determine, my Noran System Six software canít do this. Is anyone aware of a program that can do this??

Thanks
Tom


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From: jvtaylo-at-emory.edu
Date: Thu, 3 May 2007 18:40:57 -0500
Subject: [Microscopy] viaWWW: electron diffraction

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Email: jvtaylo-at-emory.edu
Name: Jeannette Taylor

Organization: IM&MF / Emory University

Title-Subject: [Filtered] electron diffraction

Question: We are beginning to explore the use of electron diffraction in the study of crystalized biological molucules. We don't have the background here. Some of these molecules are very fragile and disappear before a pattern can be captured on film. An electron diffractin pattern is seen, briefly, then it disappears. We have been following what the micropscope manual instructs on generating electron diffractions and have gotten some advice from a metallurgist. But we need some more help. Using purchased standards, we have gotten some very nice diffraction patterns.

Thanks for any help offered.

---------------------------------------------------------------------------

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From: drix00-at-gmail.com
Date: Thu, 3 May 2007 19:30:41 -0500
Subject: [Microscopy] Re: viaWWW: Simulated EDS Spectrum

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Tom,

You can use a free Monte Carlo program to simulate the EDS spectrum
like WinX-Ray: http://montecarlomodeling.mcgill.ca/download/download.html

If you have a mac with a PowerPC CPU you can use DTSA software:
http://www.cstl.nist.gov/div837/Division/outputs/DTSA/DTSA.htm

I know a couple of other software, but you need to do some coding to used them.

A nice list of Monte Carlo softwares can be found on the NISTMonte web
page by N. Ritchie:
http://www.cstl.nist.gov/div837/837.02/epq/index.html

Usual disclaimer: I am the author of WinX-Ray and I have use and test
most of the free softwares mentioned in this post.

Good luck,
Hendrix

On 5/3/07, tomw-at-uidaho.edu {tomw-at-uidaho.edu} wrote:
} Email: tomw-at-uidaho.edu
} Name: Tom Williams
}
} Organization: Univ. of Idaho
}
} Title-Subject: [Filtered] Simulated EDS Spectrum
}
} Question: One of our Geology Faculty asked me if I had software that could simulate EDS spectrums for minerals (such as a feldspars or quartz) he could use to generate a number of patterns for educational purposes (without having to actually run each sample on the SEM/EDS-itís a time-constraint issue). I have never seen software that can do this, and told him it is far from trivial. As far as I can determine, my Noran System Six software canít do this. Is anyone aware of a program that can do this??
}
} Thanks
} Tom
}
}
} ---------------------------------------------------------------------------
}
}


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From: walck-at-southbaytech.com
Date: Thu, 3 May 2007 19:47:08 -0500
Subject: [Microscopy] viaWWW: Simulated EDS Spectrum

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Tom,

First, you need a Mac. It might have to be an older Mac.

Then visit the following website,
http://www.cstl.nist.gov/div837/Division/outputs/DTSA/DTSA.htm

There you will see a complete description of DTSA which stands for Desktop
Spectrum Analyzer.

It will do exactly what you want to do.

You might want to look at my article in Microscopy Today, March 2006, on
page 34 to show examples of spectra generated by DTSA.


-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
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Email: tomw-at-uidaho.edu
Name: Tom Williams

Organization: Univ. of Idaho

Title-Subject: [Filtered] Simulated EDS Spectrum

Question: One of our Geology Faculty asked me if I had software that could
simulate EDS spectrums for minerals (such as a feldspars or quartz) he could
use to generate a number of patterns for educational purposes (without
having to actually run each sample on the SEM/EDS-itís a time-constraint
issue). I have never seen software that can do this, and told him it is far
from trivial. As far as I can determine, my Noran System Six software canít
do this. Is anyone aware of a program that can do this??

Thanks
Tom


---------------------------------------------------------------------------


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From: tivol-at-caltech.edu
Date: Thu, 3 May 2007 19:50:17 -0500
Subject: [Microscopy] Re: viaWWW: electron diffraction

Contents Retrieved from Microscopy Listserver Archives
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On May 3, 2007, at 4:41 PM, jvtaylo-at-emory.edu wrote:

} We are beginning to explore the use of electron diffraction in the
} study of crystalized biological molucules. We don't have the
} background here. Some of these molecules are very fragile and
} disappear before a pattern can be captured on film. An electron
} diffractin pattern is seen, briefly, then it disappears. We have been
} following what the micropscope manual instructs on generating electron
} diffractions and have gotten some advice from a metallurgist. But we
} need some more help. Using purchased standards, we have gotten some
} very nice diffraction patterns.

Dear Jeannette,
You will need to work in low dose conditions. If you are using film,
I highly recommend the most sensitive film available, and such films as
LoDose or MRF32 X-ray films are about 20 times more sensitive than
SO163. The only problem with these is that they must be handled in
total darkness, so it will be difficult to cut them to size. Loading
and unloading the camera, loading the racks, and developing the film
are also somewhat difficult, but the skill to do that will come after a
few times doing those tasks. Another concern is static electricity,
which will make very interesting patterns on the developed film, but
these will destroy the ED data. The simplest way to get good data is
to insert the selected area aperture that is the right size for the
crystals you are looking at, put the scope in diffraction mode, use a
very small condenser aperture, a high spot size, and underfocus the
beam until you have parallel illumination, then scan the grid in
diffraction mode, either looking for a good pattern or defocussing the
beam and looking for the distorted image of the crystal in the
zero-order spot. Blank the beam, set the lenses to the proper values
to obtain a spot pattern (if you were searching in defocussed
diffraction) start the exposure, and turn on the beam only when the
shutter opens. This happens automatically for some microscopes, but
for those that do not have a pre-specimen shutter, you must do it
yourself. Good luck.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu


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From: walck-at-southbaytech.com
Date: Thu, 3 May 2007 21:07:19 -0500
Subject: [Microscopy] viaWWW: electron diffraction

Contents Retrieved from Microscopy Listserver Archives
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Just a few comments off hand without ever doing these types of samples.

Your problem will be similar to what happened to me with glass samples. You
are putting too much energy into your sample and it is heating. There are
several things you can do to help. You have to do things quickly, i.e. low
dosage. You have to lower the amount of energy you are putting into the
sample. The best way to do this is going to higher accelerating energy.
Remember, diffraction is elastic and you don't loose energy in the sample
via that route. If your energy is lower, you have more inelastic scattering
and you are dumping energy into the sample with those types of scattering
events. You probably are working on a 100 or 120 kV machine, but you didn't
say so. And another thing that you can do is to cool your sample and use a
liquid nitrogen stage.

Having said this, I don't have any idea what the higher energy will do to
your sample in terms of radiation damage.


-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
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Sent: Thursday, May 03, 2007 4:45 PM
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Email: jvtaylo-at-emory.edu
Name: Jeannette Taylor

Organization: IM&MF / Emory University

Title-Subject: [Filtered] electron diffraction

Question: We are beginning to explore the use of electron diffraction in the
study of crystalized biological molucules. We don't have the background
here. Some of these molecules are very fragile and disappear before a
pattern can be captured on film. An electron diffractin pattern is seen,
briefly, then it disappears. We have been following what the micropscope
manual instructs on generating electron diffractions and have gotten some
advice from a metallurgist. But we need some more help. Using purchased
standards, we have gotten some very nice diffraction patterns.

Thanks for any help offered.

---------------------------------------------------------------------------

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From: nizets2-at-yahoo.com
Date: Fri, 4 May 2007 04:07:18 -0500
Subject: [Microscopy] TEM : objective aperture and EDX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Deal all!

I have a conceptual problem with my TEM (tecnai G20
(twin)). I asked help to 2 engineers from FEI, they
gave me opposite answers (!!).
Here is my problem:
The objective aperture is situated just under the
specimen itself. So it should have no influence on the
intensity of the beam hitting my sample.

In this case why does my formvar film dilate and
sometimes blow up when I forget to insert the
objective aperture but is extremely stabile when it is
inserted?

Even more strange: the EDX detector is placed at an
angle several degrees ABOVE the specimen. I cannot
detect anything when the objective aperture is
inserted, I must remove it to be able to read
something!

I have the feeling that actually the obj aperture is
at the same height as the specimen and not under it,
but one engineer told me it was just under the
specimen. Also, I dont understand why the formvar is
sensible to the presence of the obj aperture if it is
at the same height as my specimen.


Stephane

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
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8, 19 -- Subject: TEM : objective aperture and EDX
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From: keith.morris-at-ucl.ac.uk
Date: Fri, 4 May 2007 04:52:24 -0500
Subject: [Microscopy] AskAMicroscopist: Building Criteria for Optical

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Dear Sandra,

Sorry but I had missed the 'Ask-A-Miicroscpist' on the header and assumed
you were on the list-server (so I didn't email you my reply). I attach my
original musings on the subject of vibration feedback and optical
microscopes below, i.e. the microscope rooms don't really have to have
special modifications at all for light microscopy provided stout worktops
and/or standard microscope anti-vibrational systems are used as supplied my
microscope manufacturers and their support industries.

It's only high magnification electron microscopes (EM) that go to 20,000x
magnification that might benefit from more advanced modification of the room
itself for anti-vibration (and these EM devices are too big and heavy to be
used with light microscope anti-vibration systems) - but I'm mainly a light
microscopist these days and not really the person the ask about EMs, you
need an EM microscopist for this (besides it's not in your brief).

In the old days optical microscope support worktops were made of heavy
damping materials like granite and slate and these work well, but you still
need some form of rubber decoupling material as well, e.g. another heavy
plate supporting the microscope and an anti-vibrational rubber type material
underneath like squash balls or stacks of Fabreeka FabCel sheets to isolate
the microscope (plus the little £5 sticky rubber feet on the microscope help
quite a bit). Modern laminated worktops are fine provided they are 5cm+
thickness and well supported - see photos cheap £500 (bad) and
micro-dissection (much better).

For expensive confocal or live cell time-lapse microscopes the best option
is often a stand-alone air-table (powered by compressed air from a £500
electric pump). These cost around £5,000 for a standard design, which in
comparison to the £200,000 for the confocal microscope isn't that much. The
air table needs no special room modification, just a space for it and its
£500 air pump. It's a standalone device that can move with the microscope to
another room is needed. For a simple £10,000 laboratory upright microscope
used by students for fixed slides the sticky rubber feet on the microscope
are generally enough (a granite/slate support worktop might be nice though,
but a standard thick heavy laminated wooden 'woodchip' one will be fine).

My comments on the rooms air conditioning, below, are only critically
important if you wish to do time-lapse studies on the movement of living
cells (where focus and XY stage drift totally ruin the time-lapse video). It
can also affect a motorised automatic scan of a slide. However if you are
manually looking at slides or fixed samples under the microscope variation
in room temperature is less important, as you will simply refocus the optics
or move the stage when it slowly drifts - if you notice it at all (the
thermal changes being quite slow). You will still want the temperature to be
comfortable for long periods of sitting though (about 22oC), and the worktop
should be right height for comfortable microscope viewing with a fully
adjustable chair.

A few examples of Vibration isolation Air-tables
http://www.technicalmanufacturing.com/portals/lifescience.html
http://www.kineticsystems.com/page135.html
http://www.speirsrobertson.com/
http://www.nextdayscience.com/store/laboratory/anti-vibration-tables.html

Rubber isolation examples
http://www.fabreeka.com/products/fabcel_pads.htm
Some Fabcel can be seen in cheap £500 (brown in this case, but the black
looks nicer) plus there are squashballs, anti-vibration rubber feet
etc...there are loads of different types about see http://www.rswww.com and
search anti vibration.

So generally you select the type of anti-vibrational microscope support you
need, allowing for cost and whether you need high magnification for live
cell work (the culture media and living cells wobble far more than fixed
slides and need better support - fixed slides & microscopes tend to wobble
together in unison so you don't notice it so much). Other than stout
worktops most anti-vibrational devices are added after the room is built and
so can be moved to another room with the microscope. In all cases our
microscopes are used in standard laboratories with concrete floors although
they have the room to themselves.

A few examples of the anti-vibration tables/plates we use for microscopes
are attached. Note the very flimsy worktop under the cheap £500 black
in-house manufactured damping plate, and to be honest we only needed the
damping isolation plate with this microscope for our live cell work, not for
fixed slides - the microscope had been used for 5 years previously just
sitting on the worktop. The laser dissection microscope needs high
magnifications to uv laser cut out individual cells and even chromosomes
from fixed slides, but this was supplied by Zeiss with a simple steel plate
and rubber feet for 'anti-vibration' which you can see resting on the black
worktop (a better worktop than in the 'cheap £500' photo). This is all
Ziess/PALM recommend for installation (and it works fine).

My original list-server message is attached below

Hope this is all OK, you probably know a lot of this anyway but any queries
and just ask. Sorry you didn't get this earlier.

Keith

PS. Only Sandra gets the photo's I'm afraid (four photos showing a Leica SP2
confocal with a £3,000 Leica granite/squashball isolation table, a Bio-Rad
[Zeiss Microscience] Radiance confocal with a £5,000 air table, a PALM/Zeiss
Axiovert 200 micro-dissection system with PALM worktop damping isolation
plate, and our cheap £500 Fabcel/140kg plate used with a Zeiss Axiovision
100M/OpenLabs time-lapse system.

---------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology
Institute of Ophthalmology
UCL, London EC1V 9EL



-----Original Message-----
X-from: Keith Morris [mailto:keith.morris-at-ucl.ac.uk]
Sent: 02 May 2007 09:50
To: 'microscopy-at-microscopy.com'

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (sandra.filippi-at-montgomerycollege.edu) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday,
April 30, 2007 at 20:12:50
---------------------------------------------------------------------------

Email: sandra.filippi-at-montgomerycollege.edu
Name: Sandra Lee Filippi, Campus Planner

Organization: montgomery college

Education: Graduate College

Location: Rockville MD

Question:


Dear Sir or Madam Microscopist:

Montgomery College is in design development for a new academic teaching
science center. The lab planner for the design team is citing ASHRAE TC2.6
Guideline Criteria for use in Planning New Facilities and insists that we
design to the VC-B level because the biology department uses compound
microscopes with total magnification of 1,000x (oil immersion). I managed
academic science labs for a two-year college for more than 20 years. We
routinely used light microscopes at total magnification of 1,000x (oil
immersion) in a facility built in 1965 without special vibration control.
When we opened our new science center in 1995 it did not have special
vibration control either. I have studied at Georgetown University and did
student research at The NIH. Neither of these facilities required special
vibration control for a light microscope.
I have consulted with the Westover Scientific Customer Service/Product
Specialist ñ Microscopy who states, ìTo be honest with you I have never
heard of such a thing, neither have my colleagues. ÝThere are tables that
are manufactured to reduce vibrations, but Iíve never heard of a building
being designed around a microscope. ÝWe have customers who use microscopes
in old buildings all over the world. Westover recommends that a microscope
be used on a vibration free surface which typically means not locating your
microscope on a table with other equipment that can cause vibrations such as
fans and other lab equipment. Microscopes have been used for years and
years on standard benchtops and other tables without issue. It's not our
position to recommend additional support when building a new structure. In
my entire career I have never heard of this requirement.î Westover
manufactures microscopes for Fisher Scientific.
The lab planner is going to cost us dearly unless I can refute his
recommendation. What is your professional opinion in this matter? Thank
you very much for your time.
Sandra

Sandra Lee Filippi, Campus Planner
Montgomery College
Suite 200
40 West Gude Drive Room 223
Rockville MD 20850-1166
301.251.7362 vox
301.251.7379 fax
240.882.6672 cell


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From: wvrenter-at-sckcen.be
Date: Fri, 4 May 2007 07:52:13 -0500
Subject: [Microscopy] viaWWW: orientation of the diffraction

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

You have a very common problem. Firstly the objective aperture sits below
the specimen, usually in the back focal plane of the objective. The films
become damaged when you remove the aperture as the presence of the aperture
acts to neutralise the charge on the film. My guess is when you have the
objective aperture in place the x-ray signal is so great it swamps the
detector with x-rays causing the detector to go 100% dead.

Try using very small spot sizes (condenser 1 at a higher strength), or
smaller condenser apertures, or a lower emission current from the gun, all
of which will help preserve the specimen. If you still have a problem try
two of the above or even all three. The object here is to reduce the number
of electrons hitting the specimen thus reducing the damaging component. If
you still have a problem with film damage then use the largest objective
aperture that you can find, even replacing one aperture with one of the
smaller apertures used in the condenser system. I have to say that I am not
sure if your microscope uses disk apertures or an aperture strip? The idea
being that if you still need the objective aperture for stability the bigger
the hole the lower the chance of picking up its x-rays.

I hope this helps?

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com

----- Original Message -----
X-from: {nizets2-at-yahoo.com}
To: {protrain-at-emcourses.com}
Sent: Friday, May 04, 2007 10:10 AM

This Question/Comment was submitted to the Microscopy Listserver
using the WWW based Form at http://microscopy.com/MicroscopyListserver/MLFormMail.html
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Email: wvrenter-at-sckcen.be
Name: Wouter Van Renterghem

Organization: SCK-CEN

Title-Subject: [Filtered] orientation of the diffraction pattern

Question: When you defocus a diffraction pattern, you can see a small image of the selected area in the 000 spot. When you go from underfocus to overfocus the image of the selected area is rotated over 180ƒ. Can somebody tell me which of the two small images has the same orientation as the image in image mode? Do I have to underfocus the diffraction pattern or overfocus it?

Thank you,
Wouter

---------------------------------------------------------------------------


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From: microscopytoday-at-tampabay.rr.com
Date: Fri, 4 May 2007 09:52:40 -0500
Subject: [Microscopy] Microscopy Today May Table of Contents

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Listers,

Here is the May 2007 Microscopy Today table of contents. I will close
the subscription list for this issue on Wednesday, May. 9th, 2007.

Microscopists in North America and MSA members anywhere can subscribe
for free. Anyone else may subscribe for US$50 per year (to PARTIALLY
cover postage). All subscriptions at http://www.microscopy-today.com
Thank you.

Ron Anderson, Editor
================================
Illuminating the Formation of Lumens
Stephen W. Carmichael, Mayo Clinic, Rochester, MN

Moore’s Law: A Review of Feature Size Shrinkage and its Effect on
Microscopy in the Semiconductor Industry
John Mardinly, Intel Corporation, Santa Clara, CA

An SEM Analysis Of Amphipleura pellucida with New Findings
Gary Gaugler, Microtechnics, Inc., Granite Bay, CA

The Proposed Doppler Electron Velocimeter and the Need for Nanoscale
Dynamics
Phillip L. Reu, Sandia National Lab., Albuquerque, NM

Resolution and Sampling in Digital Imaging
Ian Dobbie, National University of Ireland, Galway, Ireland

Use of Astronomy Filters in Light Microscopy and Photomicrography
Jörg Piper, Clinic Meduna, Bad Bertrich, Germany

Analyzing the Growth of Magmatic Crystals – An Electron Microprobe
Analysis Study
Robert Sturm, University of Salzburg, Austria

µManager: Open Source Software for Light Microscope Imaging
Nico Stuurman, Nenad Amdodaj and Ron Vale, University of California, San
Francisco, CA

Vapor Coating: A Simple, Economical Procedure for Preparing Difficult
Specimens for Scanning Electron Microscopy
E. Ann Ellis and Michael W. Pendleton, Texas A&M University, College
Station, TX

Students and the SEM: The First Encounter
V.M.Dusevich, J.D.Eick, Univ. of Missouri, Kansas City, MO

The Magnification Myth
Sander Stoks and Bas Groen

Quality Controls on SEM Performance: A Novel Reference Sample
Brendan J. Griffin and Sharon T. Platten, Univ. of Western Australia,
Crawley, Australia

Industry News

NetNotes
SPECIMEN PREPARATION - LR White polymerization
SPECIMEN PREPARATION - pollen grains
SPECIMEN PREPARATION - tripod polishing problems
SPECIMEN PREPARATION - ruthenium tetroxide staining recipe
SPECIMEN PREPARATION – embedding mouse lens & eye
SPECIMEN PREPARATION - pre-embedding cells in agar
SPECIMEN PREPARATION - annealing tantalum
IMMUNOCTYOCHEMISTRY – BSA purity for use as blocking agent
IMAGE ANALYSIS – phase analysis
PHOTOGRAPHY – neutral density filters
DARKROOM - uneven printing
CAMERAS - CMOS versus CCD
MICROSCOPY - analysis of paper
LM - Koehler illumination
LM - calibration standard Z direction
EM - field sources
TEM - cleaning Pt apertures
TEM - free lens control
SEM - takeoff angle
SEM - shorted lead wires - current contrast?
SEM - lines on slow scan/capture
Electron beam simulation

Index of Advertisers


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From: heckman-at-bgnet.bgsu.edu
Date: Fri, 4 May 2007 10:45:16 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

HI, Tom-
In our experience, this seems a probable resultdue to re-oxidation of
the osmium in the dehydrating solutions. We never figured out what
the oxidant is in the ethanol, but we solved the problem by using
hexylene glycol as a dehydrating fluid in place of ethanol. You can
find a protocol on our web site. Just follow the procedure in the
Transmission Electron Microscopy section.
Carol


} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

--
Carol A. Heckman, Ph.D.
Director, Center for Microscopy & Microanalysis
and Professor of Biological Sciences
Bowling Green State University
Bowling Green, OH 43403
website: http://www.bgsu.edu/departments/biology/facilities/MnM

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From: cantonpa-at-unive.it
Date: Fri, 4 May 2007 11:28:30 -0500
Subject: [Microscopy] [Filtered] MicroscopyListserverviaWWW: LaB6 filament

Contents Retrieved from Microscopy Listserver Archives
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Email: cantonpa-at-unive.it
Name: Patrizia Canton

Organization: University of Venice

Title-Subject: [Filtered] LaB6 filament

Question: We had a new LaB6 filament (Denka,
M3,LKSH, tip shape 60†, 10micron mR Pointed)
installed in our Jeol 3010.
Jeol engineer had some problems during the
installation because there were some black
stripes visible in the filament image. He tried
to change te distance filament-Wehnelt three
times, to work with bias adjustment, with
filament saturation, after 5 days he had to end
his job because we finished the budget but with
no good results since he only obtained emission
current of 1 microA, and low current density. Now
we are struggling with this filament and coming
to nowhere. Is there anyone that can give some
hints, suggestions, bibliography?
Thanks
Patrizia

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From: mager-at-interchange.ubc.ca
Date: Fri, 4 May 2007 11:30:32 -0500
Subject: [Microscopy] TEM : objective aperture and EDX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Stephane,
Yes, the objective aperture sits just below the specimen and it is a
well-known phenomenon that Formvar film will "pop" if you observe the
specimen without inserting the objective aperture. The way it was explained
to me was that, when you look at your sample without the objective aperture,
all the radiation comes from one side and the charging, uneven heating and
radiation causes thermal stress and deformation that causes the film to
bulge and then, sometimes, break. When you insert the objective aperture,
the radiation reflects from the aperture and balances the heat and radiation
from the top, so there is not the stress of irradiating only one side.
For the EDX, remember that the specimen is essentially transparent to the
electron beam and the x-rays produced. The enormous flux of x-rays and
back-scattered electrons generated from the beam hitting the solid metal of
the aperture metal with 120kV electrons floods the EDX detector and "kills"
it, basically generating 100% dead time. It may take several minutes to
recover, after you remove the aperture. My sequence to switch to EDX
analysis always includes removing the objective aperture first. I find doing
EDX analysis on the TEM to be a careful balance between zero x-rays in thin
films and 100% dead time when you hit something too thick, like a grid bar.
They look the same on the EDX display.
I am mainly working in Materials Engineering, so I do lots of EDX on my TEM,
but I don't use Formvar films. I use carbon evaporated films that I make
myself and they seem to withstand 200kV TEM examination with or without an
objective aperture inserted and they are already conductive.
Good luck.
Regards,

Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: May 4, 2007 2:17 AM
To: mager-at-interchange.ubc.ca

Deal all!

I have a conceptual problem with my TEM (tecnai G20
(twin)). I asked help to 2 engineers from FEI, they
gave me opposite answers (!!).
Here is my problem:
The objective aperture is situated just under the
specimen itself. So it should have no influence on the
intensity of the beam hitting my sample.

In this case why does my formvar film dilate and
sometimes blow up when I forget to insert the
objective aperture but is extremely stabile when it is
inserted?

Even more strange: the EDX detector is placed at an
angle several degrees ABOVE the specimen. I cannot
detect anything when the objective aperture is
inserted, I must remove it to be able to read
something!

I have the feeling that actually the obj aperture is
at the same height as the specimen and not under it,
but one engineer told me it was just under the
specimen. Also, I dont understand why the formvar is
sensible to the presence of the obj aperture if it is
at the same height as my specimen.


Stephane

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
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From: tivol-at-caltech.edu
Date: Fri, 4 May 2007 12:21:51 -0500
Subject: [Microscopy] viaWWW: electron diffraction

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


On May 3, 2007, at 7:07 PM, walck-at-southbaytech.com wrote:

} Just a few comments off hand without ever doing these types of samples.
}
} Your problem will be similar to what happened to me with glass
} samples. You
} are putting too much energy into your sample and it is heating. There
} are
} several things you can do to help. You have to do things quickly,
} i.e. low
} dosage. You have to lower the amount of energy you are putting into
} the
} sample. The best way to do this is going to higher accelerating
} energy.
} Remember, diffraction is elastic and you don't loose energy in the
} sample
} via that route. If your energy is lower, you have more inelastic
} scattering
} and you are dumping energy into the sample with those types of
} scattering
} events. You probably are working on a 100 or 120 kV machine, but you
} didn't
} say so. And another thing that you can do is to cool your sample and
} use a
} liquid nitrogen stage.
}
} Having said this, I don't have any idea what the higher energy will do
} to
} your sample in terms of radiation damage.
}
Dear Scott,
Having done considerable ED at voltages from 100 to 1200 kV, I'd like
to offer a few minor corrections to your post. You are correct that
the problem is that too much energy was being deposited, but heating is
not the effect that causes the pattern to decay. Changes in the
specimen that destroy the crystalline order are responsible, and these
are caused by breaking of chemical bonds, ionization, and other
processes, such as displacement of atoms. Going to a higher energy
does give advantages for ED, principally making the scattering closer
to kinematic and getting higher resolution spots due to the flatter
Ewald sphere. As the energy of the beam increases, the total
scattering cross-section decreases, which results in less energy
deposited, but the ratio of elastic scattering to inelastic scattering
also decreases, so the damage-to-information ratio increases, but this
effect is minor, and excellent ED patterns can be obtained at 1200 kV.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu


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From: dsherman-at-purdue.edu
Date: Fri, 4 May 2007 14:22:16 -0500
Subject: [Microscopy] Carbon rod sharpener

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all,
I am looking for a carbon rod sharpened for rods with diameter of 0.12
inch diameter rods. We had a great electrical

one that rotated the rod and you just moved a small cutting edge along a
track to cut a nice long and narrow rod at the end of the larger rod. The
rotating shaft no longer is stable so rods break too easily.

Would appreciate advice as to replacements and would prefer electrical one
rather than the hand-held type.

Debby

Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.agriculture.purdue.edu/microscopy


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From: lcgould-at-med.cornell.edu
Date: Fri, 4 May 2007 14:37:39 -0500
Subject: [Microscopy] Re: Carbon rod sharpener

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Debby-
the one I have is about a gazillion years old (did they do EM then??)
but it still works beautifully. It was from Ladd. I don't know if
they still sell them, you'd have to check their online catalog.
Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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From: jd-at-laddresearch.com
Date: Fri, 4 May 2007 14:41:52 -0500
Subject: [Microscopy] Re: Carbon rod sharpener

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Debby,

Check out our's at
http://www.laddresearch.com/Key_Products/Digital_High_Vacuum_Evaporator/VacEvap/Carbon_Rod_Sharpening/carbon_rod_sharpening.html
It sounds like what you are looking for.


Disclaimer: Ladd Research sells carbon rod sharpeners, carbon rods,
carbon rod points, etc.


Debbie Sicard

Ladd Research
83 Holly Court
Williston, VT 05495

On-line Catalog: www.laddresearch.com

Telephone: 1-802-658-4961 (anywhere)
Toll Free 1-800-451-3406 (US)
Fax: 1-802-660-8859

e-mail: sales-at-laddresearch.com




At 03:29 PM 5/4/2007, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America


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From: shu-at-caltech.edu
Date: Fri, 4 May 2007 15:18:36 -0500
Subject: [Microscopy] RE: TEM : objective aperture and EDX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I agree that electrons coming from the objective aperture balance the
effect on the formvar film caused by the incident beam, as pointed out by
Dr Steve Chapman and Dr Mary Fletcher.
However, I doubt the force that breaks the film. The energy of the
incident electrons is of the order 100keV. The inelastic cross-section for
this energy loss is extremely small. So it is not likely there are a lot
incident electrons are stopped and accumulate on the film. Heat will
certainly be produced under illumination. But inelastic scattering can
happen at any depth, especially in a very thin film where most events are
single-scattering. So the heat should not differ too much between the two
surface.
I have a bold proposal. The film is broken by repulsive electrostatic
force. But the charge is possitive instead of negative. The formvar mainly
contains C, O, N and H which are light elements and easy to ionize. So
some atoms in the film lose valence electrons and become possitively
charged. Dey and Williams (J. Phys. D, 1988, vol.21, 108) studied the
EELS spectra of formvar film. They observed low binding energy peaks at
11-93eV. Scatterings in this range have considerable cross-sections and
can involve many atoms. On the other hand, the electrons coming
from the objective aperture, either back scattered or secondary electrons,
have lower energy compared to the incident beam and, hence easier to be
trapped by the film. So these electrons neutrolize the possitive charge
and stabilize the film.
This is just my personal speculation after reading posts on this topic.
Please feel free to correct me.

Shu Miao

On Fri, 4 May 2007 mager-at-interchange.ubc.ca wrote:

}
} Dear Stephane,
} Yes, the objective aperture sits just below the specimen and it is a
} well-known phenomenon that Formvar film will "pop" if you observe the
} specimen without inserting the objective aperture. The way it was explained
} to me was that, when you look at your sample without the objective aperture,
} all the radiation comes from one side and the charging, uneven heating and
} radiation causes thermal stress and deformation that causes the film to
} bulge and then, sometimes, break. When you insert the objective aperture,
} the radiation reflects from the aperture and balances the heat and radiation
} from the top, so there is not the stress of irradiating only one side.
} For the EDX, remember that the specimen is essentially transparent to the
} electron beam and the x-rays produced. The enormous flux of x-rays and
} back-scattered electrons generated from the beam hitting the solid metal of
} the aperture metal with 120kV electrons floods the EDX detector and "kills"
} it, basically generating 100% dead time. It may take several minutes to
} recover, after you remove the aperture. My sequence to switch to EDX
} analysis always includes removing the objective aperture first. I find doing
} EDX analysis on the TEM to be a careful balance between zero x-rays in thin
} films and 100% dead time when you hit something too thick, like a grid bar.
} They look the same on the EDX display.
} I am mainly working in Materials Engineering, so I do lots of EDX on my TEM,
} but I don't use Formvar films. I use carbon evaporated films that I make
} myself and they seem to withstand 200kV TEM examination with or without an
} objective aperture inserted and they are already conductive.
} Good luck.
} Regards,
}
} Mary Fletcher (nee Mager)
} Electron Microscopist
} Materials Eng. UBC
} #309 - 6350 Stores Road
} Vancouver, B.C. V6T 1Z4
} Canada
} Tel: 604-822-5648
} Fax: 604-822-3619
} email: mager-at-interchange.ubc.ca
}
} -----Original Message-----
}
}
} Deal all!
}
} I have a conceptual problem with my TEM (tecnai G20
} (twin)). I asked help to 2 engineers from FEI, they
} gave me opposite answers (!!).
} Here is my problem:
} The objective aperture is situated just under the
} specimen itself. So it should have no influence on the
} intensity of the beam hitting my sample.
}
} In this case why does my formvar film dilate and
} sometimes blow up when I forget to insert the
} objective aperture but is extremely stabile when it is
} inserted?
}
} Even more strange: the EDX detector is placed at an
} angle several degrees ABOVE the specimen. I cannot
} detect anything when the objective aperture is
} inserted, I must remove it to be able to read
} something!
}
} I have the feeling that actually the obj aperture is
} at the same height as the specimen and not under it,
} but one engineer told me it was just under the
} specimen. Also, I dont understand why the formvar is
} sensible to the presence of the obj aperture if it is
} at the same height as my specimen.
}
}
} Stephane


==============================Original Headers==============================
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From: tivol-at-caltech.edu
Date: Fri, 4 May 2007 15:47:36 -0500
Subject: [Microscopy] Re: [Filtered] MicroscopyListserverviaWWW: LaB6 filament

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


On May 4, 2007, at 9:28 AM, cantonpa-at-unive.it wrote:

} Question: We had a new LaB6 filament (Denka,
} M3,LKSH, tip shape 60†, 10micron mR Pointed)
} installed in our Jeol 3010.
} Jeol engineer had some problems during the
} installation because there were some black
} stripes visible in the filament image. He tried
} to change te distance filament-Wehnelt three
} times, to work with bias adjustment, with
} filament saturation, after 5 days he had to end
} his job because we finished the budget but with
} no good results since he only obtained emission
} current of 1 microA, and low current density. Now
} we are struggling with this filament and coming
} to nowhere. Is there anyone that can give some
} hints, suggestions, bibliography?
}
Dear Patrizia,
As the current through a LaB6 filament is increased, the first part of
the filament that emits electrons is the flat part of the tip, which is
a truncated pyramid. The next parts of the filament to emit are the
faces of the pyramid, and finally, the last parts to emit are the edges
between the flat faces. If you adjust the condenser to crossover and
put the mag to about 30 kx, so you can see the image of the filament,
you will first see a very small, round spot as the tip starts to emit,
then four larger, brighter spots will appear around the first one,
these will get larger and brighter and fill out the area until only
four dark streaks, where the emission from the edges would be, remain,
and finally, these will also fill in giving a uniform spot when the
filament is saturated. This assumes that the filament is properly
centered in the Wehnelt aperture. The failure to achieve saturation
and the low emission current sound like the filament is not getting
enough current. If the black stripes were not evenly spaced and along
radial directions in the filament image, then it is likely that the tip
was damaged, and the damaged parts are not emitting.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu



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From: tivol-at-caltech.edu
Date: Fri, 4 May 2007 15:54:56 -0500
Subject: [Microscopy] Re: viaWWW: orientation of the diffraction

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


On May 4, 2007, at 5:52 AM, wvrenter-at-sckcen.be wrote:

} Question: When you defocus a diffraction pattern, you can see a small
} image of the selected area in the 000 spot. When you go from
} underfocus to overfocus the image of the selected area is rotated over
} 180ƒ. Can somebody tell me which of the two small images has the same
} orientation as the image in image mode? Do I have to underfocus the
} diffraction pattern or overfocus it?
}
Dear Wouter,
The orientation of the pattern is not necessarily the same as that of
the image in either under or over focus, and, furthermore, it can be
different for different camera lengths. There should be information in
the manual of your instrument that can tell you the difference in
orientations. If not, put in a specimen with an asymmetric object,
take an image in normal mode, then take one in defocussed diffraction
mode and compare. An aggregate of gold beads on a carbon film is a
good test object for this.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu



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From: leunissen-at-aurion.nl
Date: Fri, 4 May 2007 16:57:27 -0500
Subject: [Microscopy] Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Good day

For OsO4 to be reduced to OsO2 (as happens during fixation) requires
the presence of hydrogen ions and an electron donor. Electrons can be
supplied from breaking up the double bonds in fatty acid alkyl chains
of unsaturated (membrane) lipids.

The reaction is reversible, so reoxidation is theoretically possible,
but only if a stronger oxidizer with a redox potential more positive
than the one associated with the OsO4/OsO2 redox couple (Eo =1.02
Volts) is present. And even then, re-oxidation will only occur under
suitable circumstances! I found Carol's comment that hexylene glycol
preserves the Osmium contrast very interesting. What could it be in
ethanol that might re-oxidize the Osmium? Peroxides? Chlorine? Oxygen
in statu nascendi? OsO4 as such is a pretty strong oxidizing agent
already. Very puzzling.

I hope I am not making any serious mistakes here, I am not a chemist
(I wish!), just trying to understand how it works, but it seems a
slightly acidic environment would be good to promote the fixation as
well as to preserve the OsO2 in the tissue after fixation. Could it
be that the ethanol is alkaline? Even though ethanol is still
somewhat polar, it may not take much in a non-aquaeous environment.

I am sure there will be proper chemists and physicists subsribing to
this great listserver. Maybe they would be willing to look into our
mostly empirically established procedures, we might be able to make a
big move forward....

Jan Leunissen



Aurion - President Electron Microscopy Research Advisor
Costerweg 5 Dept Anatomy and Structural Biology
6702 AA Wageningen Otago School of Medical Sciences
The Netherlands Dunedin, New Zealand
phone 31-317-497676 phone 64-3-4795465
fax 31-317-415955 fax 64-3-4797254
http://www.aurion.nl http://anatomy.otago.ac.nz
-----------------------------------------------------------------

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From: kraftpiano-at-gmail.com
Date: Fri, 4 May 2007 20:28:40 -0500
Subject: [Microscopy] SEM: Intermittent Vacuum Leak

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I'm completely dumbfounded by this one. I am in the process of
checking for leaks on our SEM, and it pumps down perfectly one moment,
then it won't even trigger PiG3. When I start to take things apart to
check seals and such, then I put it back together (After finding
nothing wrong) it evacuates perfectly. Then I let it sit overnight to
see if there is a leak, and when I get in the next morning, it is
completely at atmospheric pressure and then won't hold a vacuum and
trigger PiG3 anymore. I'm a little confused at how it will hold a
vacuum only the first time I pump it down after dismantling and
reassembling parts of the vacuum system. I don't have a leak
detector, but boy would I love one! (I also don't have any gauges
capable of measuring absolute vacuum vs. relative vacuum, so I have no
idea what the actual measurement in torr is.)

--Justin A. Kraft

==============================Original Headers==============================
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From: rcmoretz-at-gmail.com
Date: Fri, 4 May 2007 20:44:55 -0500
Subject: [Microscopy] Optical Microscopy

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

As soon as I saw Bruce, I thought of the Monty Python skit. Then
there was our 3 year old daughter in the back seat (of othe car) one
night as Mom & Dad (spouse and me) were discussing names for the
impending baby (this was 1972, ok?) when she (in the back seat) piped
up and said "What about Bruce?".

Roger Moretz, Ph.D.

On 5/3/07, frank.karl-at-degussa.com {frank.karl-at-degussa.com} wrote:
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Guilty!
}
} Having worn glasses since eight grade, I have come to associate optical
} with glass the photons.
}
} Phototonic microscopy? sounds like a title of a paper in search of a
} little primping
} Non-modified bright field? - sounds like a title of a paper in search of a
} little primping from a major university
} Light microscopy? - sort of suggest their might be a heavy microscopy
} Refractory microscopy - sound like microscopy in hell
} Specular microscopy - the study of the stock market in great detail..
}
} Tongue firmly implanted in cheek: Let's call it Bruce. I got the idea
} from a microscopy society bog (
} http://www.msneo.org/2006/06/blog-mail.html)
} who knows..........
}
} Stay safe..............Frank
}
}
}
}
} protrain-at-emcourse
} s.com To: frank.karl-at-degussa.com
} cc:
} 05/03/2007 07:09 Subject: [Microscopy] Optical Microscopy
} AM
} Please respond to
} protrain
}
}
}
}
}
}
}
}
} ----------------------------------------------------------------------------
}
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} ----------------------------------------------------------------------------
}
}
} Hi
}
} All the fuss over microprobes was fun but if you want to get your teeth
} into
} names used by scientists (?) what about "optical microscopy"?
}
} With my 43 years in the business I have always taught that whilst light
} microscopes have "light optics" electron microscopes have "electron
} optics",
} the principles are the same. So when I see a piece about optical
} microscopes I always look to see if it should be of interest to me?
}
} What is the general thought?
}
} Steve Chapman
} Senior Consultant Protrain
} For electron microscopy consultancy and training world wide
} Tel +44 1280 816512 Fax +44 1280 814007
} Mobile +44 7711 606967 Web www.emcourses.com
}
}
} ==============================Original
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==============================Original Headers==============================
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From: heckman-at-bgnet.bgsu.edu
Date: Sat, 5 May 2007 15:41:30 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi All

I think Shu has a very good point, the explanation I had been given 40 years
ago that I sent through to him was:-

"Yes it is a very strange phenomena. Remember the plastic film, like any
material in the electron beam, has a charge upon it, even so close to the
copper grid. People like David Joy will tell you that even the copper grid
has a charge upon it, nothing is a perfect conductor! The changes in
texture/thickness within the film take equal charge, but like charges repel
so when the charge reaches a specific level repulsion of two areas
eventually tears the film apart along the weaker areas. That is the
explanation that I was told many years ago, it does seem to fit the action
that occurs. Reduce the number of incident electrons and you reduce the
charge, coat with carbon and you increase the thickness and the conductivity
but you could spoil the resolution in relation to T/10.

For some reason the backscatter from the aperture reduces this charge
effect, or spraying electrons onto the sample with a charge neutraliser
(available in the 60s) did the same! The grid provides conductivity, the
backscatter seems to provide charge neutralisation, it is the charge that
breaks the film."

Just shows once again how the listserver can be so informative - "don't ask
and you don't get!"

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com

----- Original Message -----
X-from: {shu-at-caltech.edu}
To: {protrain-at-emcourses.com}
Sent: Friday, May 04, 2007 9:19 PM

Hi, Jan-
The question of the redox potential also puzzled me when I first
came up with this explanation for why membranes get "bleached"
after fixation with OsO4. Your idea of the alkalinity making a
difference is an appealing alternative explanation. Another aspect
of this phenomenon (problem, I mean) that was hard to
understand was that it appeared to be restricted to tissue culture
preparations, where the sample is very thin. Would the alkalinity
affect the thin sample preferentially? Using the same
reagents, we had no problem with the apparent extraction of OsO4
from membranes in bulk tissues.

Whatever we did, the hypothesis I gave was merely based on the
empirical finding that changing the solvent solved the problem. It
has no implications for the true mechanism of this effect. Now that
there are so many new instrumental methods, people are making
progress on definfing the chemical after-effects of fixation
methods!
With best regards,
Carol



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4, 14 -- Subject: Re: [Microscopy] Help with enhancing contrast of Mitochondrial
==============================End of - Headers==============================




From: protrain-at-emcourses.com
Date: Sun, 6 May 2007 04:39:06 -0500
Subject: [Microscopy] Support Film Damage in TEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

Whilst we have been discussing the failure of support films in TEM and
possible ionisation taking place, I feel it is right to indicate another
form of problem?

If there is a leak near to the specimen area of the TEM, usually the
specimen exchange mechanism, another interesting problem may appear. Whilst
working with a thin support film you may notice that the film slowly becomes
lacy, the holes growing with time until the film falls apart. What is
happening is the gas from the leak is being ionised and it is etching away
the support film. From my service background it was a give-away as a
specimen exchange leak! This usually showed up whilst I was checking the
instrument with a holey carbon film thus demonstrating that a holey film may
come in handy for more than image checking (?).

Hope this helps one of you in the future?

Regards

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


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From: cgarber-at-2spi.com
Date: Sun, 6 May 2007 10:14:35 -0500
Subject: [Microscopy] Carbon rod sharpener

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Debby Sherman wrote:
====================================================================

I am looking for a carbon rod sharpened for rods with diameter of 0.12
inch diameter rods. We had a great electrical one that rotated the rod
and you just moved a small cutting edge along a
track to cut a nice long and narrow rod at the end of the larger rod. The
rotating shaft no longer is stable so rods break too easily.


Would appreciate advice as to replacements and would prefer electrical
one rather than the hand-held type.
===================================================================
SPI Supplies has offered an electrically driven carbon rod sharpener for
a number of years, see URL
http://www.2spi.com/catalog/spec_prep/carbon-rod-sharpener.shtml

for a photo of the product and full technical description.

Our standard set up for 1/8" diameter or 3 mm should work for your
0.12" rods. The product is shipped with what would be needed if you
have carbon rods of some other diameter such as 4.8 and 6.1 mm diameter.
In that sense it is a "universal" sharpener and can sharpen all
diameters of rods found in an EM laboratory.

Chuck
===================================================

Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
President
SPI SUPPLIES FAX: 1-(610)-436-5755
PO BOX 656 e-mail: cgarber-at-2spi.com
West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com


Look for us!
############################
WWW: http://www.2spi.com
############################
==================================================


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From: cantonpa-at-unive.it
Date: Mon, 7 May 2007 08:27:58 -0500
Subject: [Microscopy] LaB6 filament:thanks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I would like to thank all the people who answered and help me to
understand what was going on.
After two days of checking it seems that our filament was not properly
centered in the Wehnelt.
Best Regards
Patrizia

--
______________________________________________
Patrizia Canton PhD
Dept. of Physical Chemistry
Via Torino 155/b
I-30170
Venezia-Mestre Italy
Phone +39-041-2346790
Fax +39-041-2346747
e-mail cantonpa-at-unive.it
______________________________________________


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From: mager-at-interchange.ubc.ca
Date: Mon, 7 May 2007 11:12:37 -0500
Subject: [Microscopy] SEM: Intermittent Vacuum Leak

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Justin,
This sounds like a very difficult problem to solve, especially if you don't
have a good high vacuum gauge. I would suggest you check the diffusion pump
and make sure it is getting hot at the bottom and cool at the top and is
staying that way. I once had an "intermittent vacuum leak" that turned out
to be that I had put the wrong diffusion pump oil in the pump and it was not
quite boiling. That only caused poor high vacuum, though, not degradation
back to atmosphere. Also check the quantity and quality (color) of the oil
in the diffusion pump. Also, some sensor or vacuum gauge may be dirty or
faulty and switching the vacuum off intermittently. I see no choice but to
sit there until it does the bad thing while you are watching.
Good luck,

Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: kraftpiano-at-gmail.com [mailto:kraftpiano-at-gmail.com]
Sent: May 4, 2007 6:34 PM
To: mager-at-interchange.ubc.ca

I'm completely dumbfounded by this one. I am in the process of
checking for leaks on our SEM, and it pumps down perfectly one moment,
then it won't even trigger PiG3. When I start to take things apart to
check seals and such, then I put it back together (After finding
nothing wrong) it evacuates perfectly. Then I let it sit overnight to
see if there is a leak, and when I get in the next morning, it is
completely at atmospheric pressure and then won't hold a vacuum and
trigger PiG3 anymore. I'm a little confused at how it will hold a
vacuum only the first time I pump it down after dismantling and
reassembling parts of the vacuum system. I don't have a leak
detector, but boy would I love one! (I also don't have any gauges
capable of measuring absolute vacuum vs. relative vacuum, so I have no
idea what the actual measurement in torr is.)

--Justin A. Kraft

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From: dac-at-research.umass.edu
Date: Mon, 7 May 2007 12:13:36 -0500
Subject: [Microscopy] SEM: Intermittent Vacuum Leak

Contents Retrieved from Microscopy Listserver Archives
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Justin,

Does it truly not have vacuum, or only indicate this?

A possibility is that the vacuum sensors or processing circuits are
giving erratic performance/faulty response and since the control of the
vacuum system is based on these it can complicate the troubleshooting.

Also, most scopes have a "Klixon" on the ODP column (or 2: one for
overtemp and one for "hot enough to function") that are continually
subjected to heat and will go flakey in time and may be shutting down
the ODP. We've had a good bit of this problem over the years.

Try to get a handle on the vacuum by some independent means if possible
- if not a sepatate gage unit, then try to read the existing raw sensor
voltage where it connects to the microscope to see what is happening
(this assumes you have/read schematics and can locate the places to read
the sensor); or your scope may have some status LEDs on the vacuum
control board that might actually be labelled?

If don't want to pay for a commercial Pirani unit and you are inclined
to "tinkering" I have some information on a DIY "Light Bulb Pirani" gage
sensor/circuit based on an opened light bulb tungsten element and a
simple driver circuit. The tungsten is heated to a preset temperature
and the voltage required to maintain that temperature is proportional to
pressure - at atmosphere the heat is conducted away so more voltage is
required; there is a spreadsheet with data, a schematic, and a couple of
photos. This system is not temperature compensated, so is best kept away
from sources of heat and for relative vacuum level sensing you probably
don't need to calibrate against another gage, just look at the
spreadsheet data. It requires only 2 feedthrough pins into the vacuum.

http://www-unix.oit.umass.edu/~dac/projects/LightBulb_Pirani/

Cheers!

Dale

mager-at-interchange.ubc.ca wrote:
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} Dear Justin,
} This sounds like a very difficult problem to solve, especially if you don't
} have a good high vacuum gauge. I would suggest you check the diffusion pump
} and make sure it is getting hot at the bottom and cool at the top and is
} staying that way. I once had an "intermittent vacuum leak" that turned out
} to be that I had put the wrong diffusion pump oil in the pump and it was not
} quite boiling. That only caused poor high vacuum, though, not degradation
} back to atmosphere. Also check the quantity and quality (color) of the oil
} in the diffusion pump. Also, some sensor or vacuum gauge may be dirty or
} faulty and switching the vacuum off intermittently. I see no choice but to
} sit there until it does the bad thing while you are watching.
} Good luck,
}
} Mary Fletcher (nee Mager)
} Electron Microscopist
} Materials Eng. UBC
} #309 - 6350 Stores Road
} Vancouver, B.C. V6T 1Z4
} Canada
} Tel: 604-822-5648
} Fax: 604-822-3619
} email: mager-at-interchange.ubc.ca
}
} -----Original Message-----
} X-from: kraftpiano-at-gmail.com [mailto:kraftpiano-at-gmail.com]
} Sent: May 4, 2007 6:34 PM
} To: mager-at-interchange.ubc.ca
} Subject: [Microscopy] SEM: Intermittent Vacuum Leak
}
}
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} I'm completely dumbfounded by this one. I am in the process of
} checking for leaks on our SEM, and it pumps down perfectly one moment,
} then it won't even trigger PiG3. When I start to take things apart to
} check seals and such, then I put it back together (After finding
} nothing wrong) it evacuates perfectly. Then I let it sit overnight to
} see if there is a leak, and when I get in the next morning, it is
} completely at atmospheric pressure and then won't hold a vacuum and
} trigger PiG3 anymore. I'm a little confused at how it will hold a
} vacuum only the first time I pump it down after dismantling and
} reassembling parts of the vacuum system. I don't have a leak
} detector, but boy would I love one! (I also don't have any gauges
} capable of measuring absolute vacuum vs. relative vacuum, so I have no
} idea what the actual measurement in torr is.)
}
} --Justin A. Kraft
}
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10, 21 -- From dac-at-research.umass.edu Mon May 7 12:13:36 2007
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From: kraftpiano-at-gmail.com
Date: Mon, 7 May 2007 15:31:42 -0500
Subject: [Microscopy] Re: SEM: Intermittent Vacuum Leak

Contents Retrieved from Microscopy Listserver Archives
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Hi

I do not know which instrument that you have as you do not say? I have
experienced a problem in the past where the customer behaved exactly as you
have. Strip and rebuild no problem, next day no good! In this case simply
turning the vacuum system off and on overcame the fault. Could this be your
problem as when you strip and rebuild you turn the system off?

If you could provide details of which instrument that you are using and how
it is pumped - diffusion pump or turbo molecular - this would help?

Please get back and I will try to help.

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


----- Original Message -----
X-from: {kraftpiano-at-gmail.com}
To: {protrain-at-emcourses.com}
Sent: Saturday, May 05, 2007 2:29 AM

Vacuum leak update:

Thanks to the many suggestions I received, the vacuum leak has been
identified. I thought I'd send in a summary of the suggestions that
ended up being the most helpful so those with similar problems in the
future may benefit. Here is the summary:

1: Using a large quantity of various sized stoppers, isolate different
parts of the vacuum system to determine where the leak may be.

2: If the system has multiple piranni gauges (As the JSM-840 does) try
switching the gauges around to make sure that it is not a bad gauge.

3: Check that the diffusion pumps are heating up. (Although this was
not the issue with mine, since it was not reaching a vacuum level that
would allow the diffusion pumps to begin to work, I thought I'd
include it just as general reference.) Also check to make sure that
the temperature sensors on the diffusion pumps are functional.

4: Short of a leak detector, spray some acetone or high purity
isopropyl alcohol on the different seals. Wait a few minutes after
each spray. If there is a leak in that spot, then the vacuum will
increase momentarily, then go back down as the alcohol or acetone
plugs the leak and then vaporizes on the inside of the system. Keep
doing this moving from the outermost portions of the vacuum system to
the pump connection until you find it.

5: Pump the system down as far as it will go, then seal all of the
valves in the closed position. Wait. The next morning, the section
with the leak will not have a vacuum in it, but the others will.

Rick Becker given this handy suggestion for repairing the leak when found:

If the leak is in a section that you can get around (i.e. the junction
between two column sections) standard electrical tape can repair it
temporarily. Make sure it is high quality electrical tape. The
thicker, the better. Decent electrical tape can hold a vacuum of 10
e-9 torr. Not bad- the tape is now holding the vacuum between the "T"
fitting under the chamber (Bottom is the small vent valve, top is the
chamber, and the leg on the side is a large butterfly valve into the
rest of the vacuum system) and the large butterfly valve. I ordered a
new gasket from Small Parts, Inc. (They really do have everything!)
but in the mean time, it's holding fine.

As it turns out (For those of you wondering) when we re-assembled the
vacuum system after carrying it piece by piece up the stairs, we must
have accidentally pinched the gasket when putting the two fittings
together. This caused a nick about the size of a couple of grains of
sand in the gasket, which would become a problem after being
reassembled when the gasket would roll slightly after tightening the
screws.

Hope this summary helps others who might need it!

--Justin.

On 5/4/07, kraftpiano-at-gmail.com {kraftpiano-at-gmail.com} wrote:
}
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} I'm completely dumbfounded by this one. I am in the process of
} checking for leaks on our SEM, and it pumps down perfectly one moment,
} then it won't even trigger PiG3. When I start to take things apart to
} check seals and such, then I put it back together (After finding
} nothing wrong) it evacuates perfectly. Then I let it sit overnight to
} see if there is a leak, and when I get in the next morning, it is
} completely at atmospheric pressure and then won't hold a vacuum and
} trigger PiG3 anymore. I'm a little confused at how it will hold a
} vacuum only the first time I pump it down after dismantling and
} reassembling parts of the vacuum system. I don't have a leak
} detector, but boy would I love one! (I also don't have any gauges
} capable of measuring absolute vacuum vs. relative vacuum, so I have no
} idea what the actual measurement in torr is.)
}
} --Justin A. Kraft
}
} ==============================Original Headers==============================
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} 2, 26 -- Subject: SEM: Intermittent Vacuum Leak
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13, 28 -- From: "Justin Kraft" {kraftpiano-at-gmail.com}
13, 28 -- To: microscopy-at-microscopy.com
13, 28 -- Subject: Re: [Microscopy] SEM: Intermittent Vacuum Leak
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From: Colin.Veitch-at-csiro.au
Date: Mon, 7 May 2007 17:49:33 -0500
Subject: [Microscopy] Many thanks for s4300 video question and imaging cell walls

Contents Retrieved from Microscopy Listserver Archives
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Just a quick not to say thank you to all of those who responded to my
recent questions on the video capture from a Hitachi S4300 and the
imaging of carrot cell walls.

Cheers

Colin Veitch

Electron Microscopist
CSIRO Textile and Fibre Technology
PO Box 21, BELMONT, Vic. 3216. Australia.
colin.veitch-at-csiro.au
http://www.tft.csiro.au

+61 (0) 3 5246 4000
0438 538 475
+61 (0) 3 5246 4811

The information contained in this e-mail message may be privileged or
confidential information. If you are not an intended recipient, you may
not copy, distribute or take any action in reliance on it. If you have
received this message in error, please telephone CSIRO Textile and Fibre
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From: dianavd-at-eye.usyd.edu.au
Date: Mon, 7 May 2007 20:43:35 -0500
Subject: [Microscopy] muddy blocks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I've recently had a problem with embedded blocks producing sections
that look "muddy". The resin part of the section is fine, but the
tissue is uniformly dark, as though it's embedded in mud, and is
sometimes quite fragile, the section (tissue area) falling apart
easily. Before I throw everything out and start with fresh
reagents....any ideas? I receive the tissue already fixed, usually in
glut/form in cacodylate buffer, not sure if it's from same operator
each time; quality hasn't been good, but is adequate. It's then
washed in cacodylate. Fixed in osmium (commercially prepared, clear
and proven to have produced good results). Uranyl acetate block stain
for an hour (same bottle I've been using for a long time). Ethanol
and acetone dehydration (both kept dry with copper sulphate). Epon/
araldite embedding. The annoying thing is that it's not constant -
one run will be fine, another ends up with the mud. I should do a run
with different tissues, but don't have spare at the moment (and
anyway I HATE embedding)....

It's probably something simple I'm doing wrong, but after all these
years perhaps I'm blind to the obvious.

Cheers,

Diana


Diana van Driel
Dept Ophthalmology
Sydney University
GPO Box 4337
Sydney, NSW
AUSTRALIA 2001

Phone 61 2 93827278
Mobile 0423 151614
FAX 61 2 93827318


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From: gmartens-at-interchange.ubc.ca
Date: Tue, 8 May 2007 10:16:12 -0500
Subject: [Microscopy] Re: muddy blocks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Diana,

When does the 'mud' appear? If it appears right after osmium
tetroxide fixation it sounds like there might be some tannic acid in
the tissue. I would ask the researcher for the complete protocol. I
have seen this before when a researcher 'forgets' to tell me they
read a paper that used tannic acid in the fix. If it appears after
polymerization it sounds like incomplete dehydration, which will
definitely lead to the fragile (I interpreted this as brittle) block.
This is usually a result of a tissue piece that is too large for the
solvent to penetrate all the way to the centre. Besides, what is to
hate about embedding? The time consumed or the exposure to the nasty
chemicals?

Good luck from Canada

Garnet

}
} I've recently had a problem with embedded blocks producing sections
} that look "muddy". The resin part of the section is fine, but the
} tissue is uniformly dark, as though it's embedded in mud, and is
} sometimes quite fragile, the section (tissue area) falling apart
} easily. Before I throw everything out and start with fresh
} reagents....any ideas? I receive the tissue already fixed, usually in
} glut/form in cacodylate buffer, not sure if it's from same operator
} each time; quality hasn't been good, but is adequate. It's then
} washed in cacodylate. Fixed in osmium (commercially prepared, clear
} and proven to have produced good results). Uranyl acetate block stain
} for an hour (same bottle I've been using for a long time). Ethanol
} and acetone dehydration (both kept dry with copper sulphate). Epon/
} araldite embedding. The annoying thing is that it's not constant -
} one run will be fine, another ends up with the mud. I should do a run
} with different tissues, but don't have spare at the moment (and
} anyway I HATE embedding)....
}
} It's probably something simple I'm doing wrong, but after all these
} years perhaps I'm blind to the obvious.
}
} Cheers,
}
} Diana
}

--
Garnet Martens

Research Manager
BioImaging Facility
University of British Columbia
6270 University Blvd.
Vancouver, B.C.
Canada
V6T 1Z4

phone 604-822-3354

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From: mdufraine-at-ebsciences.com
Date: Tue, 8 May 2007 10:19:18 -0500
Subject: [Microscopy] Re: Carbon rod sharpener

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Debbie-

Quorum Technologies has a variable size electrical carbon rod sharpener,
please review the following URL for picture and description of the unit.
http://www.quorumtech.com/Products/sc7605-carbon-rod-grinder.htm

Mike Dufraine
EM-Product Manager
Energy Beam Sciences,Inc.

dsherman-at-purdue.edu wrote:

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--
Michael F. Dufraine
EM - Product Manager
TEL 800-992-9037 X340
MDufraine-at-ebsciences.com
www.ebsciences.com


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From: peter.tomic-at-renwireless.com
Date: Tue, 8 May 2007 12:33:06 -0500
Subject: [Microscopy] Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Microscopy Folks,

I'd like to get some input on sample preparation with respect to silicon.

PROBLEM:

I have an FEI XL-50 FESEM that was originally designed to accept flat
samples, essentially silicon wafers. It does not have what one would
consider a typical exchange port. The exchange port is robotically
controlled, and the maximum sample height, including stub, must be 5 mm
or less. I must view these samples on edge, i.e. 90 degrees. This
presents a problem in that I must cleave two parallel sections very
close to each other. Perhaps diamond cutters can do this, but my hands
are too shaky.

QUESTION:

Is there a device, or method, that would allow me to make these cleaves
~ 8 mm apart with any reasonable control? I found stubs that are low
profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
still have this issue of doing two parallel cleaves very close together.

Just for info. purposes I am in the silicon MEMS development arena.

If you feel your reply is of general interest to this community, please
reply to all, or you may contact me directly.

Regards to all in this small world,

Peter Tomic
Renaissance Wireless Corp.


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From: bozhilov-at-ucr.edu
Date: Tue, 8 May 2007 13:47:06 -0500
Subject: [Microscopy] Diffracation ring problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all,

I am trying to find resolution of a problem with our CM300 TEM with a
LaB6 cathode.

When trying to get diffraction from an almost amorphous material I
have noticed a presence of relatively weak but clearly visible
perfectly circular ring of intensity around the central transmitted
spot. The ring is not centered around the transmitted spot. Its
position varies when operating the beam shift controls, its size
varies with the C2 aperture size when in diffraction mode. It is also
present when there is no sample under the beam. When there is
strongly reflecting crystalline material the ring is almost invisible
due to its weak relative intensity. It is present at any accelerating
voltage.

I am trying to get some ideas about what might be causing the ring
and how to eliminate it.

The ring is also visible when in LM imaging mode (image is formed by
the diffraction lens)
and when the beam is focused to a spot. In LM mode the ring has
strange shape. It has sharp circular outline on its outer edge and it
has irregular shape and outline on its inner edge.

Using the free lens control option I have made the following
observations:

In LM imaging mode with beam focused to a spot.
When changing the C1 lens current -the ring changes size and focus
but does not rotate.
changing C2 lens current - change in focus only, plus some rotation
changing Twin lens current - change of size and focus, no rotation
changing Objective lens current - change in size and focus, some
rotation
changing Diffraction lens current - change in size and focus, no
rotation
changing Intermediate lens current - change in size and focus, no
rotation
changing P1 lens current - change in size and focus, some rotation
changing P2 lens current - rotation only

In diffraction mode with fully spread beam:
position varies when operating the beam shift controls
size varies with the C2 aperture size.

FEI support engineers have not been succesful in identifing the
problem so far. It was suggested that it is due to undersaturated
LaB6 cathode, a possibility which we have clearly eliminated as a
possible source.

Thanks for any hints our suggestions.

Krassimir.
_______________________________________
Krassimir N. Bozhilov
Central Facility for Advanced Microscopy and Microanalysis
University of California
Riverside, CA 92521

tel 951 827 2998
fax 951 827 2489
bozhilov-at-ucr.edu
_______________________________________



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From: randerson20-at-tampabay.rr.com
Date: Tue, 8 May 2007 14:44:37 -0500
Subject: [Microscopy] Re: Diffracation ring problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Consider the possibility that it might be visible light from the hot
cathode coming down the column, i.e. like a flashlight.

This argument is abetted by the sharp outside edge, from apertures along
the way, and a diffuse inner edge and the fact that you see it without a
specimen loaded. However, it isn't immediately obvious how or why it
changes size and shape with all of your other perturbations.

Can you partly eclipse it by moving apertures around?

Ron Anderson

bozhilov-at-ucr.edu wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi all,
}
} I am trying to find resolution of a problem with our CM300 TEM with a
} LaB6 cathode.
}
} When trying to get diffraction from an almost amorphous material I
} have noticed a presence of relatively weak but clearly visible
} perfectly circular ring of intensity around the central transmitted
} spot. The ring is not centered around the transmitted spot. Its
} position varies when operating the beam shift controls, its size
} varies with the C2 aperture size when in diffraction mode. It is also
} present when there is no sample under the beam. When there is
} strongly reflecting crystalline material the ring is almost invisible
} due to its weak relative intensity. It is present at any accelerating
} voltage.
}
} I am trying to get some ideas about what might be causing the ring
} and how to eliminate it.
}
} The ring is also visible when in LM imaging mode (image is formed by
} the diffraction lens)
} and when the beam is focused to a spot. In LM mode the ring has
} strange shape. It has sharp circular outline on its outer edge and it
} has irregular shape and outline on its inner edge.
}
} Using the free lens control option I have made the following
} observations:
}
} In LM imaging mode with beam focused to a spot.
} When changing the C1 lens current -the ring changes size and focus
} but does not rotate.
} changing C2 lens current - change in focus only, plus some rotation
} changing Twin lens current - change of size and focus, no rotation
} changing Objective lens current - change in size and focus, some
} rotation
} changing Diffraction lens current - change in size and focus, no
} rotation
} changing Intermediate lens current - change in size and focus, no
} rotation
} changing P1 lens current - change in size and focus, some rotation
} changing P2 lens current - rotation only
}
} In diffraction mode with fully spread beam:
} position varies when operating the beam shift controls
} size varies with the C2 aperture size.
}
} FEI support engineers have not been succesful in identifing the
} problem so far. It was suggested that it is due to undersaturated
} LaB6 cathode, a possibility which we have clearly eliminated as a
} possible source.
}
} Thanks for any hints our suggestions.
}
} Krassimir.
} _______________________________________
} Krassimir N. Bozhilov
} Central Facility for Advanced Microscopy and Microanalysis
} University of California
} Riverside, CA 92521
}
} tel 951 827 2998
} fax 951 827 2489
} bozhilov-at-ucr.edu
} _______________________________________
}
}
}
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6, 19 -- From randerson20-at-tampabay.rr.com Tue May 8 14:44:37 2007
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From: bozhilov-at-ucr.edu
Date: Tue, 8 May 2007 16:19:35 -0500
Subject: [Microscopy] Diffracation ring problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

When the beam is focused to a probe in LM mode moving the C2 or the
diffrcation aperture does not eclipse the ring. It chages only the
intensity of the light or when moved too far blocks all the light.
Only by introducing and moving the objective aperture one can select/
block part of the ring, respectively the central spot.

Krassimir.


On May 8, 2007, at 12:46 PM, randerson20-at-tampabay.rr.com wrote:

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} Consider the possibility that it might be visible light from the hot
} cathode coming down the column, i.e. like a flashlight.
}
} This argument is abetted by the sharp outside edge, from apertures
} along
} the way, and a diffuse inner edge and the fact that you see it
} without a
} specimen loaded. However, it isn't immediately obvious how or why it
} changes size and shape with all of your other perturbations.
}
} Can you partly eclipse it by moving apertures around?
}
} Ron Anderson
}
} bozhilov-at-ucr.edu wrote:
} } ---------------------------------------------------------------------
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} }
} } Hi all,
} }
} } I am trying to find resolution of a problem with our CM300 TEM with a
} } LaB6 cathode.
} }
} } When trying to get diffraction from an almost amorphous material I
} } have noticed a presence of relatively weak but clearly visible
} } perfectly circular ring of intensity around the central transmitted
} } spot. The ring is not centered around the transmitted spot. Its
} } position varies when operating the beam shift controls, its size
} } varies with the C2 aperture size when in diffraction mode. It is also
} } present when there is no sample under the beam. When there is
} } strongly reflecting crystalline material the ring is almost invisible
} } due to its weak relative intensity. It is present at any accelerating
} } voltage.
} }
} } I am trying to get some ideas about what might be causing the ring
} } and how to eliminate it.
} }
} } The ring is also visible when in LM imaging mode (image is formed by
} } the diffraction lens)
} } and when the beam is focused to a spot. In LM mode the ring has
} } strange shape. It has sharp circular outline on its outer edge and it
} } has irregular shape and outline on its inner edge.
} }
} } Using the free lens control option I have made the following
} } observations:
} }
} } In LM imaging mode with beam focused to a spot.
} } When changing the C1 lens current -the ring changes size and focus
} } but does not rotate.
} } changing C2 lens current - change in focus only, plus some rotation
} } changing Twin lens current - change of size and focus, no rotation
} } changing Objective lens current - change in size and focus, some
} } rotation
} } changing Diffraction lens current - change in size and focus, no
} } rotation
} } changing Intermediate lens current - change in size and focus, no
} } rotation
} } changing P1 lens current - change in size and focus, some rotation
} } changing P2 lens current - rotation only
} }
} } In diffraction mode with fully spread beam:
} } position varies when operating the beam shift controls
} } size varies with the C2 aperture size.
} }
} } FEI support engineers have not been succesful in identifing the
} } problem so far. It was suggested that it is due to undersaturated
} } LaB6 cathode, a possibility which we have clearly eliminated as a
} } possible source.
} }
} } Thanks for any hints our suggestions.
} }
} } Krassimir.
} } _______________________________________
} } Krassimir N. Bozhilov
} } Central Facility for Advanced Microscopy and Microanalysis
} } University of California
} } Riverside, CA 92521
} }
} } tel 951 827 2998
} } fax 951 827 2489
} } bozhilov-at-ucr.edu
} } _______________________________________
} }
} }
} }
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From: walck-at-southbaytech.com
Date: Tue, 8 May 2007 16:51:28 -0500
Subject: [Microscopy] Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Peter,
It's much easier to control a cleave in silicon the thinner the sample.
Back thin your sample using a lapping fixture to keep the sample parallel
faced. Then simply use a fine scribe to start and propagate your cleave.

Another option for you is to use the Tripod Polisher(R) for your cross
section. You could easily polish your samples to a specific site and
control the thickness of them very accurately. It would take you a little
longer than simply cleaving, but if you have to back thin your samples, it
would be a wash. If you need any literature on the TP technique, please
contact me offline.

Disclaimer: South Bay Technology manufactures and sells the Tripod
Polisher(R).



-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
Sent: Tuesday, May 08, 2007 10:37 AM
To: Walck-at-SouthBayTech.com

Microscopy Folks,

I'd like to get some input on sample preparation with respect to silicon.

PROBLEM:

I have an FEI XL-50 FESEM that was originally designed to accept flat
samples, essentially silicon wafers. It does not have what one would
consider a typical exchange port. The exchange port is robotically
controlled, and the maximum sample height, including stub, must be 5 mm or
less. I must view these samples on edge, i.e. 90 degrees. This presents a
problem in that I must cleave two parallel sections very close to each
other. Perhaps diamond cutters can do this, but my hands are too shaky.

QUESTION:

Is there a device, or method, that would allow me to make these cleaves ~ 8
mm apart with any reasonable control? I found stubs that are low profile, 38
and 90 degrees, from the nice people at Ted Pella, but I still have this
issue of doing two parallel cleaves very close together.

Just for info. purposes I am in the silicon MEMS development arena.

If you feel your reply is of general interest to this community, please
reply to all, or you may contact me directly.

Regards to all in this small world,

Peter Tomic
Renaissance Wireless Corp.


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From: dsherman-at-purdue.edu
Date: Tue, 8 May 2007 17:21:47 -0500
Subject: [Microscopy] Plant fixation-pipes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all,

At the request of an investigator who was interested in plant microtubules,
we fixed arabidopsis hypocotyles in:

2.5% formaldehyde, 2% glutaraldehyde,  in PEMT (100mM pipes-KOH, pH
6.9, 5mM EGTA, 2 mM MgCl2, 0.05% Triton X-100) for 1h at RT

This was followed by standard 2% OsO4, ETOH dehydration and embedding in
Spurr's resin.

The results were less than pleasing. The membranes seemed soft with little
crisp clarity anywhere. In contrast we fixed maize leaves using:

2.5% glut + 2% PAF in 0.1M cacodylate, pH 6.8, as the primary fix with the
rest the same as above and got great membranes, sharp chloroplast granae
stacks, etc.

Only real difference was the buffer. Have any of you used a similar pipes
buffer and do you have any comments/ideas of why the one prep was good and
the other not?

Thanks,
Debby

Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.agriculture.purdue.edu/microscopy



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11, 22 -- Subject: Plant fixation-pipes
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From: krensing-at-ucalgary.ca
Date: Tue, 8 May 2007 17:40:14 -0500
Subject: [Microscopy] Re: Plant fixation-pipes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Debby,
The first thing that I saw was Triton X-100! I don't imagine that
including a surfactant with your primary fixative would be any good for
the membranes. I have used straight PIPES (without the additives)in
plant tissues with good results (but not better than with cacodylate or
phosphate buffers).
Kim

dsherman-at-purdue.edu wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi all,
}
} At the request of an investigator who was interested in plant microtubules,
} we fixed arabidopsis hypocotyles in:
}
} 2.5% formaldehyde, 2% glutaraldehyde, in PEMT (100mM pipes-KOH, pH
} 6.9, 5mM EGTA, 2 mM MgCl2, 0.05% Triton X-100) for 1h at RT
}
} This was followed by standard 2% OsO4, ETOH dehydration and embedding in
} Spurr's resin.
}
} The results were less than pleasing. The membranes seemed soft with little
} crisp clarity anywhere. In contrast we fixed maize leaves using:
}
} 2.5% glut + 2% PAF in 0.1M cacodylate, pH 6.8, as the primary fix with the
} rest the same as above and got great membranes, sharp chloroplast granae
} stacks, etc.
}
} Only real difference was the buffer. Have any of you used a similar pipes
} buffer and do you have any comments/ideas of why the one prep was good and
} the other not?
}
} Thanks,
} Debby
}
} Debby Sherman, Director Phone: 765-494-6666
} Life Science Microscopy Facility FAX: 765-494-5896
} Purdue University E-mail: dsherman-at-purdue.edu
} S-052 Whistler Building
} 170 S. University Street
} West Lafayette, IN 47907
} http://www.agriculture.purdue.edu/microscopy
}
}
}
} ==============================Original Headers==============================
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--
Kim Rensing Ph.D.
Assistant Research Professor
Manager, Microscopy and Imaging Facility

University of Calgary
Health Sciences Centre
B129 - 3330 Hospital Drive NW
Calgary, AB, Canada T2N 4N1
403-220-3488
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From: robinson32-at-sympatico.ca
Date: Tue, 8 May 2007 19:21:00 -0500
Subject: [Microscopy] EDS Repair

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

We have a Kevex Sigma Microanalyzer Level LPX3 which continually says that
it is receiving no x-rays. The detector has recently been rebuilt and the
system still does not work. Does anyone know of a company which repairs the
Kevex electronics on site in Canada?



==============================Original Headers==============================
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From: rosslm-at-missouri.edu
Date: Tue, 8 May 2007 23:10:52 -0500
Subject: [Microscopy] viaWWW: 5th Annual Short Course on Computer-Assisted Image

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Email: rosslm-at-missouri.edu
Name: Lou Ross

Organization: University of Missouri-Columbia

Title-Subject: [Filtered] 5th Annual Short Course on Computer-Assisted Image Analysis

Question: The Electron Microscopy Core Facility at the University of Missouri-Columbia is hosting the 5th Annual Short Course and Workshop on Computer-Assisted Image Analysis and Measurement taught by Dr. John C. Russ on June 27-29, 2007. This popular course is intended to familiarize users of image analysis hardware and software with the fundamental principles and methods available to obtain meaningful results.

Image analysis and measurement techniques are utilized in a broad range of applications and are usually concerned with extracting a numerical values (number, size, shape, etc.) or location of objects from the image. In other cases, global structural parameters such as the volume and surface of features are of interest. These measurements may require image processing to correct defects, enhance features, compare multiple images, recognize objects, or other steps. Measurements on these individual features, or on the image as a whole, must then be obtained and interpreted in a proper stereological context to obtain useful data about the objects. Statistical interpretation of the data allows comparisons of different populations, understanding of distribution plots, and other inferences about the original objects. Structural modeling and geometric probability can be used to develop models for this interpretation.

The course relies heavily on tightly coupled lectures and hands-on exercises covering a wide variety of methodologies, approaches and tools, through a set of practical, step-by-step instructions to minimize the learning curve. No specific background is assumed, although users should already be familiar with microscopy or other imaging technologies. The software platform for the examples and exercises is Adobe Photoshop utilizing a comprehensive set of plug-ins from Reindeer Graphics. Included with the registration fee is a half-day primer on Photoshop prior to the beginning of the course.

Dr. Russ is the internationally acclaimed author of innumerable articles and several books on image analysis techniques and digital imaging, including the The Image Processing Handbook. He is widely known for his workshops and short courses and has helped to develop software packages to make image analysis methods more accessible to non-specialists. Many of the examples used in the course involve light or electron microscope images, but students are invited to bring their own images (TIFF files) for discussion and analysis. Ample time is allotted at the end of the course for individual instruction.

The registration fee is $1100. Enrollment is limited to 20 attendees and there are still a few openings. More information can be found at:
http://www.emc.missouri.edu/works.htm or by contacting the course coordinator Lou Ross at 573.882.4777 or rosslm-at-missouri.edu.


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From: frederic.diana-at-gmail.com
Date: Tue, 8 May 2007 23:12:51 -0500
Subject: [Microscopy] viaWWW: Autofocusing system

Contents Retrieved from Microscopy Listserver Archives
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Email: frederic.diana-at-gmail.com
Name: Frederic

Organization: UCSB

Title-Subject: [Filtered] Autofocusing system

Question: Hi everyone!

I am looking for an autofocusing system which would be used for photoluminescence mapping. Basically I am looking for a system which can translate an objective lens to adjust the focus on the sample.

Thank you for your suggestions and help.

Frederic

---------------------------------------------------------------------------

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From: jacques.faerber-at-ipcms.u-strasbg.fr
Date: Wed, 9 May 2007 02:13:15 -0500
Subject: [Microscopy] Re : Silicon Cross-section sample preparation]

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all

Depending of the size of the wafer, cristallographic oriention and
direction you want to cut it, a solution could be to use a diamant wire
saw, either to cut the sample or, better, to cut only two groves, where
the silicon will (more or less cleanly, (100) works easier than (111))
clive. On such machines,it's very simple to cut parallel groves. Of
coarse, if you have a 8" wafer, it will be less easy to find a saw which
is big enought !

An other solution, if you have access to a power laser light, from the
kind used for PLD (pulsed laser deposition), is to draw such groves with
the laser. By that way you don't have any mechanical stress on the
sample, and it will clive or brake very easy along the groves. It's very
practical to have a wafer drawn with a grid patern, where one can brake
squares or rectangles samples on demand. Of coarse, cristallography has
its laws, and it brakes not always where one want !

Of coarse, in both cases, one makes the groves on the back side of the
sample.

Hope it helps

--
J. Faerber
IPCMS-GSI
(Institut de Physique et Chimie des Matériaux de Strasbourg
Groupe Surface et Interfaces)
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France


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Microscopy Folks,

I'd like to get some input on sample preparation with respect to silicon.

PROBLEM:

I have an FEI XL-50 FESEM that was originally designed to accept flat
samples, essentially silicon wafers. It does not have what one would
consider a typical exchange port. The exchange port is robotically
controlled, and the maximum sample height, including stub, must be 5 mm
or less. I must view these samples on edge, i.e. 90 degrees. This
presents a problem in that I must cleave two parallel sections very
close to each other. Perhaps diamond cutters can do this, but my hands
are too shaky.

QUESTION:

Is there a device, or method, that would allow me to make these cleaves
~ 8 mm apart with any reasonable control? I found stubs that are low
profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
still have this issue of doing two parallel cleaves very close together.

Just for info. purposes I am in the silicon MEMS development arena.

If you feel your reply is of general interest to this community, please
reply to all, or you may contact me directly.

Regards to all in this small world,

Peter Tomic
Renaissance Wireless Corp.


--
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==============================Original Headers==============================
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Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail Jacques.Faerber-at-ipcms.u-strasbg.fr


==============================Original Headers==============================
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From: richard.beanland-at-bookham.com
Date: Wed, 9 May 2007 04:25:50 -0500
Subject: [Microscopy] Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Peter,
I know you were working on III-V's a few years back so cleaving
Si seems a lot more difficult in comparison. I have found the best way
to cleave silicon wafers is rather different to GaAs or InP. Perhaps
this is general knowledge and you know this already, but I hope it's
worth sharing with the list if nothing else.
The problem is that Si prefers to cleave along (111) rather than (110)
and so you get an angled face on the cleave, which is usually rather
uneven and often doesn't run straight. This is even worse when you are
cleaving close to an existing edge, which attracts the crack front as it
propagates (good for making low-angle cleaved specimens, but a problem
for what you are trying to do).
It is possible to make Si cleave along (110) by cleaving the wafer
without any support. I suspect this works because the crack is
propagating supersonically or something like that; I read a description
of the technique in a paper from the 1970's but I can't remember the
reference.
So, to cleave Si along (110): make a single scribe mark on the top
surface with a good sharp diamond a couple of mm long at the edge of the
wafer. Then, hold the wafer just between forefinger and thumb in both
hands, with the top wafer surface under your fingers and the scribe mark
between the tips of your fingers. Put a thumbnail under the scribe mark
and then bend the wafer down, pulling apart slightly at the same time.
If it cleaves well, it will do so very quickly with an audible 'ping'.
It's a good idea to do this over a large clean surface in case you drop
either part.
This is not so hard to do on a whole wafer (although trying not to
drop an 8" wafer is fun, I haven't tried 300mm), but as the pieces get
smaller it gets more difficult. It should be possible to get an 8mm
wide strip, but a 5mm strip might be difficult or impossible. You could
back thin the wafer but of course this isn't straightforward for
something } 1" in diameter, and any scratches on the back might make the
cleave deviate from its path. Like a lot of these things it's a lot
easier to demonstrate than describe in text, and it takes a little
practice to get the hang of it, so I would try on a few spare wafers
first. I guess you could cleave a 10mm wide strip and grind it down to
a couple of mm before mounting it for SEM.

Good luck!

Richard

________________________________________
Richard Beanland
Materials Analysis
Bookham
Caswell
Towcester
Northants
NN12 8EQ
United Kingdom
Tel. +44 1327 356362
Fax. +44 1327 356775
http://www.bookham.com
________________________________________

-----Original Message-----
X-from: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
Sent: 08 May 2007 18:35
To: Richard Beanland

Microscopy Folks,

I'd like to get some input on sample preparation with respect to
silicon.

PROBLEM:

I have an FEI XL-50 FESEM that was originally designed to accept flat
samples, essentially silicon wafers. It does not have what one would
consider a typical exchange port. The exchange port is robotically
controlled, and the maximum sample height, including stub, must be 5 mm
or less. I must view these samples on edge, i.e. 90 degrees. This
presents a problem in that I must cleave two parallel sections very
close to each other. Perhaps diamond cutters can do this, but my hands
are too shaky.

QUESTION:

Is there a device, or method, that would allow me to make these cleaves
~ 8 mm apart with any reasonable control? I found stubs that are low
profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
still have this issue of doing two parallel cleaves very close together.

Just for info. purposes I am in the silicon MEMS development arena.

If you feel your reply is of general interest to this community, please
reply to all, or you may contact me directly.

Regards to all in this small world,

Peter Tomic
Renaissance Wireless Corp.


--
This email message is for the sole use of the intended recipient(s) and
may contain confidential and privileged information. Any unauthorized
review, use, disclosure or distribution is prohibited. If you are not
the intended recipient, please contact the sender by reply email and
destroy all copies of the original message. If you are the intended
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==============================Original Headers==============================
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From: peter.tomic-at-renwireless.com
Date: Wed, 9 May 2007 08:01:30 -0500
Subject: [Microscopy] Re: Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html



Richard,

Yes, III-V compounds cleave so much easier, but now I'm working on Si
MEMS. There's virtually no heat dissipation, therefore substrate
thickness isn't an issue.

I've gotten several good suggestions, including yours. For the list,
they are;

1. Laser serration

2. Thinning the Si [My samples are usually .025" thick.]

3. Wire Saw

4. Pre-scribing before cleaving

One other caveat in all this sample preparation is that the device
constructed on top of the substrate is amorphous.

When all else fails, use an FIB, I always say.

I never get the easy problems.

Thanks to all of you.

Peter



Richard Beanland wrote:
} Hi Peter,
} I know you were working on III-V's a few years back so cleaving
} Si seems a lot more difficult in comparison. I have found the best way
} to cleave silicon wafers is rather different to GaAs or InP. Perhaps
} this is general knowledge and you know this already, but I hope it's
} worth sharing with the list if nothing else.
} The problem is that Si prefers to cleave along (111) rather than (110)
} and so you get an angled face on the cleave, which is usually rather
} uneven and often doesn't run straight. This is even worse when you are
} cleaving close to an existing edge, which attracts the crack front as it
} propagates (good for making low-angle cleaved specimens, but a problem
} for what you are trying to do).
} It is possible to make Si cleave along (110) by cleaving the wafer
} without any support. I suspect this works because the crack is
} propagating supersonically or something like that; I read a description
} of the technique in a paper from the 1970's but I can't remember the
} reference.
} So, to cleave Si along (110): make a single scribe mark on the top
} surface with a good sharp diamond a couple of mm long at the edge of the
} wafer. Then, hold the wafer just between forefinger and thumb in both
} hands, with the top wafer surface under your fingers and the scribe mark
} between the tips of your fingers. Put a thumbnail under the scribe mark
} and then bend the wafer down, pulling apart slightly at the same time.
} If it cleaves well, it will do so very quickly with an audible 'ping'.
} It's a good idea to do this over a large clean surface in case you drop
} either part.
} This is not so hard to do on a whole wafer (although trying not to
} drop an 8" wafer is fun, I haven't tried 300mm), but as the pieces get
} smaller it gets more difficult. It should be possible to get an 8mm
} wide strip, but a 5mm strip might be difficult or impossible. You could
} back thin the wafer but of course this isn't straightforward for
} something } 1" in diameter, and any scratches on the back might make the
} cleave deviate from its path. Like a lot of these things it's a lot
} easier to demonstrate than describe in text, and it takes a little
} practice to get the hang of it, so I would try on a few spare wafers
} first. I guess you could cleave a 10mm wide strip and grind it down to
} a couple of mm before mounting it for SEM.
}
} Good luck!
}
} Richard
}
} ________________________________________
} Richard Beanland
} Materials Analysis
} Bookham
} Caswell
} Towcester
} Northants
} NN12 8EQ
} United Kingdom
} Tel. +44 1327 356362
} Fax. +44 1327 356775
} http://www.bookham.com
} ________________________________________
}
} -----Original Message-----
} From: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
} Sent: 08 May 2007 18:35
} To: Richard Beanland
} Subject: [Microscopy] Silicon Cross-section sample preparation
}
}
}
}
} ------------------------------------------------------------------------
} ----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
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} http://www.microscopy.com/MicroscopyListserver
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} ------------------------------------------------------------------------
} ----
}
} Microscopy Folks,
}
} I'd like to get some input on sample preparation with respect to
} silicon.
}
} PROBLEM:
}
} I have an FEI XL-50 FESEM that was originally designed to accept flat
} samples, essentially silicon wafers. It does not have what one would
} consider a typical exchange port. The exchange port is robotically
} controlled, and the maximum sample height, including stub, must be 5 mm
} or less. I must view these samples on edge, i.e. 90 degrees. This
} presents a problem in that I must cleave two parallel sections very
} close to each other. Perhaps diamond cutters can do this, but my hands
} are too shaky.
}
} QUESTION:
}
} Is there a device, or method, that would allow me to make these cleaves
} ~ 8 mm apart with any reasonable control? I found stubs that are low
} profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
} still have this issue of doing two parallel cleaves very close together.
}
} Just for info. purposes I am in the silicon MEMS development arena.
}
} If you feel your reply is of general interest to this community, please
} reply to all, or you may contact me directly.
}
} Regards to all in this small world,
}
} Peter Tomic
} Renaissance Wireless Corp.
}
}

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From: MCarlyle-at-veeco.com
Date: Wed, 9 May 2007 10:57:20 -0500
Subject: [Microscopy] SPM Users present cutting edge research

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Seeing at the Nanoscale V Conference

June 24-27. 2007
University of California
Santa Barbara

Register now and join other users of scanning probe microscopy as they
meet and discuss their work informally with colleagues from all over the
world at the fifth annual Seeing at the Nanoscale Conference, University
of California, Santa Barbara. Sponsored by Veeco Instruments and the
California NanoSystems Institute (CNSI), the conference includes
two-and-one-half day technical presentations and a poster session on the
following topics:

* Extending the Limits of SPM: High Speed Scanning, Ultra High
Resolution Imaging, Multiple Probe SPM

* From Single Biomolecules To Cells: Using AFM and Combined
AFM-Optical Techniques to Probe Biological Structures and Forces

* Next Generation Materials and Polymer Systems

* Beyond Topography: Measurement of Physical Properties at the
Nanoscale - Nanomechanical, Electrical, Optical, Magnetic and Thermal

* Instruments and Probes - New Tools and Techniques for
Nanoscience

For more information and to register, please go to
www.veeco.com/Nanoconference





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From: gary-at-gaugler.com
Date: Wed, 9 May 2007 11:34:25 -0500
Subject: [Microscopy] Re: Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Depending on the size of the wafer, it will have either
a flat or a notch on one place. This is the key to being
parallel with the crystal lattice. So, align this
key and use a carbide scribe (local hardware store or
Home Depot--$5) and make two linear scribes then two
lateral ones to get two small pieces of die. Mount
on the Pella stubs and you should be good to go.

gary g.



At 09:35 AM 5/8/2007, you wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America


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From: TrogadisJ-at-smh.toronto.on.ca
Date: Wed, 9 May 2007 11:49:31 -0500
Subject: [Microscopy] microscopy-macroscopy

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear List:

During the last year I have attended sevaral demonstrations of whole
slide image capture devices. For example, the TISSUEscope (Biomedical
Photometrics) describes brightfield, fluorescence and confocal imaging
and others - the Aperio - does brightfield imaging. There are others.

Does this emerging technology have a place in a core imaging facility -
in other words, do users find that it replaces some other microscope
modalities, e.g. I can see it very useful for measuring the frequency of
rare events since they can see the entire slide.

My worry is that the data files are huge. Can analysis packages handle
it easily? Do all computers have to be updated (64-bit, limitless RAM)?
High throughput analysis to me implies speedy analysis, don't want to
watch endless loading times, computer crashes.

I am curious about hearing any experience from users of whole slide
scanners.

Thank you.
Judy


Judy Trogadis
Bio-Imaging Coordinator
St. Michael's Hospital, 7Queen
30 Bond St.
Toronto, ON M5B 1W8, Canada
ph: 416-864-6060 x6337
pager: 416-685-9219
fax: 416-864-6043
trogadisj-at-smh.toronto.on.ca


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From: Susan.Kent-at-us.contiautomotive.com
Date: Wed, 9 May 2007 14:11:44 -0500
Subject: [Microscopy] XPS/ESCA: Need Help with Sun Hardware and Software

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Hi everyone!

I'm in desperate need of a copy of Sun Solaris OS 4.0.2 and a hard drive
for a Sun SPARC IPC workstation. We have a Kratos XSAM 800 which is
controlled by the SPARC computer, but unfortunately the computer has
malfunctioned and we have no backup copies of the OS (the instrument was
purchased "used"). I've already contacted Sun but they haven't been able
to help. and I haven't received a reply from Kratos.

Can anyone on the list help?

Thanks in advance.

--Sue Kent


Susan M. Kent
Principal Staff Scientist
Continental AG
Automotive Systems Division
21440 W. Lake Cook Road
Deer Park, IL 60010
847-862-0216 (Desk)
847-343-5145 (Mobile)
847-862-8330 (Fax)
Email: Susan.Kent-at-us.contiautomotive.com
www.contiautomotive.com



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From: jae5-at-lehigh.edu
Date: Wed, 9 May 2007 14:29:21 -0500
Subject: [Microscopy] 9th InterAmerican Congress on Electron Microscopy; Cusco, Peru

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Reminder: Abstracts due this month.

9th InterAmerican Congress on Electron Microscopy

Cusco Peru

September 23 - 28, 2007

One of the major electron microscopy meetings this year will
be held in the heart of the Inca empire. Machu Picchu, one
of the world's most famous and wondrous places is nearby.

Leading microscopists from across the Americas (and from
the rest of the world) will be presenting their latest work.

This is a reminder that abstracts (in a two-page format, very
similar to the M and M format) are due by the end of this
month (May, 2007).

Details of how to register (which must be done before the
abstract is submitted) and of how to submit the abstract,
can be found at the web site:
http://www.ciasem2007.com/
(To change to English click the little blue button to the right.)
The web site also lists invited speakers and gives details of the program.
--
...........
Alwyn Eades
Department of Materials Science and Engineering
Lehigh University
5 East Packer Avenue
Bethlehem
Pennsylvania 18015-3195
Phone 610 758 4231
Fax 610 758 4244
jae5-at-lehigh.edu


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From: dac-at-research.umass.edu
Date: Wed, 9 May 2007 15:29:53 -0500
Subject: [Microscopy] Gelatin subbed slides - eliminate the chromium?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi everyone,

I'm making subbed slides and just wondering if there is anyone who knows
the chemistry/function of the chromium potassium sulfate (chrome alum)
in the mix, and if there is a new-age environmentally-friendly
substitute for it, or can it be eliminated - to what effect? I'm working
on the assumption that it has a function. I know that only small
quantities of Chrome alum are used, but would like to eliminate it if
possible.

To avoid too much traffic about alternatives, let me say I've tried a
number of other things for getting epoxy semi-thins to stick to glass,
and this method works for me. I did already see a note in the archives
about a "pinch of gelatine in the water bath" and that doesn't have the
chromium compound. I've also read an alternative use of the Mayer's egg
albumin - instead of smearing the slide and drying, adding some to the
flotation water - and that also doesn't contain chromium. I like the way
water drops sit nicely on the subbed slide so rows of sequential
sections can be arrayed in the droplets without mixing.


Thanks in advance for any insights.


Dale Callaham

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From: jpchandl-at-mines.edu
Date: Wed, 9 May 2007 15:42:15 -0500
Subject: [Microscopy] Looking for recommendations about dual beam ion mills

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I need to tap in to the wisdom of the list about dual beam argon ion mills.

We have a Bal-Tec RES100 ion mill that we are having difficulty maintaining.
It had been in disuse for a couple of years, and has been repaired and
upgraded recently, at the factory, to be as close to a RES101 as possible.
We are having issues such as the guns becoming contaminated and needing
service much more frequently than we think they should. Routine maintenance
seems to be difficult, too. We think we are looking at a high maintenance
instrument here. I would like to know what kind of experience other labs
have had with this instrument.

I would also like to hear from people who have experience using/maintaining
the RES100/101 or the Gatan PIPS, or both. We need some comparative
information, so we can make a decision about how to proceed with our ion
milling needs.

Any input would be appreciated.

--John

John Chandler
Manager, EM Lab
Colorado School of Mines
Golden, CO 80401
jpchandl-at-mines.edu
303-384-2203




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From: phillipst-at-missouri.edu
Date: Wed, 9 May 2007 15:46:12 -0500
Subject: [Microscopy] Re: Gelatin subbed slides - eliminate the

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

If you haven't tried coating your slides with 1% aminopropyltriethoxysilane
(APTS) in water then you are missing out on a much better alternative. The
water really beads up on it. I used to use chrome-gel but it doesn't compare.


At 03:30 PM 05/09/07, you wrote:



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573-882-0123 (fax)
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From: lcgould-at-med.cornell.edu
Date: Wed, 9 May 2007 15:51:50 -0500
Subject: [Microscopy] Re: Gelatin subbed slides - eliminate the chromium?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dale-
Have you tried the SuperFrost Plus slides? They are pre-treated
with something proprietary that helps sections adhere. They can be
purchased from any number of vendors, although I believe that they
all come from the same manufacturer.
Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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From: larry-at-celtic.freewire.co.uk
Date: Wed, 9 May 2007 16:05:25 -0500
Subject: [Microscopy] Re: Diffracation ring problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

} } } Hi all,
} } }
} } } I am trying to find resolution of a problem with our CM300 TEM with a
} } } LaB6 cathode.
} } }
} } } When trying to get diffraction from an almost amorphous material I
} } } have noticed a presence of relatively weak but clearly visible
} } } perfectly circular ring of intensity around the central transmitted
} } } spot. The ring is not centered around the transmitted spot. Its
} } } position varies when operating the beam shift controls, its size
} } } varies with the C2 aperture size when in diffraction mode. It is also
} } } present when there is no sample under the beam. When there is
} } } strongly reflecting crystalline material the ring is almost invisible
} } } due to its weak relative intensity. It is present at any accelerating
} } } voltage.
} } }
} } } I am trying to get some ideas about what might be causing the ring
} } } and how to eliminate it.
} } }
} } } The ring is also visible when in LM imaging mode (image is formed by
} } } the diffraction lens)
} } } and when the beam is focused to a spot. In LM mode the ring has
} } } strange shape. It has sharp circular outline on its outer edge and it
} } } has irregular shape and outline on its inner edge.
} } }
} } } Using the free lens control option I have made the following
} } } observations:
} } }
} } } In LM imaging mode with beam focused to a spot.
} } } When changing the C1 lens current -the ring changes size and focus
} } } but does not rotate.
} } } changing C2 lens current - change in focus only, plus some rotation
} } } changing Twin lens current - change of size and focus, no rotation
} } } changing Objective lens current - change in size and focus, some
} } } rotation
} } } changing Diffraction lens current - change in size and focus, no
} } } rotation
} } } changing Intermediate lens current - change in size and focus, no
} } } rotation
} } } changing P1 lens current - change in size and focus, some rotation
} } } changing P2 lens current - rotation only
} } }
} } } In diffraction mode with fully spread beam:
} } } position varies when operating the beam shift controls
} } } size varies with the C2 aperture size.
} } }
} } } FEI support engineers have not been succesful in identifing the
} } } problem so far. It was suggested that it is due to undersaturated
} } } LaB6 cathode, a possibility which we have clearly eliminated as a
} } } possible source.
} } }
} } } Thanks for any hints our suggestions.
} } }
} } } Krassimir.
} } } _______________________________________
} } } Krassimir N. Bozhilov
} } } Central Facility for Advanced Microscopy and Microanalysis
} } } University of California
} } } Riverside, CA 92521
} } }
} } } tel 951 827 2998
} } } fax 951 827 2489
} } } bozhilov-at-ucr.edu
} } } _______________________________________
} } }
} On May 8, 2007, at 12:46 PM, randerson20-at-tampabay.rr.com wrote:
} }
} }
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} }
} } Consider the possibility that it might be visible light from the hot
} } cathode coming down the column, i.e. like a flashlight.
} }
} } This argument is abetted by the sharp outside edge, from apertures
} } along
} } the way, and a diffuse inner edge and the fact that you see it
} } without a
} } specimen loaded. However, it isn't immediately obvious how or why it
} } changes size and shape with all of your other perturbations.
} }
} } Can you partly eclipse it by moving apertures around?
} }
} } Ron Anderson
} }
} } bozhilov-at-ucr.edu wrote:
} } } ---------------------------------------------------------------------
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The fact that the ring changes as the lenses are adjusted suggests
that it is not anything to do with light from the filament.

Since the ring changes in dia. as the C2 aperture is changed suggests
that it is something to do with the C2 aperture.

Do you have an unusually high X-ray background?

Do you clean your C2 apertures yourself?

I'm thinking that the C2 apertures have been overheated in the
cleaning process, leading to recrystalisation and a rough edge to the
inside bore of the apertures?

You might be able to confirm this by doing convergent beam
diffraction - the ragged edge of the C2 aperture should be visible in
the CBDP.

Regards,
--
Larry Stoter
JEOL (UK) Ltd
tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoluk.com

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From: thoward-at-unm.edu
Date: Wed, 9 May 2007 18:14:55 -0500
Subject: [Microscopy] Re: Gelatin subbed slides - eliminate the

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dale -

You do not need the chrome alum, but the slides will be
"stickier" with it. I have no idea why....

I'd agree with the rest of the crowd about APS or Plus
slides, but sometimes they just aren't enough & subbing is
still the way to go, horrible as the process may be.

Tamara

On Wed, 9 May 2007 15:31:13 -0500
dac-at-research.umass.edu wrote:
}
}
}
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} Society of America
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}
} Hi everyone,
}
} I'm making subbed slides and just wondering if there is
} anyone who knows
} the chemistry/function of the chromium potassium sulfate
} (chrome alum)
} in the mix, and if there is a new-age
} environmentally-friendly
} substitute for it, or can it be eliminated - to what
} effect? I'm working
} on the assumption that it has a function. I know that
} only small
} quantities of Chrome alum are used, but would like to
} eliminate it if
} possible.
}
} To avoid too much traffic about alternatives, let me say
} I've tried a
} number of other things for getting epoxy semi-thins to
} stick to glass,
} and this method works for me. I did already see a note
} in the archives
} about a "pinch of gelatine in the water bath" and that
} doesn't have the
} chromium compound. I've also read an alternative use of
} the Mayer's egg
} albumin - instead of smearing the slide and drying,
} adding some to the
} flotation water - and that also doesn't contain
} chromium. I like the way
} water drops sit nicely on the subbed slide so rows of
} sequential
} sections can be arrayed in the droplets without mixing.
}
}
} Thanks in advance for any insights.
}
}
} Dale Callaham
}
} ==============================Original
} Headers==============================
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} 2007
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***************************
Tamara Howard
Cell Biology & Physiology
UNM-HSC
Albuquerque, NM
***************************

==============================Original Headers==============================
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From: nizets2-at-yahoo.com
Date: Thu, 10 May 2007 06:15:21 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial cristae

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Tom,

I agree with the other comments: Ferrocyanide,
decreasing dehydration times (penetration is immediate
in monolayers) and using picric acid probably can
improve membrane contrast.
(personally I already tried UAc in methanol without
satisfaction)

Please share your experience with us.

Best regards,

Stephane

--- tbargar-at-unmc.edu wrote:

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}
} Dear listers,
}
} Anyone out there have any advice on how to enhance
} the contrast and
} definition of the membranes of the cristae of
} mitochondria? The samples
} brought to me are monolayer cell cultures of cancer
} cells grown on
} Thermanox coverslips. This is how I'm currently
} processing the samples:
} Primary fixation is 2% glutaraldehyde, 2%
} paraformaldehyde, 0.5% acrolein
} in 0.1M Sorensen's phosphate buffer pH 7.2.
} Post-fixation in 1%Osmium
} Tetroxide in 0.1M Sorensen's phosphate buffer.
} Dehydration in 50, 70, 90,
} 95, 100%X3 ethanol solutions. embedding in
} Araldite. Sections are stained
} with 2% uranyl acetate aqueous 15 minutes and
} Reynold's lead citrate 10
} minutes. The density of the cytoplasm and
} mitochondrial matrix are
} similar with the result that the contrast of the
} mitochondria is similar to
} the cytoplasm. The mitochondria and it's membranes
} (outer and that of the
} cristae) don't really "stand out". The researchers
} involved want to see
} really contrasty mitochondrial cristae.
}
} The next thing I'm going to try is post-fixation
} with a mix of osmium
} tetroxide and potassium ferrocyanide.
}
} Anyone have any other ideas? Thanks in advance.
}
}
} Tom Bargar
} University of Nebraska Medical Center
} Core Electron Microscopy Research Facility
} 986395 Nebraska Medical Center
} Omaha, NE 68198-6395
} 402-559-7347
} tbargar-at-unmc.edu
}
}
} ==============================Original
} Headers==============================
} 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39
} 2007
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} Mitochondrial cristae
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From: nealzimm-at-cpinternet.com
Date: Thu, 10 May 2007 08:08:28 -0500
Subject: [Microscopy] Re: Looking for recommendations about dual beam ion

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Tom

I just thought I should mention something that I think I read years ago
in one of M.A. Hayat's books. He mentions that cacodylate and some other
buffers when incorporated into a fixative produce higher contrast
because they are more extractive whereas phosphate buffers probably
preserve cell contents better but at the expense of contrast.

I just thought that I should add this because you mention Sorenson's
buffer in the fixative. I'm sure that many of the other points have an
effect but just wondered if the buffer could be contributing to your
problems as well.

Malcolm


Malcolm Haswell
e.m. unit
School of Health, Natural and Social Sciences
Fleming Building
University of Sunderland
Tyne & Wear
SR1 3SD
UK
e-mail: malcolm.haswell-at-sunderland.ac.uk


----- Original Message -----
X-from: nizets2-at-yahoo.com

John

I work with four Gatan PIPS daily and have found them to be quite
acceptable in ease of both maintenance and use.

We currently do maintenance only when the ion guns stop functioning,
at that point the machine is taken out of service and a full cleaning
performed. This happens about every five to six months and leaves
the PIPS out of service for two days (One day to clean, pump down
over night, and alignment the next day). The diaphragm pump needs
new diaphragms once a year (about an hours work) but not too much else.

Work load is typically one to two ceramic samples per day, with an
occasional silicon or glass sample thrown in. The ceramic samples
are lapped to about six microns and mill for an hour to two hours.
Silicon and glass are left about ten microns thick and take much
longer (up to three days), not so much due to the thickness as to
the low kVs used and letting the sample "cool".

Gatan offers a digital camera/zoom lens set and an external diaphragm
pump as extras. In my opinion, they are a "must have".

If you have any further questions please feel free to contact me.

Neal R. Zimmermann



On Wed, 2007-05-09 at 15:43 -0500, jpchandl-at-mines.edu wrote:
}
}
} ----------------------------------------------------------------------------
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} I need to tap in to the wisdom of the list about dual beam argon ion mills.
}
} We have a Bal-Tec RES100 ion mill that we are having difficulty maintaining.
} It had been in disuse for a couple of years, and has been repaired and
} upgraded recently, at the factory, to be as close to a RES101 as possible.
} We are having issues such as the guns becoming contaminated and needing
} service much more frequently than we think they should. Routine maintenance
} seems to be difficult, too. We think we are looking at a high maintenance
} instrument here. I would like to know what kind of experience other labs
} have had with this instrument.
}
} I would also like to hear from people who have experience using/maintaining
} the RES100/101 or the Gatan PIPS, or both. We need some comparative
} information, so we can make a decision about how to proceed with our ion
} milling needs.
}
} Any input would be appreciated.
}
} --John
}
} John Chandler
} Manager, EM Lab
} Colorado School of Mines
} Golden, CO 80401
} jpchandl-at-mines.edu
} 303-384-2203
}
}
}
}
} ==============================Original Headers==============================
} 9, 19 -- From jpchandl-at-mines.edu Wed May 9 15:42:15 2007
} 9, 19 -- Received: from inspire.Mines.EDU (inspire.Mines.EDU [138.67.130.5])
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} 9, 19 -- Subject: Looking for recommendations about dual beam ion mills
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==============================Original Headers==============================
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From: Andrew.Bowling-at-ARS.USDA.GOV
Date: Thu, 10 May 2007 08:38:50 -0500
Subject: [Microscopy] RE: Gelatin subbed slides - eliminate the chromium?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I got my recipe for chrome-alum from an old copy of Berlin and Miksche's
"Botanical Microtechnique and Cytochemistry". In the chapter covering
microtomy, in the section under adhesives, there is a subsection for
gelatin adhesives. The first recipe is for "Haupt's adhesive". This is
prepared by dissolving 1g of gelatine in 100 cc water, smearing a thin
film of this onto slides, and then flooding with 4% formalin. The
second recipe is for the chorme-alum that we all know and love. The
paragraph which introduces the chrome-alum adhesive technique begins: "A
gelatin adhesive that does not use formalin may be prepared and used as
follows...".

I know I'm telling this poorly, but the point is that I think the
chrome-alum is used to "fix" the gelatine in place in a manner analogous
to what the formaldehyde would do in the Haupt's solution. This is the
purpose (I believe) in using chrome in the tanning industry.
Furthermore, the treatment of the gelatine layer with formaldehyde or
chrome would then leave free aldehydes/uncomplexed chrome atoms embedded
in the gelatine which might then react with your sections, adhering them
to the slide. Using glutaraldehyde instead of formaldehyde might even
provide a stronger bond, if that is what you decide to go with.

So here is a possible reason for the chrome-alum and a possible
(although old-school) alternative that still uses gelatine as the
adhesive but does not contain chromium.

Andy Bowling
Plant Physiologist
USDA-ARS-SWSRU


==============================Original Headers==============================
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From: richard.beanland-at-bookham.com
Date: Thu, 10 May 2007 09:09:28 -0500
Subject: [Microscopy] Re: Looking for recommendations about dual beam

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Neal,
I wouldn't expect a 10um Si sample to take more than 30-60 mins
to thin in a PIPS.
I can only think you are working at a relatively high milling angle
(} 5 degrees) and so have to turn the beam energy down very low so you
don't damage your sample. Why not experiment with one sample using full
power (6kV) and 3 degrees incidence angle, finishing off with 10 mins at
2kV? It could increase your throughput more than 20 times!

Richard



________________________________________
Richard Beanland
Materials Analysis
Bookham
Caswell
Towcester
Northants
NN12 8EQ
United Kingdom
Tel. +44 1327 356362
Fax. +44 1327 356775
http://www.bookham.com
________________________________________


-----Original Message-----
X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
Sent: 10 May 2007 14:10
To: Richard Beanland

John

I work with four Gatan PIPS daily and have found them to be quite
acceptable in ease of both maintenance and use.

We currently do maintenance only when the ion guns stop functioning,
at that point the machine is taken out of service and a full cleaning
performed. This happens about every five to six months and leaves
the PIPS out of service for two days (One day to clean, pump down
over night, and alignment the next day). The diaphragm pump needs
new diaphragms once a year (about an hours work) but not too much else.

Work load is typically one to two ceramic samples per day, with an
occasional silicon or glass sample thrown in. The ceramic samples
are lapped to about six microns and mill for an hour to two hours.
Silicon and glass are left about ten microns thick and take much
longer (up to three days), not so much due to the thickness as to
the low kVs used and letting the sample "cool".

Gatan offers a digital camera/zoom lens set and an external diaphragm
pump as extras. In my opinion, they are a "must have".

If you have any further questions please feel free to contact me.

Neal R. Zimmermann



On Wed, 2007-05-09 at 15:43 -0500, jpchandl-at-mines.edu wrote:
}
}
}
------------------------------------------------------------------------
----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
America
} To Subscribe/Unsubscribe --
http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
}
------------------------------------------------------------------------
----
}
} I need to tap in to the wisdom of the list about dual beam argon ion
mills.
}
} We have a Bal-Tec RES100 ion mill that we are having difficulty
maintaining.
} It had been in disuse for a couple of years, and has been repaired and
} upgraded recently, at the factory, to be as close to a RES101 as
possible.
} We are having issues such as the guns becoming contaminated and
needing
} service much more frequently than we think they should. Routine
maintenance
} seems to be difficult, too. We think we are looking at a high
maintenance
} instrument here. I would like to know what kind of experience other
labs
} have had with this instrument.
}
} I would also like to hear from people who have experience
using/maintaining
} the RES100/101 or the Gatan PIPS, or both. We need some comparative
} information, so we can make a decision about how to proceed with our
ion
} milling needs.
}
} Any input would be appreciated.
}
} --John
}
} John Chandler
} Manager, EM Lab
} Colorado School of Mines
} Golden, CO 80401
} jpchandl-at-mines.edu
} 303-384-2203
}
}
}
}
} ==============================Original
Headers==============================
} 9, 19 -- From jpchandl-at-mines.edu Wed May 9 15:42:15 2007
} 9, 19 -- Received: from inspire.Mines.EDU (inspire.Mines.EDU
[138.67.130.5])
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From: shu-at-caltech.edu
Date: Thu, 10 May 2007 09:52:34 -0500
Subject: [Microscopy] Etching of ion milled sample

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear all,
I heard some people to use a solution containing citric acid and H2O2 to
remove the damage during ion milling. Does anyone have experience with
this? What kind of damage it can remove, amorphorized pits, deposited
contamination? Does it require a following cleaning process? Can someone
with experience kindly give a detailed description of this technique?
Thanks

Shu Miao

==============================Original Headers==============================
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From: ahlst007-at-umn.edu
Date: Thu, 10 May 2007 10:11:07 -0500
Subject: [Microscopy] Re: Gelatin subbed slides - eliminate the chromium?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dale,

In your post below you stated:

"I've tried a number of other things for getting
epoxy semi-thins to stick to glass, "

You didn't state how thick nor what area your
sections are, but for me semi-thins are
nominally 2.0 microns thick and usually no
larger than 4 mm per side

I use no slide subbing method. I clean 1x3"
glass slides with an ethanol rinse, then air dry
at room temperature or blow down with hair dryer.

I then collect sections on a drop or two of
distilled water on the slide, transfered there
from the
microtome with a clean fine tipped artists
brush. I collect about 8-12 sections per drop.

Then I warm the slide beneath the water drop
from below using an alcohol lamp, fairly hot,
but not to boil, of course. After drying by
heating the sections stick quite well. I can
then stain, usually with 0.2 micron filtered
toluidine blue, again heating but gently this
time, for about a minute, until stain "develops"
the section. Then rinse that stain off with
distilled water from a squirt bottle, even
direct the spray right onto the sections to get
rid of any ppt. Dry again gently with flame -
then view on the microscope.

If you are cutting much thicker sections, say 10
micron or more, or sections have much larger
area, maybe this would not work, tho I have not
tried this method for those larger sections.

Hope this helps, but I have a sneaky feeling
that I've missed the point of what you are
trying to do and that some kind of subbing is
quite necessary for accomplishing your goals.
Anyway, thought I'd toss this out for what its
worth.

Gib
--
Gilbert (Gib) Ahlstrand, Electron Microscopist,
Imaging Center, University of Minnesota
123 Snyder Hall
St. Paul, MN 55108
http://www.cbs.umn.edu/ic

dac-at-research.umass.edu wrote:
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}
} Hi everyone,
}
} I'm making subbed slides and just wondering if there is anyone who knows
} the chemistry/function of the chromium potassium sulfate (chrome alum)
} in the mix, and if there is a new-age environmentally-friendly
} substitute for it, or can it be eliminated - to what effect? I'm working
} on the assumption that it has a function. I know that only small
} quantities of Chrome alum are used, but would like to eliminate it if
} possible.
}
} To avoid too much traffic about alternatives, let me say I've tried a
} number of other things for getting epoxy semi-thins to stick to glass,
} and this method works for me. I did already see a note in the archives
} about a "pinch of gelatine in the water bath" and that doesn't have the
} chromium compound. I've also read an alternative use of the Mayer's egg
} albumin - instead of smearing the slide and drying, adding some to the
} flotation water - and that also doesn't contain chromium. I like the way
} water drops sit nicely on the subbed slide so rows of sequential
} sections can be arrayed in the droplets without mixing.
}
}
} Thanks in advance for any insights.
}
}
} Dale Callaham
}

==============================Original Headers==============================
11, 21 -- From ahlst007-at-umn.edu Thu May 10 10:11:06 2007
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From: hyi-at-emory.edu
Date: Thu, 10 May 2007 10:40:47 -0500
Subject: [Microscopy] Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear All:

         I am so glad to see the discussion on the contrast issue
continues as I have been hearing more and more reports on this problem
and I myself have been experiencing it too. Many hypotheses have been
suggested on the causes of the problem but it seems that there were
always evidences supporting different arguments. Practically, besides
using short dehydration and infiltration times for monolayer cells, I
now HAVE TO use potassium ferrocyanide with OsO4 in order to get good
contrast (even for bulk tissue). When I find out the contrast is low
after sections are being cut, I re-counterstain sections with 15% UA in
methanol (it can give a muddy appearance if it is over done). In
addition, I make sure to use a relatively fresh OsO4 stock solution. 

         I have been working in EM core facilities for a long time and
have dealt with all kinds of samples. One question I have for people
like me is whether this is a new problem or has always been this way. I
did not remember contrast was so much an issue in the good old days
even with standard dehydration times and when OsO4 was used
alone. Personally I think this problem has something to do with the
reagents we use now days, particularly OsO4. I did notice the results
were different when I use different OsO4 (liquid vs. crystal, old vs.
fresh….). I am wondering what other people’s experiences are with OsO4
and how OsO4 solution is prepared and stored in different
laboratories. I would really love to hear from chemist more details on
how OsO4 works and from vendors if there has been any change in their
sources for OsO4.

        Thank all very much

Hong
Emory SOM EM


On May 10, 2007, at 8:10 AM, malcolm.haswell-at-sunderland.ac.uk wrote:
}
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} -----
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} America
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} Dear Tom
}
} I just thought I should mention something that I think I read years ago
} in one of M.A. Hayat's books. He mentions that cacodylate and some
} other
} buffers when incorporated into a fixative produce higher contrast
} because they are more extractive whereas phosphate buffers probably
} preserve cell contents better but at the expense of contrast.
}
} I just thought that I should add this because you mention Sorenson's
} buffer in the fixative. I'm sure that many of the other points have an
} effect but just wondered if the buffer could be contributing to your
} problems as well.
}
} Malcolm
}
}
} Malcolm Haswell
} e.m. unit
} School of Health, Natural and Social Sciences
} Fleming Building
} University of Sunderland
} Tyne & Wear
} SR1 3SD
} UK
} e-mail: malcolm.haswell-at-sunderland.ac.uk
}
}
} ----- Original Message -----
} X-from: nizets2-at-yahoo.com
} Date: Thursday, May 10, 2007 12:23 pm
} Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial
} cristae
}
} }
} }
} }
} } --------------------------------------------------------------------
} } --------
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} } AmericaTo Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserverOn-Line Help
} } http://www.microscopy.com/MicroscopyListserver/FAQ.html-------------
} } ---------------------------------------------------------------
} }
} } Dear Tom,
} }
} } I agree with the other comments: Ferrocyanide,
} } decreasing dehydration times (penetration is immediate
} } in monolayers) and using picric acid probably can
} } improve membrane contrast.
} } (personally I already tried UAc in methanol without
} } satisfaction)
} }
} } Please share your experience with us.
} }
} } Best regards,
} }
} } Stephane
} }
} } --- tbargar-at-unmc.edu wrote:
} }
} } }
} } }
} } }
} } }
} } --------------------------------------------------------------------
} } --------
} } } The Microscopy ListServer -- CoSponsor: The
} } } Microscopy Society of America
} } } To Subscribe/Unsubscribe --
} } } http://www.microscopy.com/MicroscopyListserver
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} } http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } }
} } --------------------------------------------------------------------
} } --------
} } }
} } }
} } } Dear listers,
} } }
} } } Anyone out there have any advice on how to enhance
} } } the contrast and
} } } definition of the membranes of the cristae of
} } } mitochondria? The samples
} } } brought to me are monolayer cell cultures of cancer
} } } cells grown on
} } } Thermanox coverslips. This is how I'm currently
} } } processing the samples:
} } } Primary fixation is 2% glutaraldehyde, 2%
} } } paraformaldehyde, 0.5% acrolein
} } } in 0.1M Sorensen's phosphate buffer pH 7.2.
} } } Post-fixation in 1%Osmium
} } } Tetroxide in 0.1M Sorensen's phosphate buffer.
} } } Dehydration in 50, 70, 90,
} } } 95, 100%X3 ethanol solutions. embedding in
} } } Araldite. Sections are stained
} } } with 2% uranyl acetate aqueous 15 minutes and
} } } Reynold's lead citrate 10
} } } minutes. The density of the cytoplasm and
} } } mitochondrial matrix are
} } } similar with the result that the contrast of the
} } } mitochondria is similar to
} } } the cytoplasm. The mitochondria and it's membranes
} } } (outer and that of the
} } } cristae) don't really "stand out". The researchers
} } } involved want to see
} } } really contrasty mitochondrial cristae.
} } }
} } } The next thing I'm going to try is post-fixation
} } } with a mix of osmium
} } } tetroxide and potassium ferrocyanide.
} } }
} } } Anyone have any other ideas? Thanks in advance.
} } }
} } }
} } } Tom Bargar
} } } University of Nebraska Medical Center
} } } Core Electron Microscopy Research Facility
} } } 986395 Nebraska Medical Center
} } } Omaha, NE 68198-6395
} } } 402-559-7347
} } } tbargar-at-unmc.edu
} } }
} } }
} } } ==============================Original
} } } Headers==============================
} } } 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39
} } } 2007
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} } } Mitochondrial cristae
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} } } 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu}
} } } 8, 20 -- Date: Tue, 1 May 2007 15:39:27 -0500
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} } 9, 21 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} } 9, 21 -- Subject: Re: [Microscopy] Help with enhancing contrast of
} } Mitochondrial cristae
} } 9, 21 -- To: tbargar-at-unmc.edu
} } 9, 21 -- Cc: microscopy-at-microscopy.com
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} 8, 37 -- From: Malcolm Haswell {malcolm.haswell-at-sunderland.ac.uk}
} 8, 37 -- Subject: Re: [Microscopy] Re: Help with enhancing contrast of
} Mitochondrial
} 8, 37 -- cristae
} 8, 37 -- To: Microscopy MSA {Microscopy-at-microscopy.com}
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==============================Original Headers==============================
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From: thoward-at-unm.edu
Date: Thu, 10 May 2007 11:01:42 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

More osmium questions: I still think that a lot of the
trouble with low-contrast ("missing") membranes in TC
samples is due to the containers - if you run a cell
pellet (in a polypropylene Eppendorf tube) side-by-side
with cells in their culture dishes (polystyrene), the
pelleted cells will look normal, but the dish cells will
have the "ghost" membranes....and the PP tube is still
clear, but the PS dish has browned a bit after osmication.
Using a reduced osmium (OPF) avoids the problem. I can't
figure out why the post treatments some people use reverse
this, and I've never tried those - I use OPF for TC
samples, but plan to give those a whirl in my free time
(maybe when I retire?!).

Any chemists able to hop in on the container issue?
Please?

Tamara

On Thu, 10 May 2007 10:42:01 -0500
hyi-at-emory.edu wrote:
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy
} Society of America
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} http://www.microscopy.com/MicroscopyListserver
} On-Line Help
} http://www.microscopy.com/MicroscopyListserver/FAQ.html
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}
} Dear All:
}
} I am so glad to see the discussion on the
} contrast issue
} continues as I have been hearing more and more reports
} on this problem
} and I myself have been experiencing it too. Many
} hypotheses have been
} suggested on the causes of the problem but it seems that
} there were
} always evidences supporting different
} arguments. Practically, besides
} using short dehydration and infiltration times for
} monolayer cells, I
} now HAVE TO use potassium ferrocyanide with OsO4 in
} order to get good
} contrast (even for bulk tissue). When I find out the
} contrast is low
} after sections are being cut, I re-counterstain sections
} with 15% UA in
} methanol (it can give a muddy appearance if it is over
} done). In
} addition, I make sure to use a relatively fresh OsO4
} stock solution.
}
} I have been working in EM core facilities for a
} long time and
} have dealt with all kinds of samples. One question I
} have for people
} like me is whether this is a new problem or has always
} been this way. I
} did not remember contrast was so much an issue in the
} good old days
} even with standard dehydration times and when OsO4 was
} used
} alone. Personally I think this problem has something to
} do with the
} reagents we use now days, particularly OsO4. I did
} notice the results
} were different when I use different OsO4 (liquid vs.
} crystal, old vs.
} fresh….). I am wondering what other people’s experiences
} are with OsO4
} and how OsO4 solution is prepared and stored in
} different
} laboratories. I would really love to hear from chemist
} more details on
} how OsO4 works and from vendors if there has been any
} change in their
} sources for OsO4.
}
} Thank all very much
}
} Hong
} Emory SOM EM
}
}
} On May 10, 2007, at 8:10 AM,
} malcolm.haswell-at-sunderland.ac.uk wrote:
} }
} } -----------------------------------------------------------------------
} } -----
} } The Microscopy ListServer -- CoSponsor: The Microscopy
} } Society of
} } America
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} } On-Line Help
} } http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } -----------------------------------------------------------------------
} } -----
} }
} } Dear Tom
} }
} } I just thought I should mention something that I think I
} } read years ago
} } in one of M.A. Hayat's books. He mentions that
} } cacodylate and some
} } other
} } buffers when incorporated into a fixative produce higher
} } contrast
} } because they are more extractive whereas phosphate
} } buffers probably
} } preserve cell contents better but at the expense of
} } contrast.
} }
} } I just thought that I should add this because you
} } mention Sorenson's
} } buffer in the fixative. I'm sure that many of the other
} } points have an
} } effect but just wondered if the buffer could be
} } contributing to your
} } problems as well.
} }
} } Malcolm
} }
} }
} } Malcolm Haswell
} } e.m. unit
} } School of Health, Natural and Social Sciences
} } Fleming Building
} } University of Sunderland
} } Tyne & Wear
} } SR1 3SD
} } UK
} } e-mail: malcolm.haswell-at-sunderland.ac.uk
} }
} }
} } ----- Original Message -----
} } X-from: nizets2-at-yahoo.com
} } Date: Thursday, May 10, 2007 12:23 pm
} } Subject: [Microscopy] Re: Help with enhancing contrast
} } of Mitochondrial
} } cristae
} }
} } }
} } }
} } }
} } } --------------------------------------------------------------------
} } } --------
} } } The Microscopy ListServer -- CoSponsor: The Microscopy
} } } Society of
} } } AmericaTo Subscribe/Unsubscribe --
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} } } Help
} } } http://www.microscopy.com/MicroscopyListserver/FAQ.html-------------
} } } ---------------------------------------------------------------
} } }
} } } Dear Tom,
} } }
} } } I agree with the other comments: Ferrocyanide,
} } } decreasing dehydration times (penetration is immediate
} } } in monolayers) and using picric acid probably can
} } } improve membrane contrast.
} } } (personally I already tried UAc in methanol without
} } } satisfaction)
} } }
} } } Please share your experience with us.
} } }
} } } Best regards,
} } }
} } } Stephane
} } }
} } } --- tbargar-at-unmc.edu wrote:
} } }
} } } }
} } } }
} } } }
} } } }
} } } --------------------------------------------------------------------
} } } --------
} } } } The Microscopy ListServer -- CoSponsor: The
} } } } Microscopy Society of America
} } } } To Subscribe/Unsubscribe --
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} } } } On-Line Help
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} } } http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } } }
} } } --------------------------------------------------------------------
} } } --------
} } } }
} } } }
} } } } Dear listers,
} } } }
} } } } Anyone out there have any advice on how to enhance
} } } } the contrast and
} } } } definition of the membranes of the cristae of
} } } } mitochondria? The samples
} } } } brought to me are monolayer cell cultures of cancer
} } } } cells grown on
} } } } Thermanox coverslips. This is how I'm currently
} } } } processing the samples:
} } } } Primary fixation is 2% glutaraldehyde, 2%
} } } } paraformaldehyde, 0.5% acrolein
} } } } in 0.1M Sorensen's phosphate buffer pH 7.2.
} } } } Post-fixation in 1%Osmium
} } } } Tetroxide in 0.1M Sorensen's phosphate buffer.
} } } } Dehydration in 50, 70, 90,
} } } } 95, 100%X3 ethanol solutions. embedding in
} } } } Araldite. Sections are stained
} } } } with 2% uranyl acetate aqueous 15 minutes and
} } } } Reynold's lead citrate 10
} } } } minutes. The density of the cytoplasm and
} } } } mitochondrial matrix are
} } } } similar with the result that the contrast of the
} } } } mitochondria is similar to
} } } } the cytoplasm. The mitochondria and it's membranes
} } } } (outer and that of the
} } } } cristae) don't really "stand out". The researchers
} } } } involved want to see
} } } } really contrasty mitochondrial cristae.
} } } }
} } } } The next thing I'm going to try is post-fixation
} } } } with a mix of osmium
} } } } tetroxide and potassium ferrocyanide.
} } } }
} } } } Anyone have any other ideas? Thanks in advance.
} } } }
} } } }
} } } } Tom Bargar
} } } } University of Nebraska Medical Center
} } } } Core Electron Microscopy Research Facility
} } } } 986395 Nebraska Medical Center
} } } } Omaha, NE 68198-6395
} } } } 402-559-7347
} } } } tbargar-at-unmc.edu
} } } }
} } } }
} } } } ==============================Original
} } } } Headers==============================
} } } } 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39
} } } } 2007
} } } } 8, 20 -- Received: from zixvpm02.unmc.edu
} } } } (zixvpm02.unmc.edu [192.198.54.127])
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} } } } 1 May 2007 15:39:39 -0500
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} } } } 8, 20 -- for {Microscopy-at-MSA.Microscopy.Com} ; Tue,
} } } } 1 May 2007 15:39:38 -0500 (CDT)
} } } } 8, 20 -- Subject: Help with enhancing contrast of
} } } } Mitochondrial cristae
} } } } 8, 20 -- To: Microscopy-at-MSA.Microscopy.Com
} } } } 8, 20 -- X-Mailer: Lotus Notes Release 7.0.1 January
} } } } 17, 2006
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} } } }
} } } {OFDC8670F1.FEB83264-ON862572CE.006F5771-862572CE.00717D6E-at-unmc.edu}
} } } } 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu}
} } } } 8, 20 -- Date: Tue, 1 May 2007 15:39:27 -0500
} } } } 8, 20 -- X-MIMETrack: Serialize by Router on
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} } } ==============================Original
} } } Headers==============================9, 21 -- From
} } } nizets2-at-yahoo.com Thu May 10 06:15:21 2007
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From: phillipst-at-missouri.edu
Date: Thu, 10 May 2007 11:13:36 -0500
Subject: [Microscopy] Fwd: Re: Gelatin subbed slides - eliminate the

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

This message has bounced from the listserver several times. I am trying
again but have deleted the protocol to see if this is the cause.

Yes, I am familiar with the older method requiring dry acetone. But the
half life of APTS in water pH 7 at 24 C is 8.4 hrs (see nice summary of
chemical properties and safety at
http://www.inchem.org/documents/sids/sids/919302.pdf ). So my advice is
make it fresh and skip the hassles with acetone. This means you can use
plastic slide holders to dip!

At 05:16 PM 05/09/07, you wrote:
} Hi Tom,
}
} Ok, I'm interested... I have used 3-aminopropyltriethoxysilane (same
} thing?) at 1% in acetone to treat glass so that things with aldehydes
} would stick, but the notes I had on that (method from Hans Ris) indicate
} absolutely dry acetone - that water would decompose it. So you mix it with
} water?
}
} Thanks,
}
} Dale
}
} Thomas E. Phillips wrote:
} } If you haven't tried coating your slides with 1%
} } aminopropyltriethoxysilane (APTS) in water then you are missing out on a
} } much better alternative. The water really beads up on it. I used to use
} } chrome-gel but it doesn't compare.
} }
} } At 03:30 PM 05/09/07, you wrote:
} }
} }
} } } ----------------------------------------------------------------------------
} } }
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Thomas E. Phillips, PhD
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400

573-882-4712 (office)
573-882-0123 (fax)
PhillipsT-at-missouri.edu
http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

Thomas E. Phillips, PhD
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400

573-882-4712 (office)
573-882-0123 (fax)
PhillipsT-at-missouri.edu
http://www.biology.missouri.edu/faculty/phillips.html
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==============================Original Headers==============================
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8, 19 -- From: "Thomas E. Phillips" {phillipst-at-missouri.edu}
8, 19 -- Subject: Fwd: Re: [Microscopy] Gelatin subbed slides - eliminate the
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From: hyi-at-emory.edu
Date: Thu, 10 May 2007 11:32:48 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Thank you, Tamara. I have seen what you described as well. I even
lifted cells from half of culture dish and processed it in parallel
(different resin though) in a Eppendorf tube. The contrast of cells in
tube was somewhat better than those from the dish. However, I am
puzzled by the fact that we process brain vibratome sections in dishes
routinely and have not had much problem with contrast. If the argument
for this is that the vibratome sections are not attached to the
plastic, then so is monolayer cells on glass coverslip. I have seen
low contrast in cells cultured on glass coverslip as well.

Nevertheless, I do think container could be a possible cause, and do
want to hear from explanation from a chemist.

Hong

On May 10, 2007, at 12:01 PM, Tamara A Howard wrote:

} More osmium questions: I still think that a lot of the trouble with
} low-contrast ("missing") membranes in TC samples is due to the
} containers - if you run a cell pellet (in a polypropylene Eppendorf
} tube) side-by-side with cells in their culture dishes (polystyrene),
} the pelleted cells will look normal, but the dish cells will have the
} "ghost" membranes....and the PP tube is still clear, but the PS dish
} has browned a bit after osmication. Using a reduced osmium (OPF)
} avoids the problem. I can't figure out why the post treatments some
} people use reverse this, and I've never tried those - I use OPF for TC
} samples, but plan to give those a whirl in my free time (maybe when I
} retire?!).
}
} Any chemists able to hop in on the container issue? Please?
}
} Tamara
}
} On Thu, 10 May 2007 10:42:01 -0500
} hyi-at-emory.edu wrote:
} } ----------------------------------------------------------------------
} } ------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America
} } To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------
} } ------
} } Dear All:
} } I am so glad to see the discussion on the contrast issue
} } continues as I have been hearing more and more reports on this
} } problem and I myself have been experiencing it too. Many hypotheses
} } have been suggested on the causes of the problem but it seems that
} } there were always evidences supporting different arguments.
} } Practically, besides using short dehydration and infiltration times
} } for monolayer cells, I now HAVE TO use potassium ferrocyanide with
} } OsO4 in order to get good contrast (even for bulk tissue). When I
} } find out the contrast is low after sections are being cut, I
} } re-counterstain sections with 15% UA in methanol (it can give a
} } muddy appearance if it is over done). In addition, I make sure to
} } use a relatively fresh OsO4 stock solution. I have been
} } working in EM core facilities for a long time and have dealt with
} } all kinds of samples. One question I have for people like me is
} } whether this is a new problem or has always been this way. I did not
} } remember contrast was so much an issue in the good old days even
} } with standard dehydration times and when OsO4 was used alone.
} } Personally I think this problem has something to do with the
} } reagents we use now days, particularly OsO4. I did notice the results
} } were different when I use different OsO4 (liquid vs. crystal, old
} } vs. fresh….). I am wondering what other people’s experiences are
} } with OsO4 and how OsO4 solution is prepared and stored in different
} } laboratories. I would really love to hear from chemist more details
} } on how OsO4 works and from vendors if there has been any change in
} } their sources for OsO4.
} } Thank all very much
} } Hong
} } Emory SOM EM
} } On May 10, 2007, at 8:10 AM, malcolm.haswell-at-sunderland.ac.uk wrote:
} } }
} } } ---------------------------------------------------------------------
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} } }
} } } Dear Tom
} } }
} } } I just thought I should mention something that I think I read years
} } } ago
} } } in one of M.A. Hayat's books. He mentions that cacodylate and some
} } } other
} } } buffers when incorporated into a fixative produce higher contrast
} } } because they are more extractive whereas phosphate buffers probably
} } } preserve cell contents better but at the expense of contrast.
} } }
} } } I just thought that I should add this because you mention Sorenson's
} } } buffer in the fixative. I'm sure that many of the other points have
} } } an
} } } effect but just wondered if the buffer could be contributing to your
} } } problems as well.
} } }
} } } Malcolm
} } }
} } }
} } } Malcolm Haswell
} } } e.m. unit
} } } School of Health, Natural and Social Sciences
} } } Fleming Building
} } } University of Sunderland
} } } Tyne & Wear
} } } SR1 3SD
} } } UK
} } } e-mail: malcolm.haswell-at-sunderland.ac.uk
} } }
} } }
} } } ----- Original Message -----
} } } X-from: nizets2-at-yahoo.com
} } } Date: Thursday, May 10, 2007 12:23 pm
} } } Subject: [Microscopy] Re: Help with enhancing contrast of
} } } Mitochondrial
} } } cristae
} } }
} } } }
} } } }
} } } }
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} } } } Dear Tom,
} } } }
} } } } I agree with the other comments: Ferrocyanide,
} } } } decreasing dehydration times (penetration is immediate
} } } } in monolayers) and using picric acid probably can
} } } } improve membrane contrast.
} } } } (personally I already tried UAc in methanol without
} } } } satisfaction)
} } } }
} } } } Please share your experience with us.
} } } }
} } } } Best regards,
} } } }
} } } } Stephane
} } } }
} } } } --- tbargar-at-unmc.edu wrote:
} } } }
} } } } }
} } } } }
} } } } }
} } } } }
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} } } } }
} } } } }
} } } } } Dear listers,
} } } } }
} } } } } Anyone out there have any advice on how to enhance
} } } } } the contrast and
} } } } } definition of the membranes of the cristae of
} } } } } mitochondria? The samples
} } } } } brought to me are monolayer cell cultures of cancer
} } } } } cells grown on
} } } } } Thermanox coverslips. This is how I'm currently
} } } } } processing the samples:
} } } } } Primary fixation is 2% glutaraldehyde, 2%
} } } } } paraformaldehyde, 0.5% acrolein
} } } } } in 0.1M Sorensen's phosphate buffer pH 7.2.
} } } } } Post-fixation in 1%Osmium
} } } } } Tetroxide in 0.1M Sorensen's phosphate buffer.
} } } } } Dehydration in 50, 70, 90,
} } } } } 95, 100%X3 ethanol solutions. embedding in
} } } } } Araldite. Sections are stained
} } } } } with 2% uranyl acetate aqueous 15 minutes and
} } } } } Reynold's lead citrate 10
} } } } } minutes. The density of the cytoplasm and
} } } } } mitochondrial matrix are
} } } } } similar with the result that the contrast of the
} } } } } mitochondria is similar to
} } } } } the cytoplasm. The mitochondria and it's membranes
} } } } } (outer and that of the
} } } } } cristae) don't really "stand out". The researchers
} } } } } involved want to see
} } } } } really contrasty mitochondrial cristae.
} } } } }
} } } } } The next thing I'm going to try is post-fixation
} } } } } with a mix of osmium
} } } } } tetroxide and potassium ferrocyanide.
} } } } }
} } } } } Anyone have any other ideas? Thanks in advance.
} } } } }
} } } } }
} } } } } Tom Bargar
} } } } } University of Nebraska Medical Center
} } } } } Core Electron Microscopy Research Facility
} } } } } 986395 Nebraska Medical Center
} } } } } Omaha, NE 68198-6395
} } } } } 402-559-7347
} } } } } tbargar-at-unmc.edu
} } } } }
} } } } }
} } } } } ==============================Original
} } } } } Headers==============================
} } } } } 8, 20 -- From tbargar-at-unmc.edu Tue May 1 15:39:39
} } } } } 2007
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} } } } } 8, 20 -- Subject: Help with enhancing contrast of
} } } } } Mitochondrial cristae
} } } } } 8, 20 -- To: Microscopy-at-MSA.Microscopy.Com
} } } } } 8, 20 -- X-Mailer: Lotus Notes Release 7.0.1 January
} } } } } 17, 2006
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} } } } } 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu}
} } } } } 8, 20 -- Date: Tue, 1 May 2007 15:39:27 -0500
} } } } } 8, 20 -- X-MIMETrack: Serialize by Router on
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} } } } 9, 21 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} } } } 9, 21 -- Subject: Re: [Microscopy] Help with enhancing contrast of
} } } } Mitochondrial cristae
} } } } 9, 21 -- To: tbargar-at-unmc.edu
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} } }
} } } ==============================Original
} } } Headers==============================
} } } 8, 37 -- From malcolm.haswell-at-sunderland.ac.uk Thu May 10 07:06:52
} } } 2007
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} } } 8, 37 -- Date: Thu, 10 May 2007 13:06:36 +0100
} } } 8, 37 -- From: Malcolm Haswell {malcolm.haswell-at-sunderland.ac.uk}
} } } 8, 37 -- Subject: Re: [Microscopy] Re: Help with enhancing contrast
} } } of Mitochondrial
} } } 8, 37 -- cristae
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} } ==============================Original
} } Headers==============================
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} } 9, 27 -- Subject: [Microscopy] Help with enhancing contrast of
} } Mitochondrial
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}
} ***************************
} Tamara Howard
} Cell Biology & Physiology
} UNM-HSC
} Albuquerque, NM
} ***************************



==============================Original Headers==============================
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From: scr-at-phys.uni-sofia.bg
Date: Thu, 10 May 2007 17:31:15 -0500
Subject: [Microscopy] viaWWW: SEM Hitachi S570 schematics documentation

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Email: scr-at-phys.uni-sofia.bg
Name: Stoyan Russev

Organization: Sofia University, Faculty of Physics

Title-Subject: [Filtered] SEM Hitachi S570 schematics documentation

Question: I am using Hitachi S570 SEM in the lab and I need the full documentation (schematics especially) for upgrading and repair.

Can anyone help me to find it?

Thank you in advance.
S. Russev

---------------------------------------------------------------------------

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From: heckman-at-bgnet.bgsu.edu
Date: Thu, 10 May 2007 20:53:27 -0500
Subject: [Microscopy] Re: Help with enhancing contrast of Mitochondrial

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Dr. Hong-
I only saw the problem of the membrane "disappearing" in 1978,
when I started preparing layers of epithelial cells. I found a way to
work around it and haven't (fortunately) rediscovered the problem
since then.
Carol Heckman
Department of Biological Sciences
Bowling Green State University



==============================Original Headers==============================
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From: nealzimm-at-cpinternet.com
Date: Fri, 11 May 2007 07:47:04 -0500
Subject: [Microscopy] Re: Looking for recommendations about dual beam ion

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Richard, Witold;

I'll try to reply to both your questions with this one post.

Because I use and maintain PIPS does not make me an "expert"
at ion milling. I have noticed a few things that do/do not
happen under the different milling conditions we work with.

Low kv milling seems to give lower instances of twinning and
stacking faults in the thin films I have worked with. But I
do not think that milling damage goes away at lower kVs. I
see a background of "mottling" in the films I have imaged.
This is what I assume is the damage from milling and occurs at
5kV milling as well as 2.6kV milling.

The milling times are so long for the reason that most of the
thin-films-on-glass (silicon) are created to study the behavior
as deposited - no anneal. This makes the films fragile to the
point that even the technique of encasing the sample in a brass
tube does not work - the glue bond bonding the coupons together
is greater than the bonding of the film to the substrate.

For mechanically prepared films (which our lead microscopist
prefers over FIB cut samples) I had to come up with a way
of bonding a glass cover slip to the substrate and trapping
the film between. As has been mentioned here, glass behaves
poorly in an ion beam. As it becomes very thin, it melts and
(usually) pulls the epoxy and film off of the substrate.

The only way to stop the melting is to go to low kVs and a
low duty cycle. For most samples this is 3.5kV and 1 minute
of milling for every 5 minutes. And as I am only around for
8 hours a day, this makes for a 3 day milling session.

I have been able to prepare sections of Pb (actually PbZrTiOx)
that give spectacular lattice (although disappointingly similar
to silicon). As far as reducing the time for milling silicon
samples, at ten microns thickness and 3kV milling on
non-temperature sensitive samples, milling times are usually in
the 15 to 20 minute range.

Neal

On Thu, 2007-05-10 at 09:10 -0500, richard.beanland-at-bookham.com wrote:
}
}
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} Neal,
} I wouldn't expect a 10um Si sample to take more than 30-60 mins
} to thin in a PIPS.
} I can only think you are working at a relatively high milling angle
} (} 5 degrees) and so have to turn the beam energy down very low so you
} don't damage your sample. Why not experiment with one sample using full
} power (6kV) and 3 degrees incidence angle, finishing off with 10 mins at
} 2kV? It could increase your throughput more than 20 times!
}
} Richard
}
}
}
} ________________________________________
} Richard Beanland
} Materials Analysis
} Bookham
} Caswell
} Towcester
} Northants
} NN12 8EQ
} United Kingdom
} Tel. +44 1327 356362
} Fax. +44 1327 356775
} http://www.bookham.com
} ________________________________________
}
}
} -----Original Message-----
} X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
} Sent: 10 May 2007 14:10
} To: Richard Beanland
} Subject: [Microscopy] Re: Looking for recommendations about dual beam
} ion
}
}
}
}
} ------------------------------------------------------------------------
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} John
}
} I work with four Gatan PIPS daily and have found them to be quite
} acceptable in ease of both maintenance and use.
}
} We currently do maintenance only when the ion guns stop functioning,
} at that point the machine is taken out of service and a full cleaning
} performed. This happens about every five to six months and leaves
} the PIPS out of service for two days (One day to clean, pump down
} over night, and alignment the next day). The diaphragm pump needs
} new diaphragms once a year (about an hours work) but not too much else.
}
} Work load is typically one to two ceramic samples per day, with an
} occasional silicon or glass sample thrown in. The ceramic samples
} are lapped to about six microns and mill for an hour to two hours.
} Silicon and glass are left about ten microns thick and take much
} longer (up to three days), not so much due to the thickness as to
} the low kVs used and letting the sample "cool".
}
} Gatan offers a digital camera/zoom lens set and an external diaphragm
} pump as extras. In my opinion, they are a "must have".
}
} If you have any further questions please feel free to contact me.
}
} Neal R. Zimmermann
}
}
}
} On Wed, 2007-05-09 at 15:43 -0500, jpchandl-at-mines.edu wrote:
} }
} }
} }
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} }
} } I need to tap in to the wisdom of the list about dual beam argon ion
} mills.
} }
} } We have a Bal-Tec RES100 ion mill that we are having difficulty
} maintaining.
} } It had been in disuse for a couple of years, and has been repaired and
} } upgraded recently, at the factory, to be as close to a RES101 as
} possible.
} } We are having issues such as the guns becoming contaminated and
} needing
} } service much more frequently than we think they should. Routine
} maintenance
} } seems to be difficult, too. We think we are looking at a high
} maintenance
} } instrument here. I would like to know what kind of experience other
} labs
} } have had with this instrument.
} }
} } I would also like to hear from people who have experience
} using/maintaining
} } the RES100/101 or the Gatan PIPS, or both. We need some comparative
} } information, so we can make a decision about how to proceed with our
} ion
} } milling needs.
} }
} } Any input would be appreciated.
} }
} } --John
} }
} } John Chandler
} } Manager, EM Lab
} } Colorado School of Mines
} } Golden, CO 80401
} } jpchandl-at-mines.edu
} } 303-384-2203
} }
} }
} }
} }
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From: john.mardinly-at-intel.com
Date: Fri, 11 May 2007 11:52:13 -0500
Subject: [Microscopy] Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

~ 8 mm apart? Did I read this correctly? Not microns? We use a Disco saw
routinely. If it's ~ 8 mm apart, you can use any diamond saw. If it's
microns you need a FIB.

John Mardinly
Intel Corporation
This email does not represent an opinion of Intel Corporation.

-----Original Message-----
X-from: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
Sent: Tuesday, May 08, 2007 10:33 AM
To: Mardinly, John

Microscopy Folks,

I'd like to get some input on sample preparation with respect to
silicon.

PROBLEM:

I have an FEI XL-50 FESEM that was originally designed to accept flat
samples, essentially silicon wafers. It does not have what one would
consider a typical exchange port. The exchange port is robotically
controlled, and the maximum sample height, including stub, must be 5 mm
or less. I must view these samples on edge, i.e. 90 degrees. This
presents a problem in that I must cleave two parallel sections very
close to each other. Perhaps diamond cutters can do this, but my hands
are too shaky.

QUESTION:

Is there a device, or method, that would allow me to make these cleaves
~ 8 mm apart with any reasonable control? I found stubs that are low
profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
still have this issue of doing two parallel cleaves very close together.

Just for info. purposes I am in the silicon MEMS development arena.

If you feel your reply is of general interest to this community, please
reply to all, or you may contact me directly.

Regards to all in this small world,

Peter Tomic
Renaissance Wireless Corp.


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From: LarryZ-at-fai.us
Date: Fri, 11 May 2007 12:15:37 -0500
Subject: [Microscopy] Mammalian Identified by Hair Imaging

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Can anyone recommend references on identifying mammalian species by
analysis of their hair?


Thanks,
Larry Zagorski

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From: lcgould-at-med.cornell.edu
Date: Fri, 11 May 2007 12:21:11 -0500
Subject: [Microscopy] Re: Mammalian Identified by Hair Imaging

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Larry-
That's the kind of reference that police forensics departments should
have. Try contacting libraries at places like John Jay College of
Criminal Justice here in NYC (its part of the City University of NY
system), or simialr schools near you.
Lee


}
} Can anyone recommend references on identifying mammalian species by
} analysis of their hair?
}
}
} Thanks,
} Larry Zagorski

--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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From: dsherman-at-purdue.edu
Date: Fri, 11 May 2007 12:31:34 -0500
Subject: [Microscopy] Charcoal markers

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Folks,

I have an inquiry for a source for charcoal marker beads of about 30µm in
size. These are to be used to mark plant roots to determine growth rate.
We have a paper that used this technique but they did not give a source for
the charcoal beads.

I have not been able to reach the authors so hoped one of you would have a
source.

Debby


Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.agriculture.purdue.edu/microscopy



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From: walck-at-southbaytech.com
Date: Fri, 11 May 2007 12:42:36 -0500
Subject: [Microscopy] Re: Looking for recommendations about dual beam ion

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I'd like to contribute a little to the ion milling damage issue being
discussed. I think that I have used every brand of ion mill on the market
in my career at some point or another. When I was with PPG, glass
substrates were the only cross section samples that I dealt with and they
were the most tricky. With those samples, I had the most trouble with the
PIPS(TM) instrument. If you look at the milling rates with other
instruments, the PIPS(TM) Penning ion guns are similar in ion sputtering
rates to other ion guns in other companies' ion mills (except the
Technoorg-Linda gun with the Einzel focusing lens). It is actually very
hard to compare them because everyone does it differently. I believe that
the issue wasn't one of the ion beam, but so for the design of the
clamp-type DuoPost(TM) that I was using to mill at low angles. The
DuoPost(TM) does not have a large mass, and the gripping method does not
have a great thermal path to extract heat away from the sample. Glass also
doesn't have a great thermal conductivity. If I did not operate the PIPS
with lowered ion gun performance parameters, I would soften my glass samples
and they would soften and "droop down" during milling. This did not occur
with any other ion mill that I used that had a more massive sample holder
with better heat path. These other mills included the ones from BalTec (used
extensively), EA Fischione (used extensively), and Technoorg-Linda (only a
few samples).

With respect to ion damage from ion milling, I think that Arpad Barna's
group's work is definitive on the topic. He ion milled Si and GaAs samples
at different energies at 5 degrees and then immediately evaporated Al on top
to protect the samples from oxidation when they were taken out of the ion
mill. He then prepared cross sections of those samples and measured the
thickness of the amorphous damage region on the surface due to the ion
milling by using HREM. For Si, they saw that at 3 kV (about the limit for
conventional ion guns) an amorphous region 120 Angstroms was formed. Using
the Technoorg-Linda low energy gun at 250 eV, they achieved an amorphous
region of about 10 Angstroms. (A. Barna, B. Pecz; Amorphisation and surface
morphology development at low-energy ion milling; Ultramicroscopy 70, 1998.)
In other studies and examples, they have also shown that the mottling from
ion milled and FIB samples using the low energy guns can be completely
eliminated. I have some of the data from that work if you are interested in
seeing the results.


Disclaimer: South Bay Technology, Inc. is the exclusive distributer for
Technoorg-Linda products in the United States.

-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
Sent: Friday, May 11, 2007 5:53 AM
To: Walck-at-SouthBayTech.com

Richard, Witold;

I'll try to reply to both your questions with this one post.

Because I use and maintain PIPS does not make me an "expert"
at ion milling. I have noticed a few things that do/do not happen under the
different milling conditions we work with.

Low kv milling seems to give lower instances of twinning and stacking faults
in the thin films I have worked with. But I do not think that milling
damage goes away at lower kVs. I see a background of "mottling" in the
films I have imaged.
This is what I assume is the damage from milling and occurs at 5kV milling
as well as 2.6kV milling.

The milling times are so long for the reason that most of the
thin-films-on-glass (silicon) are created to study the behavior as deposited
- no anneal. This makes the films fragile to the point that even the
technique of encasing the sample in a brass tube does not work - the glue
bond bonding the coupons together is greater than the bonding of the film to
the substrate.

For mechanically prepared films (which our lead microscopist prefers over
FIB cut samples) I had to come up with a way of bonding a glass cover slip
to the substrate and trapping the film between. As has been mentioned here,
glass behaves poorly in an ion beam. As it becomes very thin, it melts and
(usually) pulls the epoxy and film off of the substrate.

The only way to stop the melting is to go to low kVs and a low duty cycle.
For most samples this is 3.5kV and 1 minute of milling for every 5 minutes.
And as I am only around for
8 hours a day, this makes for a 3 day milling session.

I have been able to prepare sections of Pb (actually PbZrTiOx) that give
spectacular lattice (although disappointingly similar to silicon). As far
as reducing the time for milling silicon samples, at ten microns thickness
and 3kV milling on non-temperature sensitive samples, milling times are
usually in the 15 to 20 minute range.

Neal

On Thu, 2007-05-10 at 09:10 -0500, richard.beanland-at-bookham.com wrote:
}
}
} ----------------------------------------------------------------------
} ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society
} of America To Subscribe/Unsubscribe --
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} ----------------------------------------------------------------------
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}
} Neal,
} I wouldn't expect a 10um Si sample to take more than 30-60 mins to
} thin in a PIPS.
} I can only think you are working at a relatively high milling angle
} (} 5 degrees) and so have to turn the beam energy down very low so you
} don't damage your sample. Why not experiment with one sample using
} full power (6kV) and 3 degrees incidence angle, finishing off with 10
} mins at 2kV? It could increase your throughput more than 20 times!
}
} Richard
}
}
}
} ________________________________________
} Richard Beanland
} Materials Analysis
} Bookham
} Caswell
} Towcester
} Northants
} NN12 8EQ
} United Kingdom
} Tel. +44 1327 356362
} Fax. +44 1327 356775
} http://www.bookham.com
} ________________________________________
}
}
} -----Original Message-----
} X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
} Sent: 10 May 2007 14:10
} To: Richard Beanland
} Subject: [Microscopy] Re: Looking for recommendations about dual beam
} ion
}
}
}
}
} ----------------------------------------------------------------------
} --
} ----
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} America To Subscribe/Unsubscribe --
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}
} John
}
} I work with four Gatan PIPS daily and have found them to be quite
} acceptable in ease of both maintenance and use.
}
} We currently do maintenance only when the ion guns stop functioning,
} at that point the machine is taken out of service and a full cleaning
} performed. This happens about every five to six months and leaves the
} PIPS out of service for two days (One day to clean, pump down over
} night, and alignment the next day). The diaphragm pump needs new
} diaphragms once a year (about an hours work) but not too much else.
}
} Work load is typically one to two ceramic samples per day, with an
} occasional silicon or glass sample thrown in. The ceramic samples are
} lapped to about six microns and mill for an hour to two hours.
} Silicon and glass are left about ten microns thick and take much
} longer (up to three days), not so much due to the thickness as to the
} low kVs used and letting the sample "cool".
}
} Gatan offers a digital camera/zoom lens set and an external diaphragm
} pump as extras. In my opinion, they are a "must have".
}
} If you have any further questions please feel free to contact me.
}
} Neal R. Zimmermann
}
}
}
} On Wed, 2007-05-09 at 15:43 -0500, jpchandl-at-mines.edu wrote:
} }
} }
} }
} ----------------------------------------------------------------------
} --
} ----
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} America
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} }
} } I need to tap in to the wisdom of the list about dual beam argon ion
} mills.
} }
} } We have a Bal-Tec RES100 ion mill that we are having difficulty
} maintaining.
} } It had been in disuse for a couple of years, and has been repaired
} } and upgraded recently, at the factory, to be as close to a RES101 as
} possible.
} } We are having issues such as the guns becoming contaminated and
} needing
} } service much more frequently than we think they should. Routine
} maintenance
} } seems to be difficult, too. We think we are looking at a high
} maintenance
} } instrument here. I would like to know what kind of experience other
} labs
} } have had with this instrument.
} }
} } I would also like to hear from people who have experience
} using/maintaining
} } the RES100/101 or the Gatan PIPS, or both. We need some comparative
} } information, so we can make a decision about how to proceed with our
} ion
} } milling needs.
} }
} } Any input would be appreciated.
} }
} } --John
} }
} } John Chandler
} } Manager, EM Lab
} } Colorado School of Mines
} } Golden, CO 80401
} } jpchandl-at-mines.edu
} } 303-384-2203
} }
} }
} }
} }
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From: rbeavers-at-mail.smu.edu
Date: Fri, 11 May 2007 12:46:54 -0500
Subject: [Microscopy] O-ring help

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Group,

Has anyone had any experience making a custom Viton O-ring by splicing a
larger one?

Have a custom chamber operating at 10e-6 torr range and no standard
O-ring seems to fit properly.

If you have experience with type of splice (butt or diagonal) or glue
used would love to hear about it.

Thanks

Roy Beavers

Southern Methodist University
Department of Geological Sciences
P.O. Box 750395
Dallas, TX 75275
Voice: 214-768-2756
Fax: 214-768-2701
Email: rbeavers-at-smu.edu


==============================Original Headers==============================
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From: dljones-at-bestweb.net
Date: Fri, 11 May 2007 13:13:55 -0500
Subject: [Microscopy] Re: O-ring help

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Roy,
did you try using Indium wire to make a seal?
Philips used it to make UHV seals at EM 4x0 TEMs.
On the other hand the groove should not be too large.
You may get Indium wire up to 1mm diameter (or even more).

Best, Stefan Diller



----- Original Message -----
X-from: {rbeavers-at-mail.smu.edu}
To: {stefan.diller-at-t-online.de}
Sent: Friday, May 11, 2007 7:51 PM

Roy,

In one of my past lives I used to work with a lot of older vacuum
equipment that was difficult to get off the shelf O-rings for. I would
make my own O-rings with frequency. I used to get viton O-ring material of
varying diameters on a spool (like rope).

I guess you could try a larger diameter pre-made O-ring and cut it down, I
can't speak about that specifically.

As far as cutting and gluing. I did a diagonal cut using a very sharp
razor blade, cutting both ends at the same time. I would glue it with,
straight from the local hardware store, Crazy Glue. Then take fine
sandpaper - I think I used 600 grit - (but I honestly don't remember that
exact detail) and sanded the joint so it was smooth.

I tried butt joints, but never seemed to get them to work, so ended up
always doing a diagonal cut. I'm sure others have had success that way,
but I didn't.

Good luck.

dj

On Fri, 11 May 2007, rbeavers-at-mail.smu.edu wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------------
}
} Group,
}
} Has anyone had any experience making a custom Viton O-ring by splicing a
} larger one?
}
} Have a custom chamber operating at 10e-6 torr range and no standard
} O-ring seems to fit properly.
}
} If you have experience with type of splice (butt or diagonal) or glue
} used would love to hear about it.
}
} Thanks
}
} Roy Beavers
}
} Southern Methodist University
} Department of Geological Sciences
} P.O. Box 750395
} Dallas, TX 75275
} Voice: 214-768-2756
} Fax: 214-768-2701
} Email: rbeavers-at-smu.edu
}
}
} ==============================Original Headers==============================
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From: rjharris-at-uwo.ca
Date: Fri, 11 May 2007 13:42:38 -0500
Subject: [Microscopy] Mammalian Identified by Hair Imaging

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Larry
You should check ou the FBI website Forensic Science Communications Volume 6
(2004) Microscopy of Hair Part 1: A Practical Guide and Manual for Human
Hairs and Microscopy of Hair Part II: A Practical Guide and Manual for
Animal Hairs by
Douglas W. Deedrick and Sandra L. Koch

http://www.fbi.gov/hq/lab/fsc/backissu/july2004/index.htm

We use these as a principle resource in our Imaging and Analysis Course

Rick,

Richard Harris
Manager -
Imaging, Information and Data Systems
Biotron Experimental Climate Change Research Facility
University of Western Ontario,
London Ontario, CANADA.
N6A 5B7
Ph. 519-661-2111 ext. 86780
Fax 519-661-3935
e-mail rjharris-at-uwo.ca
web: www.biotron.uwo.ca


-----Original Message-----
X-from: LarryZ-at-fai.us [mailto:LarryZ-at-fai.us]
Sent: Friday, May 11, 2007 1:20 PM
To: rjharris-at-uwo.ca

Can anyone recommend references on identifying mammalian species by
analysis of their hair?


Thanks,
Larry Zagorski

==============================Original Headers==============================
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==============================Original Headers==============================
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From: jquinn-at-www.matscieng.sunysb.edu
Date: Fri, 11 May 2007 15:51:54 -0500
Subject: [Microscopy] re: O-ring help

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Folks

MarcoRubber has a full selection of o-rings.
They are online and prompt. This is their URL:
http://www.marcorubber.com/
For most rings, I find the cost for 10 to be
the same as the cost for one.

There are good charts for ISO, DIN, Italian,
British, Japanese, French, Swedish, USA, and
standard-metric at this URL:
http://mdmetric.com/or/gb01.htm
Maryland Metric

regards,

Jim

PS: No commercial connection, just a HAPPY enduser.

PPS: Can't wait for the slew of "O-o-O" messages.



} From mail-at-ns.microscopy.com Fri May 11 13:47:10 2007
} Date: Fri, 11 May 2007 12:47:38 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: rbeavers-at-mail.smu.edu
} Reply-to: rbeavers-at-mail.smu.edu
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
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}
}
}
}
} ----------------------------------------------------------------------------
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} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Group,
}
} Has anyone had any experience making a custom Viton O-ring by splicing a
} larger one?
}
} Have a custom chamber operating at 10e-6 torr range and no standard
} O-ring seems to fit properly.
}
} If you have experience with type of splice (butt or diagonal) or glue
} used would love to hear about it.
}
} Thanks
}
} Roy Beavers
}
} Southern Methodist University
} Department of Geological Sciences
} P.O. Box 750395
} Dallas, TX 75275
} Voice: 214-768-2756
} Fax: 214-768-2701
} Email: rbeavers-at-smu.edu
}
}
} ==============================Original Headers==============================
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From: yb4-at-mail.gatech.edu
Date: Fri, 11 May 2007 22:20:37 -0500
Subject: [Microscopy] focusing screen on JEOL 100CX II

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Dear Listers:
We have a malfunctioning focusing screen on a 100CX II TEM and we have been
unable to find the cause of the problem, in spite of phone help from our local
JEOL service office and on-site help from an independent service company.
Apparently, the motor is receiving about 14 V when it should be receiving 24
V. The screen doesn't lift now, because it has fallen off its track; the motor
was constantly running while it was receiving the incorrect voltage.
Has anyone seen this problem before, or has a suggestion on how it might be
corrected?

Yolande Berta
Georgia Tech
Center for Nanostructure Characterization and Fabricaton
Atlanta, GA 30332-0245

--


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From: rcsaic-at-sbcglobal.net
Date: Mon, 14 May 2007 08:29:19 -0500
Subject: [Microscopy] SEM-help finding smooth, low-Z substrates

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Listers,

We could use some suggestions/collective experience about solid substrates
for doing SEM imaging of dispersed small particles.

What we are imaging: silicate and oxide mineral grains and silicate glass
particles down to the 0.1 micron size range (maybe smaller). Objective is to
disperse the particle on a substrate and do total particle counting and
automated measuring without "losing" the smallest particles from the data
set.

What we need: A substrate for dispersing the particles that would fit two
requirements: 1) have a pretty low average Z (carbon would be ideal but I'm
willing to consider other ideas), so that my particles will stand out
strongly in BSE, and then 2) be as smooth as possible on the sub-micron
scale. This last requirement is the tricky one because although there are
lots of carbon substrates available commercially, so far we haven't found
any that don't have sub-micron scratches and other roughness such that
particles in the 0.1 micron size range don't get lost in the surface
topography.

Well, that's about all I can think of. Thanks to all of you in advance.


Roy Christoffersen
FE-STEM Facility Manager
ARES Directorate
NASA Johnson Space Center, Houston, TX
rcsaic-at-sbcglobal.net







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From: richard.beanland-at-bookham.com
Date: Mon, 14 May 2007 09:07:35 -0500
Subject: [Microscopy] Re: Looking for recommendations about dual beam

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Neal, Scott,
I think I may have missed a couple of posts on this thread, but
I'd like to clarify my comments...
There were two things - first, that three days to get a sample by ion
milling is pretty unusual, and doesn't reflect my experience with the
PIPS. Second, the angle of incidence is another parameter in addition
to beam energy which you can vary and might give a better or more
efficient way of making samples.

If you assume that the heating of the sample is due to the kinetic
energy of the ion beam, and that the most important component of the
beam's velocity is that which is perpendicular to the sample surface,
then heating of the sample should roughly increase with the square of
the angle of incidence (using E=0.5mv^2 and sin(theta)=theta for small
angles). So you can reduce the heating of the sample about 25 times by
dropping from 5 degrees incidence angle to 1 degree - which is much more
efficient than reducing the beam energy by 25 times. The milling rate
doesn't drop off as quickly as the reduction in heating. At least this
is my way of explaining why I melt my InP specimens at 6 degrees
incidence angle but get better results than my old ion mill (which had
LN2 cooling and minimum 10 degrees incidence angle) using 2 degrees in
the PIPS.

Having said all this, I don't work with glass samples and if you have a
way of making good samples which is sucessful, that's a good thing
(finding a protocol which works can be hard enough!).

Cheers

Richard


________________________________________
Richard Beanland
Materials Analysis
Bookham
Caswell
Towcester
Northants
NN12 8EQ
United Kingdom
Tel. +44 1327 356362
Fax. +44 1327 356775
http://www.bookham.com
________________________________________


-----Original Message-----
X-from: walck-at-southbaytech.com [mailto:walck-at-southbaytech.com]
Sent: 11 May 2007 18:44
To: Richard Beanland

I'd like to contribute a little to the ion milling damage issue being
discussed. I think that I have used every brand of ion mill on the
market
in my career at some point or another. When I was with PPG, glass
substrates were the only cross section samples that I dealt with and
they
were the most tricky. With those samples, I had the most trouble with
the
PIPS(TM) instrument. If you look at the milling rates with other
instruments, the PIPS(TM) Penning ion guns are similar in ion sputtering
rates to other ion guns in other companies' ion mills (except the
Technoorg-Linda gun with the Einzel focusing lens). It is actually very
hard to compare them because everyone does it differently. I believe
that
the issue wasn't one of the ion beam, but so for the design of the
clamp-type DuoPost(TM) that I was using to mill at low angles. The
DuoPost(TM) does not have a large mass, and the gripping method does not
have a great thermal path to extract heat away from the sample. Glass
also
doesn't have a great thermal conductivity. If I did not operate the
PIPS
with lowered ion gun performance parameters, I would soften my glass
samples
and they would soften and "droop down" during milling. This did not
occur
with any other ion mill that I used that had a more massive sample
holder
with better heat path. These other mills included the ones from BalTec
(used
extensively), EA Fischione (used extensively), and Technoorg-Linda (only
a
few samples).

With respect to ion damage from ion milling, I think that Arpad Barna's
group's work is definitive on the topic. He ion milled Si and GaAs
samples
at different energies at 5 degrees and then immediately evaporated Al on
top
to protect the samples from oxidation when they were taken out of the
ion
mill. He then prepared cross sections of those samples and measured the
thickness of the amorphous damage region on the surface due to the ion
milling by using HREM. For Si, they saw that at 3 kV (about the limit
for
conventional ion guns) an amorphous region 120 Angstroms was formed.
Using
the Technoorg-Linda low energy gun at 250 eV, they achieved an amorphous
region of about 10 Angstroms. (A. Barna, B. Pecz; Amorphisation and
surface
morphology development at low-energy ion milling; Ultramicroscopy 70,
1998.)
In other studies and examples, they have also shown that the mottling
from
ion milled and FIB samples using the low energy guns can be completely
eliminated. I have some of the data from that work if you are
interested in
seeing the results.


Disclaimer: South Bay Technology, Inc. is the exclusive distributer for
Technoorg-Linda products in the United States.

-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
Sent: Friday, May 11, 2007 5:53 AM
To: Walck-at-SouthBayTech.com

Richard, Witold;

I'll try to reply to both your questions with this one post.

Because I use and maintain PIPS does not make me an "expert"
at ion milling. I have noticed a few things that do/do not happen under
the
different milling conditions we work with.

Low kv milling seems to give lower instances of twinning and stacking
faults
in the thin films I have worked with. But I do not think that milling
damage goes away at lower kVs. I see a background of "mottling" in the
films I have imaged.
This is what I assume is the damage from milling and occurs at 5kV
milling
as well as 2.6kV milling.

The milling times are so long for the reason that most of the
thin-films-on-glass (silicon) are created to study the behavior as
deposited
- no anneal. This makes the films fragile to the point that even the
technique of encasing the sample in a brass tube does not work - the
glue
bond bonding the coupons together is greater than the bonding of the
film to
the substrate.

For mechanically prepared films (which our lead microscopist prefers
over
FIB cut samples) I had to come up with a way of bonding a glass cover
slip
to the substrate and trapping the film between. As has been mentioned
here,
glass behaves poorly in an ion beam. As it becomes very thin, it melts
and
(usually) pulls the epoxy and film off of the substrate.

The only way to stop the melting is to go to low kVs and a low duty
cycle.
For most samples this is 3.5kV and 1 minute of milling for every 5
minutes.
And as I am only around for
8 hours a day, this makes for a 3 day milling session.

I have been able to prepare sections of Pb (actually PbZrTiOx) that give
spectacular lattice (although disappointingly similar to silicon). As
far
as reducing the time for milling silicon samples, at ten microns
thickness
and 3kV milling on non-temperature sensitive samples, milling times are
usually in the 15 to 20 minute range.

Neal

On Thu, 2007-05-10 at 09:10 -0500, richard.beanland-at-bookham.com wrote:
}
}
} ----------------------------------------------------------------------
} ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society

} of America To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------
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}
} Neal,
} I wouldn't expect a 10um Si sample to take more than 30-60 mins
to
} thin in a PIPS.
} I can only think you are working at a relatively high milling angle
} (} 5 degrees) and so have to turn the beam energy down very low so you
} don't damage your sample. Why not experiment with one sample using
} full power (6kV) and 3 degrees incidence angle, finishing off with 10
} mins at 2kV? It could increase your throughput more than 20 times!
}
} Richard
}
}
}
} ________________________________________
} Richard Beanland
} Materials Analysis
} Bookham
} Caswell
} Towcester
} Northants
} NN12 8EQ
} United Kingdom
} Tel. +44 1327 356362
} Fax. +44 1327 356775
} http://www.bookham.com
} ________________________________________
}
}
} -----Original Message-----
} X-from: nealzimm-at-cpinternet.com [mailto:nealzimm-at-cpinternet.com]
} Sent: 10 May 2007 14:10
} To: Richard Beanland
} Subject: [Microscopy] Re: Looking for recommendations about dual beam
} ion
}
}
}
}
} ----------------------------------------------------------------------
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} ----
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} America To Subscribe/Unsubscribe --
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}
} John
}
} I work with four Gatan PIPS daily and have found them to be quite
} acceptable in ease of both maintenance and use.
}
} We currently do maintenance only when the ion guns stop functioning,
} at that point the machine is taken out of service and a full cleaning
} performed. This happens about every five to six months and leaves the

} PIPS out of service for two days (One day to clean, pump down over
} night, and alignment the next day). The diaphragm pump needs new
} diaphragms once a year (about an hours work) but not too much else.
}
} Work load is typically one to two ceramic samples per day, with an
} occasional silicon or glass sample thrown in. The ceramic samples are

} lapped to about six microns and mill for an hour to two hours.
} Silicon and glass are left about ten microns thick and take much
} longer (up to three days), not so much due to the thickness as to the
} low kVs used and letting the sample "cool".
}
} Gatan offers a digital camera/zoom lens set and an external diaphragm
} pump as extras. In my opinion, they are a "must have".
}
} If you have any further questions please feel free to contact me.
}
} Neal R. Zimmermann
}
}
}
} On Wed, 2007-05-09 at 15:43 -0500, jpchandl-at-mines.edu wrote:
} }
} }
} }
} ----------------------------------------------------------------------
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} America
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} }
} } I need to tap in to the wisdom of the list about dual beam argon ion
} mills.
} }
} } We have a Bal-Tec RES100 ion mill that we are having difficulty
} maintaining.
} } It had been in disuse for a couple of years, and has been repaired
} } and upgraded recently, at the factory, to be as close to a RES101 as
} possible.
} } We are having issues such as the guns becoming contaminated and
} needing
} } service much more frequently than we think they should. Routine
} maintenance
} } seems to be difficult, too. We think we are looking at a high
} maintenance
} } instrument here. I would like to know what kind of experience other
} labs
} } have had with this instrument.
} }
} } I would also like to hear from people who have experience
} using/maintaining
} } the RES100/101 or the Gatan PIPS, or both. We need some comparative

} } information, so we can make a decision about how to proceed with our
} ion
} } milling needs.
} }
} } Any input would be appreciated.
} }
} } --John
} }
} } John Chandler
} } Manager, EM Lab
} } Colorado School of Mines
} } Golden, CO 80401
} } jpchandl-at-mines.edu
} } 303-384-2203
} }
} }
} }
} }
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From: colijn.1-at-osu.edu
Date: Mon, 14 May 2007 09:26:18 -0500
Subject: [Microscopy] Re: SEM-help finding smooth, low-Z substrates

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Roy,

How about using a TEM carbon support film? Disperse your particles
on the support film then mount the grid on a "Faraday cup"

I made a TEM grid support by taking about a 1cm long piece of 1/4"
graphite rod, drilling out the center 6-7mm deep, and using a 1/8"
(3.1mm) end mill to make a small recess for the grid in the end of
the rod. I used carbon dag to glue the rod upright on an SEM stub
and presto... instant TEM grid holder with near zero background!

Cheers,
Henk

At 09:32 AM 05/14/07, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Hendrik O. Colijn colijn.1-at-osu.edu
Campus Electron Optics Facility Ohio State University
(614) 292-0674 http://www.ceof.ohio-state.edu
Time is that quality of nature which keeps events from happening all
at once. Lately it doesn't seem to be working.


==============================Original Headers==============================
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From: bfoster-at-mme1.com
Date: Mon, 14 May 2007 09:49:32 -0500
Subject: [Microscopy] Posting for non-MSA member Re: Microscope Job - Please respond

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Listers,

} Caleb Adams and I have been carrying on a conversation about training and jobs in microscopy. Can any of you provide him with further information? Since Caleb is not on the listserver, please respond to him directly (Caleb Adams {cannon_adamscl-at-yahoo.com} )


Thanks,
Barbara Foster, President

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com


MME is now scheduling customized, on-site courses through July. Call us today for details.

P. S.
Need a good general reference or light microscopy text for next semester? Call us today to learn more about "Optimizing LIght Microscopy". Copies still available through MME... even for class-room lots ... and we give quantity discounts. Call Ken Piel at (972)954-8011.


} Barbara,
}
} Thank you for the thorough info. I have been working
} with my Leica CME microscope for about 4 months and
} wanted to know all the equipment that would be good to
} have with my microscope. I currently have the
} following accessories: slides and cover slips,
} dissecting tool set, darkfield, cleaner, slide and
} microscope case, and petri dish. I will be done with
} my high school within 1 to 2 years and from there
} would like to work in a non medical field of
} microscopy but in the meantime is there some online
} course in microscopy that I could take?
}
} Sincerely,
} Caleb
} --- Barbara Foster {bfoster-at-mme1.com} wrote:
}
} } Hi, Caleb
} }
} } That's a pretty hard call to make because there are
} } so many factors.
} } 1. Where is microscopy done?
} } Microscopy is used universally, but your best bet
} } would be in
} } industry or as a lab tech in a university's central
} } service lab.
} } 2. What certifications are required?
} } If you do microscopy in a clinical lab, you need to
} } have the training
} } and certification to be a lab tech. I think that
} } that is usually
} } either a 2 year or 4 year program. In industry, a
} } lot of training is
} } done "on the job", so if you have a basic
} } understanding of science
} } (the simple physics of optics is important... you
} } can buy a second
} } hand high school physics book or a review book.
} } Also, our book,
} } "Optimizing Light Microscopy" has what you need) and
} } are willing to
} } start low and work your way up, that's a possibility
} } 3. What type of microscopy?
} } It would probably be easiest to start with light
} } (optical)
} } microscopy, then expand in to atomic force or
} } electron microscopy.
} } 4. Where to find jobs?
} } I'd start traveling around the industrial parks in
} } the neighborhood
} } and trying to identify businesses that might use
} } microscopy for
} } quality control. Also, Chem & Engineering News
} } (www.acs.org) has a
} } job bank... and you might try the traditional road
} } on-line
} } (Monster.com, etc.). Also, you'd be surprised at
} } what your friends
} } and family "don't know that they know." Start
} } chatting them up, to
} } see if they know someone who knows someone who knows
} } someone.
} }
} } I guess that's it. I'd suggest that you plan for
} } some advanced
} } education along the line, but if you can get into a
} } company at an
} } entry level, they might help you pay for an
} } associates degree or even
} } further. Local societies (I don't know where you
} } are, but the New
} } York Microscopical Society) offer weekend courses
} } and some colleges
} } such as LeHigh offer 1 week/intensive programs that
} } will really give
} } you a boost. Also, the Royal Microscopical Society
} } in Oxford,
} } England offers a whole series of 1 week courses
} } (that's how I got
} } started). Sounds strange, but sometimes it is less
} } expensive to
} } travel to the UK than to spend a week at a course
} } here! Also, I know
} } that San Joaquin Delta College in California has a
} } full 2 year
} } program that has a great reputation. Dr. Judy Murphy
} } runs it.... you
} } might want to investigate it further.
} }
} } I hope that all of this is helpful.
} }
} } Please let me know if you decide to make this career
} } shift.
} } Best regards,
} } Barbara
} }
} } At 11:25 AM 5/11/2007, Caleb Adams wrote:
} } } Barbara,
} } }
} } } How hard is it to get a job doing microscopy of
} } some
} } } sort when all you have is a high school education
} } and
} } } where would I look to find those jobs?
} } }
} } } Caleb
} } }
} } }
} } }
} }
} } ____________________________________________________________________________________
} } } Bored stiff? Loosen up...
} } } Download and play hundreds of games for free on
} } Yahoo! Games.
} } } http://games.yahoo.com/games/front
} }
} }
}
}
}
}
} ____________________________________________________________________________________Pinpoint customers who are looking for what you sell.
} http://searchmarketing.yahoo.com/


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From: NWWhite-at-bwxt.com
Date: Mon, 14 May 2007 09:55:16 -0500
Subject: [Microscopy] SEM-help finding smooth, low-Z substrates

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello Roy,

Can you use a high-Z background rather than low? That should give you
dark on light in place of light on dark...

If so, you might want to investigate metal-glass braze foil. Typically
it contains Si, Cr, Fe, Ni, or some combination (don't have the specs
handy). Since it is a glassy metal, there is no grain structure to
interfere with BSE imaging and the surface is relatively smooth. There
are some undulations/waves in the surface formed when frozen from the
molten state, but maybe not too much to interfere.

Woody White
BWXT Services


-----Original Message-----
X-from: rcsaic-at-sbcglobal.net [mailto:rcsaic-at-sbcglobal.net]
Sent: Monday, May 14, 2007 9:31 AM
To: White, Woody N.

Dear Listers,

We could use some suggestions/collective experience about solid
substrates
for doing SEM imaging of dispersed small particles.

What we are imaging: silicate and oxide mineral grains and silicate
glass
particles down to the 0.1 micron size range (maybe smaller). Objective
is to
disperse the particle on a substrate and do total particle counting and
automated measuring without "losing" the smallest particles from the
data
set.

What we need: A substrate for dispersing the particles that would fit
two
requirements: 1) have a pretty low average Z (carbon would be ideal but
I'm
willing to consider other ideas), so that my particles will stand out
strongly in BSE, and then 2) be as smooth as possible on the sub-micron
scale. This last requirement is the tricky one because although there
are
lots of carbon substrates available commercially, so far we haven't
found
any that don't have sub-micron scratches and other roughness such that
particles in the 0.1 micron size range don't get lost in the surface
topography.

Well, that's about all I can think of. Thanks to all of you in advance.


Roy Christoffersen
FE-STEM Facility Manager
ARES Directorate
NASA Johnson Space Center, Houston, TX
rcsaic-at-sbcglobal.net







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10, 24 -- From: "Roy Christoffersen" {rcsaic-at-sbcglobal.net}
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21, 28 -- From nwwhite-at-bwxt.com Mon May 14 09:55:16 2007
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From: peter.tomic-at-renwireless.com
Date: Mon, 14 May 2007 10:33:21 -0500
Subject: [Microscopy] Re: Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Yes, John, it's 8 mm, not microns. The issue is holding the device
while making the second cleave or cut. The sample would have to be
mounted on something to use a diamond saw, which is available. However,
I do need to keep the face of the surface I'm looking at undisturbed
with whatever method I use.

Regards,

Peter

Mardinly, John wrote:
} ~ 8 mm apart? Did I read this correctly? Not microns? We use a Disco saw
} routinely. If it's ~ 8 mm apart, you can use any diamond saw. If it's
} microns you need a FIB.
}
} John Mardinly
} Intel Corporation
} This email does not represent an opinion of Intel Corporation.
}
} -----Original Message-----
} From: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
} Sent: Tuesday, May 08, 2007 10:33 AM
} To: Mardinly, John
} Subject: [Microscopy] Silicon Cross-section sample preparation
}
}
}
}
} ------------------------------------------------------------------------
} ----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ------------------------------------------------------------------------
} ----
}
} Microscopy Folks,
}
} I'd like to get some input on sample preparation with respect to
} silicon.
}
} PROBLEM:
}
} I have an FEI XL-50 FESEM that was originally designed to accept flat
} samples, essentially silicon wafers. It does not have what one would
} consider a typical exchange port. The exchange port is robotically
} controlled, and the maximum sample height, including stub, must be 5 mm
} or less. I must view these samples on edge, i.e. 90 degrees. This
} presents a problem in that I must cleave two parallel sections very
} close to each other. Perhaps diamond cutters can do this, but my hands
} are too shaky.
}
} QUESTION:
}
} Is there a device, or method, that would allow me to make these cleaves
} ~ 8 mm apart with any reasonable control? I found stubs that are low
} profile, 38 and 90 degrees, from the nice people at Ted Pella, but I
} still have this issue of doing two parallel cleaves very close together.
}
} Just for info. purposes I am in the silicon MEMS development arena.
}
} If you feel your reply is of general interest to this community, please
} reply to all, or you may contact me directly.
}
} Regards to all in this small world,
}
} Peter Tomic
} Renaissance Wireless Corp.
}
}
}



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From: mager-at-interchange.ubc.ca
Date: Mon, 14 May 2007 10:41:03 -0500
Subject: [Microscopy] SEM-help finding smooth, low-Z substrates

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Roy,
There is a material called "glassy carbon", which is smooth, solid and can
be polished to a mirror surface. It is very hard, but is pure carbon and so
it makes a good substrate for BSE studies. Some of the EM catalogue
companies used to supply it, they probably still do. I get my glassy carbon
planchets (0.5 inch discs) from Canemco (www.canemco.com) and stick them
onto my SEM stubs.
Good luck,

Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: rcsaic-at-sbcglobal.net [mailto:rcsaic-at-sbcglobal.net]
Sent: May 14, 2007 6:38 AM
To: mager-at-interchange.ubc.ca

Dear Listers,

We could use some suggestions/collective experience about solid substrates
for doing SEM imaging of dispersed small particles.

What we are imaging: silicate and oxide mineral grains and silicate glass
particles down to the 0.1 micron size range (maybe smaller). Objective is to
disperse the particle on a substrate and do total particle counting and
automated measuring without "losing" the smallest particles from the data
set.

What we need: A substrate for dispersing the particles that would fit two
requirements: 1) have a pretty low average Z (carbon would be ideal but I'm
willing to consider other ideas), so that my particles will stand out
strongly in BSE, and then 2) be as smooth as possible on the sub-micron
scale. This last requirement is the tricky one because although there are
lots of carbon substrates available commercially, so far we haven't found
any that don't have sub-micron scratches and other roughness such that
particles in the 0.1 micron size range don't get lost in the surface
topography.

Well, that's about all I can think of. Thanks to all of you in advance.


Roy Christoffersen
FE-STEM Facility Manager
ARES Directorate
NASA Johnson Space Center, Houston, TX
rcsaic-at-sbcglobal.net







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From: jd-at-laddresearch.com
Date: Mon, 14 May 2007 10:52:12 -0500
Subject: [Microscopy] SEM-help finding smooth, low-Z substrates

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Roy,

Just to expand on Mary's suggestion, yes, most of the EM Supply
houses sell them including us at Ladd Research. See our Specially
Smooth Carbon Planchets at
http://www.laddresearch.com/General_Catalog/Chapter_4/Specimen_Mounts/plan/plan.html

John Arnott

Ladd Research
83 Holly Court
Williston, VT 05495

On-line Catalog: www.laddresearch.com

Telephone: 1-802-658-4961 (anywhere)
Toll Free 1-800-451-3406 (US)
Fax: 1-802-660-8859

e-mail: sales-at-laddresearch.co

DISCLAIMER: Ladd Research has been selling EM supplies including
carbon planchets for more than 50 years

At 11:45 AM 5/14/2007, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

m


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From: kirk-at-udel.edu
Date: Mon, 14 May 2007 11:01:55 -0500
Subject: [Microscopy] Temporary Microscopist Position

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Part-Time Temporary Microscopist

The Delaware Biotechnology Institute (DBI) is looking to fill a
part-time temporary position of microscopist in its state-of-the-art
Bioimaging Center.

DBI, a unit of the University of Delaware, was established in 1999 to
position Delaware as a leader in biotechnology, is a partnership
involving state government, the Delaware institutions of higher
education, and area industry, and is housed in a 72,000 ft2
state-of-the-art research facility located at the Delaware Technology
Park in Newark, DE. DBI’s mission is to engage in leading-edge
scientific discovery in the life sciences, provide biotechnology-based
education, and promote economic development. The major
interdisciplinary research areas include human health, agriculture,
including plant molecular biology and avian genomics, environmental
ecosystems, and biomaterials.

The Institute houses several core facilities including a microarray
center, a bioinformatics center and a bioimaging center. The bioimaging
center houses a broad range of microscopy instrumentation, including
conventional fluorescence, confocal, multiphoton, atomic force, laser
microdissection, transmission and field emission scanning microscopes
and their ancillary sample preparation equipment. Additional details
are available on the web at www.dbi.udel.edu/bioimaging.

Under the general direction of the Director of the Bioimaging Center,
the part-time microscopist will provide research support for light and
confocal microscopy as well as scanning and transmission electron
microscopy experiments in a multi-user environment. This includes, but
is not limited to training users and/or performing all steps of
experimental design, sample preparation, data acquisition and analysis.
This part-time temporary position is available immediately and at least
through December 31, 2007. Candidates should have a minimum of a
Bachelor’s degree in the life sciences. Prior microscopy experience
would be highly advantageous. Compensation is commensurate with
experience. Please inquire or send resume via email to Dr. Kirk Czymmek
(kirk-at-udel.edu).


Kirk J. Czymmek, Ph.D.
15 Innovation Way, Suite 117
Delaware Biotechnology Institute
University of Delaware
Newark DE 19711
(302) 831-3450
kirk-at-udel.edu


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From: walck-at-southbaytech.com
Date: Mon, 14 May 2007 13:07:44 -0500
Subject: [Microscopy] Re: Silicon Cross-section sample preparation

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Peter, If you thin the samples, then a perfect holding device is the
adhesive on a Post-it(TM) note. If you don't thin your sample, but can
scribe it well enough, then consider using a flat silicone sheet to hold the
sample. Both will hold the sample for your cleave and both are just
resilient enough. At 90-100 µm thick, the Post-It(TM) note works great for
the MicroCleave(TM) technique. You have to shear it off the pack so that it
doesn't have any curl and then using regular tape, tape it to a flat block
with the adhesive side up. Carefully place your samples on it, being
careful not to put the good side down on the adhesive -you won't get the
adhesive off. On the rough ground down side, the adhesive doesn't stick
when you lift the sample off.


-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
Sent: Monday, May 14, 2007 8:37 AM
To: Walck-at-SouthBayTech.com

Yes, John, it's 8 mm, not microns. The issue is holding the device while
making the second cleave or cut. The sample would have to be mounted on
something to use a diamond saw, which is available. However, I do need to
keep the face of the surface I'm looking at undisturbed with whatever method
I use.

Regards,

Peter

Mardinly, John wrote:
} ~ 8 mm apart? Did I read this correctly? Not microns? We use a Disco
} saw routinely. If it's ~ 8 mm apart, you can use any diamond saw. If
} it's microns you need a FIB.
}
} John Mardinly
} Intel Corporation
} This email does not represent an opinion of Intel Corporation.
}
} -----Original Message-----
} From: peter.tomic-at-renwireless.com [mailto:peter.tomic-at-renwireless.com]
} Sent: Tuesday, May 08, 2007 10:33 AM
} To: Mardinly, John
} Subject: [Microscopy] Silicon Cross-section sample preparation
}
}
}
}
} ----------------------------------------------------------------------
} --
} ----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America To Subscribe/Unsubscribe --
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} ----------------------------------------------------------------------
} --
} ----
}
} Microscopy Folks,
}
} I'd like to get some input on sample preparation with respect to
} silicon.
}
} PROBLEM:
}
} I have an FEI XL-50 FESEM that was originally designed to accept flat
} samples, essentially silicon wafers. It does not have what one would
} consider a typical exchange port. The exchange port is robotically
} controlled, and the maximum sample height, including stub, must be 5
} mm or less. I must view these samples on edge, i.e. 90 degrees. This
} presents a problem in that I must cleave two parallel sections very
} close to each other. Perhaps diamond cutters can do this, but my
} hands are too shaky.
}
} QUESTION:
}
} Is there a device, or method, that would allow me to make these
} cleaves ~ 8 mm apart with any reasonable control? I found stubs that
} are low profile, 38 and 90 degrees, from the nice people at Ted Pella,
} but I still have this issue of doing two parallel cleaves very close
together.
}
} Just for info. purposes I am in the silicon MEMS development arena.
}
} If you feel your reply is of general interest to this community,
} please reply to all, or you may contact me directly.
}
} Regards to all in this small world,
}
} Peter Tomic
} Renaissance Wireless Corp.
}
}
}



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From: tivol-at-caltech.edu
Date: Mon, 14 May 2007 13:14:08 -0500
Subject: [Microscopy] Re: SEM-help finding smooth, low-Z substrates

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On May 14, 2007, at 6:29 AM, rcsaic-at-sbcglobal.net wrote:

} A substrate for dispersing the particles that would fit two
} requirements: 1) have a pretty low average Z (carbon would be ideal
} but I'm
} willing to consider other ideas), so that my particles will stand out
} strongly in BSE, and then 2) be as smooth as possible on the sub-micron
} scale. This last requirement is the tricky one because although there
} are
} lots of carbon substrates available commercially, so far we haven't
} found
} any that don't have sub-micron scratches and other roughness such that
} particles in the 0.1 micron size range don't get lost in the surface
} topography

Dear Roy,
Have you tried carbon evaporated onto freshly-cleaved mica? The layer
can be floated off and deposited onto lacy carbon so that there will be
many areas several um across that have only the thin carbon substrate.
The layer can be made to be only tens of nm thick, so it is very
unlikely to have ~100 nm steps.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu


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From: TindallR-at-missouri.edu
Date: Mon, 14 May 2007 13:26:59 -0500
Subject: [Microscopy] Leica HPF EMPACT question

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Dear Listers,

We have a Leica EMPACT high pressure freezer and upon last start up the
hydraulic fluid reservoir had gone dry. I refilled with MCH and
followed the operation manual's instructions on getting air bubbles out
of the line (basically just using a long skinny rod to plunge the
sealing "bb" at the bottom of the reservoir until the bubbles stop
coming out). Now I'm getting an error message that prompts me to fill
the chamber with hydraulic fluid.

I can reboot the system and get rid of the message, but only for one
freezing run---then it returns. Many reboots, freezes, and lots of bb
plunging later, I'm still getting the message. The pressure readings
during the HPF runs have been mostly good, but there is an occasional
stinker----more than I remember on average than in past runs.

I'm guessing there's an elusive bubble in the line somewhere, but has
anyone else run across this? More to the point, does anyone have an
easy fix?

Thanks all.

Randy

Randy Tindall
Senior EM Specialist
Electron Microscopy Core Facility---We Do Small Well!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu
On-line calendar:
http://biotech.rnet.missouri.edu/cgi-bin/Calcium310.pl?Op=Splash&Amount=
Week&NavType=Both&Type=TimePlan


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From: bigelow-at-umich.edu
Date: Mon, 14 May 2007 16:23:01 -0500
Subject: [Microscopy] R: O-rings

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The Zatkoff (www.zatkoff.com) company also carries an extensive stock
of O-rings. Before I tried making an O-ring on my own I would
certainly check with them to see if they couldn't furnish something
to meet my needs.
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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From: cgarber-at-2spi.com
Date: Mon, 14 May 2007 16:47:47 -0500
Subject: [Microscopy] SEM-help finding smooth, low-Z substrates

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Roy Christoffersen wrote:
==============================================================

We could use some suggestions/collective experience about solid substrates
for doing SEM imaging of dispersed small particles.

What we are imaging: silicate and oxide mineral grains and silicate glass
particles down to the 0.1 micron size range (maybe smaller). Objective is to
disperse the particle on a substrate and do total particle counting and
automated measuring without "losing" the smallest particles from the data
set.

What we need: A substrate for dispersing the particles that would fit two
requirements: 1) have a pretty low average Z (carbon would be ideal but I'm
willing to consider other ideas), so that my particles will stand out
strongly in BSE, and then 2) be as smooth as possible on the sub-micron
scale. This last requirement is the tricky one because although there are
lots of carbon substrates available commercially, so far we haven't found
any that don't have sub-micron scratches and other roughness such that
particles in the 0.1 micron size range don't get lost in the surface
topography.

Well, that's about all I can think of. Thanks to all of you in advance.


Roy Christoffersen
FE-STEM Facility Manager
ARES Directorate
NASA Johnson Space Center, Houston, TX
rcsaic-at-sbcglobal.net
=======================================================================
In my opinion, you should consider HOPG (highly ordered pyrolytic graphite), see URL

For those who are not familiar with this unique material and its novel properties, it can be cleaved much like mica into
very thin strippings, actually more easily than mica. This is done by pressing a piece of Scotch tape, either single
sided or double sided onto the flat plate surface of the HOPG and then literally, stripping off a layer of HOPG which is
left strongly adhering to the tape. This tape can then be mounted on a conventional SEM mount, perhaps with a double
sided conductive carbon disc. Many additional strippings can be made from the resulting block until it is all consumed.

The resulting stripping is highly (mirror) reflective, demonstrating the virtually zero porosity in the HOPG stripped
layer. And depending on the grade of HOPG selected, there will be regions of varying area sizes of atomically smooth
HOPG. You will not "lose" particles in the structure of the HOPG.

There have been some suggestions by others, but even though we too offer glassy carbon, we recommend the HOPG over the
glassy carbon for several reasons: a) Glassy carbon exhibits some porosity (depending on grade) on the scale of your
fine particles that you don't want to lose and b) since one can get a number of individual strippings out of each HOPG
plate, the cost per stripping (e.g. sample) is far less than the cost of a piece of glassy carbon.

HOPG is used widely in the field of AFM, one of the reasons being exactly the reason you gave: ".......don't have
sub-micron scratches and other roughness such that particles in the 0.1 micron size range don't get lost in the surface
topography". Your specifications are consistent with the needs of AFM users looking at nano-sized particles (which can
not be cleaved).

Disclaimer: SPI Supplies offers a complete line of HOPG and glassy carbon products and therefore we have a vested
interest in seeing more of these items being used by researchers. See URL
http://www.2spi.com/catalog/mounts/vitreous.html

Chuck

PS: Remember that we are striving to be 100% paperless, therefore there are no paper copies kept of this
correspondence. Please be sure to always reply by way of "reply" on your software so that the entire string of
correspondence can be kept in one place.


===================================================
Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
President
SPI SUPPLIES FAX: 1-(610)-436-5755
PO BOX 656 e-mail: cgarber-at-2spi.com
West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com


Look for us!
############################
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From: gas19-at-daimlerchrysler.com
Date: Mon, 14 May 2007 19:00:08 -0500
Subject: [Microscopy] viaWWW: SEM-help finding smooth, low-Z substrates

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Email: gas19-at-daimlerchrysler.com
Name: Gerald Shulke

Organization: DaimlerChrysler Corporation

Title-Subject: [Filtered] Re:[Microscopy] SEM-help finding smooth, low-Z substrates

Question: Roy,
Although I normall don't work with particles that small, most of our particle analysis is performed on carbon conductive tabs from Ted Pella Inc (catalog # 16084-4). There is some texture to the tabs, but usually the particles stand proud of the tab. With a backscatter image, the higher z particles will stand out from the carbon. The problem with analyzing such a fine powder will be particle separation. We tend to sprinkle a small sample on the tab instead of pressing the tab into the powder, just to get a dispersion, provide spearation between particles, and to have the particle be proud of the tab.

Gerald Shulke
Materials Characterization Labs
DaimlerChrysler- Chrysler Group

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From: cannonmp-at-comcast.net
Date: Mon, 14 May 2007 19:50:45 -0500
Subject: [Microscopy] Smooth Low Z Substrate

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Hi Roy,

Looks like Carbon is the idea material for your research. Have you tried to
use natural graphite crystal -- it has beautiful layer structure and large
flat surface area as well. Find a geologist and ask a piece of graphic
crystal, stick it onto your SEM stub, then peel off first 3-10 layers to
have fresh surface ready for use. Use sharp knife and peel from edge -- I
saw some other people used tape to peel the top layers off. We used to make
fresh "flaw-free" graphite surface for STM analysis.

Regards,
----------------------------------------------
Xiang Yang
Electron Microscopy Center
Saint Mary's University
Science Building, Suite 135
Halifax, NS Canada B3H 3C3
Tel: (902) 496-8292 (lab)
(902) 420-5709 (off)
http://fgsr.smu.ca/emc/


----- Original Message -----
X-from: {rcsaic-at-sbcglobal.net}
To: {xyang-at-SMU.CA}
Sent: Monday, May 14, 2007 10:34 AM

Roy,

The artful use of a heat gun directed at a hot melt glue covered SEM stub
will produce a lustrous surface on the low Z, low outgassing glue substrate.
Perhaps while the glue is still warm and adhesive, you could sprinkle your
sample onto it.

Bart Cannon
Cannon Microprobe
Seattle


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From: tnkeim-at-sbcglobal.net
Date: Mon, 14 May 2007 20:01:57 -0500
Subject: [Microscopy] viaWWW: Run Zeiss 100W from Instek Power Supply

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Email: tnkeim-at-sbcglobal.net
Name: Thomas Keim

Organization: Dentist

Title-Subject: [Filtered] Run Zeiss 100W from Instek Power Supply

Question: Has anyone tried powering the Zeiss 100W halogen light source (Zeiss 467259, for Standards and Photomicroscopes) with an Intek GPR-1810HD power supply? Any issues?

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From: kya22-at-student.canterbury.ac.nz
Date: Tue, 15 May 2007 07:31:15 -0500
Subject: [Microscopy] viaWWW: Cleaning Surfaces of Polysulfone membrane for SEM

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Email: kya22-at-student.canterbury.ac.nz
Name: kuldeep

Organization: University of Canterbury

Title-Subject: [Filtered] Cleaning Surfaces of Polysulfone membrane for SEM

Question: Hi,

I am doing SEM and FESEM scanning of Polysulfone membrane.
Although I always wash the membrane with water and alchol
prior to scanning but I am getting foreign particles on new (comes sealed from industry) and casted membranes surface. I dont know what these particle are how to clean them ( I suppose they are some dust embedded on porous membrane surface)

Can anybody tell me some standard process to get rid of these foreign particles on the surface.
I can send Pics showing these particles on the surface.

Thanks and Regards

Kuldeep

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From: kraftpiano-at-gmail.com
Date: Tue, 15 May 2007 07:33:16 -0500
Subject: [Microscopy] SEM Bacterial sample prep.

Contents Retrieved from Microscopy Listserver Archives
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I have obtained a sample of freeze-dried bacteria, and would like to
prepare it for the SEM. Is there something more than just
double-stick carbon disk on a stub and sputter coating if it's already
freeze dried?

--Justin A. Kraft
Atlantis Academy

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2, 26 -- Subject: SEM Bacterial sample prep.
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From: Robert.Zonis-at-sanford.com
Date: Tue, 15 May 2007 08:08:48 -0500
Subject: [Microscopy] High-magnification imaging help and recommendations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


To all:

I'd like to get some high-magnification images of the 1.00mm tungsten carbide balls that we use in our ball point and roller ball pens. The images I've been able to get with light microscopy do not show enough surface detail. Can anyone recommend the best type of imaging for this study and a facility that will do that sort of imaging for us inexpensively?

Thanks very much,

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


This message may contain information that is confidential and/or protected by law. If the reader of this message is not the intended recipient, you are hereby notified that any dissemination, distribution, copying or communication of this message is strictly prohibited. If you have received this communication in error, please contact the sender immediately and delete the message. Please note that although we will take all commercially reasonable efforts to prevent viruses from being transmitted from our systems, it is the responsibility of the recipient to check for and prevent adverse action by viruses on its own systems.

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From: shauna_devarennes-at-phac-aspc.gc.ca
Date: Tue, 15 May 2007 09:38:33 -0500
Subject: [Microscopy] Re: Leica HPF EM PACT question

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Hi Randy,
Sounds like a problem we had recently with our EM PACT2. We tried draining
the entire hydraulic fluid chamber and refilling, as well as using a stick
to release any air bubbles. This was still not working, and the problem
ended up being a failed detector for the fluid level. We contacted Leica
and received a new detector for the instrument, which was quite easy to
install. So you may want to contact them as well to discuss the problem.
Hope that helps!

Shauna

Shauna deVarennes, B.Sc, AHT, RLAT
Research Technician
Diagnostic Imaging and Microscopy
Viral Diseases Division
National Microbiology Laboratory

Canadian Science Centre for Human and Animal Health
1015 Arlington Street, Room H4125
Winnipeg, Manitoba, R3E 3R2

Phone: (204) 789-5007
Email: shauna_devarennes-at-phac-aspc.gc.ca

Dear Listers,

We have a Leica EMPACT high pressure freezer and upon last start up the
hydraulic fluid reservoir had gone dry. I refilled with MCH and
followed the operation manual's instructions on getting air bubbles out
of the line (basically just using a long skinny rod to plunge the
sealing "bb" at the bottom of the reservoir until the bubbles stop
coming out). Now I'm getting an error message that prompts me to fill
the chamber with hydraulic fluid.

I can reboot the system and get rid of the message, but only for one
freezing run---then it returns. Many reboots, freezes, and lots of bb
plunging later, I'm still getting the message. The pressure readings
during the HPF runs have been mostly good, but there is an occasional
stinker----more than I remember on average than in past runs.

I'm guessing there's an elusive bubble in the line somewhere, but has
anyone else run across this? More to the point, does anyone have an
easy fix?

Thanks all.

Randy

Randy Tindall
Senior EM Specialist
Electron Microscopy Core Facility---We Do Small Well!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu

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From: reinhard.rachel-at-biologie.uni-regensburg.de
Date: Tue, 15 May 2007 12:47:31 -0500
Subject: [Microscopy] SEM Bacterial sample prep.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


} I have obtained a sample of freeze-dried bacteria, and would like to
} prepare it for the SEM. Is there something more than just
} double-stick carbon disk on a stub and sputter coating if it's
} already
} freeze dried?
}
} --Justin A. Kraft
} Atlantis Academy

X-from my - limited - experience with preparing Bacteria for SEM, I can
say that I would NOT recommend to just apply 'freeze-dried bacteria' to
a double-stick carbon disk, and then sputter-coat the stuff.
These freeze-dried bacteria are most likely completely collapsed and
therefore will look 'collapsed' and 'dead'.
I would suggest to recultivate the cells in a suitable culture medium,
then either fix them chemically, apply them to a filter (like
'Nuclepore'), dehydrate, followed by critical-point drying and sputter
coating, OR: apply them to a filter, fix them physically
(plunge-freezing), then dehydrate at low temperature by freeze-drying
at -80 C, then sputter-coating at low temperature or even
rotary-shadowing at low-temperature ... and then SEM.
Depending on the type of bacteria, many variations of these protocols
are possible and might be necessary.

regards
Reinhard Rachel

----------------------
PD Dr. Reinhard Rachel
Universitaet Regensburg
Centre for EM - NWF III -
Lehrstuhl fuer Anatomie
Universitaetsstr. 31
D-93053 Regensburg - Germany




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From: mager-at-interchange.ubc.ca
Date: Tue, 15 May 2007 12:58:40 -0500
Subject: [Microscopy] High-magnification imaging help and recommendations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Robert,
No one else has answered your question, so I will try. I think your best bet
would be to image the surface of your balls by Scanning Electron Microscopy
(SEM). This should be available in the closest university or at most failure
analysis consulting engineering firms. SEMs are fairly common and your
requirements are fairly simple. The SEM should be able to go from 50 times
magnification to 10,000 times with a good depth of focus, so you can see the
details on the rounded surface. You should be able to get some good images
in less than an hour.
Good luck.

Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: Robert.Zonis-at-sanford.com [mailto:Robert.Zonis-at-sanford.com]
Sent: May 15, 2007 6:13 AM
To: mager-at-interchange.ubc.ca


To all:

I'd like to get some high-magnification images of the 1.00mm tungsten
carbide balls that we use in our ball point and roller ball pens. The images
I've been able to get with light microscopy do not show enough surface
detail. Can anyone recommend the best type of imaging for this study and a
facility that will do that sort of imaging for us inexpensively?

Thanks very much,

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


This message may contain information that is confidential and/or protected
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hereby notified that any dissemination, distribution, copying or
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From: beth-at-plantbio.uga.edu
Date: Tue, 15 May 2007 13:09:42 -0500
Subject: [Microscopy] High-magnification imaging help and recommendations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

What a hoot!
Your message made me LOL...I'll be chuckling about that for the rest
of the day;-)
Thanks!

On May 15, 2007, at 1:59 PM, mager-at-interchange.ubc.ca wrote:

}
}
}
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}
} Dear Robert,
} No one else has answered your question, so I will try. I think your
} best bet
} would be to image the surface of your balls by Scanning Electron
} Microscopy
} (SEM). This should be available in the closest university or at
} most failure
} analysis consulting engineering firms. SEMs are fairly common and your
} requirements are fairly simple. The SEM should be able to go from
} 50 times
} magnification to 10,000 times with a good depth of focus, so you
} can see the
} details on the rounded surface. You should be able to get some good
} images
} in less than an hour.
} Good luck.
}
} Mary Fletcher (nee Mager)
} Electron Microscopist
} Materials Eng. UBC
} #309 - 6350 Stores Road
} Vancouver, B.C. V6T 1Z4
} Canada
} Tel: 604-822-5648
} Fax: 604-822-3619
} email: mager-at-interchange.ubc.ca
}
} -----Original Message-----
} X-from: Robert.Zonis-at-sanford.com [mailto:Robert.Zonis-at-sanford.com]
} Sent: May 15, 2007 6:13 AM
} To: mager-at-interchange.ubc.ca
} Subject: [Microscopy] High-magnification imaging help and
} recommendations
}
}
}
}
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}
} To all:
}
} I'd like to get some high-magnification images of the 1.00mm tungsten
} carbide balls that we use in our ball point and roller ball pens.
} The images
} I've been able to get with light microscopy do not show enough surface
} detail. Can anyone recommend the best type of imaging for this
} study and a
} facility that will do that sort of imaging for us inexpensively?
}
} Thanks very much,
}
} Robert Zonis
} Product Development Chemist, LMTC
} Sanford L.P. - A Newell Rubbermaid Company
} 3 Sharpie Way
} Shelbyville, TN 37160
} Direct: +1 (931) 685-6635
} Fax: +1 (931) 685-6613
} robert.zonis-at-sanford.com
} www.papermate.com
}
}
} This message may contain information that is confidential and/or
} protected
} by law. If the reader of this message is not the intended
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} hereby notified that any dissemination, distribution, copying or
} communication of this message is strictly prohibited. If you have
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} 18, 36 -- Date: Tue, 15 May 2007 10:58:07 -0700
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} and recommendations
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**********************************************************************
Beth Richardson
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Plant Biology Department
University of Georgia
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From: Sven.Terclavers-at-med.kuleuven.be
Date: Tue, 15 May 2007 13:56:32 -0500
Subject: [Microscopy] High-magnification imaging help and recommendations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

You also might consider a stereomicroscope with well-adjusted
reflected light. Zeiss' stereomicroscope LUMAR V.12 e.g. has a
magnification up to 150x and I know there's a new one coming, the
V.20, with magnifications up to 400x (if I remember well). Maybe this
could do your job! If you want I could make a few photos for you, but
than contact me off-list please!
Greets,

Sven Terclavers

-----Original Message-----
X-from: mager-at-interchange.ubc.ca [mailto:mager-at-interchange.ubc.ca]
Sent: 15 May 2007 20:01
To: sven.terclavers-at-med.kuleuven.be

Dear Robert,
No one else has answered your question, so I will try. I think your best bet
would be to image the surface of your balls by Scanning Electron Microscopy
(SEM). This should be available in the closest university or at most failure
analysis consulting engineering firms. SEMs are fairly common and your
requirements are fairly simple. The SEM should be able to go from 50 times
magnification to 10,000 times with a good depth of focus, so you can see the
details on the rounded surface. You should be able to get some good images
in less than an hour.
Good luck.

Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: Robert.Zonis-at-sanford.com [mailto:Robert.Zonis-at-sanford.com]
Sent: May 15, 2007 6:13 AM
To: mager-at-interchange.ubc.ca


To all:

I'd like to get some high-magnification images of the 1.00mm tungsten
carbide balls that we use in our ball point and roller ball pens. The images
I've been able to get with light microscopy do not show enough surface
detail. Can anyone recommend the best type of imaging for this study and a
facility that will do that sort of imaging for us inexpensively?

Thanks very much,

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


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From: bfoster-at-mme1.com
Date: Tue, 15 May 2007 15:45:38 -0500
Subject: [Microscopy] Re: High-magnification imaging help and

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I agree with Sven with one exception: you might want to try oblique
illumination or even darkfield. The off-set illumination will give a
shadowing effect that will bring out surface information.

Hope this was helpful,
Barbara Foster, President

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com


MME is now scheduling customized, on-site courses through July. Call
us today for details.

P. S.
Need a good general reference or light microscopy text for next
semester? Call us today to learn more about "Optimizing LIght
Microscopy". Copies still available through MME... even for
class-room lots ... and we give quantity discounts. Call Ken Piel at
(972)954-8011.
At 02:28 PM 5/15/2007, Sven.Terclavers-at-med.kuleuven.be wrote:



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From: donc-at-asmicro.com
Date: Tue, 15 May 2007 22:42:56 -0500
Subject: [Microscopy] Re: [a] High-magnification imaging help and recommendations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Robert,
I suggest trying Atomic Force Microscopy. This will provide true
3-dimensional information. Furthermore, we have proprietary software that
can compute Rk roughness parameters from this data. The Rk parameters are a
unique take on roughness and find use when wear and lubrication (liquid
supply) are important.
Further information about Rk is available at
http://www.asmicro.com/software/rk.htm



Disclaimer: we are in business to provide the services and software
described above.

regards,
Don
=============================================
Don Chernoff, Ph.D., President
Advanced Surface Microscopy, Inc. E-Mail: donc-at-asmicro.com
3250 N. Post Rd., Ste. 120 Voice: 317-895-5630
INDIANAPOLIS IN 46226 USA Toll free: 800-374-8557 (in USA & Canada)
web: http://www.asmicro.com Fax: 317-895-5652
[business activities: analytical services in AFM, AFM probes, consulting,
training,
calibration and test specimens, calibration and measurement software,
used NanoScope equipment.]
=============================================

----- Original Message -----
From: Robert.Zonis-at-sanford.com
To: donc-at-asmicro.com
Sent: Tuesday, May 15, 2007 9:12 AM
Subject: [a] [Microscopy] High-magnification imaging help and
recommendations





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To all:

I'd like to get some high-magnification images of the 1.00mm tungsten
carbide balls that we use in our ball point and roller ball pens. The images
I've been able to get with light microscopy do not show enough surface
detail. Can anyone recommend the best type of imaging for this study and a
facility that will do that sort of imaging for us inexpensively?

Thanks very much,

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


This message may contain information that is confidential and/or protected
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From: mbisher-at-princeton.edu
Date: Tue, 15 May 2007 22:43:56 -0500
Subject: [Microscopy] viaWWW: Free JEOL Tungsten Filaments

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Email: mbisher-at-princeton.edu
Name: Peggy Bisher

Organization: Princeton University

Title-Subject: [Filtered] Free JEOL Tungsten Filaments

Question: We are doing some spring cleaning and came across some boxes of K-type JEOL filaments. There are 14 of them total.

They are free, to the first person who emails me. Just pay for the postage to mail them.


Margaret(Peggy) E. Bisher
Electron Microscopy & Histology Core Facility Manager
Department of Molecular Biology
Princeton University
Moffett Laboratory, Room 113
Princeton, New Jersey
Office: (609) 258-7026
Fax: (609) 258-8468
mbisher-at-princeton.edu



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From: nizets2-at-yahoo.com
Date: Wed, 16 May 2007 00:01:03 -0500
Subject: [Microscopy] TEM : objective aperture and EDX - THANKS!!

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Thank you all for your numerous and very instructive
answers. I am happy to finally have clear answers and
clear solutions to this question.

Best regards,
Stephane



____________________________________________________________________________________
8:00? 8:25? 8:40? Find a flick in no time
with the Yahoo! Search movie showtime shortcut.
http://tools.search.yahoo.com/shortcuts/#news

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From: keith.morris-at-ucl.ac.uk
Date: Wed, 16 May 2007 07:34:28 -0500
Subject: [Microscopy] Re: [a] High-magnification imaging help and

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Robert,

I have tried all types of optical and atomic force microscopes for this sort
of thing and they are all plagued a limited depth of field (or just aren't
quite as good as SEM). You might even get better images using an SLR digital
camera and extension rings rather than a cheap stereo microscope simply
because you can easily turn the F stop down to 22.

However, really the only way to go is a scanning electron microscope (SEM).
The resolution is in nanometres and the incredible depth of field simply
puts it in another league (and I a light microscopist by trade). I remember
viewing a burnt pulverised fuel ash cenosphere and marvelling at the surface
detail where you could clearly see the crater holes where volatile material
had burnt off from inside. Turning the cenosphere round we found that it has
split in half and you could see the multi micro-channels left in the carbon
residue leading to surface craters. Likewise pollen & hard bodied insects
look fantastic. You have the perfect sample, it's dead, hard, unaffected by
vacuum and even conducts, you simply have to fix them to an SEM stub and
sputter coat. You don't even have any colour in the sample to worry about.
And you can get elemental analyses and size data from the specimens (but it
costs more and you probably know that already).

I would have thought any local university/college (ideally a materials
department) would image your biro balls using SEM for under $200, as it's
only a few hours work if you only have a few samples. Ring round if money is
tight as prices do vary. The SEM will have dedicated support staff to help
you. There are also many small technical support companies who will do this
type of SEM work. You probably don't need a sophisticated SEM, which will
keep the cost down. I've tried atomic force etc.. and for surface detail, &
if your sample can stand it, nothing compares.

Keith

---------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology
Institute of Ophthalmology
UCL, London EC1V 9EL



-----Original Message-----
X-from: donc-at-asmicro.com [mailto:donc-at-asmicro.com]
Sent: 16 May 2007 04:50
To: keith.morris-at-ucl.ac.uk

Dear Robert,
I suggest trying Atomic Force Microscopy. This will provide true
3-dimensional information. Furthermore, we have proprietary software that
can compute Rk roughness parameters from this data. The Rk parameters are a
unique take on roughness and find use when wear and lubrication (liquid
supply) are important.
Further information about Rk is available at
http://www.asmicro.com/software/rk.htm



Disclaimer: we are in business to provide the services and software
described above.

regards,
Don
=============================================
Don Chernoff, Ph.D., President
Advanced Surface Microscopy, Inc. E-Mail: donc-at-asmicro.com
3250 N. Post Rd., Ste. 120 Voice: 317-895-5630
INDIANAPOLIS IN 46226 USA Toll free: 800-374-8557 (in USA & Canada)
web: http://www.asmicro.com Fax: 317-895-5652
[business activities: analytical services in AFM, AFM probes, consulting,
training,
calibration and test specimens, calibration and measurement software,
used NanoScope equipment.]
=============================================

----- Original Message -----
From: Robert.Zonis-at-sanford.com
To: donc-at-asmicro.com
Sent: Tuesday, May 15, 2007 9:12 AM
Subject: [a] [Microscopy] High-magnification imaging help and
recommendations





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To all:

I'd like to get some high-magnification images of the 1.00mm tungsten
carbide balls that we use in our ball point and roller ball pens. The images
I've been able to get with light microscopy do not show enough surface
detail. Can anyone recommend the best type of imaging for this study and a
facility that will do that sort of imaging for us inexpensively?

Thanks very much,

Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
3 Sharpie Way
Shelbyville, TN 37160
Direct: +1 (931) 685-6635
Fax: +1 (931) 685-6613
robert.zonis-at-sanford.com
www.papermate.com


This message may contain information that is confidential and/or protected
by law. If the reader of this message is not the intended recipient, you are
hereby notified that any dissemination, distribution, copying or
communication of this message is strictly prohibited. If you have received
this communication in error, please contact the sender immediately and
delete the message. Please note that although we will take all commercially
reasonable efforts to prevent viruses from being transmitted from our
systems, it is the responsibility of the recipient to check for and prevent
adverse action by viruses on its own systems.

______________________________________________________________________
This email has been scanned by the MessageLabs Email Security System.
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______________________________________________________________________


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From: xiuhuixin-at-yahoo.com.cn
Date: Wed, 16 May 2007 07:57:11 -0500
Subject: [Microscopy] viaWWW: STEM-EDX

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Email: xiuhuixin-at-yahoo.com.cn
Name: Huixin Xiu

Title-Subject: [Filtered] STEM-EDX

Question: Dear all,

I have a weird problem with my STEM-EDX results. The sample is a FIB-prepared multi-layered cross-sectional sample. The structure of the sample is around 800nm Au, 50nm Ni, 50nm Au and 50nm Ni and also 500nm AlGaN. When I put the probe (the diametre about 2nm) in the Au/Ni area, there were very obvious Ga peaks. I thought it was due to X-ray scattering everywhere, but the problem is that the Ga K peak is spuriously strong in Au/Ni area compared with K peak. It seems that the Ga K intensity follows the trend of Au L intensity, but the Ga L intensity does not. When the probe was in AlGaN area, this phenomenon was not present. Could anybody kindly help me to explain why? Many thanks.

Yours sincerely,
Helen

---------------------------------------------------------------------------

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From: colijn.1-at-osu.edu
Date: Wed, 16 May 2007 09:27:00 -0500
Subject: [Microscopy] Re: viaWWW: STEM-EDX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Huixin,

The key bit of information is in the first line of your
description... "a FIB-prepared..." The FIBs use a Ga ion
source. You are seeing the ion implantation which always occurs (to
a greater or lesser extent) in every FIB sputtered foil.

I generally see ~1% Ga in most of my ex-situ liftout samples which
have been milled at 30kV. Redeposition of sputtered material can
contain up to 30% Ga.

Cheers,
Henk



At 08:59 AM 05/16/07, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Hendrik O. Colijn colijn.1-at-osu.edu
Campus Electron Optics Facility Ohio State University
(614) 292-0674 http://www.ceof.ohio-state.edu
Time is that quality of nature which keeps events from happening all
at once. Lately it doesn't seem to be working.


==============================Original Headers==============================
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From: raristau-at-ims.uconn.edu
Date: Wed, 16 May 2007 11:07:42 -0500
Subject: [Microscopy] Re: viaWWW: STEM-EDX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

You should consider doing a low-energy, low-angle "cleaning" as the last
step in the FIB process. This can be effective in removing much of the Ga
implanted from previous FIB steps.

Cheers

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5379
fax: 860-486-4745



}
} Email: xiuhuixin-at-yahoo.com.cn
} Name: Huixin Xiu
}
} Title-Subject: [Filtered] STEM-EDX
}
} Question: Dear all,
}
} I have a weird problem with my STEM-EDX results. The sample is a FIB-prepared
} multi-layered cross-sectional sample. The structure of the sample is around
} 800nm Au, 50nm Ni, 50nm Au and 50nm Ni and also 500nm AlGaN. When I put the
} probe (the diametre about 2nm) in the Au/Ni area, there were very obvious Ga
} peaks. I thought it was due to X-ray scattering everywhere, but the problem is
} that the Ga K peak is spuriously strong in Au/Ni area compared with K peak. It
} seems that the Ga K intensity follows the trend of Au L intensity, but the Ga
} L intensity does not. When the probe was in AlGaN area, this phenomenon was
} not present. Could anybody kindly help me to explain why? Many thanks.
}
} Yours sincerely,
} Helen
}


==============================Original Headers==============================
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From: rkisensee-at-ors-labs.com
Date: Wed, 16 May 2007 20:28:09 -0500
Subject: [Microscopy] viaWWW: Jeol 820 stage motors

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Email: rkisensee-at-ors-labs.com
Name: Bob Isensee

Organization: Oneida Research Services, Inc.

Title-Subject: [Filtered] Jeol 820 stage motors

Question: Does anyone know of where we might find replacement x-y stage motors for a JEOL 820 SEM.(new used or bone yard ---anything!) JEOL won't have anything until August and the motor manufacturer doesn't make this style of motor anymore.

---------------------------------------------------------------------------

==============================Original Headers==============================
6, 11 -- From zaluzec-at-microscopy.com Wed May 16 20:28:09 2007
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From: sbledsoe-at-iupui.edu
Date: Wed, 16 May 2007 20:28:23 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

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Email: sbledsoe-at-iupui.edu
Name: Sharon Bledsoe

Title-Subject: [Filtered] Need Help From Imaging Experts

Question: Here is my problem: I can only use Mac iMovie to edit my digital video. I can not do this on a PC. ONLY iMovie. To save a single frame from the video, my choices are either PICT or JPEG format.

My Question is: To save an origianl image, which is the best format, PICT or JPEG?

I'm not interested in hearing about Tiffs, BMPs or any other format as that is not one of my choices.

---------------------------------------------------------------------------

==============================Original Headers==============================
7, 11 -- From zaluzec-at-microscopy.com Wed May 16 20:28:23 2007
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From: mbisher-at-princeton.edu
Date: Wed, 16 May 2007 20:28:48 -0500
Subject: [Microscopy] viaWWW: Free JEOL Tungsten Filaments Are PROMISED

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Email: mbisher-at-princeton.edu
Name: Peggy Bisher

Organization: Princeton University

Title-Subject: [Filtered] Free JEOL Tungsten Filaments Are PROMISED

Question: To all of you who wrote and asked for the filaments, thank you.
They have been promised.
It was nice to get messages from so many interested people, I'm sorry I did not have more to go around....

Cheers, Peggy

---------------------------------------------------------------------------

==============================Original Headers==============================
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From: Elliott-at-arizona.edu
Date: Wed, 16 May 2007 22:15:37 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
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JPEG with the lowest compression you can get.
PICT is a format on the way out (it was never used much anyway).
David


On May 16, 2007, at 6:34 PM, sbledsoe-at-iupui.edu wrote:

}
}
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}
} Email: sbledsoe-at-iupui.edu
} Name: Sharon Bledsoe
}
} Title-Subject: [Filtered] Need Help From Imaging Experts
}
} Question: Here is my problem: I can only use Mac iMovie to edit my
} digital video. I can not do this on a PC. ONLY iMovie. To save a
} single frame from the video, my choices are either PICT or JPEG
} format.
}
} My Question is: To save an origianl image, which is the best
} format, PICT or JPEG?
}
} I'm not interested in hearing about Tiffs, BMPs or any other format
} as that is not one of my choices.
}
} ----------------------------------------------------------------------
} -----
}
} ==============================Original
} Headers==============================
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} Headers==============================


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From: scott.coutts-at-med.monash.edu.au
Date: Thu, 17 May 2007 00:45:48 -0500
Subject: [Microscopy] Bacterial flagella

Contents Retrieved from Microscopy Listserver Archives
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Hi,

Does anyone have any tips for preparing and imaging bacterial flagella
at reasonably high resolutions? I need to examine the flagella and
measure potential differences in their width, so I'd like to image them
at around 100-200kx. The flagella are about 25nm wide.

So far, I've been making cell-free flagella preps and adsorbing this to
carbon-coated copper or nickel grids, and then staining with PTA. They
don't look too bad, but I'd like to know if there would be better
methods for this kind of thing.

Cheers,

--
Scott J. Coutts
-------------------------------------------------------------------------
Bacterial Pathogenesis Research Group
Department of Microbiology
PO Box 53, Monash University, 3800, Australia
Phone +61 3 9905 8592, Fax +61 3 9905 4811
-------------------------------------------------------------------------


==============================Original Headers==============================
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From: reinhard.rachel-at-biologie.uni-regensburg.de
Date: Thu, 17 May 2007 05:58:38 -0500
Subject: [Microscopy] TEM: Bacterial flagella

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Quoting scott.coutts-at-med.monash.edu.au:

} Does anyone have any tips for preparing and imaging bacterial
} flagella
} at reasonably high resolutions? I need to examine the flagella and
} measure potential differences in their width, so I'd like to image
} them at around 100-200kx. The flagella are about 25nm wide.
}
} So far, I've been making cell-free flagella preps and adsorbing this
} to carbon-coated copper or nickel grids, and then staining with PTA.
} They
} don't look too bad, but I'd like to know if there would be better
} methods for this kind of thing.

Carbon-coated (copper) grids: yes
if you just want to know their width, negative staining appears
appropriate and a final magnification of 30.000 to 50.000 x (on film,
on CCD-camera) should be sufficient.
Stain: PTA might be ok, but you may also try other stains in parallel
samples, like Uranylacetate or Uranylformate. You also may add some
sugar derivatives (glucose, trehalose) for preventing / minimizing the
collapse of the flagella upon air-drying. (how much time do you have?
how many samples?how much effort do you want to put into this project?)
Ultimately, you may try cryo-TEM of unstained samples, this will give
you the samples with highest quality.

best regards,
Reinhard Rachel

----------------------
PD Dr. Reinhard Rachel
Universitaet Regensburg
Centre for EM - NWF III -
Lehrstuhl fuer Anatomie
Universitaetsstr. 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720, 1666(TEM)
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-r.de
office: VKL 3.1.29


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From: mmcheath-at-syr.edu
Date: Thu, 17 May 2007 07:25:27 -0500
Subject: [Microscopy] Excess JEOL 733 through 8600 Electron microprobes

Contents Retrieved from Microscopy Listserver Archives
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Hi,
I am looking for surplus, excess, unused JEOL 733 through
8600 vintage electron microprobes. If you have one (733, 840, 6300,
8600) that you need to get rid of or are using as an SEM only and
want to get rid of the WDS's I would like to speak with you.

Cheers
Mike


--
********************************************************************
Michael M. Cheatham
312 Heroy Geology Laboratory Phone (315)-443-1261
Syracuse University Fax (315)-443-3363
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From: michael-at-shaffer.net
Date: Thu, 17 May 2007 07:27:29 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
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David writes ...

} JPEG with the lowest compression you can get.
} PICT is a format on the way out (it was never used much anyway).

Although I don't have any problem with this suggestion --- if there were any
benefit for the OP to export single frames as PICT (e.g., avoiding lossy
compression), there are a number of softwares capable of converting PICT to
any format desired (e.g., Photoshop), which would include batch processing.

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
Inco Innovation Centre
Memorial University
St. John's, Newfoundland


} On May 16, 2007, at 6:34 PM, sbledsoe-at-iupui.edu wrote:
} } Question: Here is my problem: I can only use Mac iMovie to edit my
} } digital video. I can not do this on a PC. ONLY iMovie. To save a
} } single frame from the video, my choices are either PICT or JPEG
} } format.
} }
} } My Question is: To save an origianl image, which is the
} best format,
} } PICT or JPEG?
} }
} } I'm not interested in hearing about Tiffs, BMPs or any
} other format as that is not one of my choices.


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From: mike.reedy-at-cellbio.duke.edu
Date: Thu, 17 May 2007 09:19:56 -0500
Subject: [Microscopy] Re: Bacterial flagella

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Scott-
Hit Pubmed and look up the pioneering experts' papers, Namba,
DeRosier especially (hope your institution has e-Journal access to J
Mol Biol all the way back!). I've not read them to see how they
compensate for flattening. But in my field of interest, muscle
myosin filaments, the classic way to minimize flattening is Tannic
Acid followed by UrAc for neg stain, as in
(1986) Stewart, M. and Kensler, R. W. Arrangement of myosin
heads in relaxed thick filaments from frog skeletal muscle. J. Mol.
Biol. 192:831-851.
I think a still earlier 1985 paper about scorpion filaments may have
introduced the method.

I think Kensler has used that method ever since, certainly in
2ournal of Structural Biology 137, 154-163 (2002) where it is
described in detail. I'll send you that paper off-list.

-mike reedy-

At 12:50 AM -0500 5/17/07, scott.coutts-at-med.monash.edu.au wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America


--
-mike reedy-

************************
Michael K. Reedy, M.D.
Duke Univ. Med. Center
Dept. Cell Biology, Box 3011 (for U.S. Mail)
458 Alex Sands Bldg, Research Dr (courier)
Durham, NC 27710

Office 919-668-2534
Lab 919-684-5674
Fax 919-681-9929
mike.reedy-at-cellbio.duke.edu
http://note.cellbio.duke.edu/Faculty/Research/Reedy.html

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From: bfoster-at-mme1.com
Date: Thu, 17 May 2007 10:11:41 -0500
Subject: [Microscopy] Re: Bacterial flagella

Contents Retrieved from Microscopy Listserver Archives
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Hi, Scott

This really looks like a job for AFM. The sample prep would be a lot
easier (no staining) and the results would be more accurate.

Hope this is helpful,

Barbara Foster, President

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com


MME is now scheduling customized, on-site courses through July. Call
us today for details.

P. S.
Need a good general reference or light microscopy text for next
semester? Call us today to learn more about "Optimizing LIght
Microscopy". Copies still available through MME... even for
class-room lots ... and we give quantity discounts. Call Ken Piel at
(972)954-8011.


At 09:57 AM 5/17/2007, scott.coutts-at-med.monash.edu.au wrote:



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From: Mike.Bode-at-olympus-sis.com
Date: Thu, 17 May 2007 11:19:31 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Sharon,

This may be a bit more fundamental than what you are looking for, but
here goes:

If you go to the MSA website, you can find a statement about ethical
image processing that in essence says that images need to be stored
uncompressed. JPEG is a format that includes lossy compression, and I
believe that PICT can also contain lossy compression in the form of
JPEG. Both therefore don't seem to be the best choices.

I also found this on the web:

In the next version of its operating system, MacOS X, Apple has decided
to replace PICT by PDF. This means that the use of PICT for exchanging
data will slowly diminish.

It looks like sooner or later you will have to find a different choice
anyway. I would look at the whole chain of acquisition to see if you
cannot switch to something else.

Why can't you use anything else? Does it have to do with the original
file format that can only be read by iMovie? Or are you tied to the Mac
platform?

mike


Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
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Sent: Wednesday, May 16, 2007 19:39
To: Mike Bode

This Question/Comment was submitted to the Microscopy Listserver using
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Email: sbledsoe-at-iupui.edu
Name: Sharon Bledsoe

Title-Subject: [Filtered] Need Help From Imaging Experts

Question: Here is my problem: I can only use Mac iMovie to edit my
digital video. I can not do this on a PC. ONLY iMovie. To save a
single frame from the video, my choices are either PICT or JPEG format.


My Question is: To save an origianl image, which is the best format,
PICT or JPEG?

I'm not interested in hearing about Tiffs, BMPs or any other format as
that is not one of my choices.

------------------------------------------------------------------------
---

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From: bozhilov-at-ucr.edu
Date: Thu, 17 May 2007 11:35:53 -0500
Subject: [Microscopy] C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Looking for second condenser (C2) low-background 15, 30, 50 and 100
micron apertures for FEI CM300 TEM.

These are not the standard Pt thin foil discs.
I got a quote for a set for approx. $8000 from FEI. Any alternative
suppliers out there?

Thanks,

Krassimir.
_______________________________________
Krassimir N. Bozhilov
Central Facility for Advanced Microscopy and Microanalysis
University of California
Riverside, CA 92521

tel 951 827 2998
fax 951 827 2489
bozhilov-at-ucr.edu
_______________________________________



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From: john.mardinly-at-intel.com
Date: Thu, 17 May 2007 11:49:22 -0500
Subject: [Microscopy] C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Krassimir;
That sounds like typical prices for FIB cut apertures for field
emission TEMs, but is enough to make anyone say 'ouch!' I bought some
cheap ones from Pella (~$60 as I recall) a little smaller than what I
needed, and FIB cut them myself. If you have a FIB, it is pretty easy.

John Mardinly
Intel

This is not an opinion of Intel Corporation.

-----Original Message-----
X-from: bozhilov-at-ucr.edu [mailto:bozhilov-at-ucr.edu]
Sent: Thursday, May 17, 2007 9:36 AM
To: Mardinly, John

Looking for second condenser (C2) low-background 15, 30, 50 and 100
micron apertures for FEI CM300 TEM.

These are not the standard Pt thin foil discs.
I got a quote for a set for approx. $8000 from FEI. Any alternative
suppliers out there?

Thanks,

Krassimir.
_______________________________________
Krassimir N. Bozhilov
Central Facility for Advanced Microscopy and Microanalysis
University of California
Riverside, CA 92521

tel 951 827 2998
fax 951 827 2489
bozhilov-at-ucr.edu
_______________________________________



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16, 38 -- From john.mardinly-at-intel.com Thu May 17 11:49:21 2007
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From: bozhilov-at-ucr.edu
Date: Thu, 17 May 2007 12:14:51 -0500
Subject: [Microscopy] C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

These are not disk foils. They look like a small cylinder with 3 mm
diameter and about 5-6 mm height.

_______________________________________
Krassimir N. Bozhilov
Central Facility for Advanced Microscopy and Microanalysis
University of California
Riverside, CA 92521

tel 951 827 2998
fax 951 827 2489
bozhilov-at-ucr.edu
_______________________________________


On May 17, 2007, at 9:51 AM, john.mardinly-at-intel.com wrote:

}
}
}
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} America
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} MicroscopyListserver
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}
} Krassimir;
} That sounds like typical prices for FIB cut apertures for field
} emission TEMs, but is enough to make anyone say 'ouch!' I bought some
} cheap ones from Pella (~$60 as I recall) a little smaller than what I
} needed, and FIB cut them myself. If you have a FIB, it is pretty easy.
}
} John Mardinly
} Intel
}
} This is not an opinion of Intel Corporation.
}
} -----Original Message-----
} X-from: bozhilov-at-ucr.edu [mailto:bozhilov-at-ucr.edu]
} Sent: Thursday, May 17, 2007 9:36 AM
} To: Mardinly, John
} Subject: [Microscopy] C2 apertures
}
}
}
}
} ----------------------------------------------------------------------
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} http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------
} --
} ----
}
} Looking for second condenser (C2) low-background 15, 30, 50 and 100
} micron apertures for FEI CM300 TEM.
}
} These are not the standard Pt thin foil discs.
} I got a quote for a set for approx. $8000 from FEI. Any alternative
} suppliers out there?
}
} Thanks,
}
} Krassimir.
} _______________________________________
} Krassimir N. Bozhilov
} Central Facility for Advanced Microscopy and Microanalysis
} University of California
} Riverside, CA 92521
}
} tel 951 827 2998
} fax 951 827 2489
} bozhilov-at-ucr.edu
} _______________________________________
}
}
}
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From: jd-at-laddresearch.com
Date: Thu, 17 May 2007 12:15:36 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Since Ladd produces the vast majority of apertures in the U.S, we
surely can make them up for you. Please contact us and we'll discuss
materials, hole sizes, etc. To see samples of what we can do see our
website at:
http://www.laddresearch.com/Key_Products/Apertures/apertures.html

Disclaimer: Ladd has produced EM supplies and a wide variety of
apertures for over 50 years.

John Arnott

Ladd Research
83 Holly Court
Williston, VT 05495

On-line Catalog: www.laddresearch.com

Telephone: 1-802-658-4961 (anywhere)
Toll Free 1-800-451-3406 (US)
Fax: 1-802-660-8859

e-mail: sales-at-laddresearch.com

At 12:42 PM 5/17/2007, you wrote:



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==============================Original Headers==============================
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From: bigelow-at-umich.edu
Date: Thu, 17 May 2007 14:37:50 -0500
Subject: [Microscopy] [Microscopy}RE: O-ring

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

The Zatkoff Company (www.zatkoff.com) also carries an extensive stock
of O-rings. Before trying to make an O-ring yourself I would
recommend checking with them to see if they couldn't furnish
something to meet your needs.
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

==============================Original Headers==============================
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1, 14 -- Subject: [Microscopy}RE: O-ring
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From: Zhenquan.Liu-at-asu.edu
Date: Thu, 17 May 2007 15:58:56 -0500
Subject: [Microscopy] Short Switch for 120KV on 2010F TEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi, everyone!

One of our students wants to work on 2010F at 120 KV. At this moment the
beam is very weak if he follows a technical note from JEOL on our 2010F
TEM. He was told he can get bright beam if he has a device called Short
switch for 120KV. Local service engineer can install the short switch
easily. But JEOL USA told him that the device costs about $11,000.00 and
he has to wait for four months for delivery. But he needs it urgently.

Is there any one who has the device and our student may borrow or rent?
Or is there any idea to get bright beam at 120KV on 2010F?

Thanks a lot.

Zhenquan Liu


Zhenquan Liu
Senior Research Specialist
School of Materials
Arizona State University
901 S Palm Walk
PSA 213
Tempe, 85287-1704
Tel (480) 965 4512
Fax (480) 965 9004



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From: abansal-at-ilypsa.com
Date: Thu, 17 May 2007 19:30:22 -0500
Subject: [Microscopy] viaWWW: Sectioning polymer beads

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Krassimir;
5-6 mm height? I would suspect that the actual aperture must be
inserted in some kind of holder, because it would seem to be difficult
to form an aperture from solid material that thick. The Pella web site
shows cross-sections of Philips apertures that are 3 mm diameter, and
0.25 mm high, with a taper so that the thickness where the beam goes is
much less. However, I don't see anything 5-6 mm high in any catalogs, so
if the apertures are one piece, you might be stuck with the FEI deal.

John Mardinly
Intel

This is not an opinion of Intel Corp.

-----Original Message-----
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Sent: Thursday, May 17, 2007 10:08 AM
To: microscopy-at-microscopy.com
Cc: Mardinly, John

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Email: abansal-at-ilypsa.com
Name: Anu

Organization: Ilypsa

Title-Subject: [Filtered] Sectioning polymer beads for optical microscopy

Question: Does anyone have experience sectioning 100 to 200 micron sized polystryrene beads for optical microscopy? What are the best embedding media and microtomy conditions to get good sections?

---------------------------------------------------------------------------

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From: allan.mitchell-at-stonebow.otago.ac.nz
Date: Thu, 17 May 2007 19:53:36 -0500
Subject: [Microscopy] Cultured cells, thermanox coverslips and the PLT technique

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all

I am having some sectioning difficulties with cross sections of
cultured cells grown on Thermanox coverslips.

The monolayer of cells grown on the coverslips have been processed by
the progressive lowering of temperature technique (in an AFS), using
ethanol as the dehydration solvent, and then followed by embedding in
Lowicryl HM20.

The problem I am having is that the Thermanox coverslips are
separating from the Lowicryl HM20 resin block making cross-sectional
ultrathin sectioning (and semi-thin for that matter) that includes
the coverslip difficult.

The separation is easily seen through the binocular head of the
ultramicrotome even before I start trimming the block down. The
resin simply hasn't 'bonded' to the coverslip. (I am able to
remove the coverslip from the block and then get good ultrathin
sections of the cells embedded in the resin but we would prefer to
keep the coverslip in place if at all possible).

A parallel run of cells grown on Thermanox coverslips, processed at
room temperature and embedded in conventional epoxy resin are working
fine. I can easily get good ultrathin cross sections from these
which includes the coverslip and cells.

My question is:

Has anyone had success with ultrathin cross sections of Thermanox
coverslips and cells embedded in Lowicryl resins?

If so, any tricks or tips?


Many thanks, regards

Allan



Allan Mitchell
Technical Manager
Otago Centre for Electron Microscopy
Department of Anatomy and Structural Biology
School of Medical Sciences
P.O. Box 913
Dunedin
New Zealand

Phone (03) 479 5642 or 479 7301
Fax (03) 479 5086 or 479 7254



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From: colijn.1-at-osu.edu
Date: Thu, 17 May 2007 19:58:07 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Krassimir and John,

These sound like the "Top Hat" condenser apertures (since that is
what they look like in cross section). They have a ~250micron "brim"
which fits in the slots in the regular aperture holder rod and are
~3mm tall in the center. As with most apertures they have a tapered
hole with the small end at the top. The purpose is to reduce the
hard X-ray contribution (hole count) to the EDS spectrum. On FEG
instruments with an adjustable C1 aperture, you can get essentially
the same effect by using the C1 aperture as your beam limiter and
using a C2 aperture slightly larger than the beam to stop the brehmstrahlung.

$8000 sounds ridiculously high for an aperture, even Platinum Top
Hats. I got a package from FEI (p/n 9432 061 67001 for the
70micron) for ~$800. I don't remember offhand the number of pieces
per package.

Henk

At 07:05 PM 5/17/2007, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Hendrik O. Colijn colijn.1-at-osu.edu
OSU Campus Electron Optics Facility (614) 292-0674
040 Fontana Labs, 116 W. 19th Ave www.ceof.ohio-state.edu


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From: opmills-at-mtu.edu
Date: Thu, 17 May 2007 20:38:56 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Contact John Bishop -at- Norsam Technologies. http://www.norsam.com/ I
talked to him in 2006 about a custom aperture strip for our Hitachi FIB.
He may be able to make something for you but I don't know that it would be
any cheaper.

Owen Mills
Michigan Tech Univ.




}
}
}
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}
} Krassimir;
} 5-6 mm height? I would suspect that the actual aperture must be
} inserted in some kind of holder, because it would seem to be difficult
} to form an aperture from solid material that thick. The Pella web site
} shows cross-sections of Philips apertures that are 3 mm diameter, and
} 0.25 mm high, with a taper so that the thickness where the beam goes is
} much less. However, I don't see anything 5-6 mm high in any catalogs, so
} if the apertures are one piece, you might be stuck with the FEI deal.
}
} John Mardinly
} Intel
}
} This is not an opinion of Intel Corp.
}
} -----Original Message-----
} X-from: KN Bozhilov [mailto:bozhilov-at-ucr.edu]
} Sent: Thursday, May 17, 2007 10:08 AM
} To: microscopy-at-microscopy.com
} Cc: Mardinly, John
} Subject: Re: [Microscopy] RE: C2 apertures
}
} These are not disk foils. They look like a small cylinder with 3 mm
} diameter and about 5-6 mm height.
}
} _______________________________________
} Krassimir N. Bozhilov
} Central Facility for Advanced Microscopy and Microanalysis
} University of California
} Riverside, CA 92521
}
} tel 951 827 2998
} fax 951 827 2489
} bozhilov-at-ucr.edu
} _______________________________________
}
}
} On May 17, 2007, at 9:51 AM, john.mardinly-at-intel.com wrote:
}
} }
} }
} }
} } ----------------------------------------------------------------------
}
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} }
} } Krassimir;
} } That sounds like typical prices for FIB cut apertures for field
} } emission TEMs, but is enough to make anyone say 'ouch!' I bought some
} } cheap ones from Pella (~$60 as I recall) a little smaller than what I
} } needed, and FIB cut them myself. If you have a FIB, it is pretty easy.
} }
} } John Mardinly
} } Intel
} }
} } This is not an opinion of Intel Corporation.
} }
} } -----Original Message-----
} } X-from: bozhilov-at-ucr.edu [mailto:bozhilov-at-ucr.edu]
} } Sent: Thursday, May 17, 2007 9:36 AM
} } To%3


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From: walck-at-southbaytech.com
Date: Thu, 17 May 2007 22:45:14 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I agree with Henk here, that it is probably a "Top Hat" aperture. But these
are typically about 1 mm in height (and hard to get a hold of for a JEOL
2000FX). My question to the group is, isn't there a hard, fixed aperture
below the C2 aperture that is in the column and thicker like the one he is
descibing?


-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: colijn.1-at-osu.edu [mailto:colijn.1-at-osu.edu]
Sent: Thursday, May 17, 2007 6:01 PM
To: Walck-at-SouthBayTech.com

Hi Krassimir and John,

These sound like the "Top Hat" condenser apertures (since that is what they
look like in cross section). They have a ~250micron "brim"
which fits in the slots in the regular aperture holder rod and are ~3mm tall
in the center. As with most apertures they have a tapered hole with the
small end at the top. The purpose is to reduce the hard X-ray contribution
(hole count) to the EDS spectrum. On FEG instruments with an adjustable C1
aperture, you can get essentially the same effect by using the C1 aperture
as your beam limiter and using a C2 aperture slightly larger than the beam
to stop the brehmstrahlung.

$8000 sounds ridiculously high for an aperture, even Platinum Top Hats. I
got a package from FEI (p/n 9432 061 67001 for the
70micron) for ~$800. I don't remember offhand the number of pieces per
package.

Henk

At 07:05 PM 5/17/2007, you wrote:



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Hendrik O. Colijn colijn.1-at-osu.edu
OSU Campus Electron Optics Facility (614) 292-0674
040 Fontana Labs, 116 W. 19th Ave www.ceof.ohio-state.edu


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From: colijn.1-at-osu.edu
Date: Fri, 18 May 2007 07:19:07 -0500
Subject: [Microscopy] Re: C2 apertures

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Hi Scott,

In some of the older JEOL scopes (e.g. 200CX, 2000EX??) with a
conventional (non C-O) objective lens you could get an optional HAX
kit. This fit in the large upper bore of the pole-piece to reduce
the brehmstrahlung contribution. In a scope with a
condenser-objective, there is no room to put an additional
absorber. There are no apertures between the C2 and the sample
unless you want to call the upper bore of the objective an
"aperture". {...grin...}

Krassimir was looking for an aperture for a CM300. There aren't any
apertures between C2 and the sample.

Cheers,
Henk

At 11:47 PM 05/17/07, you wrote:



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Hendrik O. Colijn colijn.1-at-osu.edu
Campus Electron Optics Facility Ohio State University
(614) 292-0674 http://www.ceof.ohio-state.edu
Time is that quality of nature which keeps events from happening all
at once. Lately it doesn't seem to be working.


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From: dljones-at-bestweb.net
Date: Fri, 18 May 2007 07:55:47 -0500
Subject: [Microscopy] Re: [Microscopy}RE: O-ring

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dr. Bigelow,

With all due respect, making an O-ring is not a particularly difficult
adventure. In fact, the very reference you are citing below has a kit
Zatkoff sells to make your own custom O-rings.

https://www.zatkoff.com/catalog/download_pdf.asp?file=1-252_splicing_kits.pdf

(I don't know if one has to register with Zatkoff to see that link, but
the registration is simple and quick if one is interested in looking into
this.)

I certainly agree that it is better to just buy the proper O-ring, but
sometimes the size you need is just not available easily without having to
buy far more than you could need in the foreseeable future. I have had
numerous occassions where just making the specific sized O-ring needed is
just the simplest, most direct way to go.

dj

On Thu, 17 May 2007, bigelow-at-umich.edu wrote:

}
}
}
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} The Zatkoff Company (www.zatkoff.com) also carries an extensive stock
} of O-rings. Before trying to make an O-ring yourself I would
} recommend checking with them to see if they couldn't furnish
} something to meet your needs.
} --
} Wilbur C. Bigelow, Professor Emeritus
} Materials Sci. & Engr., Univ. of Michigan
} Ann Arbor, Michigan 48109-2136
} e-mail: bigelow-at-umich.edu;
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From: pforcen-at-unizar.es
Date: Fri, 18 May 2007 08:27:05 -0500
Subject: [Microscopy] viaWWW: TEM and RuO4

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Email: pforcen-at-unizar.es
Name: Patricia

Organization: University of Zaragoza

Title-Subject: [Filtered] TEM and RuO4

Question: Dear Microscopist,
my question is about TEM and the staining with RuO4.
I am using this method to enhance the contrast of my sample and it is working properly, but I am trying to find some bibliography or some explanation about the mechanism, it is not very well described in any of the papers and book I could have a look.
I know it is an oxydation proccess but how it is attaching to the molecule and more important to me, how it is modificating the size!!

Thank you for your help in advance!
Kind Regards,
patricia

---------------------------------------------------------------------------

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From: thatch-at-mit.edu
Date: Fri, 18 May 2007 08:27:37 -0500
Subject: [Microscopy] viaWWW: automatic grid stainer

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Email: thatch-at-mit.edu
Name: Tori Hatch

Organization: HHMI

Title-Subject: [Filtered] automatic grid stainer

Question: I need to replace my Leica EM stainer. Leica is coming out with a new machine soon, but I'd like to hear from anyone who has experience with other automatic grid stainers.

Thanks,
Tori Hatch
HHMI at MIT


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From: Mike.Bode-at-olympus-sis.com
Date: Fri, 18 May 2007 09:59:36 -0500
Subject: [Microscopy] RE: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
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In that case I would suggest to ask Apple. Contact them and tell them
that for reasons of scientific Ethics you need the images in TIFF format
(or any other format that you think might work), and ask them if they
have a converter for their iMovie Product. They claim to be super
user-friendly. Put them to the test.

mike

Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
X-from: Sharon Bledsoe [mailto:sbledsoe-at-iupui.edu]
Sent: Friday, May 18, 2007 06:12
To: Mike Bode

} to replace PICT by PDF. This means that the use of PICT for exchanging
} data will slowly diminish.
}
} It looks like sooner or later you will have to find a different choice
} anyway. I would look at the whole chain of acquisition to see if you
} cannot switch to something else.
}
} Why can't you use anything else? Does it have to do with the original
} file format that can only be read by iMovie? Or are you tied to the Mac

} platform?
}
} mike
}
}
} Michael Bode, Ph.D.
}
} General Manager
}
} OLYMPUS SOFT IMAGING SOLUTIONS
} 12596 West Bayaud Ave #300
} Lakewood, CO 80228
} USA
} Tel.: +1 (303) 234-9270
} Fax.: +1 (303) 234-9271
} E-mail: Mike.Bode-at-olympus-sis.com
} www.olympus-sis.com
}
} -----Original Message-----
} X-from: sbledsoe-at-iupui.edu [mailto:sbledsoe-at-iupui.edu]
} Sent: Wednesday, May 16, 2007 19:39
} To: Mike Bode
} Subject: [Microscopy] viaWWW: Need Help From Imaging Experts
}
}
}
}
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From: gary.m.brown-at-exxonmobil.com
Date: Fri, 18 May 2007 10:12:56 -0500
Subject: [Microscopy] Re: viaWWW: TEM and RuO4

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Patricia,

The best reference I know is Trent, J.S. et al, Macromolecules, 1983, 16,
589. Trent deals with a variety of issues including specificity of
staining by RuO4 and OsO4 and the chemistry of the reactions.

I suggest you also look in the Polymer Microscopy, by Sawyer and Grubbs,
2nd ed for other references.

Regards.

Disclaimer: The comments and opinions are those of the author alone and do
not reflect any official position by Exxon Mobil Chemical Company.

Gary M. Brown
Microscopy Specialist
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

"Happiness equals reality minus expectations." Tom Magliozzi





pforcen-at-unizar
.es
To
gary.m.brown-at-exxonmobil.com
05/18/07 08:29 cc
AM
Subject
[Microscopy] viaWWW: TEM and RuO4
Please respond
to
pforcen-at-unizar
.es









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Email: pforcen-at-unizar.es
Name: Patricia

Organization: University of Zaragoza

Title-Subject: [Filtered] TEM and RuO4

Question: Dear Microscopist,
my question is about TEM and the staining with RuO4.
I am using this method to enhance the contrast of my sample and it is
working properly, but I am trying to find some bibliography or some
explanation about the mechanism, it is not very well described in any of
the papers and book I could have a look.
I know it is an oxydation proccess but how it is attaching to the molecule
and more important to me, how it is modificating the size!!

Thank you for your help in advance!
Kind Regards,
patricia

---------------------------------------------------------------------------

==============================Original
Headers==============================
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==============================Original Headers==============================
30, 19 -- From gary.m.brown-at-exxonmobil.com Fri May 18 10:12:55 2007
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From: lamiller-at-uiuc.edu
Date: Fri, 18 May 2007 10:17:27 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I see what he means , just tried iMovie on OS X 10.3.


It seems to me though, that about 5 years ago, it DID let me save to
TIFF and there were more options, perhaps trying to get an Old copy of
iMovie? or try on an old machine if you still have OS 9?

Also can you open the clip in Quicktime Pro? Maybe Quicktime Pro will
let you pull a tiff off of it, it too will save a frame.

Not a great solution, but if you are in a pinch.....



Lou Ann

Mike.Bode-at-olympus-sis.com wrote:
} ----------------------------------------------------------------------------
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}
} In that case I would suggest to ask Apple. Contact them and tell them
} that for reasons of scientific Ethics you need the images in TIFF format
} (or any other format that you think might work), and ask them if they
} have a converter for their iMovie Product. They claim to be super
} user-friendly. Put them to the test.
}
} mike
}
} Michael Bode, Ph.D.
}
} General Manager
}
} OLYMPUS SOFT IMAGING SOLUTIONS
} 12596 West Bayaud Ave #300
} Lakewood, CO 80228
} USA
} Tel.: +1 (303) 234-9270
} Fax.: +1 (303) 234-9271
} E-mail: Mike.Bode-at-olympus-sis.com
} www.olympus-sis.com
}
} -----Original Message-----
} X-from: Sharon Bledsoe [mailto:sbledsoe-at-iupui.edu]
} Sent: Friday, May 18, 2007 06:12
} To: Mike Bode
} Subject: [Microscopy] RE: viaWWW: Need Help From Imaging Experts
}
} IF I had a choice I would choose to save my images as TIFF's, however,
} the iMovie program will not let me. iMoive will only let me save as
} PICT or JPEG.
}
} And I am not tied to the Mac, I'm chained to it.
}
}
}
}
}
} } -----------------------------------------------------------------------
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} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } -----------------------------------------------------------------------
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} }
} } Hi Sharon,
} }
} } This may be a bit more fundamental than what you are looking for, but
} } here goes:
} }
} } If you go to the MSA website, you can find a statement about ethical
} } image processing that in essence says that images need to be stored
} } uncompressed. JPEG is a format that includes lossy compression, and I
} } believe that PICT can also contain lossy compression in the form of
} } JPEG. Both therefore don't seem to be the best choices.
} }
} } I also found this on the web:
} }
} } In the next version of its operating system, MacOS X, Apple has decided
} }
}
}
} } to replace PICT by PDF. This means that the use of PICT for exchanging
} } data will slowly diminish.
} }
} } It looks like sooner or later you will have to find a different choice
} } anyway. I would look at the whole chain of acquisition to see if you
} } cannot switch to something else.
} }
} } Why can't you use anything else? Does it have to do with the original
} } file format that can only be read by iMovie? Or are you tied to the Mac
} }
}
}
} } platform?
} }
} } mike
} }
} }
} } Michael Bode, Ph.D.
} }
} } General Manager
} }
} } OLYMPUS SOFT IMAGING SOLUTIONS
} } 12596 West Bayaud Ave #300
} } Lakewood, CO 80228
} } USA
} } Tel.: +1 (303) 234-9270
} } Fax.: +1 (303) 234-9271
} } E-mail: Mike.Bode-at-olympus-sis.com
} } www.olympus-sis.com
} }
} } -----Original Message-----
} } X-from: sbledsoe-at-iupui.edu [mailto:sbledsoe-at-iupui.edu]
} } Sent: Wednesday, May 16, 2007 19:39
} } To: Mike Bode
} } Subject: [Microscopy] viaWWW: Need Help From Imaging Experts
} }
} }
} }
} }
} } -----------------------------------------------------------------------
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} } ---
} }
} } Email: sbledsoe-at-iupui.edu
} } Name: Sharon Bledsoe
} }
} } Title-Subject: [Filtered] Need Help From Imaging Experts
} }
} } Question: Here is my problem: I can only use Mac iMovie to edit my
} } digital video. I can not do this on a PC. ONLY iMovie. To save a
} } single frame from the video, my choices are either PICT or JPEG format.
} }
} }
} } My Question is: To save an origianl image, which is the best format,
} } PICT or JPEG?
} }
} } I'm not interested in hearing about Tiffs, BMPs or any other format as
} } that is not one of my choices.
} }
} } -----------------------------------------------------------------------
} } -
} } ---
} }
} } ==============================Original
} } Headers==============================
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} } ==============================End of -
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} }
} } ==============================Original
} } Headers==============================
} } 26, 25 -- From Mike.Bode-at-olympus-sis.com Thu May 17 11:19:31 2007 26,
} } 25 -- Received: from mail.soft-imaging.de (ns.soft-imaging.de
} } [62.180.61.130])
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} ==============================Original Headers==============================
} 16, 25 -- From Mike.Bode-at-olympus-sis.com Fri May 18 09:59:35 2007
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} 16, 25 -- References: {200705171631.l4HGVqWh015155-at-ns.microscopy.com} {a06240800c27347fa66ce-at-[134.68.251.140]}
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}
}

--
~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Lou Ann Miller, MT(ASCP)
Service Supervisor
Center for Microscopic Imaging
College of Veterinary Medicine
Rm 1204 VMBSB
2001 S Lincoln Ave
Urbana, IL 61802

217/244-1567
http://treefrog.cvm.uiuc.edu


==============================Original Headers==============================
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From: gmartens-at-interchange.ubc.ca
Date: Fri, 18 May 2007 10:21:47 -0500
Subject: [Microscopy] Re: Cultured cells,

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Allan,

We have had the same problem in that only the epoxy resins will bind
to the thermanox, and aclar for that matter. Are you doing PLT and
HM20 embedding for immunolabeling, or just to avoid room temp
dehydration? I know some people who have had success immunolabeling
after etching the epoxy resin away (just an idea). We have had the
most success growing cells on a Transwell Plate (sold through Corning
here in Canada) that has a porous polyester membrane on the bottom.
The HM20 resin is able to penetrate through the pores and will hold
everything together during sectioning allowing for decent thin
sectioning. Expect some wrinkles and getting a perfect flat cross
section is difficult because the membrane is pretty flexible, but it
works. Why is it so important to have the coverslip present?

Hope this helps,

Garnet


} Hi all
}
} I am having some sectioning difficulties with cross sections of
} cultured cells grown on Thermanox coverslips.
}
} The monolayer of cells grown on the coverslips have been processed by
} the progressive lowering of temperature technique (in an AFS), using
} ethanol as the dehydration solvent, and then followed by embedding in
} Lowicryl HM20.
}
} The problem I am having is that the Thermanox coverslips are
} separating from the Lowicryl HM20 resin block making cross-sectional
} ultrathin sectioning (and semi-thin for that matter) that includes
} the coverslip difficult.
}
} The separation is easily seen through the binocular head of the
} ultramicrotome even before I start trimming the block down. The
} resin simply hasn't 'bonded' to the coverslip. (I am able to
} remove the coverslip from the block and then get good ultrathin
} sections of the cells embedded in the resin but we would prefer to
} keep the coverslip in place if at all possible).
}
} A parallel run of cells grown on Thermanox coverslips, processed at
} room temperature and embedded in conventional epoxy resin are working
} fine. I can easily get good ultrathin cross sections from these
} which includes the coverslip and cells.
}
} My question is:
}
} Has anyone had success with ultrathin cross sections of Thermanox
} coverslips and cells embedded in Lowicryl resins?
}
} If so, any tricks or tips?
}
}
} Many thanks, regards
}
} Allan
}
}
}
} Allan Mitchell
} Technical Manager
} Otago Centre for Electron Microscopy
} Department of Anatomy and Structural Biology
} School of Medical Sciences
} P.O. Box 913
} Dunedin
} New Zealand
}
} Phone (03) 479 5642 or 479 7301
} Fax (03) 479 5086 or 479 7254
}
}
}
} ==============================Original Headers==============================
} 18, 20 -- From allan.mitchell-at-stonebow.otago.ac.nz Thu May 17 19:53:36 2007
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} 18, 20 -- To: microscopy-at-msa.microscopy.com
} 18, 20 -- From: Allan Mitchell {allan.mitchell-at-stonebow.otago.ac.nz}
} 18, 20 -- Subject: Cultured cells, thermanox coverslips and the PLT technique
} 18, 20 -- Date: Fri, 18 May 2007 12:40:58 +1200
} 18, 20 -- X-Mailer: Apple Mail (2.752.2)
} ==============================End of - Headers==============================


--
Garnet Martens

Research Manager
BioImaging Facility
University of British Columbia
6270 University Blvd.
Vancouver, B.C.
Canada
V6T 1Z4

phone 604-822-3354

==============================Original Headers==============================
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10, 31 -- Date: Fri, 18 May 2007 08:21:42 -0700
10, 31 -- From: Garnet Martens {gmartens-at-interchange.ubc.ca}
10, 31 -- Subject: Re: [Microscopy] Cultured cells,
10, 31 -- thermanox coverslips and the PLT technique
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From: trazy-at-umn.edu
Date: Fri, 18 May 2007 10:26:14 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


The PICT files captured through iMovie are lossless, so this would be
the flavor of choice. You can then easily convert PICT to TIFF if
you wish.

Cheers,

--
Tracy E. Anderson
Microscopist and digital imaging specialist
Imaging Center
University of Minnesota
College of Biological Sciences
Phone: 612.624.3454
Fax: 612.624.1799
http://www.cbs.umn.edu/ic/
--
"Science and art belong to the whole world, and before them vanish
the barriers of nationality." - Goethe




On 18 May 2007, Mike.Bode-at-olympus-sis.com wrote:
}
}
}
}
----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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}
}
} In that case I would suggest to ask Apple. Contact them and tell them
} that for reasons of scientific Ethics you need the images in TIFF format
} (or any other format that you think might work), and ask them if they
} have a converter for their iMovie Product. They claim to be super
} user-friendly. Put them to the test.
}
} mike
}
} Michael Bode, Ph.D.
}
} General Manager
}
} OLYMPUS SOFT IMAGING SOLUTIONS
} 12596 West Bayaud Ave #300
} Lakewood, CO 80228
} USA
} Tel.: +1 (303) 234-9270
} Fax.: +1 (303) 234-9271
} E-mail: Mike.Bode-at-olympus-sis.com
} www.olympus-sis.com
}
} -----Original Message-----
} X-from: Sharon Bledsoe [mailto:sbledsoe-at-iupui.edu]
} Sent: Friday, May 18, 2007 06:12
} To: Mike Bode
} Subject: [Microscopy] RE: viaWWW: Need Help From Imaging Experts
}
} IF I had a choice I would choose to save my images as TIFF's, however,
} the iMovie program will not let me. iMoive will only let me save as
} PICT or JPEG.
}
} And I am not tied to the Mac, I'm chained to it.
}
}
}
}
} } -----------------------------------------------------------------------
} } ----- The Microscopy ListServer -- CoSponsor: The Microscopy Society
} } of America To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } -----------------------------------------------------------------------
} } -----
} }
} } Hi Sharon,
} }
} } This may be a bit more fundamental than what you are looking for, but
} } here goes:
} }
} } If you go to the MSA website, you can find a statement about ethical
} } image processing that in essence says that images need to be stored
} } uncompressed. JPEG is a format that includes lossy compression, and I
} } believe that PICT can also contain lossy compression in the form of
} } JPEG. Both therefore don't seem to be the best choices.
} }
} } I also found this on the web:
} }
} } In the next version of its operating system, MacOS X, Apple has decided
}
} } to replace PICT by PDF. This means that the use of PICT for exchanging
} } data will slowly diminish.
} }
} } It looks like sooner or later you will have to find a different choice
} } anyway. I would look at the whole chain of acquisition to see if you
} } cannot switch to something else.
} }
} } Why can't you use anything else? Does it have to do with the original
} } file format that can only be read by iMovie? Or are you tied to the Mac
}
} } platform?
} }
} } mike
} }
} }
} } Michael Bode, Ph.D.
} }
} } General Manager
} }
} } OLYMPUS SOFT IMAGING SOLUTIONS
} } 12596 West Bayaud Ave #300
} } Lakewood, CO 80228
} } USA
} } Tel.: +1 (303) 234-9270
} } Fax.: +1 (303) 234-9271
} } E-mail: Mike.Bode-at-olympus-sis.com
} } www.olympus-sis.com
} }
} } -----Original Message-----
} } X-from: sbledsoe-at-iupui.edu [mailto:sbledsoe-at-iupui.edu]
} } Sent: Wednesday, May 16, 2007 19:39
} } To: Mike Bode
} } Subject: [Microscopy] viaWWW: Need Help From Imaging Experts
} }
} }
} }
} }
} } -----------------------------------------------------------------------
} } -
} } ----
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } -----------------------------------------------------------------------
} } -
} } ----
} }
} } This Question/Comment was submitted to the Microscopy Listserver using
} } the WWW based Form at
} } http://microscopy.com/MicroscopyListserver/MLFormMail.html
} } -----------------------------------------------------------------------
} } -
} } ---
} } Remember this posting is most likely not from a Subscriber, so when
} } replying
} } please copy both sbledsoe-at-iupui.edu as well as the MIcroscopy
} } Listserver
} } -----------------------------------------------------------------------
} } -
} } ---
} }
} } Email: sbledsoe-at-iupui.edu
} } Name: Sharon Bledsoe
} }
} } Title-Subject: [Filtered] Need Help From Imaging Experts
} }
} } Question: Here is my problem: I can only use Mac iMovie to edit my
} } digital video. I can not do this on a PC. ONLY iMovie. To save a
} } single frame from the video, my choices are either PICT or JPEG format.
} }
} }
} } My Question is: To save an origianl image, which is the best format,
} } PICT or JPEG?
} }
} } I'm not interested in hearing about Tiffs, BMPs or any other format as
} } that is not one of my choices.
} }
} } -----------------------------------------------------------------------
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} } ==============================Original
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From: bozhilov-at-ucr.edu
Date: Fri, 18 May 2007 10:31:41 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Scott,

These are straightforward C2 apertures and sit in the C2 aperture
holder rod. They do have the function of the "top hat" apertures but
they do not have the "top hat" shape. It is a simple cylinder with
3mm diameter and (after I measured again) 4 mm height. I do not know
what is the material it is made of. I am surprised that these
apertures turn out to be so esotheric objects, after all CM300 is not
that of a surreptitious instrument.

Krassimir.
_______________________________________
Krassimir N. Bozhilov
Central Facility for Advanced Microscopy and Microanalysis
University of California
Riverside, CA 92521

tel 951 827 2998
fax 951 827 2489
bozhilov-at-ucr.edu
_______________________________________


On May 17, 2007, at 8:47 PM, walck-at-southbaytech.com wrote:

}
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} I agree with Henk here, that it is probably a "Top Hat" aperture.
} But these
} are typically about 1 mm in height (and hard to get a hold of for a
} JEOL
} 2000FX). My question to the group is, isn't there a hard, fixed
} aperture
} below the C2 aperture that is in the column and thicker like the
} one he is
} descibing?
}
}
} -Scott
}
} Scott D. Walck, Ph.D.
} Technical Director
} South Bay Technology, Inc.
} 1120 Via Callejon
} San Clemente, CA 92673
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} walck-at-southbaytech.com
}
} -----Original Message-----
} X-from: colijn.1-at-osu.edu [mailto:colijn.1-at-osu.edu]
} Sent: Thursday, May 17, 2007 6:01 PM
} To: Walck-at-SouthBayTech.com
} Subject: [Microscopy] Re: C2 apertures
}
}
}
}
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} Hi Krassimir and John,
}
} These sound like the "Top Hat" condenser apertures (since that is
} what they
} look like in cross section). They have a ~250micron "brim"
} which fits in the slots in the regular aperture holder rod and are
} ~3mm tall
} in the center. As with most apertures they have a tapered hole
} with the
} small end at the top. The purpose is to reduce the hard X-ray
} contribution
} (hole count) to the EDS spectrum. On FEG instruments with an
} adjustable C1
} aperture, you can get essentially the same effect by using the C1
} aperture
} as your beam limiter and using a C2 aperture slightly larger than
} the beam
} to stop the brehmstrahlung.
}
} $8000 sounds ridiculously high for an aperture, even Platinum Top
} Hats. I
} got a package from FEI (p/n 9432 061 67001 for the
} 70micron) for ~$800. I don't remember offhand the number of
} pieces per
} package.
}
} Henk
}
} At 07:05 PM 5/17/2007, you wrote:
}
}
}
} } ---------------------------------------------------------------------
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} }
} } Krassimir;
} } 5-6 mm height? I would suspect that the actual aperture must
} } be inserted in some kind of holder, because it would seem to be
} } difficult to form an aperture from solid material that thick. The
} } Pella
} } web site shows cross-sections of Philips apertures that are 3 mm
} } diameter, and
} } 0.25 mm high, with a taper so that the thickness where the beam
} } goes is
} } much less. However, I don't see anything 5-6 mm high in any catalogs,
} } so if the apertures are one piece, you might be stuck with the FEI
} } deal.
} }
} } John Mardinly
} } Intel
} }
} } This is not an opinion of Intel Corp.
} }
} } -----Original Message-----
} } X-from: KN Bozhilov [mailto:bozhilov-at-ucr.edu]
} } Sent: Thursday, May 17, 2007 10:08 AM
} } To: microscopy-at-microscopy.com
} } Cc: Mardinly, John
} } Subject: Re: [Microscopy] RE: C2 apertures
} }
} } These are not disk foils. They look like a small cylinder with 3 mm
} } diameter and about 5-6 mm height.
} }
} } _______________________________________
} } Krassimir N. Bozhilov
} } Central Facility for Advanced Microscopy and Microanalysis University
} } of California Riverside, CA 92521
} }
} } tel 951 827 2998
} } fax 951 827 2489
} } bozhilov-at-ucr.edu
} } _______________________________________
} }
} }
} } On May 17, 2007, at 9:51 AM, john.mardinly-at-intel.com wrote:
} }
} } }
} } }
} } }
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} } }
} } } Krassimir;
} } } That sounds like typical prices for FIB cut apertures for
} } } field emission TEMs, but is enough to make anyone say 'ouch!' I
} } } bought some cheap ones from Pella (~$60 as I recall) a little
} } } smaller than what I needed, and FIB cut them myself. If you have
} } } a FIB,
} it is pretty easy.
} } }
} } } John Mardinly
} } } Intel
} } }
} } } This is not an opinion of Intel Corporation.
} } }
} } } -----Original Message-----
} } } X-from: bozhilov-at-ucr.edu [mailto:bozhilov-at-ucr.edu]
} } } Sent: Thursday, May 17, 2007 9:36 AM
} } } To: Mardinly, John
} } } Subject: [Microscopy] C2 apertures
} } }
} } }
} } }
} } }
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} } }
} } } Looking for second condenser (C2) low-background 15, 30, 50 and 100
} } } micron apertures for FEI CM300 TEM.
} } }
} } } These are not the standard Pt thin foil discs.
} } } I got a quote for a set for approx. $8000 from FEI. Any alternative
} } } suppliers out there?
} } }
} } } Thanks,
} } }
} } } Krassimir.
} } } _______________________________________
} } } Krassimir N. Bozhilov
} } } Central Facility for Advanced Microscopy and Microanalysis
} } } University of California Riverside, CA 92521
} } }
} } } tel 951 827 2998
} } } fax 951 827 2489
} } } bozhilov-at-ucr.edu
} } } _______________________________________
} } }
} } }
} } }
} } } ==============================Original
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From: lamiller-at-uiuc.edu
Date: Fri, 18 May 2007 10:45:23 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

In that case, PICT would indeed be the format of choice. I found some
information on the web that said the PICT format CAN include JPEG
compression. That may be a user defined option, then.

Do you know if the iMovie format that is used before extracting he
inividual frames uses mPEG compression?

mike


Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
X-from: tracy [mailto:trazy-at-umn.edu]
Sent: Friday, May 18, 2007 09:26
To: Mike Bode; microscopy-at-microscopy.com

OK...

Now I'm a little confused.........

Because I was taught that a Pic file, like a screen shot is only a
rasterized 72 dpi image.


So I threw a 6400 dpi image into iMovie, and saved as a pic file, and
then opened in photoshop, and it says it's a 640 by 480 .............72
dpi image


Was the conversion on the import into iMovie, or is this what I've
always found with pic files, 72 dpi cause that's what the Mac does with
a pic file?

So I save as JPEG, and it's a 243x144 pixel image and still only 72 dpi.

If I use the OS X grab-it function for a screen shot, it actually saves
it in TIFF, but again, only 72 bit dpi.

In which case it wouldn't make any difference just to use the System
application "Grab-it", select to capture selection, and save using
Grab-it, as both would be 72 dpi, and there would be no more conversion
involved



Lou Ann


trazy-at-umn.edu wrote:
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}
} The PICT files captured through iMovie are lossless, so this would be
} the flavor of choice. You can then easily convert PICT to TIFF if
} you wish.
}
} Cheers,
}
} --
} Tracy E. Anderson
}
}


==============================Original Headers==============================
16, 19 -- From lamiller-at-uiuc.edu Fri May 18 10:45:23 2007
16, 19 -- Received: from expredir5.cites.uiuc.edu (expredir5.cites.uiuc.edu [128.174.5.96])
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16, 19 -- Date: Fri, 18 May 2007 10:45:22 -0500
16, 19 -- From: Lou Ann Miller {lamiller-at-uiuc.edu}
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From: trazy-at-umn.edu
Date: Fri, 18 May 2007 11:04:23 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

iMovie deals with NTSC (TV) resolution (unless you're working in HDTV), which
is 640x480 at 72dpi. So yes, any large image file brought into iMovie will
be scaled down accordingly.

Presumably, the original poster is working with a video captured with NTSC
resolution, so there would be no loss when capturing individual frames.

Cheers,

-
Tracy E. Anderson
Microscopist and digital imaging specialist
Imaging Center
University of Minnesota
College of Biological Sciences
Phone: 612.624.3454
Fax: 612.624.1799
http://www.cbs.umn.edu/ic/
--
"Science and art belong to the whole world, and before them vanish the
barriers of nationality." - Goethe
--


On 18 May 2007, lamiller-at-uiuc.edu wrote:
}
}
}
}
----------------------------------------------------------------------------
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} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
}
----------------------------------------------------------------------------
}
} OK...
}
} Now I'm a little confused.........
}
} Because I was taught that a Pic file, like a screen shot is only a
} rasterized 72 dpi image.
}
}
} So I threw a 6400 dpi image into iMovie, and saved as a pic file, and
} then opened in photoshop, and it says it's a 640 by 480 .............72
} dpi image
}
}
} Was the conversion on the import into iMovie, or is this what I've
} always found with pic files, 72 dpi cause that's what the Mac does with
} a pic file?
}
} So I save as JPEG, and it's a 243x144 pixel image and still only 72 dpi.
}
} If I use the OS X grab-it function for a screen shot, it actually saves
} it in TIFF, but again, only 72 bit dpi.
}
} In which case it wouldn't make any difference just to use the System
} application "Grab-it", select to capture selection, and save using
} Grab-it, as both would be 72 dpi, and there would be no more conversion
} involved
}
}
}
} Lou Ann
}
}
} trazy-at-umn.edu wrote:
} }
}
----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
America
} } To Subscribe/Unsubscribe --
http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} }
}
----------------------------------------------------------------------------
} }
} }
} } The PICT files captured through iMovie are lossless, so this would be
} } the flavor of choice. You can then easily convert PICT to TIFF if
} } you wish.
} }
} } Cheers,
} }
} } --
} } Tracy E. Anderson
} }
} }
}
}
} ==============================Original
Headers==============================
} 16, 19 -- From lamiller-at-uiuc.edu Fri May 18 10:45:23 2007
} 16, 19 -- Received: from expredir5.cites.uiuc.edu (expredir5.cites.uiuc.edu
} [128.174.5.96])
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} l4IFjMCi007315
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} (CDT)
} 16, 19 -- Message-ID: {464DCA12.9010409-at-uiuc.edu}
} 16, 19 -- Date: Fri, 18 May 2007 10:45:22 -0500
} 16, 19 -- From: Lou Ann Miller {lamiller-at-uiuc.edu}
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} 16, 19 -- User-Agent: Thunderbird 2.0.0.0 (Macintosh/20070326)
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} Experts
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}



==============================Original Headers==============================
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From: lamiller-at-uiuc.edu
Date: Fri, 18 May 2007 11:18:19 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Oh,

In that case, Grab-it can save directly to TIFF but would it be the
same or worse than pict as a screen shot?


Lou Ann



trazy-at-umn.edu wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} iMovie deals with NTSC (TV) resolution (unless you're working in HDTV), which
} is 640x480 at 72dpi. So yes, any large image file brought into iMovie will
} be scaled down accordingly.
}
} Presumably, the original poster is working with a video captured with NTSC
} resolution, so there would be no loss when capturing individual frames.
}
} Cheers,
}
} -
} Tracy E. Anderson
} Microscopist and digital imaging specialist
} Imaging Center
} University of Minnesota
} College of Biological Sciences
} Phone: 612.624.3454
} Fax: 612.624.1799
} http://www.cbs.umn.edu/ic/
} --
} "Science and art belong to the whole world, and before them vanish the
} barriers of nationality." - Goethe
} --
}
}
} On 18 May 2007, lamiller-at-uiuc.edu wrote:
}
} }
} }
} }
} ----------------------------------------------------------------------------
}
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} }
} }
} ----------------------------------------------------------------------------
}
} } OK...
} }
} } Now I'm a little confused.........
} }
} } Because I was taught that a Pic file, like a screen shot is only a
} } rasterized 72 dpi image.
} }
} }
} } So I threw a 6400 dpi image into iMovie, and saved as a pic file, and
} } then opened in photoshop, and it says it's a 640 by 480 .............72
} } dpi image
} }
} }
} } Was the conversion on the import into iMovie, or is this what I've
} } always found with pic files, 72 dpi cause that's what the Mac does with
} } a pic file?
} }
} } So I save as JPEG, and it's a 243x144 pixel image and still only 72 dpi.
} }
} } If I use the OS X grab-it function for a screen shot, it actually saves
} } it in TIFF, but again, only 72 bit dpi.
} }
} } In which case it wouldn't make any difference just to use the System
} } application "Grab-it", select to capture selection, and save using
} } Grab-it, as both would be 72 dpi, and there would be no more conversion
} } involved
} }
} }
} }
} } Lou Ann
} }
} }
} } trazy-at-umn.edu wrote:
} }
} ----------------------------------------------------------------------------
}
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } }
} America
}
} } } To Subscribe/Unsubscribe --
} } }
} http://www.microscopy.com/MicroscopyListserver
}
} } } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } }
} } }
} ----------------------------------------------------------------------------
}
} } } The PICT files captured through iMovie are lossless, so this would be
} } } the flavor of choice. You can then easily convert PICT to TIFF if
} } } you wish.
} } }
} } } Cheers,
} } }
} } } --
} } } Tracy E. Anderson
} } }
} } }
} } }
} }
} } ==============================Original
} }
} Headers==============================
}
} } 16, 19 -- From lamiller-at-uiuc.edu Fri May 18 10:45:23 2007
} } 16, 19 -- Received: from expredir5.cites.uiuc.edu (expredir5.cites.uiuc.edu
} } [128.174.5.96])
} } 16, 19 -- by ns.microscopy.com (8.12.11.20060308/8.12.8) with ESMTP id
} } l4IFjMCi007315
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} } 16, 19 -- Received: from redkite.cvm.uiuc.edu (redkite.cvm.uiuc.edu
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} } 16, 19 -- Message-ID: {464DCA12.9010409-at-uiuc.edu}
} } 16, 19 -- Date: Fri, 18 May 2007 10:45:22 -0500
} } 16, 19 -- From: Lou Ann Miller {lamiller-at-uiuc.edu}
} } 16, 19 -- Reply-To: lamiller-at-uiuc.edu
} } 16, 19 -- User-Agent: Thunderbird 2.0.0.0 (Macintosh/20070326)
} } 16, 19 -- MIME-Version: 1.0
} } 16, 19 -- To: microscopy-at-microscopy.com
} } 16, 19 -- Subject: Re: [Microscopy] Re: viaWWW: Need Help From Imaging
} } Experts
} } 16, 19 -- References: {200705181526.l4IFQg4i007630-at-ns.microscopy.com}
} } 16, 19 -- In-Reply-To: {200705181526.l4IFQg4i007630-at-ns.microscopy.com}
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} }
} }
}
}
}
} ==============================Original Headers==============================
} 8, 20 -- From trazy-at-umn.edu Fri May 18 11:04:23 2007
} 8, 20 -- Received: from mta-w2.tc.umn.edu (mta-w2.tc.umn.edu [134.84.119.6])
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--
~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Lou Ann Miller, MT(ASCP)
Service Supervisor
Center for Microscopic Imaging
College of Veterinary Medicine
Rm 1204 VMBSB
2001 S Lincoln Ave
Urbana, IL 61802

217/244-1567
http://treefrog.cvm.uiuc.edu


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From: oshel1pe-at-cmich.edu
Date: Fri, 18 May 2007 11:18:31 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Download Graphic Converter from http://www.lemkesoft.com
It will convert PICT files to TIFF or any of a plethora of other file
formats. $30 shareware, free 30 day trial, nearly as powerful as
Photoshop and *much* easier to use.

Phil

} I see what he means , just tried iMovie on OS X 10.3.
}
}
} It seems to me though, that about 5 years ago, it DID let me save to
} TIFF and there were more options, perhaps trying to get an Old copy of
} iMovie? or try on an old machine if you still have OS 9?
}
} Also can you open the clip in Quicktime Pro? Maybe Quicktime Pro will
} let you pull a tiff off of it, it too will save a frame.
}
} Not a great solution, but if you are in a pinch.....
}
}
}
} Lou Ann
}
} Mike.Bode-at-olympus-sis.com wrote:
}
} } In that case I would suggest to ask Apple. Contact them and tell them
} } that for reasons of scientific Ethics you need the images in TIFF format
} } (or any other format that you think might work), and ask them if they
} } have a converter for their iMovie Product. They claim to be super
} } user-friendly. Put them to the test.
} }
} } mike
} }
} } Michael Bode, Ph.D.
} }
} } General Manager
} }
} } OLYMPUS SOFT IMAGING SOLUTIONS
} } 12596 West Bayaud Ave #300
} } Lakewood, CO 80228
} } USA
} } Tel.: +1 (303) 234-9270
} } Fax.: +1 (303) 234-9271
} } E-mail: Mike.Bode-at-olympus-sis.com
} } www.olympus-sis.com
} }
} } -----Original Message-----
} } X-from: Sharon Bledsoe [mailto:sbledsoe-at-iupui.edu]
} } Sent: Friday, May 18, 2007 06:12
} } To: Mike Bode
} } Subject: [Microscopy] RE: viaWWW: Need Help From Imaging Experts
} }
} } IF I had a choice I would choose to save my images as TIFF's, however,
} } the iMovie program will not let me. iMoive will only let me save as
} } PICT or JPEG.
} }
} } And I am not tied to the Mac, I'm chained to it.
} } } Hi Sharon,
} } }
} } } This may be a bit more fundamental than what you are looking for, but
} } } here goes:
} } }
} } } If you go to the MSA website, you can find a statement about ethical
} } } image processing that in essence says that images need to be stored
} } } uncompressed. JPEG is a format that includes lossy compression, and I
} } } believe that PICT can also contain lossy compression in the form of
} } } JPEG. Both therefore don't seem to be the best choices.
} } }
} } } I also found this on the web:
} } }
} } } In the next version of its operating system, MacOS X, Apple has decided
} } }
} }
} }
} } } to replace PICT by PDF. This means that the use of PICT for exchanging
} } } data will slowly diminish.
} } }
} } } It looks like sooner or later you will have to find a different choice
} } } anyway. I would look at the whole chain of acquisition to see if you
} } } cannot switch to something else.
} } }
} } } Why can't you use anything else? Does it have to do with the original
} } } file format that can only be read by iMovie? Or are you tied to the Mac
} } }
} }
} }
} } } platform?
} } }
} } } mike
} } }
} } }
} } } Michael Bode, Ph.D.
} } }
} } } General Manager
} } }
} } } OLYMPUS SOFT IMAGING SOLUTIONS
} } } 12596 West Bayaud Ave #300
} } } Lakewood, CO 80228
} } } USA
} } } Tel.: +1 (303) 234-9270
} } } Fax.: +1 (303) 234-9271
} } } E-mail: Mike.Bode-at-olympus-sis.com
} } } www.olympus-sis.com
}
} } } Email: sbledsoe-at-iupui.edu
} } } Name: Sharon Bledsoe
} } }
} } } Title-Subject: [Filtered] Need Help From Imaging Experts
} } }
} } } Question: Here is my problem: I can only use Mac iMovie to edit my
} } } digital video. I can not do this on a PC. ONLY iMovie. To save a
} } } single frame from the video, my choices are either PICT or JPEG format.
} } }
} } }
} } } My Question is: To save an origianl image, which is the best format,
} } } PICT or JPEG?
} } }
} } } I'm not interested in hearing about Tiffs, BMPs or any other format as
} } } that is not one of my choices.

} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~
} Lou Ann Miller, MT(ASCP)
} Service Supervisor
} Center for Microscopic Imaging
} College of Veterinary Medicine
} Rm 1204 VMBSB
} 2001 S Lincoln Ave
} Urbana, IL 61802
}
} 217/244-1567
} http://treefrog.cvm.uiuc.edu

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
voice: (989) 774-3576
dept. fax: (989) 774-3462

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From: lamiller-at-uiuc.edu
Date: Fri, 18 May 2007 11:23:35 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I guess I should qualify the question...... I can make the image
bigger on my screen while in iMovie.


Would then taking a bigger image as TIFF than one can with the save
frame command, with Grab-it be of any benefit?

Such as making it smaller and increasing the dpi in photoshop mathmatically?




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From: paul_hazelton-at-umanitoba.ca
Date: Fri, 18 May 2007 11:47:36 -0500
Subject: [Microscopy] Re: viaWWW: TEM and RuO4

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Patricia

The key to understanding the mechanism is that the structure of
Ruthenium Tetroxide is the same as that of Osmium Tetroxide, a central
heavy metal bound by double bonds to 4 oxygens. The reaction is the
same as for Osmium when it interacts with organic material - breaking of
one of the double bonds by the target and then cross linking of the
target with the RuO4. Contrast is the result of added density of the
heavy metal to the target.

Hayat is relatively quiet about the chemical reaction in his Fixation
for Electron Microscopy. I seem to have my copy of his Positive
Staining for Electron Microscopy at home so I don't know if he even
covers it as a stain, my memory is that he doesn't. It is also not
covered in Lewis and Knight's Staining Methods for Sectioned Material.


Paul



Paul R. Hazelton, PhD
Electron Microscope Unit
University of Manitoba
Department of Medical Microbiology
531 Basic Medical Sciences Building
730 William Avenue
Winnipeg, Manitoba, Canada, R3E 0W3
e-mail: paul_hazelton-at-umanitoba.ca
Phone:204-789-3313
Pager:204-931-9354
Cell:204-781-1502
Fax:204-789-3926





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From: wesaia-at-iastate.edu
Date: Fri, 18 May 2007 12:26:23 -0500
Subject: [Microscopy] viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

As this discussion proceeds, please remember the pixel count is what's
important and not the size of the image in inches or the DPI.

If you make that image bigger on the screen it will give you a few more
pixels in the grabbed image, but where did those pixels come from? You
have not changed anything in the native resolution of the movie images.
If the image was originally 640x480, new pixels will not magically
multiple.

In fact, if you don't increase the pixel count an integral number of
times (e.g., 1280x960) you may introduce questions due to mapping. A new
pixel will likely consist of contributions of multiple old pixels, or if
you prefer, a single old pixel will contribute to multiple new pixels.
Teh details depend on how the software magnifies small images. Still,
you are not creating new information, so leave the image size at
640x480.

Personally, I still have little use for the DPI (actually PPI) value.
Sure it will interact with some document programs to insert an image at
the "proper size". I still prefer just knowing the pixel count and the
image width and I can do the math myself. DPI just seems to confuse the
matter for most folks.

Warren

-----Original Message-----
X-from: lamiller-at-uiuc.edu [mailto:lamiller-at-uiuc.edu]
Sent: Friday, May 18, 2007 11:26 AM
To: wesaia-at-iastate.edu

I guess I should qualify the question...... I can make the image
bigger on my screen while in iMovie.


Would then taking a bigger image as TIFF than one can with the save
frame command, with Grab-it be of any benefit?

Such as making it smaller and increasing the dpi in photoshop
mathmatically?



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From: gary.m.brown-at-exxonmobil.com
Date: Fri, 18 May 2007 13:00:48 -0500
Subject: [Microscopy] viaWWW: TEM and RuO4

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Patricia,

As Paul, I'm working from memory. However, I recall (hopefully accurate)
information on mechanisms from Sawyer and Grubbs or Trent. I believe that
the mechanisms for OsO4 and RuO4 oxidation of polyolefins is different.
OsO4 adds across C=C bonds and does not react with C-C bonds at all. RuO4,
on the other hand, scissions C-C or C=C bonds to produce oxygenated chain
ends.

Regards,

Disclaimer: The comments and opinions are those of the author alone and do
not reflect any official position by his employer.

Gary M. Brown
Microscopy Specialist
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

"Happiness equals reality minus expectations." Tom Magliozzi





paul_hazelton-at-
umanitoba.ca
To
gary.m.brown-at-exxonmobil.com
05/18/07 11:50 cc
AM
Subject
[Microscopy] Re: viaWWW: TEM and
Please respond RuO4
to
paul_hazelton-at-
umanitoba.ca










----------------------------------------------------------------------------

The Microscopy ListServer -- CoSponsor: The Microscopy Society of America


Patricia

The key to understanding the mechanism is that the structure of
Ruthenium Tetroxide is the same as that of Osmium Tetroxide, a central
heavy metal bound by double bonds to 4 oxygens. The reaction is the
same as for Osmium when it interacts with organic material - breaking of
one of the double bonds by the target and then cross linking of the
target with the RuO4. Contrast is the result of added density of the
heavy metal to the target.

Hayat is relatively quiet about the chemical reaction in his Fixation
for Electron Microscopy. I seem to have my copy of his Positive
Staining for Electron Microscopy at home so I don't know if he even
covers it as a stain, my memory is that he doesn't. It is also not
covered in Lewis and Knight's Staining Methods for Sectioned Material.


Paul



Paul R. Hazelton, PhD
Electron Microscope Unit
University of Manitoba
Department of Medical Microbiology
531 Basic Medical Sciences Building
730 William Avenue
Winnipeg, Manitoba, Canada, R3E 0W3
e-mail: paul_hazelton-at-umanitoba.ca
Phone:204-789-3313
Pager:204-931-9354
Cell:204-781-1502
Fax:204-789-3926





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From: lamiller-at-uiuc.edu
Date: Fri, 18 May 2007 13:46:49 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Yes, that is the good question "What does iMovie do with the image when
it is spread on the screen?"

An automatic .... it just spreads the pixels, actually may not be accurate.


I have a small box when I marginalize iMovie on my computer screen, say
4-5 inches ( also I do have a $4K Sony high resolution Pro-video
camera), yet, as I do a lot of movies to DVD using iMovie, on a large TV
screen, the product of iMovie does not appear to be pulling apart a 72
dpi small box., that's going from a 4 inch box to a 23 inch screen with
only a slight loss of resolution, and that doesn't seem to fit the
pulling apart of what we think of as a normal, email, web page weight
images we know as 72 dpi.


So does iMovie just spit out the 72 dpi on the frame picture export
work, and give back better stuff with more resolution with it's movie
processing when the source was higher resolution? Or is the eye being
fooled by the ~33 frames per second presentation?


I'll have to play with that when I get home,


Lou Ann









Straszheim, Warren E [CCE E] wrote:
} As this discussion proceeds, please remember the pixel count is what's
} important and not the size of the image in inches or the DPI.
}
} If you make that image bigger on the screen it will give you a few more
} pixels in the grabbed image, but where did those pixels come from? You
} have not changed anything in the native resolution of the movie images.
} If the image was originally 640x480, new pixels will not magically
} multiple.
}
} In fact, if you don't increase the pixel count an integral number of
} times (e.g., 1280x960) you may introduce questions due to mapping. A new
} pixel will likely consist of contributions of multiple old pixels, or if
} you prefer, a single old pixel will contribute to multiple new pixels.
} Teh details depend on how the software magnifies small images. Still,
} you are not creating new information, so leave the image size at
} 640x480.
}
} Personally, I still have little use for the DPI (actually PPI) value.
} Sure it will interact with some document programs to insert an image at
} the "proper size". I still prefer just knowing the pixel count and the
} image width and I can do the math myself. DPI just seems to confuse the
} matter for most folks.
}
} Warren
}
}


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From: gary.m.brown-at-exxonmobil.com
Date: Fri, 18 May 2007 15:33:01 -0500
Subject: [Microscopy] Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Folks,

I would appreciate any insight that the microscopy community might have
regarding safe practices and tools for cutting and trimming polymer
samples. Our lab members generally use single- and double-edge razor
blades for small jobs. Any suggestions on these or other tools, blade
guards, etc. will be appreciated.

Thanks,

Disclaimer: The comments and opinions are those of the author alone and do
not reflect any official position by ExxonMobil Chemical Company.

Gary M. Brown
Microscopy Specialist
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

"Happiness equals reality minus expectations." Tom Magliozzi



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From: paul_hazelton-at-umanitoba.ca
Date: Fri, 18 May 2007 15:33:42 -0500
Subject: [Microscopy] Re: viaWWW: TEM and RuO4

Contents Retrieved from Microscopy Listserver Archives
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Patricia

Gary's right, the reaction of the C=O with unsaturated C=C double bonds
- such as seen with olefins results in a five atom circular structure
between the original 2 carbons, and two O's and the Os from the OsO4.
Also, as he points out, with olefins, as more OsO4 gets into the act you
can see scissions of the C-C bond. At least that is the theory, I'm
just a scope jockey/structural/molecular virologist. In my humble
opinion it takes a very different brain than mine to understand that
organic chemistry stuff.

Paul


Paul R. Hazelton, PhD
Electron Microscope Unit
University of Manitoba
Department of Medical Microbiology
531 Basic Medical Sciences Building
730 William Avenue
Winnipeg, Manitoba, Canada, R3E 0W3
e-mail: paul_hazelton-at-umanitoba.ca
Phone:204-789-3313
Pager:204-931-9354
Cell:204-781-1502
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From: colijn.1-at-osu.edu
Date: Fri, 18 May 2007 20:04:37 -0500
Subject: [Microscopy] Re: C2 apertures

Contents Retrieved from Microscopy Listserver Archives
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Hi Krassimir,

Apparently at one time, FEI sold sort of a super-TopHat condenser
aperture that was essentially a number of condenser apertures stacked
on each other with some sort of low-Z material in between. Perhaps
that is the aperture you are describing. The amazingly high cost
could be due to the obvious difficulty of aligning the stacked apertures.

Unless you need incredibly clean EDS spectra, try a normal Pt TopHat
aperture or even a regular 250micron thick Pt condenser aperture. Of
course, you could always try stacking some apertures yourself!

Cheers,
Henk

At 07:05 PM 5/17/2007, you wrote:



} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

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From: Sander.Stoks-at-fei.com
Date: Sat, 19 May 2007 05:29:00 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Lou Ann,

Please, forget about the "dpi" rating of the image. What is important (as has been pointed out in a previous reply) is simply the _total number_ of pixels. After all, you can increase the dpi very simply by displaying the image on a tiny screen. The same number of pixels on fewer inches, so higher dpi. Of course, that did not improve the actual image content at all.

In a previous life, I wrote consumer level non-linear video editing software.

Usually, video editing software works in such a way that the final resolution is determined by the format you're exporting the movie to. If you're targetting a low-bandwidth "web movie", it is rendered to 320x240 or thereabouts. All source clips and images are re-rendered in this target resolution. If you added a high-resolution still image (from an 8 megapixel photo camera) in the middle of your movie, this image will be downsampled to the final image resolution, and not somehow stored in the movie in its original resolution.

If you target a more high-quality format such as DVD or even HD, all source clips and images are rendered into this higher resolution format. If your originals were in low quality, the resulting images are simply "stretched to fit". This is the same effect as when you enlarge your media player to full screen: there is no "extra information" - (after all, the media player would somehow have to "make it up" since it is not present in the movie data); the existing information is simply interpolated to give it a "smooth appearance". Added to the way human perception works for moving images, this gives an excellent _perceived_ quality.

But there's more.

If you're capturing movies with a digital camera, chances are very high that there is already some compression being performed during the capture. If it's a DV camera, the quality of these images is quite good; but if it's a modern MPEG-2 camera, you can expect lots of compression artefacts. Movies taken this way are simply not meant to be dissected frame-by-frame, as human perception for moving images and for still images is very different. This is extra visible if frames are captured interlaced (i.e. one field will contain all the odd lines, the next field all the even ones). This is great for moving images as this will actually increase the perceived quality, but looking at one such a field in a still will show hideous jagged edges.

So again, if you extract a single frame from a digitized, compressed movie, exporting it to uncompressed TIFF will not magically increase its quality. It will only not make it _worse_.

You will probably get better suggestions from the list if you don't limit our choices to "A or B", but tell us exactly what you're trying to accomplish.

Regards,
Sander Stoks (not speaking for FEI)


________________________________

X-from: lamiller-at-uiuc.edu [mailto:lamiller-at-uiuc.edu]
Sent: Fri 5/18/2007 8:52 PM
To: Stoks, Sander

Yes, that is the good question "What does iMovie do with the image when
it is spread on the screen?"

An automatic .... it just spreads the pixels, actually may not be accurate.


I have a small box when I marginalize iMovie on my computer screen, say
4-5 inches ( also I do have a $4K Sony high resolution Pro-video
camera), yet, as I do a lot of movies to DVD using iMovie, on a large TV
screen, the product of iMovie does not appear to be pulling apart a 72
dpi small box., that's going from a 4 inch box to a 23 inch screen with
only a slight loss of resolution, and that doesn't seem to fit the
pulling apart of what we think of as a normal, email, web page weight
images we know as 72 dpi.


So does iMovie just spit out the 72 dpi on the frame picture export
work, and give back better stuff with more resolution with it's movie
processing when the source was higher resolution? Or is the eye being
fooled by the ~33 frames per second presentation?


I'll have to play with that when I get home,


Lou Ann




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From: zaluzec-at-microscopy.com
Date: Sat, 19 May 2007 10:16:30 -0500
Subject: [Microscopy] Digital Imaging, Digital Movies and Compression

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Colleagues

There is a thread currently discussing digital imaging and movies etc...

Let me take a moment to remind you (or reinforce with others have
said or implied) namely : if you save a file in any "lossy" compressed
format your changing as well as loosing data. This must be reported
if you use any of this processed data for any scientific analysis.

The last time I checked there are some 60+ different image file formats.

Lossless formats include (but are not limited to ) the following:
RAW, TIFF, BMP, PICT, PNG, PCX, EXR, SVG, TGA, JPEG2000

Lossy formats include: GIF, JPEG, MPEG, MOV, H263, Video

On top of this there are also lossy and lossless COMPRESSION methods, which
may be encoded within/upon some of these formats. So life can get
very complicated , you might use a lossless method and then compress
the data using a lossy alogrithm.

The Microscopy Society of America (MSA) has defined a policy on ethical
digital imaging, which is available on line under Reference/Education section
of their WWW site (http://www.microscopy.org) . This ethical
position applies to all digital images, be they: still , time series or movies.

While making movies from time series events etc.. is a valuable
mechanism of looking through your data, (and I also do this frequently).
Remember that digital imaging ethics requires that you store the
original data using a procedure similiar to that documented below.
If you analyze your data you should refer back to the original uncompressed
data, never to the compressed data.

You also need to report these operation particuliarly if you downsample,
compress etc your data. Consumer grade software does this routinely
and likely does not tell you so. So be extremely careful
when using some of this software for analysis your scientific data!

If you are recording directly to digital video, I would recommend that
you record in RAW format store that data and then after the fact convert
your images to some alternative display.

-----------------------------------------------------------------------------------
Here is the MSA statement verbatum.
-----------------------------------------------------------------------------------

The MSA position on digital image processing has been approved as follows:

"Ethical digital imaging requires that the original uncompressed image file be stored on archival media (e.g., CD-R) without any image manipulation or processing operation. All parameters of the production and acquisition of this file, as well as any subsequent processing steps, must be documented and reported to ensure reproducibility.

Generally, acceptable (non-reportable) imaging operations include gamma correction, histogram stretching, and brightness and contrast adjustments. All other operations (such as Unsharp-Masking, Gaussian Blur, etc.) must be directly identified by the author as part of the experimental methodology. However, for diffraction data or any other image data that is used for subsequent quantification, all imaging operations must be reported."

This policy was formulated by the Digital Image Processing & Ethics Group of the MSA Education Committee and was adopted as MSA policy at the Summer Council meeting August 2-3, 2003.




Nestor
Your Friendly Neighborhood SysOp

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From: lamiller-at-uiuc.edu
Date: Sat, 19 May 2007 10:20:06 -0500
Subject: [Microscopy] viaWWW: Imaging

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Email: lamiller-at-uiuc.edu
Name: Lou Ann Miller

Organization: U of Illinois

Title-Subject: [Filtered] Imaging

Question: Thanks Sander and Warren,


Those answers helped explain what happens with the video, which was what my question was if not stated very clearly, you don't have to beg me to give up on dpi, ;-)

But I was curious as to what exactly the video software was actually doing to the image when, and it's good to know what mixed resolution inputs and expected outputs will do, and that the camera is actually doing some compression ( like scanners might ) to the image.


I'm a videographer on weekends, basically putting our church services to DVD for shut ins ( we do pay a license for copyright) and the sermon movies online (stjohnsermons.org). What one does with what is dependent upon what works best for the task, and not the dpi. I don't use grab-it, but do use the export on the software.

For example, I capture action shots and dim lit areas far off (balcony is a long LONG ways back) images off the video frames than I do with my very very nice 8 mega pixel camera even with the zoom lens, better meaning I got the data I wanted with out blurring...... confirmation, school graduations etc etc. Also, since even my digital camera is a reflex, and sounds like the paparazzi in a place with excellent sound carrying capability and tiled floor place, video suits better than even the 8 mega pixel very nice lens system in some situations. For other uses, the camera wins over the video frames hands down.


I'm sure in microscopy it is the same, what imaging technique is used, very much dependent on what works best for the best results.

So usage and results, and what is being done to the image, do win over the dpi. I know people who buy pocket cameras the size of 1/2 a wallet with huge mega pixel value, but their images don't turn out as well as they expect for buying that mega pixel value ...... because they don't use a good lens system...and probably other reasons you can tell me about for sure :-)

Thanks for the discussion! I've enjoyed it.



Lou Ann

(email failed from my home computer so submitting via web)

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From: Sander.Stoks-at-fei.com
Date: Sat, 19 May 2007 14:27:54 -0500
Subject: [Microscopy] Digital Imaging, Digital Movies and Compression

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Hello all,

Note that BMP _used_ to be a lossless format, but since (I think) Windows 2000 a valid BMP file _can_ in fact contain a JPEG compressed (i.e., lossy) image. I've yet to run into one of those "in the wild".

Also: GIF does not use lossy compression (but is never used for serious image processing).

Regarding "RAW video": Suppose you are capturing full NTSC frames (there are different opinions on what the exact resolution is, but let's take 720x540 for the example) at 30 full frames per second (actually 29.97), in a raw format with three bytes per pixel this would take more than 32MB per second of footage. This would fill up a normal-sized hard disk quite rapidly. What's worse, up to a few years ago consumer grade hard disks couldn't even store data at this speed. Compression wasn't added to movie formats just to save space and costs; it was mainly because there simply wasn't any other way of doing real-time image acquisition otherwise. I don't know of any current (affordable) video cameras which can store their captured data in a raw format, but perhaps the state of the art has advanced more than I know.

Also, there's the problem that even if you pick a well-known image format and stay away from the "obscure" ones, you'll still run into spotty support even on major software platforms. For example, I regulartly get "complaints" from people claiming my TIFF exporter is broken because Word can't read the resulting TIFFs. Usually, that's because they've chosen 16-bit export, and Word only partially implements the (huge) TIFF specification and simply can't read all valid TIFFs. Likewise, Windows Explorer won't show any thumbnails for those images, because it only handles 8 bit-per-channel TIFFs.

The whole of "image postprocessing" is a bit of a gray area. If you use postprocessing to downsample an image you should report it, but if you select "binning 2" on your camera when doing the acquisition, you're in effect doing the same thing. Even worse, some consumer-grade cameras perform a host of image post processing _in the camera itself_ to make their images look better. It's not uncommon to see an "artificial sharpening filter" being performed to get away with a cheaper lens system.

I agree with reporting on the entire post-processing setup though, and with storing your original images in a lossless format, preferably TIFF, with sufficient bit depth (if you have a 10 bit TEM camera, don't store your images as BMP even if that format is listed as "lossless"). Even if some of your software can't correctly handle these original files, there are plenty of (free) conversion libraries around (many open source, so they will _stay_ around) which can convert it to any format you currently need.

Regards,
Sander Stoks (not speaking for FEI)

________________________________

X-from: zaluzec-at-microscopy.com [mailto:zaluzec-at-microscopy.com]
Sent: Sat 5/19/2007 5:27 PM
To: Stoks, Sander

Colleagues

There is a thread currently discussing digital imaging and movies etc...

Let me take a moment to remind you (or reinforce with others have
said or implied) namely : if you save a file in any "lossy" compressed
format your changing as well as loosing data. This must be reported
if you use any of this processed data for any scientific analysis.

The last time I checked there are some 60+ different image file formats.

Lossless formats include (but are not limited to ) the following:
RAW, TIFF, BMP, PICT, PNG, PCX, EXR, SVG, TGA, JPEG2000

Lossy formats include: GIF, JPEG, MPEG, MOV, H263, Video

On top of this there are also lossy and lossless COMPRESSION methods, which
may be encoded within/upon some of these formats. So life can get
very complicated , you might use a lossless method and then compress
the data using a lossy alogrithm.

The Microscopy Society of America (MSA) has defined a policy on ethical
digital imaging, which is available on line under Reference/Education section
of their WWW site (http://www.microscopy.org) . This ethical
position applies to all digital images, be they: still , time series or movies.

While making movies from time series events etc.. is a valuable
mechanism of looking through your data, (and I also do this frequently).
Remember that digital imaging ethics requires that you store the
original data using a procedure similiar to that documented below.
If you analyze your data you should refer back to the original uncompressed
data, never to the compressed data.

You also need to report these operation particuliarly if you downsample,
compress etc your data. Consumer grade software does this routinely
and likely does not tell you so. So be extremely careful
when using some of this software for analysis your scientific data!

If you are recording directly to digital video, I would recommend that
you record in RAW format store that data and then after the fact convert
your images to some alternative display.

-----------------------------------------------------------------------------------
Here is the MSA statement verbatum.
-----------------------------------------------------------------------------------

The MSA position on digital image processing has been approved as follows:

"Ethical digital imaging requires that the original uncompressed image file be stored on archival media (e.g., CD-R) without any image manipulation or processing operation. All parameters of the production and acquisition of this file, as well as any subsequent processing steps, must be documented and reported to ensure reproducibility.

Generally, acceptable (non-reportable) imaging operations include gamma correction, histogram stretching, and brightness and contrast adjustments. All other operations (such as Unsharp-Masking, Gaussian Blur, etc.) must be directly identified by the author as part of the experimental methodology. However, for diffraction data or any other image data that is used for subsequent quantification, all imaging operations must be reported."

This policy was formulated by the Digital Image Processing & Ethics Group of the MSA Education Committee and was adopted as MSA policy at the Summer Council meeting August 2-3, 2003.




Nestor
Your Friendly Neighborhood SysOp

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From: john_mackenzie-at-ncsu.edu
Date: Sun, 20 May 2007 00:57:04 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

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Folks

The problem here is that saving in iMovie has compressed the data to 640
by 480 (or as several have pointed out 10 or 15 variants of this) So you
start out with really awful resolution. THEN the data is compressed by
imovie to be able to handle the movie without needing terabyte drives.
The compression of the movie is in MPEG 2 if it is DVD compliant. The
codec can vary here if you are not trying to have a format that can be
played on your TV. (At last count my computer has about 50 different codecs)

Before we start talking about frames, your data has thrown away anything
bigger than 640 by 480 and compressed it alot.( This is an
understatement.. ALOT!!!) The data that you grab will be 640 by 480
compressed. If you use a lossless format as many have suggested then you
won't lose any more. Any method that you now use will never, never,
never make the data better than 640 by 480.

All you are doing is interpolating the data up. This will not improve
anything. If you are not careful the interpolation may include
sharpening and/or smoothing. You would therefore need to document this.
The reason that this all gets so complicated is that many of the
suggestions are trying to "fix" the loss of resolution that occured when
the data was aquired in imovie and this just can't be done.

The 72 dpi is meaningless for alot of reasons
1. The 72 dpi came from the screen resolution of computers in 1990.
2. The dpi is only really descriptive when you are talking about output.
3. Dpi is meaningless as the dot is defined so many different ways that
you really have no idea what the dot is at any given time.

If you are restricted to movie format then best way to do it is to
record on mini dv format. The sony cameras have an internal hardware
compressor that is DVD compliant. It records data at 40 Megabytes per
second. This is about 10 times the data rate that "direct digital"
methods can achieve. The digital tape can record MUCH faster than a hard
drive. After recording you can bring the data into thew computer over
the IEEE 1394 (firewire) input. This is a frame accurate data transfer
that does not change the data in any way. You do not lose one bit of data.
The frames that you then can save need to be saved in a lossless manner.
I would suggest you use Adobe Premier because it is a Mac program and
you can save in a large variety of lossless formats.

Please understand that the chip that does the compression in the Sony
cameras is one of those best kept secrets. Sony developed an
unbelievably powerful chip so that they could own the mini dv format.
They won but unfortuneately for them consumers are much more interested
in BluRay and HD DVD

The documentation for publication needs to describe the acquisition
chain as you are not recording anything that is uncompressed. The MSA
guidelines that Nestor pointed you to would require that you report that
hardware/software system so that others can reproduce your results

A HD tv camera would improve things greatly but I have not had time to
run any tests to see exactly what the bandwidth of the system is. Since
you would be putting 4 times as much data onto the tape you might have
to use a chip that compresses it more than the Sony does for NTSC/PAL
format.

John

John M. Mackenzie, Jr., PhD

Professor of Microbiology
Coordinator, Center for Electron Microscopy
North Carolina State University
Phone (919) 515-2664 Fax (919) 515-8293



lamiller-at-uiuc.edu wrote:
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} I guess I should qualify the question...... I can make the image
} bigger on my screen while in iMovie.
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} Would then taking a bigger image as TIFF than one can with the save
} frame command, with Grab-it be of any benefit?
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} Such as making it smaller and increasing the dpi in photoshop mathmatically?
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From: kraftpiano-at-gmail.com
Date: Sun, 20 May 2007 00:58:45 -0500
Subject: [Microscopy] SEM: Quick vacuum question.

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Let's say you have a scale of 0-100 for your vacuum gauge. You need
95 in order to get a full emission reading on the SEM. The roughing
pump works fine, brings it up to about 30, and the diffusion pump
kicks in and rapidly brings it up to 90.

Here's the symptom:

The emission current never comes on. I can hear the high pitched
sound of the high voltage, and occasionally the emission will peak at
20-30%, but then quickly die down again. There are two things that I
can think of that could be wrong. First, there is a capacitor in the
emission circuitry that is going a little haywire. Second, the vacuum
system is fluctuating just above and then just below optimal levels.

Would low diffusion pump oil create a symptom like that where the
vacuum fluctuates (Incidentally, the vacuum meter doesn't move, but
I'm guessing that it isn't stable because of the fluctuating emission
current). I thought that having a low amount of diffusion pump oil
would just result in a lower vacuum, not necessarily a fluctuating
one.

Thanks in advance,

Justin A. Kraft

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From: keith.morris-at-ucl.ac.uk
Date: Sun, 20 May 2007 08:16:34 -0500
Subject: [Microscopy] Re: viaWWW: Need Help From Imaging Experts

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Dear Justin,
some possibilities:

1- diffusion pump oil may be too low; use at main diff pump 100cc, column
diff pump 35cc oil filling. Warm up pumps, drain, check liquid (should be
white/ yellow, not brownish). Use same oil as had been inside the pump. If
you do not know if for ex. Santovac had been used, you have to clean the
pumps thoroughly. Use NEW O-rings (special silicon? because of heat) to seal
pumps again. Never use the old O-rings on the diff pump drains again!!!

2- You rebuild the complete column / pumping lines. There will be a lot of
water inside all lines. This may take some days of continuos pumping to get
rid of.

3-I suppose you checked the filament. Is it OK? Did you get a reading from
the socket of the wehnelt? Is this OK? Should be some ohms... If not, clean
the "feets" ofr the filament with very fine sandpaper. Sometimes long stored
old filaments build up insulating layers...

4- Try some very low accelerating voltage, 2 KV or so. Are the fluctuations
gone? If so, check for cleanliness of cathode / anode and keep the
instrument pumping for some days...

Good luck,
Stefan Diller



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----- Original Message -----
X-from: {kraftpiano-at-gmail.com}
To: {stefan.diller-at-t-online.de}
Sent: Sunday, May 20, 2007 8:08 AM

Hi

I am afraid on first sight your problem does not look like a "quick vacuum
question".

Lets look at your comments regarding the vacuum and DP. With a level of
fluid that is too small the DP will pump in "spurts" as the fluid boils off
and then has to wait for condensation before it may re cycle. In most cases
this will be adequate to hold a specific vacuum but it may take longer to
reach a good operating pressure. The position of most manufacturers vacuum
gauges means that they only give a rough clue as to what is going on in the
gun so they will not help here. Removing the gun cable and replacing it
with a gauge is the only way to really find out what is happening in the gun
area - an engineer task in most laboratories.

Much more to the point, and in my view the problem, is the high voltage
situation. If the vacuum is not good enough for the high voltage to operate
it will switch off. If a component is breaking down in the high voltage
circuitry it will switch off. To isolate the high voltage system from the
vacuum may help in the investigation.

Switch off the high voltage and remove the cable from the tank. Wrap the
cable end in aluminium foil and cover the tank connection socket. Switch on
the high voltage watch the emission meter and listen. Same sounds? Same
meter reactions? Nothing changes then it is a tank problem. Different
symptoms then it is a high voltage cable or gun vacuum problem.

Can we find out more?

Steve Chapman
Protrain for EM training & consultancy world wide
www.emcourses.com
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967


----- Original Message -----
X-from: {kraftpiano-at-gmail.com}
To: {protrain-at-emcourses.com}
Sent: Sunday, May 20, 2007 7:00 AM

Hi all,

Here when we produce a 'video' time-lapse on our optical microscope systems
we are actually collecting a series of uncompressed TIF 'photographs' at
12-bit (4096 grey levels) or similar in a propriety format. These images are
therefore discrete and if we want to 'extract' a quality still image we
simply export/save one from the list, normally in .jpg hi-res compressed
format for articles etc.. where the slight loss of image quality is
unimportant given the image size in print, or as original uncompressed TIF
format for image analysis if that program can't read the original file
format. The cameras or detectors we use are typically things like an £10,000
Hammamatsu Orca ER B&W one. Technically at maximum resolutions of around
1280x1280 this is only just over a 1 megapixel camera, but of course the
camera has excellent sensitivity at low light for fluorescence samples and
has very low noise (I can see room for future improvement in resolution
though). Our 'video' image sequences will thus be created from a series of
microphotographs of live cells obtained using a £80k to £250k optical
time-lapse wide-field or confocal optical microscope system. Images being
captured at a maximum rate of one every few seconds unless we are prepared
to take a hit in resolution (e.g. binning pixels) and/or noise for capturing
rapid intra-cellular activity.

Thus to produce a 'proper' DVD type video, we export these TIF image
sequences into a .mov (Apple) or .avi (PC) format. We are often only
concerned that this exported video looks very similar to the original TIF
sequence collected by say Kinetic Imaging AQM, OpenLabs, Leica LCS, Bio-Rad
Lasersharp etc. For lab presentations we go for maximum quality during
compression (or even lossless) as a movie of 100 Mb+ is no problem for a
modern PC during presentation, and generally we just use the 'export to
movie/video' option on the microscope/camera image capture software. As
Microsoft Office on the PC now inconveniently doesn't support .mov video any
more we mainly use £25 Quicktime Pro [v6] to convert the Apple .mov to PC
.avi format afterwards (normally with the setting to minimum compression &
maximum quality as our video sequences aren't that long, say up 1,000 images
maximum and often far far less). On the odd occasions that we require a
100Mb+ video to be compressed to under 4Mb (say for web support of a
published article) we then turn to our single £200 Adobe Premiere Pro (v1.5)
software and play around with the export settings until we just hit 4Mb
limit and image quality is adequate to illustrate the point (and we still
have the original lossless image sequences should anyone wish to view them).


QuickTime Pro (and probably iMovie) can't compete with professional video
software for this type of thing, but as it's so simple to use we stick to
QuickTime Pro for routine video conversion of .mov to .avi (only I really
know anything about using Adobe Premiere Pro here). For Apple users you
would probably benefit with something like Final Cut Studio 2 as the
professional video editing option, or indeed Adobe Premiere CS3 that runs on
Apples (to get all features you will need boot-camp as well, but I doubt
that's necessary for single frame export or further video compression). Note
that you can run QuickTime .mov in PowerPoint 2003, but you have to insert
it as a link to QuickTime viewer, and most users here prefer the simplicity
of converting the video sequence to .avi which runs natively within
PowerPoint. Plus you have to add the .jpg, .avi or .mov suffix on export
from an Apple to get the PC to notice what type of file it is [e.g. for
auto-play and preview].

So as we often want to routinely export our 'video' images into hi-res
12-bit uncompressed TIFF [from a microscope], we got a decent (and
expensive) microscope camera and capture in lossless TIFF type format in the
first place as a video time-lapse sequence (rather than use a cheap NTSC/PAL
video camera). Generally these cameras were supplied by the microscope
manufactures and/or the image capture/processing software company (e.g.
OpenLabs, Zeiss, Leica), and we were happy with their recommendations (it’s
a small percent of the system total cost and they often write the camera
drivers).

I would add that rather like applying complex image processor filters to
images, it's far easier just to play around with the export quality settings
in something like £25 QuickTime Pro or better still £200 Adobe Premiere CS3
and see how far you have to go to get an acceptable image. This is easier
than trying to think too much and too deeply about what would work best. The
software normally offers a big guide hint with terms like 'maximum quality'
and 'maximum compression' or 'minimum file size & minimum quality'. What is
acceptable naturally varies widely depending on what the image is for, but
normally for us it will look very very much like the original TIF image on
casual inspection.

But if you have a compressed video image sequence to start with, or you have
them archived as uncompressed TIFF sequences elsewhere in an OpenLabs
propriety .Liff or Leica LCS .Lei file/folder, you have far less worries
about what you do - in both cases you have the unadulterated original anyway
should anyone raise questions and you are often just providing the small
article photo or highly compressed on-line video. Thus the exported frames
quality is just set to simply what is acceptable for the task in hand (in
terms of image quality, file storage size and what you want to show). It's
very easy to see when you have gone too far in terms of compression, bit
depth etc - but make sure your VDU monitor and graphics card are set to a
correct high bit depth, brightness/contrast and large pixel number screen
'size' that can show the exported image/video correctly. Plus 12-bit B&W
images need to be exported converted to 256 grey levels (8-bit) or 16-bit
for Photoshop to work with them correctly (off-line image analysis programs
expect 12-bit and so will have no problems with the original).

In most cases we only really need to use our uncompressed 12-bit TIFF
sequences for image analysis as it's pretty essential for image intensity
measurements and it's known pixel area is easier to calibrate for things
like manual count per unit area and object area (plus it's in that format
anyway). For home use I would love to keep all my colour SLR photographs in
.raw or TIF format, but in practice have to store them in hi-quality jpeg to
prevent the hard drive filling up in a few weeks (don't edit and save .jpg
images repeatedly though as you will lose detail with each re-compression
file save - that’s another advantage of TIF/RAW and a patient disposition
when working on them).

With 'proper' DVD/Camcorder video, I do find that working with a Sony MiniDV
Camcorder video using cheap [Nero 7 type] authoring software to capture and
write a DVD in a standard home PC always seems to stick in some compression
and nasty pixelisation somewhere, even when I say please don't do that at
all (compared to the original tape image quality on a nominally 625 line PAL
TV). It's also worse captured to PC via its USB2 station compared to
Firewire lead (as stated in the manual). However when I use the analogue PAL
output via S-Video lead from the MiniDV camcorder straight into the analogue
AV input of a dedicated TV DVD recorder the DVD image quality in XP/SP is
excellent (as good as standard PAL TV on casual viewing). Seems strange
given the two times analogue to digital conversion involved, so I assume
it's just that the dedicated £300 TV DVD recorder is doing a far better job
with writing DVDs than my old PC, as that’s all it does for the money (it
can't play Doom 3 or run Photoshop though). John's just gone into this
aspect in more detail (and I'm not NTSC friendly except for making sure I
can play Region 1 DVD film releases on my TV). I can't use my works copy of
Premiere Pro on this home PC with my Sony camcorder to test lossless or
lower compression rates (as suggested by John) as it's got an AMD older
style Athlon CPU chip (not compatible).

I know the above is the exact inverse of the question (and tends to ignore
it), but hopefully it will be on interest to a few list-servers
(particularly to those of us who find the majority of the electron
microscope threads merely 'quite interesting').

Regards

Keith J Morris

PS Adobe Premiere is great value as Create Suite three (CS3) Production
Premium, where you get:

Photoshop Adobe After Effects (graphics and visual effects)
Adobe Premiere® Pro CS3 (video editing)
Adobe Photoshop® CS3 Extended (photo-editing)
Adobe Flash CS3 Professional (interactive web and instructional content),
Adobe Illustrator CS3 (drawing/illustration)
Adobe Soundbooth™ CS3 (film audio)
Adobe Encore® CS3 (DVD authoring, but I tend to use Nero 7).

Normally available very cheaply as the complete CS3 package (so getting the
less useful programs is just a free bonus). CS3 isn't released yet though.

Just a shame that Adobe, unlike Microsoft, have no concept of user friendly
software - e.g. Adobe help always discusses a command in almost enough
detail without ever saying where you can actually find it, and in Indesign
(DT Publisher) they put the spell checker on the left under Edit, instead of
on the right somewhere like Office and everyone else, so that it takes 5
minutes to find it first time (sure they aren't related to Apple?).

-----------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology
Institute of Ophthalmology
UCL, London EC1V 9EL














-----Original Message-----
X-from: john_mackenzie-at-ncsu.edu [mailto:john_mackenzie-at-ncsu.edu]
Sent: 20 May 2007 07:07
To: keith.morris-at-ucl.ac.uk

Folks

The problem here is that saving in iMovie has compressed the data to 640
by 480 (or as several have pointed out 10 or 15 variants of this) So you
start out with really awful resolution. THEN the data is compressed by
imovie to be able to handle the movie without needing terabyte drives.
The compression of the movie is in MPEG 2 if it is DVD compliant. The
codec can vary here if you are not trying to have a format that can be
played on your TV. (At last count my computer has about 50 different codecs)

Before we start talking about frames, your data has thrown away anything
bigger than 640 by 480 and compressed it alot.( This is an
understatement.. ALOT!!!) The data that you grab will be 640 by 480
compressed. If you use a lossless format as many have suggested then you
won't lose any more. Any method that you now use will never, never,
never make the data better than 640 by 480.

All you are doing is interpolating the data up. This will not improve
anything. If you are not careful the interpolation may include
sharpening and/or smoothing. You would therefore need to document this.
The reason that this all gets so complicated is that many of the
suggestions are trying to "fix" the loss of resolution that occured when
the data was aquired in imovie and this just can't be done.

The 72 dpi is meaningless for alot of reasons
1. The 72 dpi came from the screen resolution of computers in 1990.
2. The dpi is only really descriptive when you are talking about output.
3. Dpi is meaningless as the dot is defined so many different ways that
you really have no idea what the dot is at any given time.

If you are restricted to movie format then best way to do it is to
record on mini dv format. The sony cameras have an internal hardware
compressor that is DVD compliant. It records data at 40 Megabytes per
second. This is about 10 times the data rate that "direct digital"
methods can achieve. The digital tape can record MUCH faster than a hard
drive. After recording you can bring the data into thew computer over
the IEEE 1394 (firewire) input. This is a frame accurate data transfer
that does not change the data in any way. You do not lose one bit of data.
The frames that you then can save need to be saved in a lossless manner.
I would suggest you use Adobe Premier because it is a Mac program and
you can save in a large variety of lossless formats.

Please understand that the chip that does the compression in the Sony
cameras is one of those best kept secrets. Sony developed an
unbelievably powerful chip so that they could own the mini dv format.
They won but unfortuneately for them consumers are much more interested
in BluRay and HD DVD

The documentation for publication needs to describe the acquisition
chain as you are not recording anything that is uncompressed. The MSA
guidelines that Nestor pointed you to would require that you report that
hardware/software system so that others can reproduce your results

A HD tv camera would improve things greatly but I have not had time to
run any tests to see exactly what the bandwidth of the system is. Since
you would be putting 4 times as much data onto the tape you might have
to use a chip that compresses it more than the Sony does for NTSC/PAL
format.

John

John M. Mackenzie, Jr., PhD

Professor of Microbiology
Coordinator, Center for Electron Microscopy
North Carolina State University
Phone (919) 515-2664 Fax (919) 515-8293



lamiller-at-uiuc.edu wrote:
}
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}
} I guess I should qualify the question...... I can make the image
} bigger on my screen while in iMovie.
}
}
} Would then taking a bigger image as TIFF than one can with the save
} frame command, with Grab-it be of any benefit?
}
} Such as making it smaller and increasing the dpi in photoshop
mathmatically?
}
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From: gary.m.brown-at-exxonmobil.com
Date: Mon, 21 May 2007 09:26:15 -0500
Subject: [Microscopy] Re: Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Friends,

I have gotten several off-line replies to my request for information on
cutting tools and hazards. I appreciate any and all information and
encourage replies to sent to the listserver for all to benefit. If, for
privacy concerns, you choose to reply off-line, I will summarize your
anonymous contribution with other feedback.

One last note: I will appreciate any comments, suggestions, concerns
regarding any kind of cutting or preparation tool. This in an area where
many of us have expertise that should be shared with the community at
large.

Thanks,

Gary M. Brown
Microscopy Specialist
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

"Happiness equals reality minus expectations." Tom Magliozzi



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From: frank.karl-at-degussa.com
Date: Mon, 21 May 2007 10:25:25 -0500
Subject: [Microscopy] Re: Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Gary,
You're not going to like this, the best safety tool is the operator.
Having cut, hacked, dissected and whacked tires, hoses, belts, plastic
bottles and all things rubber and polymer for over 25 years, I have only a
few "guards" for you.

Rule one: before any cut is made the operator needs to look at the sample
and ask: if the blade slips, where will it go? If the answer involves
finger or other body parts change the configuration of the cut.

Rule two: when working under a stereomicroscope take a second to take the
eyes away from the image and check the positions of the fingers, sample and
blades. See rule one.

Rule three: always use sharp edges. Edges are sharp or they are not. One
cut may dull an edge, but it's cheaper to replace an edge then pay
emergency room bills. My worse cuts have always been with a dull knife.

Rule four: Never rush. Keep the supervisor from breathing down the
operators neck, never tell a person working with a knife to hurry up.

Rule five: use the right tool for the sample. Cutting a wire reinforced
hose may require mechanical saws or embedding material and a polishing
wheel or a vice and a number 11 scalpel blade held by a pair of vice grips.

Rule six: always ask for assistance if you need it. Let someone else pull
back a ply or component while you concentrate on the cut. Helper can were
kevlar gloves and/or should use pliers, vice grips, forceps of other
mechanical devices to keep there fingers out of the way.

These rules form the basic "guard" when cutting. Have I cut myself? Yes,
usually when I break one of these rules.

Stay safe.............Frank





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From: bliss5-at-llnl.gov
Date: Mon, 21 May 2007 10:32:19 -0500
Subject: [Microscopy] electrochemical polishing

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello All:

Has anyone out there ever done electrochemical polishing on cerium? I
have consulted two or three folks here with no luck. I also have
Petzow's Metallographic Etching and a link to an online database of
etchants. (Sorry, I cannot remember the author's name offhand and I'm
pretty sure he monitors the list. Am I in trouble or what?)
--

+++++++++++++++++++++++++++++

R. Ann Bliss
Technologist, Chemistry Materials and Life Science
Materials Science and Technology Division
Lawrence Livermore National Laboratory

_____________________________

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From: henriks-at-cox.net
Date: Mon, 21 May 2007 11:34:18 -0500
Subject: [Microscopy] electrochemical polishing

Contents Retrieved from Microscopy Listserver Archives
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Ann:

The online database you are referring to is the Metallographic Etchants
Database by Henrik Kaker. It is available from South Bay Technology at

http://www.southbaytech.com/shop/med1.shtml.

I hope this helps.

Best regards-

David

David Henriks
henriks-at-cox.net
cell: 949-533-9100

-----Original Message-----
X-from: bliss5-at-llnl.gov [mailto:bliss5-at-llnl.gov]
Sent: Monday, May 21, 2007 8:38 AM
To: Henriks-at-cox.net

Hello All:

Has anyone out there ever done electrochemical polishing on cerium? I
have consulted two or three folks here with no luck. I also have
Petzow's Metallographic Etching and a link to an online database of
etchants. (Sorry, I cannot remember the author's name offhand and I'm
pretty sure he monitors the list. Am I in trouble or what?)
--

+++++++++++++++++++++++++++++

R. Ann Bliss
Technologist, Chemistry Materials and Life Science
Materials Science and Technology Division
Lawrence Livermore National Laboratory

_____________________________

==============================Original Headers==============================
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==============================Original Headers==============================
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From: Henrik.Kaker-at-guest.arnes.si
Date: Mon, 21 May 2007 13:50:27 -0500
Subject: [Microscopy] electrochemical polishing

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Ann,

Please the link http://www.kaker.com/etch/demo/index.html.

Henrik

henriks-at-cox.net wrote:
} ----------------------------------------------------------------------------
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} ----------------------------------------------------------------------------
}
} Ann:
}
} The online database you are referring to is the Metallographic Etchants
} Database by Henrik Kaker. It is available from South Bay Technology at
}
} http://www.southbaytech.com/shop/med1.shtml.
}
} I hope this helps.
}
} Best regards-
}
} David
}
} David Henriks
} henriks-at-cox.net
} cell: 949-533-9100
}
} -----Original Message-----
} X-from: bliss5-at-llnl.gov [mailto:bliss5-at-llnl.gov]
} Sent: Monday, May 21, 2007 8:38 AM
} To: Henriks-at-cox.net
} Subject: [Microscopy] electrochemical polishing
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hello All:
}
} Has anyone out there ever done electrochemical polishing on cerium? I
} have consulted two or three folks here with no luck. I also have
} Petzow's Metallographic Etching and a link to an online database of
} etchants. (Sorry, I cannot remember the author's name offhand and I'm
} pretty sure he monitors the list. Am I in trouble or what?)
}


--
Henrik Kaker Ph.D.
Metal Ravne d.o.o.
SEM-EDS Laboratory
Koroska cesta 14
SI-2390 Ravne
Slovenia
Phone: +386 2 870 7076
GSM: +386 31 380 875
http://www2.arnes.si/~sgszmera1/index.html


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From: c.r.vandenbrom-at-bham.ac.uk
Date: Mon, 21 May 2007 16:46:58 -0500
Subject: [Microscopy] Supporting lipid bilayers with holey carbon?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear all,

A few of my colleagues are trying to study transport behaviour of lipid bilayer membranes. This requires a membrane to be deposited over a (tiny) hole between two aqeous compartments. The (in)stability of the suspended part of such a membrane is a major complication. I am wondering whether a support technique from the microscopy field might be helpful here. In particular I am curious about the possibility of using a holey carbon-like approach. Not being a holey-carbon expert myself, I would much appreciate if anyone could answer the questions underneath or provide similar or alternative suggestions.

1. What materials are typically used to make holey or lacey films. Just amorphous carbon and formvar?

2. What are the typical dimensions attainable for holey or lacey films, in terms of film thickness and hole size. For our purposes, 20-40 nm hole diameter would be ideal, but 100 - 200 nm would be very helpful already.

3. Are there good descriptions of the preparation or availability of such holey films?

4. Does anyone have experience in supporting lipid bilayers in different ways, e.g. for TEM investigation?

Thank you all for your help!


With kind regards,

Coen van den Brom



----------------------------

dr CR van den Brom
Department of Nanoscale Chemistry
School of Chemistry
University of Birmingham
Edgbaston
Birmingham B15 2TT
United Kingdom

T: +44 121 4144426
E: c.r.vandenbrom-at-bham.ac.uk



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From: mpease-at-jhmi.edu
Date: Mon, 21 May 2007 18:19:38 -0500
Subject: [Microscopy] viaWWW: monkey perfusion methodology

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Email: mpease-at-jhmi.edu
Name: Mary Ellen Pease

Organization: Johns Hopkins Wilmer Eye Institute

Title-Subject: [Filtered] monkey perfusion methodology

Question: What is the recommended flow rate for perfusion of monkey brain and eyes when using a pump rather than gravity feed of the solutions?

Thank you,
Mary Ellen

---------------------------------------------------------------------------

==============================Original Headers==============================
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From: jrminter-at-rochester.rr.com
Date: Mon, 21 May 2007 20:33:21 -0500
Subject: [Microscopy] RE: Supporting lipid bilayers with holey carbon?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Coen van den Brom,

I have made holey carbon films using various recipes. I much prefer the
commercial "C-Flat" films by ProtoChips.

I have no interest in ProtoChips, other than as a satisfied customer.

John Minter


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From: rcmoretz-at-gmail.com
Date: Mon, 21 May 2007 21:28:04 -0500
Subject: [Microscopy] Cutting tools and hazards

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One very important rule that I always check: make sure the sharp side
is down! Sounds stupid, but I can't tell you how many times people
have cut themselves because the sharp edge was up! So, to prevent
such disasters, I always ask "Is the sharp side down?" Maybe it's
just that I'm getting older and catch myself every now and then with
the sharp side up. And I will strongly second the rule about not
rushing or allowing other distraction (conversations, etc)--when I
have hurt myself the worst it has been someone standing there
yammering at me.

Roger Moretz

On 5/21/07, frank.karl-at-degussa.com {frank.karl-at-degussa.com} wrote:
}
}
}
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} ----------------------------------------------------------------------------
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} Gary,
} You're not going to like this, the best safety tool is the operator.
} Having cut, hacked, dissected and whacked tires, hoses, belts, plastic
} bottles and all things rubber and polymer for over 25 years, I have only a
} few "guards" for you.
}
} Rule one: before any cut is made the operator needs to look at the sample
} and ask: if the blade slips, where will it go? If the answer involves
} finger or other body parts change the configuration of the cut.
}
} Rule two: when working under a stereomicroscope take a second to take the
} eyes away from the image and check the positions of the fingers, sample and
} blades. See rule one.
}
} Rule three: always use sharp edges. Edges are sharp or they are not. One
} cut may dull an edge, but it's cheaper to replace an edge then pay
} emergency room bills. My worse cuts have always been with a dull knife.
}
} Rule four: Never rush. Keep the supervisor from breathing down the
} operators neck, never tell a person working with a knife to hurry up.
}
} Rule five: use the right tool for the sample. Cutting a wire reinforced
} hose may require mechanical saws or embedding material and a polishing
} wheel or a vice and a number 11 scalpel blade held by a pair of vice grips.
}
} Rule six: always ask for assistance if you need it. Let someone else pull
} back a ply or component while you concentrate on the cut. Helper can were
} kevlar gloves and/or should use pliers, vice grips, forceps of other
} mechanical devices to keep there fingers out of the way.
}
} These rules form the basic "guard" when cutting. Have I cut myself? Yes,
} usually when I break one of these rules.
}
} Stay safe.............Frank
}
}
}
}
}
} ==============================Original Headers==============================
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3, 28 -- To: frank.karl-at-degussa.com, "Microscopy Listserv" {Microscopy-at-microscopy.com}
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From: tina-at-pbrc.hawaii.edu
Date: Mon, 21 May 2007 22:24:21 -0500
Subject: [Microscopy] Arabidopsis GFP in TEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi, All-

I am participating in a workshop (again) where they want me to train 14
people to find GFP in Arabidopsis in both confocal and TEM. I have tried
fixing young leaves and cotyledons (cut into {1mm strips) in 2%
paraformaldehyde/0.1% glutaraldehyde in 0.1M Na cacodylate with 2mM CaCl2,
washing, dehydrating, embedding in LR White, polymerized at 50C. I'm
getting holes in the resin (I didn't particularly rush through
dehydration, but I can prolong it, and I did not pull a vacuum, but I
can), and I'm not seeing any GFP in 1 um sections in the confocal. I am
reluctant to keep going with these samples and then try to do anti-GFP for
TEM if I'm not seeing GFP (but the organizers of the workshop don't seem
to care, as long as we go through the motions)(Sigh). I would be grateful
for any tips about any step along the way from anybody!

Mahalo!
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************


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From: cgarber-at-2spi.com
Date: Tue, 22 May 2007 02:20:20 -0500
Subject: [Microscopy] Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Roger Moretz wrote:
===============================================================
One very important rule that I always check: make sure the sharp side
is down! Sounds stupid, but I can't tell you how many times people
have cut themselves because the sharp edge was up! So, to prevent
such disasters, I always ask "Is the sharp side down?" Maybe it's
just that I'm getting older and catch myself every now and then with
the sharp side up. And I will strongly second the rule about not
rushing or allowing other distraction (conversations, etc)--when I
have hurt myself the worst it has been someone standing there
yammering at me.
===============================================================
In our laboratory and production facilities, we do not permit the use of (the cheaper) double sided razor blades because
of the reasons cited above. A single edge razor blade (we prefer the GEM type, see URL
http://www.2spi.com/catalog/tools/smtol14.shtml is all that is permitted.

People would probably be amazed at the number of eye injuries that occur (not necessarily in EM labs) when one uses a
razor (or scalpel) blade (of any type) not so much for cutting but for "lifting" or "prying", now the forces are not
within the plane of the blade but are perpendicular to it, and as a result of the shear stresses against which the blade
was never designed, a corner of the blade can easily "fly" off at high speed and injure the worker if not wearing eye
protection. This is also a good reason why eye protection should be worn by anyone in the vicinity of anyone using any
kind of a razor blade.

Over the years, we have been asked to do quite a number of failure analysis studies and reports on such blade fragments
that caused eye injury (or even blindness). I don't recall that we have ever found a "material defect" in such studies,
but there was apparently more than enough evidence to suggest that the user was improperly using the blade (in shear).

I also agree with Roger that idle "chit-chat" can be hazardous when one is trying to do work with potentially dangerous
tools at the same time.

Disclaimer: SPI Supplies distributes the GEM single edge "scientific" razor blades so we would have a vested interest
in seeing more of our customers using this type of blade.

Chuck

===================================================
Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
President
SPI SUPPLIES FAX: 1-(610)-436-5755
PO BOX 656 e-mail: cgarber-at-2spi.com
West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com


Look for us!
############################
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############################
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From: nizets2-at-yahoo.com
Date: Tue, 22 May 2007 04:46:12 -0500
Subject: [Microscopy] basic SEM: imaging salt crystals

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear listers,

Here are some basic questions from the basic SEM user
I am: I would like to visualize salt crystals by SEM
and make EDX analysis if possible. I just leave a drop
of salt solution on a carbon tab dry, that's all!
My questions:

- Do I need to carbon-coat the crystals?
- Is there a risk to evaporate the crystals under the
electron beam and make the column dirty? (I usually
don't use HV higher than 15 kV). Would carbon coating
help dissipate the heat in this regard?

Now another question related to SEM but not to
crystals:
When I analyse fine particles by EDX, I never know if
I analyse only the particle or also part of the
material which is behind the particle. Is there a way
to determine the depth of analysis of the material
studied? I suppose that it depends on the HV and on
the nature of the material itself, but I already would
be happy to have a order of magnitude. Is this 10 nm
or 2 µm deep?
Lets say I analyze at 15kV (1) NaCl crystals (2)
Quartz (3) Feldpar (4) Copper, what would be the
expected "depth of analysis" in these samples?
Does it depend on the size of the beam?

Best regards,

Stephane








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From: nizets2-at-yahoo.com
Date: Tue, 22 May 2007 07:09:31 -0500
Subject: [Microscopy] Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I completely share this opinion.
Seriousness and concentration are not an art, it is
available to anybody wanting them.
Correct training is also most important.
I met a girl who embedded her Epon in eppendorf tubes.
Then she took the tube in one hand (bare hands!) and
cut through the plastic with the other hand using a
razor blade! I mean, you don't need to be a genius to
expect an accident one day or another.

I don't cut blocks for 25 years, but still for 15 good
years and I never had the smallest accident. I don't
think it is luck.
Now I understand that one cannot always control what
others do. In this case ask the person to wear
plexiglas goggles, bullet-proof vest, kevlar gloves
and there is still the risk that he cuts his ear!

Frank's rules are excellent, but most of them could be
all replaced by one only rule: use your common sense.
Rules 3 and 4 are critical and should be part of a
correct training.

Best regards,
Stephane



--- frank.karl-at-degussa.com wrote:

}
}
}
}
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}
} Gary,
} You're not going to like this, the best safety tool
} is the operator.
} Having cut, hacked, dissected and whacked tires,
} hoses, belts, plastic
} bottles and all things rubber and polymer for over
} 25 years, I have only a
} few "guards" for you.
}
} Rule one: before any cut is made the operator needs
} to look at the sample
} and ask: if the blade slips, where will it go? If
} the answer involves
} finger or other body parts change the configuration
} of the cut.
}
} Rule two: when working under a stereomicroscope take
} a second to take the
} eyes away from the image and check the positions of
} the fingers, sample and
} blades. See rule one.
}
} Rule three: always use sharp edges. Edges are sharp
} or they are not. One
} cut may dull an edge, but it's cheaper to replace an
} edge then pay
} emergency room bills. My worse cuts have always been
} with a dull knife.
}
} Rule four: Never rush. Keep the supervisor from
} breathing down the
} operators neck, never tell a person working with a
} knife to hurry up.
}
} Rule five: use the right tool for the sample.
} Cutting a wire reinforced
} hose may require mechanical saws or embedding
} material and a polishing
} wheel or a vice and a number 11 scalpel blade held
} by a pair of vice grips.
}
} Rule six: always ask for assistance if you need it.
} Let someone else pull
} back a ply or component while you concentrate on the
} cut. Helper can were
} kevlar gloves and/or should use pliers, vice grips,
} forceps of other
} mechanical devices to keep there fingers out of the
} way.
}
} These rules form the basic "guard" when cutting.
} Have I cut myself? Yes,
} usually when I break one of these rules.
}
} Stay safe.............Frank
}
}
}
}
}
} ==============================Original
} Headers==============================
} 13, 18 -- From frank.karl-at-degussa.com Mon May 21
} 10:25:25 2007
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} and hazards
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} microscopy-at-msa.microscopy.com
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From: Walter.Bobrowski-at-pfizer.com
Date: Tue, 22 May 2007 07:28:03 -0500
Subject: [Microscopy] viaWWW: monkey perfusion methodology

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

300 ml/min. Perfuse either through the carotid artery or left ventricle
(occlude descending aorta).

Best regards,
Walter F. Bobrowski
Sr. Scientist
Pfizer Global R&D
2800 Plymouth Rd.
Ann Arbor, MI 48105

TEL: 734-622-7814
FAX: 734-622-5718

"Like the strength of a steel rod, the true character of a person can
only be known under extreme stress." - Leslie Fieger




-----Original Message-----
X-from: mpease-at-jhmi.edu [mailto:mpease-at-jhmi.edu]
Sent: Monday, May 21, 2007 7:25 PM
To: Bobrowski, Walter

This Question/Comment was submitted to the Microscopy Listserver
using the WWW based Form at
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Email: mpease-at-jhmi.edu
Name: Mary Ellen Pease

Organization: Johns Hopkins Wilmer Eye Institute

Title-Subject: [Filtered] monkey perfusion methodology

Question: What is the recommended flow rate for perfusion of monkey
brain and eyes when using a pump rather than gravity feed of the
solutions?

Thank you,
Mary Ellen

------------------------------------------------------------------------
---

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Headers==============================
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From: dsherman-at-purdue.edu
Date: Tue, 22 May 2007 07:35:10 -0500
Subject: [Microscopy] Re: Arabidopsis GFP in TEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Tina,

Can you screen for GFP by fluorescence after fixation, then dehydration, and
finally embedding to see where the signal is lost?

You may have better luck using a water soluble resin such as Durcupan. I
have not tried this resin for ICC but it is relatively soft so may work.
Advantage is that you do not have to dehydrate in ETOH or acetone and this
may help preserve the GFP signal.

Debby

Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907



} From: {tina-at-pbrc.hawaii.edu}
} Reply-To: {tina-at-pbrc.hawaii.edu}
} Date: Mon, 21 May 2007 22:26:18 -0500
} To: {dsherman-at-purdue.edu}
} Subject: [Microscopy] Arabidopsis GFP in TEM
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------------
}
} Hi, All-
}
} I am participating in a workshop (again) where they want me to train 14
} people to find GFP in Arabidopsis in both confocal and TEM. I have tried
} fixing young leaves and cotyledons (cut into {1mm strips) in 2%
} paraformaldehyde/0.1% glutaraldehyde in 0.1M Na cacodylate with 2mM CaCl2,
} washing, dehydrating, embedding in LR White, polymerized at 50C. I'm
} getting holes in the resin (I didn't particularly rush through
} dehydration, but I can prolong it, and I did not pull a vacuum, but I
} can), and I'm not seeing any GFP in 1 um sections in the confocal. I am
} reluctant to keep going with these samples and then try to do anti-GFP for
} TEM if I'm not seeing GFP (but the organizers of the workshop don't seem
} to care, as long as we go through the motions)(Sigh). I would be grateful
} for any tips about any step along the way from anybody!
}
} Mahalo!
} Tina
}
} ****************************************************************************
} * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
} * Biological Electron Microscope Facility * (808) 956-6251 *
} * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
} ****************************************************************************
}
}
} ==============================Original Headers==============================
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9, 23 -- Subject: Re: [Microscopy] Arabidopsis GFP in TEM
9, 23 -- From: Debby Sherman {dsherman-at-purdue.edu}
9, 23 -- To: Tina Carvalho {tina-at-pbrc.hawaii.edu} ,
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From: jquinn-at-www.matscieng.sunysb.edu
Date: Tue, 22 May 2007 09:30:23 -0500
Subject: [Microscopy] re: SEM and EDAX

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Stephane

0) Purchase a copy of Electron Flight Simulator
or download a copy of WinXray (from McGill).
You should run simulations to estimate your
excitation volume.

1) Personally, I would not put a drop of liquid
on carbon tabs or double sided tape. It is
bad practice, as the water does interact with
the adhesive.

Instead, use a silicon wafer (you can purchase
precut wafers), carbon planchet, Be planchet, etc....

2) If the substrates are conducting (as are the ones
mentioned), then you will not need much overcoating.
Personally, a few nm of gold on an incline sample
will do the trick.

regards,

Jim

PS: OoO away...............



} From mail-at-ns.microscopy.com Tue May 22 05:45:52 2007
} Return-Path: {mail-at-ns.microscopy.com}
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} Date: Tue, 22 May 2007 04:47:09 -0500
} Message-Id: {200705220947.l4M9l9uH018525-at-ns.microscopy.com}
} To: jquinn-at-www.matscieng.sunysb.edu
} From: nizets2-at-yahoo.com
} Reply-to: nizets2-at-yahoo.com
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
} Subject: [Microscopy] basic SEM: imaging salt crystals
} Errors-To: MicroscopyListSpamFilter-at-microscopy.com
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}
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} Dear listers,
}
} Here are some basic questions from the basic SEM user
} I am: I would like to visualize salt crystals by SEM
} and make EDX analysis if possible. I just leave a drop
} of salt solution on a carbon tab dry, that's all!
} My questions:
}
} - Do I need to carbon-coat the crystals?
} - Is there a risk to evaporate the crystals under the
} electron beam and make the column dirty? (I usually
} don't use HV higher than 15 kV). Would carbon coating
} help dissipate the heat in this regard?
}
} Now another question related to SEM but not to
} crystals:
} When I analyse fine particles by EDX, I never know if
} I analyse only the particle or also part of the
} material which is behind the particle. Is there a way
} to determine the depth of analysis of the material
} studied? I suppose that it depends on the HV and on
} the nature of the material itself, but I already would
} be happy to have a order of magnitude. Is this 10 nm
} or 2 µm deep?
} Lets say I analyze at 15kV (1) NaCl crystals (2)
} Quartz (3) Feldpar (4) Copper, what would be the
} expected "depth of analysis" in these samples?
} Does it depend on the size of the beam?
}
} Best regards,
}
} Stephane
}
}
}
}
}
}
}
}
} ____________________________________________________________________________________
} Need Mail bonding?
} Go to the Yahoo! Mail Q&A for great tips from Yahoo! Answers users.
} http://answers.yahoo.com/dir/?link=list&sid=396546091
}
} ==============================Original Headers==============================
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} 13, 19 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} 13, 19 -- Subject: basic SEM: imaging salt crystals
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} From mail-at-ns.microscopy.com Tue May 22 05:45:52 2007
} Date: Tue, 22 May 2007 04:47:09 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: nizets2-at-yahoo.com
} Reply-to: nizets2-at-yahoo.com
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
} Subject: [Microscopy] basic SEM: imaging salt crystals
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}
}
}
}
} ----------------------------------------------------------------------------
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} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Dear listers,
}
} Here are some basic questions from the basic SEM user
} I am: I would like to visualize salt crystals by SEM
} and make EDX analysis if possible. I just leave a drop
} of salt solution on a carbon tab dry, that's all!
} My questions:
}
} - Do I need to carbon-coat the crystals?
} - Is there a risk to evaporate the crystals under the
} electron beam and make the column dirty? (I usually
} don't use HV higher than 15 kV). Would carbon coating
} help dissipate the heat in this regard?
}
} Now another question related to SEM but not to
} crystals:
} When I analyse fine particles by EDX, I never know if
} I analyse only the particle or also part of the
} material which is behind the particle. Is there a way
} to determine the depth of analysis of the material
} studied? I suppose that it depends on the HV and on
} the nature of the material itself, but I already would
} be happy to have a order of magnitude. Is this 10 nm
} or 2 µm deep?
} Lets say I analyze at 15kV (1) NaCl crystals (2)
} Quartz (3) Feldpar (4) Copper, what would be the
} expected "depth of analysis" in these samples?
} Does it depend on the size of the beam?
}
} Best regards,
}
} Stephane
}
}
}
}
}
}
}
}
} ____________________________________________________________________________________
} Need Mail bonding?
} Go to the Yahoo! Mail Q&A for great tips from Yahoo! Answers users.
} http://answers.yahoo.com/dir/?link=list&sid=396546091
}
} ==============================Original Headers==============================
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} 13, 19 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} 13, 19 -- Subject: basic SEM: imaging salt crystals
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}

==============================Original Headers==============================
13, 12 -- From jquinn-at-www.matscieng.sunysb.edu Tue May 22 09:30:23 2007
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13, 12 -- Date: Tue, 22 May 2007 10:28:58 -0400
13, 12 -- From: Jim Quinn {jquinn-at-www.matscieng.sunysb.edu}
13, 12 -- Message-Id: {200705221428.l4MESwq17328-at-www.matscieng.sunysb.edu}
13, 12 -- To: microscopy-at-microscopy.com
13, 12 -- Subject: re: SEM and EDAX
==============================End of - Headers==============================




From: gcouger-at-science-info.net
Date: Tue, 22 May 2007 11:40:17 -0500
Subject: [Microscopy] Re: Cutting tools and hazards

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

cgarber-at-2spi.com wrote:
} {snip}
}
} I also agree with Roger that idle "chit-chat" can be hazardous when one is trying to do work with potentially dangerous tools at the same time.
}
} Disclaimer: SPI Supplies distributes the GEM single edge "scientific" razor blades so we would have a vested interestin seeing more of our customers using this type of blade.
}
} Chuck
}
} ===================================================
} Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
} President
} SPI SUPPLIES FAX: 1-(610)-436-5755
} PO BOX 656 e-mail: cgarber-at-2spi.com
} West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
}
} WWW: http://www.2spi.com
}
You are the key to your own safety. Not the safety officer, not the
rules in the lab, not the no slip floors, no spill containers, hoods or
anything else in the end your safety rests in on you. Nothing can make
you safe from you if your not safe in the way you do things. Just take
the case of Lucky. He a man that has cut off the fingers of one hand,
ruined the joints in the other arm , his back and more. He managed to
blow up two guns reloading ammunition and runs his boat on the rocks
nearly every time he goes out. Of all the fishing rods I have broken
Lucky stepped on all 3. Nothing can be made safe for Lucky to use. I
call him Lucky because he turned his head just a bolt snapped off a
machine he was abusing and knocked out all his front teeth. Had he not
turned around to look at the machine the bolt would have hit the base of
his scull and killed him. I don't get close to Lucky when he is working
or hunting and we fish in my boat.

When cutting proper tools for the job make it safer and easier. I have
too many tools. I am obsessive about tools being sharp and properly
shaped for the job. But I don't have many scars on my hands from knives.
I do use double edged razor blade cut into 4 pieces an mounted in a
handle for some things as described by Anna Teetsov at McCrone
Associates in "Quarter-Razor Blade for Hand Sectioning" in the Prep
School Section of Modern Microscopy
www.modernmicroscopy.com/main.asp?article=50
{http://www.modernmicroscopy.com/main.asp?article=50} . I sure wouldn't
use them with out a handle. I also find Schick razor blades useful in a
handle thought somewhat expensive and hard to find. I sure wouldn't use
either one with out a good handle on it.

In 30 years of working at hazardous work, farming, machine shops,
welding and such before coming into a slightly safer lab environment
every injury I was involved with was from not paying attention to what I
was doing. Trying to hurry was the worst offender. Safety is in the
person doing the work. Taking my time and planing ahead have got me
though life with all my fingers and the only broken bone is my nose when
I came out of the bath room and tripped over the kids dog we were
keeping for them while they went to China 2 years ago.


The only serious accident I have been involved in was when I was taking
a neighbor with brain damaged to the brain stem to a meeting and he
fell and broke his pelvis as he got out of the car. It was years after I
retired and we were talking and I failed to stand behind him as I
normally do with anyone that has trouble with getting in and out of a
car so I can catch them should they fall. I use a cane when I have that
much trouble getting around and that's 2 or 3 times a year but many
people let pride get in the way of good sense on things like that. Pride
has no place where safety is concerned. Anything that make you less safe
needs to be dealt with once an for all. If it is miss placed pride,
contempt for a naive safety officers, cultural custom, urgency or
anything that distracts you from the the task at hand you need to deal
with it now.

No one can make you safe but you.

Gordon
Gordon Couger
Microscope documentation Archive http://science-info.net/


==============================Original Headers==============================
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From: tivol-at-caltech.edu
Date: Tue, 22 May 2007 12:03:13 -0500
Subject: [Microscopy] Re: Supporting lipid bilayers with holey carbon?

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On May 21, 2007, at 2:47 PM, c.r.vandenbrom-at-bham.ac.uk wrote:

} 1. What materials are typically used to make holey or lacey films.
} Just amorphous carbon and formvar?
}
} 2. What are the typical dimensions attainable for holey or lacey
} films, in terms of film thickness and hole size. For our purposes,
} 20-40 nm hole diameter would be ideal, but 100 - 200 nm would be very
} helpful already.
}
} 3. Are there good descriptions of the preparation or availability of
} such holey films?
}
} 4. Does anyone have experience in supporting lipid bilayers in
} different ways, e.g. for TEM investigation?
}
Dear Coen,
1. Typically holey formvar is produced, carbon is evaporated onto it,
and then the formvar is dissolved away, leaving a holey carbon support
film. This is unsuitable for supporting lipid bilayers for at least
two reasons; first, the carbon will not be hydrophobic enough that
lipid would wet it, and second, the areas of the holes are not
well-defined enough to measure ion current as a function of bilayer
area accurately. I know of techniques that can change the nature of
the carbon--glow-discharging to increase hydrophyllicity, and letting
the carbon age to increase hydrophobicity--but I do not think that the
carbon ever gets truly hydrophobic. Once one has a holey formvar film,
one can either leave it uncoated or evaporate or otherwise coat it with
a variety of substances other than carbon, so perhaps a suitable
hydrophobic coating can be found.
2. One can vary the hole size during preparation. Generally, one
suspends hydrophyllic droplets in the formvar, so that when the formvar
dries, the droplets leave holes where the formvar has been excluded.
This results in holes of about the size of the droplets, which are
typically of um dimension, not nm. However, holes can be produced by
irradiation, so perhaps holes in your desired size range are possible.
3. Yes, but I don't have one at hand.
4. I have just dissolved the lipid in a suitable solvent, put a drop
across a hole in a teflon wafer, let the dissolved lipid spread until a
bilayer formed across the hole, and lowered the wafer into the water so
that the wafer served to separate the two sides of a compartment or
added water to both sides of the compartment after the wafer was in
place. This way the area of the bilayer is known, and one could use a
wafer with many holes of the desired total area. There are likely to
be companies that can construct plastic wafers with appropriately sized
holes, and maybe one or more of them will contact you through the list.
Good luck.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu


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From: tivol-at-caltech.edu
Date: Tue, 22 May 2007 12:31:53 -0500
Subject: [Microscopy] Re: basic SEM: imaging salt crystals

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On May 22, 2007, at 2:46 AM, nizets2-at-yahoo.com wrote:

} Here are some basic questions from the basic SEM user
} I am: I would like to visualize salt crystals by SEM
} and make EDX analysis if possible. I just leave a drop
} of salt solution on a carbon tab dry, that's all!
} My questions:
}
} - Do I need to carbon-coat the crystals?
} - Is there a risk to evaporate the crystals under the
} electron beam and make the column dirty? (I usually
} don't use HV higher than 15 kV). Would carbon coating
} help dissipate the heat in this regard?
}
} Now another question related to SEM but not to
} crystals:
} When I analyse fine particles by EDX, I never know if
} I analyse only the particle or also part of the
} material which is behind the particle. Is there a way
} to determine the depth of analysis of the material
} studied? I suppose that it depends on the HV and on
} the nature of the material itself, but I already would
} be happy to have a order of magnitude. Is this 10 nm
} or 2 µm deep?
} Lets say I analyze at 15kV (1) NaCl crystals (2)
} Quartz (3) Feldpar (4) Copper, what would be the
} expected "depth of analysis" in these samples?
} Does it depend on the size of the beam?
}
Dear Stephane,
If the crystals are conductive, you do not need to coat them, and if
they are not, you could examine them at a voltage for which the
production and escape of secondary electrons is the same as the beam
electrons deposited in the crystal, giving no net change in charge.
This might not be sufficient voltage to generate particular X-ray
lines, however. Heat conductivity is generally high in substances for
which charge conductivity is high, and for substances with low heat
conductivity, using a low beam current with longer times of analysis
can ameliorate specimen heating, so I would advise against carbon
coating. In any case, carbon coating will not help with heat generated
too far below the surface of the sample, but this may not be an issue.
David Joy has written programs to determine the analysis volumes for
various substances using electrons of various energies, and I think
that these are freely available. For 15 kV or below, the depth would
be closer to 10 nm than to 2 um--I don't have access to my tables of
electron stopping powers, so I can't give you a better answer. The
depth does not depend strongly on the size of the beam; each electron
acts independently. If, however, excess charge is not dissipated, and
the extent of the charged volume is larger and the charge per unit
volume is smaller for a wider, but less intense beam, there may be a
difference in the voltage at the surface of the specimen that could
lead to changes in the depth of analysis. In the absence of charging
effects--which will obviously be different for quartz than for
copper--the penetration depth of electrons is approximately the same
when measured in units of mg/cm^2 for all materials. It is greater in
higher-Z materials, since they have a higher neutron/proton ratio,
which adds mass but not stopping power.
Yours,
Bill Tivol, PhD
EM Scientist
Electron Cryo-Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu



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From: nholson-at-ucsd.edu
Date: Tue, 22 May 2007 13:16:28 -0500
Subject: [Microscopy] Supporting lipid bilayers with holey carbon?

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--
Coen: Try this as a technique for making holey carbon films:
http://cryoem.ucsd.edu/procedures/H-carbonfilm.shtm

I wouldn't call it the final and absolute way to do it but it is the
one we teach to our students and it works well. The holes we get are
of varying size from less than a micron in diameter on up to several
microns. If you want the smallest holes possible I would suggest
sonnicating for a much longer time than suggested in the procedure.

Norm

______________________________________________________________
Norm Olson
Cryoelectron Microscopy Facilities Manager
1510 Bonner Hall
Department of Chemistry & Biochemistry, MC-0378
University of California San Diego
La Jolla, CA 92093-0378
nholson-at-ucsd.edu
http://cryoem.ucsd.edu
Cell: (858)220-2183
(858)822-6718 - Office; (858)534-5846 - Fax
______________________________________________________________



3. Are there good descriptions of the preparation or availability of
such holey films?

Thank you all for your help!


With kind regards,

Coen van den Brom



----------------------------

dr CR van den Brom
Department of Nanoscale Chemistry
School of Chemistry
University of Birmingham
Edgbaston
Birmingham B15 2TT
United Kingdom

T: +44 121 4144426
E: c.r.vandenbrom-at-bham.ac.uk


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From: gary-at-gaugler.com
Date: Tue, 22 May 2007 13:26:37 -0500
Subject: [Microscopy] Re: basic SEM: imaging salt crystals

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Why a salt solution? You can just drop salt
crystals onto a sticky tab on a stub and image
a bunch of crystals. Different salts look
different. Table salt is like a face centered
cubic while marine salt is layered as is Kosher
salt.

For EDS, the Cl shells are the ones to get
set up for. Na Ka is at 1.041KeV while Cl Ka
is at 2.621KeV. The only other transitions
are K beta. Thus, about 6-8KV should be enough
KV to get all peaks and do quant. To get just
the/a crystal, do spot mode and put the spot
on a crystal.

I would coat the crystals with Au/Pd or Pt or Pd.
Au is probably best since its La peak is at 9.710KeV
and its Ma 2.120KeV which puts that peak in between
Na Ka and Cl Ka. I would not coat with C since
if your beam is off of the crystal, you would not
necessarily know if the C is from the coating or
the sticky tab. It probably would be a giveaway
since the tab C peak would be higher (much) than
the coating C peak. But just eliminate this issue
by not using C.

With coating and relatively low KV, you can get nice
surface morphology. Counts will probably be low
but if beam position is stable, just collect for
enough time for good results.

The Electron Flight Simulator will tell you how
deep a particular KV beam will penetrate into a
specified material.

gary g.


At 01:47 AM 5/22/2007, you wrote:
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From: hopechen-at-u.washington.edu
Date: Tue, 22 May 2007 21:30:03 -0500
Subject: [Microscopy] AskAMicroscopist:High Speed microscope illumination

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I have already seen a number of good answers, but I will chime in anyway.

I would suggest coating the samples unless you are operating in variable pressure mode (which may not be an option for you). I recall that NaCl charges as well as most any insulator. I don't imagine you would be doing quant on the sample since the lack of a known geometry would lead to substantial uncertainty in the results. Gary made a good point that it would be nice to use an element for coating that is only present in the coating. Otherwise, there will be uncertainty in the source of the peaks. I would expect some difference in the relative peak intensities depending on which element is used.

I wouldn't worry much about vaporizing the salt. (Maybe I should.) I don't expect you will be using nanoamps of current. Once again, VP mode would help minimize or prevent contamination.

I don't have the latest and greatest Monte Carlo programs. I do have an old one derived from David Joy's work. I ran a simulation on S at 15 kV and I estimate the excitation depth to be slightly less than 1 um. Running the same simulation for Na, I found the depth to be more like 2 um. For Cu, I found the depth to be on the order of 0.3 um. Therefore, I generally quote penetration depths of hundreds of nm up to a couple of microns. The dimensions of the interaction volume dwarf the spot size so much that I don't often bother considering it in calculating interaction volume.

As a fun little exercise, you might lay down a gold layer on copper or some other metal. The interaction volume for gold appears to be slightly less than 0.2 um (200 nm). Since we routinely coat samples with 10 nm of gold, we need a deeper penetration depth if we are ever going to see the sample through the gold. We don't have a problem seeing the sample unless we are using a very thick layer of gold or a low voltage. A 3 kV beam would have difficulty penetrating 10 nm of gold, but there would not be much excitation of sample x-rays at that low a voltage anyway.

Warren

-----Original Message-----
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Sent: Tuesday, May 22, 2007 4:47 AM
To: wesaia-at-iastate.edu

This Question was submitted to Ask-A-Microscopist by (hopechen-at-u.washington.edu)
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Email: hopechen-at-u.washington.edu
Name: Hope Chen

Organization: University of Washington

Education: Graduate College

Location: Seattle. WA US

Title: microscope illumination

Question: Dear Microscopist,

I try to set up a high-speed microphotography system using an ultra-high speed camera and an inverted microscope. I'm now facing a problem that is how to incorporate flash tube light source with the microscope. The light source we has is Photogenic Radio Powerlight 2500DRUV and the flash tube is Photogenic C4-19 Flashtube. The flash tube is annular with an outside diameter about 1.8 inch and inner diameter about 1 inch. Our requirements to the microscope illumination system are: 1) collect as much light as possible to increase the illumination light intensity, which hopefully could allow us to adjust our high-speed camera's exposure time to as small as 10 ns, and 2) help us to get high resolution images.
Iím thinking of using Kohler illumination, but due to the light source is a large annular light source I couldnít get idea Kohler illumination. Nelsonian illumination focus light to the specimen, so this illumination method might be better than Kohler illumination in terms of light intensity, but then I need to focus an annular light to a point.
May I ask for your suggestions?

send with thanks,
Hope

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From: mthrapp-at-grapevinegroup.com
Date: Tue, 22 May 2007 21:31:28 -0500
Subject: [Microscopy] viaWWW: Polymer Research Scientist, Microscopy Job Opportunity

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Email: mthrapp-at-grapevinegroup.com
Name: Mark Thrapp

Organization: The Grapevine Group

Title-Subject: [Filtered] Polymer Research Scientist, Microscopy Job Opportunity

Question: Research Scientist, Microscopy

The Microscopy Scientist is responsible for operating the laboratory; preparing and analyzing samples for Transmission Electron Microscopy (TEM), Scanning Electron Microscopy (SEM), and Light Microscopy (LM); and performing TEM/SEM/LM imaging of materials in support of Research & Development, Manufacturing, and Technical Service projects.

The individual will be interfacing with customers, providing insights into a variety of polymer issues (i.e., surface adhesion, morphology, fracture analysis, dispersion, etc.) Communication skills are extremely important.

Job Specifications
Education

ï Graduate degree in Polymer Science, Materials Science, Chemistry or related field of study. (PhD preferred). Bachelors is acceptable, but only those with 3+ years of definitively focused microscopy/polymer research background will be considered.

Minimum Experience
ï 3 years of related laboratory research experience.

Minimum Field-of-Expertise
ï Experience in the plastics/chemicals industry strongly preferred.
ï Operation and maintenance of TEM, SEM, and LM.

More information may be obtained by contacting Mark Thrapp, mthrapp-at-grapevinegroup.com, 404-365-9796.


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From: gcouger-at-science-info.net
Date: Wed, 23 May 2007 01:23:33 -0500
Subject: [Microscopy] Re: AskAMicroscopist:High Speed microscope illumination

Contents Retrieved from Microscopy Listserver Archives
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Dear Hope,

If no one come up with a better idea, you may consider trying a
truncated cone of Plexiglas that has one end large enough to capture
most of the light from the flash tube and the other the size you need
for the light source of the scope and fitted in the proper place over
the condenser may work. You may have to frost the small end of the cone
or cut the large end at an angle of the central axis of the microscope
lighting to get the light as even as you want.

The cone works on the principle of Total Internal Reflection. I would
use Plexiglas because it can be fire polished by finishing it with 400
grit wet/dry silicon carbide sandpaper and then a flame will melt the
surface to a high polish. It takes a few tires to get the fire polishing
down but you can clean up the mess and do it over a lot of times. Just
keep the flame moving and the part spinning. The fire polishing happens
very fast at about 180-200 C.

Using plastic any machine shop or anyone with a lathe can do the job in
a few minutes and you don't need a optical shop to do the work.

Plexiglas is much more transparent to light than glass and doesn't block
Ultra Violet. So it works with florescent light as well.

Good luck
Gordon
Gordon Couger
Microscope documentation Archive http://science-info.net/



hopechen-at-u.washington.edu wrote:
}
} Email: hopechen-at-u.washington.edu
} Name: Hope Chen
}
} Organization: University of Washington
}
} Education: Graduate College
}
} Location: Seattle. WA US
}
} Title: microscope illumination
}
} Question: Dear Microscopist,
}
} I try to set up a high-speed microphotography system using an ultra-high speed camera and an inverted microscope. I'm now facing a problem that is how to incorporate flash tube light source with the microscope. The light source we has is Photogenic Radio Powerlight 2500DRUV and the flash tube is Photogenic C4-19 Flashtube. The flash tube is annular with an outside diameter about 1.8 inch and inner diameter about 1 inch. Our requirements to the microscope illumination system are: 1) collect as much light as possible to increase the illumination light intensity, which hopefully could allow us to adjust our high-speed camera's exposure time to as small as 10 ns, and 2) help us to get high resolution images.
} Iím thinking of using Kohler illumination, but due to the light source is a large annular light source I couldnít get idea Kohler illumination. Nelsonian illumination focus light to the specimen, so this illumination method might be better than Kohler illumination in terms of light intensity, but then I need to focus an annular light to a point.
} May I ask for your suggestions?
}
} send with thanks,
} Hope
}
}
}




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From: tbargar-at-unmc.edu
Date: Wed, 23 May 2007 08:44:23 -0500
Subject: [Microscopy] Cacodylate buffer vs. phosphate buffer

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Dear listers,

I have a faculty member who wants to do a perfusion fix of brain (it's a
rat or mouse I forget which). Normally I avoid using cacodylate buffer
because of the arsenic. However, in the dim recesses of my memory I seem
to recall that phosphate buffer is very prone to precipitate formation in
brain tissue. Now I don't know if I'm remembering this correctly. So,
what are your opinions? Would it be better to use a cacodylate buffer or
should I stick with the safer less toxic phosphate buffer? As always
thanks in advance for your help.

Tom Bargar
University of Nebraska Medical Center
Core Electron Microscopy Research Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu


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From: phillipst-at-missouri.edu
Date: Wed, 23 May 2007 10:14:43 -0500
Subject: [Microscopy] Re: Cacodylate buffer vs. phosphate buffer

Contents Retrieved from Microscopy Listserver Archives
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I have never used it for perfusion fixation but HEPES buffer works well in
my standard fixes with 2% paraformaldehyde or 2% paraformaldehyde + 2.5%
glutaraldehyde. Alternatively, PIPES is used by other researchers with
success. I see no argument for retaining use of cacodylate. Good luck, Tom


At 08:46 AM 05/23/07, you wrote:



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Thomas E. Phillips, PhD
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400

573-882-4712 (office)
573-882-0123 (fax)
PhillipsT-at-missouri.edu
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http://www.biotech.missouri.edu/mcc/


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From: jacques.faerber-at-ipcms.u-strasbg.fr
Date: Wed, 23 May 2007 11:11:51 -0500
Subject: [Microscopy] basic SEM: imaging salt crystals

Contents Retrieved from Microscopy Listserver Archives
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Hi all

One or two cent more!

For EDS analysis, if you coat your crystals with a high Z material, and
your analysis software dont' have an "sample coating" option, you'll
probably will have wrong results on crystals like NaCl. It's always
difficult to have accurate results on Na (it evaoprate under the beam,
but not enough to polluate the SEM !), but with gold coating, absorption
from the Na-k by the gold layer will be much stronger than that from
Cl-k, and the balance Na-Cl will be worse. I think that carbon is the
less problematic coating for such analysis, and I would not use Au or
Pt. Of coarse, imaging can be less nice than with Au, Pt or Ir coating.

For Monte Carlo simulation, you can download Casino
(http://www.gel.usherbrooke.ca/casino/), with has some nice possibility,
like multiple energies simulation. I use it complementary to Electron
Flight Simulator.

A simple test in particule (or thin layer) analysis, is to put the
particules on a substrate which don't have any element from the
particules (of coarse, you must have some idees about these particules,
and yes, you don't have any, that's what your are looking for !) and to
perform measurement with different energies, from say 5 to 20 keV. As
long as you see the x-ray lines of the subtrate in the spectrum, the
primary energie is too hight. By the way, it is suffisent that one
element from the substrate does not exist in the particule, if possible
with a line at low energie, to be able to see it in the spectrum at all
energies (Cu-L par exemple).

Hope it helps

J. Faerber
IPCMS-GSI
(Institut de Physique et Chimie des Matériaux de Strasbourg
Groupe Surface et Interfaces)
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail Jacques.Faerber-at-ipcms.u-strasbg.fr



wesaia-at-iastate.edu a écrit :
}
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} I have already seen a number of good answers, but I will chime in anyway.
}
} I would suggest coating the samples unless you are operating in variable pressure mode (which may not be an option for you). I recall that NaCl charges as well as most any insulator. I don't imagine you would be doing quant on the sample since the lack of a known geometry would lead to substantial uncertainty in the results. Gary made a good point that it would be nice to use an element for coating that is only present in the coating. Otherwise, there will be uncertainty in the source of the peaks. I would expect some difference in the relative peak intensities depending on which element is used.
}
} I wouldn't worry much about vaporizing the salt. (Maybe I should.) I don't expect you will be using nanoamps of current. Once again, VP mode would help minimize or prevent contamination.
}
} I don't have the latest and greatest Monte Carlo programs. I do have an old one derived from David Joy's work. I ran a simulation on S at 15 kV and I estimate the excitation depth to be slightly less than 1 um. Running the same simulation for Na, I found the depth to be more like 2 um. For Cu, I found the depth to be on the order of 0.3 um. Therefore, I generally quote penetration depths of hundreds of nm up to a couple of microns. The dimensions of the interaction volume dwarf the spot size so much that I don't often bother considering it in calculating interaction volume.
}
} As a fun little exercise, you might lay down a gold layer on copper or some other metal. The interaction volume for gold appears to be slightly less than 0.2 um (200 nm). Since we routinely coat samples with 10 nm of gold, we need a deeper penetration depth if we are ever going to see the sample through the gold. We don't have a problem seeing the sample unless we are using a very thick layer of gold or a low voltage. A 3 kV beam would have difficulty penetrating 10 nm of gold, but there would not be much excitation of sample x-rays at that low a voltage anyway.
}
} Warren
}
} -----Original Message-----
} X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Tuesday, May 22, 2007 4:47 AM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] basic SEM: imaging salt crystals
}
} Dear listers,
}
} Here are some basic questions from the basic SEM user
} I am: I would like to visualize salt crystals by SEM
} and make EDX analysis if possible. I just leave a drop
} of salt solution on a carbon tab dry, that's all!
} My questions:
}
} - Do I need to carbon-coat the crystals?
} - Is there a risk to evaporate the crystals under the
} electron beam and make the column dirty? (I usually
} don't use HV higher than 15 kV). Would carbon coating
} help dissipate the heat in this regard?
}
} Now another question related to SEM but not to
} crystals:
} When I analyse fine particles by EDX, I never know if
} I analyse only the particle or also part of the
} material which is behind the particle. Is there a way
} to determine the depth of analysis of the material
} studied? I suppose that it depends on the HV and on
} the nature of the material itself, but I already would
} be happy to have a order of magnitude. Is this 10 nm
} or 2 µm deep?
} Lets say I analyze at 15kV (1) NaCl crystals (2)
} Quartz (3) Feldpar (4) Copper, what would be the
} expected "depth of analysis" in these samples?
} Does it depend on the size of the beam?
}
} Best regards,
}
} Stephane
}
}
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From: gary-at-gaugler.com
Date: Wed, 23 May 2007 11:27:32 -0500
Subject: [Microscopy] Re: basic SEM: imaging salt crystals

Contents Retrieved from Microscopy Listserver Archives
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What is a "sample coating option?" I don't see a need
for this. Just don't tag the coating peak(s) with their
element nomenclature. Why one would want to quant NaCl
is another and separate issue.

An 8KV beam on thin coated NaCl will produce nice pix
and generate the desired peaks (slowly). With a Zeiss
Supra 55VP, 30u aperture, low current, 10KV, the crystals
do not melt. If one is worried about it, use smaller
aperture and squeeze down the beam if your SEM is set up
that way.

gary g.




At 08:13 AM 5/23/2007, you wrote:




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From: larry.ackerman-at-ucsf.edu
Date: Wed, 23 May 2007 12:41:45 -0500
Subject: [Microscopy] Re: monkey perfusion methodology

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Mary Ellen,
The rate that Walter suggested seemed extremely high compared to my
experience with smaller animals so I asked an anatomist with many
decades of experience with monkeys. He has always used an initial flush
of 500ml in 5 minutes time with buffer followed by 1.5 liters of
fixative over 20 minutes of time for a typical 5kg animal. The
appropriate size tubing and cannula in collaboration with the setting on
the pump will enable an excellent fixation. Walter's method may well
provide good results. I would love to see some brain slices and TEM
micrographs at 10,000X. I'm open to different methods.

Larry

Walter.Bobrowski-at-pfizer.com wrote:
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} 300 ml/min. Perfuse either through the carotid artery or left ventricle
} (occlude descending aorta).
}
} Best regards,
} Walter F. Bobrowski
} Sr. Scientist
} Pfizer Global R&D
} 2800 Plymouth Rd.
} Ann Arbor, MI 48105
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} -----Original Message-----
} X-from: mpease-at-jhmi.edu [mailto:mpease-at-jhmi.edu]
} Sent: Monday, May 21, 2007 7:25 PM
} To: Bobrowski, Walter
} Subject: [Microscopy] viaWWW: monkey perfusion methodology
}
}
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} Email: mpease-at-jhmi.edu
} Name: Mary Ellen Pease
}
} Organization: Johns Hopkins Wilmer Eye Institute
}
} Title-Subject: [Filtered] monkey perfusion methodology
}
} Question: What is the recommended flow rate for perfusion of monkey
} brain and eyes when using a pump rather than gravity feed of the
} solutions?
}
} Thank you,
} Mary Ellen
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--
Larry Ackerman, Associate Specialist
UCSF, Dept. of Anatomy, Rm S1347
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu


==============================Original Headers==============================
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From: oshel1pe-at-cmich.edu
Date: Wed, 23 May 2007 14:40:35 -0500
Subject: [Microscopy] Re: Unknown creature

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear All,
can somebody help me out with the name and species of a tiny little animal I
found on an absinthe leaf?
See http://www.elektronenmikroskopie.info/absinthe_animal.jpg

:-)

Thank you,
Stefan Diller



----------------------------------------------------------------------------
-----------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49 - 931 - 7848700 Phone
++49 - 931 - 7848701 Fax
++49 - 175 - 717 70 51 Cell-Phone

Websites:
www.stefan-diller.com
www.elektronenmikroskopie.info
www.assisi.de
Anfahrt: http://mail.map24.com/stefan.diller
----------------------------------------------------------------------------
-----------------------------------------
----- Original Message -----
X-from: {gary-at-gaugler.com}
To: {stefan.diller-at-t-online.de}
Sent: Wednesday, May 23, 2007 6:33 PM

Stefan,

Do you have any more images? Different angles of view would help

Phil

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From: Phil.Ahrenkiel-at-sdsmt.edu
Date: Wed, 23 May 2007 14:49:01 -0500
Subject: [Microscopy] Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Greetings-

Our nanoscience program received some information about the Phenom-Ed
from FEI, a tablettop SEM designed for education. We are considering
whether it would useful for students to perform quick analysis of
nanomaterials. Sounds like the instrument is still in the prototype
stage, but we are wondering if anyone has any impressions they could
share on this. From the price we were quoted, it might actually be
within our budget.

Some info on this product is available at:
http://investor.fei.com/phoenix.zhtml?c=60978&p=irol-newsArticle&ID=9682
99&highlight=

Thanks in advance for your feedback.
-Phil
------------------------------------------
Phil Ahrenkiel, Assistant Professor
Nanoscience and Nanoengineering Ph.D. Program
South Dakota School of Mines and Technology
501 East St. Joseph Street
Rapid City, South Dakota 57701
Office: EEP 221
Phone: (605)394-5238, Fax: (605)394-2365
Email: Phil.Ahrenkiel-at-sdsmt.edu



==============================Original Headers==============================
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From: Sander.Stoks-at-fei.com
Date: Wed, 23 May 2007 15:30:29 -0500
Subject: [Microscopy] Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello,

I would not call it "prototype stage" anymore. As coincidence would have it, I was at the official introduction of the Phenom at the Fontys University of Applied Sciences in Eindhoven, The Netherlands, earlier today. The instrument will be used there exactly for what you suggest.

There is (much) more information available at http:// {http://www.fei.com/phenom} www.fei.com/phenom.

I won't say more about the instrument here on the list (unless requested) since I am part of the team who actually designed it, and hence would come across as rather biased (in a positive way!). There are links on the website with contact information. Please click on them, we love talking about it!

Regards,
Sander

________________________________

X-from: Phil.Ahrenkiel-at-sdsmt.edu [mailto:Phil.Ahrenkiel-at-sdsmt.edu]
Sent: Wed 5/23/2007 9:53 PM
To: Stoks, Sander


Greetings-

Our nanoscience program received some information about the Phenom-Ed
from FEI, a tablettop SEM designed for education. We are considering
whether it would useful for students to perform quick analysis of
nanomaterials. Sounds like the instrument is still in the prototype
stage, but we are wondering if anyone has any impressions they could
share on this. From the price we were quoted, it might actually be
within our budget.

Some info on this product is available at:
http://investor.fei.com/phoenix.zhtml?c=60978&p=irol-newsArticle&ID=9682
99&highlight=

Thanks in advance for your feedback.
-Phil
------------------------------------------
Phil Ahrenkiel, Assistant Professor
Nanoscience and Nanoengineering Ph.D. Program
South Dakota School of Mines and Technology
501 East St. Joseph Street
Rapid City, South Dakota 57701
Office: EEP 221
Phone: (605)394-5238, Fax: (605)394-2365
Email: Phil.Ahrenkiel-at-sdsmt.edu



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From: jquinn-at-www.matscieng.sunysb.edu
Date: Wed, 23 May 2007 18:58:55 -0500
Subject: [Microscopy] Re: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

} From mail-at-ns.microscopy.com Wed May 23 15:48:05 2007
} Date: Wed, 23 May 2007 14:49:29 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: Phil.Ahrenkiel-at-sdsmt.edu
} Reply-to: Phil.Ahrenkiel-at-sdsmt.edu
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
} Subject: [Microscopy] Phenom-Ed SEM
} Errors-To: MicroscopyListSpamFilter-at-microscopy.com
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}
} Greetings-
}
} Our nanoscience program received some information about the Phenom-Ed
} from FEI, a tablettop SEM designed for education. We are considering
} whether it would useful for students to perform quick analysis of
} nanomaterials. Sounds like the instrument is still in the prototype
} stage, but we are wondering if anyone has any impressions they could
} share on this. From the price we were quoted, it might actually be
} within our budget.
}
} Some info on this product is available at:
} http://investor.fei.com/phoenix.zhtml?c=60978&p=irol-newsArticle&ID=9682
} 99&highlight=
}
} Thanks in advance for your feedback.
} -Phil
} ------------------------------------------
} Phil Ahrenkiel, Assistant Professor
} Nanoscience and Nanoengineering Ph.D. Program
} South Dakota School of Mines and Technology
} 501 East St. Joseph Street
} Rapid City, South Dakota 57701
} Office: EEP 221
} Phone: (605)394-5238, Fax: (605)394-2365
} Email: Phil.Ahrenkiel-at-sdsmt.edu
}
}
}
} ==============================Original Headers==============================
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} 7, 20 -- From: "Ahrenkiel, Scott \(Phil\) P." {Phil.Ahrenkiel-at-sdsmt.edu}
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}

Phil and others -

Tell me the cost and I will give you an answer.

It certainly looks like a great entry level unit.

But a few caveats.......

1) I would question the use of the word "nanoscale".

2) You will never get EDS out of it.

3) You cannot use large samples.

The last two would exclude 3/4 of my users.

However, I eagerly look forward to the cost.

kind regards,

Jim

PS: OoO away.........


} From mail-at-ns.microscopy.com Wed May 23 15:48:05 2007
} Date: Wed, 23 May 2007 14:49:29 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: Phil.Ahrenkiel-at-sdsmt.edu
} Reply-to: Phil.Ahrenkiel-at-sdsmt.edu
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
} Subject: [Microscopy] Phenom-Ed SEM
} Errors-To: MicroscopyListSpamFilter-at-microscopy.com
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}
}
}
}
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}
}
} Greetings-
}
} Our nanoscience program received some information about the Phenom-Ed
} from FEI, a tablettop SEM designed for education. We are considering
} whether it would useful for students to perform quick analysis of
} nanomaterials. Sounds like the instrument is still in the prototype
} stage, but we are wondering if anyone has any impressions they could
} share on this. From the price we were quoted, it might actually be
} within our budget.
}
} Some info on this product is available at:
} http://investor.fei.com/phoenix.zhtml?c=60978&p=irol-newsArticle&ID=9682
} 99&highlight=
}
} Thanks in advance for your feedback.
} -Phil
} ------------------------------------------
} Phil Ahrenkiel, Assistant Professor
} Nanoscience and Nanoengineering Ph.D. Program
} South Dakota School of Mines and Technology
} 501 East St. Joseph Street
} Rapid City, South Dakota 57701
} Office: EEP 221
} Phone: (605)394-5238, Fax: (605)394-2365
} Email: Phil.Ahrenkiel-at-sdsmt.edu
}
}
}
} ==============================Original Headers==============================
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From: scott.coutts-at-med.monash.edu.au
Date: Thu, 24 May 2007 01:11:05 -0500
Subject: [Microscopy] Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

jquinn-at-www.matscieng.sunysb.edu wrote:
} Phil and others -
}
} Tell me the cost and I will give you an answer.
} It certainly looks like a great entry level unit.
} But a few caveats.......
} 1) I would question the use of the word "nanoscale".
} 2) You will never get EDS out of it.
} 3) You cannot use large samples.
} The last two would exclude 3/4 of my users.
} However, I eagerly look forward to the cost.
} kind regards,
} Jim
} PS: OoO away.........
}

Well, 62,000 Euro is the RRP from the website.


--
Scott J. Coutts
-------------------------------------------------------------------------
Bacterial Pathogenesis Research Group
Department of Microbiology
Box 53, Monash University, 3800, Australia
Phone +61 3 9905 8592, Fax +61 3 9905 4811
-------------------------------------------------------------------------

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From: michael-at-shaffer.net
Date: Thu, 24 May 2007 06:36:19 -0500
Subject: [Microscopy] RE: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Phil writes ...

} Our nanoscience program received some information about the
} Phenom-Ed from FEI, a tablettop SEM designed for education.
} We are considering whether it would useful for students to
} perform quick analysis of nanomaterials. ...

Regarding "nanomaterials", keep in mind that magnifications are not what
they used to be. For example, it used to be that a SEM's mag number was
based on a 4x5 polaroid, whereas now the mag number is based on a 10" window
on the computer display. The white paper describes the Phenom capable of
mag=20,000x which, relative to SEMs we are used to, would equate to 10,000x
(1 cm = 1,000 nM on a 4x5).

Not to take away from what should be a very valuable and interesting tool in
a science program for many high schools and colleges, and I only comment
relative to the term "nanomaterials".

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
Inco Innovation Centre
Memorial University
St. John's, Newfoundland



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From: jacques.faerber-at-ipcms.u-strasbg.fr
Date: Thu, 24 May 2007 07:15:03 -0500
Subject: [Microscopy] basic SEM: imaging salt crystals

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all again

Gary, in answer to your question, the last time I made a round robin to
buy an EDS system (back 6 years ago or more), one manufacturer, but I
don't remember who, had that option, in which you give your coating
material and an estimated thickness. It takes the coating in account to
correct the selective absorbtion of the X-ray lines from the sample by
the coating. Perheps someone has informations on that possibility.

Of coarse, it has not much meaning to quantify NaCl by EDS (only to test
your quanti software !). But, Na is a typical exemple of that problem :
low energy lines will be absorbed selectively, in comparaison to higher
energy lines. This particulary the case with O-k, l lines from
transition metals, Na, Mg, etc. The quantification will not be correct,
if you only ignore the presence of gold. The balance between Ca, K on
one side, and Na or Mg (and O if one use the measured intensities) on
the other side will be biased.

Of coarse, with 5 nm of a grainy gold coating (blue at daylight), where
electrical conducitvity is done mostly by tunneling between gold
islands, letting a large aera of the sample uncoated, you will not see a
big error. But with a nice yellowish } 25 nm coating, which cover
uniformely the sample, the effect can be very effective. I have recently
seen pictures/samples with a uniform grainy surface from the coating, in
a case where one was looking after Cl and S in a polymer. No hope to see
something in such a case. And such situations are unfortunatly not
unfrequent.

As said, Na is soon difficult to quantify correctly, in glasses for
exemple. So I prefer to be carefull, and not add a source of errors.

Do you think I'm overcarefull ? Maybe !!!

jcqs

J. Faerber
IPCMS-GSI
(Institut de Physique et Chimie des Matériaux de Strasbourg
Groupe Surface et Interfaces)
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail Jacques.Faerber-at-ipcms.u-strasbg.fr



gary-at-gaugler.com a écrit :
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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}
} What is a "sample coating option?" I don't see a need
} for this. Just don't tag the coating peak(s) with their
} element nomenclature. Why one would want to quant NaCl
} is another and separate issue.
}
} An 8KV beam on thin coated NaCl will produce nice pix
} and generate the desired peaks (slowly). With a Zeiss
} Supra 55VP, 30u aperture, low current, 10KV, the crystals
} do not melt. If one is worried about it, use smaller
} aperture and squeeze down the beam if your SEM is set up
} that way.
}
} gary g.
}
}
}
}
} At 08:13 AM 5/23/2007, you wrote:
}
}
}
}
}
} } ----------------------------------------------------------------------------
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} } ----------------------------------------------------------------------------
} }
} } Hi all
} }
} } One or two cent more!
} }
} } For EDS analysis, if you coat your crystals with a high Z material, and
} } your analysis software dont' have an "sample coating" option, you'll
} } probably will have wrong results on crystals like NaCl. It's always
} } difficult to have accurate results on Na (it evaoprate under the beam,
} } but not enough to polluate the SEM !), but with gold coating, absorption
} }
} } from the Na-k by the gold layer will be much stronger than that from
}
} } Cl-k, and the balance Na-Cl will be worse. I think that carbon is the
} } less problematic coating for such analysis, and I would not use Au or
} } Pt. Of coarse, imaging can be less nice than with Au, Pt or Ir coating.
} }
} } For Monte Carlo simulation, you can download Casino
} } (http://www.gel.usherbrooke.ca/casino/), with has some nice possibility,
} } like multiple energies simulation. I use it complementary to Electron
} } Flight Simulator.
} }
} } A simple test in particule (or thin layer) analysis, is to put the
} } particules on a substrate which don't have any element from the
} } particules (of coarse, you must have some idees about these particules,
} } and yes, you don't have any, that's what your are looking for !) and to
} } perform measurement with different energies, from say 5 to 20 keV. As
} } long as you see the x-ray lines of the subtrate in the spectrum, the
} } primary energie is too hight. By the way, it is suffisent that one
} } element from the substrate does not exist in the particule, if possible
} } with a line at low energie, to be able to see it in the spectrum at all
} } energies (Cu-L par exemple).
} }
} } Hope it helps
} }
} } J. Faerber
} } IPCMS-GSI
} } (Institut de Physique et Chimie des Matériaux de Strasbourg
} } Groupe Surface et Interfaces)
} } 23, rue de Loess ; BP43
} } 67034 Strasbourg CEDEX 2
} } France
} }
} } Tel 00 33(0)3 88 10 71 01
} } Fax 00 33(0)3 88 10 72 48
} } E-mail Jacques.Faerber-at-ipcms.u-strasbg.fr
} }
} }
} }
} } wesaia-at-iastate.edu a écrit :
} }
} } }
} } ----------------------------------------------------------------------------
} }
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} } }
} } }
} } ----------------------------------------------------------------------------
} }
} } } I have already seen a number of good answers, but I will chime in anyway.
} } }
} } } I would suggest coating the samples unless
} } }
} } you are operating in variable pressure mode
} } (which may not be an option for you). I recall
} } that NaCl charges as well as most any
} } insulator. I don't imagine you would be doing
} } quant on the sample since the lack of a known
} } geometry would lead to substantial uncertainty
} } in the results. Gary made a good point that it
} } would be nice to use an element for coating
} } that is only present in the coating. Otherwise,
} } there will be uncertainty in the source of the
} } peaks. I would expect some difference in the
} } relative peak intensities depending on which element is used.
} }
} } } I wouldn't worry much about vaporizing the
} } }
} } salt. (Maybe I should.) I don't expect you will
} } be using nanoamps of current. Once again, VP
} } mode would help minimize or prevent contamination.
} }
} } } I don't have the latest and greatest Monte
} } }
} } Carlo programs. I do have an old one derived
} } from David Joy's work. I ran a simulation on S
} } at 15 kV and I estimate the excitation depth to
} } be slightly less than 1 um. Running the same
} } simulation for Na, I found the depth to be more
} } like 2 um. For Cu, I found the depth to be on
} } the order of 0.3 um. Therefore, I generally
} } quote penetration depths of hundreds of nm up
} } to a couple of microns. The dimensions of the
} } interaction volume dwarf the spot size so much
} } that I don't often bother considering it in calculating interaction volume.
} }
} } } As a fun little exercise, you might lay down
} } }
} } a gold layer on copper or some other metal. The
} } interaction volume for gold appears to be
} } slightly less than 0.2 um (200 nm). Since we
} } routinely coat samples with 10 nm of gold, we
} } need a deeper penetration depth if we are ever
} } going to see the sample through the gold. We
} } don't have a problem seeing the sample unless
} } we are using a very thick layer of gold or a
} } low voltage. A 3 kV beam would have difficulty
} } penetrating 10 nm of gold, but there would not
} } be much excitation of sample x-rays at that low a voltage anyway.
} }
} } } Warren
} } }
} } } -----Original Message-----
} } } X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} } } Sent: Tuesday, May 22, 2007 4:47 AM
} } } To: wesaia-at-iastate.edu
} } } Subject: [Microscopy] basic SEM: imaging salt crystals
} } }
} } } Dear listers,
} } }
} } } Here are some basic questions from the basic SEM user
} } } I am: I would like to visualize salt crystals by SEM
} } } and make EDX analysis if possible. I just leave a drop
} } } of salt solution on a carbon tab dry, that's all!
} } } My questions:
} } }
} } } - Do I need to carbon-coat the crystals?
} } } - Is there a risk to evaporate the crystals under the
} } } electron beam and make the column dirty? (I usually
} } } don't use HV higher than 15 kV). Would carbon coating
} } } help dissipate the heat in this regard?
} } }
} } } Now another question related to SEM but not to
} } } crystals:
} } } When I analyse fine particles by EDX, I never know if
} } } I analyse only the particle or also part of the
} } } material which is behind the particle. Is there a way
} } } to determine the depth of analysis of the material
} } } studied? I suppose that it depends on the HV and on
} } } the nature of the material itself, but I already would
} } } be happy to have a order of magnitude. Is this 10 nm
} } } or 2 µm deep?
} } } Lets say I analyze at 15kV (1) NaCl crystals (2)
} } } Quartz (3) Feldpar (4) Copper, what would be the
} } } expected "depth of analysis" in these samples?
} } } Does it depend on the size of the beam?
} } }
} } } Best regards,
} } }
} } } Stephane
} } }
} } }
} } } ==============================Original
} } }
} } Headers==============================
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From: gas19-at-daimlerchrysler.com
Date: Thu, 24 May 2007 08:40:56 -0500
Subject: [Microscopy] viaWWW: Phenom-Ed SEM

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Email: gas19-at-daimlerchrysler.com
Name: Gerald Shulke

Organization: DaimlerChrysler Corporation

Title-Subject: [Filtered] Re: [Microscopy] Phenom-Ed SEM

Question: Just to throw in another option. I saw a quick review of a Hitachi tabletop SEM. It's always good to see a comparison. I'm really not interested in either one. They are limited in function. Hitachi's model is BSE only and had limited stage manipulation capability. They maybe a good learning tool for someone who has never been exposed to an SEM before. It's not a replacement for a true SEM, just a fun toy.

http://www.hitachi-hta.com/pageloader~type~product~id~450~orgid~42.html

Gerald Shulke
Materials Engineering Specialist
DaimlerChrysler Corporation

---------------------------------------------------------------------------

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From: Karen_Bentley-at-URMC.Rochester.edu
Date: Thu, 24 May 2007 09:32:43 -0500
Subject: [Microscopy] Myelin Osmication Problem

Contents Retrieved from Microscopy Listserver Archives
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Hello Fellow Listers:

I have an issue regarding perfusion fixed (glut & paraform. in
cacodylate buffer) adult rat spinal cord.

I have cut cord cross sections approx. 1-1.5 mm. thick which were
osmicated in a 1.0% OsO4/cacodylate buffer for 1.5 hrs. When I cut into
the middle of the tissue block it was completely white.......the osmium
did not penetrate beyond the surface of the tissue. I know that the
heavily myelinated spinal cord can be a barrier to complete diffusion of
osmium post fixation. These blocks will be sectioned for light
microscopy stained with Toluidine Blue to access myelin fiber numbers,
then potentially TEM examination.

Question: I have researched the literature and one paper suggested
using 2.0% osmium and leaving the spinal cord blocks in overnight.
Another suggested using potassium ferrocyanide with the osmium. One of
our university EM experienced researchers suggested warming the osmium
to 37 C. Any help would be greatly appreciated as I need an answer
quickly.

Karen Bentley

Karen L. Bentley, M.S.
Technical Director
Electron Microscope Research Core
University of Rochester Medical Center
575 Elmwood Avenue, Box 626
Rochester, NY 14642
585-275-1954


==============================Original Headers==============================
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From: ahlst007-at-umn.edu
Date: Thu, 24 May 2007 09:48:53 -0500
Subject: [Microscopy] Re: Myelin Osmication Problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Karen,

Is is possible for you to cut thinner
cross-sections? I am working with a project that
hands over to me fixed (similar fix to yours)
rat spinal and ganglia sections around 40
micron. They are fully osmicated, black all the
way through. I think they use a vibratome to cut
those sections.
Gib
--
Gilbert (Gib) Ahlstrand, Electron Microscopist,
Imaging Center, University of Minnesota
123 Snyder Hall
St. Paul, MN 55108
http://www.cbs.umn.edu/ic

Karen_Bentley-at-URMC.Rochester.edu wrote:
}
} Hello Fellow Listers:
}
} I have an issue regarding perfusion fixed (glut & paraform. in
} cacodylate buffer) adult rat spinal cord.
}
} I have cut cord cross sections approx. 1-1.5 mm. thick which were
} osmicated in a 1.0% OsO4/cacodylate buffer for 1.5 hrs. When I cut into
} the middle of the tissue block it was completely white.......the osmium
} did not penetrate beyond the surface of the tissue. I know that the
} heavily myelinated spinal cord can be a barrier to complete diffusion of
} osmium post fixation. These blocks will be sectioned for light
} microscopy stained with Toluidine Blue to access myelin fiber numbers,
} then potentially TEM examination.
}
} Question: I have researched the literature and one paper suggested
} using 2.0% osmium and leaving the spinal cord blocks in overnight.
} Another suggested using potassium ferrocyanide with the osmium. One of
} our university EM experienced researchers suggested warming the osmium
} to 37 C. Any help would be greatly appreciated as I need an answer
} quickly.
}
} Karen Bentley
}
} Karen L. Bentley, M.S.
} Technical Director
} Electron Microscope Research Core
} University of Rochester Medical Center
} 575 Elmwood Avenue, Box 626
} Rochester, NY 14642
} 585-275-1954


==============================Original Headers==============================
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From: rjharris-at-uwo.ca
Date: Thu, 24 May 2007 10:10:51 -0500
Subject: [Microscopy] Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Phil
You might also want to check out Hitachi's Tabletop Microscope TM-1000
"Designed to fulfill the needs of applications that require more information
than can be provided by a light microscope but doesn't require the expertise
of a dedicated SEM operator"
Don't own one, but have been impressed with the instrument at a couple of
shows. Seems robust and reliable.
Rick,

Richard Harris
Manager -
Imaging, Information and Data Systems
Biotron Experimental Climate Change Research Facility
University of Western Ontario,
London Ontario, CANADA.
N6A 5B7
Ph. 519-661-2111 ext. 86780
Fax 519-661-3935
e-mail rjharris-at-uwo.ca
web: www.biotron.uwo.ca

-----Original Message-----
X-from: Phil.Ahrenkiel-at-sdsmt.edu [mailto:Phil.Ahrenkiel-at-sdsmt.edu]
Sent: Wednesday, May 23, 2007 3:53 PM
To: rjharris-at-uwo.ca


Greetings-

Our nanoscience program received some information about the Phenom-Ed
from FEI, a tablettop SEM designed for education. We are considering
whether it would useful for students to perform quick analysis of
nanomaterials. Sounds like the instrument is still in the prototype
stage, but we are wondering if anyone has any impressions they could
share on this. From the price we were quoted, it might actually be
within our budget.

Some info on this product is available at:
http://investor.fei.com/phoenix.zhtml?c=60978&p=irol-newsArticle&ID=9682
99&highlight=

Thanks in advance for your feedback.
-Phil
------------------------------------------
Phil Ahrenkiel, Assistant Professor
Nanoscience and Nanoengineering Ph.D. Program
South Dakota School of Mines and Technology
501 East St. Joseph Street
Rapid City, South Dakota 57701
Office: EEP 221
Phone: (605)394-5238, Fax: (605)394-2365
Email: Phil.Ahrenkiel-at-sdsmt.edu



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From: mcauliff-at-umdnj.edu
Date: Thu, 24 May 2007 10:38:36 -0500
Subject: [Microscopy] Re: Myelin Osmication Problem

Contents Retrieved from Microscopy Listserver Archives
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Hi Karen:

Osmium will penetrate 1 mm maximum but if both surfaces of a cross
section of sp. cord are exposed you 'should' be OK with 2 exposed surfaces.
Are the sections really 1-1.5 mm thick? Are you agitating the tissue? Is
the osmium reasonably fresh?
I don't see how 2% osmium will solve the problem and ferrocyanide won't
help if the osmium does not arrive at the 'target'.
Warming the osmium might help but I don't see it making a huge difference.
I suggest fresh osmium, longer times in osmium, frequent (continuous?)
agitation. If that fails try using a Vibratome to cut 0.5 -0.8 mm sections.
If you are sectioning for TEM are you really going to cut all the way
through a 1.5 mm slice? If not, make the sections thinner.

Geoff

Karen_Bentley-at-URMC.Rochester.edu wrote:

} ----------------------------------------------------------------------------
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--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583; fax: -4029
mcauliff-at-umdnj.edu
**********************************************



==============================Original Headers==============================
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8, 35 -- 24 May 2007 11:38:01 -0400 (EDT)
8, 35 -- Date: Thu, 24 May 2007 11:38:01 -0400
8, 35 -- From: Geoff McAuliffe {mcauliff-at-umdnj.edu}
8, 35 -- Subject: Re: [Microscopy] Myelin Osmication Problem
8, 35 -- In-reply-to: {200705241433.l4OEXkQ3014514-at-ns.microscopy.com}
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==============================End of - Headers==============================




From: mcauliff-at-umdnj.edu
Date: Thu, 24 May 2007 10:56:31 -0500
Subject: [Microscopy] telephone solicitations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Lists:

I got another telephone call at my lab yesterday from a solicitor.
My phone number here is unlisted and does not show up on any caller id.
I am getting 1 or 2 of these calls a week, too many to be random hits. I
think some companies are harvesting my number from Listserv postings.
Has anyone else had this experience?

Geoff

--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583; fax: -4029
mcauliff-at-umdnj.edu
**********************************************



==============================Original Headers==============================
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6, 33 -- 24 May 2007 11:45:19 -0400 (EDT)
6, 33 -- Date: Thu, 24 May 2007 11:45:18 -0400
6, 33 -- From: Geoff McAuliffe {mcauliff-at-umdnj.edu}
6, 33 -- Subject: telephone solicitations
6, 33 -- To: MicroscopyListserver {microscopy-at-msa.microscopy.com} ,
6, 33 -- Histonet {histonet-at-pathology.swmed.edu}
6, 33 -- Message-id: {4655B30E.6030502-at-umdnj.edu}
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From: TJJ-at-stowers-institute.org
Date: Thu, 24 May 2007 11:10:33 -0500
Subject: [Microscopy] RE: telephone solicitations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Geoff, this is why I don't list my phone number in my Histonet posts any
more. I was getting unsolicited phone calls.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110



Dear Lists:

I got another telephone call at my lab yesterday from a solicitor.
My phone number here is unlisted and does not show up on any caller id.
I am getting 1 or 2 of these calls a week, too many to be random hits. I

think some companies are harvesting my number from Listserv postings.
Has anyone else had this experience?

Geoff

--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583; fax: -4029
mcauliff-at-umdnj.edu
**********************************************



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May 2007 11:45:18 -0400 6, 33 -- From: Geoff McAuliffe
{mcauliff-at-umdnj.edu} 6, 33 -- Subject: telephone solicitations 6, 33 --
To: MicroscopyListserver {microscopy-at-msa.microscopy.com} ,
6, 33 -- Histonet {histonet-at-pathology.swmed.edu}
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From: lcgould-at-med.cornell.edu
Date: Thu, 24 May 2007 11:15:55 -0500
Subject: [Microscopy] Re: telephone solicitations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Geoff-
I haven't been getting calls, but I do know that if I "Google"
myself (don't we all do that sometimes?), I get pages of information,
some of which is from my facilities' websites, some reference
publications, but many hits are postings to this list. I am also
frequently asked about my pricing for services, based on a price list
that hasn't been on the facility's website in years, but persists out
there on the www. We all must remember that there is no way to
"erase" things once they hit cyberspace: a fact many new college
grads are finding out when prospective employers find their
embarrassing "My Space" postings.
If these calls bother you, may I suggest that you create a alternate
"signature" for your emails to the list, omitting your phone number.
MSA members can find you from the member directory if we need to.
That listing is password protected so outsiders can't get the info
from there. Just remember, those of us who are frequent users of the
listserver have put ourselves out in public through no fault of our
wonderful Sysop, or anyone else. Its just the nature of the beast.
Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

==============================Original Headers==============================
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2, 24 -- Date: Thu, 24 May 2007 12:15:39 -0400
2, 24 -- From: "Leona Cohen-Gould" {lcgould-at-med.cornell.edu}
2, 24 -- Subject: Re: [Microscopy] telephone solicitations
2, 24 -- In-reply-to: {200705241557.l4OFv9Y7028726-at-ns.microscopy.com}
2, 24 -- Sender: "Leona Cohen-Gould" {lcgould-at-med.cornell.edu}
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From: beth-at-plantbio.uga.edu
Date: Thu, 24 May 2007 11:54:44 -0500
Subject: [Microscopy] Photonics telephone solicitations

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

On a similar note:
I received a strange call from the publication Photonics Spectra two
days ago.
They read all my address info to me then asked me "What color are
your eyes?".
I told the woman that was a totally inappropriate question and she
said they made them ask.
I hung up - it was too weird.

Made me wonder if anyone else has had a bizarre exchange with them.

Beth

On May 24, 2007, at 11:57 AM, mcauliff-at-umdnj.edu wrote:

}
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} Dear Lists:
}
} I got another telephone call at my lab yesterday from a solicitor.
} My phone number here is unlisted and does not show up on any caller
} id.
} I am getting 1 or 2 of these calls a week, too many to be random
} hits. I
} think some companies are harvesting my number from Listserv postings.
} Has anyone else had this experience?
}
} Geoff
}
} --
} --
} **********************************************
} Geoff McAuliffe, Ph.D.
} Neuroscience and Cell Biology
} Robert Wood Johnson Medical School
} 675 Hoes Lane, Piscataway, NJ 08854
} voice: (732)-235-4583; fax: -4029
} mcauliff-at-umdnj.edu
} **********************************************
}
}
}
} ==============================Original
} Headers==============================
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} id l4OFuU5T027644
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} 10:56:31 -0500
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} 11:56:30 -0400 (EDT)
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} 11:45:51 -0400 (EDT)
} 6, 33 -- Received: from ([130.219.34.131])
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} 6, 33 -- Thu, 24 May 2007 11:45:19 -0400
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} Polaris.umdnj.edu
} 6, 33 -- (iPlanet Messaging Server 5.2 HotFix 2.02 (built Oct 21
} 2004))
} 6, 33 -- id {0JIJ00L01Y9LKS-at-Polaris.umdnj.edu} (original mail from
} mcauliff-at-umdnj.edu)
} 6, 33 -- for microscopy-at-msa.microscopy.com; Thu, 24 May 2007
} 11:45:19 -0400 (EDT)
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} 6, 33 -- 2004)) with ESMTP id {0JIJ00EFGYFI9N-at-Polaris.umdnj.edu} ;
} Thu,
} 6, 33 -- 24 May 2007 11:45:19 -0400 (EDT)
} 6, 33 -- Date: Thu, 24 May 2007 11:45:18 -0400
} 6, 33 -- From: Geoff McAuliffe {mcauliff-at-umdnj.edu}
} 6, 33 -- Subject: telephone solicitations
} 6, 33 -- To: MicroscopyListserver {microscopy-at-msa.microscopy.com} ,
} 6, 33 -- Histonet {histonet-at-pathology.swmed.edu}
} 6, 33 -- Message-id: {4655B30E.6030502-at-umdnj.edu}
} 6, 33 -- MIME-version: 1.0
} 6, 33 -- Content-type: text/plain; format=flowed; charset=ISO-8859-1
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} 6, 33 -- X-Accept-Language: en-us, en
} 6, 33 -- User-Agent: Mozilla/5.0 (Windows; U; Windows NT 5.1; en-
} US; rv:1.7.2)
} 6, 33 -- Gecko/20040804 Netscape/7.2 (ax)
} ==============================End of -
} Headers==============================

**********************************************************************
Beth Richardson
Electron Microscopy Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

http://www.plantbio.uga.edu/emlab/

Phone - (706) 542-1790 & FAX - (706) 542-1805

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***
The Friends of the Marine Institute - Join Today!
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==============================Original Headers==============================
14, 21 -- From beth-at-plantbio.uga.edu Thu May 24 11:54:43 2007
14, 21 -- Received: from dogwood.plantbio.uga.edu (dogwood.plantbio.uga.edu [128.192.26.2])
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14, 21 -- Cc: microscopy microscopy {microscopy-at-microscopy.com}
14, 21 -- Content-Transfer-Encoding: 7bit
14, 21 -- From: Beth Richardson {beth-at-plantbio.uga.edu}
14, 21 -- Subject: Photonics telephone solicitations
14, 21 -- Date: Thu, 24 May 2007 12:54:42 -0400
14, 21 -- To: mcauliff-at-umdnj.edu
14, 21 -- X-Mailer: Apple Mail (2.752.3)
==============================End of - Headers==============================




From: mager-at-interchange.ubc.ca
Date: Thu, 24 May 2007 11:54:47 -0500
Subject: [Microscopy] RE: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
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Dear Michael,
I would check on that. I know that my Hitachi S-3000N, although it has no
Polaroid camera, still bases its magnification on a 4" X 5" image printout.
I always remember that a full-screen image is showing about twice the stated
magnification.
Regards,


Mary Fletcher (nee Mager)
Electron Microscopist
Materials Eng. UBC
#309 - 6350 Stores Road
Vancouver, B.C. V6T 1Z4
Canada
Tel: 604-822-5648
Fax: 604-822-3619
email: mager-at-interchange.ubc.ca

-----Original Message-----
X-from: michael-at-shaffer.net [mailto:michael-at-shaffer.net]
Sent: May 24, 2007 4:44 AM
To: mager-at-interchange.ubc.ca

Phil writes ...

} Our nanoscience program received some information about the
} Phenom-Ed from FEI, a tablettop SEM designed for education.
} We are considering whether it would useful for students to
} perform quick analysis of nanomaterials. ...

Regarding "nanomaterials", keep in mind that magnifications are not what
they used to be. For example, it used to be that a SEM's mag number was
based on a 4x5 polaroid, whereas now the mag number is based on a 10" window
on the computer display. The white paper describes the Phenom capable of
mag=20,000x which, relative to SEMs we are used to, would equate to 10,000x
(1 cm = 1,000 nM on a 4x5).

Not to take away from what should be a very valuable and interesting tool in
a science program for many high schools and colleges, and I only comment
relative to the term "nanomaterials".

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
Inco Innovation Centre
Memorial University
St. John's, Newfoundland



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From: bozzola-at-siu.edu
Date: Thu, 24 May 2007 12:37:42 -0500
Subject: [Microscopy] Re: Photonics telephone solicitations

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Although I no longer subscribe to a number of "freebie" magazines,
this question is one that is very commonly asked. Other questions
might include: place of birth, hair color, height, etc. I suppose it
is some way to moitor if the solicitors are connecting with the
clients. I have never had a followup call from a supervisor, however,
verifying the data.
}
} On a similar note:
} I received a strange call from the publication Photonics Spectra two
} days ago.
} They read all my address info to me then asked me "What color are
} your eyes?".
} I told the woman that was a totally inappropriate question and she
} said they made them ask.
} I hung up - it was too weird.
}
} Made me wonder if anyone else has had a bizarre exchange with them.
}
} Beth
}
} On May 24, 2007, at 11:57 AM, mcauliff-at-umdnj.edu wrote:
}
} }
} }
} }
} } ----------------------------------------------------------------------

--
+++++++++++++++++++++++++++++++++++++++++++++++++++++++

John J. Bozzola, Ph.D., Director
Integrated Microscopy & Graphics Expertise (IMAGE)
Southern Illinois University
750 Communications Drive - MC 4402
Carbondale, IL 62901
Telephone: 618-453-3730

+++++++++++++++++++++++++++++++++++++++++++++++++++++++

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From: stefan.diller-at-t-online.de
Date: Thu, 24 May 2007 13:04:41 -0500
Subject: [Microscopy] Absinthe animal - follow-up...

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Dear All,
see some new images at
www.elektronenmikroskopie.info/absinthe_animal/index.htm

Thanks for all your emails helping me solve this riddle. :-))


Kind regards,
Stefan


----------------------------------------------------------------------------
-----------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49 - 931 - 7848700 Phone
++49 - 931 - 7848701 Fax
++49 - 175 - 717 70 51 Cell-Phone

Websites:
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From: john.mardinly-at-intel.com
Date: Thu, 24 May 2007 13:07:56 -0500
Subject: [Microscopy] Re: telephone solicitations

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I get telephone solicitations from mortgage brokers on a daily basis. I
am on the no-call list at home, but I don't think this applies to a work
phone. Sure is annoying.

John Mardinly
Intel

This does not represent an opinion of Intel Corporation.

-----Original Message-----
X-from: lcgould-at-med.cornell.edu [mailto:lcgould-at-med.cornell.edu]
Sent: Thursday, May 24, 2007 9:16 AM
To: Mardinly, John

Geoff-
I haven't been getting calls, but I do know that if I "Google"
myself (don't we all do that sometimes?), I get pages of information,
some of which is from my facilities' websites, some reference
publications, but many hits are postings to this list. I am also
frequently asked about my pricing for services, based on a price list
that hasn't been on the facility's website in years, but persists out
there on the www. We all must remember that there is no way to
"erase" things once they hit cyberspace: a fact many new college
grads are finding out when prospective employers find their
embarrassing "My Space" postings.
If these calls bother you, may I suggest that you create a alternate
"signature" for your emails to the list, omitting your phone number.
MSA members can find you from the member directory if we need to.
That listing is password protected so outsiders can't get the info
from there. Just remember, those of us who are frequent users of the
listserver have put ourselves out in public through no fault of our
wonderful Sysop, or anyone else. Its just the nature of the beast.
Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Weill-Cornell Medical College

voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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From: oshel1pe-at-cmich.edu
Date: Thu, 24 May 2007 13:39:46 -0500
Subject: [Microscopy] Re: Absinthe animal - follow-up...

Contents Retrieved from Microscopy Listserver Archives
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Not a Tardigrade.
The features I'd really like to see are obscured, but 04 looks to
show the prosoma of an eriophyid mite. The other images fit this,
especially the setation and cuticular decorations.
A good example is at
http://www.insectimages.org/browse/detail.cfm?imgnum=1318030
(Not by me.) There are other good images on this site as well.

The hand-like feature is really nifty, but if it were claws of a
tardigrade, they'd be on the ends of all limbs, and there would be 2
pairs of limbs at the posterior end. 04 looks to show

Phil

} Dear All,
} see some new images at
} www.elektronenmikroskopie.info/absinthe_animal/index.htm
}
} Thanks for all your emails helping me solve this riddle. :-))
}
}
} Kind regards,
} Stefan
}
}
} ----------------------------------------------------------------------------
} -----------------------------------------
} Stefan Diller - Scientific Photography
} Arndtstrasse 22
} D - 97072 Wuerzburg Germany
} ++49 - 931 - 7848700 Phone
} ++49 - 931 - 7848701 Fax
} ++49 - 175 - 717 70 51 Cell-Phone
}
} Websites:
} www.stefan-diller.com
} www.elektronenmikroskopie.info
} www.assisi.de
} Anfahrt: http://mail.map24.com/stefan.diller
} ----------------------------------------------------------------------------
--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859

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From: bozzola-at-siu.edu
Date: Thu, 24 May 2007 17:49:52 -0500
Subject: [Microscopy] RE: magnification and Phenom-Ed SEM

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That's an excellent point about magnification. I hadn't been seriously shopping for an SEM in a while and just noticed that issue for the first time at MSA last year. Our two older scopes are both setup to record the magnification as for a 120-mm Polaroid print. I noticed that features in 50kx images in the new scopes were not as big as they were in our scopes. It was explained that magnification was now calculated according to displayed size which was seldom only 120 mm across.
 
When did the new rule take effect? I have long been accustomed to interpreting SEM images as 120mm across as for the old Polaroid film. It didn't matter that the image was displayed on a CRT twice as large. I just assumed magnification was calculated according to the print. Maybe it phased in as Polaroids phased out.
 
Warren
________________________________________
X-from: michael-at-shaffer.net [mailto:michael-at-shaffer.net]
Sent: Thu 5/24/2007 6:37 AM
To: wesaia-at-iastate.edu

Hi Warren,

I don't think that there was a "rule" that changed. As long as the first document from a microscope was relatively inflexible (a negative or Polaroid), it was quite easy to go back and get good numbers with information like "mag: 10,000x". This already changed when people started to use computers to look at images. If you reduce the image size by a factor of 2 (to insert in a publication, for example), you have already gone from 10,000x to 5,000x. Now, with digital acquisition and the flexibility to acquire a certain number of "pixels", the "magnification" has become a dangerous concept. If you received an image in the mail and it says "10,000x", you would have to know it's entire history to judge the validity of the magnification and make measurements.

A better number is probably one that does not assume a certain size of the final image to specify dimensions. We used to specify the "full width" of the image in microns or nm, today with the digital images, it is better to specify the dimension of a single pixel. That can then be used to directly measure or to display a scale bar, or even to calcualte the real magnification on paper.

The advantage of using magnification is that it immediately gives you an idea of size. If you look at something that was acquired at 100,000x, you know that it must be small.


Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
X-from: wesaia-at-iastate.edu [mailto:wesaia-at-iastate.edu]
Sent: Thursday, May 24, 2007 15:23
To: Mike Bode

That's an excellent point about magnification. I hadn't been seriously shopping for an SEM in a while and just noticed that issue for the first time at MSA last year. Our two older scopes are both setup to record the magnification as for a 120-mm Polaroid print. I noticed that features in 50kx images in the new scopes were not as big as they were in our scopes. It was explained that magnification was now calculated according to displayed size which was seldom only 120 mm across.
 
When did the new rule take effect? I have long been accustomed to interpreting SEM images as 120mm across as for the old Polaroid film. It didn't matter that the image was displayed on a CRT twice as large. I just assumed magnification was calculated according to the print. Maybe it phased in as Polaroids phased out.
 
Warren
________________________________________
X-from: michael-at-shaffer.net [mailto:michael-at-shaffer.net]
Sent: Thu 5/24/2007 6:37 AM
To: wesaia-at-iastate.edu

Magnification, as you have indicated, can be a
dangerous concept especially when one deals with
digital images (in TEM, SEM, and LM). In our
beginning microscopy classes, we try to emphasize
that the micrometer bar is what is to be
displayed on the image, never the magnifaction.
However, as a relative means of orientation, one
could state in the legend that the original image
was examined at a instrumental mangification of
10,000x, etc.

JB


} Hi Warren,
}
} I don't think that there was a "rule" that
} changed. As long as the first document from a
} microscope was relatively inflexible (a negative
} or Polaroid), it was quite easy to go back and
} get good numbers with information like "mag:
} 10,000x". This already changed when people
} started to use computers to look at images. If
} you reduce the image size by a factor of 2 (to
} insert in a publication, for example), you have
} already gone from 10,000x to 5,000x. Now, with
} digital acquisition and the flexibility to
} acquire a certain number of "pixels", the
} "magnification" has become a dangerous concept.
} If you received an image in the mail and it says
} "10,000x", you would have to know it's entire
} history to judge the validity of the
} magnification and make measurements.
}
} A better number is probably one that does not
} assume a certain size of the final image to
} specify dimensions. We used to specify the "full
} width" of the image in microns or nm, today with
} the digital images, it is better to specify the
} dimension of a single pixel. That can then be
} used to directly measure or to display a scale
} bar, or even to calcualte the real magnification
} on paper.
}
} The advantage of using magnification is that it
} immediately gives you an idea of size. If you
} look at something that was acquired at 100,000x,
} you know that it must be small.
}
}
} Michael Bode, Ph.D.
}
} General Manager
}
} OLYMPUS SOFT IMAGING SOLUTIONS
} 12596 West Bayaud Ave #300
} Lakewood, CO 80228
} USA
} Tel.: +1 (303) 234-9270
} Fax.: +1 (303) 234-9271
} E-mail: Mike.Bode-at-olympus-sis.com
} www.olympus-sis.com
}
} -----Original Message-----
} X-from: wesaia-at-iastate.edu [mailto:wesaia-at-iastate.edu]
} Sent: Thursday, May 24, 2007 15:23
} To: Mike Bode
} Subject: [Microscopy] Phenom-Ed SEM
}
}
}
}
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From: vlk7-at-alfred.edu
Date: Thu, 24 May 2007 18:09:34 -0500
Subject: [Microscopy] AskAMicroscopist: help with Image-Pro plus

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (vlk7-at-alfred.edu) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Thursday, May 24, 2007 at 09:18:07
---------------------------------------------------------------------------

Email: vlk7-at-alfred.edu
Name: Victoria Knox

Organization: Alfred University

Education: Undergraduate College

Location: Alfred, NY USA

Question: I've been working with a program called Image-Pro plus to measure the diameter of micron-sized fibers from SEM micrographs. I need some assistance in how the program is measureing these fibers because I am getting inaccurate data that is off by at least 25% each time. I have tried to get the data using different measurements but it is still off. Any help you can provide would be greatly appreciated.

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From: kay-at-natural-immunogenics.com
Date: Thu, 24 May 2007 18:16:30 -0500
Subject: [Microscopy] viaWWW: TEM PHILLIPS EM400 help

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Email: kay-at-natural-immunogenics.com
Name: Kay Mitchen

Title-Subject: [Filtered] TEM PHILLIPS EM400

Question: We are looking for training of 2 additional operators and maintenance of a TEM reference Phillips EM 400

Please contact me urgently - feel free to call at 954/979-0885

PS: We are located in Pompano Beach, Florida

Thanks

Kay E. Mitchen

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==============================Original Headers==============================
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From: dianavd-at-eye.usyd.edu.au
Date: Thu, 24 May 2007 20:17:20 -0500
Subject: [Microscopy] lead citrate and block staining

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I found your message particularly interesting since you thought that the
results deviated from an already known value by 25%.

As you seem to have found out rather quickly, image processing packages
rarely if ever divulge the methods they employ to obtain a result.

Fibers can be difficult to work with unless some assumptions are made about
the shape of the fiber. A fiber with an elliptical cross section can appear
circular, and a circular cross section appears to be elliptical in all but
one possible intersection. Non-convex fibers can be a real dilemma.

Suppose that you could measure the diameters accurately - right to the last
possible digit of precision. If the fibers are slightly different in size,
then you see a range of sizes. If you happen to look at only a few fibers or
possibly a set of fibers with a particular diameter, then you might be led
to think that the measurements are off.

The problem you are investigating may not be as simple or as knowable as you
might have hoped. So let's see what you are doing and maybe collectively we
can assist you in finding out what is happening.

Cheers
----- Original Message -----
X-from: {vlk7-at-alfred.edu}
To: {rboehrin-at-vt.edu}
Sent: Thursday, May 24, 2007 7:14 PM

I've just read a paper where the author talks about block staining
with uranyl acetate and lead citrate. UAc yes, but Pbcitrate? Has
anyone heard of this? Does it work? The pictures in the paper were
very nice; good contrast and detail. With the recent talk about
general lack of contrast in specimens these days (I quite agree on
that; I find contrast poorer now than in the past), could this be a
new method?

All opinions please!

Cheers,

Diana

==============================Original Headers==============================
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From: Elliott-at-arizona.edu
Date: Thu, 24 May 2007 21:13:19 -0500
Subject: [Microscopy] Re: lead citrate and block staining

Contents Retrieved from Microscopy Listserver Archives
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Hi Diana
I use Pb in block and not on the grid. Works great. NO lead
pepper. See my article in Microscopy Today, January 2007.
David

_____________________

David Elliott Ph.D.
Assistant Professor - Department of Cell Biology and Anatomy
Director, Research Microscopy Core Service
University of Arizona College of Medicine
PO Box 245004
Tucson, AZ 85724

Voice: 520-626-7870
Fax: 520-626-2097



On May 24, 2007, at 6:20 PM, dianavd-at-eye.usyd.edu.au wrote:

}
}
}
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}
} I've just read a paper where the author talks about block staining
} with uranyl acetate and lead citrate. UAc yes, but Pbcitrate? Has
} anyone heard of this? Does it work? The pictures in the paper were
} very nice; good contrast and detail. With the recent talk about
} general lack of contrast in specimens these days (I quite agree on
} that; I find contrast poorer now than in the past), could this be a
} new method?
}
} All opinions please!
}
} Cheers,
}
} Diana
}
} ==============================Original
} Headers==============================
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} 4, 18 -- Date: Fri, 25 May 2007 11:17:10 +1000
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From: yamawaki-at-stanford.edu
Date: Thu, 24 May 2007 23:05:12 -0500
Subject: [Microscopy] viaWWW: EM service technician

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Email: yamawaki-at-stanford.edu
Name: Ruth Yamawaki

Organization: Stanford University

Title-Subject: [Filtered] EM service technician

Question: I am looking for a an EM service technician. If you know of an independent EM service technician for a JEOL electron microscope who services EMs in California please contact me.

Thank you,

Ruth Yamawaki

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From: gas19-at-daimlerchrysler.com
Date: Thu, 24 May 2007 23:07:00 -0500
Subject: [Microscopy] viaWWW: AskAMicroscopist: help with Image-Pro plus

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Email: gas19-at-daimlerchrysler.com
Name: Gerald Shulke

Organization: DaimlerChrysler

Title-Subject: [Filtered] Re: [Microscopy] AskAMicroscopist: help with Image-Pro plus

Question: Victoria,

We use Image Pro Plus for measurements too. Feel free to send me an e-mail off line with a reference picture, and I can try to help you. Between the 6 of us that use it here, we could figure it out. It could be the aspect ratio for pixels in your calibration set-up, so that a fiber not parallel wth the x-axis of the picture would have an incorrect measurement. it could also be that you are transforming the picture (different size or resolution of the image) which is different from your calibration.

Gerald Shulke
Materials Engieering Specialist
gas19-at-dcx.com

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From: nizets2-at-yahoo.com
Date: Fri, 25 May 2007 02:51:46 -0500
Subject: [Microscopy] Re: Absinthe animal - follow-up...

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The problem with your beast is that I cannot recognize
feet. This suggests me that it may be not be an adult
but a larva. In thise case "Viel Glück" ;-)

LG
Stephane

--- stefan.diller-at-t-online.de wrote:

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www.elektronenmikroskopie.info/absinthe_animal/index.htm
}
} Thanks for all your emails helping me solve this
} riddle. :-))
}
}
} Kind regards,
} Stefan
}
}
}
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From: nizets2-at-yahoo.com
Date: Fri, 25 May 2007 02:59:42 -0500
Subject: [Microscopy] Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Sure, but if you see a scale bar next to your
feature, you don't know it is small, you know its size
directly! (and if the full scale bar is 50 nm, you
know it is
very small)

Stephane


} --- Mike.Bode-at-olympus-sis.com wrote:
}
} }
} } The advantage of using magnification is that it
} } immediately gives you an idea of size. If you look
} } at something that was acquired at 100,000x, you
} know
} } that it must be small.
} }
} }
} } Michael Bode, Ph.D.
} }
} } General Manager
} }
} } OLYMPUS SOFT IMAGING SOLUTIONS
} } 12596 West Bayaud Ave #300
} } Lakewood, CO 80228
} } USA
} } Tel.: +1 (303) 234-9270
} } Fax.: +1 (303) 234-9271
} } E-mail: Mike.Bode-at-olympus-sis.com
} } www.olympus-sis.com
} }
}
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}
}
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From: Walter.Bobrowski-at-pfizer.com
Date: Fri, 25 May 2007 07:34:58 -0500
Subject: [Microscopy] Myelin Osmication Problem

Contents Retrieved from Microscopy Listserver Archives
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Hi Karen! Long time, no chat!
In my experience with rat sciatic nerve, my pieces were several
millimeters long (oriented for x-section view), but tended to be less
wide than spinal cord, so I typically had great osmium penetration.
However, I noted that several samples that were very wide tended to have
poor osmium penetration throughout, particularly if I sectioned too
deeply.

Keeping that in mind, I suggest thinner tissue samples (no longer than
0.5mm), either by hand or vibratome. Secondly, minimize tissue loss
during your initial block face trimming and obtain your sections ASAP.

Best regards,

Walter F. Bobrowski
Sr. Scientist
Pfizer Global R&D
2800 Plymouth Rd.
Ann Arbor, MI 48105

TEL: 734-622-7814
FAX: 734-622-5718

"Keep away from small people who try to belittle your ambitions. Small
people always do that, but the really great make you feel that you, too,
can become great."
Mark Twain
1835-1910, Humorist and Writer



-----Original Message-----
X-from: Karen_Bentley-at-URMC.Rochester.edu
[mailto:Karen_Bentley-at-URMC.Rochester.edu]
Sent: Thursday, May 24, 2007 10:37 AM
To: Bobrowski, Walter



Hello Fellow Listers:

I have an issue regarding perfusion fixed (glut & paraform. in
cacodylate buffer) adult rat spinal cord.

I have cut cord cross sections approx. 1-1.5 mm. thick which were
osmicated in a 1.0% OsO4/cacodylate buffer for 1.5 hrs. When I cut into
the middle of the tissue block it was completely white.......the osmium
did not penetrate beyond the surface of the tissue. I know that the
heavily myelinated spinal cord can be a barrier to complete diffusion of
osmium post fixation. These blocks will be sectioned for light
microscopy stained with Toluidine Blue to access myelin fiber numbers,
then potentially TEM examination.

Question: I have researched the literature and one paper suggested
using 2.0% osmium and leaving the spinal cord blocks in overnight.
Another suggested using potassium ferrocyanide with the osmium. One of
our university EM experienced researchers suggested warming the osmium
to 37 C. Any help would be greatly appreciated as I need an answer
quickly.

Karen Bentley

Karen L. Bentley, M.S.
Technical Director
Electron Microscope Research Core
University of Rochester Medical Center
575 Elmwood Avenue, Box 626
Rochester, NY 14642
585-275-1954


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From: oshel1pe-at-cmich.edu
Date: Fri, 25 May 2007 07:53:43 -0500
Subject: [Microscopy] Re: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Stephane,

No, you don't know the size directly. Not unless the measured object
is exactly orthogonal to the projected sight line.
Quick experiment: project a bright light on the wall, and hold up a
pencil (or similar). Hold the pencil so its long axis is exactly at
90 degrees to the central axis of the light path. Pretend the
enlarged size of the pencil shadow is magnification.
Now, tip the pencil into different orientations. Notice how the size
of the shadow changes? Projective geometry.
Same thing with an SEM image. It's a 3D object whose image is
projected onto a 2D plane. Unless the dimension desired is exactly
orthongonal to the line of sight, then the projected size will be
different than the true size by an unknown amount.
This can be solved by tilting the image, taking stereopairs, and
doing trigonometry.
But it will still be wrong unless the instrument has been properly
calibrated *and* unless the all of the effects the specimen
processing steps have on the sample are exactly known.
Mind, this all ignores the more subtle question of "where's the
edge"? Given the signal that forms an SEM image come from an
interaction volume, not a dimensionless point, then the edge of a
feature is essentially smeared out over the size of that volume. This
is not much of an issue, or even no issue, for most imaging, but at
high magnifications and resolutions, it is.

Phil

} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

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From: david.wilbur-at-tufts.edu
Date: Fri, 25 May 2007 08:04:12 -0500
Subject: [Microscopy] Can I use my SEM sputter coater for metals other than

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I have a Polaron SC-502 sputter coater in the SEM lab. One of the
Engineering faculty wants to coat samples with chromium, titanium and
aluminum for an application other than SEM. Assuming I can get targets
that fit the sputter coater, are there other issues that would prevent
using these other metals?

Thank You

Dave Wilbur
--
__________________________________
David J. Wilbur, Ph.D.
Instrumentation Specialist
Department of Chemistry
Tufts University
62 Talbot Ave.
Medford, MA 02155
voice: 617-627-2163
Fax: 617-627-3443
email: david.wilbur-at-tufts.edu
__________________________________

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From: Robert.Zonis-at-sanford.com
Date: Fri, 25 May 2007 08:09:46 -0500
Subject: [Microscopy] Re: AskAMicroscopist: help with Image-Pro plus

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

We've been working rather intensively with PAX-it here for microscopy. I've done some image analysis work with Image-Pro, but I can't answer definitively about how they are doing their measurements. PAX-it's method for determining measurements is quite simple and easy to understand, however, and I'm sure that Chris Jahns up at MIS [mailto:chrisj-at-paxit.com] would be happy to take a look at your images and attempt to give you some dimensions.

PAX-it works quite simply by counting pixels and comparing them to a known reference that you have calibrated the program with at a specific "magnification" that you set, i.e., for light microscopy, you put a stage micrometer onto the microscope stage, choose your objective, focus, and capture the image. You tag that image with your parameters, and input those parameters into the database. Then you calibrate the measurement module by indicating precisely where your units of measurements are according to the micrometer. This gives PAX-it the number of pixels per micron, inch, etc. both horizontally and vertically, and the program will simply count pixels between designated measurement points and apply that specific conversion factor every time you present it with an image that you have designated as being captured with those particular parameters.

I'm not sure how people calibrate measurement distances with a SEM, but I do know that there are reference target particles used for calibrating particle size analyzers. If you could capture an image from silica beads, for example, with a known diameter of 10 microns, for example, you should be able to tell the accuracy of your system.


Robert Zonis
Product Development Chemist, LMTC
Sanford L.P. - A Newell Rubbermaid Company
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From: TindallR-at-missouri.edu
Date: Fri, 25 May 2007 08:26:25 -0500
Subject: [Microscopy] Can I use my SEM sputter coater for metals other

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

We use our Emitech coater to coat with a variety of metals, including
chromium, platinum, tantalum, and others. You will have to check on
sputtering currents and times and how they relate to coating thickness
for whatever metal you're using, but I'm not personally aware of any
reason you can't use the Polaron for a variety of metals.

My real reason for replying is to get one of the best-kept secrets in
the World of Sputtering out there. You can get custom-made targets of
virtually any metal from Refining Systems, Inc. in Las Vegas, NV. These
targets can be made to your specified diameters, shapes and thicknesses,
usually at substantially less cost than targets from coater
manufacturers. The company is run by Mr. Abe Dayani, who has always
answered the phone himself when I've called. Website is at
http://www.refiningsystems.com/, phone is 702-368-0579.

I have heard that the purity of the metals used by the OEMs may be
higher (like 99.998 vs. 99.99, for example), but I can say that we have
never had any problem with the coatings from the RFI targets. It may be
a factor for super-critical applications in engineering, so you may want
to consider this.

I have no connection with RFI other than as a satisfied customer.

Cheers,
Randy

Randy Tindall
Senior EM Specialist
Electron Microscopy Core Facility---We Do Small Well!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu
On-line calendar:
http://biotech.rnet.missouri.edu/cgi-bin/Calcium310.pl?Op=Splash&Amount=
Week&NavType=Both&Type=TimePlan

-----Original Message-----
X-from: david.wilbur-at-tufts.edu [mailto:david.wilbur-at-tufts.edu]
Sent: Friday, May 25, 2007 8:05 AM
To: Tindall, Randy D.

I have a Polaron SC-502 sputter coater in the SEM lab. One of the
Engineering faculty wants to coat samples with chromium, titanium and
aluminum for an application other than SEM. Assuming I can get targets
that fit the sputter coater, are there other issues that would prevent
using these other metals?

Thank You

Dave Wilbur
--
__________________________________
David J. Wilbur, Ph.D.
Instrumentation Specialist
Department of Chemistry
Tufts University
62 Talbot Ave.
Medford, MA 02155
voice: 617-627-2163
Fax: 617-627-3443
email: david.wilbur-at-tufts.edu
__________________________________

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Can I use my SEM sputter coater for metals other than 3, 20 -- gold?
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From: dljones-at-bestweb.net
Date: Fri, 25 May 2007 08:32:22 -0500
Subject: [Microscopy] Re: Can I use my SEM sputter coater for metals other than

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dave,

I am not familiar with the specific sputter coater you are using. So here
are some questions. Do you know if it is a straight DC sputter coater
or magnitron sputter coater? What is the pumping system, just a mechanical
pump?

Most sputter coaters for SEM sample prep only use a mechanical pump for
vacuum generation. This means you are only going to about 10-3 torr of
vacuum before backfilling and then coating. For most thin film
depositions, this is quite poor vacuum. Most thin film sputter coaters
will go to somewhere around 10-6 to 10-8 torr before backfilling for
sputter deposition. They also usually have a way to clean the substrate
prior to deposition, which your little sputter coater I imagine does not
have. The amount of contamination you will have, both on the first coat
and then when switching between targets will be much larger than usually
considered acceptable for thin film deposition.

All that being said, I don't see any reason you can't use your machine to
deposit the materials you are mentioning, but I do question how useful the
resulting layered structure will end up being. I would have to know a lot
more about what the desired end product is to give you my thoughts on
that.

dj

On Fri, 25 May 2007, david.wilbur-at-tufts.edu wrote:

}
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} I have a Polaron SC-502 sputter coater in the SEM lab. One of the
} Engineering faculty wants to coat samples with chromium, titanium and
} aluminum for an application other than SEM. Assuming I can get targets
} that fit the sputter coater, are there other issues that would prevent
} using these other metals?
}
} Thank You
}
} Dave Wilbur
} --
} __________________________________
} David J. Wilbur, Ph.D.
} Instrumentation Specialist
} Department of Chemistry
} Tufts University
} 62 Talbot Ave.
} Medford, MA 02155
} voice: 617-627-2163
} Fax: 617-627-3443
} email: david.wilbur-at-tufts.edu
} __________________________________
}
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From: Andrew.Bowling-at-ars.usda.gov
Date: Fri, 25 May 2007 08:35:40 -0500
Subject: [Microscopy] AskAMicroscopist: help with Image-Pro plus

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I recommend that you download and install NIH Image (a.k.a. ImageJ).
Using the same calibration image you used for ImagePro, does it give you
the same numbers? If so, you might want to image a new calibration
specimen. If it's still not what you expected, at least you'll know it
isn't an error in the software. You might be seeing shrinkage due to
specimen processing (critical point drying, etc.), exposure to vacuum,
the 3d projection errors mentioned in another post, etc.

-----Original Message-----
X-from: vlk7-at-alfred.edu [mailto:vlk7-at-alfred.edu]
Sent: Thursday, May 24, 2007 5:15 PM
To: Bowling, Andrew

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (vlk7-at-alfred.edu) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
Thursday, May 24, 2007 at 09:18:07
------------------------------------------------------------------------
---

Email: vlk7-at-alfred.edu
Name: Victoria Knox

Organization: Alfred University

Education: Undergraduate College

Location: Alfred, NY USA

Question: I've been working with a program called Image-Pro plus to
measure the diameter of micron-sized fibers from SEM micrographs. I need
some assistance in how the program is measureing these fibers because I
am getting inaccurate data that is off by at least 25% each time. I have
tried to get the data using different measurements but it is still off.
Any help you can provide would be greatly appreciated.

------------------------------------------------------------------------
---

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From: Walter.Bobrowski-at-pfizer.com
Date: Fri, 25 May 2007 08:53:20 -0500
Subject: [Microscopy] Re: monkey perfusion methodology

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Thanks Larry. Actually I concur with your experience with small animals
(mice: 5-8 ml/min; rats: 10-20 ml/min). Interestingly I supported a
colleague who needed monkey brain perfusion assistance and we simply
followed a recommended protocol they had in hand (~300 ml/min). They
were delighted with the histology results. Even I was shocked as the
quality of preservation in spite of the high flow rate, as we typically
recommend ~100 ml/min for large animals. I'm attempting to track down
the reference.

In addition, Microscopy Today (May 2006) had a very relevant article on
animal brain perfusion, advocating precise pressures, rather than flow
rate, including a short bolus at 300 mm Hg, followed by 100 mm Hg.

Best regards,
Walt


-----Original Message-----
X-from: larry.ackerman-at-ucsf.edu [mailto:larry.ackerman-at-ucsf.edu]
Sent: Wednesday, May 23, 2007 1:49 PM
To: Bobrowski, Walter

Mary Ellen,
The rate that Walter suggested seemed extremely high compared to my
experience with smaller animals so I asked an anatomist with many
decades of experience with monkeys. He has always used an initial flush
of 500ml in 5 minutes time with buffer followed by 1.5 liters of
fixative over 20 minutes of time for a typical 5kg animal. The
appropriate size tubing and cannula in collaboration with the setting on

the pump will enable an excellent fixation. Walter's method may well
provide good results. I would love to see some brain slices and TEM
micrographs at 10,000X. I'm open to different methods.

Larry

Walter.Bobrowski-at-pfizer.com wrote:
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}
} 300 ml/min. Perfuse either through the carotid artery or left
ventricle
} (occlude descending aorta).
}
} Best regards,
} Walter F. Bobrowski
} Sr. Scientist
} Pfizer Global R&D
} 2800 Plymouth Rd.
} Ann Arbor, MI 48105
}
} TEL: 734-622-7814
} FAX: 734-622-5718
}
} "Like the strength of a steel rod, the true character of a person can
} only be known under extreme stress." - Leslie Fieger
}
}
}
}
} -----Original Message-----
} X-from: mpease-at-jhmi.edu [mailto:mpease-at-jhmi.edu]
} Sent: Monday, May 21, 2007 7:25 PM
} To: Bobrowski, Walter
} Subject: [Microscopy] viaWWW: monkey perfusion methodology
}
}
}
}
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} Email: mpease-at-jhmi.edu
} Name: Mary Ellen Pease
}
} Organization: Johns Hopkins Wilmer Eye Institute
}
} Title-Subject: [Filtered] monkey perfusion methodology
}
} Question: What is the recommended flow rate for perfusion of monkey
} brain and eyes when using a pump rather than gravity feed of the
} solutions?
}
} Thank you,
} Mary Ellen
}
}
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} ==============================Original
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--
Larry Ackerman, Associate Specialist
UCSF, Dept. of Anatomy, Rm S1347
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu


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From: fschamber-at-aspexcorp.com
Date: Fri, 25 May 2007 10:12:02 -0500
Subject: [Microscopy] RE: magnification and Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

In fear if you have a conventional sputter coater - not the high vacuum
versions that are designed for more difficult materials like chromium - you
will not be able to coat other than with the simple materials - gold,
platinum or gold-palladium.

Basically the everyday SEM sputter coater does not have the power.

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


----- Original Message -----
X-from: {david.wilbur-at-tufts.edu}
To: {protrain-at-emcourses.com}
Sent: Friday, May 25, 2007 2:05 PM

The convention of identifying magnification is one that seems to warrant some new attention in our digitized world. As several people have pointed out, there are several good conventions for reliably labeling the true scale for micrographs, the most common being the micron bar. That should probably remain the "gold standard" for publications, etc.

The familiar concept of "X" magnification is, of course, loaded with problems for digital images. It's not just the size of the instrument's viewing screen, but just what is the "magnification" of an image stored in a digital file? There has to be a point of reference, and historically that was the size of Polaroid film.

However, I think we should not casually dismiss the use of a "magnification" value as an identifier on our micrographs and our instruments. First of all, the use of "X" terminology is so solidly entrenched in our field that even if we don't like it, it's not going to go away anytime soon since it's just too convenient. The instrument's magnification knob has long been used as the means for setting up consistent operation, and was (in the Polaroid days) a pretty solid convention that you could count on -- a micrograph taken at 1000X on two different instruments would be generally comparable at least. Though I'm sure it's possible to write a procedure that specifies how to set the image scale in terms of the micron bar, it's much simpler to just say "set the control to 1000X." And if you're taking a series of micrographs at different magnifications, don't most of us label them like "10KX", "500X" etc.?

More fundamentally, it seems to me that the "X" concept speaks to something intrinsic about the way our brain perceives visual size. Think about a pair of binoculars that is specified as having 10X magnification. In that case, it is certainly not the absolute size of the recorded image that is being referenced (just what is the "size" of an image in one's mind?) but it relates to the perceived size of things. The convention for magnification in light optics is formally based on the angular deflection produced by the optics, but we perceive it as how much of the visual field is filled. I'll argue that the "X" value of SEM micrographs performs the same function. Thus, even if I see a "500X" micrograph reduced to a "thumbnail" or projected on a screen, I don't find it confusing since I'm gauging the size of the contents to the size of its frame. In other words, a term like "500X," though not sufficient for accurate size calibration in publications and the like, is still meaningful for making rapid assessments and comparisons, and I think we would all be well served if the convention of "X" magnification were better standardized. I'd propose two modest conventions:

(1) We adopt a standard terminology (perhaps "relative magnification") to refer to such "X" values, so everyone knows it is relative to the size of the field (not the viewing medium); and
(2) We adopt a standard reference field size convention such as 100mm for the smaller field dimension (close to the four inch dimension of Polaroid film) to which an "X" value always refers.

This should not replace micron bars or other more quantitative metrics, but an agreed convention would, I think, eliminate a lot of confusion that currently exists.


Fred Schamber
Aspex Corporation



-----Original Message-----
X-from: bozzola-at-siu.edu [mailto:bozzola-at-siu.edu]
Sent: Thursday, May 24, 2007 5:55 PM
To: Fred Schamber

Magnification, as you have indicated, can be a
dangerous concept especially when one deals with
digital images (in TEM, SEM, and LM). In our
beginning microscopy classes, we try to emphasize
that the micrometer bar is what is to be
displayed on the image, never the magnifaction.
However, as a relative means of orientation, one
could state in the legend that the original image
was examined at a instrumental mangification of
10,000x, etc.

JB


} Hi Warren,
}
} I don't think that there was a "rule" that
} changed. As long as the first document from a
} microscope was relatively inflexible (a negative
} or Polaroid), it was quite easy to go back and
} get good numbers with information like "mag:
} 10,000x". This already changed when people
} started to use computers to look at images. If
} you reduce the image size by a factor of 2 (to
} insert in a publication, for example), you have
} already gone from 10,000x to 5,000x. Now, with
} digital acquisition and the flexibility to
} acquire a certain number of "pixels", the
} "magnification" has become a dangerous concept.
} If you received an image in the mail and it says
} "10,000x", you would have to know it's entire
} history to judge the validity of the
} magnification and make measurements.
}
} A better number is probably one that does not
} assume a certain size of the final image to
} specify dimensions. We used to specify the "full
} width" of the image in microns or nm, today with
} the digital images, it is better to specify the
} dimension of a single pixel. That can then be
} used to directly measure or to display a scale
} bar, or even to calcualte the real magnification
} on paper.
}
} The advantage of using magnification is that it
} immediately gives you an idea of size. If you
} look at something that was acquired at 100,000x,
} you know that it must be small.
}
}
} Michael Bode, Ph.D.
}
} General Manager
}
} OLYMPUS SOFT IMAGING SOLUTIONS
} 12596 West Bayaud Ave #300
} Lakewood, CO 80228
} USA
} Tel.: +1 (303) 234-9270
} Fax.: +1 (303) 234-9271
} E-mail: Mike.Bode-at-olympus-sis.com
} www.olympus-sis.com
}
} -----Original Message-----
} X-from: wesaia-at-iastate.edu [mailto:wesaia-at-iastate.edu]
} Sent: Thursday, May 24, 2007 15:23
} To: Mike Bode
} Subject: [Microscopy] Phenom-Ed SEM
}
}
}
}
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--
+++++++++++++++++++++++++++++++++++++++++++++++++++++++

John J. Bozzola, Ph.D., Director
Integrated Microscopy & Graphics Expertise (IMAGE)
Southern Illinois University
750 Communications Drive - MC 4402
Carbondale, IL 62901
Telephone: 618-453-3730

+++++++++++++++++++++++++++++++++++++++++++++++++++++++


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From: Mike.Bode-at-olympus-sis.com
Date: Fri, 25 May 2007 10:24:22 -0500
Subject: [Microscopy] Re: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I think, you are both right :-)

Unless you have the scale bar right on an object, it will give you only
a good idea of the size and dimensions of an object, but the same is
true for "magnification". The difference is that a scale bar gives you
an immediate idea of the real size of an object, while magnification is
indirect and you have to convert it into a "virtual scale bar" first
("Let's see, the mag is 10,000x, so 1 cm on the image corresponds to 1
micron, so the object is roughly 4 microns long). When it comes to
actual measurements, you have to take additional steps.

As far as edges for measurement go, Philip os of course right. SEMs in
particular show edge effects that need to be modeled if accurate
measurements are desired at high resolution.

mike


Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
X-from: oshel1pe-at-cmich.edu [mailto:oshel1pe-at-cmich.edu]
Sent: Friday, May 25, 2007 06:58
To: Mike Bode

Stephane,

No, you don't know the size directly. Not unless the measured object is
exactly orthogonal to the projected sight line.
Quick experiment: project a bright light on the wall, and hold up a
pencil (or similar). Hold the pencil so its long axis is exactly at 90
degrees to the central axis of the light path. Pretend the enlarged size
of the pencil shadow is magnification.
Now, tip the pencil into different orientations. Notice how the size of
the shadow changes? Projective geometry.
Same thing with an SEM image. It's a 3D object whose image is projected
onto a 2D plane. Unless the dimension desired is exactly orthongonal to
the line of sight, then the projected size will be different than the
true size by an unknown amount.
This can be solved by tilting the image, taking stereopairs, and doing
trigonometry.
But it will still be wrong unless the instrument has been properly
calibrated *and* unless the all of the effects the specimen processing
steps have on the sample are exactly known.
Mind, this all ignores the more subtle question of "where's the edge"?
Given the signal that forms an SEM image come from an interaction
volume, not a dimensionless point, then the edge of a feature is
essentially smeared out over the size of that volume. This is not much
of an issue, or even no issue, for most imaging, but at high
magnifications and resolutions, it is.

Phil

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From: amich-at-ufl.edu
Date: Fri, 25 May 2007 10:48:11 -0500
Subject: [Microscopy] flat section release

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I would greatly appreciate advice on the following: I have flat
Epon embedded section sandwiched between Kapton (polyimide) film
and Permanox plastic slide. It seems that excess of epoxy run over
the slide sides and I have not been successful in film removal
which I never encountered previously with Aclar films. Can anyone
suggest an approach to release the section? Solvent, epoxy
removal, dip in liquid nitrogen?

Thank you in advance,

Albina


--
MIKHAYLOVA,ALBINA, PhD
Materials Science and Engineering
University of Florida
PO Box 116400
Gainesville, Florida 32611
Phone: (352) 392-6533
Fax: (352) 392-3771
E-Mail: amich-at-ufl.edu


==============================Original Headers==============================
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From: jquinn-at-www.matscieng.sunysb.edu
Date: Fri, 25 May 2007 11:50:13 -0500
Subject: [Microscopy] re: Fiber diameter

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Victoria

Post a typical image to a public web page.

Then resubmit your message to the listserver,
and include a link to the URL for the image.

Make sure to include a micron marker or
some other length calibration.

Do not send the image as an attachment.

In this manner, everyone will be able
to make measurements for you.

regards,

Jim


PS: OoO away........

} From mail-at-ns.microscopy.com Thu May 24 19:08:35 2007
} Date: Thu, 24 May 2007 18:10:04 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: vlk7-at-alfred.edu
} Reply-to: vlk7-at-alfred.edu
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
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} Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (vlk7-at-alfred.edu) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Thursday, May 24, 2007 at 09:18:07
} ---------------------------------------------------------------------------
}
} Email: vlk7-at-alfred.edu
} Name: Victoria Knox
}
} Organization: Alfred University
}
} Education: Undergraduate College
}
} Location: Alfred, NY USA
}
} Question: I've been working with a program called Image-Pro plus to measure the diameter of micron-sized fibers from SEM micrographs. I need some assistance in how the program is measureing these fibers because I am getting inaccurate data that is off by at least 25% each time. I have tried to get the data using different measurements but it is still off. Any help you can provide would be greatly appreciated.
}
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From: walck-at-southbaytech.com
Date: Fri, 25 May 2007 12:24:53 -0500
Subject: [Microscopy] Re: Can I use my SEM sputter coater for metals other

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Let me add a little to this discussion. Even if the coater would have a
high vacuum, there still are issues with these three materials. All of them
have an oxide coating on the targets. Ti and Al are very thin. Cr takes
longer to build a thicker oxide coating. You have to sputter the oxide away
before you can use it to deposit the film. Ti is a getter material and so
is Al to a lesser degree. What this means is that the thin film deposited
will act as a pump to pump reactive species such as O2, H2O, H2, from the
vacuum system. So you have to sputter for a significant time prior to
actually laying the film down so that these species are pumped from the
chamber and your film on your substrate does not act as a getter pump. Ti
and Al will instantaneously oxidize when the sample is brought up to air,
but it will be a very thin protective coat. This is important with respect
to the properties and the thickness of your film. Cr will take longer to
fully oxidize, but it will unless protected by something like our
SampleSaver(TM) container.

It would really help everything if your base pressure for your coater is at
least in the 10^-7 torr range or better.

You might consider putting a top layer of a very thin Ti onto these other
materials to act as a protective oxide layer.

Disclaimer: SBT makes and sells the IBS/e ion beam sputter and etch system
and the SampleSaver(TM) storage container.


-Scott

Scott D. Walck, Ph.D.
Technical Director
South Bay Technology, Inc.
1120 Via Callejon
San Clemente, CA 92673

US Toll Free: 1-800-728-2233
Tel: (949) 492-2600
Fax: (949) 492-1499

www.southbaytech.com
walck-at-southbaytech.com

-----Original Message-----
X-from: dljones-at-bestweb.net [mailto:dljones-at-bestweb.net]
Sent: Friday, May 25, 2007 6:36 AM
To: Walck-at-SouthBayTech.com

Dave,

I am not familiar with the specific sputter coater you are using. So here
are some questions. Do you know if it is a straight DC sputter coater or
magnitron sputter coater? What is the pumping system, just a mechanical
pump?

Most sputter coaters for SEM sample prep only use a mechanical pump for
vacuum generation. This means you are only going to about 10-3 torr of
vacuum before backfilling and then coating. For most thin film depositions,
this is quite poor vacuum. Most thin film sputter coaters will go to
somewhere around 10-6 to 10-8 torr before backfilling for sputter
deposition. They also usually have a way to clean the substrate prior to
deposition, which your little sputter coater I imagine does not have. The
amount of contamination you will have, both on the first coat and then when
switching between targets will be much larger than usually considered
acceptable for thin film deposition.

All that being said, I don't see any reason you can't use your machine to
deposit the materials you are mentioning, but I do question how useful the
resulting layered structure will end up being. I would have to know a lot
more about what the desired end product is to give you my thoughts on that.

dj

On Fri, 25 May 2007, david.wilbur-at-tufts.edu wrote:

}
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} I have a Polaron SC-502 sputter coater in the SEM lab. One of the
} Engineering faculty wants to coat samples with chromium, titanium and
} aluminum for an application other than SEM. Assuming I can get
} targets that fit the sputter coater, are there other issues that would
} prevent using these other metals?
}
} Thank You
}
} Dave Wilbur
} --
} __________________________________
} David J. Wilbur, Ph.D.
} Instrumentation Specialist
} Department of Chemistry
} Tufts University
} 62 Talbot Ave.
} Medford, MA 02155
} voice: 617-627-2163
} Fax: 617-627-3443
} email: david.wilbur-at-tufts.edu
} __________________________________
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From: mnesta-at-ebsciences.com
Date: Fri, 25 May 2007 12:43:08 -0500
Subject: [Microscopy] Re: Can I use my SEM sputter coater for metals other

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Dave,

The Polaron SC-502 is a great old coater, but unfortunately it only has
the ability to sputter Noble (non-oxidizing) Metals like Au, Pt and Pd.
If you wish to sputter oxidizing metals like Al, Cr, Ir, etc., you
require a turbo pumped coater such as the Emitech K575X,
http://www.ebsstore.com/ecommerce/control/product/~category_id=F1/~product_id=K575X,
which is manufactured by Quorum Technologies, the same Company that
manufactures the Polaron Range of instruments.

Best regards,
Mike Nesta

Michael R. Nesta
Managing Director
Energy Beam Sciences, Inc.
Tel: 860 653-0411
Fax: 860 653-0422
MNesta-at-ebsciences.com
www.ebsciences.com
“ADDING BRILLIANCE TO YOUR VISION”


DISCLAIMER: Energy Beam Sciences, Inc. is the US distributor for Quorum
Technologies products, which include the Polaron and Emitech ranges of
SEM specimen preparation instruments.



david.wilbur-at-tufts.edu wrote:
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} I have a Polaron SC-502 sputter coater in the SEM lab. One of the
} Engineering faculty wants to coat samples with chromium, titanium and
} aluminum for an application other than SEM. Assuming I can get targets
} that fit the sputter coater, are there other issues that would prevent
} using these other metals?
}
} Thank You
}
} Dave Wilbur
}


==============================Original Headers==============================
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From: jquinn-at-www.matscieng.sunysb.edu
Date: Fri, 25 May 2007 12:43:11 -0500
Subject: [Microscopy] re: Can I use my SEM sputter coater for metals other than

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Folks

Aluminum and titanium will deposit as alumina and titania
unless you've got a turbopump or better. There is no
easy solution. Even then, it will instantly oxidize
upon exposure to air.

If the SC-502 is a magnetron sputter unit, then the chrome
may shunt the field.

regards,

Jim

} From mail-at-ns.microscopy.com Fri May 25 09:03:20 2007
} Date: Fri, 25 May 2007 08:04:51 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: david.wilbur-at-tufts.edu
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} I have a Polaron SC-502 sputter coater in the SEM lab. One of the
} Engineering faculty wants to coat samples with chromium, titanium and
} aluminum for an application other than SEM. Assuming I can get targets
} that fit the sputter coater, are there other issues that would prevent
} using these other metals?
}
} Thank You
}
} Dave Wilbur
} --
} __________________________________
} David J. Wilbur, Ph.D.
} Instrumentation Specialist
} Department of Chemistry
} Tufts University
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From: vlk7-at-alfred.edu
Date: Fri, 25 May 2007 13:37:16 -0500
Subject: [Microscopy] AskAMicroscopist: Image Pro Analysis Question - URL with an Image

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (vlk7-at-alfred.edu) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Friday, May 25, 2007 at 12:19:24
---------------------------------------------------------------------------

Email: vlk7-at-alfred.edu
Name: Victoria Knox

Organization: Alfred University

Education: Undergraduate College

Location: Alfred, NY USA

Question: This is a re-post with a URL to an example image in question:

http://i100.photobucket.com/albums/m16/ceramicchick7/362M-B.jpg

I've been working with a program called Image-Pro plus to measure the diameter of micron-sized fibers from SEM micrographs. I need some assistance in how the program is measureing these fibers because I am getting inaccurate data that is off by at least 25% each time. I have tried to get the data using different measurements but it is still off. Any help you can provide would be greatly appreciated.

---------------------------------------------------------------------------

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From: jquinn-at-www.matscieng.sunysb.edu
Date: Fri, 25 May 2007 14:00:16 -0500
Subject: [Microscopy] Re: AskAMicroscopist: Image Pro Analysis Question - URL with an Image

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Victoria

Your magnification is too low. The answer is that simple.

If the fiber size is only a few pixels wide, then how
will you get good results?

Also, I assume that you have use a spatial calibration.

Jim


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} Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (vlk7-at-alfred.edu) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Friday, May 25, 2007 at 12:19:24
} ---------------------------------------------------------------------------
}
} Email: vlk7-at-alfred.edu
} Name: Victoria Knox
}
} Organization: Alfred University
}
} Education: Undergraduate College
}
} Location: Alfred, NY USA
}
} Question: This is a re-post with a URL to an example image in question:
}
} http://i100.photobucket.com/albums/m16/ceramicchick7/362M-B.jpg
}
} I've been working with a program called Image-Pro plus to measure the diameter of
micron-sized fibers from SEM micrographs. I need some assistance in how the program
is measureing these fibers because I am getting inaccurate data that is off by
at least 25% each time. I have tried to get the data using different measurements
but it is still off. Any help you can provide would be greatly appreciated.
}
} ---------------------------------------------------------------------------
}
} ==============================Original Headers==============================
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==============================Original Headers==============================
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From: NWWhite-at-bwxt.com
Date: Fri, 25 May 2007 14:41:40 -0500
Subject: [Microscopy] Test Post Don't Read

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Sorry, but testing posting ability. Seems to have quit!

Woody


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From: beaurega-at-westol.com
Date: Fri, 25 May 2007 17:22:45 -0500
Subject: [Microscopy] AskAMicroscopist: Image Pro Analysis

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I agree with Jim after seeing the image.

A good rule of thumb in image analysis is to have about 30 features in each
field of view that you are counting. However, some of that depends on the
degree of dispersion (separation) of the features, their sizes, and in your
case the aspect ratios. If you have a very good dispersion of smaller
(rounder) features, then higher numbers of features can be counted per
field but you risk flocculation affects.

Raise the mag to start with and count multiple fields of view.

Paul

At 02:00 PM 5/25/07 -0500, you wrote:
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From: Mike.Bode-at-olympus-sis.com
Date: Fri, 25 May 2007 18:12:02 -0500
Subject: [Microscopy] AskAMicroscopist: Image Pro Analysis

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I, too, agree with Jim and Paul.

However, seeing your image, you could try an automated approach for your
measurements. You could have the software detect the fibers and measure
their areas, then do a skeletonization and calculate the length. Divide
the area by the length, and you'll have a measurement of the width. If
you do that for multiple fibers, you should get some reasonable
statistics of the fiber width and length. Unfortunately I do not know
how that works in Image-Pro.

mike

Michael Bode, Ph.D.

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-mail: Mike.Bode-at-olympus-sis.com
www.olympus-sis.com

-----Original Message-----
X-from: beaurega-at-westol.com [mailto:beaurega-at-westol.com]
Sent: Friday, May 25, 2007 16:28
To: Mike Bode

I agree with Jim after seeing the image.

A good rule of thumb in image analysis is to have about 30 features in
each field of view that you are counting. However, some of that depends
on the degree of dispersion (separation) of the features, their sizes,
and in your case the aspect ratios. If you have a very good dispersion
of smaller
(rounder) features, then higher numbers of features can be counted per
field but you risk flocculation affects.

Raise the mag to start with and count multiple fields of view.

Paul

At 02:00 PM 5/25/07 -0500, you wrote:
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} results?
}
} Also, I assume that you have use a spatial calibration.
}
} Jim
}
}
} } From mail-at-ns.microscopy.com Fri May 25 14:36:32 2007
} } Date: Fri, 25 May 2007 13:38:05 -0500
} } To: jquinn-at-www.matscieng.sunysb.edu
} } From: vlk7-at-alfred.edu
} } Reply-to: vlk7-at-alfred.edu
} } X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
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} } Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (vlk7-at-alfred.edu) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
Friday, May 25, 2007 at 12:19:24
} }
------------------------------------------------------------------------
---
} }
} } Email: vlk7-at-alfred.edu
} } Name: Victoria Knox
} }
} } Organization: Alfred University
} }
} } Education: Undergraduate College
} }
} } Location: Alfred, NY USA
} }
} } Question: This is a re-post with a URL to an example image in
question:
} }
} } http://i100.photobucket.com/albums/m16/ceramicchick7/362M-B.jpg
} }
} } I've been working with a program called Image-Pro plus to measure
} } the
diameter of
} micron-sized fibers from SEM micrographs. I need some assistance in how
the program
} is measureing these fibers because I am getting inaccurate data that is
off by
} at least 25% each time. I have tried to get the data using different
measurements
} but it is still off. Any help you can provide would be greatly
appreciated.
} }
} }
------------------------------------------------------------------------
---
} }
} } ==============================Original
Headers==============================
} } 9, 11 -- From zaluzec-at-ultra5.microscopy.com Fri May 25 13:37:16 2007

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20, 25 -- From Mike.Bode-at-olympus-sis.com Fri May 25 18:12:02 2007
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From: nizets2-at-yahoo.com
Date: Sat, 26 May 2007 01:47:11 -0500
Subject: [Microscopy] Re: Phenom-Ed SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Phil,

I am not sure to understand what you mean.
The electron beam IS orthogonal to the object
visualized.
For the "edge smearing" I can understand the problem
though.

Stephane


--- Philip Oshel {oshel1pe-at-cmich.edu} wrote:

} Stephane,
}
} No, you don't know the size directly. Not unless the
} measured object
} is exactly orthogonal to the projected sight line.
} Quick experiment: project a bright light on the
} wall, and hold up a
} pencil (or similar). Hold the pencil so its long
} axis is exactly at
} 90 degrees to the central axis of the light path.
} Pretend the
} enlarged size of the pencil shadow is magnification.
} Now, tip the pencil into different orientations.
} Notice how the size
} of the shadow changes? Projective geometry.
} Same thing with an SEM image. It's a 3D object whose
} image is
} projected onto a 2D plane. Unless the dimension
} desired is exactly
} orthongonal to the line of sight, then the projected
} size will be
} different than the true size by an unknown amount.
} This can be solved by tilting the image, taking
} stereopairs, and
} doing trigonometry.
} But it will still be wrong unless the instrument has
} been properly
} calibrated *and* unless the all of the effects the
} specimen
} processing steps have on the sample are exactly
} known.
} Mind, this all ignores the more subtle question of
} "where's the
} edge"? Given the signal that forms an SEM image come
} from an
} interaction volume, not a dimensionless point, then
} the edge of a
} feature is essentially smeared out over the size of
} that volume. This
} is not much of an issue, or even no issue, for most
} imaging, but at
} high magnifications and resolutions, it is.
}
} Phil
}
}
} ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The
} Microscopy Society of America
} } To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} } On-Line Help
}
http://www.microscopy.com/MicroscopyListserver/FAQ.html
}
} ----------------------------------------------------------------------------
} }
} } Sure, but if you see a scale bar next to your
} } feature, you don't know it is small, you know its
} size
} } directly! (and if the full scale bar is 50 nm, you
} } know it is
} } very small)
} }
} } Stephane
} }
} }
} } } --- Mike.Bode-at-olympus-sis.com wrote:
} } }
} } } }
} } } } The advantage of using magnification is that
} it
} } } } immediately gives you an idea of size. If you
} look
} } } } at something that was acquired at 100,000x,
} you
} } } know
} } } } that it must be small.
} } } }
} } } }
} } } } Michael Bode, Ph.D.
} } } }
} } } } General Manager
} } } }
} } } } OLYMPUS SOFT IMAGING SOLUTIONS
} } } } 12596 West Bayaud Ave #300
} } } } Lakewood, CO 80228
} } } } USA
} } } } Tel.: +1 (303) 234-9270
} } } } Fax.: +1 (303) 234-9271
} } } } E-mail: Mike.Bode-at-olympus-sis.com
} } } } www.olympus-sis.com
} --
} Philip Oshel
} Microscopy Facility Supervisor
} Biology Department
} 024C Brooks Hall
} Central Michigan University
} Mt. Pleasant, MI 48859
}




____________________________________________________________________________________Need a vacation? Get great deals
to amazing places on Yahoo! Travel.
http://travel.yahoo.com/

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9, 21 -- From: Stephane Nizet {nizets2-at-yahoo.com}
9, 21 -- Subject: Re: [Microscopy] Phenom-Ed SEM
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From: zaluzec-at-microscopy.com
Date: Sat, 26 May 2007 09:28:33 -0500
Subject: [Microscopy] Administrivia: Subject Line / Topic Drift

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Colleagues....

Remember when the subject has drifting off the original topic, as it has in the
discussion on of the FEI table top SEM, into a new area it would
be advisable to modify the subject line so that it reflects the
discussion. It will help others (as well as the search engine)
in the long run.

Nestor
Your FriendlyNeighborhood SysOp



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From: kraftpiano-at-gmail.com
Date: Sat, 26 May 2007 14:28:07 -0500
Subject: [Microscopy] SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
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Just out of curiosity, what should the vacuum level be to get a steady
electron beam in an older SEM? I grabbed a vacuum gauge off of my
sputter coater and measured the vacuum at 1.2 x 10e-5. I don't think
that's enough, though. Any thoughts?

--Justin.

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2, 26 -- Date: Sat, 26 May 2007 15:28:06 -0400
2, 26 -- From: "Justin Kraft" {kraftpiano-at-gmail.com}
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2, 26 -- Subject: SEM: Vacuum level.
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From: michael-at-shaffer.net
Date: Sat, 26 May 2007 16:04:52 -0500
Subject: [Microscopy] RE: SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Justin writes ...

} Just out of curiosity, what should the vacuum level be to get
} a steady electron beam in an older SEM? I grabbed a vacuum
} gauge off of my sputter coater and measured the vacuum at 1.2
} x 10e-5. I don't think that's enough, though. Any thoughts?

10-5 would be good enough if we were speaking Pascals, and only if this
vacuum could be assumed to be in the region of the gun.

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
Inco Innovation Centre
Memorial University
St. John's, Newfoundland



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6, 21 -- References: {200705261928.l4QJSx6V018940-at-ns.microscopy.com}
6, 21 -- Subject: RE: [Microscopy] SEM: Vacuum level.
6, 21 -- Date: Sat, 26 May 2007 18:34:52 -0230
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From: kraftpiano-at-gmail.com
Date: Sat, 26 May 2007 16:08:04 -0500
Subject: [Microscopy] Re: SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Sorry, I forgot to include the units in my vacuum measurement! I just
committed the cardinal sin that I drill into my students- I wrote a
number without a unit of measure!

Anyway, the vacuum measurement is 1.2x10e-5 torr, and is measured in
the chamber itself. I can only assume that it is the same throughout
the gun, as the sensor is in a location directly between the two
locations, but on the vacuum system side.

--Justin.

On 5/26/07, Justin Kraft {kraftpiano-at-gmail.com} wrote:
} Just out of curiosity, what should the vacuum level be to get a steady
} electron beam in an older SEM? I grabbed a vacuum gauge off of my
} sputter coater and measured the vacuum at 1.2 x 10e-5. I don't think
} that's enough, though. Any thoughts?
}
} --Justin.
}

==============================Original Headers==============================
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4, 28 -- Date: Sat, 26 May 2007 17:08:03 -0400
4, 28 -- From: "Justin Kraft" {kraftpiano-at-gmail.com}
4, 28 -- To: microscopy-at-microscopy.com
4, 28 -- Subject: Re: SEM: Vacuum level.
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From: michael-at-shaffer.net
Date: Sun, 27 May 2007 09:04:49 -0500
Subject: [Microscopy] SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Justin writes ...

} Anyway, the vacuum measurement is 1.2x10e-5 torr, and is
} measured in the chamber itself. I can only assume that it is
} the same throughout the gun, as the sensor is in a location
} directly between the two locations, but on the vacuum system side.

I would consider this vacuum borderline, only because I've seen slightly
better across a broad range of SEMs and probes that I have some experience
with. Your vacuum might be considered average for the chamber region which
is generally a large volume and usually subject to a small amount of
outgassing. However, take the samples out and give the pumping 12 hours and
10-6 Torr should be achievable (e.g., my SEM achieves 1e-6 Torr easily.

However, your question is primarily about gun stability, and I might
imagine a small leak in the region of the gun "could be" the reason for a
relatively poor vacuum elsewhere. What exactly are your symptoms of
instability, and what are your filament lifetimes? If instabilities aren't
short term, abrupt and acute, and if your filament lifetimes are normal, I
might believe the small leak isn't near the gun and that your problem can be
lived with while you inspect, clean or replace o-rings in one region at a
time.

Genuinely, Michael Shaffer :o)

SEM/MLA Research Coordinator
http://www.mun.ca/creait/maf/

Inco Innovation Centre
Memorial University
St. John's, Newfoundland



} On 5/26/07, Justin Kraft {kraftpiano-at-gmail.com} wrote:
} } Just out of curiosity, what should the vacuum level be to
} get a steady
} } electron beam in an older SEM? I grabbed a vacuum gauge off of my
} } sputter coater and measured the vacuum at 1.2 x 10e-5. I
} don't think
} } that's enough, though. Any thoughts?
} }
} } --Justin.


==============================Original Headers==============================
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11, 21 -- From: "michael shaffer" {michael-at-shaffer.net}
11, 21 -- To: {kraftpiano-at-gmail.com}
11, 21 -- Cc: "MSA Microscopy list" {Microscopy-at-microscopy.com}
11, 21 -- References: {200705262108.l4QL8OW2003987-at-ns.microscopy.com}
11, 21 -- Subject: RE: [Microscopy] Re: SEM: Vacuum level.
11, 21 -- Date: Sun, 27 May 2007 11:34:07 -0230
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From: michael-at-shaffer.net
Date: Sun, 27 May 2007 12:52:03 -0500
Subject: [Microscopy] Yet another Photoshop pitfall (YAPP)

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I believe that most of us learned some time ago to disable color management
in Photoshop for avoiding side-effects due to color space conversions.
Indeed, most of us do not need CM enabled, until it's time to actually use
it (e.g., accurate prints).

For those of you, like me, who end up with instrument image TIFs that are
"indexed", you may have noticed rather extreme histogram shifts when you
converted to "grayscale". This problem is eliminated by disabling CM;
however, I've just noticed another problem even when CM is disabled. There
is another option available under color settings only if you enable "more
options". It is under "conversion options" and called "use dither
(8bit/channel)", and used for randomizing 8bit quantization effects (usually
a good thing). I never suspected this came into effect for
indexed-to-grayscale conversions, but I can see significant changes to the
histogram while the changes to the image are not significant at all.

This is not the end of it; i.e., I can still see small changes at the low
end of the histogram even while CM and dithering are disabled. Has anyone
found a remedy for this? That is, a simple way of stripping the LUT from an
indexed TIF?

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
Inco Innovation Centre
Memorial University
St. John's, Newfoundland



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From: dljones-at-bestweb.net
Date: Sun, 27 May 2007 14:12:19 -0500
Subject: [Microscopy] SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
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Justin,

While this is not a particularly "good" vacuum for your SEM as others have
pointed out and you should likely look for leaks in order for that to
improve, it should be fine for running your instrument.

I think, but I'll have to check, that you should be able to run your
electron beam and image with an order of magnitude less vacuum (10-4 torr
range).

Certainly, your instrument should be achieving better vacuum levels and
you should look for small leaks. There could be out-gassing going on,
there could be small leaks or any combination. Are you able to leave the
instrument running under vacuum for a couple days and take the measurement
at that point?

But if you are getting into the low 10-5 torr range, you should have no
problem with beam stability or imaging. If you are, the cause is not your
vacuum level...

dj

On Sat, 26 May 2007, kraftpiano-at-gmail.com wrote:

} Date: Sat, 26 May 2007 16:10:48 -0500
} From: kraftpiano-at-gmail.com
} To: dljones-at-bestweb.net
} Subject: [Microscopy] Re: SEM: Vacuum level.
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} Sorry, I forgot to include the units in my vacuum measurement! I just
} committed the cardinal sin that I drill into my students- I wrote a
} number without a unit of measure!
}
} Anyway, the vacuum measurement is 1.2x10e-5 torr, and is measured in
} the chamber itself. I can only assume that it is the same throughout
} the gun, as the sensor is in a location directly between the two
} locations, but on the vacuum system side.
}
} --Justin.
}
} On 5/26/07, Justin Kraft {kraftpiano-at-gmail.com} wrote:
} } Just out of curiosity, what should the vacuum level be to get a steady
} } electron beam in an older SEM? I grabbed a vacuum gauge off of my
} } sputter coater and measured the vacuum at 1.2 x 10e-5. I don't think
} } that's enough, though. Any thoughts?
} }
} } --Justin.
} }
}
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} 4, 28 -- From: "Justin Kraft" {kraftpiano-at-gmail.com}
} 4, 28 -- To: microscopy-at-microscopy.com
} 4, 28 -- Subject: Re: SEM: Vacuum level.
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From: r.sims-at-auckland.ac.nz
Date: Sun, 27 May 2007 15:54:59 -0500
Subject: [Microscopy] Re: SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


Hi Justin

That vacuum should be fine for a stable beam, provided, as Mike has said, that
the vacuum in the gun is good. It would be worthwhile, IMHO, to make up a
gauge-mounting flange right by the gun, there is a suitable place immediately
above V3. A gauge near the gun is really useful for troubleshooting.

Do you still have any wds spectros mounted? Those things can leak a bit.

cheers

rtch

}
} Just out of curiosity, what should the vacuum level be to get a steady
} electron beam in an older SEM? I grabbed a vacuum gauge off of my
} sputter coater and measured the vacuum at 1.2 x 10e-5. I don't think
} that's enough, though. Any thoughts?
}
} --Justin.
}

--
Ritchie Sims Ph D Phone : 64 9 3737599 ext
87713
Microanalyst Fax : 64 9 3737435
Department of Geology email : r.sims-at-auckland.ac.nz
The University of Auckland
Private Bag 92019
Auckland
New Zealand


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From: gary-at-gaugler.com
Date: Sun, 27 May 2007 20:29:43 -0500
Subject: [Microscopy] Re: Yet another Photoshop pitfall (YAPP)

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

I don't see the big deal about this. Zeiss TIFF images
are indexed color. but the header contains specific info
so the image files can be used with the separate TIFF editor.

I programmed F2 as a Photoshop action to convert the Zeiss
indexed color files (8-bit and 16-bit) to 8-bit b/w and
Save. It is really important to do this conversion since
the indexed color images are quite worse than pure b/w.

No problem.

gary g.


At 09:53 AM 5/27/2007, you wrote:




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From: dianavd-at-eye.usyd.edu.au
Date: Sun, 27 May 2007 22:03:45 -0500
Subject: [Microscopy] lead citrate and block staining - more info

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Geoff McA asked for the reference to the paper I mentioned. It's
Peters S et al; American Journal of Ophthalmology (2007)
143:995-1002; Ultrastructural findings in the primate eye after
intravitreal injection of Bevacizumab. All it says is

"postfixed with 1% OsO4 at room temperature
in 0.1 M cacodylate buffer (pH 7.4) for three hours,
bloc-stained with uranyl acetate and lead citrate, and embedded
in Epon after dehydration in a graded series of acetones"

I've Emailed asking for more details. I have the PDF if anyone would
like to see the pics.


Diana

} ----------------------------------------------------------------------
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} I've just read a paper where the author talks about block staining
} with uranyl acetate and lead citrate. UAc yes, but Pbcitrate? Has
} anyone heard of this? Does it work? The pictures in the paper were
} very nice; good contrast and detail. With the recent talk about
} general lack of contrast in specimens these days (I quite agree on
} that; I find contrast poorer now than in the past), could this be
} a new method?
}
} All opinions please!
}
} Cheers,
}
} Diana
}

==============================Original Headers==============================
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From: keith.morris-at-ucl.ac.uk
Date: Mon, 28 May 2007 05:30:25 -0500
Subject: [Microscopy] AskAMicroscopist: Image Pro Analysis

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Victoria,

The advice given by others is sound on measuring length and diameter (e.g.
by area/length). Your fibres look very uniform in diameter though, so a lot
of manual 'measure distance' clicks on a highly magnified live SEM image may
be fine, and quicker, if your SEM PC can do it - save to Excel.

You can try binary 'erode' options on a thresholded fibre as well as
skeletize (which can leave branches, but your fibres look ideal in this
case). At Harwell Biomedical (now defunct) we used dedicated 'fibre' image
analysis 'plug-ins' to measure length and diameter to help through-put (used
with equally defunct software from Magiscan). We did a lot of work for the
MMVF fibre manufacturers back then, and had sophisticated fibre activation
techniques for in-vivo radio-tracing. I don't have access to my Image Pro
(Bio-Rad LaserPix) key any more but look out for things like 'chord' and
'arc' length if offered for the skeletized line, and just make sure your
length isn't the shortest distance [feret] between the fibre beginning and
end co-ordinates. Do chat to your local Image Pro rep who will have almost
certainly been asked this question before - www.mediacy.com. Not all
programs are curvy fibre friendly, so you can sometimes use perimeter to
roughly estimate length on virgin 'rectangular' curved fibres (divided by 2
and corrected for two diameters at the ends), unless the fibre surface has
eroded.

Once you get a binary squiggly 'line' object on the overlay (say after
skeletize), the fibre length measurement should be straightforward with any
decent image analysis package. If you can't threshold the fibres correctly
from the support background, try editing the thresholded binary image
(exported to Photoshop if necessary as most modern image analysis packages
seem the lack the obviously useful full binary editor - MetaMorph has a
rather poor stab at it and promptly forgets your object editing for 'field'
measurements). Editing images via a cheap Wacom Graphire tablet and stylus
can be easier, but ImagePro did have the 'magnify' option to zoom into the
image when using a mouse (or stylus). I notice that your fibres threshold
well, but background 'particles' needed removing (probably simple with an
object classifier removing say all objects below 5um in length or are more
circular in shape (circularity)). Plus you will have to edit for length and
possibly diameter measurements with overlapping fibres.

By the way, you must use the correct W.H.O. counting rules for counting
fibres as they aren't as easy to score as rounder particles, e.g.

http://www.mtas.es/insht/en/MTA/MA_051_A04_en.htm#7321

which is biased towards optical microscopy, but it's easily applied to SEM.
Here you do need the low magnification of your original SEM image you showed
us. Essentially you select random fields, and a countable fibre is a
particle longer than 5ìm, with width less than 3ìm, and with a
length:diameter ratio greater than 3:1 (for Health & Safety atmospheric
monitoring - you probably know exactly which are your fibres). A countable
fibre with both ends in the graticule area is counted as one fibre, a
countable fibre with only one end in the area counts as half. A fibre
completely crossing the graticule, with no ends in, is not counted. (An
easy, practical method is to count the number of fibre ends in the graticule
area and divide by two, having regard to the proper treatment of split
fibres and clumps).... and so on. Fibre counting is always far quicker by
eye, viewing captured images onscreen or down the microscope, and can be
done live by SEM.

Just zoom in on random fibres or all those in the field counted to get your
diameters (from 30+ fibres - we always did hundreds as our fibres were often
recovered from lungs in in-vivo durability studies and fibre diameter could
be highly variable). This generally done quickest while on the SEM, and we
used a CamScan SEM with a full Quantimet image analysis software package
on-line (it saves capturing loads of images, other than a few 'typical
views') - and in those days our hard drives were in Mb. Naturally make sure
all fields are chosen randomly to prevent bias.

I'll send another pdf that adds in a lot of useful fibre counting info (but
check out the latest WHO (world health organisation) counting rules as I
left the fibre toxicology field behind 8 years ago.

Regards

Keith

-----------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology
Institute of Ophthalmology
UCL, London EC1V 9EL











-----Original Message-----
X-from: beaurega-at-westol.com [mailto:beaurega-at-westol.com]
Sent: 25 May 2007 23:27
To: keith.morris-at-ucl.ac.uk

I agree with Jim after seeing the image.

A good rule of thumb in image analysis is to have about 30 features in each
field of view that you are counting. However, some of that depends on the
degree of dispersion (separation) of the features, their sizes, and in your
case the aspect ratios. If you have a very good dispersion of smaller
(rounder) features, then higher numbers of features can be counted per
field but you risk flocculation affects.

Raise the mag to start with and count multiple fields of view.

Paul

At 02:00 PM 5/25/07 -0500, you wrote:
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From: kraftpiano-at-gmail.com
Date: Mon, 28 May 2007 08:07:17 -0500
Subject: [Microscopy] SEM: Vacuum level.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Again

I am sorry to pursue this question one more time, but WHY are you looking at
the vacuum?

If the high voltage will switch on there is absolutely NO reason to look at
the vacuum at that stage.
Once the high voltage is operable if you then have discharge problems at
high kV then you MAY have a leak. Lowering the kV the problem will be
reduced but of course the strain on the high voltage system is also reduced
so we do not necessarily prove it is a vacuum problem. It is almost
impossible in the lab to check a high voltage cable as it may only fail at
very high voltages, this is only a task for the experienced high voltage
technician! The next phase sould be to look at filament life because it may
surprise people to know that the instruments will run with a poor vacuum.
Levels inferior to five times ten minus four torr have been quite normal in
SEM and TEM in the past without high voltage discharge problems.

With over 40 years experience with the problems that beset TEM, SEM and EDS
one of the major problems we all suffer from is pre judging a fault. When a
fault occurs the question should be "what did you do last?". I have
discussed this problem with Justin over a few weeks and I fail to see that
the vacuum is at fault. The problem seems to me to be a high voltage cable
breaking down.

There are ways of checking this out and finding a reduced cost fix rather
than going to the original equipment manufacturer!

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


----- Original Message -----
X-from: {kraftpiano-at-gmail.com}
To: {protrain-at-emcourses.com}
Sent: Saturday, May 26, 2007 8:29 PM

Actually, the reason I was thinking it was the vacuum is exactly the
"What happened last" question- there was a major leak, and in the
process of repairing it, I think I may have lost some of the diffusion
pump oil because it wasn't fully cooled down when I opened up the
valve head above it to replace the O-ring. Then, the O-ring that I
got was slightly larger than the O-ring that I removed. Other than
that, nothing has changed a bit. Everything else was working
perfectly until the aforementioned vacuum leak. I also discovered (In
the process of troubleshooting things yesterday) that the vacuum gauge
on the SEM is wildly off base. Late last night I was able to get an
absolute vacuum reading from a gauge (Rather than convert the 0-200
scale on the built-in gauge) and the vacuum is actually only at about
120 millitorr, not the 1.2x10e-5 I stated earlier (That number was
gained by a false conversion factor between the built-in gauge and
reality).

I'm still not seeing, however, how removing the high voltage cable
shows that it is the cable breaking down. The HV switches on, and no
emission is measured. When the cable is plugged in, the HV switches
on, and emission is measured at 20-30 (Out of 200, using the same
arbitrary meter that the vacuum uses) then slowly dies down again. As
to it not switching on if the vacuum level is low, I discovered that
the high voltage will switch on as long as the vacuum meter registers
high enough, which doesn't necessarily have anything to do with the
actual vacuum, as I found out at about 1:00 AM this morning.

It seems that since the emission current is measured by how much
current is actually flowing, removing the HV cable just proves that no
current is flowing at all without it, and with it some flows, but then
something breaks down. I'm not sure how that limits it to the HV
cable.

I'm sorry if it seems like I'm just giving out information that should
have been given out a while ago, (Or cancels out previous information)
but I just got the vacuum gauge setup properly late last night.

--Justin.

On 5/28/07, protrain-at-emcourses.com {protrain-at-emcourses.com} wrote:
}
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} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} Hi Again
}
} I am sorry to pursue this question one more time, but WHY are you looking at
} the vacuum?
}
} If the high voltage will switch on there is absolutely NO reason to look at
} the vacuum at that stage.
} Once the high voltage is operable if you then have discharge problems at
} high kV then you MAY have a leak. Lowering the kV the problem will be
} reduced but of course the strain on the high voltage system is also reduced
} so we do not necessarily prove it is a vacuum problem. It is almost
} impossible in the lab to check a high voltage cable as it may only fail at
} very high voltages, this is only a task for the experienced high voltage
} technician! The next phase sould be to look at filament life because it may
} surprise people to know that the instruments will run with a poor vacuum.
} Levels inferior to five times ten minus four torr have been quite normal in
} SEM and TEM in the past without high voltage discharge problems.
}
} With over 40 years experience with the problems that beset TEM, SEM and EDS
} one of the major problems we all suffer from is pre judging a fault. When a
} fault occurs the question should be "what did you do last?". I have
} discussed this problem with Justin over a few weeks and I fail to see that
} the vacuum is at fault. The problem seems to me to be a high voltage cable
} breaking down.
}
} There are ways of checking this out and finding a reduced cost fix rather
} than going to the original equipment manufacturer!
}
} Steve Chapman
} Senior Consultant Protrain
} For electron microscopy consultancy and training world wide
} Tel +44 1280 816512 Fax +44 1280 814007
} Mobile +44 7711 606967 Web www.emcourses.com
}
}
} ----- Original Message -----
} X-from: {kraftpiano-at-gmail.com}
} To: {protrain-at-emcourses.com}
} Sent: Saturday, May 26, 2007 8:29 PM
} Subject: [Microscopy] SEM: Vacuum level.
}
}
} }
} }
} }
} } ----------------------------------------------------------------------------
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} }
} } Just out of curiosity, what should the vacuum level be to get a steady
} } electron beam in an older SEM? I grabbed a vacuum gauge off of my
} } sputter coater and measured the vacuum at 1.2 x 10e-5. I don't think
} } that's enough, though. Any thoughts?
} }
} } --Justin.
} }
} } ==============================Original
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} } 2, 26 -- From: "Justin Kraft" {kraftpiano-at-gmail.com}
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} 11, 27 -- From protrain-at-emcourses.com Mon May 28 05:41:20 2007
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From: mjacobs04-at-sbcglobal.net
Date: Mon, 28 May 2007 08:28:55 -0500
Subject: [Microscopy] Unsubscribe

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Thanks
jakeers


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From: mike.reedy-at-cellbio.duke.edu
Date: Mon, 28 May 2007 10:27:42 -0500
Subject: [Microscopy] Re: lead citrate and block staining - more info

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

The crunch here is not what is the vacuum BUT, can you get a beam?

I have run Electron Guns under a good rotary pump pressure, if the vacuum is
not good enough the high voltage will short out to the walls of the system
and you will hear high voltage discharge in the gun. Forget the vacuum and
try to get a beam. When you draw a current, if the cable is at fault, the
system will breakdown, currents will fall to zero! Only by trying to obtain
a beam will you find out what is going on in your gun. You say that with
the cable removed you do not obtain a reading, good no breakdown. You then
say with the cable fitted you obtain a reading which falls to zero, is this
the natural surge or a sign that the cable has broken down? Now if you try
to heat the filament can you obtain an emission reading that will hold and
then find the beam?

Just forget about the vacuum and run the gun. If the gun will not work you
have proven that the high voltage system is the problem and a check on the
cable in my mind will prove that the cable is at fault.

Steve Chapman
Senior Consultant Protrain
For electron microscopy consultancy and training world wide
Tel +44 1280 816512 Fax +44 1280 814007
Mobile +44 7711 606967 Web www.emcourses.com


----- Original Message -----
X-from: "Justin Kraft" {kraftpiano-at-gmail.com}
To: {protrain-at-emcourses.com} ; {microscopy-at-microscopy.com}
Sent: Monday, May 28, 2007 2:07 PM

Diana,

David Elliott kindly emailed me his Microscopy Today article and
protocol at my request. After aqueous Glut-OsO4 fix and UrAc bloc
stain,and EtOH dehydration, he uses the lead as Pb acetate,
saturated, in 50:50 EtOH:Acetone, then shifts into acetone to perform
epoxy resin infiltration.

In the ref you cited, Dr Peters uses Pb Citrate, apparently aqueous,
in a UrAc --} PbCit sequence before acetone dehydration.

So there seems a little leeway as to which lead salt and what stage
to apply it, but only acetone reported as the transition solvent to
epoxy resin.

The staining looks fine in both papers. I'll be trying it to compare
with the contrast levels we get using Tannic Acid in the primary fix,
and KMnO4-SatoPb as section stain sequence.

(I was clearly mistaken in my off-list warnings to you about curing
and cutting problems I thought likely with the double bloc stain.)

-mike reedy-

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Geoff McA asked for the reference to the paper I mentioned. It's
Peters S et al; American Journal of Ophthalmology (2007)
143:995-1002; Ultrastructural findings in the primate eye after
intravitreal injection of Bevacizumab. All it says is

"postfixed with 1% OsO4 at room temperature
in 0.1 M cacodylate buffer (pH 7.4) for three hours,
bloc-stained with uranyl acetate and lead citrate, and embedded
in Epon after dehydration in a graded series of acetones"

I've Emailed asking for more details. I have the PDF if anyone would
like to see the pics.


Diana

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} I've just read a paper where the author talks about block staining
} with uranyl acetate and lead citrate. UAc yes, but Pbcitrate? Has
} anyone heard of this? Does it work? The pictures in the paper were
} very nice; good contrast and detail. With the recent talk about
} general lack of contrast in specimens these days (I quite agree on
} that; I find contrast poorer now than in the past), could this be
} a new method?
}
} All opinions please!
}
} Cheers,
}
} Diana
}

==============================Original Headers==============================
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6, 18 -- Content-Transfer-Encoding: 7bit
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6, 18 -- Subject: lead citrate and block staining - more info
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--
-mike reedy-

************************
Michael K. Reedy, M.D.
Duke Univ. Med. Center
Dept. Cell Biology, Box 3011 (for U.S. Mail)
458 Alex Sands Bldg, Research Dr (courier)
Durham, NC 27710

Office 919-668-2534
Lab 919-684-5674
Fax 919-681-9929
mike.reedy-at-cellbio.duke.edu
http://note.cellbio.duke.edu/Faculty/Research/Reedy.html

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From: kehler-at-twu.ca
Date: Mon, 28 May 2007 17:11:21 -0500
Subject: [Microscopy] TEM available

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

We have a Philips EM300, TEM with various user, service & installation
manuals plus some accessories, including single-tilt goniometer
available.

This instrument was initially installed in the University of Victoria
(Biology), BC Canada in the early 1970's.

It was moved to Simon Fraser University (Physics), sometime in the
1990's, and was acquired by Trinity Western (Biology) in 2000.

This instrument was demonstrated to be functional in its TWU location.
Actual use has been extremely light.

We are interested in offering the instrument "free - as is where is".
If assistance in disassembly/packing/shipping is required, those costs
will be negotiated.

Please direct your inquiries to me, as convenient.

Darcy

Mr. Darcy Kehler, B.Sc.
Sr. Lab Supervisor, Biology
Faculty of Natural and Applied Sciences
Trinity Western University
p: 604.888.7511 ext. 3249
F: 604.513.2018
Langley, BC,
Canada, V2Y 1Y1


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From: oshel1pe-at-cmich.edu
Date: Tue, 29 May 2007 07:32:22 -0500
Subject: [Microscopy] Re: SEM beam-specimen relations (subject line change)

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Stephan,

The beam is orthogonal only if the specimen is e.g. perfectly flat on
the specimen stub and the stub is exactly at zero degrees tilt. Like
looking down on a pencil on your desktop. Push down on the point of
the pencil and the opposite end pops up. Still looking *straight
down* on the pencil, the image of the pencil will now be shorter. Try
the shadow trick to see what I mean. The image on the view CRT, or as
collected by the imaging system, is the 2D projection of the 3D image.
And, the beam is only orthogonal at the exact center of the scan
raster. Anywhere else, and it is approaching the specimen at some
angle.

_/___ versus __|__

This is only important on perfectly flat specimens, though. On most
normal specimens, and anything biological, the surface topography
means the beam is very rarely perfectly orthogonal to the specimen
surface. And, it may not affect the signal enough to matter. So we
ignore the effect. For high-precision metrology or imaging of things
like single molecules, I suspect this effect could become important.

Phil

} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

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From: jae5-at-lehigh.edu
Date: Tue, 29 May 2007 08:19:35 -0500
Subject: [Microscopy] Changed submission date CIASEM 2007 Cusco

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Change of dates: JUNE 30 ABSTRACT DEADLINE

The submission date for abstracts (in a two-page format, very
similar to the M and M format) has been changed to June 30, 2007.
There is still a month to get your paper in to this exciting meeting.

9th InterAmerican Congress on Electron Microscopy

Cusco Peru

September 23 - 28, 2007

One of the major electron microscopy meetings this year will
be held in the heart of the Inca empire. Machu Picchu, one
of the world's most famous and wondrous places is nearby.

Leading microscopists from across the Americas (and from
the rest of the world) will be presenting their latest work.

Details of how to register (which must be done before the
abstract is submitted) and of how to submit the abstract,
can be found at the web site:
http://www.ciasem2007.com/
(To change to English click the little blue button to the right.)


--
...........
Alwyn Eades
Department of Materials Science and Engineering
Lehigh University
5 East Packer Avenue
Bethlehem
Pennsylvania 18015-3195
Phone 610 758 4231
Fax 610 758 4244
jae5-at-lehigh.edu


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From: chunmin.wang-at-ingeniapolymers.com
Date: Tue, 29 May 2007 20:38:41 -0500
Subject: [Microscopy] viaWWW: Leitz weitzlar heating stage 1350

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Email: chunmin.wang-at-ingeniapolymers.com
Name: Chunmin Wang

Organization: Ingenia Polymer Corp

Title-Subject: [Filtered] Leitz weitzlar heating stage 1350

Question: Dear All,

I just got one leitz 1350 heating stage from ebay and I want to connect it to our lab microscope. However, I cannot find the right temperature controller for the stage.
Can anyone who has the same stage kindly tell me the model of the controller?
Thank you,

Chunmin Wang

Research Engineer
Ingenia Polymers Corp.
565 Greenwich Street
Brantford, Ontario N3T 5M8 CANADA
Tel: 519-758-8941 ext: 1020
Fax: 519-758-1254


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From: bressan-at-smt.zeiss.com
Date: Tue, 29 May 2007 20:39:23 -0500
Subject: [Microscopy] viaWWW: 2 OPEN POSITIONS- FIB Applications and La Manager

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Email: bressan-at-smt.zeiss.com
Name: Beth Bressan

Organization: Carl Zeiss SMT Inc.

Title-Subject: [Filtered] 2 OPEN POSITIONS- FIB Applications and La Manager

Question:
Due to growth, Carl Zeiss SMT Inc. is seeking two experienced FIB Product Managers, to be located at our new headquarters in Peabody, MA.

Responsibilities include overall FIB product management-related sales support activities, specifically instrument demonstrations, sample processing, prospect presentations, preparation and development of promotional items and sales tools, and customer training. Responsibilities also include overall FIB applications expertise in order to assist the Companyís sales efforts in achieving our instrument orders and sales targets, to equip the Companyís prospects and customers with in-depth product knowledge and capabilities enabling them to achieve high levels of operator satisfaction, to ensure the optimum product performance of our FIB instruments, and to assist the Companyís technical service efforts by disseminating appropriate product information and providing technical support.
An advanced degree in a related field and extensive prior experience with FIBs in a corporate setting is required. One position requires frequent travel (both domestic and international to our facilities in Germany), a customer-focused mentality, and excellent communication skills. The Lab Manager position requires successful prior experience managing a lab. Compensation is commensurate with experience and Carl Zeiss SMT Inc. provides a comprehensive benefits package. An Affirmative Action Employer M/F/D/V
Contact Beth Bressan, bressan-at-smt.zeiss.com All replies are confidential.





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From: nizets2-at-yahoo.com
Date: Wed, 30 May 2007 03:54:05 -0500
Subject: [Microscopy] Nitrocellulose membranes and SEM

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dears friends,

I blotted a suspension of particles in a salt solution
on a nitrocellulose membrane (those used for protein
blotting, 0,2 µm pore size) and washed it extensively
under vacuum to remove the salts. Then I dried the
membrane, glued it on a alu stub and carbon coated it
(5 sec at 4 V).
Now when I observe in SEM at 12 kV the image moves a
lot, especially the luminosity. The region around the
scanned area (I see it when I zoom out) is saturated
(white). I suspect it is charged.
In EDX the sensitivity is very low, much lower than
expected. This is specific for this sample, other
samples work good in EDX.
I tried to observe in low vacuum mode (10 Pa) but at
12 kV I can see nothing. At 20 kV I get an image
although the luminosity still is very unstable and the
quality is far worse than in high vacuum mode. However
it is possible to get an EDX spectrum in these
conditions.

I was very surprised the way this membrane works in
SEM and particularly in EDX. I already analyzed the
same suspension directly deposited on a carbon tabs
and I had good conditions to work.

1) Should I expected heavy loading in such a membrane,
even carbon coated?
2) Does (charge) loading of the sample hamper the EDX
analysis?
3) What would be a solution? coat more with carbon? I
would like to avoid gold since it has a band that
superimposes on a band of interest in EDX and could
absorb some signal.

Stephane




____________________________________________________________________________________Ready for the edge of your seat?
Check out tonight's top picks on Yahoo! TV.
http://tv.yahoo.com/

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From: dls_douglasl-at-yahoo.com.br
Date: Wed, 30 May 2007 08:22:21 -0500
Subject: [Microscopy] AskAMicroscopist: surface densities of particles

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dls_douglasl-at-yahoo.com.br) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Tuesday, May 29, 2007 at 00:41:27
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Email: dls_douglasl-at-yahoo.com.br
Name: Douglas da Silva

Organization: Universidade federal do Rio Grande do Sul

Education: Graduate College

Location: Porto Alegre, Rio Grande do Sul, Brazil

Question: Why is not possible to obtain surface densities of particles from cross sectional samples (XTEM)?
There is any reference explaining this?

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From: mukesh_ag7-at-rediffmail.com
Date: Wed, 30 May 2007 08:23:12 -0500
Subject: [Microscopy] AskAMicroscopist: image analysis without caoting of plant samples

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mukesh_ag7-at-rediffmail.com) from http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Wednesday, May 30, 2007 at 06:32:52
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Email: mukesh_ag7-at-rediffmail.com
Name: Mukesh Kumar

Organization: University of Agricultural Science

Education: Graduate College

Location: Bangalore, Karnataka, India

Question: Hi,

I want to know whether using SEM with BSE,LVSTD can we do image analysis without caoting of plant samples to view cellullar structural & details like Virus,bacteria etc., at very low resolution

Also is that only that you can use ESEM system for this analysis OR any other system is available in world

Please clarify me this doubt and also sent some image of virus on plant samples if available with protocol for analysis

Thanks
Mukesh Kumar

---------------------------------------------------------------------------

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From: cyrus.shaoul-at-ualberta.ca
Date: Wed, 30 May 2007 16:59:20 -0500
Subject: [Microscopy] Seeking a functional, obsoleted SEM system for educational use.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Microscopy List Members:

I am a graduate student at the University of Alberta, and I am hoping to
pass my interest in science on to my children. My son Darius is very
much interested in "the small", and we talked about setting up an
electron microscope in our garage to do experiments.

I have some experience with vacuum equipment, high voltage equipment and
running scientific software. Setting up an SEM and getting images with
it would be fun for both of us.

If anyone knows of a functional, obsoleted SEM system that needs a new
home, please let me know. I will arrange transport for it from
wherever it is to my home, at my expense.

I will also take full responsibility for learning how to operate it, and
operating it safely.

Thanks for your time and generosity,

Cyrus Shaoul
http://www.ualberta.ca/~cshaoul/

--
=[=]={=}=[=]={=}=[=]={=}=[=]={=}=[=]={=}
Cyrus Shaoul
http://www.psych.ualberta.ca/~westburylab/
University of Alberta
780-492-5843
=[=]={=}=[=]={=}=[=]={=}=[=]={=}=[=]={=}




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From: john.brealey-at-imvs.sa.gov.au
Date: Wed, 30 May 2007 21:37:45 -0500
Subject: [Microscopy] Recommendations for a book on Ultrastructural Pathology

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi,

Are there any recent publications on this subject?
My most recent book is Erlandson's "Diagnostic Transmission Electron
Microscopy of Tumors" from 1994.
While this is an excellent reference, I was wondering if there is anything
similar published recently.

Thanks,

John Brealey
EM Unit
Queen Elizabeth Hospital
Adelaide, South Australia


==============================Original Headers==============================
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From: hyi-at-emory.edu
Date: Wed, 30 May 2007 23:39:13 -0500
Subject: [Microscopy] Workshop announcement

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Researchers:

There are still a couple of spots available in our upcoming
hands-on Cryo-technique and Immunogold Labeling workshop (See details
below). Please contact Hong Yi at hyi-at-emory.edu or (404) 712-8491
asap if you would like to participate. This workshop will be led by
world renowned Mr. Helmut Gnagi for cryo-ultramicrotomy and Dr. Jan
Leunissen for immunogold labeling. Dr. Leunissen will also be
lecturing on cryo-fixation and introduce a new freezing method that
produces high quality cryo-fixation.

Thank you and looking forward to seeing you in Atlanta.

================

1. Date and Curriculum

Aug. 12-13:
· Cryo-ultramicrotomy
· A new cryo-fixation method
· Set up for cryo-substitution
Aug. 14-16:
· The properties of gold particles and their protein conjugates.
· Theories underlying immunogold labeling protocols.
· Silver enhancement of gold particles
· Imunogold labeling on a variety of sample preparations for LM.
· Immunogold labeling for EM
a. Pre-embedding immunogold labeling using ultrasmall gold conjugates
and silver enhancement.
b. Post-embedding immunogold labeling using conventional colloidal
gold conjugates and ultrasmall gold conjugates.

Numerous biological microscopy techniques will be also
demonstrated during the workshop. Details TBA

2. Main Instructors and Sponsors

Dr. Jan Leunissen, Aurion
Mr. Helmut Gnägi, Diatome
Hong Yi, Emory University

Leica Microsystems
Aurion ImmunoGold Reagents
Electron Microscopy Sciences/Diatome
Hitachi High Technologies America, Inc

3. Fees

Session A: Cryo-technique: $500
Session B: Immunogold: $500
Session A and B: $800

Participants can sign up for either the entire workshop
or a particular session of the workshop. If desired, participants
who sign up for cryo-techniques will have additional practice time
after lectures and training during the first two days. Applicants
signing up for both sessions will be given first priority for
enrollment.

4. Participants

The enrollment is open to anyone with interest to learn
regardless of previous experience. However, due to limited space
availability, the number of participants will be limited to 12 for
the cryo-techniques and 20 for immunogold labeling.

5. Lodging

Participants are responsible for making hotel
reservation themselves. The workshop will block a number of rooms at
the following hotels

Villa International: (404) 633-6783, $24/night/person (double
occupancy), or $36/night/person (single occupancy)

This hotel is cozy and clean and often used by Emory to house
temporary or visiting employees. However, TV and phone are only
available in the hotel common room.

Emory Inn: (800) 933-6679, $107/night or higher


Hong Yi
Emory SOM EM

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From: richard.beanland-at-bookham.com
Date: Thu, 31 May 2007 05:16:34 -0500
Subject: [Microscopy] AskAMicroscopist: surface densities of particles

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi Douglas,
There is no a priori reason why you can't measure densities of
particles using cross section TEM. However there are two things you
need to be aware of:


1) The specimen is seen in transmission, so it is possible that you have
two (or more) particles along the 'line of sight' of the electron beam.
This can make it difficult to get an accurate count of the number of
particles.

2) The specimen has a finite thickness (which might not be immediately
apparent from a cross section image). To work out how many particles
you have per unit area of the surface you need to know the specimen
thickness.


Both of these problems can be overcome by tilting the specimen so that
the surface is not seen 'edge-on'. You can then work out the specimen
thickness from trigonometry and the particles will not overlap. However
the image may look more complicated and need more careful
interpretation. If you want you can also take a stereo pair and then
you can see the true 3D structure.

Best of luck

Richard



________________________________________
Richard Beanland
Materials Analysis
Bookham
Caswell
Towcester
Northants
NN12 8EQ
United Kingdom
Tel. +44 1327 356362
Fax. +44 1327 356775
http://www.bookham.com
________________________________________

-----Original Message-----
X-from: dls_douglasl-at-yahoo.com.br [mailto:dls_douglasl-at-yahoo.com.br]
Sent: 30 May 2007 14:24
To: Richard Beanland

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (dls_douglasl-at-yahoo.com.br) from
http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
Tuesday, May 29, 2007 at 00:41:27
------------------------------------------------------------------------
---

Email: dls_douglasl-at-yahoo.com.br
Name: Douglas da Silva

Organization: Universidade federal do Rio Grande do Sul

Education: Graduate College

Location: Porto Alegre, Rio Grande do Sul, Brazil

Question: Why is not possible to obtain surface densities of particles
from cross sectional samples (XTEM)?
There is any reference explaining this?

------------------------------------------------------------------------
---

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From: dljones-at-bestweb.net
Date: Thu, 31 May 2007 07:59:26 -0500
Subject: [Microscopy] wood identification

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Good morning,

I am wondering if anyone knows a good reference for wood identifcation
using microscopy. I have sometimes run into the need to know exactly what
species of wood was used in a piece that I have to analyze. A number of
years ago, I had a part to identify and gave the wood piece to someone I
knew who was able to look at the cell structure and tell me what the wood
was. I have run across this same identification problem several times
since (I no longer have someone I can use for this) and have one such
problem right now in house. But I do not know where or how to identify the
exact wood species used in this part.

Does anyone know of a good reference(s) I could get on wood
identification?

Thank you in advance.

dj




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From: dsherman-at-purdue.edu
Date: Thu, 31 May 2007 08:12:17 -0500
Subject: [Microscopy] Cryo-SEM technique review

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi all,

There are numerous references for cryo-SEM in the literature but I am
continually asked for just one that will best explain the technique to those
not familiar with it.

Does anyone have a favorite review or general reference that would suit this
need?

Thanks,
Debby

Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.agriculture.purdue.edu/microscopy


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From: TindallR-at-missouri.edu
Date: Thu, 31 May 2007 09:11:25 -0500
Subject: [Microscopy] wood identification

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi,

First places I would check would be with a university department of
anthropology/archaeology or forestry, or with your local USDA Forest
Service. Archaeologists are always identifying wood from samples of
charcoal, for example.

Good luck,
Randy

Randy Tindall
President, Officers, and Member
Half-Norwegian (on my mother's side) Microscopy Society of America---Ve
Do Small Real Good!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu
On-line calendar:
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Week&NavType=Both&Type=TimePlan


-----Original Message-----
X-from: dljones-at-bestweb.net [mailto:dljones-at-bestweb.net]
Sent: Thursday, May 31, 2007 8:01 AM
To: Tindall, Randy D.

Good morning,

I am wondering if anyone knows a good reference for wood identifcation
using microscopy. I have sometimes run into the need to know exactly
what species of wood was used in a piece that I have to analyze. A
number of years ago, I had a part to identify and gave the wood piece to
someone I knew who was able to look at the cell structure and tell me
what the wood was. I have run across this same identification problem
several times since (I no longer have someone I can use for this) and
have one such problem right now in house. But I do not know where or how
to identify the exact wood species used in this part.

Does anyone know of a good reference(s) I could get on wood
identification?

Thank you in advance.

dj




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18, 26 -- From TindallR-at-missouri.edu Thu May 31 09:11:24 2007
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From: jquinn-at-www.matscieng.sunysb.edu
Date: Thu, 31 May 2007 09:16:19 -0500
Subject: [Microscopy] re: wood

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html


DLJ and Folks

Wood Structure and Indentification
Core, Cote, Day
Syracuse
1980


Purham and Gray
Practical Indentification of Wood Pulp Fibers
Atlanta
Tappi Press

Hoadley
Indentifying Wood
Taunton Press

regards,

JQuinn

PS: OoO away...........

} From mail-at-ns.microscopy.com Thu May 31 08:58:13 2007
} Date: Thu, 31 May 2007 08:00:13 -0500
} To: jquinn-at-www.matscieng.sunysb.edu
} From: dljones-at-bestweb.net
} Reply-to: dljones-at-bestweb.net
} X-Resent-From: "Microscopy Listserver" {microscopy-at-microscopy.com}
} Subject: [Microscopy] wood identification
} Errors-To: MicroscopyListSpamFilter-at-microscopy.com
} X-lewp: MicroscopyListSpam NAGS
}
}
}
}
} ----------------------------------------------------------------------------
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} ----------------------------------------------------------------------------
}
} Good morning,
}
} I am wondering if anyone knows a good reference for wood identifcation
} using microscopy. I have sometimes run into the need to know exactly what
} species of wood was used in a piece that I have to analyze. A number of
} years ago, I had a part to identify and gave the wood piece to someone I
} knew who was able to look at the cell structure and tell me what the wood
} was. I have run across this same identification problem several times
} since (I no longer have someone I can use for this) and have one such
} problem right now in house. But I do not know where or how to identify the
} exact wood species used in this part.
}
} Does anyone know of a good reference(s) I could get on wood
} identification?
}
} Thank you in advance.
}
} dj
}

==============================Original Headers==============================
10, 12 -- From jquinn-at-www.matscieng.sunysb.edu Thu May 31 09:16:19 2007
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10, 12 -- To: dljones-at-bestweb.net, microscopy-at-microscopy.com
10, 12 -- Subject: re: wood
==============================End of - Headers==============================




From: kwon.ohkyung-at-gmail.com
Date: Thu, 31 May 2007 09:26:21 -0500
Subject: [Microscopy] [RE] Wood identification references

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello,

There are many books for wood identification, but the following books
would be most helpful to you.

**
IAWA Committee on Nomenclature. 1964. Multilingual glossary of terms
used in wood anatomy. Konkordia, Winterthur, 186pp.

IAWA Committee. 1989. IAWA list of microscopic features for hardwood
identification. IAWA Bull., n.s. 10(3) : 219 $B!A(B 332.

IAWA Committee. 2004. IAWA list of microscopic features for softwood
identification. IAWA J. 25(1): 1 $B!A(B70.

Panshin, A. J. and C. de Zeeuw. 1980. Textbook of wood technology. 4th
ed., McGraw-Hill Book Co., 722pp.

Core, H. A., Cote, W. A., and A. C. Day. 1979. Wood structure and
identification. 2nd ed., Syracuse Univ. Press, 182pp.

Cote, W. A. 1965. Cellular ultrastructure of woody plants. Syracuse
Univ. Press, New York, 603pp.

**

Regards,

--
Ohkyung Kwon, Ph D

http://www.meso.or.kr/portfolio
http://www.wpskorea.org

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From: dac-at-research.umass.edu
Date: Thu, 31 May 2007 09:33:29 -0500
Subject: [Microscopy] Re: wood identification

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Bruce Hoadley's books from Tauton Press are superb. Always well written
and illustrated. He was (is?) a professor at University of
Massachusetts, Amherst.

http://www.amazon.com/Identifying-Wood-Accurate-Results-Simple/dp/0942391047/ref=pd_sim_b_3/105-9189572-8702064

or try:

http://www.amazon.com/What-Wood-That-Manual-Identification/dp/0670759074


Hope this helps,

Dale


dljones-at-bestweb.net wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Good morning,
}
} I am wondering if anyone knows a good reference for wood identifcation
} using microscopy. I have sometimes run into the need to know exactly what
} species of wood was used in a piece that I have to analyze. A number of
} years ago, I had a part to identify and gave the wood piece to someone I
} knew who was able to look at the cell structure and tell me what the wood
} was. I have run across this same identification problem several times
} since (I no longer have someone I can use for this) and have one such
} problem right now in house. But I do not know where or how to identify the
} exact wood species used in this part.
}
} Does anyone know of a good reference(s) I could get on wood
} identification?
}
} Thank you in advance.
}
} dj
}
}
}
}
} ==============================Original Headers==============================
} 8, 16 -- From dljones-at-bestweb.net Thu May 31 07:59:26 2007
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9, 22 -- From dac-at-research.umass.edu Thu May 31 09:33:29 2007
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From: Karen_Bentley-at-URMC.Rochester.edu
Date: Thu, 31 May 2007 09:43:03 -0500
Subject: [Microscopy] Recommendations for a book on Ultrastructural

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

John:

After spending 20 years diagnosing patient tumor biopsies I am
recommending these books:

1)Ultrastructural Pathology of the Cell and Matrix(1997) by Feroze N.
Ghadially is a must have 2 volume reference book.
2)Diagnostic Electron Microscopy, A text /atlas (2000) by G. Richard
Dickersin has excellent examples (whole page electron images of almost
every neoplasm, metabolic, renal and skeletal/nerve biopsy
interpretation).


Karen L. Bentley, M.S.
Technical Director
Electron Microscope Research Core
University of Rochester Medical Center
575 Elmwood Avenue, Box 626
Rochester, NY 14642
585-275-1954


-----Original Message-----
X-from: john.brealey-at-imvs.sa.gov.au [mailto:john.brealey-at-imvs.sa.gov.au]
Sent: Wednesday, May 30, 2007 10:46 PM
To: Bentley, Karen

Hi,

Are there any recent publications on this subject?
My most recent book is Erlandson's "Diagnostic Transmission Electron
Microscopy of Tumors" from 1994.
While this is an excellent reference, I was wondering if there is
anything similar published recently.

Thanks,

John Brealey
EM Unit
Queen Elizabeth Hospital
Adelaide, South Australia


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From: mcgeejj-at-kapl.gov
Date: Thu, 31 May 2007 10:12:00 -0500
Subject: [Microscopy] Open Position - Experienced TEM microscopist for materials science

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Lockheed Martin- KAPL, Inc. has an open position for an experienced TEM
materials scientist/engineer.
Brief description of job below.
US CITIZENS ONLY.

Interested people may contact me or go to the Lockheed Martin Careers
section to apply (job ID included below).

************************************
James J. McGee
Materials Engineer, Test Operations
Lockheed Martin, KAPL, Inc.
Mail Bin 149
PO Box 1072
Schenectady, NY 12301-1072

Tel: 518-395-4612
Fax: 518-395-4340
email: mcgeejj-at-kapl.gov
************************************



Open Position: Lockheed Martin, KAPL, Inc., Schenectady NY

Req ID 35139BR

Industry Job Title: Materials Engineer Stf

Required skills: PhD plus substantial experience in transmission electron
microscopy techniques (high resolution, EDX, Electron diffraction) applied
to engineering materials. Demonstrated materials problem solving ability.
Excellent oral and written communication skills. Desired skills Material
problem solving experience includes metals and various types of metal
corrosion. Metalographic sample preparation experience (electropolishing,
ion milling, FIB.) PEELS experience. Previous work with nuclear or
radioactive materials.
Additional background in this or other microchemical or microstructural
analysis tools.

Specific Job Description: Analytical electron microscopy(AEM): Transmission
electron microscopy analysis for materials used in nuclear reactors systems.
Skills in high resolution, use of FEG instruments, EDX techniques, and
electron diffraction interpretation required.
Work with materials personnel to understand issues, plan tests, prepare
specimens, perform necessary analyses and issue reports that draw
conclusions about the relevance of microscopy work to the problem.
Collaborate with other analysts.

Work may involve some radioactive specimens and may require radiological
training / qualification to perform this work.

Applicants selected will be subject to a Federal background investigation
and must meet eligibility requirements for access to classified matter. U.S.
citizenship is required.


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14, 28 -- From: "J.J. McGee" {mcgeejj-at-kapl.gov}
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14, 28 -- Subject: Open Position - Experienced TEM microscopist for materials science
14, 28 -- applications
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14, 28 -- X-imss-settings: Baseline:2 C:2 M:3 S:4 R:4 (0.1500 0.1500)
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From: dac-at-research.umass.edu
Date: Thu, 31 May 2007 10:55:28 -0500
Subject: [Microscopy] TEM Fixation for botanical leaf sample (Switchgrass, Panicum virgatum)

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Dear Microscopists,

I have tried to prepare Switchgrass leaf samples twice with less than
desirable outcomes in both cases. I used 2% glutaraldehyde and 2%
paraformaldehyde in 50 mM sodium cacodylate, pH 7.0, for 6 hours at 22C
on a rotator. With a sharp razor blade I cut cross-sections of the leaf
blade while immersed in the fixative, and re-cut the samples to various
sizes of {1mm to 3mm in length. I tried some with and without aspiration
to 0.5 atm. Samples were washed in the same buffer, and post-fixed in 1%
OsO4 in the same buffer (2 hr), acetone dehydrated, and embedded in
Spurr's following very gradual infiltration with the same.

In all cases, every piece of tissue has intercellular air bubbles that
are not removed by the aspiration in fixative, and they remain into the
final resin. In addition, the bundle sheath cells "deflate" to some
extent, the collape causing distortion of the bundle sheath cells and
also mechanical disruption and breakage of the thinner, attached,
mesophyll cell walls. The bundle sheath cells (very thick-walled) are
also poorly infiltrated with resin. Some very few BS cells appear well
preserved and are fully infiltrated, but this is not the norm and the
tissue is difficult to work with due to the extensive poor infiltration
of the bundle sheath ring. The mesophyll cells look good aside from
physical disruption from the shrinking bundle sheath cells. I am
interested in the general ultrastructure, and the chloroplasts of the
mesophyll/bundle sheath cells in particular.

Does anyone have experience with tough grass samples that could help me
improve this preparation? It seems that conditions for good preservation
of the mesophyll and bundle sheath cells are quite different. Can anyone
suggest a reference protocol that has been successful with similar samples?

Thanks in advance.

Dale Callaham
University of Massachusetts, Amherst

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From: mdienelt-at-msn.com
Date: Thu, 31 May 2007 15:08:34 -0500
Subject: [Microscopy] viaWWW: Advice on critical point dryers

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Email: mdienelt-at-msn.com
Name: Margaret Dienelt

Organization: USDA, ARS, National Arboretum

Title-Subject: [Filtered] Advice on critical point dryers

Question: Hello everyone,

I've just learned we might have funds to replace our ancient,
tempermental critical point dryer and would greatly appreciate any
feedback anyone can give me on their CPD. I'm especially interested
in reliability - the valves in our current unit have been a problem
for years, requiring numerous repairs (usually after destroying a few
samples).

In addition to information anyone can share on specific models, I
also have two general questions:

What the pros and cons are to the two profiles, i.e. horizontal (like
Tousimis) and vertical (like Polaron)?

What are pros and cons to automatic vs manual models?

Basically, any wisdom you'd like to share about purchasing a new
critical point dryer will be more than welcome. Our new one will be
used in a plant pathology lab and will be used to process primarily
plant tissue. We don't need one for a clean room or one with the
jumbo chamber.

I'm sending this from my personal e-mail account since I'm having
problems with my government e-mail, but if replies are sent to
mdienelt-at-msn.com or margaret.dienelt-at-ars.usda.gov. I'll receive them
one way or the other.

Thank you!

Margaret



Margaret Dienelt
Plant Pathologist/Electron Microscopist
Floral and Nursery Plants Research Unit
National Arboretum, ARS, USDA

10300 Baltimore Avenue
Beltsville, MD 20705

(301)504-6097


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From: yitianp-at-ece.arizona.edu
Date: Thu, 31 May 2007 21:52:36 -0500
Subject: [Microscopy] viaWWW: Question about image on Glass

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Email: yitianp-at-ece.arizona.edu
Name: Yitian Peng

Organization: University of Arizona

Title-Subject: [Filtered] Question about image on Glass.

Question: Hi everyone,
I have some patterned Cr structure(width:100micro, height:60nm) on Glass.
Then I spin 200nm PMMA on the surface.
Then I Sputter 10nm Cr on the PMMA.
I want to using JSM 6400 SEM to look at the structure.
But I can't see anything?/
What's the problem?/
Thank you very much.
Yitian


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From: oddioeng-at-aol.com
Date: Thu, 31 May 2007 21:53:09 -0500
Subject: [Microscopy] viaWWW: Old Union Carbide UC-4 Ln2 dewar

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Email: oddioeng-at-aol.com
Name: J. Allen Williams, Jr

Organization: ORDELA, Inc.

Title-Subject: [Filtered] Old Union Carbide UC-4 Ln2 dewar

Question: Hello,
I have recently obtained an old UC-4 dewar and need to re-evacuate the vacuum chamber. My question is this; has anyone carried out this procedure? I am a little stumpped because there is a metal ball inserted in the port. Was this a poor attempt at a fix, or is it a membrane type flow valve?
Any answers, comments or suggestions would be nice.
Thanks,
Allen

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