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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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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 -- From: Dorothy Sorenson {dsoren-at-umich.edu}
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.microscopy.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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} 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 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.microscopy.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


==============================Original Headers==============================
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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.microscopy.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.microscopy.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.microscopy.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

Contents Retrieved from Microscopy Listserver Archives
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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

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

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8, 11 -- Subject: viaWWW: Hitachi SEM 2460N Service Contract
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==============================End of - Headers==============================




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
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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.microscopy.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
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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.microscopy.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==============================
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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.microscopy.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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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.microscopy.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








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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.microscopy.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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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

Contents Retrieved from Microscopy Listserver Archives
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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.microscopy.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

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

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

Contents Retrieved from Microscopy Listserver Archives
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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
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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.microscopy.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
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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: 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.microscopy.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.microscopy.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 -----
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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.microscopy.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.microscopy.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.microscopy.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

__________________________________________________
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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.microscopy.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.microscopy.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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6, 21 -- Subject: Carbon rod sharpener
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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
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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
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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
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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.microscopy.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.microscopy.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.microscopy.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
-----------------------------------------------------------------

==============================Original Headers==============================
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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.microscopy.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.microscopy.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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}

==============================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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

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
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

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 -- To: microscopy-at-microscopy.com
} 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
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

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
Technology on +61 3 5246 4000.



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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.microscopy.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.microscopy.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.microscopy.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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--
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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.microscopy.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.microscopy.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.microscopy.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
} _______________________________________
}
}
}
} ==============================Original Headers==============================
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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.microscopy.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.microscopy.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.microscopy.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.microscopy.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:
} ----------------------------------------------------------------------------
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} 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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--
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Calgary, AB, Canada T2N 4N1
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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
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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.microscopy.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.microscopy.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
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the intended recipient, please contact the sender by reply email and
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==============================Original Headers==============================
23, 34 -- From richard.beanland-at-bookham.com Wed May 9 04:25:50 2007
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23, 34 -- Subject: RE: [Microscopy] Silicon Cross-section sample preparation
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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.microscopy.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
} 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: 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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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.microscopy.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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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.microscopy.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.microscopy.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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==============================Original Headers==============================
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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.microscopy.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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} http://www.microscopy.com/MicroscopyListserver
} On-Line Help
} http://www.microscopy.com/MicroscopyListserver/FAQ.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
}
} ==============================Original
} Headers==============================
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} 2007
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***************************
Tamara Howard
Cell Biology & Physiology
UNM-HSC
Albuquerque, NM
***************************

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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.microscopy.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.microscopy.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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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.microscopy.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.microscopy.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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14:42:13 -0600
} 9, 19 -- From: "John Chandler" {jpchandl-at-mines.edu}
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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.microscopy.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.microscopy.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.microscopy.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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} 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 --
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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
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} } } Mitochondrial cristae
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} } } 8, 20 -- From: Tom W Bargar {tbargar-at-unmc.edu}
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} } 9, 21 -- Subject: Re: [Microscopy] Help with enhancing contrast of
} } Mitochondrial cristae
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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
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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.microscopy.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:
}
}
}
} ----------------------------------------------------------------------------
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} Society of America
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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
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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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} } } The Microscopy ListServer -- CoSponsor: The Microscopy
} } } Society of
} } } AmericaTo Subscribe/Unsubscribe --
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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:
} } }
} } } }
} } } }
} } } }
} } } }
} } } --------------------------------------------------------------------
} } } --------
} } } } The Microscopy ListServer -- CoSponsor: The
} } } } Microscopy Society of America
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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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} } } } 8, 20 -- Subject: Help with enhancing contrast of
} } } } Mitochondrial cristae
} } } } 8, 20 -- To: Microscopy-at-MSA.Microscopy.Com
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} } } } 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
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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.microscopy.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 -- 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.microscopy.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:
} } ---------------------------