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Good afternoon/evening,
Dear Dr. Ross,

I would like to be short....."glow discharging" is a kind of creating a
kind of "air plasma" in order to get formvar-coated grids stickier (= more
hydrophilic, discharging of surface charge).

This process necessarily /usually is done in a sputter coater or "plasma
etcher" or the like (must have a recipient and vacuum pump(s) rotary, and -
if (perhaps) you need higher vac, diffusion oil or tubomolecular pump...)
as well as an inbuilt ionizing source.

Often such a technique is used for getting formvar-coated grids (used for
negative staining procedure on virus particles) again hydrophilic as well
as stickier for the adsorption of virus particles to the formvar film.
I enclose here - FY convenience - a thread on that issue published in
Microscopy Today (and therefore was a theme via the MSA Listserver in
2005.........

NetNotes aus MICROSCOPY TODAY Vol. 13, No 3, May 2005, p. 64-65

Formvar Grids (de-charging by glow discharging and other methods)
----------------
Q: We have been using Formvar/carbon coated grids for spreading magnetic
nano-particles in water and for negative staining. But as you know, an
aqueous solution does not spread well on grids because of the charge
characteristics on the film surface. The only way I know of for making the
film surface more hydrophilic is to do a glow discharge in a sputter
coater, but we do not have such a device. I heard treating coated grids
with ethanol vapor works, but not for me. Does anyone have any other tricks
or suggestions? Hong Yi {hyi-at-emory.edu} 20 Feb 2005
-----------
Answers:
Do you have a vacuum evaporator? It's simple to rig it for glow discharge
with an inexpensive Tesla coil. A plastic vacuum desiccator, the same Tesla
coil, and a rough vacuum source will do the job also.
Caroline Schooley {schooley-at-mcn.org} 21 Feb 2005
------------
I don't think that a splitter coater, old or new, is going to solve your
problem. I have not heard of ethanol vapors making a carbon grid more
hydrophilic. Carbon coated grids lose their hydrophilic nature as they age
and they become more hydrophobic. The process can be "reversed" by
(a) exposure to an RF "air" plasma in a small plasma etcher (effect will
last 60-90 days) or
(b) a thin evaporation of VictawetO onto the grids. Our own studies would
suggest that Victawet can keep the grids highly hydrophilic essentially
forever (e.g. more than one year).
Just remember that it is a phosphate based surfactant so if you are doing
elemental analysis work, you might not want to have P showing up in your
data. But if you have an ordinary vacuum evaporator and tungsten baskets,
and don't have a plasma etcher, you can solve your problem with Victawet.
The best bet for having carbon coated grids with the greatest hydrophilic
characteristics is to make or purchase your carbon coated grids always
"fresh': If the grids are purchased, and their age is uncertain, contact
the manufacturer of the carbon coated grids, give them the lot number and
then you will know.
Charles A. Garber {cgarber-at-2spi.com} 21 Feb 2005
----------
Another method occasionally used to make C-coated grids hydrophilic is to
expose them for some minutes to the direct illumination of a UV lamp. This
is done at normal atmospheric pressure, so it needs no vacuum technology.
James Chalcroft {jchalcro-at-neuro.mpg.de} 21 Feb 2005
----------
I personally don't like glow discharge at all: it's very difficult to
reproduce. It depends on the equipment and there is no way to control the
"amount" of discharge. I use 0.5-1 % poly-lysine from any of the EM
suppliers (don't try to make the solution yourself since there is some
trick required). Place the EM grid on a 10 ?l poly-lysine drop for 5-10
min, wash on a few drops of deionized water, air dry - good for at least a
month. Alcian Blue works in the similar way with a similar result. Of
course, these methods will only work for positively charged molecules.
Sergey Ryazantsev {sryazant-at-ucla.edu} 22 Feb 2005

hope this answers your question...
best regards

Wolfgang MUSS
Salzburg, Austria
=============================} } { {========================
----------
Von: RossLM-at-missouri.edu[SMTP:RossLM-at-missouri.edu]
Antwort an: RossLM-at-missouri.edu
Gesendet: Donnerstag, 01. Juni 2006 20:50
An: W.Muss-at-salk.at
Betreff: [Microscopy] Re: Sample prep for disbersing powder samples

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Thanks,
Lou Ross

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--
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Bill, We too have used the method you've described, but I am not
familiar with "glow discharge the grids". What is tis and how does it
work?

Thanks,
Lou Ross

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

Hello Lou,

Essentially, glow discharge is a procedure that you can carry out on
a standard vacuum evaporator that is equipped with a high voltage AC
input. Unfortunately, not many vac evaporators have this accessory.
In operation, specimens are loaded into the chamber and one end of
about 8-12 inches of high purity aluminum wire (3-5 mm) is plugged
into the HV feedthrough and a large loop is formed to surround the
grids. The chamber is pumped down using only the rotary pump until
you get approximately 50-100 millitorr vacuum. At this point, you
turn up the AC high voltage (using a variAC) and you will generate a
plasma similar to what one sees in a sputter coater. The plasma is
thought to both clean the grid surface as well as imparting a charge
that renders the grid surface (of carbon-coated grids) hydrophilic.
The effect lasts about a day--less if in a high humidity environment.

We have been able to approach this technique by placing the grids
inside a sputter coater and shielding them from direct line of sight
with the target (to prevent coating with metal). The plasma generated
will impart a hydrophobic character to the grids. But you may also
get metal deposition, so do some trial runs first.

I have been depositing a lot of nanocrystals as you describe by
directly depositing them on carbon or silicon substrates. The powders
are suspended in acetone, shaken and (after allowing the large
particles to settle) a small volume taken up in a micropitettor
(about 10-20 microliters). This is then dropped directly onto the
grid surface from a distance of 10 or so mm.

I can take a picture of the glow discharge setup if you like, Lou.

JB

} }
} } On Jun 1, 2006, at 8:40 AM, lkrupp-at-us.ibm.com wrote:
} }
} } } I need to make samples disbersed on carbon-coated grids for collecting
} } } some
} } } EELS standards of various oxides in powder form. I have Maganese,
} } } Titiatium, and Ruthenium oxides in various states. Anyone know a good
} } } prep
} } } that will evenly disburse a thin layer onto a carbon-coated grid? My
} } } first
} } } thought would be to mix with a solvent and put a drop on a grid, but I
} } } figured someone here has probably already done this.
} } }
} } Dear Leslie,
} } My choice would be your first thought--it worked very well for
} } river-bottom sediment. I would try various dilutions to find the one
} } that gives the desired distribution of particles, and I would
} } glow-discharge the grids. If using water leads to aggregation, try an
} } organic solvent, and if the particles still have a tendency to
} } aggregate, try putting the grid with the drop of solvent in an
} } oven--faster evaporation might overcome that tendency.
} } Yours,
} } Bill Tivol, PhD
} } EM Scientist and Manager
} } Cryo-Electron 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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}
} --
} Senior Electron Microscope Specialist
} Electron Microscopy Core Facility
} W136 Veterinary Medicine
} University of Missouri
} Columbia, MO 65211-5120
} (573) 882-4777, fax 884=2227
} email: rosslm-at-missouri.edu
} http://www.emc.missouri.edu
}
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Vendors who recondition LKB 7801 B knife makers please contact me.
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Hi:

I have a couple of old SEM's looking for a place to land. An ISI WB-6
and a Hitachi S-570.

Thanks

Jon
--

Jonathan Krupp
Microscopy & Imaging Lab
C230 Earth & Marine Science
University of California
Santa Cruz, CA 95064
(831) 459-2477
jmkrupp-at-ucsc.edu

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} ### apologies for multiple postings
}
}
}
}
} The Department of Sedimentology and Environmental Geology,
} Geoscience Center of the University
} of Goettingen, is offering a
}
}
} post-doc position in Sedimentology / Sedimentary Petrology
}
}
} beginning 1st October, 2006. The appointment is for one year with an
} option for another six month.
} Salary is based on the wage agreement of the German civil service
} (75% BAT IIa, approx. 30.000 EUR p.a.).
}
} The Geoscience Center in Goettingen offers a wide range of
} state-of-the-art analytical facilities and an excellent environment
} for geological, sedimentological and geochemical research (
} http://www.gzg.uni-goettingen.de ).
} We are seeking for a gifted and motivated scientist with a several
} years perspective for our research team.
}
} The successful candidate has finished his/her excellent doctoral
} thesis no longer than two years ago. Ongoing and future research
} should focus on one of the major research topics of the department,
} such as exhumation & erosion, weathering processes, sediment
} geochemistry and/or
} provenance analysis. During the one-year period offered here a
} research proposal should be set up to continue research on soft money.
}
} Teaching duties comprise sedimentary petrology as well as
} sedimentary facies and basin analysis at
} various levels, as well as field trips. The University of Goettingen
} is attempting to increase the
} proportion of women on the scientific staff and strongly encourages
} qualified women to apply.
} Disabled candidates with equal qualification will be preferred.
}
} Applicants should send a letter of interest, curriculum vitae, a
} brief statement of research interests, list of publications and
} contact information for two referees, no later than June 30, 2006,
} to Prof. Dr. Hilmar von Eynatten, Department of Sedimentology and
} Environmental Geology, Geoscience Center Goettingen,
} Goldschmidtstrasse 3, D-37077 Goettingen, Germany
} (hilmar.von.eynatten-at-geo.uni-goettingen.de,
} http://www.sediment.uni-goettingen.de ).
}

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Microscopists,

We cure blocks in an oven at 65 degrees. Is there any information
regarding the evolution of toxic fumes during curing?

--

____________________

Bob Josephs
773 702-1077

http://gingi.uchicago.edu

Most People learn form their own mistakes -the fortunate learn from
those of others.

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Colleagues

The Listserver Archives are now updated through
May 31, 2006 and can be accessed at

http://www.microscopy.com

Nestor
Your Friendly Neighborhood SysOp

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Hello,
Do you know guys where to get magnetron sputtered tantalum? No thickness
restriction, I just wanna get a flat tantalum substrate.

Thanks

Hany

--
**********************************************************
Hany Ramadan
Graduate student
Chemistry department
McMaster university, Hamilton, Ontario, Canada
905-525-9140 x: 26322
elsayeh-at-mcmaster.ca
**********************************************************

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This Question/Comment was submitted to the Microscopy Listserver
using the WWW based Form at http://microscopy.com/MicroscopyListserver/MLFormMail.html
---------------------------------------------------------------------------
Remember this posting is most likely not from a Subscriber, so when replying
please copy both lynda-at-biotech.ufl.edu as well as the MIcroscopy Listserver
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Email: lynda-at-biotech.ufl.edu
Name: Lynda Schneider

Organization: University of Florida

Title-Subject: [Filtered] SEM of bacteria in milk

Question: Hello all,

I have a SEM sample of bacteria in milk. Does anyone have any suggestions on how to reduce the milk debris/fat leaving the bacteria intact? I have centrifuged the sample, took from the formed pellet and deposited onto a millipore filter on a pump, buffer washed several times and followed a standard fixation protocol though HMDS. Bacteria of other samples (not in milk) looked great, but the milk sample left way too much debris behind.

Thanks in advance for any suggestions.
Lynda Schneider
University of Florida

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

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Hello to all,

Am looking for the schematic for the above TC Controller, can anybody help
please?

Varian does not have the info.

Cheers
Peter

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It's that time again!
We are in need for volunteers for the M&M meeting
in Chicago.

Student volunteers will be paid through a voucher system.
Other volunteers will receive benefits according to the
hours provided. As soon as you are accepted as a volunteer,
your name will be provided to meeting registration. All
efforts will be made to place volunteers for sessions and
booths that they are interested in.

For more information and a schedule of sessions/jobs, please
contact John Shields at jshields-at-cb.uga.edu (706-542-4080),
or to Bill Monroe at monroe-at-emcenter.msstate.edu (662-325-
3019).
John P. Shields
Center for Ultrastructural Research
151 Barrow Hall
University of Georgia
Athens, GA 30602

706-542-4080

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Dear Fellow Microscopists,

We have a grad student user who is interested in thin sectioning carbon
nanotubes. The tubes are in clusters and suspended in some buffer. Is it
possible to embed the clumps in epoxy resin and section them? Is diamond knife
going to survive after sectioning nanotubes? The average diameter of those tubes
is approx. 1.4 nm. Any suggestion will be greatly appreciated.

Thanks in advance,

Soumitra

******************************************************************************
Soumitra Ghoshroy
College Associate Professor, Biology
Director, Electron Microscopy Lab
New Mexico State University
Las Cruces, NM 88003
Tel: 505-646-3268 (office)
505-646-3283 (lab)
Fax: 505-646-3282
http://biology-web.nmsu.edu/ghoshroy/ghoshroy.htm
http://emldata.nmsu.edu

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Dear Colleagues...

We are planning to examine some cellulose made microspheres containing
albumin with transmission EM. Is there anybody experienced with this
material before? Are routine EM tissue processing methods suitable with this
material? Thanks for any kind of helps...

Dr. Necat Yilmaz
Mersin Universitesi Tip Fakultesi
Histoloji ve Embriyoloji Anabilim Dali

==============================Original Headers==============================
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Dear listers,

We solved our contrasting problems by just following
simple rules found in this list. If it solved our
problems, I thought it could be useful for others.

- We prepared "carbonated lead citrate" (the time
required to "burn" it was much shorter as proposed
though)
- We bought decarbonated NaOH to a big american
supplier

These are very simple rules which make all the
difference between "I see nothing" and "oh my god I
never had such a nice staining". And that's true: I've
never ever had such a beautiful staining!

Thank you again for your support.

St�phane

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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7, 18 -- Subject: Lead citrate staining: Thank you again and again....
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Group

Need some help on preparing samples of the following algae for
observation in SEM and the take-up of heavy metals in cell structure.

Spirogyra Green Algae
Chlorella Green Algae
Fischerella Blue-Green Algae

EDS X-ray maps will be attempted to look at metal distribution.

Have not looked at this type of material before so any help would be
useful.

Thanks

Roy Beavers

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

==============================Original Headers==============================
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Dear listers,

Here is another weird idea from me :-D
I would like to localize a fluorescent molecule in the
transversal plane of a cell monolayer(not from above).
We have no material for the preparation of
histological sections, but we have all necessary for
TEM. I wonder if I could not embed the monolayer in
Epon, then cut transversal semi-thin sections and
observe in epifluroscence.
Would Epon hinder fluorescence?
Could it be done in another resin than Epon?

Regards,

St�phane

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Dear All,

I would appreciate any comments, references, or first-hand experiences from
those using a combined FIB/SEM+Cryo instrument with & without a field
emission gun - where biological or polymer samples can be sliced by the ion
beam then imaged. I would be interested to know things such as:

Why you chose the instrument you use?
How easy is it in practice to do this sort of ion beam slicing & is it
restricted to particular types of biological or polymer samples?
Does the FIB mode impact on the column cleanliness & resolution over time
particularly if used to thin samples for TEM?
Is the instrument expensive to run & does it require an experienced
operator?
Anything else you feel would help those looking at this type of instrument
for a multi-user facility.

Many thanks
Ursula
-------------------

Ursula J. Potter
Centre for Electron Optical Studies (CEOS)
Building 3 West 2.15
The University of Bath
Claverton Down
Bath BA2 7AY
UK
Tel: 01225 385651
Email: U.J.Potter-at-bath.ac.uk

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Hello:

Are there anyone out there using this model? I would need specially the
low level protocol on commanding it's RS232C port for lab automation
purposes on our lab set-up at the university. Your help is greatly
appreciated.

Regards,

Berns B.

--
--------------------------------------------
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University Research Associate
National Institute of Physics
University of the Philippines
Diliman Campus 1101 Quezon City
Philippines
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Stephane,
Not so weird, but might not work. The two
things to worry about are autofluorescence and
loss of your fluorochrome. If you have some of
your cells (unlabeled) in epon you can check some
sections under a fluorescence microscope and see
about the autofuor. I think the loss of
fluorescence is a more serious problem, either
because it is extracted during
dehydration/infiltration or quenched by the
embedment. You may have to embed a labeled sample
and find out the hard way!

Tobias
}
} Dear listers,
}
} Here is another weird idea from me :-D
} I would like to localize a fluorescent molecule in the
} transversal plane of a cell monolayer(not from above).
} We have no material for the preparation of
} histological sections, but we have all necessary for
} TEM. I wonder if I could not embed the monolayer in
} Epon, then cut transversal semi-thin sections and
} observe in epifluroscence.
} Would Epon hinder fluorescence?
} Could it be done in another resin than Epon?
}
} Regards,
}
} St�phane
}
}
} __________________________________________________
} Do You Yahoo!?
} Tired of spam? Yahoo! Mail has the best spam protection around
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} ==============================Original Headers==============================
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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

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Regarding the idea of localizing fluorescence label in epon, I would suggest
that you not use Epon...there are autoflourescence problems (at least in our
hands, using Spurrs resin). However, I think you may be happy using LR
White, polymerized at 60C. We've done some similar work here and have been
very impressed with the results.

Good luck,

Doug

Douglas R. Keene
Assistant Investigator
Micro-Imaging Center
Shriners Hospital for Children
3102 S.W. Sam Jackson Park Road
Portland, Oregon 97239
503-221-3434
drk-at-shcc.org
-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Wednesday, June 07, 2006 1:10 AM
To: drk-at-SHCC.org

Dear listers,

Here is another weird idea from me :-D
I would like to localize a fluorescent molecule in the
transversal plane of a cell monolayer(not from above).
We have no material for the preparation of
histological sections, but we have all necessary for
TEM. I wonder if I could not embed the monolayer in
Epon, then cut transversal semi-thin sections and
observe in epifluroscence.
Would Epon hinder fluorescence?
Could it be done in another resin than Epon?

Regards,

St�phane

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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St�phane

Don't ever hesitate. some of my best things have happened because of an
offbeat idea or insane shot in the dark question during discussions. Of
course, I don't want to admit that to my wife. She hates to be referred
to as an offbeat idea, just as the best thing that ever happened to me
O:-) .

Frankly, you're keeping the rest of us fresh and on our toes.

Paul

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Dear without an I,
Success will depend upon the fluorophore. FITC and TRITC will lose
their fluorescence. Alexa fluor dyes and cyanine dyes will work
well. Actually, I think with reduced photobleaching.
Autofluorescence will depend upon imaging modality. Spurr's looks
great under the confocal, but the autolfuorescence creates an
impossible haze with epi-fluorescence on thick samples.
Deconvolution will clean up the haze. 3 micron sections looked very
good. the new formulation of spurr's is too brittle so I'm going to
try Eponate 812 or something similar for my next round of embedded
fluorescence. I tried Histo-Resin and found the autofluorescence was
noticeable under confocal, but didn't try LR-White.

Hardie, MacDonald, Rubel, Brain Res. 1000:200-210, 2003

Regards,
Glen
On Jun 7, 2006, at 10:49 AM, drk-at-SHCC.org wrote:

}
}
}
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} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
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}
} Regarding the idea of localizing fluorescence label in epon, I
} would suggest
} that you not use Epon...there are autoflourescence problems (at
} least in our
} hands, using Spurrs resin). However, I think you may be happy
} using LR
} White, polymerized at 60C. We've done some similar work here and
} have been
} very impressed with the results.
}
} Good luck,
}
} Doug
}
} Douglas R. Keene
} Assistant Investigator
} Micro-Imaging Center
} Shriners Hospital for Children
} 3102 S.W. Sam Jackson Park Road
} Portland, Oregon 97239
} 503-221-3434
} drk-at-shcc.org
} -----Original Message-----
} X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Wednesday, June 07, 2006 1:10 AM
} To: drk-at-SHCC.org
} Subject: [Microscopy] weird idea: fluorescence in Epon
}
}
}
}
} ----------------------------------------------------------------------
} ------
} 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,
}
} Here is another weird idea from me :-D
} I would like to localize a fluorescent molecule in the
} transversal plane of a cell monolayer(not from above).
} We have no material for the preparation of
} histological sections, but we have all necessary for
} TEM. I wonder if I could not embed the monolayer in
} Epon, then cut transversal semi-thin sections and
} observe in epifluroscence.
} Would Epon hinder fluorescence?
} Could it be done in another resin than Epon?
}
} Regards,
}
} St�phane
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Email: wim.vandenbroeck-at-UGent.be
Name: Wim Van den Broeck

Organization: Morphology, Ghent University

Title-Subject: [Filtered] TEM - virus particles in cell culture

Question: Dear Friends,

I need to visualize viral particles grown in cell culture, but I do not have any experience with that (only with tissue samples). Could you give me any advice (collection of cells, fixation, embedding, �.).

Thanks in advance,

Wim Van den Broeck.

Wim Van den Broeck, DVM, MSc, PhD
Professor in Cytology and Histology
Department of Morphology
Faculty of Veterinary Medicine, Ghent University
Salisburylaan 133, B-9820 Merelbeke,
BELGIUM
tel.: +32 (0)9 264 77 16
fax: +32 (0)9 264 77 90
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13, 14 -- From zaluzec-at-microscopy.com Wed Jun 7 19:36:50 2006
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Hello,

While on the Medical School Faculty of USC, Los
Angeles I did extensive studies of hepatitis B virus
in liver explants. The technique is straightforward.

Remove cells using a soft spatula or a pipette
depending on whether or not the cells are attached to
the surface of the culture container.

Place the cells in a centrifuge tube containg
fixative. Carry out the dehydration and embedding
fluid impregnation process in the centrifuge tube
spinning between each stage. Finally transfer the
embedding mixture with the cells into a Beem-type
embedding capsule and spin the cells down into the
tip.

If you wish I can send you the reference for one of my
publications that outlines the techniques employed.

Best wishes,

Ted Dunn
The EMscope Company Ltd.
Thailand

--- wim.vandenbroeck-at-UGent.be wrote:

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} Organization: Morphology, Ghent University
}
} Title-Subject: [Filtered] TEM - virus particles in
} cell culture
}
} Question: Dear Friends,
}
} I need to visualize viral particles grown in cell
} culture, but I do not have any experience with that
} (only with tissue samples). Could you give me any
} advice (collection of cells, fixation, embedding,
} �.).
}
} Thanks in advance,
}
} Wim Van den Broeck.
}
} Wim Van den Broeck, DVM, MSc, PhD
} Professor in Cytology and Histology
} Department of Morphology
} Faculty of Veterinary Medicine, Ghent University
} Salisburylaan 133, B-9820 Merelbeke,
} BELGIUM
} tel.: +32 (0)9 264 77 16
} fax: +32 (0)9 264 77 90
} Email: wim.vandenbroeck-at-UGent.be
}
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} 13, 14 -- From zaluzec-at-microscopy.com Wed Jun 7
} 19:36:50 2006
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} MicroscopyListserver)
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12, 20 -- From drteddunne-at-yahoo.com Wed Jun 7 23:57:20 2006
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Dear Wim,

It depends if you need to visualize the viral
particles inside the cells or not. If so, I would
consider usual protocol for classical cell morphology.

I already observed viral particles in cells I observed
for other purposes.
If you need the viral particles alone, you have 2
solutions depending on the concentration of the virus
in medium.

1) If it is concentrated, you just collect the
supernatant, centrifuge 5 min at 5000 RPM to pellet
the cells and cell debris, and collect the supernatant
again.

2) If it is diluted, you have to find an ultrafast
centrifuge and perform an additional centrifugation to
pellet your virus particles and concentrate them.

Then you can just do a negative staining (PTA worked
well with rhinoviruses for me). You will find tons of
protocols on the net.

Good luck.

Stephane

--- wim.vandenbroeck-at-UGent.be wrote:

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} Email: wim.vandenbroeck-at-UGent.be
} Name: Wim Van den Broeck
}
} Organization: Morphology, Ghent University
}
} Title-Subject: [Filtered] TEM - virus particles in
} cell culture
}
} Question: Dear Friends,
}
} I need to visualize viral particles grown in cell
} culture, but I do not have any experience with that
} (only with tissue samples). Could you give me any
} advice (collection of cells, fixation, embedding,
} �.).
}
} Thanks in advance,
}
} Wim Van den Broeck.
}
} Wim Van den Broeck, DVM, MSc, PhD
} Professor in Cytology and Histology
} Department of Morphology
} Faculty of Veterinary Medicine, Ghent University
} Salisburylaan 133, B-9820 Merelbeke,
} BELGIUM
} tel.: +32 (0)9 264 77 16
} fax: +32 (0)9 264 77 90
} Email: wim.vandenbroeck-at-UGent.be
}
}
}
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} 19:36:50 2006
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} MicroscopyListserver)
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} cell culture
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13, 20 -- From nizets2-at-yahoo.com Thu Jun 8 01:33:06 2006
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For those interested in the technique, it is simply
available in the internet site of EMS. Look in
Technical-} tips and articles--} "A stable lead
staining solution".

For the very practical side of the preparation of
carbonated lead citrate, here is what I exactly did:
I heated 2g at 250�C for 3h in a ceramic beaker
(called "melting pot" on EMS site) with a glass cover.
I have the chance to work with chemists so the
equipment is not a problem: I used a
temperature-controlled special oven connected to a
mobile hood.

I controlled the color of the powder. It turns grey,
then yellowish grey, a little bit like humid sand. I
stopped then. If one is unsure, one can still heat
several beakers each containing 1g of lead citrate and
heat them for different times (2h, 3h, 4h).

I think it is pretty straithforward because it worked
the first time I tried. I stored the powder in a
sealed glass flask (don't know if it is the best
though).

Now IMHO, seeing the easiness of the technique and the
cheap price of carbonate-free NaOH, there is really no
reason NOT to do it :-D

Stephane

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8, 19 -- From nizets2-at-yahoo.com Thu Jun 8 01:55:43 2006
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Hello,

Can anyone suggest good scanner to scan TEM negatives
and prints and and latest Imange Analysis software.

Regards
Shrunali Kulkarni
Scientist
IMTech

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Dear listers,

Thank you for your numerous answers.
Lots of them show a concern about autofluorescence,
but I must say that I was more concerned about the
quenching by Epon and the processing steps:
dehydration and embedding. I wondered if and how it
could "damage" the fluorochrome.

We use Epon 812 (glycidether 100) and we just cut
empty block at 300 to 500nm thickness to observe the
autofluorescence with the "green" filter (our
fluorochrome is Alexa488). Actually there IS some
autofluorescence, but not dramatic even with a
thickness of 500nm. I don't think it will perturb the
observation, but it depends of course how well the
alexa will sustain the processing.

An excellent remark was made, pointing out that
glutaraldehyde must be avoided because it brings
autofluorescence. I knew it but recalling me was not
bad :-D Anyway glutar is not necessary for LM
observation.

I will definitely try light fixation (4% PFA for 20
min), fast dehydration in alcohol and direct embedding
in Epon.

Another concern is the localisation of the cells and
cell comparments. Do some of you have an idea how well
work DIC or phase contrast with cells embedded in
Epon?
Could I just use general staining protocols
hematoxylin-eosin before embedding in Epon?

Stephane

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Dear Stephane,
Don't use eosin...it fluoresces very intensely (in the usual
"Rhodamine" wavelengths). In fact, the slide I use when teaching
people how to use the confocal is a standard paraffin section stained
with H&E. It never seems to bleach, and it fluoresces like mad at
488 (the connective tissue), 543, and 633.

I've looked at semi-thin resin sections with DIC...it works.
Lee
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I put my TEM negatives on a lightbox and photograph them with a Canon G3
digital camera. Then I convert to a positive with PhotoShop.

Geoff

aarti_harle-at-yahoo.co.in wrote:

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Hi Shrunali,

I'll send a pdf of a short article on scanners I wrote for Microscopy Today
(May 2006). My personal favourite scanner is the Epson V750 Pro (�600) and
the Cheaper sister the Epson V700 (�400). The older model Epson 4990 Photo
is not quite as good but cheaper at �280. All these scan film up to full A4
size, and have an easy to use twain interface. They also do reflective scans
for paper and photographs. My favourite scanner review site for pro-sumer
(i.e. cheap) scanners is http://www.photo-i.co.uk.

As mentioned you can use a camera and stand to copy film (used to be called
an epidiascope in my Quantimet image analyser days) - and you can get pretty
good optical microscope images just by resting a compact digital camera lens
against the eyepiece. These days though I think for film, 6,400 dpi
pro-sumer flatbed scanners do it better, more conveniently and relatively
cheaply.

For image analysis there's lot to choose from, e.g.
Improvision OpenLabs and Velocity (http://www.improvision.co.uk)
ImageProPlus (http://www.mediacy.com)
Kinetic Imaging (now http://www.andor.com/products)
MetaMorph (http://www.moleculardevices.com/pages/software/metamorph.html)

These packages are expensive though (typically �3k+). They all deal with
scanned tif images (and much more).

However if you don't have �3,000 for the above software, have at look at the
freeware ImageJ (for the PC) at http://rsb.info.nih.gov/ij - you often need
some of the plug-ins as well (there is also the sister program NIHImage for
Apple fans at http://rsb.info.nih.gov/nih-image). Not always an easy program
to get to grips with quickly (like most image analysis software), but quite
powerful with plug-ins. For simple image manipulation, Photoshop CS2 is the
obvious choice - otherwise the cut-down Photoshop Elements is pretty good
and is supplied free with the above scanners (as a slightly older version).

Regards

Keith

----------------------------------------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology Division
Institute of Ophthalmology
University College London
11-43 Bath Street
London EC1V 9EL

Tel: 020 7608 4050
Fax: 020 7608 4034
email: keith.morris-at-ucl.ac.uk

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X-from: aarti_harle-at-yahoo.co.in [mailto:aarti_harle-at-yahoo.co.in]
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Hello,

Can anyone suggest good scanner to scan TEM negatives
and prints and and latest Imange Analysis software.

Regards
Shrunali Kulkarni
Scientist
IMTech

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Wonderful weird idea!

I wonder if the autofluorescence might be due
primarily to the MNA (aka NMA) in the usual Luft
formulation?

When i found that NMA caused the cured resin to
react horribly with MnO4 section stain ---
(useful because the Mn04-Pb staining sequence
gives much more contrast than any other stain
we've ever tried) --- I learned to avoid MNA and
just empirically readjusted DDSA and Epon ratios
empirically for a firm enough cured resin,
needing no MNA, yet compatible with MnO4
staining. However, I soon abandoned Epon and
Spurr of all kinds in favor of an Araldite
506-DDSA-DER 736 mixture (10:15:2 gm by weight)
because it sections thinner and accepts Mn-Pb
staining with less graininess. So that Araldite
mix is what i will try soon with Alexa
488-phalloidin.

-mike reedy-
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Michael K. Reedy, M.D.
Duke Univ. Med. Center
Dept. Cell Biology, Box 3011 (for U.S. Mail)
458 Alex Sands Bldg, Research Dr (courier)
Durham, NC 27710

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Hi Mike,
Interesting thought about the NMA, and your Araldite recipe looks
worth a try. I had dropped the DMAE in Spurr's to 1/2 and then to
1/4 of the original recipe, thinking that it might generate radicals
reacting with the fluorophores. The result was slightly reduced
autofluorescence after a 3 day cure time at 60 C. No apparent effect
on the fluorophores themselves, but didn't spend any time measuring.

FYI, alcohols will cause phalloidin to dissociate from the actin.
Maybe it will survive a higher alcohol, propyl or butyl, but an actin
antibody would be a better choice if needing to dehydrate the sample.

Regards,
Glen

Glen MacDonald
Core for Communication Research
Virginia Merrill Bloedel Hearing Research Center
Box 357923
University of Washington
Seattle, WA 98195-7923 USA
(206) 616-4156
glenmac-at-u.washington.edu

************************************************************************
******
The box said "Requires Windows 95 or better", so I bought a Macintosh.
************************************************************************
******

On Jun 8, 2006, at 9:17 AM, Mike Reedy wrote:

} Wonderful weird idea!
}
} I wonder if the autofluorescence might be due primarily to the MNA
} (aka NMA) in the usual Luft formulation?
}
} When i found that NMA caused the cured resin to react horribly with
} MnO4 section stain --- (useful because the Mn04-Pb staining
} sequence gives much more contrast than any other stain we've ever
} tried) --- I learned to avoid MNA and just empirically readjusted
} DDSA and Epon ratios empirically for a firm enough cured resin,
} needing no MNA, yet compatible with MnO4 staining. However, I soon
} abandoned Epon and Spurr of all kinds in favor of an Araldite 506-
} DDSA-DER 736 mixture (10:15:2 gm by weight) because it sections
} thinner and accepts Mn-Pb staining with less graininess. So that
} Araldite mix is what i will try soon with Alexa 488-phalloidin.
}
} -mike reedy-
} } ---------------------------------------------------------------------
} } -------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America
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} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
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} }
} } Dear without an I,
} } Success will depend upon the fluorophore. FITC and TRITC will lose
} } their fluorescence. Alexa fluor dyes and cyanine dyes will work
} } well. Actually, I think with reduced photobleaching.
} } Autofluorescence will depend upon imaging modality. Spurr's looks
} } great under the confocal, but the autolfuorescence creates an
} } impossible haze with epi-fluorescence on thick samples.
} } Deconvolution will clean up the haze. 3 micron sections looked very
} } good. the new formulation of spurr's is too brittle so I'm going to
} } try Eponate 812 or something similar for my next round of embedded
} } fluorescence. I tried Histo-Resin and found the autofluorescence was
} } noticeable under confocal, but didn't try LR-White.
} }
} } Hardie, MacDonald, Rubel, Brain Res. 1000:200-210, 2003
} }
} } Regards,
} } Glen
} } On Jun 7, 2006, at 10:49 AM, drk-at-SHCC.org wrote:
} }
} } }
} } }
} } }
} } }
} } } --------------------------------------------------------------------
} } } --
} } } ------
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} } } America
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} } } FAQ.html
} } }
} } } --------------------------------------------------------------------
} } } --
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} } }
} } } Regarding the idea of localizing fluorescence label in epon, I
} } } would suggest
} } } that you not use Epon...there are autoflourescence problems (at
} } } least in our
} } } hands, using Spurrs resin). However, I think you may be happy
} } } using LR
} } } White, polymerized at 60C. We've done some similar work here and
} } } have been
} } } very impressed with the results.
} } }
} } } Good luck,
} } }
} } } Doug
} } }
} } } Douglas R. Keene
} } } Assistant Investigator
} } } Micro-Imaging Center
} } } Shriners Hospital for Children
} } } 3102 S.W. Sam Jackson Park Road
} } } Portland, Oregon 97239
} } } 503-221-3434
} } } drk-at-shcc.org
} } } -----Original Message-----
} } } X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} } } Sent: Wednesday, June 07, 2006 1:10 AM
} } } To: drk-at-SHCC.org
} } } Subject: [Microscopy] weird idea: fluorescence in Epon
} } }
} } }
} } }
} } }
} } }
} } } --------------------------------------------------------------------
} } } --
} } } ------
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } } America
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} } } --
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} } }
} } } Dear listers,
} } }
} } } Here is another weird idea from me :-D
} } } I would like to localize a fluorescent molecule in the
} } } transversal plane of a cell monolayer(not from above).
} } } We have no material for the preparation of
} } } histological sections, but we have all necessary for
} } } TEM. I wonder if I could not embed the monolayer in
} } } Epon, then cut transversal semi-thin sections and
} } } observe in epifluroscence.
} } } Would Epon hinder fluorescence?
} } } Could it be done in another resin than Epon?
} } }
} } } Regards,
} } }
} } } St�phane
} } }

==============================Original Headers==============================
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Shrunali
Could I suggest you check out the Microscopy List Server Archive? There has
been extensive and intelligent discussion about this subject quite
frequently over the last couple of years

Richard Harris
Laboratory Supervisor,
Imaging and Analysis
Department of Biology,
University of Western Ontario,
London Ontario, CANADA.
N6A 5B7
Ph. 519-661-2111 ext. 86780
Fax 519-661-3935

-----Original Message-----
X-from: aarti_harle-at-yahoo.co.in [mailto:aarti_harle-at-yahoo.co.in]
Sent: June 8, 2006 7:48 AM
To: rjharris-at-uwo.ca

Hello,

Can anyone suggest good scanner to scan TEM negatives and prints and and
latest Imange Analysis software.

Regards
Shrunali Kulkarni
Scientist
IMTech

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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4, 18 -- From aarti_harle-at-yahoo.co.in Thu Jun 8 06:42:41 2006 4, 18 --
Received: from web8324.mail.in.yahoo.com (web8324.mail.in.yahoo.com
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-0700 (PDT) 4, 18 -- From: Aarti Harle {aarti_harle-at-yahoo.co.in} 4, 18 --

You can purchase many different scanners for prices under $500 that will do
the job for routine negatives. What is critical is that they are capable of
both reflective and transmittal illumination. For negatives you need to be
able to transmit the light through the negative while normal scanning of
documents utilizes reflective light.

We normally scan routine negatives at 600dpi. However, if we need to
enlarge the image substantially, we will often scan at 1200dpi or higher.

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

} From: {mcauliff-at-umdnj.edu}
} Reply-To: {mcauliff-at-umdnj.edu}
} Date: Thu, 8 Jun 2006 08:37:52 -0500
} To: {dsherman-at-purdue.edu}
} Subject: [Microscopy] Re: Scanner
}
}
}
}
} ----------------------------------------------------------------------------
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}
} I put my TEM negatives on a lightbox and photograph them with a Canon G3
} digital camera. Then I convert to a positive with PhotoShop.
}
} Geoff
}
}
} aarti_harle-at-yahoo.co.in wrote:
}
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} } ----------------------------------------------------------------------------
} }
} } Hello,
} }
} } Can anyone suggest good scanner to scan TEM negatives
} } and prints and and latest Imange Analysis software.
} }
} } Regards
} } Shrunali Kulkarni
} } Scientist
} } IMTech
} }
} } __________________________________________________
} } Do You Yahoo!?
} } Tired of spam? Yahoo! Mail has the best spam protection around
} } http://mail.yahoo.com
} }
} }
} }
}
}
} --
} --
} **********************************************
} Geoff McAuliffe, Ph.D.
} Neuroscience and Cell Biology
} Robert Wood Johnson Medical School
} 675 Hoes Lane, Piscataway, NJ 08854
} voice: (732)-235-4583; fax: -4029
} mcauliff-at-umdnj.edu
} **********************************************
}
}
}
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Seeing at the Nanoscale IV, July 17-20, University of Pennsylvania, Philadelphia.

All users of SPM and metrology instruments are invited to Philadelphia next month to meet and discuss their work informally with colleagues and nanoscience pioneers from around the world!

The three-day, event-filled "Seeing at the Nanoscale" international conference includes two-and-a-half days of technical presentations, an evening poster session, and a reception at the National Constitution Center.

Technical Program:
� Nanomechanical and Local Property Measurements
� Visualization I: Biomolecules and Biological Processes
� Visualization II: Materials and Polymer Systems
� Measurements of Electrical, Optical, Magnetic, and Thermal Properties of Materials at the Nanoscale
� Instrumentation: New Tools and Techniques for Nanoscience

For more information and to register online, please go to: www.veeco.com/nanoconference

____________________________
Marlene Carlyle
Veeco Instruments
Conference Coordinator
112 Robin Hill Road
Santa Barbara, CA 93117
Tel: 805-967-1400 (ext. 2312)
Fax: 805-967-7717
Email: mcarlyle-at-veeco.com
____________________________

==============================Original Headers==============================
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Yes, this has been a recurring issue. The archives
are a good source for reflection.

Also, one should scan as a transparency rather than as a
negative. Then invert in PS or whatever image app
you use. Maximum D value is highly desirable and this
is engaged with transparencies.

gary g.

At 10:54 AM 6/8/2006, you wrote:

} ----------------------------------------------------------------------------
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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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Remember this posting is most likely not from a Subscriber, so when replying
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Email: fisher.phyllis-at-mayo.edu
Name: Phyllis Fisher

Organization: Mayo Foundation, Rochester, MN USA

Title-Subject: [Filtered] cathode luminescence

Question: I am considering adding a cathode luminescence tip to an old Centaurus backscatter detector on an Hitachi 4700 FESEM. Does anyone have experience with this tip and/or suggestions or comments regarding such an installation?

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

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Dear Group,
I am looking for someone who is going to participate the meeting on
Microscopy and Microanalysis 2006 in Chicago and wants to share a room
(hotel, hostel or guest house). Please inform me offline!

Grzegorz Tylko
Department of Cytology and Histology
Institute of Zoology
Jagiellonian Univeristy
Ingardena 6, 30-060 Krakow
Poland
tel.: +48 12 663 24 25
fax: +48 12 634 49 51
mobile: +48 698 55 55 85
e-mail: tylko-at-zuk.iz.uj.edu.pl

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Hi Richard,

The only problem with archived scanner info is that it goes out of date the
minute a new scanner hits the streets. I bought a �250 Canon 9550F at home
[Xmas 2005] under the mistaken view that consumer flatbeds can't get any
better than this for the price - I was very wrong (and annoyingly it was my
money). Going back 5 years a scanner that produced images like the 9950F
would have cost �4,000.

And I remember paying �5,000 for an office 386 computer with 8Mb of RAM and
a 120Mb hard drive back in 1989 - I wouldn't recommend it as particularly
good value these days. It did allow me to ditch the expensive IBM central
mainframe and run Fortran FTN77 independently on my PC though.

Keith

----------------------------------------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology Division
Institute of Ophthalmology
University College London
11-43 Bath Street
London EC1V 9EL

Tel: 020 7608 4050
Fax: 020 7608 4034
email: keith.morris-at-ucl.ac.uk

-----Original Message-----
X-from: rjharris-at-uwo.ca [mailto:rjharris-at-uwo.ca]
Sent: 08 June 2006 18:57
To: keith.morris-at-ucl.ac.uk

Shrunali
Could I suggest you check out the Microscopy List Server Archive? There has
been extensive and intelligent discussion about this subject quite
frequently over the last couple of years

Richard Harris
Laboratory Supervisor,
Imaging and Analysis
Department of Biology,
University of Western Ontario,
London Ontario, CANADA.
N6A 5B7
Ph. 519-661-2111 ext. 86780
Fax 519-661-3935

-----Original Message-----
X-from: aarti_harle-at-yahoo.co.in [mailto:aarti_harle-at-yahoo.co.in]
Sent: June 8, 2006 7:48 AM
To: rjharris-at-uwo.ca

Hello,

Can anyone suggest good scanner to scan TEM negatives and prints and and
latest Imange Analysis software.

Regards
Shrunali Kulkarni
Scientist
IMTech

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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Hi all,

I have been attempting to prepare cross sectional TEM samples of anodic tantala
(Ta2O5) by mechanical polishing. The substrate is tantalum metal with an
approximate thickness of 200um. The oxide film is grown anodically on the Ta
using electrochemistry.

In the literature, I have seen cross sectional TEM images of anodic tantala
grown on sputter deposited tantalum; an aluminum or silicon substrate is
typically used. I am curious if anyone has had success in preparing anodic
Ta2O5 samples on a tantalum substrate. (not a sputtered tantalum layer)

My main problem seems to be with de-lamination of the cross section sandwich.
Here is my attempted procedure:

I first bound two faces of my sample together and placed the sandwich in a press
while the epoxy cured (I have tried M-Bond and G1 from Gatan), but the sandwich
frequently breaks apart when I begin polishing. To address this problem, I
encased my sandwich in a brass tube and fitted the tube with supporting brass
shims, all held in place with epoxy. Using this technique, I was able to thin
some parts of the specimen to 20-50 um, but the epoxy was no longer present at
this point. When I attempted to ion-mill, the sandwiches splintered apart, and
any of the oxide that may have been surviving was obliterated in the ion mill.

Any suggestions or advice would be greatly appreciated.

Thank You,

Jennifer Ray Sloppy
Materials Science & Engineering Department
Ph.D. Candidate
Pennsylvania State University

==============================Original Headers==============================
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Jennifer Ray Sloppy wrote:
====================================================
I have been attempting to prepare cross sectional TEM samples of anodic tantala
(Ta2O5) by mechanical polishing. The substrate is tantalum metal with an
approximate thickness of 200um. The oxide film is grown anodically on the Ta
using electrochemistry.

In the literature, I have seen cross sectional TEM images of anodic tantala
grown on sputter deposited tantalum; an aluminum or silicon substrate is
typically used. I am curious if anyone has had success in preparing anodic
Ta2O5 samples on a tantalum substrate. (not a sputtered tantalum layer)

My main problem seems to be with de-lamination of the cross section sandwich.
Here is my attempted procedure:

I first bound two faces of my sample together and placed the sandwich in a press
while the epoxy cured (I have tried M-Bond and G1 from Gatan), but the sandwich
frequently breaks apart when I begin polishing. To address this problem, I
encased my sandwich in a brass tube and fitted the tube with supporting brass
shims, all held in place with epoxy. Using this technique, I was able to thin
some parts of the specimen to 20-50 um, but the epoxy was no longer present at
this point. When I attempted to ion-mill, the sandwiches splintered apart, and
any of the oxide that may have been surviving was obliterated in the ion mill.

Any suggestions or advice would be greatly appreciated.
============================================================
Some number of years ago, when our firm was first getting active in surface analysis and the need for calibrating etch rates became apparent, and the "standard" at the time for calibrating etching rates was to etch through an anodic film grown on a thin tantalum foil of known thickness, we found this could be cross-sectioned by diamond knife ultramicrotomy. Other than possessing a lot of experience with the ultramicrotomy of "difficult and hard" samples, we vacuum embedded the sample with our own SPI-Pon� 812 epoxy embedding resin, and did the thin sectioning with a 45 deg angle SPI Supplies� Brand Materials Science Diamond Knife but with a fairly ordinary (e.g. not very new) ultramicrotome. Separation of the anodized layer from the embedding resin could be minimized by using a vacuum impregnation. We were not always successful in keeping the anodized layer/substrate tantalum interface intact. We could get less separation by using a 35 deg knife but this wore out the knife very quickly and since we did not mind some separation, we used mainly a 45 deg. knife.

But one could get good TEM views of the ultramicrotome-produced cross-section for making (anodized layer) thickness measurements and high resolution images showing the morphology and fine structure of the anodized layer.

Disclaimer: SPI Supplies offers both the above-mentioned embedding resin and diamond knife and we also offer, through Structure Probe, Inc. this kind of a laboratory service for others.

Chuck

==================================================
Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400
President
1-(800)-2424-SPI
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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I have a colleague who wants to do in situ hybridizations on sponge
larvae (yes, sponges are animals and they have larval forms!). The
larvae are less than a millimeter long and quite delicate.

She tried fixing some in 4% paraformaldehyde in sea water, and the
larvae immediately exploded. She also tried Carnoy's fix, and 4% PFA
in Millonig's
phosphate buffer, and the same thing happened.

The only thing I could think of to suggest was dropping to 1 or 2%
PFA in sea water. I've seen references to using Bouins for light
microscopy or osmium tetroxide for TEM, but I don't think either of
these approaches would be good for in situs.

Any other suggestions?

Gary P. Radice
Department of Biology 804-289-8107 (voice)
University of Richmond 804-289-8233 (FAX)
Richmond VA 23173

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Jennifer,

There is a trick that I pulled with MoS2 films on Si in which I had
similar problems as you are having. What I did was to first deposit the
film through a gridded mask (I think that it was 400 mesh, but can't
remember for sure.) Then I prepared the samples as normal. What happens
is that you have enough Si-epoxy-Si areas when the samples are put
together to hold the sample stack together. It worked quite well. For
you, you will have to make a mask to prevent the anodic Ta2O5 from
forming and then remove your mask prior to making your cross section.

I also have another solution that might work for you. These are
Stainless steel rods that were electrospark cut with slits in them. You
must precisely cut the width of two strips of material and remove the
burrs and put them together between the rods. This gives you more
support with less glue interfaces. If you send me your address, I can
send you some 3mm tubes and some rods to try out.

The third method that might work for you is the Technoorg-Linda method.
This is illustrated in the Arpad Barna article: Barna, A. (1992)
"Topographic kinetics and practice of low-angle ion beam thinning",
Specimen Preparation for TEM of Materials. III, Mat. Res. Soc. Symp.
Proc. Vol. 254. R. Anderson, B. Tracy, and J. Bravman, eds. pp. 3�22.
You can also check the following articles: "Preparation of
Cross-Sectional TEM Samples for Low-Angle Ion Milling", J.P. MCCAFFREY
AND A. BARNA, MICROSCOPY RESEARCH AND TECHNIQUE 36:362�367 (1997). I
can also send you a copy of a video that shows how to do it. Basically,
two pieces of the sample are cut down to precise sizes and put in a
special titanium grid face-to-face. Then the grid is manipulated with
tools to clamp the samples in place tightly. The assembly is then put
into an epoxy. You can use M-bond 610, but T-L recommends a special
Araldit mixture with carbon that avoids the differential sputter removal
problem with non-filled epoxies. The sample is then ground down with
600 grit paper to a thickness of about 30-50 �m. You polish with 1 �m
paste to finish. It is then ion milled at a high voltage and a low
angle, say about 5 to 7 degrees with full rotation for say about an hour
depending on the sputter rate. This will give a dimple in the epoxy
joint. Then go to a very low angle, ~2 degrees, on one side of the
sample and mill until the topography is pushed to one side of the
sample. Not all commercially available ion mills are capable of going
to such a low angle. You must use a holder that is capable of it. Some
mills have holders that are more massive that allow the heat to be
dissipated more readily. Use the holder that is made to be milled from
one side and at very low angles. You will see the topography of the
sample pushed to the opposite side of the sample from where the gun is.
Turn the sample over and mill in such a way that the ion direction would
appear to oppose the direction from the first side milling. When you
are done, the topography will again be pushed away from the ion gun and
it will be on the opposite side of the sample of the glue joint than
where it is on the first side. Finish the sample using the same angles,
but low energy. Unless you have a special low energy gun such as the
one available from Technoorg-Linda, most commercially available guns are
limited to about 3 kV because they do not efficiently ionize the gas and
the beam spread. The low energy guns available from T-L will go as low
as 100 eV.

If you have any questions, please give me a call.

-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: jds451-at-psu.edu [mailto:jds451-at-psu.edu]
Sent: Friday, June 09, 2006 9:49 AM
To: Walck-at-SouthBayTech.com

Hi all,

I have been attempting to prepare cross sectional TEM samples of anodic
tantala
(Ta2O5) by mechanical polishing. The substrate is tantalum metal with
an approximate thickness of 200um. The oxide film is grown anodically
on the Ta using electrochemistry.

In the literature, I have seen cross sectional TEM images of anodic
tantala grown on sputter deposited tantalum; an aluminum or silicon
substrate is typically used. I am curious if anyone has had success in
preparing anodic Ta2O5 samples on a tantalum substrate. (not a sputtered
tantalum layer)

My main problem seems to be with de-lamination of the cross section
sandwich. Here is my attempted procedure:

I first bound two faces of my sample together and placed the sandwich in
a press while the epoxy cured (I have tried M-Bond and G1 from Gatan),
but the sandwich frequently breaks apart when I begin polishing. To
address this problem, I encased my sandwich in a brass tube and fitted
the tube with supporting brass shims, all held in place with epoxy.
Using this technique, I was able to thin some parts of the specimen to
20-50 um, but the epoxy was no longer present at this point. When I
attempted to ion-mill, the sandwiches splintered apart, and any of the
oxide that may have been surviving was obliterated in the ion mill.

Any suggestions or advice would be greatly appreciated.

Thank You,

Jennifer Ray Sloppy
Materials Science & Engineering Department
Ph.D. Candidate
Pennsylvania State University

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Hi

For cross sectional samples of metal/metal oxide, a rocking motion in ion mill (or single
section mode in PIPS) serves best. Align the glue line perpendicular with the ion beam,
allowing the beam ONLY bombarding from the backside of the film. With such arrangement, the
substrate is acting as a shield for the oxide film, protecting it from being overly milled.

Chengyu Song
National Center for Electron Microscopy
Lawrence Berkeley National Laboratory

jds451-at-psu.edu wrote:

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} Hi all,
}
} I have been attempting to prepare cross sectional TEM samples of anodic tantala
} (Ta2O5) by mechanical polishing. The substrate is tantalum metal with an
} approximate thickness of 200um. The oxide film is grown anodically on the Ta
} using electrochemistry.
}
} In the literature, I have seen cross sectional TEM images of anodic tantala
} grown on sputter deposited tantalum; an aluminum or silicon substrate is
} typically used. I am curious if anyone has had success in preparing anodic
} Ta2O5 samples on a tantalum substrate. (not a sputtered tantalum layer)
}
} My main problem seems to be with de-lamination of the cross section sandwich.
} Here is my attempted procedure:
}
} I first bound two faces of my sample together and placed the sandwich in a press
} while the epoxy cured (I have tried M-Bond and G1 from Gatan), but the sandwich
} frequently breaks apart when I begin polishing. To address this problem, I
} encased my sandwich in a brass tube and fitted the tube with supporting brass
} shims, all held in place with epoxy. Using this technique, I was able to thin
} some parts of the specimen to 20-50 um, but the epoxy was no longer present at
} this point. When I attempted to ion-mill, the sandwiches splintered apart, and
} any of the oxide that may have been surviving was obliterated in the ion mill.
}
} Any suggestions or advice would be greatly appreciated.
}
} Thank You,
}
} Jennifer Ray Sloppy
} Materials Science & Engineering Department
} Ph.D. Candidate
} Pennsylvania State University
}
}
} ==============================Original Headers==============================
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Email: fisher.phyllis-at-mayo.edu
Name: Phyllis Fisher

Organization: Mayo Foundation, Rochester, MN USA

Title-Subject: [Filtered] SEM - cathode luminescence detector

Question: I am considering adding a cathode luminescence tip to an old Centaurus backscatter detector on an Hitachi 4700 FESEM. Do you have any suggestions or comments on such a move?

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Email: warner.rr-at-pg.com
Name: Ron Warner

Organization: Procter & Gamble Co.

Title-Subject: [Filtered] Reichert CS Auto, free to a good home

Question: We have the ancient Reichert CS Auto freeze substitution unit available to a good home. It was working when we last used it, about 3 years ago. The body shows some of its years (so does mine), but the internals should be fine. You pay the shipping.

Ron

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Email: aetmicro-at-optonline.net
Name: Andrew Thelian

Organization: Nanoprobes, Inc.

Title-Subject: [Filtered] Philips EM 300 camera chamber issues

Question: Hi All :),

I had an issues with the plate chambers on my Philips EM 300... The venting valve jammed... So I naturally took it upon myself to attempt a repair... the overall repair seemed to have worked, the valve knob no longer jams or sticks like it had...

my problem now is...

when turning the knob(clockwise) to 10/4 o clock... the vacuum pump initiates like it is suppose to... i then press the knob in creating a seal(i wait on the vacuum light sequence)... then the last step is to finish the circuit and leave the knob at its resting position at 9/3 o clock... the rough pump shuts off as i attempt to complete the circuit...

does anyone know how to troubleshoot this issue?

Thanks in advance :)

Andrew

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Dear Gary,

I really doubt that PFA in itseld could make a
structure explode. Your colleague should look at the
osmoticity of the medium used for the fixation.
I don't know sponges very well, but there is a chance
that the intracellular osmoticity is regulated and is
not the same as the extracellular medium (sea water).
If the fixative stops the ion pumps present in the
membrane, it may be possible that the high salt
concentration of sea water provokes an osmotic shock
(even though in this case one should expect a
shrinkage of the cell, but you never know).
If the intracellular osmotocity of sponge cells is
unknown, he'd better try fixation media with different
osmotic forces.

Stephane

--- gradice-at-richmond.edu wrote:

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}
} I have a colleague who wants to do in situ
} hybridizations on sponge
} larvae (yes, sponges are animals and they have
} larval forms!). The
} larvae are less than a millimeter long and quite
} delicate.
}
}
} She tried fixing some in 4% paraformaldehyde in sea
} water, and the
} larvae immediately exploded. She also tried Carnoy's
} fix, and 4% PFA
} in Millonig's
} phosphate buffer, and the same thing happened.
}
} The only thing I could think of to suggest was
} dropping to 1 or 2%
} PFA in sea water. I've seen references to using
} Bouins for light
} microscopy or osmium tetroxide for TEM, but I don't
} think either of
} these approaches would be good for in situs.
}
}
} Any other suggestions?
}
}
} Gary P. Radice
} Department of Biology 804-289-8107 (voice)
} University of Richmond 804-289-8233 (FAX)
} Richmond VA 23173
}
}
} ==============================Original
} Headers==============================
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} 13:48:42 2006
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} hybridization
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==============================Original Headers==============================
8, 20 -- From nizets2-at-yahoo.com Mon Jun 12 02:48:31 2006
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Fluid mounting 'film' scanning,

I notice that the Epson V750 Pro flatbed scanner now has an optional fluid
mounting accessory that involves a bottom sealed lift out tray (that
'prevents' fluid seepage into the scanner and is 'easy clean'). See it at:

http://www.photo-i.co.uk/Reviews/interactive/Epson%20V750/page_7.htm

Professional drum scanners use mounting fluid to improve film scan quality
if you really need the best possible scan, and have a private income (most
of us don't). If you are flatbed scanning odd specimens or really want to
use oil or mounting fluid, this accessory looks very useful. I don't think
the kit's generally available yet though.

Keith

----------------------------------------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology Division
Institute of Ophthalmology
University College London
11-43 Bath Street
London EC1V 9EL

Tel: 020 7608 4050
Fax: 020 7608 4034
email: keith.morris-at-ucl.ac.uk

-----Original Message-----
X-from: mey-at-amnh.org [mailto:mey-at-amnh.org]
Sent: 25 May 2006 20:47
To: keith.morris-at-ucl.ac.uk

Hi Mike,
I have been exploring and having success using a flatbed scanner for
particle analysis, but
actually started out in this endeavor by scanning rocks and rock cores.
We now use an Epson XL 1640 scanner, which has a focus function,
essentially allowing us to get right on the surface of the samples.
However, a pretty flat cut is needed to get overall excellent quality. I
found that this works much better than greasing up the bed with oils,
glycerin or other sticky stuff. We used a thin plastic sheet (same
material as thin clear view-foils) to protect the glass from the rocks.
As a side note, I too saw some strange artifacts with oils, which could
(?) be due to "internal" reflections or interference patterns between
the glass-plastic or micro-bubbles on or close the sample surface.
My major concern was contaminating the samples with oils, especially
when doing trace element work or isotope analysis for age dating
purposes of the rock cores. Not that the oil itself would be the major
culprit, but more the rock dust flying around everywhere in the cutting lab
sticking to the greasy surfaces. Rock cores sometimes have oil on them
from the
drilling process and is usually washed off with sulfo. However, the
porosity of the sample may
contribute to deeper contamination, especially is if the sample shows
cracks or micro-fractures. I had to consider this for archival purposes
and potential future analysis on the same samples as new technologies
emerge.

All the best

Jacob

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

Jacob Louis Mey
Microscopy and Imaging Facility
American Museum of Natural History
Central Park west at 79th Street
New York, NY 10024

Office: 1-212-313-7975
fax: 1-212-496-3480

http://research.amnh.org/users/mey

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

michael-at-Shaffer.net wrote:
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} We have been using a flatbed scanner for documenting rock core before
} processing it for other analytical techniques. Now, we want to explore
the
} technique a bit further, possibly towards applied image analysis, but need
} one last ingredient for better quality scans. We want to use some akin to
} immersion oil for the interface between rock and flatbed, but want to find
} something inexpensive, non-toxic and easy to clean. Any thoughts?
}
} TIA & genuinely :o)
} michael shaffer
}
} SEM/MLA Research Coordinator
} (709) 737-6799 (ofc)
} (709) 737-6790 (lab)
} (709) 737-6193 (FAX)
} {http://www.mun.ca/creait/maf/}
} {http://www.esd.mun.ca/epma/}
}
} Inco Innovation Centre
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--
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

Jacob Louis Mey
Microscopy and Imaging Facility
American Museum of Natural History
Central Park west at 79th Street
New York, NY 10024

Office: 1-212-313-7975
fax: 1-212-496-3480

http://research.amnh.org/users/mey

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

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Greetings listers. We have an image analysis problem, and I could use your
help. We are frequently asked to characterize the cell size and structure
of various polymer foams for our customers. To date we have relied upon an
ASTM method that involves counting the number of cell intersections with a
reference line drawn on the image to compute average cell size. The method
assumes a degree of homogeneity and cell density that is often an imperfect
representation of our customer's samples, and in any event it would be far
more desirable to select and measure the cells directly with our
ImageProPlus software. There's the rub. We're having the devil of a time
getting good segmentation of the open cells against the background matrix.
We've tried a number of filters and operations, but thus far without
success. I have posted two examples through the link listed below (I
disabled it to get through the spam filter). To download, just right click
on the file name and "save target as".

**http://www.impactanalytical.com/data/**

If any of you experts out there can set us on the right track, or
alternatively explain why we're wasting our time, I would GREATLY appreciate
it. My head hurts and the walls are in need of repair!

Incidentally, I have tried varying kV, and collecting in backscatter as well
as backscatter plus/minus for topography. Thus far these efforts have not
solved the fundamental problem that contrast along the rim of a given cell
shifts from light to dark, confounding segmentation. Thanks for your
consideration - there's a windy city beer in the bargain if my travel
request goes through!
Yours,
Matt
Matthew Stephenson
Impact Analytical/MMI
1910 W. Saint Andrews Rd.
Midland, MI 48640
(989)832-5555 X506

==============================Original Headers==============================
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Your images are nearly identical to one that I use as a lab experiment
in the image analysis courses I teach.

If you examine the image visually, how do you "see" the edges of the
pores? There is always some local brightness change associated with the
edge. A good way to highlight these edges is to duplicate the image,
and perform a morphological erosion to replace each pixel with its
darkest neighbor. Then subtract that image from the original, which
will leave the locallly lightest pixels. These highlight the edges. Of
course, you will want to do a bit of cleanup on the images first (a
median filter or one of its derivatives is probably better than a
smoothing convolution).

Your major problem is not ** measuring ** the circular features in your
image, it is relating those measurements to the pores. This is not an
imaging problem but a stereological one. The sections through the pores
are NOT the pore sizes. There is a lot of literature on the
relationships between the sizes of 2D sections through features and the
sizes of the 3D structures.

John Russ

-----Original Message-----
X-from: stephenson-at-impactanalytical.com
To: DrJohnRuss-at-AOL.com
Sent: Mon, 12 Jun 2006 07:20:35 -0500

Greetings listers. We have an image analysis problem, and I could use
your
help. We are frequently asked to characterize the cell size and
structure
of various polymer foams for our customers. To date we have relied
upon an
ASTM method that involves counting the number of cell intersections
with a
reference line drawn on the image to compute average cell size. The
method
assumes a degree of homogeneity and cell density that is often an
imperfect
representation of our customer's samples, and in any event it would be
far
more desirable to select and measure the cells directly with our
ImageProPlus software. There's the rub. We're having the devil of a
time
getting good segmentation of the open cells against the background
matrix.
We've tried a number of filters and operations, but thus far without
success. I have posted two examples through the link listed below (I
disabled it to get through the spam filter). To download, just right
click
on the file name and "save target as".

**http://www.impactanalytical.com/data/**

If any of you experts out there can set us on the right track, or
alternatively explain why we're wasting our time, I would GREATLY
appreciate
it. My head hurts and the walls are in need of repair!

Incidentally, I have tried varying kV, and collecting in backscatter as
well
as backscatter plus/minus for topography. Thus far these efforts have
not
solved the fundamental problem that contrast along the rim of a given
cell
shifts from light to dark, confounding segmentation. Thanks for your
consideration - there's a windy city beer in the bargain if my travel
request goes through!
Yours,
Matt
Matthew Stephenson
Impact Analytical/MMI
1910 W. Saint Andrews Rd.
Midland, MI 48640
(989)832-5555 X506

==============================Original
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________________________________________________________________________
Check out AOL.com today. Breaking news, video search, pictures, email
and IM. All on demand. Always Free.

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Hi Jennifer-

There have been several good suggestions already, so I will try to add a
few hints that have worked for me. First of all, do make sure you are not

milling in the same direction as the epoxy line, as that will
preferentially mill the epoxy away. Use beam modulation or the equivalent

to restrict the beam to milling perpendicular to the interface. Also,
I've noticed the thinner the epoxy line, the better the milling. When
gluing your two sample pieces together, make sure the surfaces are
absolutely clean, check the epoxy (I always use G-1) for specks of
'stuff', and use the strongest clamp/vise you can. This will protect the
epoxy line as well. Finally, when you do have a hard-to-glue surface,
sometimes this can be improved by coating it with a thin layer of a
material that will adhere to the G-1, such as chromium. You didn't
mention dimpling in your sample prep, but this might also help. In my
experience dimpling is less preferential than ion milling, and can help
you get a thinner sample that requires less milling.

Leslie

Leslie Krupp (Thompson)
IBM Almaden Research
650 Harry Road, K19/D1
San Jose, CA 95120-6099
(408) 927-3856

jds451-at-psu.edu
06/09/2006 09:49 AM
Please respond to
jds451-at-psu.edu

To
Leslie E Krupp/Almaden/IBM-at-IBMUS
cc

Subject
[Microscopy] Tantalum / Ta Oxide Cross Section TEM Sample Prep

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Hi all,

I have been attempting to prepare cross sectional TEM samples of anodic
tantala
(Ta2O5) by mechanical polishing. The substrate is tantalum metal with an
approximate thickness of 200um. The oxide film is grown anodically on the

Ta
using electrochemistry.

In the literature, I have seen cross sectional TEM images of anodic
tantala
grown on sputter deposited tantalum; an aluminum or silicon substrate is
typically used. I am curious if anyone has had success in preparing
anodic
Ta2O5 samples on a tantalum substrate. (not a sputtered tantalum layer)

My main problem seems to be with de-lamination of the cross section
sandwich.
Here is my attempted procedure:

I first bound two faces of my sample together and placed the sandwich in a

press
while the epoxy cured (I have tried M-Bond and G1 from Gatan), but the
sandwich
frequently breaks apart when I begin polishing. To address this problem,
I
encased my sandwich in a brass tube and fitted the tube with supporting
brass
shims, all held in place with epoxy. Using this technique, I was able to
thin
some parts of the specimen to 20-50 um, but the epoxy was no longer
present at
this point. When I attempted to ion-mill, the sandwiches splintered
apart, and
any of the oxide that may have been surviving was obliterated in the ion
mill.

Any suggestions or advice would be greatly appreciated.

Thank You,

Jennifer Ray Sloppy
Materials Science & Engineering Department
Ph.D. Candidate
Pennsylvania State University

==============================Original
Headers==============================
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9, 17 -- To: microscopy-at-microscopy.com
9, 17 -- Subject: Tantalum / Ta Oxide Cross Section TEM Sample Prep
9, 17 -- From: "Jen Sloppy" |--jds451-at-psu.edu--|
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|--br--||--font size=2 face="sans-serif"--|Hi Jennifer-|--/font--|
|--br--|
|--br--||--font size=2 face="sans-serif"--|There have been several good
suggestions
already, so I will try to add a few hints that have worked for me.
First
of all, do make sure you are not milling in the same direction as the
epoxy
line, as that will preferentially mill the epoxy away. Use beam
modulation
or the equivalent to restrict the beam to milling perpendicular to the
interface. Also, I've noticed the thinner the epoxy line, the better
the milling. When gluing your two sample pieces together, make sure
the surfaces are absolutely clean, check the epoxy (I always use G-1) for
specks of 'stuff', and use the strongest clamp/vise you can. This
will protect the epoxy line as well. Finally, when you do have a
hard-to-glue surface, sometimes this can be improved by coating it with
a thin layer of a material that will adhere to the G-1, such as chromium.
You didn't mention dimpling in your sample prep, but this might also
help. In my experience dimpling is less preferential than ion
milling,
and can help you get a thinner sample that requires less
milling.|--/font--|
|--br--|
|--br--||--font size=2 face="sans-serif"--|Leslie|--/font--|
|--br--|
|--br--||--font size=2 face="sans-serif"--|Leslie Krupp (Thompson)|--br--|
IBM Almaden Research|--br--|
650 Harry Road, K19/D1|--br--|
San Jose, CA 95120-6099|--br--|
(408) 927-3856|--/font--|
|--br--|
|--br--|
|--br--|
|--table width=100%--|
|--tr valign=top--|
|--td width=40%--||--font size=1
face="sans-serif"--||--b--|jds451-at-psu.edu|--/b--| |--/font--|
|--p--||--font size=1 face="sans-serif"--|06/09/2006 09:49 AM|--/font--|
|--table border--|
|--tr valign=top--|
|--td bgcolor=white--|
|--div align=center--||--font size=1 face="sans-serif"--|Please respond
to|--br--|
jds451-at-psu.edu|--/font--||--/div--||--/table--|
|--br--|
|--td width=59%--|
|--table width=100%--|
|--tr valign=top--|
|--td--|
|--div align=right--||--font size=1
face="sans-serif"--|To|--/font--||--/div--|
|--td--||--font size=1 face="sans-serif"--|Leslie E
Krupp/Almaden/IBM-at-IBMUS|--/font--|
|--tr valign=top--|
|--td--|
|--div align=right--||--font size=1
face="sans-serif"--|cc|--/font--||--/div--|
|--td--|
|--tr valign=top--|
|--td--|
|--div align=right--||--font size=1
face="sans-serif"--|Subject|--/font--||--/div--|
|--td--||--font size=1 face="sans-serif"--|[Microscopy] Tantalum / Ta
Oxide Cross
Section TEM Sample Prep|--/font--||--/table--|
|--br--|
|--table--|
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|--br--|
Hi all,|--br--|
|--br--|
I have been attempting to prepare cross sectional TEM samples of anodic
tantala|--br--|
(Ta2O5) by mechanical polishing. The substrate is tantalum metal
with an|--br--|
approximate thickness of 200um. The oxide film is grown anodically
on the Ta|--br--|
using electrochemistry. |--br--|
|--br--|
In the literature, I have seen cross sectional TEM images of anodic
tantala|--br--|
grown on sputter deposited tantalum; an aluminum or silicon substrate
is|--br--|
typically used. I am curious if anyone has had success in preparing
anodic|--br--|
Ta2O5 samples on a tantalum substrate. (not a sputtered tantalum
layer)|--br--|
|--br--|
My main problem seems to be with de-lamination of the cross section
sandwich.|--br--|
Here is my attempted procedure: |--br--|
|--br--|
I first bound two faces of my sample together and placed the sandwich in
a press|--br--|
while the epoxy cured (I have tried M-Bond and G1 from Gatan), but the
sandwich|--br--|
frequently breaks apart when I begin polishing. To address this
problem,
I|--br--|
encased my sandwich in a brass tube and fitted the tube with supporting
brass|--br--|
shims, all held in place with epoxy. Using this technique, I was able to
thin|--br--|
some parts of the specimen to 20-50 um, but the epoxy was no longer
present
at|--br--|
this point. When I attempted to ion-mill, the sandwiches splintered
apart, and|--br--|
any of the oxide that may have been surviving was obliterated in the ion
mill.|--br--|
|--br--|
Any suggestions or advice would be greatly appreciated.|--br--|
|--br--|
Thank You,|--br--|
|--br--|
Jennifer Ray Sloppy|--br--|
Materials Science & Engineering Department|--br--|
Ph.D. Candidate|--br--|
Pennsylvania State University|--br--|
|--br--|
|--br--|
==============================Original
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9, 17 -- To: microscopy-at-microscopy.com|--br--|
9, 17 -- Subject: Tantalum / Ta Oxide Cross Section TEM Sample
Prep|--br--|
9, 17 -- From: "Jen Sloppy" <jds451-at-psu.edu>|--br--|
9, 17 -- X-Mailer: Penn State WebMail|--br--|
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Hello listers-

Many thanks to all the folks who responded to my post. Here is a summary
of the suggestions.

At least 4 or 5 people suggested sonicating the powders in a solvent prior
to disbursing. After this step, some suggested putting a drop on a filmed
grid, or using some type of atomizer to spray the dispersion onto a grid.
(Perfume sprayer, air can held in a jar, nebulizer)

A couple people recommended simply mixing the powder with a solvent
without sonicating and either letting a drop dry on the grid, or heating
it to aid evaporation.

There were two mentions of dry loading the grid, which is dumping some of
the powder on the grid and tapping off the excess. This is the method I
tried first as it was the easiest, but I have not looked at the grid yet
to see if there is enough material to do EELS on.

Thanks again,
Leslie

Leslie Krupp (Thompson)
IBM Almaden Research
650 Harry Road, K19/D1
San Jose, CA 95120-6099
(408) 927-3856

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Great suggestions, John, I will give them a try. And point well taken
regarding the indirect relation of the 2D feature to the 3D pore size. The
ASTM method we use (#3576) makes that dimensional correction, but only by
assuming close-packed, spherical features. That said, our customers have
real use for comparative data from their less ideal samples. If we can
select their 2D open cells, they can compare one region to another as a
function of molding conditions, one sample to another as a function of
processing or R&D variations, or see how they stack up with the competition.
But if I can get your suggestions to work, we can offer them stereological
analysis as well. Just have to see if they're willing to pay...Thanks
again!
Yours,
Matt

-----Original Message-----
X-from: drjohnruss-at-aol.com [mailto:drjohnruss-at-aol.com]
Sent: Monday, June 12, 2006 9:10 AM
To: microscopy-at-sparc5.microscopy.com
Cc: stephenson-at-impactanalytical.com

Greetings listers. We have an image analysis problem, and I could use
your
help. We are frequently asked to characterize the cell size and
structure
of various polymer foams for our customers. To date we have relied
upon an
ASTM method that involves counting the number of cell intersections
with a
reference line drawn on the image to compute average cell size. The
method
assumes a degree of homogeneity and cell density that is often an
imperfect
representation of our customer's samples, and in any event it would be
far
more desirable to select and measure the cells directly with our
ImageProPlus software. There's the rub. We're having the devil of a
time
getting good segmentation of the open cells against the background
matrix.
We've tried a number of filters and operations, but thus far without
success. I have posted two examples through the link listed below (I
disabled it to get through the spam filter). To download, just right
click
on the file name and "save target as".

**http://www.impactanalytical.com/data/**

If any of you experts out there can set us on the right track, or
alternatively explain why we're wasting our time, I would GREATLY
appreciate
it. My head hurts and the walls are in need of repair!

Incidentally, I have tried varying kV, and collecting in backscatter as
well
as backscatter plus/minus for topography. Thus far these efforts have
not
solved the fundamental problem that contrast along the rim of a given
cell
shifts from light to dark, confounding segmentation. Thanks for your
consideration - there's a windy city beer in the bargain if my travel
request goes through! Yours, Matt Matthew Stephenson Impact Analytical/MMI
1910 W. Saint Andrews Rd. Midland, MI 48640 (989)832-5555 X506

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1, 14 -- From stephenson-at-impactanalytical.com Mon Jun 12 07:19:49 2006 1, 14
-- Received: from mitconexch.mitcon.org (mitconexch.mitcon.org
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Hi All,

I have a user who has presented a very tough problem.

They are looking at a polymer tube (ID ~ .1 - 1 micron) which is
filled with a liquid with CNT suspended in it. The tube is the
result of drawing from a larger tube and they are hoping to have
aligned CNT. They would like to verify this in some way.

I have access to a FESEM and a conventional TEM. The SEM is not
environmental. I could not come up with a way to image these. The
SEM will not be able to resolve the CNT in any case I think. The TEM
could but the sample would have to be thinned to the point that it is
likely the inner core of the tube is penetrated. I don't think
liquid in the TEM is viable.

I have suggested XRD as a possibility, but the CNT signal on top of
the tube and liquid signal may be problematic.

Any thoughts?

Thanks,

Tom

------------------------------------------------------------------------
---------------------------------------------
Thomas M Murray email:
murraytm-at-u.washington.edu
Electron Microscopy Center Manager Phone: (206)543-2836
Materials Science & Engineering Fax: (206)543-3100
Box 352120 302 Roberts Hall Cell: (425)345-0083
University of Washington
Seattle, WA 98195

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Any suggestions on the best way to image protein crystals? They are
20-100 um in length and very stable in high molarity salt solutions or PEG
solutions, but dissolve readily in less dilute aqueous solutions. We
would like to avoid chemical fixation.

Joe Neilly

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Greetings,
If the goal is not to image the tubes per se but to tell
whether they have a net alignment, polarized light microscopy could
work. I think sensitivity is ok (With a good instrument, you can
detect birefringent retardation of 1 nm and with a really good
instrument you can get down to almost 0.1 nm). I expect the problem
will come from the polymer tube. Its material could be birefringent
and if the inner diameter is really 0.1 micron (rather than 1 micron)
then that will probably be too small. But if your i.d.s are closer to
1 micron and the polymer tube itself behaves in the light, it could
work...

Good luck!
Tobias
}
}
} Hi All,
}
} I have a user who has presented a very tough problem.
}
} They are looking at a polymer tube (ID ~ .1 - 1 micron) which is
} filled with a liquid with CNT suspended in it. The tube is the
} result of drawing from a larger tube and they are hoping to have
} aligned CNT. They would like to verify this in some way.
}
} I have access to a FESEM and a conventional TEM. The SEM is not
} environmental. I could not come up with a way to image these. The
} SEM will not be able to resolve the CNT in any case I think. The TEM
} could but the sample would have to be thinned to the point that it is
} likely the inner core of the tube is penetrated. I don't think
} liquid in the TEM is viable.
}
} I have suggested XRD as a possibility, but the CNT signal on top of
} the tube and liquid signal may be problematic.
}
} Any thoughts?
}
} Thanks,
}
} Tom
}
} ------------------------------------------------------------------------
} ---------------------------------------------
} Thomas M Murray email:
} murraytm-at-u.washington.edu
} Electron Microscopy Center Manager Phone: (206)543-2836
} Materials Science & Engineering Fax: (206)543-3100
} Box 352120 302 Roberts Hall Cell: (425)345-0083
} University of Washington
} Seattle, WA 98195
}
}
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--
_ ____ __ ____
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/ / / / \ \ \ Biology Department
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On Jun 12, 2006, at 1:33 PM, joe.p.neilly-at-abbott.com wrote:

} Any suggestions on the best way to image protein crystals? They are
} 20-100 um in length and very stable in high molarity salt solutions or
} PEG
} solutions, but dissolve readily in less dilute aqueous solutions. We
} would like to avoid chemical fixation.
}
Dear Joe,
CryoTEM is a good method if the crystals are thin enough--10s of nm.
They would appear lighter against the darker buffer if the salt
concentration is high enough, but they might have very little contrast
in a PEG buffer. If the crystals are well-ordered, you should see
strong diffraction spots and strong spots in the FFT of an image.
Plunge freezing would be good for thin crystals.
Yours,
Bill Tivol, PhD
EM Scientist and Manager
Cryo-Electron 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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Why not try an ESEM in the near liquid state?

They sound more than large enough.

Nestor

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Tel: 630-252-7901, Fax: 630-252-4798
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The box said ...
"This program requires Win 95/98/NT or better..."
So I bought a Mac !

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (mhunt-at-flinnsci.com) from http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Friday, May 26, 2006 at 09:59:18
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Email: mhunt-at-flinnsci.com
Name: Maureen Hunt

Organization: Flinn

Education: 9-12th Grade High School

Location: Batavia, Illinois, USA

Question: I am a former materials microscopist and teacher who is trying to find a more exciting way to teach mitosis and meiosis. Currently, everyone uses prepared slides of whitefish blastula to teach animal mitosis and either prepared slides or unprepared slides of allium root tips to teach plant mitosis. Is there a protist or other "animal-like" eukaryote that can be given to the students, squashed and stained to show animal mitosis? I have googled and come up short. Any suggestions?
Thank-you for your time.

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7, 13 -- mitosis
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (sullivanmassi-at-comcast.net) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday, June 12, 2006 at 09:12:29
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Email: sullivanmassi-at-comcast.net
Name: Martha Sullivan

Organization: St. John's College High School

Education: 9-12th Grade High School

Location: Washington, DC 20015

Question: Do you have any recommendations on purchasing microscopes for high school biology? Is there a particular manufacturer that you would go with in terms of value, reliability, etc? Are there new technologies that would fall within a typical high school budget to consider?

Thanks in advance for any advice.
- Martha S.

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Since MItosis and Meiosis are both dynamic processes, I would
recommend that you look for movies of the processes.I did a Google
search on "mitosis video", and came up with about 300,000 hits. The
first few looked quite impressive.

I would also recommend that you teach the function rather than the
structure. That is, mitosis functions in normal organismal growth,
while meiosis has a significant role only in reproduction.

I can't resist one story. After a lecture that I gave on mitosis in
a non-majors Biology course, a student came up to me and said "You
have finally cleared up a major mystery for me. I've always wondered
how it is that the skin completely covers a little baby, and that it
still completely covers the adult that the baby becomes. Now I
understand"

Date sent: Mon, 12 Jun 2006 22:04:50 -0500
To: jbs-at-temple.edu
X-from: mhunt-at-flinnsci.com
Send reply to: mhunt-at-flinnsci.com

This is an extremely broad and unbounded question.
How much money are you willing to spend and what
analysis methods do you want (LM, DF, DIC, PH, POL, etc.)?

I would recommend querying the usenet group about your
question, as it comes up frequently.

sci.techniques.microscopy

Many Web apps have Usenet avenues. There is a large number of
informed and experienced folks who deal with LMs on a
daily basis. Many are quite helpful for non-professional
applications. This is good since the opposite can get
out of hand for some folks (i.e., too pricey).

gary g.

At 08:07 PM 6/12/2006, you wrote:

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12, 21 -- high school biology
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This Question was submitted to Ask-A-Microscopist by (Goinoutonalamb-at-yahoo.com)
from http://www.microscopy.org/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday, June 12, 2006 at 23:04:35
Remember to consider the Grade/Age of the student when considering the Question
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Email: Goinoutonalamb-at-yahoo.com
Name: Michael Lamb

Organization: Nassau Community College

Education: Undergraduate College

Location: Upton, New York

Title: Tomographic Reconstruction

Question: I am collaborating with a group on a 3D visualization project through Brookhaven National Labs and I have a question pertaining to 3D reconstruction by means of electron tomography. From the literature that I have read, I've gathered that it's recommended that a 200 kv beam or higher is used when working with sections of 120 nm or thicker. Unfortunately, the philips EM-300 that we are working with is only capable of producing a 100 kv beam. Because of the time constraints of our project (10 weeks) it will be very difficult to determine what section thicknesses/maximum tilt angles will work with 100kv with trial and error. Is there a formula, or a guideline of some sort to follow to determine how much mass a 100kv beam can pass with minimal loss of resolution? We are planning on doing a tilt series from -60 to +60 degrees, in 2 degree increments, at 120 nm sections (hopefully). If anyone could lend me some insight as to whether or not this is feasible, and how far we can go, I would greatly appreciate it!

Thanks!

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I'm sure there are several articles (also in books, like the one from
Joachim Frank on electron tomography) in which the problem of specimen
thickness vs. resolution has been discussed.

see e.g. the following article:

Biophys J, February 1998, p. 1031-1042, Vol. 74, No. 2

Electron Tomography of Ice-Embedded Prokaryotic Cells
Rudo Grimm,* Hapreet Singh,* Reinhard Rachel,# Dieter Typke,* Wolfram
Zillig,* and Wolfgang Baumeister*

best regards,
Reinhard Rachel
---------------------
PD Dr.Reinhard Rachel
Universit�t Regensburg
Zentrum f�r EM der NWF III am
Lehrstuhl fuer Anatomie
D-93053 Regensburg - Germany
fon +49 941 943 1666, 1720
fax +49 941 943 2868

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Hi,

Probably the most interesting way to teach mitosis is
to synchronize cells in culture. It is probably not
the easiest though ;-). It is probable that
unsynchronized but very actively growing cells will
show enough mitotic events to cover the whole cycle.
Most of the time they are easy to recognize by normal
phase constrast, although fluorescent staining like
DAPI are very fast to perform and very impressive to
observe.
I must admit that after working now for more than 10
years with cells in culture, seeing them split in late
telophase is still an exciting event. I witness events
which are actually occuring in my own body at that
very moment!!!

I agree that you should insist on the functional part
though. It is not easy to comprehend these phenomenon
in themself, but understand the difference between
mitosis and meiosis is really a hard task.

Good luck

Stephane

--- mhunt-at-flinnsci.com wrote:

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} Email: mhunt-at-flinnsci.com
} Name: Maureen Hunt
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} Education: 9-12th Grade High School
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} Location: Batavia, Illinois, USA
}
} Question: I am a former materials microscopist and
} teacher who is trying to find a more exciting way to
} teach mitosis and meiosis. Currently, everyone uses
} prepared slides of whitefish blastula to teach
} animal mitosis and either prepared slides or
} unprepared slides of allium root tips to teach plant
} mitosis. Is there a protist or other "animal-like"
} eukaryote that can be given to the students,
} squashed and stained to show animal mitosis? I have
} googled and come up short. Any suggestions?
} Thank-you for your time.
}
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10, 20 -- Subject: Re: [Microscopy] AskAMicroscopist: trying to find a more exciting way to teach
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Email: iztok.dogsa-at-ijs.si
Name: iztok dogsa

Organization: IJS

Title-Subject: [Filtered] confocal spinning disk microscope

Question: Has anyone experince with confocal spinning disk microscopy?

Which system would you suggest?

We'd like to achieve really high frame rates, but don't want to buy the laser. The arc lamp is Ok for us.

What about BD CARV II imaging system? Is it better than olympus DSU?

Thanks & regards to you all,

Iztok

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Hi Martha,

We get asked this fairly regularly by schools and parents (so we have stock
replies). Often professional microscopists aren't the best source of
information for cheap microscope's as all our optical systems are always
over �10,000, and most lab mainstays like our three confocal microscopes
come in at nearer �200,000 each (and users still complain about image
quality).

Cheap microscopes under �100 are always a disappointment and toy (often
called student) microscopes can be very poor. You can easily buy second-hand
via ebay, but again there are risks that the optics will be damaged or
simply very dirty and difficult to clean (look for old branded 'laboratory'
microscopes e.g. Bausch & Lomb) and besides you can't really fill a
classroom quickly that way, just get a good one for demos. Local labs may
have a good old microscope they no longer need and they may give it to
schools (or sell them cheaply) if you acknowledge them - do ask parents and
write begging letters if you any spare time (but don't expect anything -
look for old establishments that will have years of stuff accumulated). Also
try any local microscope enthusiast clubs (they may even be persuaded to
come in and talk/demo to pupils) and visit other schools for info and a
try-out.

There are suppliers geared up to providing microscopes for schools, so ask
around at other school's science departments, but expect to pay nearer �500
each for a quality setup (although with those like bottom end Meade
[www.meade.com] at around �100 you can see something at low mag (~20x
objective i.e. around 180x mag) with a quality stained section. For pond
life etc. a stereoscopic 'dissecting' microscope (40x to 120x mag) is ideal,
and of course you can get a really long way with a good magnifying glass
(not the really small hi-mag cheap lens ones, try before you buy) - I have a
few excellent ones at home for �1 and a good low mag Osram one that includes
an illuminating halogen bulb at �8.

Generally prepared slides can rapidly get a bit boring for under 14s, but
living or unusual things always attract an audience. Also try your flatbed
film scanner (not LiDe and from around �60 upwards) that will be good for
looking at soft static things: leaves, fruit, nuts, household objects (scan
at max resolution and try reflected and 'film' mode). In the UK there are
sites like http://www.wedgwood-group.com/microscopes_digital.htm that cater
for schools and colleges, providing standard compound and stereo microscopes
as well as cheap PC video based microscope solutions where the whole class
can look at a computer screen with some pushing and shoving. Best to try
them on 'approval' as many cheap microscopes can be disappointing if you
expect too much.

Excellent pre-prepared stained slides of plant stems and leaves or bits of
rats, insects etc.. can be bought, but they tend to be expensive and are
easily broken by any age-group. Mounted slides keep well though, so
'vintage' ones even from 50 years ago can still look OK - most schools will
have some specimen slides knocking about.

At home and our Primary school (under 11) we use the Digital Blue QX-5 (�70)
- it's fun but pretty useless for serious microscope work as it's so low res
(but at 640 x 480 better than the old Intel QX-3 it replaced). It puts the
image on a PC screen. See
http://micro.magnet.fsu.edu/optics/intelplay/index.html (not updated for the
QX-5 but all applies - the site even discusses ways of contrast enhancement
etc..). Once on the PC the 640x480 images can be manipulated and pasted etc,
and the QX-5 does time-lapse for things like crystal growth. living plants
growing and small animals. The similar but far better built Olympus MIC-D
was great but being over �500 it was just too expensive for most schools and
is now discontinued - there are other similar budget systems about though.

The macro on a good digital camera (like the image stabilised 5MP Canon S2IS
going cheap at �200 over here - http://www.dpreview.com/reviews/canons2is)
is also worth a try, particularly with a small tripod and halogen bendy desk
lamp if very close-up, but I'm not sure I'd like a class with 20 boys near
my Olympus E500 digital SLR system though. You can get quite reasonable
pictures by resting a small compact digital camera lens against the eyepiece
of a microscope.

By the way do try growing crystals on a slide, a few drops of a saturated
solution of salt (NaCl) or copper sulphate will grow superb crystals on the
surface of a slide when viewed under a microscope (but it takes a few hours
for the crystals to form and they often look best before the liquids all
gone). Just make sure they don't drive the objective tips into the solution.
It's not biology but its fun.

Regards

Keith

Try looking in Amazon.com for decent microscope books with lots of pictures
(and they have a good customer review system). Plus try web searches for
general sites like these (and for more specific topics) :

http://www.101science.com/Microscope.htm
http://micro.magnet.fsu.edu/
http://www.microscopy-uk.org.uk/index.html
http://www.btinternet.com/~stephen.durr/
http://www.mccroneatlas.com
http://www.diatoms.co.uk [for fun images made of diatoms]

----------------------------------------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology Division
Institute of Ophthalmology
University College London
11-43 Bath Street
London EC1V 9EL

Tel: 020 7608 4050
Fax: 020 7608 4034
email: keith.morris-at-ucl.ac.uk

-----Original Message-----
X-from: gary-at-gaugler.com [mailto:gary-at-gaugler.com]
Sent: 13 June 2006 05:12
To: keith.morris-at-ucl.ac.uk

This is an extremely broad and unbounded question.
How much money are you willing to spend and what
analysis methods do you want (LM, DF, DIC, PH, POL, etc.)?

I would recommend querying the usenet group about your
question, as it comes up frequently.

sci.techniques.microscopy

Many Web apps have Usenet avenues. There is a large number of
informed and experienced folks who deal with LMs on a
daily basis. Many are quite helpful for non-professional
applications. This is good since the opposite can get
out of hand for some folks (i.e., too pricey).

gary g.

At 08:07 PM 6/12/2006, you wrote:

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for
12, 21 -- high school biology
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39, 27 -- From keith.morris-at-ucl.ac.uk Tue Jun 13 08:34:16 2006
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On topic: The May issue of Microscopy Today contained the following joke:

"Scientifically, maybe body cells -do- replace themselves completely in
seven years − but, legally, you’re still married."

Ron Anderson, MT Editor

nizets2-at-yahoo.com wrote:
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} Hi,
}
} Probably the most interesting way to teach mitosis is
} to synchronize cells in culture. It is probably not
} the easiest though ;-). It is probable that
} unsynchronized but very actively growing cells will
} show enough mitotic events to cover the whole cycle.
} Most of the time they are easy to recognize by normal
} phase constrast, although fluorescent staining like
} DAPI are very fast to perform and very impressive to
} observe.
} I must admit that after working now for more than 10
} years with cells in culture, seeing them split in late
} telophase is still an exciting event. I witness events
} which are actually occuring in my own body at that
} very moment!!!
}
} I agree that you should insist on the functional part
} though. It is not easy to comprehend these phenomenon
} in themself, but understand the difference between
} mitosis and meiosis is really a hard task.
}
} Good luck
}
} Stephane
}
} --- mhunt-at-flinnsci.com wrote:
}
}
} }
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} } Below is the result of your feedback form
} } (NJZFM-ultra-55). It was submitted by
} } (mhunt-at-flinnsci.com) from
} }
} }
} http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html
}
} } on Friday, May 26, 2006 at 09:59:18
} }
} }
} ---------------------------------------------------------------------------
}
} } Email: mhunt-at-flinnsci.com
} } Name: Maureen Hunt
} }
} } Organization: Flinn
} }
} } Education: 9-12th Grade High School
} }
} } Location: Batavia, Illinois, USA
} }
} } Question: I am a former materials microscopist and
} } teacher who is trying to find a more exciting way to
} } teach mitosis and meiosis. Currently, everyone uses
} } prepared slides of whitefish blastula to teach
} } animal mitosis and either prepared slides or
} } unprepared slides of allium root tips to teach plant
} } mitosis. Is there a protist or other "animal-like"
} } eukaryote that can be given to the students,
} } squashed and stained to show animal mitosis? I have
} } googled and come up short. Any suggestions?
} } Thank-you for your time.
} }
} }
} }
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}
} }
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} } Ask-A-Microscopist)
} } 7, 13 -- Subject: AskAMicroscopist: trying to find
} } a more exciting way to teach
} } 7, 13 -- mitosis
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The Olympus with the EM camera is great for high frame rates. No
laser. The spinning slits are not as confocal as spinning pinholes,
but it is quite bright. I was recently torturing the EM camera and
were getting rates that I did not think were possible. Check it out.
David

On Jun 13, 2006, at 5:21 AM, iztok.dogsa-at-ijs.si wrote:

}
}
}
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} Email: iztok.dogsa-at-ijs.si
} Name: iztok dogsa
}
} Organization: IJS
}
} Title-Subject: [Filtered] confocal spinning disk microscope
}
} Question: Has anyone experince with confocal spinning disk microscopy?
}
} Which system would you suggest?
}
} We'd like to achieve really high frame rates, but don't want to buy
} the laser. The arc lamp is Ok for us.
}
} What about BD CARV II imaging system? Is it better than olympus DSU?
}
} Thanks & regards to you all,
}
} Iztok
}
}
} ----------------------------------------------------------------------
} -----
}
} ==============================Original
} Headers==============================
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Thanks to everyone who responded to my question regarding sectioning carbon
nanotubes. This listserve is a great resource.

Best wishes,

Soumitra

**********************************************************************************
Soumitra Ghoshroy
College Associate Professor, Biology
Director, Electron Microscopy Lab
New Mexico State University
Las Cruces, NM 88003
Tel: 505-646-3268 (office)
505-646-3283 (lab)
Fax: 505-646-3282
http://biology-web.nmsu.edu/ghoshroy/ghoshroy.htm
http://emldata.nmsu.edu

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Hello all,

I'm in the market for a stand alone, oil-less, glow discharge unit.
Recommendations would be greatly appreciated.

Thanks, Kim

Kimberly A. Riddle
Florida State University
Biological Science Imaging Resource
119 Bio Unit I, 4370
Tallahassee, FL 32306
tel: 850.644.6519
fax: 850.644.0481
http://bsir.bio.fsu.edu

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Any recommendations for someone to upgrade a PGT EDS detector from standard
Be window to thin window? The instrument is in the Houston, Texas area.

Thanks in advance for any leads or suggestions.

Regards,
Andrew T. Werner
Shaped Charge Research Metallurgy Laboratory
Schlumberger Reservoir Completions Technology Center
14910 Airline Road, Rosharon, TX 77583-1590
Voice (281) 285-5272 Fax (281) 285-5228

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On Jun 13, 2006, at 9:07 AM, riddle-at-bio.fsu.edu wrote:

} I'm in the market for a stand alone, oil-less, glow discharge unit.
} Recommendations would be greatly appreciated.
}
Dear Kim,
We have used our Harrick Basic Plasma Cleaner for several trouble-free
years, and they also make an Extended Plasma Cleaner, which is a larger
model. You would have to buy an oil-free pump to go with either unit,
but a diaphragm pump would be adequate for glow-discharging.
Yours,
Bill Tivol, PhD
EM Scientist and Manager
Cryo-Electron 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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Dear Listers;

I have an individual learning TEM. He will be doing a lot of mouse brain
tissue infected with HIV. What sort of safety or handling procedures
should we follow? I don't have experience in this area with this sort of
pathogen. The tissue is arriving in my lab fixed in 2%glutaraldehyde,
2%paraformaldehyde, 0.5%acrolein in .1M phosphate buffer which I provided
to his lab. So, for processing up through embedding can we just handle the
samples the same as any other tissue? When sectioning the blocks do the
shavings from trimming the block, sectioning, etc. have to be specially
collected? If he nicks himself with the razor blade while trimming the
block is there an increased risk? Should I have him wearing exam gloves
while trimming? Any and all advice will be greatly appreciated.

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

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Does anyone have a copy of a service manual for the Sorvall MT-2B
microtomes? I have the (original) manuals that came with the things,
but not a take-it-apart-and-fix-it service manual. If such things
still exist, I would appreciate a photocopy.
Many thanks!

Phil
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Tom

Glutaraldehyde and Formaldehyde are both very good inactivators. When I
do level 4 stuff they treat in 4%PF for 48hr to 4 weeks, depending on
the protocol they follow at the time, so maybe you want to increase to a
more Karnovsky like fix and make the PF 4%. That works for them, it
should work for you. Under either condition, by the time you get your
stuff it will be well inactivated, and safe.

Paul

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Email: emily.wiesner-at-medecine.unige.ch
Name: Emily Wiesner

Organization: Uni. of Geneva

Title-Subject: [Filtered] Blood vessel staining

Question: Hi All,
I was wondering if anyone call suggest a marker for endothelial cells in the rat brain (cryostat sections). I wish to count the density of blood vessels in the brains of rat pups who have been exposed to undernutrition or glucocorticoid treatment. I have tried CD31 from Chemicon with little success. I am looking for a structural marker of blood vessels- a marker which will stain all of the blood vessels and not be influenced or altered by undernutrition or hypoxia.
Thanks in advance,
Emily

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Email: Dandersen-at-optimedica.com
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Organization: OptiMedica

Title-Subject: [Filtered] Histo Prep of ophthalmic samples for EM

Question: I am looking for a private laboraory to process some ocular samples I wish to analyze under EM.

Any suggestions?

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On Jun 12, 2006, at 9:23 PM, Goinoutonalamb-at-yahoo.com wrote:

} Question: I am collaborating with a group on a 3D visualization
} project through Brookhaven National Labs and I have a question
} pertaining to 3D reconstruction by means of electron tomography. From
} the literature that I have read, I've gathered that it's recommended
} that a 200 kv beam or higher is used when working with sections of 120
} nm or thicker. Unfortunately, the philips EM-300 that we are working
} with is only capable of producing a 100 kv beam. Because of the time
} constraints of our project (10 weeks) it will be very difficult to
} determine what section thicknesses/maximum tilt angles will work with
} 100kv with trial and error. Is there a formula, or a guideline of
} some sort to follow to determine how much mass a 100kv beam can pass
} with minimal loss of resolution? We are planning on doing a tilt
} series from -60 to +60 degrees, in 2 degree increments, at 120 nm
} sections (hopefully). If anyone could lend me some insight as to
} whether or not this is feasible, and how !
} far we can go, I would greatly appreciate it!
}
Dear Michael,
Some issues not answered by the other person who responded to your
post are what the specimen consists of, and what resolution you want.
For biological materials and polymers consisting of low-Z materials,
the penetration of the beam will be greater than for electronics
components consisting of silicon or germanium, and minerals containing
high-Z atoms will be more restricted still. On the other hand, some of
the restriction on thickness comes from the fact that at high tilt, the
effective thickness is greater than the specimen thickness--you mention
60 degrees, at which angle it is twice the thickness. You may be able
to get significant information by taking stereo pairs; these are
obtained at small tilts, usually {10 degrees, so a thicker specimen can
be observed in 3-D through a stereo viewer, which may be all you need,
and 3-D reconstructions can be made with such software as sterecon,
which was used at the Wadsworth Center in Albany NY when I was there.
If your specimen exceeds 120 nm by a small amount, you can still try
tomography with an angular range less that +/-60 degrees and obtain
useful data (although artifacts from the missing wedge will be more
pronounced the smaller the angular range), and you can take dual-axis
tilt series, which will reduce the missing wedge to a missing pyramid
and lessen the missing data artifacts. Finally, if the Wadsworth
Center is still a Regional Resource, you might be able to get time on
one of their higher-voltage EMs, and they have a 1.2 MV scope, which
should provide sufficient penetrating power even for roughly 1 um
sections and/or for higher-Z materials. Good luck.
Yours,
Bill Tivol, PhD
EM Scientist and Manager
Cryo-Electron 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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Email: aribaudo-at-triprinceton.org
Name: Anthony Ribaudo

Organization: TRI Princeton

Title-Subject: [Filtered] TEM Sample Preparation of Paper using Foraldehyde

Question: Dear All,

Can anyone suggest the experimental setup, i.e. the apparatus needed to prepare paper samples that utilizes formaldehyde to crosslink the paper fibers. Formaldehyde imparts water-resistant qualities and increases the dimensional stability against moisture. The formaldehyde reaction conditions recommended are as follow:
Temperature 140��C.
Reaction period 0.5-48 hours.
Formaldehyde concentration introduced in vapor phase 2.19 x 10-2 mol/L.

After treatment the paper sample is embedded in epoxy resin and ultrasectioning.

Thank you,

Anthony Ribaudo
TRI Princeton
601 Prospect Avenue
Princeton, NJ 08542

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Hi,

I was asked to look into the morphology and cause of tin whiskers in high
tin content lead free solders by a guy I meet. Some of the following
questions have been asked for 50 years. Maybe someone out there can share
some more recent expertise about tin whiskers.

* What is the root cause of crystalline tin whiskers? (It's probably not
resolved yet. There are lots of conflicting opinions and observations on
Google.)

* How do you prevent tin whiskers from growing out of almost pure tin
solder joints?
(He is not that interested in a thin film coating solution.)
* What is the mechanism(s) that can cause 2 micron diameter or less
"fibers" (et al) to grow right out of the surface of solder joins to a
length of up to one observed to be 10 mm long?
* What are the EDS results of tin whisker analysis?
* Are the EDS analysis results settled science?
* Have solder joints been ion milled to explore the interior of the solder
or is this strictly a compression/stress surface phenomena? Or both?

He is NOT interested in high humidity dendritic growth of tin on PCB
surfaces reputed to be caused by electric lines and associated
equipotential lines. He is interested in the shorts caused by tin whiskers
bridging solder joints through the air or a vacuum.

This raises the following big question.
* How do you accelerate the growth of tin whiskers so that they can be
studied without waiting 1-15 years for them to develop?

Please reply off list with any insights you have on this problem. I don't
think this subject will have wide appeal. Thank you in advance for any
perspectives that can be shared with him.
If you are speculating on a solution or theory, please say so in your reply.

Paul

Disclaimers: I do not work for this guy's company, the guy, or any company
or organization working on tin whiskers.
I was astonished at the phenomenon, morphology, and how they ruined an
obiting satellite.
Tin whiskers grow in air or vacuum.

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Paul,

You're right. There is not a clear concensus on the formation of tin
whiskers. A couple of great sites on the subject are
http://www.inemi.org/cms/newsroom/Presentations/TinWhiskerWorkshop_ECTC0
6.html and http://nepp.nasa.gov/whisker/

If you wander around on these sites, you'll find enough reports to read
to keep you busy for quite a while. I notice that an April 2006 report
from NASA (
http://nepp.nasa.gov/whisker/reference/tech_papers/2006-Leidecker-Tin-Wh
isker-Failures.pdf ) states that "If you can't live with tin whiskers,
then "Don't Use Tin"."

There are things that will keep whiskers from growing, most notably
adding lead (but that probably doesn't help much).

Scratching the surface of the tin will influence more whisker growth
(but I'm not sure if it'll be faster whisker growth). There are some
extensive studies that have been done to try to find out what
accelerates whisker growth, so I'd read those before starting my own
study.

Hope this helps.

Diane
_____________________________
Diane Ciaburri
Principal Materials Engineer
General Dynamics
100 Plastics Ave., Rm 2552A
Pittsfield MA 01201
phone: (413) 494-3430
email: diane.ciaburri-at-gd-ais.com

-----Original Message-----
X-from: beaurega-at-westol.com [mailto:beaurega-at-westol.com]
Sent: Thursday, June 15, 2006 10:55 AM
To: Ciaburri, Diane A.

Hello,

Anyone with contact info for a 3rd party service person/organization that
can provide support services for a Rigaku Model: Ultra-18 XRD instrument,
please contact Mr. Ross Potoff at the University of Arizona (520) 621-8101
or email to Potoff-at-email.arizona.edu.

Regards,

Bob Roberts
EM Lab Services, Inc.
449 NW 62nd St.
Topeka, Kansas 66617-1780
785.246.1232 voice
785.246.0168 fax
www.emlabservices.com

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Hi,

I would be curious to see any feedback on this subject, in the name of
education.

I have a couple questions. Some of the lead-free solders have silver in
them. Are you (or he) sure the whiskers are tin, or could they be silver?
When you say "vacuum", are you referring to the low pressures in orbit, or
a clean, high vacuum in a chamber of some sort? (or, perhaps, vacuum tube
electronic component)

I believe the fields associated with a bias have something to do with it,
but don't have any facts, or know what the mechanism is. I wonder if a
high (strong) bias, in a vacuum chamber, would accelerate the phenomenon.

Looking forward to learning more.
Thanks,
Darrell

beaurega-at-westol.com wrote on 06/15/2006 10:55:59 AM:

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} Hi,
}
} I was asked to look into the morphology and cause of tin whiskers in high
} tin content lead free solders by a guy I meet. Some of the following
} questions have been asked for 50 years. Maybe someone out there can
share
} some more recent expertise about tin whiskers.
}
} * What is the root cause of crystalline tin whiskers? (It's probably not
} resolved yet. There are lots of conflicting opinions and observations on
} Google.)
}
} * How do you prevent tin whiskers from growing out of almost pure tin
} solder joints?
} (He is not that interested in a thin film coating solution.)
} * What is the mechanism(s) that can cause 2 micron diameter or less
} "fibers" (et al) to grow right out of the surface of solder joins to a
} length of up to one observed to be 10 mm long?
} * What are the EDS results of tin whisker analysis?
} * Are the EDS analysis results settled science?
} * Have solder joints been ion milled to explore the interior of the
solder
} or is this strictly a compression/stress surface phenomena? Or both?
}
} He is NOT interested in high humidity dendritic growth of tin on PCB
} surfaces reputed to be caused by electric lines and associated
} equipotential lines. He is interested in the shorts caused by tin
whiskers
} bridging solder joints through the air or a vacuum.
}
} This raises the following big question.
} * How do you accelerate the growth of tin whiskers so that they can be
} studied without waiting 1-15 years for them to develop?
}
} Please reply off list with any insights you have on this problem. I
don't
} think this subject will have wide appeal. Thank you in advance for any
} perspectives that can be shared with him.
} If you are speculating on a solution or theory, please say so in your
reply.
}
} Paul
}
} Disclaimers: I do not work for this guy's company, the guy, or any
company
} or organization working on tin whiskers.
} I was astonished at the phenomenon, morphology, and how they ruined an
} obiting satellite.
} Tin whiskers grow in air or vacuum.
}
}
}
}
}
}
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The whiskers are tin. Silver is added to lead solder to reduce
the melting point.

Whiskering occurs at room temperature, high and low humidity and
all sorts of conditions. It is a nasty problem for commercial
and especially MIL applications. Alternate solders are being
studied (different combinations of elements) but their melting
point is sufficiently higher than Pb solder to be a real problem.

gary g.

At 01:35 PM 6/15/2006, you wrote:

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I am in the middle of efforts to transfer manufacturing operations from
Lead:Tin solder to Lead Free solder. Presently Silver and copper are used
with the tin to reduce the melting point to about 220C. There are many different theories about why tin whiskers form including a compressive stress on the tin layer. The stress may be caused by copper diffusing into the tin at grain boundaries and forming copper:tin complexes which take up more volume.

There is a historical paper covering the history of the research entitled:
A History of Tin Whisker Theory: 1946 to 2004
George T. Galyon
IBM eSG Group
Poughkeepsie, New York

I'm sorry I can't find the original source for the article. It isn't mentioned on my document. But George T. Gaylon has been (and may still be) associated with iNEMI. Some of his material is available through them at
http://www.inemi.org/cms/newsroom/NEMI_NIST_TMS_workshop.html

Hope this helps. The electronics industry has done a lot of work in this area in the past five or so years.

Mark Woolley
PTRL Lab
Avaya, Inc.
1300 West 120th Ave.
Westminster, CO 80234
(303) 538-9179�� Office
(303) 538-2166�� Lab
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Hi all.
I would like to take a look
a the moth eye structure or a fly,
but I'm completely new to
biological sample...
What kind of sample preparation is recommended
(any essiccation procedure)?
Or it is simple possible to coat it with a
metal layer and this will avoid it to explode and splatter
around the column?
what will happen in the latter case?
(just will have to wait longer for a good vacuum
next time, or ppermanent damage to any parts?)

Please let me know.
Any help is welcome.
All the best,
Marco.

xxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxx
Dr. Marco Salerno, Ph.D.:
marco.salerno-at-unile.it
CNR-INFM Center NNL:
www.nnl.it
c/o Palazzine Garrisi,
via per Arnesano 16 (km 5)
73100 Lecce - Italy
Off. +39-0832-29.8236
Lab.+39-0832-29.8336/8347
Mob.+39-349-26.75.277
Fax +39-0832-29.8238
Home +39-0832-32.51.45
xxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxx

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Dan,

you will find some EM analytical labs at this site:
http://microanalysis.mikroanalytik.de/service%20microanalysis%20e.phtml

Regards

Frank

Dandersen-at-optimedica.com wrote:

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

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Marco,

I routinely image insects in the SEM without any preparation other
than sputter coating. I kill the insects first with ethyl acetate
fumes or CO2. Then mount using a double-sticky carbon conductive tab,
and coat. Note: if the insect/etc. only contacts the mounting tab in
a small area because of convexity, you may have to also use some
silver paint to improve the electrical ground. This is commonly
needed (something I do automatically). The paint requires a wait to
dry. Nor is it always needed -- the University president brought his
6 year-old son to our facility with a beetle to see the SEM. It was a
quick mount, no paint, 3 minute Au coat, and in the SEM. Good points
for the facility, too.
If the critters aren't particularly hairy or you have access to an
ESEM, you might be able to leave off the coating.
Some insects and arachnids can be imaged at low kVs without coating,
and will survive the experience.
Note though, that soft-bodied arthropods can (and will) shrink and
distort in the vacuum. Robust ones hold up better. Beetles are
particularly good for this.
Make certain your insect is *intact*. Any missing limbs, antennae,
etc. will allow for haemolymph to be sucked out into the chamber.
This will cause the body to collapse and coat the innards of your
'scope with bug goo. Not good.
I wouldn't do this at all in an instrument used for high-resolution
or analytical work.
Personally, though, external eye structure is a bit boring. Feet and
especially mouthparts are much more interesting. Plus, their
morphology is directly related to ecology. Eyes are, yes, but less
obviously in external SEM-level anatomy.
There are lots of examples on the web, as well as books of example
images, with technique chapters.

Phil

} Hi all.
} I would like to take a look
} a the moth eye structure or a fly,
} but I'm completely new to
} biological sample...
} What kind of sample preparation is recommended
} (any essiccation procedure)?
} Or it is simple possible to coat it with a
} metal layer and this will avoid it to explode and splatter
} around the column?
} what will happen in the latter case?
} (just will have to wait longer for a good vacuum
} next time, or ppermanent damage to any parts?)
}
} Please let me know.
} Any help is welcome.
} All the best,
} Marco.
}
}
} xxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxx
} Dr. Marco Salerno, Ph.D.:
} marco.salerno-at-unile.it
} CNR-INFM Center NNL:
} www.nnl.it
} c/o Palazzine Garrisi,
} via per Arnesano 16 (km 5)
} 73100 Lecce - Italy
} Off. +39-0832-29.8236
} Lab.+39-0832-29.8336/8347
} Mob.+39-349-26.75.277
} Fax +39-0832-29.8238
} Home +39-0832-32.51.45
} xxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxxx
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Here are my thoughts:

If this phenomenon can be reliably reproduced, the
effect from bias should be easily confirmed or denied.

This appears to be some type of transport mechanism...
vapor transport, surface transport, bulk diffusion
from the body of the metal, or grain boundary
diffusion. It would be interesting to find where tin
is being depleted in the solder joint, perhaps using
EDS methods and metallographic cross sections. The
challenge would be to distinguish tin depletion from
normal solder casting segregation.

At the moment I'm assuming the whisker is a single
crystal. Perhaps the whisker can be studied using
x-ray crystallographic methods - either Laue or TEM
methods. This may give a clue to the growth
mechanism.

Stu Smalinskas, P.E.
Metallurgist
SKF
Plymouth, Michigan

--- gary-at-gaugler.com wrote:

The whiskers are tin. Silver is added to lead solder
to reduce the melting point.

Whiskering occurs at room temperature, high and low
humidity and all sorts of conditions. It is a nasty
problem for commercial and especially MIL
applications. Alternate solders are being studied
(different combinations of elements) but their melting
point is sufficiently higher than Pb solder to be a
real problem.

gary g.

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

At 01:35 PM 6/15/2006, you wrote:

Hi,

I would be curious to see any feedback on this
subject, in the name of education.

I have a couple questions. Some of the lead-free
solders have silver in them. Are you (or he) sure the
whiskers are tin, or could they be silver? When you
say "vacuum", are you referring to the low
pressures in orbit, or a clean, high vacuum in a
chamber of some sort? (or, perhaps, vacuum tube
electronic component)

I believe the fields associated with a bias have
something to do with it, but don't have any facts, or
know what the mechanism is. I wonder if a high
(strong) bias, in a vacuum chamber, would accelerate
the phenomenon.

Looking forward to learning more.
Thanks,
Darrell

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

beaurega-at-westol.com wrote on 06/15/2006 10:55:59
AM:

Hi,
I was asked to look into the morphology and cause of
tin whiskers in high tin content lead free solders by
a guy I meet. Some of the following questions have
been asked for 50 years. Maybe someone out there can
share some more recent expertise about tin whiskers.

* What is the root cause of crystalline tin whiskers?
(It's probably not resolved yet. There are lots of
conflicting opinions and observations on Google.)

* How do you prevent tin whiskers from growing out of
almost pure tin solder joints? (He is not that
interested in a thin film coating solution.)

* What is the mechanism(s) that can cause 2 micron
diameter or less "fibers" (et al) to grow right out of
the surface of solder joins to a length of up to one
observed to be 10 mm long?

* What are the EDS results of tin whisker analysis?

* Are the EDS analysis results settled science?

* Have solder joints been ion milled to explore the
interior of the solder or is this strictly a
compression/stress surface phenomena? Or both?

He is NOT interested in high humidity dendritic growth
of tin on PCB surfaces reputed to be caused by
electric lines and associated equipotential lines. He
is interested in the shorts caused by tin whiskers
bridging solder joints through the air or a vacuum.

This raises the following big question.
* How do you accelerate the growth of tin whiskers so
that they can be studied without waiting 1-15 years
for them to develop?

Please reply off list with any insights you have
on this problem. I don't think this subject will have
wide appeal. Thank you in advance for any
perspectives that can be shared with him. If you are
speculating on a solution or theory, please say so in
your reply.

Paul

Disclaimers: I do not work for this guy's company,
the guy, or any company or organization working on tin
whiskers. I was astonished at the phenomenon,
morphology, and how they ruined an obiting satellite.
Tin whiskers grow in air or vacuum.

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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Ralph,

I recall that there is a MT-2B in rm 110. What I don't remember is
whether or not you have anywhere a *service* manual for one.
We have the manuals that were shipped with the microtomes, even one
old enough to be in a binder. We don't have a service manual.
Do you have one, and if yes, can I get a photocopy of it? We have 3
of them, one horrible that's at best parts, and one I'm trying to
resurrect before classes start this fall.
Thanks.

Phil
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Hi,

If anyone out there has been doing any immunogold labeling for GFP, I would be interested in what primary you're using and how well it labels, depending on fixation.

Have a good weekend....

Lesley

Lesley S. Bechtold
Senior Manager, Histopathology & Microscopy Sciences
The Jackson Laboratory
600 Main St.
Bar Harbor, ME 04609
207-288-6322

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Sorry. My mousing finger glitched and picked the wrong name in my address book.
Phil
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Stu,

This is a well known problem that goes back to the early space programs.
It has shown-up again since the "world" is switching to no-lead solders
due the EU RoHS initiatives. Components cannot contain lead (Pb)in
their lead (frame) finishes, so pure tin is the easiest solution.
Unfortunately, component manufacturers are not aware of or forgot about
the whisker problem also.

One of the most plausible scenarios was presented by Dr. Tu of UCLA at
the 2003 ISTFA conference (See "Solder Failure Mechanisms, K.N. Tu, 29th
International Symposium for Testing and Failure Analysis -
Microelectronics Seminar, ASM International, 2003, P.187). He asserts
that the whiskers grow in response to induced stresses in the tin plate
deposit, caused by the volumetric changes due to Cu-Sn intermetallic
compound formation. A nickel underplate slows this process due to
slower Ni-Sn intermetallic compound growth kinetics.

For more info on tin whisker failures, go to
http://nepp.nasa.gov/whisker/

Joseph

Joseph M. Oparowski
Center for Materials Science
Bose Corporation Joseph_Oparowski-at-bose.com
The Mountain, M/S 415 Phone: (508) 766-1371
Framingham, MA 01701-9168 Fax: (508) 766-1313

-----Original Message-----
X-from: smalinskas-at-yahoo.com [mailto:smalinskas-at-yahoo.com]
Sent: Friday, June 16, 2006 10:59 AM
To: Oparowski, Joseph

Here are my thoughts:

If this phenomenon can be reliably reproduced, the
effect from bias should be easily confirmed or denied.

This appears to be some type of transport mechanism...
vapor transport, surface transport, bulk diffusion
from the body of the metal, or grain boundary
diffusion. It would be interesting to find where tin
is being depleted in the solder joint, perhaps using
EDS methods and metallographic cross sections. The
challenge would be to distinguish tin depletion from
normal solder casting segregation.

At the moment I'm assuming the whisker is a single
crystal. Perhaps the whisker can be studied using
x-ray crystallographic methods - either Laue or TEM
methods. This may give a clue to the growth
mechanism.

Stu Smalinskas, P.E.
Metallurgist
SKF
Plymouth, Michigan

--- gary-at-gaugler.com wrote:

The whiskers are tin. Silver is added to lead solder
to reduce the melting point.

Whiskering occurs at room temperature, high and low
humidity and all sorts of conditions. It is a nasty
problem for commercial and especially MIL
applications. Alternate solders are being studied
(different combinations of elements) but their melting
point is sufficiently higher than Pb solder to be a
real problem.

gary g.

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

At 01:35 PM 6/15/2006, you wrote:

Hi,

I would be curious to see any feedback on this
subject, in the name of education.

I have a couple questions. Some of the lead-free
solders have silver in them. Are you (or he) sure the
whiskers are tin, or could they be silver? When you
say "vacuum", are you referring to the low
pressures in orbit, or a clean, high vacuum in a
chamber of some sort? (or, perhaps, vacuum tube
electronic component)

I believe the fields associated with a bias have
something to do with it, but don't have any facts, or
know what the mechanism is. I wonder if a high
(strong) bias, in a vacuum chamber, would accelerate
the phenomenon.

Looking forward to learning more.
Thanks,
Darrell

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

beaurega-at-westol.com wrote on 06/15/2006 10:55:59
AM:

Hi,
I was asked to look into the morphology and cause of
tin whiskers in high tin content lead free solders by
a guy I meet. Some of the following questions have
been asked for 50 years. Maybe someone out there can
share some more recent expertise about tin whiskers.

* What is the root cause of crystalline tin whiskers?
(It's probably not resolved yet. There are lots of
conflicting opinions and observations on Google.)

* How do you prevent tin whiskers from growing out of
almost pure tin solder joints? (He is not that
interested in a thin film coating solution.)

* What is the mechanism(s) that can cause 2 micron
diameter or less "fibers" (et al) to grow right out of
the surface of solder joins to a length of up to one
observed to be 10 mm long?

* What are the EDS results of tin whisker analysis?

* Are the EDS analysis results settled science?

* Have solder joints been ion milled to explore the
interior of the solder or is this strictly a
compression/stress surface phenomena? Or both?

He is NOT interested in high humidity dendritic growth
of tin on PCB surfaces reputed to be caused by
electric lines and associated equipotential lines. He
is interested in the shorts caused by tin whiskers
bridging solder joints through the air or a vacuum.

This raises the following big question.
* How do you accelerate the growth of tin whiskers so
that they can be studied without waiting 1-15 years
for them to develop?

Please reply off list with any insights you have
on this problem. I don't think this subject will have
wide appeal. Thank you in advance for any
perspectives that can be shared with him. If you are speculating on a
solution or theory, please say so in your reply.

Paul

Disclaimers: I do not work for this guy's company,
the guy, or any company or organization working on tin whiskers. I was
astonished at the phenomenon, morphology, and how they ruined an obiting
satellite.
Tin whiskers grow in air or vacuum.

__________________________________________________
Do You Yahoo!?
Tired of spam? Yahoo! Mail has the best spam protection around
http://mail.yahoo.com

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We are starting a project looking at otolith microstructure, and I was
wondering if folks have recommendations for mounting resins. We have
one reference that recommends spurr's resin. Does anybody have
experience with this type of preparation or a recommendation?

Karl Hagglund
Biological Sciences, SC-102B
Northern Kentucky University, Nunn Drive
Highland Heights, KY 41099
859-572-5238
http://semlab.nku.edu

==============================Original Headers==============================
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It's not too late to enter the International Metallographic Contest and
Exhibit co-sponsored since 1972 by the International Metallographic Society
and ASM International. The contest will be held in conjunction with the 39th
annual technical meeting of the IMS and the M&M 2006 meeting this summer in
Chicago. Twelve categories of competition. Best in show receives $3000 and
the prestigious Jaquet-Lucas Award. Cash awards for first, second, and third
place winners in eleven of the categories. Entries are prominently displayed
during the M&M 2006 meeting and again in the fall during the ASM Annual
Event. Deadline for entries is July 17. For additional information including
rules, tips for creating a winning entry, judging guidelines, and examples
of winning entries contact me or visit
http://www.internationalmetallographicsociety.org/contest.html.

Jeff Stewart
International Metallographic Contest Chair
Metallographic Laboratory Manager
Stern-Leach Co.
49 Pearl Street
Attleboro, MA 02703 USA
Phone: 508-222-7400 extension 1329
FAX: 508-699-4030

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Hi,

Our Hitachi S-4700 is suffering from 60 Hz EMF's in our lab. We see
jagged lines in the horizontal direction at mags } /= to 150kX. I
just had a vibration survey done and the results are within Hitachi
specifications.

Do any of you pay particular attention to movement (of people, people
in chairs, talking, etc.) when you are operating the SEM } 200kX? We
pay particular attention to this in our HR-TEM lab. Just wondering
if it is an issue in the FESEM lab too.

I'm looking for solutions. Cheap(er) DYI solutions. Any ideas?

Thanks!

OWen

Owen P. Mills
Director, Materials Characterization & Fabrication Facilities
Electron Optics Engineer, Applied Chemical & Morphological Analysis
Laboratory

Materials Science & Engineering
Michigan Technological University
Rm 512 M&M Bldg.
Houghton, MI 49931
PH 906-369-1875
FAX 906-487-2934
mailto:opmills-at-mtu.edu
http://www.mm.mtu.edu/~opmills

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Hi

I am sorry to say but round the world one of the problems that we all have
is electromagnetic fields. To prove your problem compare working at a long
WD with working at a very short WD. If the problem changes for the better
at short WD it is a field problem. If it stays the same its most probably
vibration or an instrument fault.

Check laboratories above, below and on all sides to track down the equipment
that may be causing the problem, try switching equipment off.

As a warning to all, magnetic fields now prove to be the biggest FEG
installation problem so always have a professional check your site.
Remember the field will change by the inverse square so in some cases to
move the instrument in the room may help. Field reduction systems work if a
field is not over strong but the best route is to find a room which is truly
field free.

Good luck

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: {opmills-at-mtu.edu}
To: {protrain-at-emcourses.com}
Sent: Monday, June 19, 2006 4:37 PM

Hi All,
I have been trying unsuccessfully to immuno-gold label HUVEC cells
that have been embedded in LR White resin. The cells were plated and
grown to supposed confluence, incubated with primary antibodies to
PECAM and another adherence molecule, then fixed in 2% pfa, 0.5%
glut., dehydrated, & embedded. The resin was cured at 50 deg. C
overnight. En face sections were hydrated & blocked with buffer,
then incubated with gold-tagged secondary Ab (anti-mouse or
anti-rabbit). The sections were contrasted with aqueous Ur. Ac. then
Pb citrate. I"ve seen nothing.
For secondaries, I tried both F(ab)2s and whole IgG. I have been
very successful with the same F(ab)2 secondaries on other samples
processed the same way, but on which both primary and secondary Ab
incubations were done on grid.
In the past, we have successfully localized each of these primary Abs
with peroxidase-DAB labelling prior to stringent fixation and
embedding in Spurr's resin.

One of the problems is that HUVECs (human umbilical cord endothelial
cells) are so darned flat...(less than 0.3 micrometers except for the
nucleus) that its easy to cut right through them just trying to get a
smooth section with no holes off of the raw block face. Another
problem is that the proteins of interest are located at the cell
junctions, in little vesicles and its not always easy to find
cell-cell contacts.

My question (finally) is this: Has anyone tried to do an immuno-gold
approach after immuno-peroxidase? We thought we'd like to try
repeating the successful labelling of PECAM with DAB, pre-embedding,
then try for the other Ab post-embedding.

Any ideas? (I'm sure there are....)
thanks in advance,
Lee
--
Leona Cohen-Gould, M.S.
Sr. Staff Associate
Director, Electron Microscopy & Histology Core Facility
Manager, Optical Microscopy Core Facility
Joan & Sanford I. Weill Medical College
of Cornell University
voice (212)746-6146
fax (212)746-8175
http://www.cornellcelldevbiology.org
http://www.cornellbiochem.org

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1) I would worry that the DAB black mass would obscure the gold. I
have not tried it, but it seems like this would be a problem.
2) Why not do primary and secondary Ab before fix, then fix hard and
go into Epon? Your gold would be in the Epon. No post-cutting Ab work.
3) I would recommend cutting cross-sections of the cells. Then you
do not have to worry about missing the flat cells when facing the
block. Also, cell-cell junctions are much easier to see. I have
protocols to do this if you want them.

David

_____________________

David Elliott Ph.D.
Assistant Professor
Department of Cell Biology and Anatomy
University of Arizona College of Medicine
PO Box 245004
Tucson, AZ 85724

Voice: 520-626-7870
Fax: 520-626-2097

On Jun 19, 2006, at 11:54 AM, lcgould-at-med.cornell.edu wrote:

}
} Hi All,
} I have been trying unsuccessfully to immuno-gold label HUVEC cells
} that have been embedded in LR White resin. The cells were plated and
} grown to supposed confluence, incubated with primary antibodies to
} PECAM and another adherence molecule, then fixed in 2% pfa, 0.5%
} glut., dehydrated, & embedded. The resin was cured at 50 deg. C
} overnight. En face sections were hydrated & blocked with buffer,
} then incubated with gold-tagged secondary Ab (anti-mouse or
} anti-rabbit). The sections were contrasted with aqueous Ur. Ac. then
} Pb citrate. I"ve seen nothing.
} For secondaries, I tried both F(ab)2s and whole IgG. I have been
} very successful with the same F(ab)2 secondaries on other samples
} processed the same way, but on which both primary and secondary Ab
} incubations were done on grid.
} In the past, we have successfully localized each of these primary Abs
} with peroxidase-DAB labelling prior to stringent fixation and
} embedding in Spurr's resin.
}
} One of the problems is that HUVECs (human umbilical cord endothelial
} cells) are so darned flat...(less than 0.3 micrometers except for the
} nucleus) that its easy to cut right through them just trying to get a
} smooth section with no holes off of the raw block face. Another
} problem is that the proteins of interest are located at the cell
} junctions, in little vesicles and its not always easy to find
} cell-cell contacts.
}
} My question (finally) is this: Has anyone tried to do an immuno-gold
} approach after immuno-peroxidase? We thought we'd like to try
} repeating the successful labelling of PECAM with DAB, pre-embedding,
} then try for the other Ab post-embedding.
}
} Any ideas? (I'm sure there are....)
} thanks in advance,
} Lee
} --
} Leona Cohen-Gould, M.S.
} Sr. Staff Associate
} Director, Electron Microscopy & Histology Core Facility
} Manager, Optical Microscopy Core Facility
} Joan & Sanford I. Weill Medical College
} of Cornell University
} voice (212)746-6146
} fax (212)746-8175
} http://www.cornellcelldevbiology.org
} http://www.cornellbiochem.org

==============================Original Headers==============================
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Hi, All-

A friend wants to upgrade/keep going their Zeiss DSM 940A SEM.

First, they would like to know of any recommendations for a digital image
acquisition upgrade available for this model. Contact off-list by
commercial companies welcome.

Also, they are interested in anyone they can contact who might have the
technical service manual, electrical schematics, etc., for this machine
and/or spare parts that they might purchase or have donated. They are in
Maine.

Mahalo!
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

==============================Original Headers==============================
8, 19 -- From tina-at-pbrc.hawaii.edu Mon Jun 19 16:35:13 2006
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Tina,

My, isn't this a popular instrument to keep up and running... I have asked
the same in the past few months...

Tell your friend in Maine to contact me, as I have been actively looking
for replacement parts for this instrument, I have run across a fair amount
of information. In fact, there is a group in Maine that has a Zeiss DM940A
that they may be interested in getting to know....

Also, I would be interested in any information you may get from your
request for digital image acquisition.

dj

On Mon, 19 Jun 2006, tina-at-pbrc.hawaii.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-
}
} A friend wants to upgrade/keep going their Zeiss DSM 940A SEM.
}
} First, they would like to know of any recommendations for a digital image
} acquisition upgrade available for this model. Contact off-list by
} commercial companies welcome.
}
} Also, they are interested in anyone they can contact who might have the
} technical service manual, electrical schematics, etc., for this machine
} and/or spare parts that they might purchase or have donated. They are in
} Maine.
}
} Mahalo!
} Tina
}
} ****************************************************************************
} * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
} * Biological Electron Microscope Facility * (808) 956-6251 *
} * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
} ****************************************************************************
}
}
}
} ==============================Original Headers==============================
} 8, 19 -- From tina-at-pbrc.hawaii.edu Mon Jun 19 16:35:13 2006
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==============================Original Headers==============================
8, 19 -- From dljones-at-bestweb.net Tue Jun 20 07:33:03 2006
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (cloverbags-at-yahoo.com) from http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Tuesday, June 20, 2006 at 09:34:21
---------------------------------------------------------------------------

Email: cloverbags-at-yahoo.com
Name: raffaela

Organization: cebu doctors' university

Education: Undergraduate College

Location: Cebu City, Philippines,Asia

Question: what is the pH of a cedarwood oilJuniperus virginiana) for oil immersion objective?
what is the pH of lemongrass oil(Cymbopogon citratus)?

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

==============================Original Headers==============================
7, 12 -- From zaluzec-at-ultra5.microscopy.com Wed Jun 21 07:16:09 2006
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Dear friends,
please find the following CALL for PAPERS for Journal of Biomedical
Optics
All the best.
Alby and Peri

----
May/June 2007

Visible Fluorescent Proteins

Guest Editors:

Alberto "Alby" Diaspro
University of Genoa
Department of Physics
MicroScoBio-LAMBS/IFOM-NANOMED
Via Dodecaneso 33
16146 Genova, Italy
Tel: +39 10 353 6426
Fax: +39 10 314 218
E-mail: diaspro-at-fisica.unige.it

Ammasi Periasamy
University of Virginia
Keck Center for Cellular Imaging
Biology Department
Gilmer Hall (064), McCormick Rd
Charlottesville, VA 22904
Tel: 434-243-7602
Fax: 434-982-5210
E-mail: ap3t-at-virginia.edu

Call for Papers: This special section will focus on the utilization
of visible fluorescent proteins (VFPs) in fluorescence microscopy and
spectroscopy. The use of fluorescent proteins and live-cell imaging
has greatly improved our understanding of cell functioning in recent
years. Discoveries in different organisms and development of
fluorescent proteins are revolutionizing the study of cell behavior
by providing convenient in vivo markers for gene expression and
protein targeting in intact cells and organisms. VFPs coupled to
three-dimensional fluorescence imaging methods such as confocal and
multiphoton excitation microscopy are very effective in live-cell
imaging. Moreover, the need to understand the fluorescence mechanism
under different conditions has inspired a renaissance in single-
molecule methods. Areas of interest also include papers related to
fluorescence resonance energy transfer (FRET) and fluorescence
recovery after photobleaching (FRAP) applications, VFP photophysical
characterization, and photoconvertible and photactivatable mutants.
In vivo applications are welcome as well as comparisons with
potential competitors such as new fluorescent molecules and quantum
dots.

Manuscripts due September 1, 2006

---------

---------------------------------------------
[...] Dance with wolves and count the stars, including the
unseen...(L.Ferlinghetti, Challenges to young poet, 2001)
---------------------------------------------
Alberto Diaspro, MicroScoBIO Research Center, LAMBS-IFOM,
Department of
Physics, University of Genoa, Via Dodecaneso 33, 16146 Genova, Italy
facsimile +39-010314218 - voice +39-0103536426/480/309;
URL: http://www.lambs.it
http://www.wiley.com/WileyCDA/WileyTitle/productCd-0471409200.html
----------------------------------------------

==============================Original Headers==============================
15, 31 -- From diaspro-at-fisica.unige.it Wed Jun 21 18:27:29 2006
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15, 31 -- To: mplsm-users-at-yahoogroups.com,
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15, 31 -- microscopy-at-microscopy.com, Ronzitti Emiliano {ronzi81-at-virgilio.it} ,
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15, 31 -- Mondal Partha Pratim {partha-at-fisica.unige.it} ,
15, 31 -- Magrassi Raffaella {magrassi-at-fisica.unige.it} ,
15, 31 -- Caorsi Valentina {caorsi-at-fisica.unige.it} ,
15, 31 -- Diaspro Alberto {diaspro-at-fisica.unige.it} ,
15, 31 -- Bianchini Paolo {bianchini-at-ge.infm.it} ,
15, 31 -- Gattodub Gattodub {gattodub-at-email.it} ,
15, 31 -- Mazza Davide {mazza-at-fisica.unige.it} ,
15, 31 -- Morotti Federica {morottina-at-yahoo.it} ,
15, 31 -- Testa Ilaria {ilaria.testa-at-email.it} ,
15, 31 -- Vicidomini Giuseppe {vicidomini-at-fisica.unige.it} ,
15, 31 -- Testa Ilaria {testa-at-fisica.unige.it} ,
15, 31 -- Mario Faretta {mario.faretta-at-ifom-ieo-campus.it}
15, 31 -- From: diaspro {diaspro-at-fisica.unige.it}
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Colleagues....

A enthusiastic construction worker cut a major telecom trunk
line out here in the suburbs of Chicago a little over 24 hours ago.
Took out ALL telecom links, phone lines, networks the whole system. It
made for a very quiet and annoying 24 hours here. Amazing how
rapidly wedded we become to communications links.

Anyway, although the Listserver was running it was not able to "talk to anyone".

If you attempted posting anything during the las 24 hours there is a
good likelihood it was held in queue somewhere on the net and it
may get resent. If your not sure then feel free to repost
any messages.

Cheers...

Nestor
Your Friendly Neighborhood SysOp
(who had a very quiet 24 + hours )

==============================Original Headers==============================
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---------------------------------------------------------------------------
Remember this posting is most likely not from a Subscriber, so when replying
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Email: kjl226-at-vt.edu
Name: Kathy

Organization: Virginia Tech

Title-Subject: [Filtered] Salary ranges

Question: What is a fair salary for a supervisory position, where the person will run an Electron Microscope Service Laboratory? This lab will offer all phases of processing of SEM amd TEM samples. Sectioning of the TEM samples, thick and/or thins, negative staining. Running both scopes and assist with use of the scopes including capturing digital images.

This person will also be responible for ordering all supplies, undating SOP's as needed, keeping labortories maintained, helping students with projects, keeping inventory of equipment, keeping all equipment in good working condition, calling service personnel when scopes on service contracts need work, and calulating bills for EM services.

Thanks in advance for your opinions.

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

==============================Original Headers==============================
10, 12 -- From zaluzec-at-microscopy.com Wed Jun 21 20:03:12 2006
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10, 12 -- To: microscopy-at-microscopy.com
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10, 12 -- Subject: viaWWW: Salary ranges
10, 12 -- Content-Type: text/plain; charset="us-ascii"
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Check out the Microscopy Today archives for the salary survey done in 2004
by Ron Anderson and Barb Foster. It contains a lot of valuable comparison
data that should help you determine an appropriate salary range for your
geographical location and responsibilities.

Debby

Debby Sherman, Manager 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

On 6/21/06 9:04 PM, "kjl226-at-vt.edu" {kjl226-at-vt.edu} wrote:

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}
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}
} Email: kjl226-at-vt.edu
} Name: Kathy
}
} Organization: Virginia Tech
}
} Title-Subject: [Filtered] Salary ranges
}
} Question: What is a fair salary for a supervisory position, where the person
} will run an Electron Microscope Service Laboratory? This lab will offer all
} phases of processing of SEM amd TEM samples. Sectioning of the TEM samples,
} thick and/or thins, negative staining. Running both scopes and assist with use
} of the scopes including capturing digital images.
}
} This person will also be responible for ordering all supplies, undating SOP's
} as needed, keeping labortories maintained, helping students with projects,
} keeping inventory of equipment, keeping all equipment in good working
} condition, calling service personnel when scopes on service contracts need
} work, and calulating bills for EM services.
}
} Thanks in advance for your opinions.
}
}
}
} ---------------------------------------------------------------------------
}
} ==============================Original Headers==============================
} 10, 12 -- From zaluzec-at-microscopy.com Wed Jun 21 20:03:12 2006
} 10, 12 -- Received: from [206.69.208.22] (mac22.zaluzec.com [206.69.208.22])
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Regarding:

mail: cloverbags-at-yahoo.com
Name: raffaela

Organization: cebu doctors' university

Education: Undergraduate College

Location: Cebu City, Philippines,Asia

Question: what is the pH of a cedarwood oilJuniperus virginiana) for oil immersion objective? what is the pH of lemongrass oil(Cymbopogon citratus)?

1. There are three types of cedar oil:

Virginia cedarwood oil (Juniperus virginiana), Texas cedarwood oil (Juniperus ashei or mexicana), and Western red cedar (Thuja plicata)

I have cedar oil (Texas) from Polysciences. Their MSDS does not list a pH. Nor does Merck list one. The Sigma Aldrich MSDS does not list one, nor does the Libety Nautral Products (Virginia), nor does JT Baker, nor does Well, Naturally Products (Virginia & Texas & Western).

I will try to get a pH on my Texas cedar oil next week and report this at least.

The constituents are mainly cedrene (a terpene) and cedral (cedar camphor).

2. regarding lemongrass oil:

I have found no pH. Well, Naturally Products does not have it in their MSDS.

Constituents (as listed by Merck): 75-85% citral, methylheptenone, citronellel, geraniol, limonene

3. I have a client that extracts, sells, distributes these oils and will check for their data as well next week.

Tony

..........................................................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
pH2, LLC
PO Box 34140
Indianapolis, IN 46234
(317) 752-6386
(317) 409-3238 cell

90% of Risk Management is knowing where to place the decimal point...any consultant can give you the other 10%℠

This message is from pH2. This message and any attachments may contain legally privileged or confidential information, and are intended only for the individual or entity identified above as the addressee. If you are not the addressee, or if this message has been addressed to you in error, you are not authorized to read, copy, or distribute this message and any attachments, and we ask that you please delete this message and attachments (including all copies) and notify the sender by return e-mail or by phone at 317-752-6386. Delivery of this message and any attachments to any person other than the intended recipient(s) is not intended in any way to waive confidentiality or a privilege. All personal messages express views only of the sender, which are not to be attributed to pH2 and may not be copied or distributed without this statement.

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Greetings,
It is likely that the pH of any oil is 7,
almost by definition. I am sure a real chemist
will correct me but pH expresses the relative
abundance of protons over hydroxyls. As oils are
non polar, they presumably repel a positive and
negative charge equally and hence the pH is
'neutral', or a value of 7. Furthermore, the
whole pH system is developed for aqueous
solutions and there is presumably very little
water indeed in the cedar oil, so it may not even
be sensible to speak of pH for oils. Presumably
that is why you can't find values in MSDSs, ehgo.
Perhaps there is some solid-state notion of
'proton activity' that applies? Again maybe a
chemist or physicist will have a more rigorous
answer. Why do you care about the pH of these
oils?

Hope this helps,
Tobias
}
} Regarding:
}
} mail: cloverbags-at-yahoo.com
} Name: raffaela
}
} Organization: cebu doctors' university
}
} Education: Undergraduate College
}
} Location: Cebu City, Philippines,Asia
}
} Question: what is the pH of a cedarwood
} oilJuniperus virginiana) for oil immersion
} objective? what is the pH of lemongrass
} oil(Cymbopogon citratus)?
}
} 1. There are three types of cedar oil:
}
} Virginia cedarwood oil (Juniperus virginiana),
} Texas cedarwood oil (Juniperus ashei or
} mexicana), and Western red cedar (Thuja plicata)
}
} I have cedar oil (Texas) from Polysciences.
} Their MSDS does not list a pH. Nor does Merck
} list one. The Sigma Aldrich MSDS does not list
} one, nor does the Libety Nautral Products
} (Virginia), nor does JT Baker, nor does Well,
} Naturally Products (Virginia & Texas & Western).
}
} I will try to get a pH on my Texas cedar oil
} next week and report this at least.
}
} The constituents are mainly cedrene (a terpene) and cedral (cedar camphor).
}
} 2. regarding lemongrass oil:
}
} I have found no pH. Well, Naturally Products does not have it in their MSDS.
}
} Constituents (as listed by Merck): 75-85%
} citral, methylheptenone, citronellel, geraniol,
} limonene
}
} 3. I have a client that extracts, sells,
} distributes these oils and will check for their
} data as well next week.
}
} Tony
}
} ..........................................................................
} Andrew Anthony "Tony" Havics, CHMM, CIH, PE
} pH2, LLC
} PO Box 34140
} Indianapolis, IN 46234
} (317) 752-6386
} (317) 409-3238 cell
}
} 90% of Risk Management is knowing where to place
} the decimal point...any consultant can give you
} the other 10%��
}
} This message is from pH2. This message and any
} attachments may contain legally privileged or
} confidential information, and are intended only
} for the individual or entity identified above as
} the addressee. If you are not the addressee, or
} if this message has been addressed to you in
} error, you are not authorized to read, copy, or
} distribute this message and any attachments, and
} we ask that you please delete this message and
} attachments (including all copies) and notify
} the sender by return e-mail or by phone at
} 317-752-6386. Delivery of this message and any
} attachments to any person other than the
} intended recipient(s) is not intended in any way
} to waive confidentiality or a privilege. All
} personal messages express views only of the
} sender, which are not to be attributed to pH2
} and may not be copied or distributed without
} this statement.
}
}
}
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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

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Email: rpowell-at-nanoprobes.com
Name: Rick Powell

Organization: Nanoprobes, Incorporated

Title-Subject: [Microscopy] Re: EM: immuno labelling DAB & gold (Commercial Vendor Reply)

Question: Hello Leona and Group:

Not sure how flexible you are in terms of your staining and preparation procedure... a PubMed search with the terms "post-embedding AND gold AND peroxidase" gave me 35 results, citing a variety of combinations of methods.

We have encountered several papers where silver-enhanced Nanogold (disclaimer - Nanogold is our product!) was used with DAB for double, or in the most recent case triple immunolabeling. We have summarized a couple of these in our newsletter - here are the references and links:

(1) Triple labeling, pre-embedding procedure:

Feng, Y.-P.; Li, Y.-Q.; Wang, W.; Wu, S.-X.; Chen, T.; Shigemoto, R., and Mizuno, N.: Morphological evidence for GABA/glycine-cocontaining terminals in synaptic contact with neurokinin-1 receptor-expressing neurons in the sacral dorsal commissural nucleus of the rat. Neurosci. Lett., 18, 144-148.

Reference describing methods in more detail:

Li, J. L.; Wang, D.; Kaneko, T.; Shigemoto, R.; Nomura, S., and Mizuno, N.: Relationship between neurokinin-1 receptor and substance P in the striatum: light and electron microscopic immunohistochemical study in the rat. J. Comp. Neurol., 418, 156�163 (2000).

Our article:

http://www.nanoprobes.com/Vol6_Iss11.html#3

(2) Pre-embedding localization of m2R and either ChAT or VAChT in the nucleus basalis magnocellularis (NBM) in rat brain sections where m2R was detected using pre-embedding ultrasmall (0.8 nm) gold enhanced with HQ Silver, and ChAT or VAChT by pre-embedding immunoperoxidase using the peroxidase anti-peroxidase (PAP) technique:

Decossas, M.; Doudnikoff, E.; Bloch, B., and Bernard, V.: Aging and subcellular localization of m2 muscarinic autoreceptor in basalocortical neurons in vivo. Neurobiol. Aging, 26, 1061-1072 (2005).

Our article:

http://www.nanoprobes.com/Vol6_Iss3.html#5

(3) Similar (pre-embedding labeling of vesicular glutamate transporter 1 (VGLUT1) and choline acetyltransferase (ChAT) immuno-reactivity respectively in the rat lumbar spinal cord):

Wu, S.-X.; Koshimizu, Y.; Feng, Y. P.; Okamoto, K.; Fujiyama, F.; Hioki, H.; Li, Y. Q.; Kaneko, T., and Mizuno, N.: Vesicular glutamate transporter immunoreactivity in the central and peripheral endings of muscle-spindle afferents. Brain Res.,} 1011, 247-251 (2004).

Other pre-embedding method:

Gutekunst, C. A.; Torre, E. R.; Sheng, Z.; Yi, H.; Coleman, S. H.; Riedel, I. B., and Bujo, H.: Stigmoid Bodies Contain Type I Receptor Proteins SorLA/LR11 and Sortilin. New perspectives on their function. J. Histochem. Cytochem., 51, 841-852 (2003).

Our article:

http://www.nanoprobes.com/Vol5_Iss7.html#4

Hope some of these are helpful,

Rick Powell
Nanoprobes, Incorporated
www.nanoprobes.com

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Hi All,
I have been trying unsuccessfully to immuno-gold label HUVEC cells
that have been embedded in LR White resin. The cells were plated and
grown to supposed confluence, incubated with primary antibodies to
PECAM and another adherence molecule, then fixed in 2% pfa, 0.5%
glut., dehydrated, & embedded. The resin was cured at 50 deg. C
overnight. En face sections were hydrated & blocked with buffer,
then incubated with gold-tagged secondary Ab (anti-mouse or
anti-rabbit). The sections were contrasted with aqueous Ur. Ac. then
Pb citrate. I"ve seen nothing.
For secondaries, I tried both F(ab)2s and whole IgG. I have been
very successful with the same F(ab)2 secondaries on other samples
processed the same way, but on which both primary and secondary Ab
incubations were done on grid.
In the past, we have successfully localized each of these primary Abs
with peroxidase-DAB labelling prior to stringent fixation and
embedding in Spurr's resin.

One of the problems is that HUVECs (human umbilical cord endothelial
cells) are so darned flat...(less than 0.3 micrometers except for the
nucleus) that its easy to cut right through them just trying to get a
smooth section with no holes off of the raw block face. Another
problem is that the proteins of interest are located at the cell
junctions, in little vesicles and its not always easy to find
cell-cell contacts.

My question (finally) is this: Has anyone tried to do an immuno-gold
approach after immuno-peroxidase? We thought we'd like to try
repeating the successful labelling of PECAM with DAB, pre-embedding,
then try for the other Ab post-embedding.

Any ideas? (I'm sure there are....)
thanks in advance,
Lee

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To the list:

The histotechnologist in our group is having difficulty keeping formalin-fixed,
paraffin-embedded sections of pig hooves on the slide for staining. Does anyone
have any suggestions for her? She uses superfrost slides which works for all
other tissues. If you have any hints for changes in fixation, etc. please pass
those along.

Thanks!
Peggy
Peggy Sherwood
Lab Associate, Photopathology
Wellman Center for Photomedicine (W224)
Massachusetts General Hospital
55 Fruit Street
Boston, MA 02114
617-724-4839 (voice mail)
617-726-6983 (lab)
617-726-3192 (fax)
msherwood-at-partners.org

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Hi Raffaela

I was under the impression that, although oils can contain acidic
contaminates, these are often removed during the refining process (not being
good for engines etc.). Acidity (the acid-type constituents) "is usually
defined in terms of total acid number (TAN). These constituents vary in
nature and may or may not markedly influence the behaviour of the
lubricant". Low acidity oils can oxidise during use and become more acidic
over time (years) - e.g. the acid number goes to over 0.4 - while the newly
refined 'non-corrosive' oil would be expected to be nearer to 0.05. The
acids can be weak organic and strong inorganic. Acidity in oil can be useful
for things like improved water resistance. Oils don't have an aqueous pH as
such though, just the TAN number (total acidic number).

Olive oil for example is also graded for consumption on it's oleic acid
acidity - around 3% in cheaper olive oil, and down to below 1% in 'extra
virgin'. Oils are generally insoluble in water, which rather helps reduce
it's corrosive properties during use.

Cargille use the Neutralization Number for their immersion oils. This is a
measure of the acidity or alkalinity of the oil and is the mass in
milligrams of the amount of acid (HCl) or base (KOH) required to neutralize
one gram of the oil. For their standard immersion oils and type FF the value
is very low [acidity] at 0.01 [KOH] (whereas the type DF is a rather high at
0.15 and the bottle states that it may damage objectives in constant use -
it has lower auto-fluorescence than FF though). Cargille states that
Cedarwood oil has a KOH neutralisation number greater than these values (ie.
it is more acidic).

When used for objective immersion, the main things of interest in an oil
would be things like : refractive index, colour, fluorescent properties,
likely damage to the lens cement, crystallisation when cold (reversible),
and whether the oil evaporates leaving a hard residue. The metal outside
parts of the objective generally have an anti-corrosion finish, although
here with our inverted microscopes spillage of corrosive culture media has
sometimes caused some damage at the base of the objective, in particular
corrosion of the area around the screw thread. The immersion oils generally
dissolve in things like ethyl alcohol. Rather like engine oils for your car
one tends to stick with the manufacturers [typically expensive] recommended
one and not look too deeply into it's chemistry (microscope manufacturer's
will soak test their objectives immersed for 6 months or more in an
immersion oil before recommending it).

Oil acids can attack the lens mounting cement [see Cargille's comments
below]. Plus the organic solvents used to remove lens immersion oil (like
Xylene and ethanol) have been implicated in lens cement damage - I use ether
generally for cleaning (and only clean rarely, when the lens is too dirty to
be cleaned by just a lens tissue). Modern [low acid content] immersion oils
supplied by the microscope manufacturer seem to be safe for longer term
contact with the objective and don't evaporate to a hard residue (unlike say
corn oil).

Cargille quote immersion oil shelf lives of 10 years (FF) and 5 years (DF) -
and half that when opened.

On Cargille's site [http://www.cargille.com/immersionoilmicroscope.shtml]
John Cargille states:

"To the knowledge of the author, no immersion oil matches the optics of the
microscope perfectly. One of the closest matches is thickened cedarwood oil
which, for many years, was the most widely used, if not the only immersion
oil available. The disadvantageous properties of cedarwood oil are: high
absorption of blue and UV light, yellowing with age, a tendency to harden on
lenses due to uneven volatility, acidity, and changing viscosity (diluting
with solvent changes the index and dispersion). The synthetic immersion oils
eliminate many of the disadvantages cited above.

The acid value of immersion oil should be very low. The synthetics usually
have acid values lower than cedarwood oil.

High acidity can, in time, affect the condition of the metal parts of the
objective, and possibly more important, can cause deterioration of lens
cements. The lens cement is also a seal that prevents oil from penetrating
to the back of the lens. A crack or perforation in the cement draws oil by
capillary attraction and a thin film of oil slowly creeps over the back of
the objective lens; a poor image may develop without being immediately
noticed. When the microscopist realizes the image has deteriorated, he may
not realize the cause unless he has faced this problem before."

See Cargille's excellent site for more immersion oil info.

Regards

Keith

----------------------------------------------------------
Dr Keith J Morris
Imaging Facilities Manager
Cell Biology Division
Institute of Ophthalmology
University College London
11-43 Bath Street
London EC1V 9EL

Tel: 020 7608 4050
Fax: 020 7608 4034
email: keith.morris-at-ucl.ac.uk

-----Original Message-----
X-from: ph2-at-sprynet.com [mailto:ph2-at-sprynet.com]
Sent: 22 June 2006 04:40
To: keith.morris-at-ucl.ac.uk

Regarding:

mail: cloverbags-at-yahoo.com
Name: raffaela

Organization: cebu doctors' university

Education: Undergraduate College

Location: Cebu City, Philippines,Asia

Question: what is the pH of a cedarwood oilJuniperus virginiana) for oil
immersion objective? what is the pH of lemongrass oil(Cymbopogon citratus)?

1. There are three types of cedar oil:

Virginia cedarwood oil (Juniperus virginiana), Texas cedarwood oil
(Juniperus ashei or mexicana), and Western red cedar (Thuja plicata)

I have cedar oil (Texas) from Polysciences. Their MSDS does not list a pH.
Nor does Merck list one. The Sigma Aldrich MSDS does not list one, nor does
the Libety Nautral Products (Virginia), nor does JT Baker, nor does Well,
Naturally Products (Virginia & Texas & Western).

I will try to get a pH on my Texas cedar oil next week and report this at
least.

The constituents are mainly cedrene (a terpene) and cedral (cedar camphor).

2. regarding lemongrass oil:

I have found no pH. Well, Naturally Products does not have it in their
MSDS.

Constituents (as listed by Merck): 75-85% citral, methylheptenone,
citronellel, geraniol, limonene

3. I have a client that extracts, sells, distributes these oils and
will check for their data as well next week.

Tony

..........................................................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
pH2, LLC
PO Box 34140
Indianapolis, IN 46234
(317) 752-6386
(317) 409-3238 cell

90% of Risk Management is knowing where to place the decimal point...any
consultant can give you the other 10%℠

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