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Email: leforestier-at-lps.u-psud.fr
Name: leforestier amelie

Organization: CNRS Laboratoire de Physique des Solides, Orsay

Title-Subject: [Filtered] engineer position in cryo-electron microscopy

Question: We are seeking an engineer in cryo-electron microscopy:
The candidate will be in charge of a 200 kV LaB6 electron microscope equipped with a cryo-specimen holder and a post-column energy filter (GIF). He/she will conduct cryo-TEM experiments in biology as well as in physics of complex systems (polymers, colloids, amphiphiles, �), from specimen preparation to interpretation and image analysis. He/she must possess good abilities to design new and delicate experiments.
Theoretical and practical background in transmission electron microscopy is required. Cryo-EM expertise will be appreciated.
The candidate will integrate a multidisciplinary team "Structure and function of condensed DNA" located at the Solid State Physics Laboratory of the Paris-Sud University (Orsay, France). The position could suit either a physicist or a biologist with strong motivation for interdisciplinarity.
Mastering of the English and French languages is required (the interview will be in French). Application deadline May 22.

Contact: Fran�oise Livolant
Livolant-at-lps.u-psud.fr {mailto:Livolant-at-lps.u-psud.fr}
Tel: 33 1 69 15 53 92
http://www.lps.u-psud.fr/rubrique.php3?id_rubrique=9&lang=fr

Inscription: http://www.education.gouv.fr/personnel/administratif_technique/itrf/inscrire.htm

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Hi Listers,

Just another day in multi-user world.

Today's challenge is trying to image platinum nanoparticles in an
agarose matrix, in order to see how they distribute themselves and to
get a size distribution. TEM and/or SEM are possibilities. So far, my
check of the literature finds tons of stuff on nanoparticles in
electrophoresis gels, but none of it is relevant, since the particles
are removed from the gels and put back into a liquid medium. The one
article I found that is comparable to our problem used thin-sectioning
and we have tried that.

We have also tried melting the agarose, dipping grids into the
particle/agarose mix, then rinsing the grid in hot water to thin the
gelatin out. We can get images in the TEM, but the results are
inconsistent when repeating with the same sample. Also, there is the
chance that the hot water and melting are re-arranging the particles.

We have tried thin sectioning the dehydrated agarose with particles, but
finding the particles in a given thin section is a crap shoot with long
odds. Also, we may be cutting through aggregations we want to see.

We have tried viewing carbon-coated dehydrated agarose with particles
using backscattered electrons in our FESEM. This gives images with
particles, but only those on or right at the surface are imaged clearly
enough for good size data. Plus, we can't see far into the agarose for
good distribution data.

We are going to try increasing the concentration of the particles to
increase chances of getting them reliably in thin sections, and we will
also try putting the melted mixture on cover slips in a thin layer and
re-trying the BSE imaging after carbon coating. (The latter still has
the potential problem of redistributing the particles, however.) We
could also try doing large thick or semi-thin sections and viewing them
in BSE imaging.

However, if someone out there has viewing nanos in agarose down to a
fine art, we, as usual, would be delighted to hear about it. In the
meantime, I will continue to search the databases.

Thanks to all!

Randy

Randy Tindall
EM Specialist
Electron Microscopy Core Facility---We Do Small Immediately----Nano
Takes a Little Longer!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu

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Dear microscopists

I have recently had a discussion with a colleague about the best
protocol to follow when staining cells during a time course experiment.
I don't think there is a single correct answer, however, would like to
know current thinking on the following issue:

Live cells were treated with a compound and observed at various time
points during a period of 48 hours. At each time point, cells were fixed
and immunofluorescently stained for the protein of interest.

Is it a less artefactual procedure to fix cells at each time point and
keep in a buffer until the end of 48 hours to stain them all at the same
time or fix and stain at each sampling time point? To stain at the same
time may would reduce staining differences, however, keeping cells in
buffer for different times may induce changes in the protein.

I look forward to hearing your opinions.
Thank you.
Judy

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

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Judy,
An end run around the problem is to start off the treatments
at different times so they all end together and then fix and stain is
all done at the same time. This doesn't answer your question but
maybe a useful wrinkle?

Good luck,
Tobias
}
}
} Dear microscopists
}
} I have recently had a discussion with a colleague about the best
} protocol to follow when staining cells during a time course experiment.
} I don't think there is a single correct answer, however, would like to
} know current thinking on the following issue:
}
} Live cells were treated with a compound and observed at various time
} points during a period of 48 hours. At each time point, cells were fixed
} and immunofluorescently stained for the protein of interest.
}
} Is it a less artefactual procedure to fix cells at each time point and
} keep in a buffer until the end of 48 hours to stain them all at the same
} time or fix and stain at each sampling time point? To stain at the same
} time may would reduce staining differences, however, keeping cells in
} buffer for different times may induce changes in the protein.
}
} I look forward to hearing your opinions.
} Thank you.
} Judy
}
} Judy Trogadis
} Bio-Imaging Coordinator
} St. Michael's Hospital, 7Queen
} 30 Bond St.
} Toronto, ON M5B 1W8, Canada
} ph: 416-864-6060 x6337
} pager: 416-685-9219
} fax: 416-864-6043
} trogadisj-at-smh.toronto.on.ca
}
}
} ==============================Original Headers==============================
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} 7, 19 -- From: "Judy Trogadis" {TrogadisJ-at-smh.toronto.on.ca}
} 7, 19 -- To: {Microscopy-at-microscopy.com}
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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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There are some other things to consider. First off, a great deal of this
debate will depend on what you are looking for and how it reacts with
your fixative. If the cells are 'lightly fixed' there may be some
reversal of fixation with prolonged buffer storage. Does that effect the
staining? Tobias offered a good suggestion but there might be some
chrono effects, cells fixed at different times of the day or night
depending on your experimental design. I suggest avoiding all problems
and debate by keeping all of the fixation, buffer wash times and
staining times the same. Your staining proceedure should be sufficiently
standardized so that it is not a variable, or is the least problematic
of the potential variables. Finally, people looking for something to
criticize in your proceedures will always find something 'wrong'.

Geoff

TrogadisJ-at-smh.toronto.on.ca wrote:

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--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583; fax: -4029
mcauliff-at-umdnj.edu
**********************************************

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Dear Judy,

I don't know in detail what you want to observe, but isn't there a
fluorescent tracker-molecule (such as a Lyso-tracker) available for your
purpose? Another more ideal solution might be creating a, for that one
protein GFP-positive cell-line?! Than you could make a continuous
time-lapse without the need of fixation etc., all depending on your
experiment's requests of course!

Anyway, if the fixation is strong enough, does the protein still show
activity / are changes still induced? To my opinion and experience, if
fixed strong enough and there are no/little changes, it should not matter
whether the cells are stained immediately or a few hours later, especially
when also stored cold.
Best regards,

Sven Terclavers

-----Original Message-----
X-from: TrogadisJ-at-smh.toronto.on.ca [mailto:TrogadisJ-at-smh.toronto.on.ca]
Sent: dinsdag 2 mei 2006 22:02
To: sven.terclavers-at-med.kuleuven.be

Dear microscopists

I have recently had a discussion with a colleague about the best
protocol to follow when staining cells during a time course experiment.
I don't think there is a single correct answer, however, would like to
know current thinking on the following issue:

Live cells were treated with a compound and observed at various time
points during a period of 48 hours. At each time point, cells were fixed
and immunofluorescently stained for the protein of interest.

Is it a less artefactual procedure to fix cells at each time point and
keep in a buffer until the end of 48 hours to stain them all at the same
time or fix and stain at each sampling time point? To stain at the same
time may would reduce staining differences, however, keeping cells in
buffer for different times may induce changes in the protein.

I look forward to hearing your opinions.
Thank you.
Judy

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

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7, 19 -- From: "Judy Trogadis" {TrogadisJ-at-smh.toronto.on.ca}
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Dear TEM users,

This is a specimen prep question for everyone out there with EM expertise. We are having terrible success with Lead Citrate contrast staining. Principally, we suffer from precipitates showing up all over the specimen. On the advice of EM science technical support, we are double distilling our own water (they say Milli-Q is too pure and also is de-ionized which we don't want for EM). Then we make it CO2 free by autoclaving and capping directly upon removal from the autoclave. We do this the morning of reagent prep--so it doesn't sit in the bottle for longer than it takes to cool down before we begin making up the Reynolds.

We make the Reynolds Lead citrate according to the protocol listed in Ch 5 of Bozzola and Russell's Electron Microscopy (2nd edition) (mix 1.33g lead nitrate, 1.76g sodium citrate, and 30ml CO2 free double distilled water...shake vigorously for a few minutes and then again 5-6 times over the next 30 minutes). Ensure solution is milky white and free of particles. Add 8.0 ml commercially prepared, titrated 1.0 N NaOH--from EM science--solution turns clear. Adjust pH strictly to 12.0+/-0.1 unit. Bring volume to 50ml with CO2-free double distilled water. Stopper tightly with rubber stopper and parafilm until use later that day.

When we stain the grids with lead nitrate, we make sure to wash well before and after with CO2 free double distilled water in addition to surrounding the staining plate (Hiraoka kit) with NaOH pellets. In addition, we centrifuge the Reynolds at 5000xg for 8 minutes prior to use AND we 0.2um filter it into staining plate.

ANY advice or thoughts are welcome...what are we doing wrong?? What can we change about this protocol to ensure ppt free staining?

A thousand thanks in advance!

Danielle Crippen
Morphology and Imaging Core Manager
Buck Institute for Age Research
8001 Redwood Blvd.
Novato, CA 94945
415-209-2046
dcrippen-at-buckinstitute.org

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Danielle,

Are you SURE it's lead citrate precip? There are many other sources of
precipitates and "pepper", as I'm sure you're aware. What kind of
sample are you preparing? What buffer is being used? Are you
osmicating your samples?

We fought a pepper problem for over two years, before finally
discovering that adding 2-mercaptoethanol to our buffer solved the
problem.

Feel free to email an image, if you like.

Randy

Randy Tindall
EM Specialist
Electron Microscopy Core Facility---We Do Small Well!
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65211
Tel: (573) 882-8304
Fax: (573) 884-2227
Email: tindallr-at-missouri.edu
Web: http://www.emc.missouri.edu

-----Original Message-----
X-from: dcrippen-at-buckinstitute.org [mailto:dcrippen-at-buckinstitute.org]
Sent: Tuesday, May 02, 2006 5:47 PM
To: Tindall, Randy D.

Dear TEM users,

This is a specimen prep question for everyone out there with EM
expertise. We are having terrible success with Lead Citrate contrast
staining. Principally, we suffer from precipitates showing up all over
the specimen. On the advice of EM science technical support, we are
double distilling our own water (they say Milli-Q is too pure and also
is de-ionized which we don't want for EM). Then we make it CO2 free by
autoclaving and capping directly upon removal from the autoclave. We do
this the morning of reagent prep--so it doesn't sit in the bottle for
longer than it takes to cool down before we begin making up the
Reynolds.

We make the Reynolds Lead citrate according to the protocol listed in Ch
5 of Bozzola and Russell's Electron Microscopy (2nd edition) (mix 1.33g
lead nitrate, 1.76g sodium citrate, and 30ml CO2 free double distilled
water...shake vigorously for a few minutes and then again 5-6 times over
the next 30 minutes). Ensure solution is milky white and free of
particles. Add 8.0 ml commercially prepared, titrated 1.0 N NaOH--from
EM science--solution turns clear. Adjust pH strictly to 12.0+/-0.1
unit. Bring volume to 50ml with CO2-free double distilled water.
Stopper tightly with rubber stopper and parafilm until use later that
day.

When we stain the grids with lead nitrate, we make sure to wash well
before and after with CO2 free double distilled water in addition to
surrounding the staining plate (Hiraoka kit) with NaOH pellets. In
addition, we centrifuge the Reynolds at 5000xg for 8 minutes prior to
use AND we 0.2um filter it into staining plate.

ANY advice or thoughts are welcome...what are we doing wrong?? What can
we change about this protocol to ensure ppt free staining?

A thousand thanks in advance!

Danielle Crippen
Morphology and Imaging Core Manager
Buck Institute for Age Research
8001 Redwood Blvd.
Novato, CA 94945
415-209-2046
dcrippen-at-buckinstitute.org

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Danielle,

I haven't worked in biological electron microscopy for over 25 years.
However, I vividly remember a colleague having a terrible time with
precipitates from lead citrate staining. Turned out that the problem was
his eye-sight. He was extremely near-sighted. To watch his work, his face
was only several inches from the staining grid and water rinse. The source
of the problem was CO2 from his breath. Another colleague happened to see
his proximity to the stain and suggested the source of the problem. The
precipitates disappeared once he isolated his exhaust from the process.

Food for thought.

Regards,

Gary M. Brown
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

dcrippen-at-bucki
nstitute.org
To
gary.m.brown-at-exxonmobil.com
05/02/06 05:48 cc
PM
Subject
[Microscopy] TEM--Lead
Please respond Citrate--HELP!!
to
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nstitute.org

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Dear TEM users,

This is a specimen prep question for everyone out there with EM expertise.
We are having terrible success with Lead Citrate contrast staining.
Principally, we suffer from precipitates showing up all over the specimen.
On the advice of EM science technical support, we are double distilling our
own water (they say Milli-Q is too pure and also is de-ionized which we
don't want for EM). Then we make it CO2 free by autoclaving and capping
directly upon removal from the autoclave. We do this the morning of
reagent prep--so it doesn't sit in the bottle for longer than it takes to
cool down before we begin making up the Reynolds.

We make the Reynolds Lead citrate according to the protocol listed in Ch 5
of Bozzola and Russell's Electron Microscopy (2nd edition) (mix 1.33g lead
nitrate, 1.76g sodium citrate, and 30ml CO2 free double distilled
water...shake vigorously for a few minutes and then again 5-6 times over
the next 30 minutes). Ensure solution is milky white and free of
particles. Add 8.0 ml commercially prepared, titrated 1.0 N NaOH--from EM
science--solution turns clear. Adjust pH strictly to 12.0+/-0.1 unit.
Bring volume to 50ml with CO2-free double distilled water. Stopper tightly
with rubber stopper and parafilm until use later that day.

When we stain the grids with lead nitrate, we make sure to wash well before
and after with CO2 free double distilled water in addition to surrounding
the staining plate (Hiraoka kit) with NaOH pellets. In addition, we
centrifuge the Reynolds at 5000xg for 8 minutes prior to use AND we 0.2um
filter it into staining plate.

ANY advice or thoughts are welcome...what are we doing wrong?? What can we
change about this protocol to ensure ppt free staining?

A thousand thanks in advance!

Danielle Crippen
Morphology and Imaging Core Manager
Buck Institute for Age Research
8001 Redwood Blvd.
Novato, CA 94945
415-209-2046
dcrippen-at-buckinstitute.org

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On May 2, 2006, at 12:26 PM, TindallR-at-missouri.edu wrote:

} Today's challenge is trying to image platinum nanoparticles in an
} agarose matrix, in order to see how they distribute themselves and to
} get a size distribution. TEM and/or SEM are possibilities. So far, my
} check of the literature finds tons of stuff on nanoparticles in
} electrophoresis gels, but none of it is relevant, since the particles
} are removed from the gels and put back into a liquid medium. The one
} article I found that is comparable to our problem used thin-sectioning
} and we have tried that.
}
} We have also tried melting the agarose, dipping grids into the
} particle/agarose mix, then rinsing the grid in hot water to thin the
} gelatin out. We can get images in the TEM, but the results are
} inconsistent when repeating with the same sample. Also, there is the
} chance that the hot water and melting are re-arranging the particles.
}
} We have tried thin sectioning the dehydrated agarose with particles,
} but
} finding the particles in a given thin section is a crap shoot with long
} odds. Also, we may be cutting through aggregations we want to see.
}
} We have tried viewing carbon-coated dehydrated agarose with particles
} using backscattered electrons in our FESEM. This gives images with
} particles, but only those on or right at the surface are imaged clearly
} enough for good size data. Plus, we can't see far into the agarose for
} good distribution data.
}
} We are going to try increasing the concentration of the particles to
} increase chances of getting them reliably in thin sections, and we will
} also try putting the melted mixture on cover slips in a thin layer and
} re-trying the BSE imaging after carbon coating. (The latter still has
} the potential problem of redistributing the particles, however.) We
} could also try doing large thick or semi-thin sections and viewing them
} in BSE imaging.
}
} However, if someone out there has viewing nanos in agarose down to a
} fine art, we, as usual, would be delighted to hear about it. In the
} meantime, I will continue to search the databases.
}
Dear Randy,
Thick or semi-thick sections in TEM would be my choice--probably since
I have a 300 kV TEM. If you can get to a high-pressure freezer, I
would suggest using that to prepare your specimens, followed by
freeze-substitution and resin embedding. Assuming that you do not need
to image the strands of agarose, just section the embedded specimen and
observe.
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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Reaching into my past......We used special analytical grade NaOH,
since it seemed to us that the NaOH was picking up carbonate from the
air. The analytical grade stuff came in a sealed glass vial, and
made quite a difference.

Joel

Date sent: Tue, 2 May 2006 17:45:41 -0500
To: jbs-at-temple.edu
X-from: dcrippen-at-buckinstitute.org
Send reply to: dcrippen-at-buckinstitute.org

This may be shear luck, but I've never had trouble with precipitate
(other things yes, but ppte no) - I keep my lead citrate (made up
with ordinary distilled water, not specially CO2 free) in a
volumetric flask (50ml), which sits in the same place month after
month and is never moved. I don't use the stain for 24 hours after
it's prepared, but then just carefully take off what's needed from
close to the surface using a glass pipette. I wipe the end of the
pipette with a tissue before dispensing the stain and then discard
the first drop. I put the drops onto Parafilm in a covered glass
petri dish. No need for NaOH pellets. Finally wash the grids for 5
seconds in a gentle stream of water from a wash bottle. I probably
shouldn't admit this but I've had a bottle of stain last over 2 years
(the surface of the bottle becomes cloudy with ppte) and still
produce perfect results.

Cheers,

Diana

On 3 May 2006, at 8:47 AM, dcrippen-at-buckinstitute.org wrote:

}
}
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} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
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} Dear TEM users,
}
} This is a specimen prep question for everyone out there with EM
} expertise. We are having terrible success with Lead Citrate
} contrast staining. Principally, we suffer from precipitates
} showing up all over the specimen. On the advice of EM science
} technical support, we are double distilling our own water (they say
} Milli-Q is too pure and also is de-ionized which we don't want for
} EM). Then we make it CO2 free by autoclaving and capping directly
} upon removal from the autoclave. We do this the morning of reagent
} prep--so it doesn't sit in the bottle for longer than it takes to
} cool down before we begin making up the Reynolds.
}
} We make the Reynolds Lead citrate according to the protocol listed
} in Ch 5 of Bozzola and Russell's Electron Microscopy (2nd edition)
} (mix 1.33g lead nitrate, 1.76g sodium citrate, and 30ml CO2 free
} double distilled water...shake vigorously for a few minutes and
} then again 5-6 times over the next 30 minutes). Ensure solution is
} milky white and free of particles. Add 8.0 ml commercially
} prepared, titrated 1.0 N NaOH--from EM science--solution turns
} clear. Adjust pH strictly to 12.0+/-0.1 unit. Bring volume to
} 50ml with CO2-free double distilled water. Stopper tightly with
} rubber stopper and parafilm until use later that day.
}
} When we stain the grids with lead nitrate, we make sure to wash
} well before and after with CO2 free double distilled water in
} addition to surrounding the staining plate (Hiraoka kit) with NaOH
} pellets. In addition, we centrifuge the Reynolds at 5000xg for 8
} minutes prior to use AND we 0.2um filter it into staining plate.
}
} ANY advice or thoughts are welcome...what are we doing wrong??
} What can we change about this protocol to ensure ppt free staining?
}
} A thousand thanks in advance!
}
} Danielle Crippen
} Morphology and Imaging Core Manager
} Buck Institute for Age Research
} 8001 Redwood Blvd.
} Novato, CA 94945
} 415-209-2046
} dcrippen-at-buckinstitute.org
}
}
}
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Dear Danielle,

We keep lead citrate prepared by reynold's method for
months in a volumetric flask, Use Whatman 1 to filter
and make drops on a parafilm in a petridish containing
NaOH pellets. After staining for 1-3 min, wash the
grids in water, then water with about 0.01% NaOH and
again water. Never had any precipitates.

regrads
shashi
--- dcrippen-at-buckinstitute.org wrote:

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} Dear TEM users,
}
} This is a specimen prep question for everyone out
} there with EM expertise. We are having terrible
} success with Lead Citrate contrast staining.
} Principally, we suffer from precipitates showing up
} all over the specimen. On the advice of EM science
} technical support, we are double distilling our own
} water (they say Milli-Q is too pure and also is
} de-ionized which we don't want for EM). Then we
} make it CO2 free by autoclaving and capping directly
} upon removal from the autoclave. We do this the
} morning of reagent prep--so it doesn't sit in the
} bottle for longer than it takes to cool down before
} we begin making up the Reynolds.
}
} We make the Reynolds Lead citrate according to the
} protocol listed in Ch 5 of Bozzola and Russell's
} Electron Microscopy (2nd edition) (mix 1.33g lead
} nitrate, 1.76g sodium citrate, and 30ml CO2 free
} double distilled water...shake vigorously for a few
} minutes and then again 5-6 times over the next 30
} minutes). Ensure solution is milky white and free
} of particles. Add 8.0 ml commercially prepared,
} titrated 1.0 N NaOH--from EM science--solution turns
} clear. Adjust pH strictly to 12.0+/-0.1 unit.
} Bring volume to 50ml with CO2-free double distilled
} water. Stopper tightly with rubber stopper and
} parafilm until use later that day.
}
} When we stain the grids with lead nitrate, we make
} sure to wash well before and after with CO2 free
} double distilled water in addition to surrounding
} the staining plate (Hiraoka kit) with NaOH pellets.
} In addition, we centrifuge the Reynolds at 5000xg
} for 8 minutes prior to use AND we 0.2um filter it
} into staining plate.
}
} ANY advice or thoughts are welcome...what are we
} doing wrong?? What can we change about this
} protocol to ensure ppt free staining?
}
} A thousand thanks in advance!
}
} Danielle Crippen
} Morphology and Imaging Core Manager
} Buck Institute for Age Research
} 8001 Redwood Blvd.
} Novato, CA 94945
} 415-209-2046
} dcrippen-at-buckinstitute.org
}
}
}
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Doesn't anybody else use or recommend Sato's lead stain as a more
stable replacement for Reynolds Pb citrate? We've used it since the
1970s.
1968 Sato, T.: J. Electron Microsc. 17:158, 1968.
1968 Sato and others: Proc. XIth Int. Cong. on Electron Microscopy.
Kyoto. 1986, pp. 2181-2182. [

-mike reedy-

At 5:49 PM -0500 5/2/06, dcrippen-at-buckinstitute.org wrote:
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************************
Michael K. Reedy, M.D.
Duke Univ. Med. Center
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Durham, NC 27710

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Dear Danielle,
dear Listers,

since it was not an issue in the replies so far, I would like to ask wether
your double distilling apparatus overall is made of quartz glass or does
have a destillate } container { bin made from metal (e. g. copper).

I only would like to add this since we had - several years ago - a problem
when our destillation apparatus was out of function and on repair for some
month and we used } bidistilled { water obtained from our hospital pharmacy.
We had a lot of precipitation problems then, which ended not before we
changed to the ddH2O from the repaired quartz-glass still used formerly.

When checking the quality of the "pharmacy"-water later on it turned out to
contain a high amount of copper-ions (storage bin was made from copper
sheets), which IMO perhaps might have had a detrimental precipitating
action on the lead-staining performance.

By the way: we use Lead Citrate according to Venable&Coggeshall (1965),
store solutions in "ultraclean" snap cap-glass-vials [and also ultraclean
plastic snap cap!] which are used only for that purpose, that means we take
care of any traces of cleaning substances by washing /cleaning also with
chrome-sulfuric acid or a modern substitute but take care by ourselves (not
a washing machine) to get rid of any resting traces of substances by
vigorously washing several times with bidistilled hot water and a final
step with ultrapure water (UHQ). We found also that intermittent air drying
of glass vial/bottle creates probably otherwise insoluble incrustations, so
we always keep the stuff in wet condition until the final step of cleaning.

Another point we found is that "freshly" made lead citrate solution
(Venable&Coggeshall) -if used the same day - will be "more aggressive/more
reactive", that means, we decrease staining times (say 30 sec when freshly
prepared instead of 2-3 min -at-room temperature, e.g. after one week storage
in the dark).

Avoiding or at least some sort of control for the CO2-reaction is
obligatory in our lab (NaOH-pellets in a petridish filled with dental wax,
the latter always being melted and } flamed/singed { after a staining cycle,
but perhaps use of virgin, clean? { {Parafilm} } sheets is the better choice)
knowing that some/severely disturbing precipitation nuclei also could be
present in previousely uncleaned, and therefore } oily { injection needles,
syringes, (plastic) tips, rubber stoppers (especially if always one and the
same is used) as well as the surface areas where you are staining/handling
your grids.

In general our experience is / was: the more steps you are introducing in
your schedule to reduce an anticipated precipitate (or to inhibit the
formation of such one) the more you (likely) will initiate precipitation
due to unexpected particle impurities.

All best wishes for an excellent result of your next staining series,

Wolfgang Muss
Salzburg Austria
----------
Von: dcrippen-at-buckinstitute.org[SMTP:dcrippen-at-buckinstitute.org]
Antwort an: dcrippen-at-buckinstitute.org
Gesendet: Mittwoch, 03. Mai 2006 00:50
An: W.Muss-at-salk.at
Betreff: [Microscopy] TEM--Lead Citrate--HELP!!

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Dear TEM users,

This is a specimen prep question for everyone out there with EM expertise.
We are having terrible success with Lead Citrate contrast staining.
Principally, we suffer from precipitates showing up all over the specimen.
On the advice of EM science technical support, we are double distilling
our own water (they say Milli-Q is too pure and also is de-ionized which we
don't want for EM). Then we make it CO2 free by autoclaving and capping
directly upon removal from the autoclave. We do this the morning of
reagent prep--so it doesn't sit in the bottle for longer than it takes to
cool down before we begin making up the Reynolds.

We make the Reynolds Lead citrate according to the protocol listed in Ch 5
of Bozzola and Russell's Electron Microscopy (2nd edition) (mix 1.33g lead
nitrate, 1.76g sodium citrate, and 30ml CO2 free double distilled
water...shake vigorously for a few minutes and then again 5-6 times over
the next 30 minutes). Ensure solution is milky white and free of
particles. Add 8.0 ml commercially prepared, titrated 1.0 N NaOH--from EM
science--solution turns clear. Adjust pH strictly to 12.0+/-0.1 unit.
Bring volume to 50ml with CO2-free double distilled water. Stopper
tightly with rubber stopper and parafilm until use later that day.

When we stain the grids with lead nitrate, we make sure to wash well before
and after with CO2 free double distilled water in addition to surrounding
the staining plate (Hiraoka kit) with NaOH pellets. In addition, we
centrifuge the Reynolds at 5000xg for 8 minutes prior to use AND we 0.2um
filter it into staining plate.

ANY advice or thoughts are welcome...what are we doing wrong?? What can we
change about this protocol to ensure ppt free staining?

A thousand thanks in advance!

Danielle Crippen
Morphology and Imaging Core Manager
Buck Institute for Age Research
8001 Redwood Blvd.
Novato, CA 94945
415-209-2046
dcrippen-at-buckinstitute.org

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24, 28 -- From W.Muss-at-salk.at Wed May 3 03:00:12 2006
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Hi folks,
Strictly this isn't a microscopy question but I think I have a
fair chance of catching someone who might be able to help!

I was looking through our boxes of 'historical' samples (there has been
a materials analysis lab here for more than 50 years), and I have
examples of 1", 2", 3", 4", 6" and 8" Si wafers. Since we stopped
getting Si samples to analyse about 5 years ago I don't have any 12" (or
300mm, I should say) wafers. Is there anyone out there willing to swap
a 12" wafer for a 1" one? I know it doesn't seem like good value for
the amount of material but I hope that would be more than compensated
for by the historical interest. Or I could swap odd bits of Ge, GaAs,
InP, sapphire, etc. etc. if Si isn't your bag. Perhaps I should start
up a little 'Caswell museum'..

Many thanks

Richard

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

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} This may be shear luck, but I've never had trouble with
} precipitate (other things yes, but ppte no) - I keep my lead
} citrate (made up with ordinary distilled water, not specially
} CO2 free) in a volumetric flask (50ml), which sits in the
} same place month after month and is never moved. I don't use
} the stain for 24 hours after it's prepared, but then just
} carefully take off what's needed from close to the surface
} using a glass pipette. I wipe the end of the pipette with a
} tissue before dispensing the stain and then discard the first
} drop. I put the drops onto Parafilm in a covered glass petri
} dish. No need for NaOH pellets. Finally wash the grids for 5
} seconds in a gentle stream of water from a wash bottle. I
} probably shouldn't admit this but I've had a bottle of stain
} last over 2 years (the surface of the bottle becomes cloudy
} with ppte) and still produce perfect results.
}
} Cheers,
}
} Diana

I use similar protocol, but I do use DI-} distilled-} boiled water
and I do put NaOH pellets in the petry dish. Never tried to keep
stain for 2 years, but for 6 month it works fine.

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Manager
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

Phone: (816) 235-2072
Fax: (816) 235-5524
Web: http://www.umkc.edu/dentistry/microscopy

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I have been using "calcined lead citrate" (apparently a modification of
Sato's lead citrate) for a couple of years, and it certainly is much
more stable than traditional Reynold's lead citrate.
Takamasa Hanaichi et al. (1986) A Stable Lead by Modification of
Sato's Method. J. Electron Microsc., Vol. 35. No. 3. 304-306.
--Jan Factor

---------------------------------------
Jan Robert Factor, Ph.D.
Professor of Biology
---------------------------------------
Natural Sciences
Purchase College, State University of New York
735 Anderson Hill Rd.
Purchase, NY 10577
USA
---------------------------------------
Office Tel: 914-251-6659
Office Fax: 914-251-6635
E-mail: jfactor-at-ns.purchase.edu
or- jan.factor-at-purchase.edu
---------------------------------------

mike.reedy-at-cellbio.duke.edu wrote:
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--|
--| Doesn't anybody else use or recommend Sato's lead stain as a more
--| stable replacement for Reynolds Pb citrate? We've used it since the
--| 1970s.
--| 1968 Sato, T.: J. Electron Microsc. 17:158, 1968.
--| 1968 Sato and others: Proc. XIth Int. Cong. on Electron Microscopy.
--| Kyoto. 1986, pp. 2181-2182. [
--|
--| -mike reedy-
--|
--| At 5:49 PM -0500 5/2/06, dcrippen-at-buckinstitute.org wrote:
--|
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--|--|
--|--| Dear TEM users,
--|--|
--|--| This is a specimen prep question for everyone out there with EM
--|--| expertise. We are having terrible success with Lead Citrate
--|--| contrast staining. Principally, we suffer from precipitates showing
--|--| up all over the specimen. On the advice of EM science technical
--|--| support, we are double distilling our own water (they say Milli-Q is
--|--| too pure and also is de-ionized which we don't want for EM). Then
--|--| we make it CO2 free by autoclaving and capping directly upon removal
--|--|
--| --|from the autoclave. We do this the morning of reagent prep--so it
--|
--|--| doesn't sit in the bottle for longer than it takes to cool down
--|--| before we begin making up the Reynolds.
--|--|
--|--| We make the Reynolds Lead citrate according to the protocol listed
--|--| in Ch 5 of Bozzola and Russell's Electron Microscopy (2nd edition)
--|--| (mix 1.33g lead nitrate, 1.76g sodium citrate, and 30ml CO2 free
--|--| double distilled water...shake vigorously for a few miutes and then
--|--| again 5-6 times over the next 30 minutes). Ensure solution is milky
--|--| white and free of particles. Add 8.0 ml commercially prepared,
--|--| titrated 1.0 N NaOH--from EM science--solution turns clear. Adjust
--|--| pH strictly to 12.0+/-0.1 unit. Bring volume to 50ml with CO2-free
--|--| double distilled water. Stopper tightly with rubber stopper and
--|--| parafilm until use later that day.
--|--|
--|--| When we stain the grids with lead nitrate, we make sure to wash well
--|--| before and after with CO2 free double distilled water in addition to
--|--| surrounding the staining plate (Hiraoka kit) with NaOH pellets. In
--|--| addition, we centrifuge the Reynolds at 5000xg for 8 minutes prior
--|--| to use AND we 0.2um filter it into staining plate.
--|--|
--|--| ANY advice or thoughts are welcome...what are we doing wrong?? What
--|--| can we change about this protocol to ensure ppt free staining?
--|--|
--|--| A thousand thanks in advance!
--|--|
--|--| Danielle Crippen
--|--| Morphology and Imaging Core Manager
--|--| Buck Institute for Age Research
--|--| 8001 Redwood Blvd.
--|--| Novato, CA 94945
--|--| 415-209-2046
--|--| dcrippen-at-buckinstitute.org
--|--|
--|--|

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I've had a couple of inquiries about calcined lead citrate, so I
thought I'd send this to the list in case anyone else is interested. (To
be fair, I learned about this method by perusing the EMS Catalog, which
has the formulation.) I've pasted the relevant page from my in-house lab
manual (below). To prepare the calcined the lead citrate, the unusual
step, I simply went up to our chemistry program and asked them to fire
up their high-temp oven (which is in a fume hood) for the day. Once you
get a successful batch of calcined lead citrate, and I suggest making a
good deal more than you need immediately, it can be stored as a powder
in a vial for some time (perhaps indefinitely?). This way, you only have
to bake it once, and you can make enough for multiple batches of lead
citrate stain. I still use the usual precautions when handling lead
stain, such as using NaOH pellets, and I spin down the stain in a
table-top centrifuge before each use.
Hope this is helpful.
--Jan

---------------------------------------
Jan Robert Factor, Ph.D.
Professor of Biology
---------------------------------------
Natural Sciences
Purchase College
State University of New York
735 Anderson Hill Rd.
Purchase, NY 10577
USA
---------------------------------------
Office Tel: 914-251-6659
Office Fax: 914-251-6635
E-mail: jfactor-at-ns.purchase.edu
or- jan.factor-at-purchase.edu
---------------------------------------

Calcined lead citrate (Hanaichi et al., 1986)

Calcined lead citrate is a stable, non-precipitating replacement for
Reynolds’ lead citrate, which is reportedly free from precipitates for
over one year when kept at room temperature.
Takamasa Hanaichi et al. (1986) A Stable Lead by Modification of Sato's
Method. J. Electron Microsc., Vol. 35. No. 3. 304-306.
Calcined lead citrate: Heat crystal lead citrate for several hours in a
melting pot (200-300̊C) until the color changes to a light brownish
yellow. This takes ~6.5 hrs at 250̊C. Note: Check the color periodically,
as overheated lead citrate with a dark brownish or black color can't be
used. The calcined lead citrate can be stored and used for repeated
batches of stain.

The stock lead solution:
1. The following reagents are placed in a 50 ml volumetric flask and
mixed well to produce a yellowish milky solution:
CALCINED LEAD CITRATE........0.20 g Prepared from lead citrate
Lead nitrate......................................0.15 g
Lead acetate....................................0.15 g EMS #17600-25, 25g
Sodium citrate.................................1.00 g
Distilled water................................ 41.00 ml

2. Then, add: 1.0N NaOH.....................................9.0 ml
Carbonate free, EMS #21170-01, 225ml
Then 9.0 ml of 1N NaOH (use carbonate free solution) is added to the
solution and mixed well until the solution becomes clear with a light
yellowish color. The solution is then transferred to an amber glass with
screw cap bottle for storage, and can be stored at room temperature or
in the refrigerator (recommended) for over 1 year.

-

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Hi, All-

I have a client who would like to do some confocal on GFP-transformed
Arabidopsis (no problem) and then some TEM immunolabeling. What do I need
to know about buying an anti-GFP antibody conjugated to gold? Any tricks,
or is it straightforward? Does anyone have a favorite vendor? Any tips
gratefully accepted!

Aloha,
Tina

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

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Dear listers,

I am observing the internalization of a fluorescent
component in the cells. I thought it would be a good
idea to quench the extracellular fluorescence after
several hours of incubation. This way I would see only
the fluorescence coming from inside the cells.
Would you know a substance which quenches Alexa488
dyes and which is not toxic to the cells (and do not
enter the cells)?
St�phane-without-an-i

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Tina,

I am not aware of anti-GFP gold conjugate (although it may be somewhere
out there), but you can use the indirect method, i.e. primary
antibody/secondary conjugate (protein A/gold or secondary
antibody/gold). For the primary, we use Torrey Pines Biolabs purified
rabbit Anti-GFP, that works well even after 2% formaldehyde/0.2%
glutaraldehyde (that is what we use for Tokuyashu's technique).

Hope this helps,

Michal

tina-at-pbrc.hawaii.edu wrote:
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}
} I have a client who would like to do some confocal on GFP-transformed
} Arabidopsis (no problem) and then some TEM immunolabeling. What do I need
} to know about buying an anti-GFP antibody conjugated to gold? Any tricks,
} or is it straightforward? Does anyone have a favorite vendor? Any tips
} gratefully accepted!
}
} Aloha,
} Tina
}
} ****************************************************************************
} * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
} * Biological Electron Microscope Facility * (808) 956-6251 *
} * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
} ****************************************************************************
}
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Email: rpowell-at-nanoprobes.com
Name: Rick Powell

Organization: Nanoprobes, Incorporated

Title-Subject: [Filtered] Re: [Microscopy] Anti-GFP for TEM?

Question: Hello Tina and Everyone:

We don't have a gold probe for this (yet), and can't comment on the best antibodies, but the question comes up occasionally, and we keep an eye out for references. We have reported on several that might be helpful in back issues of our newsletter:

http://www.nanoprobes.com/Newsletter_Archive.html

(1) Follet�Gueye and co-workers have reported an improved fixation method for the immunogold localization of green fluorescent protein (GFP). The method is a simple one-step procedure which consists of fixing in the dark for 30 min, 60 min, or 120 min at 4�C in a solution consisting of 1% glutaraldehyde and 1% osmium tetroxide in 0.1 M Na cacodylate buffer, pH 7.2. This mixture was prepared and used immediately. After washing in distilled water, the samples were gradually dehydrated in 10%, 20%, and 40% aqueous ethanol (10 min in each bath), then in 60% and 80% (20 min in each bath), and finally in anhydrous ethanol for 30 min, and embedded in LR White or Spurr resin. GFP-tagged Golgi glycosyltransferase was localized in transgenic BY-2 cells using rabbit anti-GFP primary and 10 nm gold anti-rabbit secondary antibodies; this method allowed improved structural preservation of the endomembrane system, and anti-GFP antibodies bound with high specificity, allowing the localization of GFP-tagged glycosyltransferases within individual Golgi cisternae.

Reference:

Follet-Gueye, M. L.; Pagny, S.; Faye, L.; Gomord, V., and Driouich, A.: An improved chemical fixation method suitable for immunogold localization of green fluorescent protein in the Golgi apparatus of tobacco Bright Yellow (BY-2) cells. J. Histochem. Cytochem., 51, 931-940 (2003).

Abstract (courtesy of the Journal of Histochemistry and Cytochemistry):
http://www.jhc.org/cgi/content/abstract/51/7/931

Reprint (courtesy of the Journal of Histochemistry and Cytochemistry):
http://www.jhc.org/cgi/reprint/51/7/931

(2) Luby-Phelps and co-workers describe a procedure for labeling green fluorescent protein (GFP)-expressing cells in 1-micron zebrafish sections for fluorescence and electron microscope observation using a 10 nm gold-labeled secondary antibody. GFP fluorescence survived fixation in 4% paraformaldehyde/0.1% glutaraldehyde and can be visualized directly by fluorescence microscopy. For electron microscopy, thin sections of silver�gold color were collected on nickel grids and were incubated in anti-GFP antibody at a 1:25 dilution for 2 hr at room temperature, followed by 10-nm immunogold-conjugated goat anti-rabbit IgG for 1 hr. After contrasting with 3% aqueous uranyl acetate solution, the grids were dried and viewed with a Hitachi 600 transmission electron microscope operated at 75 kV.

Reference:

Luby�Phelps, K.; Ning, G.; Fogerty, J., and Besharse, J. C.: Visualization of Identified GFP-expressing Cells by Light and Electron Microscopy. J. Histochem. Cytochem., 51, 271-274 (2003).

Abstract (courtesy of the Journal of Histochemistry and Cytochemistry):
http://www.jhc.org/cgi/content/abstract/51/3/271

Reprint (courtesy of the Journal of Histochemistry and Cytochemistry):
http://www.jhc.org/cgi/reprint/51/3/271

(3) Joanne Buchanan and co-workers investigated immuno-EM of GFP with Nanogold� secondary antibody conjugates, and described some results at Microscopy & Microanalysis 2002: use of microwave processing greatly shortened all the steps.

Reference:

Buchanan, J.; Micheva, K. D., and Smith, S. J.; Microwave Processing and Pre-embedding Nanogold Immunolabeling for Electron Microscopy. Microsc. Microanal., 8, (Suppl. 2: Proceedings); Lyman, C. E.; Albrecht, R. M.; Carter, C. B.; Dravid, V. P.; Herman, B., and Schatten, H. (Eds.); Cambridge University Press, New York, NY, 2002, 160.

Prior and co-workers prepared their own 5nm gold anti-GFP:

Prior, I. A.; Muncke, C.; Parton, R. G., and Hancock J. F.: Direct visualization of Ras proteins in spatially distinct cell surface microdomains. J. Cell. Biol., 160, 165-170 (2003).

Reprint (Journal of Cell Science):
http://www.jcb.org/cgi/reprint/160/2/165

Hope some of this is helpful,

Rick Powell

********************************************************
Richard D. Powell
rpowell-at-nanoprobes.com * www.nanoprobes.com
NANOPROBES, Incorporated
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At 11:57 PM 5/3/2006, you wrote:

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

Hi, All-

I have a client who would like to do some confocal on GFP-transformed
Arabidopsis (no problem) and then some TEM immunolabeling. What do I need
to know about buying an anti-GFP antibody conjugated to gold? Any tricks,
or is it straightforward? Does anyone have a favorite vendor? Any tips
gratefully accepted!

Aloha,
Tina

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

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Hi Tina,
I have not had very good luck with localizing GFP for on TEM sectios.
Hope you will share any good suggestions that you get

Greg

tina-at-pbrc.hawaii.edu wrote:
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} Hi, All-
}
} I have a client who would like to do some confocal on GFP-transformed
} Arabidopsis (no problem) and then some TEM immunolabeling. What do I need
} to know about buying an anti-GFP antibody conjugated to gold? Any tricks,
} or is it straightforward? Does anyone have a favorite vendor? Any tips
} gratefully accepted!
}
} Aloha,
} Tina
}
} ****************************************************************************
} * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
} * Biological Electron Microscope Facility * (808) 956-6251 *
} * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
} ****************************************************************************
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--
Gregory W. Erdos, Ph.D.
Assistant Director, Biotechnology Program
Scientific Director, EM Core Lab
P.O. Box 118525
University of Florida
Gainesville, FL 32611
gwe-at-ufl.edu
Phone: 352-392-1295
Fax: 352-846-0251

--
Gregory W. Erdos, Ph.D.
Assistant Director, Biotechnology Program
Scientific Director, EM Core Lab
P.O. Box 118525
University of Florida
Gainesville, FL 32611
gwe-at-ufl.edu
Phone: 352-392-1295
Fax: 352-846-0251

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Dear All,
I am looking for a flange to mount a KEVEX EDS detector on my Philips 525
SEM (conical lens).
Or for a drawing to help manufacturing the flange.

Thanks,
Stefan Diller

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Stefan Diller - Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49 - 931 - 7848700 Phone
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++49 - 175 - 717 70 51 Cell-Phone
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Hi Tina. We published a detailed protocol : Micheva, Buchanan, Hotz
and Smith Nature Neuroscience 2003 Vol 6 #9 p925-32.
I have had some good results with anti GFP antibodies from MBL and
Roche for TEM. We used Fluro-Nanogold secondaries and silver enhanced
and looked at hippocampal neurons in culture, Drosophila salivary
glands and mouse brain -all pre-embedding labeling.
I don't have any experience with aradadopsis.
Good luck let me know if you need more info.
JoAnn Buchanan

Department of Molecular and Cellular Physiology
Stanford University School of Medicine
Stanford, CA 94305
650-723-5856

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I am looking for a method to identify Type IIB skeletal muscle fibers at
the electron microscopy level in human muscle biopsies. There are well
established techniques for light microscopy, typically detecting ATPase,
but I need good fixation and embedding for immunoelectron microscopy. A
reasonable fixation compromise is 2% formaldehyde and 0.1% --0.2%
glutaraldehyde with embedding in LR White/Gold or one of the Lowicryl
resins. Any suggestions? Any experience with a particular antibody for
human type IIB fibers?

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

larry.ackerman-at-ucsf.edu

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Email: staffan-at-physto.se
Name: Staffan Wachtmeister

Organization: Stockholm University, Stockholm, Sweden

Title-Subject: [Filtered] TEM EELS; any old/used spectrometer available?

Question: Hello everyone!

I have a question that I would like to ask and Gerald Kothleitner at Graz recommended this forum to me, i.e. this is my first posting here.

At Stockholm University, we are buying some new TEM and spectrometer equipment. In the purchase process, however, we have an extra microscope. My idea is, if possible, to buy an additional spectrometer, a used one, for example GIF or PEELS to use together with our extra microscope (our old JEOL 3010 LaB6 instrument) so that young scientists can get more access to the instruments as well. So my question is, if you have, or if you know someone who has an old GIF or PEELS or similar that is not or will not be needed anymore at your lab due to for example an upgrade, or if is not needed anymore for any reason. Then "I" would be very interested in somehow purchasing this old/used spectrometer system for an affordable price (because it is not part of our current budget). I am asking this question as a PhD student, but I will happily forward your proposals or give you the contact information to the people who are making the decisions at my university.

With kind regards

Staffan Wachtmeister

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Dear Ferri,
We have the same instrument and I have seen the
problem you are experiencing. We have found it is
usually a no more than a specimen clamping problem.You
must make sure that the clamping mechanism is
correctly located over the grid hole. We always use a
pointed wooden spill to gently push it into place ,
you will hear a 'click' as it
locates. It could be that the holder and clamping
mechanism just need cleaning so that a good contact
can be made.. This is not a unique
problem to the H7600 in any TEM if the grid is not
firmly clamped this will happen.
Hope this solves it for you.
Regards

Christine.

Christine Richardson
Experimental Officer
School of Biological and Biomedical Science
Centre for Molecular Imaging
University of Durham
Science site
South Rd
Durham
England
DH1 3LE
Tel: 0191 3341285\3341321
Fax:0191 3341201
E-mail: a.c.richardson-at-dur.ac.uk

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Hi, All-

Thanks to all who responded with tips. We will try an indirect labeling
process, and I will fix more lightly than usual. If we are dramatically
successful, I'll post our procedure!

Mahalo,
Tina

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

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Listers,

Here is the May 2006 Microscopy Today table of contents. I will close
the subscription list for this issue on Thursday May 11, 2006.

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

Ron Anderson, Editor
=======================================================

Teaching Old Microscopes New Tricks
Stephen W. Carmichael, Mayo Clinic

VisBio: a Flexible Open-Source Visualization Package for
Multidimensional Image Data
Curtis T. Rueden and Kevin W. Eliceiri, U. Wisconsin-Madison

BioImageXD � New Open Source Free Software for the Processing, Analysis
and Visualization of Multidimensional Microscopic Images
P. Kankaanp��1, K. Pahajoki1, V. Marjom�ki2, J. Heino2, D. White2
1University Turku, 2University Jyv�skyl�, Finland

Novel Developments in High-Frequency Micro-Ultrasound Imaging
Tom Little, VisualSonics Inc., Toronto, Ontario, Canada

Precise SEM Cross Section Polishing via Argon Beam Milling
N. Erdman, R. Campbell, and S. Asahina,* JEOL USA Inc., Peabody,
Massachusetts, *JEOL Ltd., Japan

Perfusion Fixation of Research Animals
C. W. Scouten,* R. O�Connor,** & M. Cunningham** *MyNeuroLab.com, St
Louis, MO, ** Harvard U., Belmont, MA

Three-Dimensional Crystallographic Analysis Beyond EBSD Mapping: The
Next Dimension
J.J.L. Mulders*, and A. Gholinia**, * FEI, Eindhoven, The Netherlands,
** HKL Technology, Hobro, Denmark

Improved Cutting of One-Micron Plastic Sections using Qwick Glass Knives
J. T. McMahon and J. Alsouss, Cleveland Clinic Foundation, Ohio

The Scanning of Colour and B&W Film and Photographs for Image
Processing, Analysis and Archiving - On a Tight Budget
Keith J. Morris, The Institute of Ophthalmology, UCL, UK

A Comment on AFM vs. Replicas for High Resolution Imaging
Don Chernoff, Advanced Surface Microscopy, Inc.

Embedding Cultured Cells Grown in Well Plates
Leona Cohen-Gould, Cornell University, Ithaca, NY

Flies in a Box
P. S. Connelly & L. Ruggiero,* *Oregon Health and Science U. Portland, OR

A Comment on using FLIM with FRET
Karl Garsha, Roper Scientific, Tucson, AZ

Microscopy for Children
Carolyn Schooley, MSA Project MICRO

New and Interesting at PITTCON & Industry News

NetNotes Topics

--LM - floaters

--SAMPLE PREPARATION - viral particles

--SAMPLE PREPARATION � cell culture preparation

--SAMPLE PREPARATION - propylene oxide

--SAMPLE PREPARATION - MgO preparation and Fe oxidation

--MICROTOMY � cleaning grids

--MICROTOMY - section thickness

--IMMUNOCYTOCHEMISTRY - nanogold in semithin sections

--EM - microscope cooling lines

--EM � operating voltage

--TEM - Replicas

--TEM - carbon post-coating

--EDS - Low Z peak pileup

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I am new to the listserve and was hoping somebody could give me some advise
and/or guidance. I am looking for the best way to embed, cut, and adhere to
microscope slides wool/hair fibers. I have tried nitrocellulose/paraffin,
Spurr's Resin, Epon, and LR white. I am having an issue with the fibers popping
out of the resin during sectioning and/or during the sample processing. I have
tried cutting the resin dry and wet - floating off into water. I have tried
infiltrating with acetone/resin and ethanol/resin mixtures with the Spurr's
resin, acetone/resin infiltration with the Epon and LR white. I seem to get the
best cuts with LR white (5 microns) dry but then I have a problem adhering them
to a slide for processing. To improve sample adhesion, I have tried albumin, gelatin, and neoprene. Of the three, neoprene seems to work the best. If anybody can offer advice for any or all of these
issues, I would greatly appreciate it.

Thank you,
Felicia Dixon

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Microscopy can be used for investigation? Really?

If I recall my history of optics properly, people in the Middle Ages knew
precisely what things looked like at the microscopical level but just
couldn't draw accurate pictures of them. So they made the technological
decision to develop the microscope so they could show each other what they
already knew God had created. Really, it wasn't investigation, it was just
confirmation of what they already knew was there. Thus, they were able to
illustrate that ontogeny really does recapitulate philogeny. They
"illustrated", not "investigated".

I hope we've settled this issue.

-Michael the OHR (Optics Historian of Record)

At 03:22 PM 04/20/06 -0500, you wrote:

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____________________________________________________________________________
Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med.
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What!? There are several ideas in there that are hard for me to swallow.
Of course, I know that my ability to swallow alone does not determine
the truth of concept.

I am not so familiar with the early history of microscopy and the
thinking surrounding it. (I would not think of challenging you for the
position of OHR.) I doubt there was a decision made to develop
microscopy to only verify what was suspected of being there. Their
preconceptions might be considered hypotheses in need of verification,
but we certainly get surprised enough about hypotheses on a macro scale.
Microscopy would hardly be more of a sure thing.

I thought that microscopy would have developed more out of a desire to
explore. I can guess what might be out there on a microscopic scale, but
until I actually go and take a look, it is only conjecture - no matter
how accurate the conjecture might be.

Maybe I am misunderstanding Mr. Cammer's point on Ontogeny
recapitulating phylogeny. Are you saying that _those_ folks used
microscopy to "prove" it? Perhaps they "proved" it to themselves. Or
are you saying you still accept the idea yourself? If so, my
understanding is that the concept has been discredited for some time. I
am not a great fan of Stephen Gould but I found the following comment in
an Amazon review of his book, _Ontogeny and Phylogeny_.

"Ontogeny recapitulates phylogeny" was Haeckel's answer--the wrong
one--to the most vexing question of nineteenth-century biology: what is
the relationship between individual development (ontogeny) and the
evolution of species and lineages (phylogeny)? In this, the first major
book on the subject in fifty years, Stephen Gould documents the history
of the idea of recapitulation from its first appearance among the
pre-Socratics to its fall in the early twentieth century. -Ernst Mayr.

FWIW, I consider microscopy an important tool for investigation.
However, it must be used carefully as many of our observations are quite
few. We need to pay attention to the statistics if we are going to
generalize.

Warren Straszheim
Iowa State University

-----Original Message-----
X-from: cammer-at-aecom.yu.edu [mailto:cammer-at-aecom.yu.edu]
Sent: Monday, May 08, 2006 10:26 AM
To: wesaia-at-iastate.edu

Microscopists,
Please please forgive me for leaving out the word "only"
in my original post. I meant to write ..."it is worth remembering
that microscopy can be used for demonstration not ONLY investigation.

I was simply trying to point out that it is reasonable to
demand one kind of thing when we are investigating but also have room
for another purpose, namely the demonstration, for which demands are
different. In no way shape or form was I meaning to suggest that
microscopy cannot be used to investigate. Indeed, that suggestion is
ludicrous.

I did send an apology after my post and it seems once is not
enough. So once more, I am sorry to have wasted time and bandwith
with my failure to proofread.

As ever,
Tobias

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I agree that EM or LM can and are used for investigation.
They are also used for discovery. In the context of IC
reverse engineering and technology evaluation, discovery
and investigation are biggies. For failure analysis, I
suspect that investigation fits the bill better than
does discovery--unless investigation leads to the discovery
of the failure mechanism.

IMO, I suspect that the early scientists and researchers
had a clue that microbes existed but did not know what they
looked like. Along comes the microscope. Now they could get
a good idea of what they looked like and confirm that they
existed. Is this investigation, discovery or both?

gary g.

At 10:23 AM 5/8/2006, you wrote:

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Sorry Mike,

You're too clever by half. I assume that "philogeny" was not a
misspelling.

Now, where did I hear that ontology recapitulates phylogeny?

Joel

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} Microscopy can be used for investigation? Really?
}
} If I recall my history of optics properly, people in the Middle Ages
} knew precisely what things looked like at the microscopical level but
} just couldn't draw accurate pictures of them. So they made the
} technological decision to develop the microscope so they could show
} each other what they already knew God had created. Really, it wasn't
} investigation, it was just confirmation of what they already knew was
} there. Thus, they were able to illustrate that ontogeny really does
} recapitulate philogeny. They "illustrated", not "investigated".
}
} I hope we've settled this issue.
}
} -Michael the OHR (Optics Historian of Record)
}
}
} At 03:22 PM 04/20/06 -0500, you wrote:
}
}
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Joel B. Sheffield, Ph.D.
Biology Department, Temple University
1900 North 12th Street
Philadelphia, PA 19122
jbs-at-temple.edu
(215) 204 8839, fax (215) 204 0486
http://astro.temple.edu/~jbs

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Email: dotys-at-hss.edu
Name: Steve Doty

Organization: Hospital for Special Surgery

Title-Subject: [Filtered] Billing for core services.

Question: As director of a microscopy core, I am also responsible for sending bills/invoices for services provided by core. We have external as well as internal users. We provide TEM, SEM, LM, morphometric analysis, paraffin embedding and methacrylate embedding and sectioning, immuno and enzyme histochemistry. And we have a mix of clinical and basic research to accomodate. My problem is trying to keep the user and techniques together for billing purposes since any particular technique may take several days and different technician's support. Does anyone have a "system" that will help me keep track of everything that is happening around me!
Many thanks, Steve Doty, PhD.

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Email: crnl_srbu-at-yahoo.com
Name: Corneliu Sarbu

Organization: Natl.Inst.for Materials Physics, Bucharest, Romania

Title-Subject: [Filtered] Zr70Ni10Pd20 metalic glasses - how can they be acid dissolved

Question: Dear colleagues,

Is there anybody who could give me a hint concerning the acid or acid mixture that would be effective in surface etching a piece of ternary metallic glass having the composition Zr70Ni10Pd20 ? I need to reveal the dimensions of grains which are most probably formed in the material after a short aannealing during which an icosahedral quasicrystalline phase was formed (it was revealed by X-ray diffraction). Any suggestion will be very welcome.

Thank you in advance.

Dr. Corneliu Sarbu
National Institute for Materials Physics
Magurele-Bucharest
Romania

e-mail: crnl_srbu-at-yahoo.com

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Title-Subject: [Filtered] Porter - Blum Microtome JB-4

Question: Does anyone have an operators manual for the
SORVALL Porter - Blum JB-4 Microtome please?
Thank you
Dainis

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Although we are an academic lab in materials, our problems with
billing are very similar. The data, of course, has to feed into the
corporate financial management system (SAP, in our case).

After a long period of wondering what to do, and a false start with
an almost institute-wide system that promised to be totally
comprehensive but was in fact far too ambitious for our resources, we
are currently developing an in-house system which will enable us to
track usage by instrument, user, date, project, etc. etc., as well,
of course, as providing the data for our billing.

It has a MySQL heart, with Visual Basic and Filemaker Pro front-ends
(for historical reasons). The data collection part of the system is
up and running, and in use for most of our systems. All the
instruments should be using it by the end of next month. Then we
have to develop the interface to the billing system, and the analysis
tools (currently we extract the data from the database by manual SQL
calls and generate a text file).

Tony

At 08:40 PM 5/8/2006, you wrote:
Email: dotys-at-hss.edu
Name: Steve Doty

Organization: Hospital for Special Surgery

Title-Subject: [Filtered] Billing for core services.

Question: As director of a microscopy core, I am also responsible for
sending bills/invoices for services provided by core. We have
external as well as internal users. We provide TEM, SEM, LM,
morphometric analysis, paraffin embedding and methacrylate embedding
and sectioning, immuno and enzyme histochemistry. And we have a mix
of clinical and basic research to accomodate. My problem is trying
to keep the user and techniques together for billing purposes since
any particular technique may take several days and different
technician's support. Does anyone have a "system" that will help me
keep track of everything that is happening around me!
Many thanks, Steve Doty, PhD.

***********************************************
Anthony J. Garratt-Reed, M.A., D.Phil.
MIT Room 13-1027
77 Massachusetts Avenue
Cambridge, MA 02139-4307
USA

Tel: 617-253-4622
Fax: 617-258-6478
************************************************

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Good day, Corneliu,
Considering Ti as chemically similar to Zr, I found some etchant
solutions for Ti alloys in Metals Handbook from ASM. That these might
work for Zr alloys is supported by comments in Cotton and Wilkinson, Adv.
Inorg. Chem. Anyway, here are a couple that ASM suggests as general purpose
etchants:

10 ml HF, 5 ml HNO3, 85ml water
1-3 ml HF, 2-6 ml HNO3, water to 1000 ml

You can also try contacting ATI Wah Chang in Oregon, USA. They've been
producing Zr alloys for decades and have plenty of expertise in that area.
phone: 1-541-926-4211
www.wahchang.com

Good luck, and BE CAREFUL if you use HF.

Rob Bowen

--
Robert C. Bowen
Research Scientist
Caddock Electronics, Inc
rob.bowen-at-caddock.com
http://www.caddock.com

} From: {crnl_srbu-at-yahoo.com}
} Reply-To: {crnl_srbu-at-yahoo.com}
} Date: Mon, 8 May 2006 19:37:44 -0500
} To: {rob.bowen-at-caddock.com}
} Subject: [Microscopy] viaWWW: Zr70Ni10Pd20 metalic glasses - how can they be
} acid
}
}
}
}
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} Email: crnl_srbu-at-yahoo.com
} Name: Corneliu Sarbu
}
} Organization: Natl.Inst.for Materials Physics, Bucharest, Romania
}
} Title-Subject: [Filtered] Zr70Ni10Pd20 metalic glasses - how can they be acid
} dissolved
}
} Question: Dear colleagues,
}
} Is there anybody who could give me a hint concerning the acid or acid mixture
} that would be effective in surface etching a piece of ternary metallic glass
} having the composition Zr70Ni10Pd20 ? I need to reveal the dimensions of
} grains which are most probably formed in the material after a short aannealing
} during which an icosahedral quasicrystalline phase was formed (it was revealed
} by X-ray diffraction). Any suggestion will be very welcome.
}
} Thank you in advance.
}
} Dr. Corneliu Sarbu
} National Institute for Materials Physics
} Magurele-Bucharest
} Romania
}
} e-mail: crnl_srbu-at-yahoo.com
}
}
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} 11, 13 -- From zaluzec-at-microscopy.com Mon May 8 19:32:38 2006
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==============================Original Headers==============================
11, 20 -- From Rob.Bowen-at-caddock.com Tue May 9 10:39:16 2006
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11, 20 -- they be acid
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Hi,
Might any of you know if polyurethane (Festo blue PU) tubing
is compatible with P-10 gas. I currently have blue PU 3/16" tubing
plumbed from the P-10 tank regulator to the first gfpc WDS on my
uProbe. Is anyone 100% sure that this type of tubing should not be
used to delivered P-10 gas to gas floow proportional counter
detectors?

TIA
Mike
--
********************************************************************
Michael M. Cheatham
312 Heroy Geology Laboratory Phone (315)-443-1261
Syracuse University Fax (315)-443-3363
Syracuse, NY 13244-1070

email:mmcheath-at-mailbox.syr.edu
http://www.geochemistry.syr.edu/cheatham/InstrPages.html

owner of PLASMACHEM-L: http://listserv.syr.edu/archives/plasmachem-l.html
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Folks

StereoScan260 up for grabs.
Contact Gary Halada at:
ghalada-at-ms.cc.sunysb.edu

I shall not be acting as a "go between".

You pack.......you ship....looking for good home.

regards,

JQuinn

==============================Original Headers==============================
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TECHNICAL OFFICER

CSIRO Plant Industry
Black Mountain, Canberra
Australia

Reference: 2006/411

We require an experienced microscopist with demonstrated knowledge of plant
structure and developmental biology to assist in the Microscopy Centre in
the Division of Plant Industry. Ideally, the appointee would have skills in
preparation of plant material for light and electron microscopy, especially
in cryo-scanning electron microscopy and x-ray microanalysis. The
Microscopy Centre currently has light, fluorescence and confocal
microscopes, a cryo-SEM with EDX, image analysis software and other
ancilliary equipment.

The successful applicant will assist the Manager in training other staff in
use of the instruments and in microscopy techniques, and in carrying out
microscopy work for specific research projects. He/she will have at least a
Bachelor's degree or equivalent training in plant structure and functional
plant anatomy and experience in working in a research laboratory.
Additional skills required are the ability to collaborate effectively with
scientists and members of a research team and strong communication and
computer skills.

This position is indefinite after a probationary period, salary $44K - $57K
p.a plus Superannuation .
To obtain selection documentation or details on how to apply visit
http://recruitment.csiro.au/asp/job_details.asp?RefNo=2006%2F411. For
further information contact Dr Rosemary White - rosemary.white-at-csiro.au .
Responses to the selection criteria accompanied by a CV, must be received by
close of business 4 June 2006.

Unfortunately, the powers-that-be have decided it's open to Australian
residents only, largely so they don't have to fly people in for interviews.
However, if you're interested, or know anyone who might be, contact me
anyway.

cheers,
Rosemary

Dr. Rosemary White rosemary.white-at-csiro.au
CSIRO Plant Industry ph. 02-6246 5475
GPO Box 1600 mob. 0402 835 973
Canberra, ACT 2601 fax. 02-6246 5334
Australia

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Hi,

We are considering buying EDX system integrated with EBSD for
elemental and "structural" analysis of inorganic materials in SEM. I hope
therefore that vendors of such equipment will contact me off the list.
However, I would be grateful also for any comments from listers on the
possibilities of EBSD as the method of structure identification of
individual nanocrystals in composite materials.

Thank you,

Leszek

Leszek Kepinski
Institute of Low Temperature and Structure Research,
Polish Academy of Sciences,
P.O.Box 1410,
50-950 Wroclaw, Poland
e-mail: L.Kepinski-at-int.pan.wroc.pl

==============================Original Headers==============================
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Hello,
I'm using an older Pelco CPD 2 critical point drier. Anyone use this? It
has these gnurled steel knobs for opening/closing the valves. Controlling
the valves kills your fingers since the rough metal is hard on your skin.
Anyone modified it with plastic knobs or rubber covers or something to
make it less of a literal pain to do a critical point drying run?

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\
Gordon Ante Vrdoljak Electron Microscope Lab
AOL/IM rakhasha http://nature.berkeley.edu/~gvrdolja 26 Giannini Hall
gvrdolja-at-nature.berkeley.edu UC Berkeley
phone (510) 642-2085 Berkeley CA 94720-3330
fax (510) 643-6207 cell (510) 290-6793

==============================Original Headers==============================
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When we had one of those we used pliers to turn the knobs.

gvrdolja-at-nature.berkeley.edu wrote:
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} Hello,
} I'm using an older Pelco CPD 2 critical point drier. Anyone use this? It
} has these gnurled steel knobs for opening/closing the valves. Controlling
} the valves kills your fingers since the rough metal is hard on your skin.
} Anyone modified it with plastic knobs or rubber covers or something to
} make it less of a literal pain to do a critical point drying run?
}
} \/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\
} Gordon Ante Vrdoljak Electron Microscope Lab
} AOL/IM rakhasha http://nature.berkeley.edu/~gvrdolja 26 Giannini Hall
} gvrdolja-at-nature.berkeley.edu UC Berkeley
} phone (510) 642-2085 Berkeley CA 94720-3330
} fax (510) 643-6207 cell (510) 290-6793
}
} ==============================Original Headers==============================
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--
Greg Erdos
5410 SE 185th Ave.
Micanopy, FL 32667

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Contents Retrieved from Microscopy Listserver Archives
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Sending to list--direct msgs bounce. If you
have a better address, please advise and I will
send any other material off-list.

-------

What SEM do you plan on using? What type of
gun does it use?

What feature sizes are you interested in resolving?
Ta and Cu seed layers are not possible to resolve
even at .01u step size. I think that this is because
no discernable grains have formed as yet. The electrodep Cu for
damascene interconnects works well.

I've done single crystal Sapphire, Silicon and micro
crystal Si. What would your nano crystals be made of?
If the material is non-conductive, this may pose a
charging issue. Since EBSD penetrates only about 50nm,
this does not leave much depth for coating. The
preferred coating is C. However, I rarely coat any
insulating material even at 20KV.

There are basically two EBSD choices. TSL/EDAX or
HKL/PGT. TSL and EDAX have been integrated for much
longer than HKL and PGT (I think HKL connected just this
year). TSL's camera/phosphor is round while HKL's camera end
is square. This allows the HKL camera screen to get
closer to the specimen. But I think that the TSL
software complement is much better. AFAIK, only TSL
has drift correction during data collection. At high
mag and small step size, this is critical. However TSL's
drift correction is not always repeatable and its own
set of problems. But at least it is there and does
usually work.

The SEM is going to be at issue too since higher frames
per second need more probe current. However, this is
at the expense of probe diameter and lattice resolution.

I'm using EDAX/TSL Pegasus with EDAX Genesis EDS and like it.
The SEM is a Zeiss Supra 55VP, which has its own set
of issues.

gary g.

At 03:34 AM 5/10/2006, you wrote:

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==============================Original Headers==============================
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Hi Gordon - we have that same unit and have well developed calluses!
Two simple tricks to keep your skin more or less intact:
- I never adjust the coarse vent valve. Permanently leave it
halfway open and control venting with the needle valve.
- When going through the fill-drain cycles - open the fill valve
one-half turn or so and leave it there until you're finished. Then you
just open the drain valve to let fluid out, and close it to let fluid
in. This cuts the number of knob-turning operations down by half. On
our unit, the fill valve is the hardest to operate so this trick is very
helpful.

I'd avoid pliers unless you absolutely can't get the knobs to turn.
Pliers can strip the knurled texture and just make things harder in the
future. And too much force can damage the valves. I do have one user
that uses vise-grip pliers on the fill valve, but she is under stern
warnings to pad the knob with cloth and not use force to close the valve.

Rick

gvrdolja-at-nature.berkeley.edu wrote:
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}
} Hello,
} I'm using an older Pelco CPD 2 critical point drier. Anyone use this? It
} has these gnurled steel knobs for opening/closing the valves. Controlling
} the valves kills your fingers since the rough metal is hard on your skin.
} Anyone modified it with plastic knobs or rubber covers or something to
} make it less of a literal pain to do a critical point drying run?
}
} \/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\
} Gordon Ante Vrdoljak Electron Microscope Lab
} AOL/IM rakhasha http://nature.berkeley.edu/~gvrdolja 26 Giannini Hall
} gvrdolja-at-nature.berkeley.edu UC Berkeley
} phone (510) 642-2085 Berkeley CA 94720-3330
} fax (510) 643-6207 cell (510) 290-6793
}
}

--
{} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {}
Richard C. Hugo, Ph.D
Geomicrobiology and Electron Microscopy Laboratory
Portland State University
Ph# 503-725-3356
FAX 503-725-3025
{} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {}

==============================Original Headers==============================
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All this talk of forcing needle valves and pliers, etc. on CPD systems
is scaring me. There used to be a homemade CPD in the Berkeley Microlab
which sounds very similar to the unit currently being discussed. It was
retired in favor of a Tousimis unit chiefly because of safety concerns.
The pressures and explosive-release volumes on these systems are
sufficient to cause serious injury in the event of a failed valve or
fitting. Trust me - It's Not Worth It. If you find you are using hand
tools to adjust needle valves on these systems, you DO have a safety
problem.

-------- Original Message --------

Right, it is HKL+Oxford now. Thanks for the correction.

TSL uses a Digiview 1612 Firewire camera that does
a good job.

Depending on the grain size you are examining, probe
diameter will be critical. Too large and small grains
will be missed. Probe current increases frames per
second--which is good.

TSL will do multi-phase EBSD. Their expansion of this
is their Delphi option. I do not have this. I figure
it is more for those trying to discover new things than
sorting out what is basically known but not quantified.
The plain OIM system comes with the TSL database and
accepts the AMCS database....a very huge set of materials
indeed.

EDAX EDS has drift correction (maps). This is the same basic
shell used by TSL. Without correction, long scans at
high mag are IMO useless and impossible. These are easy
to spot since the resulting scans are wavy rather than
consistent. Coating with C pretty much eliminates that
element from analysis and also reduces signal by about
20-30%. High Z coating is not in the cards--too much
absorption to produce diffraction patterns. VP is
workable. Since one is not doing EDS maps, drift correction
during EBSD collection is the key. EDAX/TSL also offers
off-line dataset analysis for EBSD like they do for EDS.
I'm not sure if HKL has the same option. I would ask
about this when shopping.

gary g.

At 10:11 AM 5/11/2006, you wrote:
} Gary,
}
} We're also considering a new EBSD system for an old LaB6 JEOL 840 that
} delivers lots of beam current. It's to replace an old TSL system with a
} deteriorated SIT camera. We're interested in the phase ID capabilities as
} well as grain orientation analysis.
}
} Your comments on drift correction are especially interesting. I don't know
} that a PGT/HKL connection exists any more. (Is PGT still in business?) HKL
} is now part of Oxford, and they have at least partially integrated the EBSD
} operation with Oxford's EDS software (INCA). INCA does has drift
} correction, but I don't believe the combined system does yet.
}
} Larry
}
}
} Larry Thomas
} Pacific Northwest National Laboratory
} Richland, WA 99352
}
} email: Larry.Thomas-at-pnl.gov
} phone: 509 372-0793
} --
}
}
}
}
} On 5/11/06 9:23 AM, "gary-at-gaugler.com" {gary-at-gaugler.com} wrote:
}
} }
} }
} }
} }
} ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} }
} ----------------------------------------------------------------------------
} }
} } Sending to list--direct msgs bounce. If you
} } have a better address, please advise and I will
} } send any other material off-list.
} }
} } -------
} }
} } What SEM do you plan on using? What type of
} } gun does it use?
} }
} } What feature sizes are you interested in resolving?
} } Ta and Cu seed layers are not possible to resolve
} } even at .01u step size. I think that this is because
} } no discernable grains have formed as yet. The electrodep Cu for
} } damascene interconnects works well.
} }
} } I've done single crystal Sapphire, Silicon and micro
} } crystal Si. What would your nano crystals be made of?
} } If the material is non-conductive, this may pose a
} } charging issue. Since EBSD penetrates only about 50nm,
} } this does not leave much depth for coating. The
} } preferred coating is C. However, I rarely coat any
} } insulating material even at 20KV.
} }
} } There are basically two EBSD choices. TSL/EDAX or
} } HKL/PGT. TSL and EDAX have been integrated for much
} } longer than HKL and PGT (I think HKL connected just this
} } year). TSL's camera/phosphor is round while HKL's camera end
} } is square. This allows the HKL camera screen to get
} } closer to the specimen. But I think that the TSL
} } software complement is much better. AFAIK, only TSL
} } has drift correction during data collection. At high
} } mag and small step size, this is critical. However TSL's
} } drift correction is not always repeatable and its own
} } set of problems. But at least it is there and does
} } usually work.
} }
} } The SEM is going to be at issue too since higher frames
} } per second need more probe current. However, this is
} } at the expense of probe diameter and lattice resolution.
} }
} } I'm using EDAX/TSL Pegasus with EDAX Genesis EDS and like it.
} } The SEM is a Zeiss Supra 55VP, which has its own set
} } of issues.
} }
} } gary g.
} }
} }
} } At 03:34 AM 5/10/2006, you wrote:
} }
} }
} }
} } }
} ----------------------------------------------------------------------------
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } } To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} } } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } }
} ----------------------------------------------------------------------------
} } }
} } } Hi,
} } }
} } } We are considering buying EDX system integrated with EBSD for
} } } elemental and "structural" analysis of inorganic materials in SEM. I hope
} } } therefore that vendors of such equipment will contact me off the list.
} } } However, I would be grateful also for any comments from listers on the
} } } possibilities of EBSD as the method of structure identification of
} } } individual nanocrystals in composite materials.
} } }
} } } Thank you,
} } }
} } }
} } }
} } } Leszek
} } }
} } }
} } }
} } }
} } }
} } } Leszek Kepinski
} } } Institute of Low Temperature and Structure Research,
} } } Polish Academy of Sciences,
} } } P.O.Box 1410,
} } } 50-950 Wroclaw, Poland
} } } e-mail: L.Kepinski-at-int.pan.wroc.pl
} } }
} } }
} } }
} } } ==============================Original
} Headers==============================
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10, 21 -- To: "Thomas, Larry (PNNL)" {Larry.Thomas-at-pnl.gov}
10, 21 -- From: Gary Gaugler {gary-at-gaugler.com}
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The first question I would ask:
WHY are the knobs hard to turn?

Generally it's a couple reasons.

#1 If the threads are lubricated and the grease/oil and enough of the
VOCs (even if they aren't very volatile) dissipate the lubricant can
turn to glue.
-Solution:
Disassemble, clean (with WD-40, CRC or other solvating
lubricants), and re-apply some lightweight grease (lithium or wheel
bearing would be fine), or a touch of heavy oil.

#2 If the threads/valve are damaged, you probably should replace the
valve.

These all are extremely simple devices. And as such parts are
relatively easy to repair/replace or fix. Assuming you have the
patience to source the proper valves. The pressures the CPDs the
microscopy community utilizes are relatively insignificant to some of
the devices folks in Physics and engineering play with daily, not to
mention, McMaster-Carr has a great selection of valves and what not that
are rated well above the typical 1500-3000 psi burst limit (safety) on
the CPDs.

If the valve was always hard to turn that's one thing, if it is now,
make it easy to turn.

Try even spraying the threads with some WD-40 or liquid Wrench and run a
cycle or two and see if that helps anything.

I am a big fan of the Polaron type CPD. I've never been let down by
one, and it is the best model for 'teaching' the principles... no black
boxes and although manual, no sane minded individual should start a CPD
run and let it go un-attended... so why not be manual? My apologies to
those who sell automatic ones, this is just my opinion.

Geoff Williams
Leduc Bioimaging Facility Manager
Brown University

http://www.brown.edu/Facilities/Leduc_Bioimaging_Facility/

-----Original Message-----
X-from: gvrdolja-at-nature.berkeley.edu [mailto:gvrdolja-at-nature.berkeley.edu]

Sent: Wednesday, May 10, 2006 7:10 PM
To: Williams, Geoffrey

Hello,
I'm using an older Pelco CPD 2 critical point drier. Anyone use this?
It
has these gnurled steel knobs for opening/closing the valves.
Controlling
the valves kills your fingers since the rough metal is hard on your
skin.
Anyone modified it with plastic knobs or rubber covers or something to
make it less of a literal pain to do a critical point drying run?

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/
\/\/\
Gordon Ante Vrdoljak Electron
Microscope Lab
AOL/IM rakhasha http://nature.berkeley.edu/~gvrdolja 26 Giannini
Hall
gvrdolja-at-nature.berkeley.edu UC Berkeley
phone (510) 642-2085 Berkeley CA
94720-3330
fax (510) 643-6207 cell (510) 290-6793

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Colleagues,

I am writing to get information on web-pages which have compiled links to bio-imaging and microscopy facilities both within the US and internationally.

I have searched and located a few such web-sites, but they are not very comprehensive and many of the links are dead.

If anyone in the community may have web-addresses or information on sites of this nature, I would appreciate their input.

Thank you,

David Lowry
School of Life Sciences
Arizona State University
Tempe, AZ 85287-4501
office: 480-727-0725
lab: 480-965-2463

==============================Original Headers==============================
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This Question was submitted to Ask-A-Microscopist by (cheetham.3-at-osu.edu)
from http://www.microscopy.org/Ask-A-Microscopist/Ask-A-Microscopist.html on Thursday, May 11, 2006 at 16:42:15
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Email: cheetham.3-at-osu.edu
Name: Sonia Cheetham

Organization: The Ohio State University

Education: Graduate College

Location: Wooster , Ohio

Title: question on photomicrophotography

Question: I have being trying to find out what photomicrophotography is about. I came across this term in a paper regarding confirmation of PMO getting into the cells. Can you give me a brief exlanation of what is this technique? Thanks

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Try this one. http://www.magma.ca/~scimat/

Ann Fook Yang
EM Unit/ Unite EM
AAFC/AAC
960 Carling Ave,
Ottawa,Ontario
Canada K1A 0C6
yanga-at-agr.gc.ca
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada

-----Original Message-----
X-from: dlowry-at-asu.edu [mailto:dlowry-at-asu.edu]
Sent: Thursday, May 11, 2006 4:18 PM
To: Yang, Ann-Fook

Colleagues,

I am writing to get information on web-pages which have compiled links to bio-imaging and microscopy facilities both within the US and internationally.

I have searched and located a few such web-sites, but they are not very comprehensive and many of the links are dead.

If anyone in the community may have web-addresses or information on sites of this nature, I would appreciate their input.

Thank you,

David Lowry
School of Life Sciences
Arizona State University
Tempe, AZ 85287-4501
office: 480-727-0725
lab: 480-965-2463

==============================Original Headers==============================
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Email: nyilmaz-at-mersin.edu.tr
Name: nejat yilmaz

Organization: mersin university medical school

Title-Subject: [Filtered] Cutting latex for TEM

Question: Hello Everybody...

We're trying to investigate bacteria biofilms on a latex material (like surgical gloves) with TEM. We couldn't cut the latex material with our regular glass knives. Is there any suggestion about this problem?

Thanks for any comment...

Dr. Necat Yilmaz
Mersin University Medical School
Histology & Embryolgy Dept.

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Dear Listers:

We are looking for a lab to do some contract work, embedding,
sectioning and H&E staining of zebrafish using JB4/GMA plastics. We will
provide fixed specimens. Ultimately we would like serial sections but
for now we just want to see what sort of results we can expect. Please
contact me directly. Nothe that I will be out of the office next week
(15th-19th) so my response may be delayed.
Thanks.

Geoff

--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583; fax: -4029
mcauliff-at-umdnj.edu
**********************************************

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At the risk of putting my foot in my oversized mouth - - - You need a
cryo-microtome and work around -40. This gets you below the glass
transistion temperature for most elastomers and the latex will no longer
behave like an elastic material, but a hard brittle glass. You may need to
fiddle with temperature and pick-up technique. I've sectioned polymer and
and used both glycerin/water, mineral spirits/xylene and DMSO/water
depending on the temperature and my end goal (Light, SEM or TEM). A
diamond knife and boat would be my prefered method. I like to pick up with
a "perfect loop" and place on carbon coated grid.

good luck and have fun........

Frank Karl
Degussa Corporation
Akron Technical Center
3500 Embassy Parkway
Suite 100
Akron, Ohio 44333

330-668-2235 Ext. 238

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nyilmaz-at-mersin.ed
u.tr To: frank.karl-at-degussa.com
cc:
05/12/2006 08:44 Subject: [Microscopy] viaWWW: Cutting latex for TEM
AM
Please respond to
nyilmaz

Title-Subject: [Filtered] Cutting latex for TEM

Question: Hello Everybody...

We're trying to investigate bacteria biofilms on a latex material (like
surgical gloves) with TEM. We couldn't cut the latex material with our
regular glass knives. Is there any suggestion about this problem?

Thanks for any comment...

Dr. Necat Yilmaz
Mersin University Medical School
Histology & Embryolgy Dept.

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Dear Colleagues...

Thanks for your kindly helps about our latex cutting problem.
Best regards...

Dr. Necat Y�lmaz
Mersin �niversitesi T�p Fak�ltesi
Histoloji ve Embriyoloji Anabilim Dal�

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using the WWW based Form at
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Email: AMCGroup2-at-aol.com
Name: James Glossinger

Organization: AMC Group

Title-Subject: TEM Contractor

We are currently seeking an independent TEM analyst, academic/non-profit TEM facility or, commercial TEM lab for contracting ongoing advanced TEM imaging and AEM projects, involving semiconductor materials and devices.

If interested and qualified, please contact me off-line via email.

Thanks,
Jim

---------------------
James Glossinger, Ph.D.
Principal Scientist
AMC Group

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

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Email: palladineus-at-yahoo.com
Name: Randy Khoo

Title-Subject: [Filtered] Masson-Fontana staining for melanin

Question: I am wondering if anybody can help me out on a poser that I have.

Melanin is able to reduce the ammoniacal silver nitrate solution without the addition of an external reducing agent. However, as I understand it, melanin exists in 2 forms, both in oxidized form and reduced form. So, is the reduced form only responsible for reducing the silver nitrate? If so, can the addition of a reducing agent result in both forms reducing the silver nitrate and show a better staining profile? Can this method be used to semi-quantify both the reduced and oxidized forms?

Thanks in advance.

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

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I have a project where I need to positively identify human cells that
may have mouse cells mixed with them in a tumor. Does anyone have a
suggestion for a protocol that will positively identify a cell as mouse
or as human. Antibodies that I have tried so far have not worked,
post-embedding.

Thanks, Greg
--
Gregory W. Erdos, Ph.D.
Assistant Director, Biotechnology Program
Scientific Director, EM Core Lab
P.O. Box 118525
University of Florida
Gainesville, FL 32611
gwe-at-ufl.edu
Phone: 352-392-1295
Fax: 352-846-0251

==============================Original Headers==============================
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Ah... I'm disappointed. I thought we were going to have a nice
discussion about the transition from human adjustments (knobs) to using
the mouse (bar sliders)....

I was already to pull out the link to
http://www.griffintechnology.com/products/powermate/
... suggesting that maybe it is the answer we old school knob
aficionados have been waiting for.

Alas I have no suggestions relating to the topic. So please accept or
excuse this mild attempt at being funny on a Monday morning.

Geoff Williams
Leduc Bioimaging Facility Manager
Brown University

http://www.brown.edu/Facilities/Leduc_Bioimaging_Facility/

-----Original Message-----
X-from: gwe-at-ufl.edu [mailto:gwe-at-ufl.edu]
Sent: Monday, May 15, 2006 9:22 AM
To: Williams, Geoffrey

I have a project where I need to positively identify human cells that
may have mouse cells mixed with them in a tumor. Does anyone have a
suggestion for a protocol that will positively identify a cell as mouse
or as human. Antibodies that I have tried so far have not worked,
post-embedding.

Thanks, Greg
--
Gregory W. Erdos, Ph.D.
Assistant Director, Biotechnology Program
Scientific Director, EM Core Lab
P.O. Box 118525
University of Florida
Gainesville, FL 32611
gwe-at-ufl.edu
Phone: 352-392-1295
Fax: 352-846-0251

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Does anyone where to find one of the spatial calibration slides that used to
be sold by a company called Richardson Technologies? The company appears to
be gone but I'm trying to get hold of one the slides.

Thank you!

--David Hitrys
QImaging Corporation

==============================Original Headers==============================
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Hi, Dave

Try EMS. They were selling Richardson's products.

Best regards,
Barbara Foster

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com

MME is now scheduling customized, on-site courses through September. Call us today for details.

P. S.
Need a good general reference or light microscopy text for the Spring semester? Call us today to learn more about "Optimizing LIght Microscopy". Copies still available through MME... even for class-room lots ... and we give quantity discounts. Call Ken Piel at (972)954-8011.

At 08:42 AM 5/15/2006, dhitrys-at-qimaging.com wrote:

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

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ListServer Subscribers:

In response to a recent question on the ListServer which, in part, asked about the status of PGT Microanalysis, I would like to reply:

On Nov 17, 2005 Bruker AXS announced the acquisition of Princeton Gamma-Tech Microanalysis and R�ntec AG. These two organizations were merged to form Bruker AXS Microanalysis. Bruker AXS develops and manufactures a broad range of analytical X-ray systems including XRF, XRD and now, X-ray Microanalysis.

Customer service and support for both PGT Microanalysis and R�ntec have been expanded as part of the Bruker AXS worldwide operation. PGT and R�ntec customers should contact Bruker AXS for continued product and applications support.

Bruker AXS Microanalysis is a proud Sustaining Member of MSA and MAS, carrying on the long time support given by PGT Microanalysis and R�ntec to these fine professional organizations.

Doug Skinner
Assistant Vice President
Bruker-AXS Microanalysis
609-771-4400
Doug.Skinner-at-Bruker-AXS.com
www.bruker-axs-ma.com

==============================Original Headers==============================
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Listers,

Does anybody have any experience with this digital microscope camera?
I don't see on the webpage I found that the chip is cooled.
How opinions about the image quality? Reliability? The price is attractive.
Thanks.

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

==============================Original Headers==============================
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Email: lewryan-at-gmail.com
Name: Ryan

Organization: Dalhousie University

Title-Subject: [Filtered] SEM imaging on sample with magnetic substrate

Question: I have a saple with deposits of an alloy of Sn-Co-C on a magnetic stainless steel substrate (430 stainless steel). I'm using a cold field emission microscope and am having trouble getting high resolution images. Could you suggest what setting I should use to get started?

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No experience, but it looks good, except that on the polaroid website:

http://www.polaroid.com/global/detail.jsp;jsessionid=Eo1VHNEfv4zBiHQO811vhkPjhU2i
9bsgrpjdBKtwD1ZlyI25lSFe!191862641!-1979950386!7005!8005!1446422789!-
1979950377!7005!8005?PRODUCT%3C%3Eprd_id=845524441760112&FOLDER%3
C%3Efolder_id=2534374302028681&bmUID=1147731925565&bmLocale=en_US

(honest, that's the page URL, I got it from Googling polaroid dmc2)

is says that it's no longer manufactured.

seems a pity

cheers

rtch

On 15 May 2006 at 16:16, oshel1pe-at-cmich.edu wrote:

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} Listers,
}
} Does anybody have any experience with this digital microscope camera?
} I don't see on the webpage I found that the chip is cooled.
} How opinions about the image quality? Reliability? The price is attractive.
} Thanks.
}
} Phil
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} Philip Oshel
} Microscopy Facility Supervisor
} Department of Biology
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Ritchie Sims Ph D Phone : 64 9 3737599 ext 87713
Microanalyst Fax : 64 9 3737435
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Dear Greg,
If you could culture and clone these cells. do
karyotyping.
shashi

--- gwe-at-ufl.edu wrote:

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} I have a project where I need to positively identify
} human cells that
} may have mouse cells mixed with them in a tumor.
} Does anyone have a
} suggestion for a protocol that will positively
} identify a cell as mouse
} or as human. Antibodies that I have tried so far
} have not worked,
} post-embedding.
}
} Thanks, Greg
} --
} Gregory W. Erdos, Ph.D.
} Assistant Director, Biotechnology Program
} Scientific Director, EM Core Lab
} P.O. Box 118525
} University of Florida
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} gwe-at-ufl.edu
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Hi!

If your purpose is just to identify mouse cells in a
human culture, you don't need EM. Actually it would be
much easier in LM. Any way, you could think about
using HLA marker, they are external (it's just an idea
I have no experience with that).

regards,

Stephane

--- gwe-at-ufl.edu wrote:

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} I have a project where I need to positively identify
} human cells that
} may have mouse cells mixed with them in a tumor.
} Does anyone have a
} suggestion for a protocol that will positively
} identify a cell as mouse
} or as human. Antibodies that I have tried so far
} have not worked,
} post-embedding.
}
} Thanks, Greg
} --
} Gregory W. Erdos, Ph.D.
} Assistant Director, Biotechnology Program
} Scientific Director, EM Core Lab
} P.O. Box 118525
} University of Florida
} Gainesville, FL 32611
} gwe-at-ufl.edu
} Phone: 352-392-1295
} Fax: 352-846-0251
}
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9, 20 -- From nizets2-at-yahoo.com Tue May 16 03:56:59 2006
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We are considering Fovea Pro as a set of quantitation plug-ins for
Photoshop. Can I ask those who use FP, and who also teach IA, to contact me
off list? I'd like to get an idea of the user base, and accumulate any
thoughts on re-establishing an FP forum.

Genuinely, Michael Shaffer :o)

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
c/o Memorial University
230 Elizabeth Avenue
P.O. Box 4200
St. John's, NL A1C 5S7

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Several years ago many (most?) of the EM supply vendors switched the
formula for carbon tabs to a "stiff", non-flexible formula. I was
told at that time that the old formula used typewriter tape as a
component.

These new stiff tabs are unsuitable for many of the preps we do. I
was able to find the old formula at Fullam and now they have switched
too.

Does anyone know where to find truly flexible carbon tabs. The ones
Fullam had were from Nisshin EM, very expensive but quite high quality.

Thanks in advance.

Owen Mills
MTU, Houghton MI

==============================Original Headers==============================
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5, 31 -- From: "Owen P. Mills" {opmills-at-mtu.edu}
5, 31 -- Subject: dbl stick carbon tab blues
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http://www.pelcoint.com/technote_html/Conductive%20Tab%20Form%20Letter.p
df

Have you looked at Ted Pella's assortment (Pelco)?
Some answers to your questions may be found in his letter above.
Best wishes,

Jim

-----Original Message-----
X-from: opmills-at-mtu.edu [mailto:opmills-at-mtu.edu]
Sent: Tuesday, May 16, 2006 2:56 PM
To: James Chalcroft

Several years ago many (most?) of the EM supply vendors switched the
formula for carbon tabs to a "stiff", non-flexible formula. I was
told at that time that the old formula used typewriter tape as a
component.

These new stiff tabs are unsuitable for many of the preps we do. I
was able to find the old formula at Fullam and now they have switched
too.

Does anyone know where to find truly flexible carbon tabs. The ones
Fullam had were from Nisshin EM, very expensive but quite high quality.

Thanks in advance.

Owen Mills
MTU, Houghton MI

==============================Original Headers==============================
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Conductive double sided carbon adhesive tape is more flexible and could perhaps solve your problem. This tape can be obtained from several vendors.

Best regards,
J�rgen.

= = = = = = = = = = = = = = = = =

Joergen B. Bilde-Soerensen
Senior Research Scientist, Ph. D.
Materials Research Department
Risoe National Laboratory
DK-4000 Roskilde
Denmark

e-mail: j.bilde-at-risoe.dk
phone: +45 4677 5802 (direct)
phone: +45 4677 4677 (switchboard)
fax: +45 4677 5758
website: http://www.risoe.dk/afm/Personal/jqbi/jqbi.htm

-----Original Message-----
X-from: opmills-at-mtu.edu [mailto:opmills-at-mtu.edu]
Sent: Tuesday, May 16, 2006 2:50 PM
To: j.bilde-at-risoe.dk

Several years ago many (most?) of the EM supply vendors switched the
formula for carbon tabs to a "stiff", non-flexible formula. I was
told at that time that the old formula used typewriter tape as a
component.

These new stiff tabs are unsuitable for many of the preps we do. I
was able to find the old formula at Fullam and now they have switched
too.

Does anyone know where to find truly flexible carbon tabs. The ones
Fullam had were from Nisshin EM, very expensive but quite high quality.

Thanks in advance.

Owen Mills
MTU, Houghton MI

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Hi Ryan

I suppose that you have not only a cold FEG SEM, but a so called
"semi-in-lens" type of OL on it too. In fact the cold or schottky type
of FEG doesn't play a role in the problem. It's only a OL question. That
type of OL has a strong magnetic field coming out of it, which enveloppe
the sample at short working distance, alouding to work virtuusely at WD0
and to get nice high res images of ..... non magnetic materials, with
the "trough the lens" SE detector. The field coming out of the OL can be
very strong ; I have measured values such as 3kG at WD2 and 10keV
primary energie. With lower beam energy the field decreases and at 3keV,
I measured a value of 1.2kG at WD3 mm, and 0.3kG at WD8 mm.

Pratically, you have a few solutions, but all are half solutions.

First, you must avoid to work with short WD. Look at the shortes WD
possible without too much astigmatisme. Ask the SEM manufacturer at
which WD the sample is quite out of the field. If your shortest WD is
2mm or so, it will bee something like 6-8mm.

Second, you must find a primary energie not to low, which would give
a poor resolution, due to the perturbation of the primary beam by the
field lignes in the sample, and not too high, to minimase the filed
coming out of the OL, wich increases with increasing primary energy. In
most cases, you 'll find a good compromise between 5 and 10 keV.

Third, you must play very very much with the astigmatism
corrections, at replay again if you move your sample a little bit. And
depending of the electronic, you may touch the range limits of the
astigmatism corrections. In that case, if you have a CL astigmatism
correction setting, you can play on it a little bit, puting
astigmatisme at the Cl, in the opposit direction, to gain some margin of
the OL astigmatism settings. It's not a clean way to work, but in may help.

Fourth, take the smalest sample possible, not too thick, not too
wide, and fix it very securly on the holder. It may land from it and
stick to the OL !

Fifth solutions : buy an other SEM for that kind of samples !!!

Don't wait as well resolved images than with ..... gold particles on
silicon. If you have a nice picture at x50000, you can be satisfied !
More depends on your particular situation, sample and SEM.

Hope it helps

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

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

lewryan-at-gmail.com a �crit :

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Dear List

I have a faculty member at UIC who is looking for a one off image of a
purified protein. She has seen this done by rotary shadowing and while we
have the instrumentation at the university we do not have the necessary
experience. As it is a one off experiment (she needs an image showing to
confirm if there is bend in the middle of a mutant) I was wondering if
there was a lab out there that regularly uses rotary shadowing who could
help us?

Alan

Alan W Nicholls, PhD
Director of Research Service Facility (Electron Microscopy)
Research Resources Center - East (M/C 337)
Room 100 Science and Engineering South Building
The University of Illinois at Chicago
845 West Taylor St
Chicago, IL 60607-7058

Tel: 312 996 1227
Fax: 312 996 8091
Office: Room 110

Web site www.rrc.uic.edu

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Hi, Allen

We have used the AFM to image DNA conformation and think that it might also work for your protein, without all the elaborate preparation.

If you are interested in having us give it a try, contact Dr. Kim Kangasniemi (just call him "Kim") at (972)954-8014 or kim-at-nt-america.com

Hope this was helpful,

Best regards,
Barbara Foster

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com

MME is now scheduling customized, on-site courses through September. Call us today for details.

P. S.
Need a good general reference or light microscopy text for the Spring semester? Call us today to learn more about "Optimizing LIght Microscopy". Copies still available through MME... even for class-room lots ... and we give quantity discounts. Call Ken Piel at (972)954-8011.

At 08:54 AM 5/16/2006, nicholls-at-uic.edu wrote:

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Check the MSA list archives for sticky tabs.

Ted Pella has good ones as does EMS. No experience
with SPI tabs.

gary g.

At 05:51 AM 5/16/2006, you wrote:

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We have been doing our pre-embedding immunogold localizations
incubations for animal tissue in PBS, 0.2% BSA and 10mM Na Azide.

I am working now on some em pre-embedding localizations on plant
tissue, for which light microscopy localizations have been done using
MTSB buffer (50mM PIPES, 5mM MgSO4, 5mM EGTA, pH~7.0). Is there any
reason for which I should not be using this same MTSB buffer for the
em work?

Thank you very much for your suggestions. They will definitely save
me trial time.

Thanks.

Tea

--
***************************************
Tea Meulia, PhD
Director
Molecular and Cellular Imaging Center
Ohio State University/OARDC
1680 Madison Ave.
Wooster OH 44691

tel.: 330-263-3836 or -3828
fax: 330-202-3563

http://www.oardc.ohio-state.edu/mcic

*****************************************

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Hi,
In dealing with plant tissue, EGTA is included in buffers
because calcium chelation removes calcium cross bridges and probably
some pectin from the cell wall and allows antiboody access. Although
protocols for doing this originally called for having EGTA in the
fixation buffer, in our hands, we get better preservation if the EGTA
is included after the fixation as a separate incubation. The removal
of the calcium by the same token makes the cell wall weaker. You may
find quite distorted tissue. It may be possible to minimize this
distortion by including an incubation in mM CaCl2 after the 2nd
antibody and before dehydration.

Note that buffers with pipes, magnesium, and EGTA are not
microtubule stablizing (if that is what you mean by MTSB). They are
the standard buffers for studying microtubule dynamics in vitro.

Hope this helps.

Tobias
}
}
} We have been doing our pre-embedding immunogold localizations
} incubations for animal tissue in PBS, 0.2% BSA and 10mM Na Azide.
}
} I am working now on some em pre-embedding localizations on plant
} tissue, for which light microscopy localizations have been done using
} MTSB buffer (50mM PIPES, 5mM MgSO4, 5mM EGTA, pH~7.0). Is there any
} reason for which I should not be using this same MTSB buffer for the
} em work?
}
} Thank you very much for your suggestions. They will definitely save
} me trial time.
}
} Thanks.
}
} Tea
}
}
}
} --
} ***************************************
} Tea Meulia, PhD
} Director
} Molecular and Cellular Imaging Center
} Ohio State University/OARDC
} 1680 Madison Ave.
} Wooster OH 44691
}
} tel.: 330-263-3836 or -3828
} fax: 330-202-3563
}
} http://www.oardc.ohio-state.edu/mcic
}
} *****************************************
}
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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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Hi

I would suggest setting the specimen at a working distance of at least 15mm
so as to be outside the field of the lens. What may also help is to lower
the kV as this will also lower the lens field. Of course the kV level will
depend upon what you are asking of the specimen?

If you wish to examine the TRUE surface {5kV will be ideal. If you wish to
investigate the sub surface detail 15kV backscatter would probably be a good
starting point.

Do not try to use an upper detector if fitted tot he instrument as this will
require you being close to the lens. Lower detectors in a twin detector
system require a higher probe current (weaker C1) than that used for an
upper detector.

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 -----
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Sent: Monday, May 15, 2006 11:04 PM

I need to look at some RBC's using SEM. anyone have a favorite protocol for
preparing them? Thanks, Tom

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

573-882-4712 (office)
573-882-0123 (fax)
PhillipsT-at-missouri.edu

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Tom,

We've used poly-l-lysine cover slips and standard fixation/dehydration
procedures with good success. I believe some people have used HMDS
successfully, too.

Randy

-----Original Message-----
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Sent: Tuesday, May 16, 2006 1:55 PM
To: Tindall, Randy D.

I need to look at some RBC's using SEM. anyone have a favorite protocol
for preparing them? Thanks, Tom

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

573-882-4712 (office)
573-882-0123 (fax)
PhillipsT-at-missouri.edu

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Hi, All-

Someone wants to send me isolated, fixed rhinovirus on a grid for negative
staining and imaging. I have not worked with rhinovirus before. Does
anyone have a favorite protocol for fixation and staining? Glut or
PFA? Uranyl acetate or PTA?

Mahalo!
Tina

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

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Sorry to bring this topic up again. I know it has been discussed:

What are the additives that can go in a water recirculator for a TEM?
I use to use a product called cool-prep and someone said ethylene
glycol in the same ratio.
Any thoughts on that?

thanks,
Beth

**********************************************************************
Beth Richardson
EM Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

Phone - (706) 542-1790 & FAX - (706) 542-1805
http://www.plantbio.uga.edu/emlab

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***

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Colleagues....

For those few of us that are not in the life science arena
what is an RBC?

I'm just curious as to what the abrievations mean.

Your Friendly Neighborhood SysOp.

Nestor
--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Lab
Electron Microscopy Center
Materials Science Division/Bldg 212
9700 S. Cass Ave
Argonne, Illinois 60439 USA
Tel: 630-252-7901, Fax: 630-252-4798
Email: Zaluzec-at-aaem.amc.anl.gov
===========================================
TPMLab: http://tpm.amc.anl.gov
MMSite: http://www.amc.anl.gov
===========================================

The box said ...
"This program requires Win 95/98/NT or better..."
So I bought a Mac !

===========================================

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O.k., folks here's an odd problem:

I've got a user who wants to characterize 60-atom Carbon
Buckyballs. Now He wants to start by determining if they clump,
form larger groupsing etc. when mixed into ultra pure water (Q-
water), and then other quality waters. So, any one out there have
any idea on how to determine simple sizing? In a hydrated state?
In suspension? At potentially nanoscale? Obviously, going EM
requires drying them down - which artifically modifys the samples.
Cryo- might work but we don't have the capability here. SPM might
work but how to adhere to a surface for imaging? Diffraction of
some sort?

Any ideas?

Thanks in advance!

Richard E. Edelmann, Ph.D.
Associate Editor of EXPO, Microscopy and Microanalysis Supplement
Electron Microscopy Facility Director
364 Pearson Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.emf.muohio.edu

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Nestor, Finally a question I can answer for someone!
RBC = Red Blood Cell (~7 microns or so in size)

Not to be confused with TLA (three letter acronym) or DAP (Parents against
Dyslexia) ;o)

Paul

----------------------------------------------------------------
Though the boys throw stones at the frogs in sport, the frogs
do not die in sport, but in earnest. - Plutarch

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Beth,

I use deionized water only (no additives) and have had no problems over the
past twenty years. An important note is that we prevent the growth of
algae by plumbing all our chillers and microscopes with opaque water hose.
We never have algae, or any other, growth in our chillers.

Regards,

Gary M. Brown
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

beth-at-plantbio.
uga.edu
To
gary.m.brown-at-exxonmobil.com
05/16/06 03:24 cc
PM
Subject
[Microscopy] TEM water recirculator
Please respond additives
to
beth-at-plantbio.
uga.edu

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Sorry to bring this topic up again. I know it has been discussed:

What are the additives that can go in a water recirculator for a TEM?
I use to use a product called cool-prep and someone said ethylene
glycol in the same ratio.
Any thoughts on that?

thanks,
Beth

**********************************************************************
Beth Richardson
EM Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

Phone - (706) 542-1790 & FAX - (706) 542-1805
http://www.plantbio.uga.edu/emlab

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***

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I would suggest small-angle x-ray scattering (SAXS). Can be done
hydrated.

See, for example:

http://www.uni.aps.anl.gov/usaxs/

Jeffrey A. Fortner, Ph.D.
Chemical Engineering Division
Argonne National Laboratory
Argonne, IL 60439
fortner(at)cmt.anl.gov

-----Original Message-----
X-from: edelmare-at-muohio.edu [mailto:edelmare-at-muohio.edu]
Sent: Tuesday, May 16, 2006 3:33 PM
To: Fortner, Jeffrey A.

O.k., folks here's an odd problem:

I've got a user who wants to characterize 60-atom Carbon
Buckyballs. Now He wants to start by determining if they clump, form
larger groupsing etc. when mixed into ultra pure water (Q- water), and
then other quality waters. So, any one out there have any idea on how
to determine simple sizing? In a hydrated state?
In suspension? At potentially nanoscale? Obviously, going EM requires
drying them down - which artifically modifys the samples.
Cryo- might work but we don't have the capability here. SPM might work
but how to adhere to a surface for imaging? Diffraction of some sort?

Any ideas?

Thanks in advance!

Richard E. Edelmann, Ph.D.
Associate Editor of EXPO, Microscopy and Microanalysis Supplement
Electron Microscopy Facility Director
364 Pearson Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.emf.muohio.edu

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Nester,

RBC is the acronym for "red blood cells".

Cheers,

Gary M. Brown
ExxonMobil Chemical Company
Baytown Technology & Engineering - West
5200 Bayway Drive
Baytown, Texas 77520-2101
phone: 281 834 2387
fax: 281 834 2395
e-mail: Gary.M.Brown-at-ExxonMobil.com

zaluzec-at-aaem.a
mc.anl.gov
To
gary.m.brown-at-exxonmobil.com
05/16/06 03:31 cc
PM
Subject
[Microscopy] Hmmm... what is RBC
Please respond
to
zaluzec-at-aaem.a
mc.anl.gov

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Colleagues....

For those few of us that are not in the life science arena
what is an RBC?

I'm just curious as to what the abrievations mean.

Your Friendly Neighborhood SysOp.

Nestor
--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Lab
Electron Microscopy Center
Materials Science Division/Bldg 212
9700 S. Cass Ave
Argonne, Illinois 60439 USA
Tel: 630-252-7901, Fax: 630-252-4798
Email: Zaluzec-at-aaem.amc.anl.gov
===========================================
TPMLab: http://tpm.amc.anl.gov
MMSite: http://www.amc.anl.gov
===========================================

The box said ...
"This program requires Win 95/98/NT or better..."
So I bought a Mac !

===========================================

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Human red blood cells respond nicely to fairly routine fixatives. We use
cacodylate with Ca, Mg, and NaCl added as buffer system but biological
buffers such as PIPES should work also.

However, be aware the RBC's from other animals have different osmotic
requirements and may crenellate easily when the H-RBCs are fine. We ran
into real problems with mouse RBC's a while back. Only sure way to avoid
problems is to check and balance osmolarity of fixatives for the specific
host RBCs.

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://www3.agriculture.purdue.edu/microscopy

} From: {TindallR-at-missouri.edu}
} Reply-To: {TindallR-at-missouri.edu}
} Date: Tue, 16 May 2006 14:11:02 -0500
} To: {dsherman-at-purdue.edu}
} Subject: [Microscopy] RE: SEM of RBC's
}
}
}
}
} ----------------------------------------------------------------------------
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}
} Tom,
}
} We've used poly-l-lysine cover slips and standard fixation/dehydration
} procedures with good success. I believe some people have used HMDS
} successfully, too.
}
} Randy
}
} -----Original Message-----
} X-from: phillipst-at-missouri.edu [mailto:phillipst-at-missouri.edu]
} Sent: Tuesday, May 16, 2006 1:55 PM
} To: Tindall, Randy D.
} Subject: [Microscopy] SEM of RBC's
}
}
}
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} I need to look at some RBC's using SEM. anyone have a favorite protocol
} for preparing them? Thanks, Tom
}
}
}
} Thomas E. Phillips, PhD
} Professor of Biological Sciences
} Director, Molecular Cytology Core
} 2 Tucker Hall
} University of Missouri
} Columbia, MO 65211-7400
}
} 573-882-4712 (office)
} 573-882-0123 (fax)
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On May 16, 2006, at 1:22 PM, beth-at-plantbio.uga.edu wrote:

} Sorry to bring this topic up again. I know it has been discussed:
}
} What are the additives that can go in a water recirculator for a TEM?
} I use to use a product called cool-prep and someone said ethylene
} glycol in the same ratio.
} Any thoughts on that?
}
Dear Beth,
I don't know what is in cool-prep, but if ethylene glycol is a good
substitute, it sounds like anti-freeze. The important things for a TEM
water additive are to prevent corrosion and growth of bacteria and
algae, but the cooling water is not so cold that anti-freeze is
necessary. I have added a molybdenum-based corrosion inhibitor and the
chemical 2,2'-Methylenebis(4-chloro-phenol), AKA dichlorophene, to
inhibit growth with good effect. It is also important to keep the pH
at ~8 to prevent Cu from dissolving.
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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Hello listers,

Please suggest a sodium compound soluble in hydrocarbons such as light oil
or kerosene. Must be non-corrosive. Non-toxic preferred.

Vitaly Feingold
SIA
2773 Heath Line
Duluth GA 30096
Ph. 770-232-7785
Fax 770-232-1791
www.sia-cam.com

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Thanks, all, for the suggestions about negative staining of rhinovirus.

With one notorious exception, I have never had to fix virus before
staining, but these people want to send me fixed rhinovirus on commercial
non-glow-discharged grids (sigh). And it looks like I'll try my usual
stable of stains; UrAc, PTA, and NH4M.

Mahalo,
Tina

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

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On May 16, 2006, at 2:50 PM, Yang, Ann-Fook wrote:

} Please let us know how much you put in. Thank you.
}
Hi Ann-Fook,
I aim for 100 ppm of the Mo, as determined by a test kit, K1805, from
Taylor Technonogies, Inc. This is somewhat complicated due to not
knowing how much water is in the Haskris and how much is in the tubing,
lenses, etc., so I add a few hundred ml of the corrosion inhibitor,
check it monthly, and add more when necessary. After a few months, you
will get a pretty good idea of how much inhibitor it takes to increase
the ppm Mo by a given amount. I just float a small amount of the
dichlorophene on top of the water in the Haskris--it's not very
soluble--and add more when little solid remains.
Yours,
Bill

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Just wanted to thank everyone for the responses to my posting about the
TEM water chiller additives.
I'll try just using distilled water and monitoring the system more
frequently.
Thank you!
Beth

**********************************************************************
Beth Richardson
EM Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

Phone - (706) 542-1790 & FAX - (706) 542-1805
http://www.plantbio.uga.edu/emlab

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***

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8, 18 -- From beth-at-plantbio.uga.edu Tue May 16 17:45:55 2006
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Quite a while ago now I did some pre-embedding immunogold labelling, and
fixed in phosphate rather than Pipes buffer, using 2 mM rather than 5 mM Mg
and EGTA. I think use the fixative that works best for what you're after,
I'd suggest using the same fixative/buffer combination as for the LM work,
taking on board Tobias' comments about how EGTA softens the walls causing
some tissue distortion if you're not careful.

When I did this, I then cut frozen sections, rinsed in PBS then labelled the
sections on slides with antibodies in PBS, after the usual blocking in BSA
or gelatin. Then embedded the sections in Spurr's (messy) before sectioning
for TEM. I did this to get greater penetration of label into tissue, while
avoiding the original cut surface of the tissue block.

good luck,
cheers,
Rosemary

Rosemary White rosemary.white-at-csiro.au
CSIRO Plant Industry ph. 02-6246 5475
GPO Box 1600 mob. 0402 835 973
Canberra, ACT 2601 fax. 02-6246 5334
Australia

} From: meulia.1-at-osu.edu
} Reply-To: meulia.1-at-osu.edu
} Date: Tue, 16 May 2006 13:19:55 -0500
} To: rosemary.white-at-csiro.au
} Subject: [Microscopy] em immunolocalizations
}
}
}
}
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} We have been doing our pre-embedding immunogold localizations
} incubations for animal tissue in PBS, 0.2% BSA and 10mM Na Azide.
}
} I am working now on some em pre-embedding localizations on plant
} tissue, for which light microscopy localizations have been done using
} MTSB buffer (50mM PIPES, 5mM MgSO4, 5mM EGTA, pH~7.0). Is there any
} reason for which I should not be using this same MTSB buffer for the
} em work?
}
} Thank you very much for your suggestions. They will definitely save
} me trial time.
}
} Thanks.
}
} Tea
}
}
}
} --
} ***************************************
} Tea Meulia, PhD
} Director
} Molecular and Cellular Imaging Center
} Ohio State University/OARDC
} 1680 Madison Ave.
} Wooster OH 44691
}
} tel.: 330-263-3836 or -3828
} fax: 330-202-3563
}
} http://www.oardc.ohio-state.edu/mcic
}
} *****************************************
}
} ==============================Original Headers==============================
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7, 22 -- From Rosemary.White-at-csiro.au Tue May 16 17:51:17 2006
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Dear Tea,

The buffer you indicated may be fine, although a word of warning may be
in place: be careful when using divalent cations like Mg2+ . We've
never actually tested this for gold conjugates but such ions cause
aggregates of gold particles even at very low concentrations. The
coating proteins should help preventing that but it may still happen.
If that buffer, as Tobias Baskin indicates, is used to open cell walls
to antibodies, then it may be sufficient if it is applied only to
obtain that effect, i.e. fix, treat with the permeabilising buffer and
then wash with PBS a few times before proceeding using the same
protocol that was used for your animal tissue.

Good luck

Jan

On May 17, 2006, at 6:17 AM, meulia.1-at-osu.edu wrote:

}
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} We have been doing our pre-embedding immunogold localizations
} incubations for animal tissue in PBS, 0.2% BSA and 10mM Na Azide.
}
} I am working now on some em pre-embedding localizations on plant
} tissue, for which light microscopy localizations have been done using
} MTSB buffer (50mM PIPES, 5mM MgSO4, 5mM EGTA, pH~7.0). Is there any
} reason for which I should not be using this same MTSB buffer for the
} em work?
}
} Thank you very much for your suggestions. They will definitely save
} me trial time.
}
} Thanks.
}
} Tea
}
}
}
} --
} ***************************************
} Tea Meulia, PhD
} Director
} Molecular and Cellular Imaging Center
} Ohio State University/OARDC
} 1680 Madison Ave.
} Wooster OH 44691
}
} tel.: 330-263-3836 or -3828
} fax: 330-202-3563
}
} http://www.oardc.ohio-state.edu/mcic
}
} *****************************************
}
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7, 22 -- From leunissen-at-aurion.nl Tue May 16 19:23:02 2006
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Hi, Richard

This should be another interesting AFM experiment, done with liquid cell. It would be helpful, however, to get them to adhere to a substrate first, so that they don't roll around. Contact me off-line and we can probably put together a protocol for you. Also, if you can get samples, we can try to run them here. We have an AFM that can run in liquid.

Hope this is helpful,

Barbara Foster

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com

MME is now scheduling customized, on-site courses through September. Call us today for details.

P. S.
Need a good general reference or light microscopy text for the Spring semester? Call us today to learn more about "Optimizing LIght Microscopy". Copies still available through MME... even for class-room lots ... and we give quantity discounts. Call Ken Piel at (972)954-8011.

At 03:35 PM 5/16/2006, edelmare-at-muohio.edu wrote:

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This Question was submitted to Ask-A-Microscopist by (Dstekl-at-frontiernet.net)
from on Tuesday, May 16, 2006 at 19:19:11
Remember to consider the Grade/Age of the student when considering the Question
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Email: Dstekl-at-frontiernet.net
Name: Dave Steklenski

Organization: Catherine McAuley School, Rochester NY

Education: K-8 Grade Grammar School

Location: Rochester NY

Question: We are planning an elementary microscopy lab using a variety of natural and synthetic fibers. I would like to locat some photomicrographs of sample fibers so the kids could compare their observations to know appearance samples. I have been unable to find a good set of micrographs on the web.......are there any common references that might have some useable pictures.

THANK you for the help!!!

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (milindphd-at-gmail.com) from http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Wednesday, May 17, 2006 at 00:52:35
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Email: milindphd-at-gmail.com
Name: Milind Redkar

Organization: University institute of chemical technology

Education: Graduate College

Location: mumbai,India

Question: please let me know, why multilamellar vesicle show maltese crosses when veiwed with crosspolarizing microscopy.?

what is the difference between conoscopy and microscopy with resoect to multilamellar vesicles?

Thank you.

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I have a small box of new JEOL 35C electronic parts. Potentiometers,
lamps, IC's, transistors,push-button switches, etc. I'll send it to
anyone that wants it. Email me at directly.

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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I have 2 Jeol M3 cathodes I'll give anyone that wants them. They
were bought for a 100CX TEM. Both are new, but have been on a shelf
for 20yrs. Let me know if you want them.

Owen

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Hello TEM community: I have LEO 912AB TEM and need to order a new LaB6
filament. I can't for the life of me remember or find in my files what I
ordered the last time except that it was a Denka. If anyone has this
information and would pass it on I would be truly grateful. bob harris

Guelph Regional Imaging Facility
Dept.of Molecular and Cellular
Biology
New Science Complex
488 Gordon St.
Univ of Guelph
Guelph,On, Canada
N1G 2W1
ph: 519-824-4120 ext. 56409/58962
Fax: 519-837-1802

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Hi Dave,

Have a look at
http://www.microscopy-uk.org.uk/index.html
under MENU

It's a bit of struggle with all the links, but under Best Image Galleries &
Collections have a look at the SEM (scanning electron microscope) - the
first one. They aren't light microscopy but SEMs are great for fibre
morphology surface details.

I found things like Cellulose fibers (fibres) in toilet paper, paper towels,
T shirt cotton with dirt, asbestos (best to view that on-line), woven silk
etc.. [In 'Dennis Kunkel Microscopy, Inc.' link.]

Also search through the microscopy primer site, they have loads of stock
images

http://www.microscopy.fsu.edu

e.g.

http://www.microscopy.fsu.edu/primer/techniques/polarized/gallery/pages/silv
ercottonsmall.html

I found rabbit hair, silk, nylon (all under polarised light microscopy so
they appear brightly coloured - rather unlike standard microscope images).

The microscopy primer also tells you loads about microscopes (and you can
operate them virtually).

Plus try the hobby site
http://www.btinternet.com/~stephen.durr/
as it has links that may provide something (but most sites are interested in
things like plants, animals and pond life).

You may be able to get a library loan of an old book that has suitable
pictures for scanning, e.g.

Textile fiber atlas : A collection of photomicrographs of old and new
textile fibers, by Werner Von Bergen (Jan 1, 1949).

The microscopy of animal textile fibres,: Including methods for the complete
analysis of fibre blends. 235 half-tone and ll colour photomicrographs, 88
line drawings by Alec Blakey Wildman (Jan 1, 1954)

I found the book links on amazon.com (both are out of print).

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: Dstekl-at-frontiernet.net [mailto:Dstekl-at-frontiernet.net]
Sent: 17 May 2006 13:03
To: keith.morris-at-ucl.ac.uk

This Question was submitted to Ask-A-Microscopist by
(Dstekl-at-frontiernet.net)
from on Tuesday, May 16, 2006 at 19:19:11
Remember to consider the Grade/Age of the student when considering the
Question
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Listserver
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Email: Dstekl-at-frontiernet.net
Name: Dave Steklenski

Organization: Catherine McAuley School, Rochester NY

Education: K-8 Grade Grammar School

Location: Rochester NY

Question: We are planning an elementary microscopy lab using a variety of
natural and synthetic fibers. I would like to locat some photomicrographs of
sample fibers so the kids could compare their observations to know
appearance samples. I have been unable to find a good set of micrographs on
the web.......are there any common references that might have some useable
pictures.

THANK you for the help!!!

==============================Original Headers==============================
31, 26 -- From keith.morris-at-ucl.ac.uk Wed May 17 09:12:43 2006
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Ryan,
In addition to all the other suggestions about small sample size, out of the
lens field, etc., also try running your sample through a degausser just
before putting it in the SEM. Any residual magnetism is going to adversely
affect a high mag image. I can remember (too many years ago) swearing at
the SEM I operated (and didn't particularly like) because my resolution was
terrible, then remembering that my sample was a piece of carbon steel.
After degaussing, the resolution was fine.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
Fax 207-647-2688
kenconverse-at-qualityimages.biz
qualityimages.biz

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Email: lewryan-at-gmail.com
Name: Ryan

Organization: Dalhousie University

Title-Subject: [Filtered] SEM imaging on sample with magnetic substrate

Question: I have a saple with deposits of an alloy of Sn-Co-C on a magnetic
stainless steel substrate (430 stainless steel). I'm using a cold field
emission microscope and am having trouble getting high resolution images.
Could you suggest what setting I should use to get started?

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Dave,

Dave,

I forwarded your request to Scott Stoeffler, a member of our Optical
Microscopy group, and he provided these suggestions:

Any good basic book on textile science will have fiber photomicrographs.
Margery Joseph's Introductory Textile Science is a good one and
available pretty cheaply.

http://www.amazon.com/gp/product/0030507235/002-1547224-5402436?v=glance
&n=283155

There is also a CD available with a lot of fiber pictures, but it's
pretty expensive:

http://www.atexinc.com/digital_textiles_(cd).htm

Natural fibers such as cotton, linen, wool and silk are easier to
distinguish based on morphology alone. Without polarized light,
synthetics aren't easily distinguishable, although you can look at
cross-sections.

You might also take a look at our on-line Atlas of Microscopic Particles
at www.mccroneatlas.com. If you do a basic search on fibers, you'll get
some examples of cloth and paper fiber images with some background
information.

Hope this helps.

Regards,

Elaine

*********************************************************************
Elaine F.Schumacher
Senior Research Scientist
McCrone Associates, Inc.
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7417 (fax)
E-mail: eschumacher-at-mccrone.com
Web Site: www.mccrone.com

*********************************************************************
This message and any attachments are solely for the
intended recipient. If you are not the intended recipient,
disclosure, copying, use or distribution of the information
included in this message is prohibited.
*********************************************************************

-----Original Message-----
X-from: Dstekl-at-frontiernet.net [mailto:Dstekl-at-frontiernet.net]
Sent: Wednesday, May 17, 2006 6:55 AM
To: Elaine F. Schumacher

This Question was submitted to Ask-A-Microscopist by
(Dstekl-at-frontiernet.net)
from on Tuesday, May 16, 2006 at 19:19:11
Remember to consider the Grade/Age of the student when considering the
Question
------------------------------------------------------------------------
---
Please reply to both Dstekl-at-frontiernet.net as well as to the
Microscopy Listserver
------------------------------------------------------------------------
---

Email: Dstekl-at-frontiernet.net
Name: Dave Steklenski

Organization: Catherine McAuley School, Rochester NY

Education: K-8 Grade Grammar School

Location: Rochester NY

Question: We are planning an elementary microscopy lab using a variety
of natural and synthetic fibers. I would like to locat some
photomicrographs of sample fibers so the kids could compare their
observations to know appearance samples. I have been unable to find a
good set of micrographs on the web.......are there any common references
that might have some useable pictures.

THANK you for the help!!!

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

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Hello Bob,

The standard configuration of Denka cathode for Leo Microscopes is the
M3-CA, which has a 15 micron round tip. For greater brightness,
especially in TEM applications, many users prefer the M3-CA sharp 60/10
or M3-CA sharp 60/5. Please feel free to contact me off line if you
would like to discuss the differences.

Sincerely,
Mike Nesta

Michael R. Nesta
Managing Director
Energy Beam Sciences, Inc.
Tel: 860 653-0411
Fax: 860 653-0422
MNesta-at-ebsciences.com
www.ebsciences.com
�ADDING BRILLIANCE TO YOUR VISION�

bharris-at-uoguelph.ca wrote:
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} Hello TEM community: I have LEO 912AB TEM and need to order a new LaB6
} filament. I can't for the life of me remember or find in my files what I
} ordered the last time except that it was a Denka. If anyone has this
} information and would pass it on I would be truly grateful. bob harris
}
} Guelph Regional Imaging Facility
} Dept.of Molecular and Cellular
} Biology
} New Science Complex
} 488 Gordon St.
} Univ of Guelph
} Guelph,On, Canada
} N1G 2W1
} ph: 519-824-4120 ext. 56409/58962
} Fax: 519-837-1802
}
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Hi

A question to whose who have made some recent tests on FEG-SEM.

I would have advices about the differences which can be practically seen
between the Zeiss Supra 40 et the Supra 55. Resolutions annonced are
quite different. I want to know if it's only a spec difference, or if
practically it's easy to see it. We have made tests on the 55, but the
budget doesn't follow...
All other advices are welcome.

Thanks

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

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

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----Register Before May 19th and Save!---

Seeing at the Nanoscale IV, July 17-20, University of Pennsylvania, Philadelphia.

All users of SPM and metrology instruments are invited to Philadelphia, this July 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 a 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

We are very honored to have Dr. Paul Hansma, University of California, Santa Barbara, as our keynote speaker.

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

Be an early bird! -- Register by May 19 and receive a conference discount!

____________________________
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
____________________________

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Hi Beth,

Someone had said that distilled water will cause copper tubing to ionize, and correction will be faster than tap water. Has anybody heard this?

Ann Fook Yang
EM Unit/ Unite EM
AAFC/AAC
960 Carling Ave,
Ottawa,Ontario
Canada K1A 0C6
yanga-at-agr.gc.ca
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada

-----Original Message-----
X-from: beth-at-plantbio.uga.edu [mailto:beth-at-plantbio.uga.edu]
Sent: Tuesday, May 16, 2006 6:50 PM
To: Yang, Ann-Fook

Just wanted to thank everyone for the responses to my posting about the
TEM water chiller additives.
I'll try just using distilled water and monitoring the system more
frequently.
Thank you!
Beth

**********************************************************************
Beth Richardson
EM Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

Phone - (706) 542-1790 & FAX - (706) 542-1805
http://www.plantbio.uga.edu/emlab

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***

==============================Original Headers==============================
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The only additive I use is colloidal silver, sold as a health food item by Natural Immunogenics 888-328-8840. This method, suggested to me by Vitaly Feingold has worked very well. I also add a quart or so of tap water to the DI water so it is not so aggressive on the copper.

I have no connection with the silver supplier.

David Rothbard

beth-at-plantbio.uga.edu wrote:

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Ann:

Distilled water is fine. It is the deionized water that cause problems to the copper tubing due to the lack of ions in it.

Zhaojie Zhang
Microscopy Core Facility
University of Wyoming
Laramie, WY 82071

-----Original Message-----
X-from: YANGA-at-AGR.GC.CA [mailto:YANGA-at-AGR.GC.CA]
Sent: Wednesday, May 17, 2006 8:47 AM
To: Z.J. Zhang

Hi Beth,

Someone had said that distilled water will cause copper tubing to ionize, and correction will be faster than tap water. Has anybody heard this?

Ann Fook Yang
EM Unit/ Unite EM
AAFC/AAC
960 Carling Ave,
Ottawa,Ontario
Canada K1A 0C6
yanga-at-agr.gc.ca
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada

-----Original Message-----
X-from: beth-at-plantbio.uga.edu [mailto:beth-at-plantbio.uga.edu]
Sent: Tuesday, May 16, 2006 6:50 PM
To: Yang, Ann-Fook

Just wanted to thank everyone for the responses to my posting about the
TEM water chiller additives.
I'll try just using distilled water and monitoring the system more
frequently.
Thank you!
Beth

**********************************************************************
Beth Richardson
EM Lab Coordinator
Plant Biology Department
University of Georgia
Athens, GA 30602-7271

Phone - (706) 542-1790 & FAX - (706) 542-1805
http://www.plantbio.uga.edu/emlab

"Between the two evils,
I always pick the one I never tried before". Mae West (1893-1980)
*******************************************************************

"And it's only the giving that makes you what you are".
Wond'ring Aloud, Jethro Tull (Aqualung)

************************************************************************
***

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That is a difficult situation. First off, the Supra 55
was discontinued for some unknown reason, leaving the
Supra 55VP still viable. IMO, the 55 is superior to the
55VP. Why the 40 has lower resolution than the 55 is
unknown to me. At 200KX-350KX, I rather doubt that one
could tell the difference in resolution between the
40 and 55VP. But one probably could see a difference
between the 40 and 55. this assumes identical conditions
(low current, 30u center aperture, 3mm WD, 20KV, in-lens
detector).

gary g.

At 08:37 AM 5/17/2006, you wrote:

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Dear Milind,

The questions you have asked require a whole day lecture on polarized light, but here are the fundamentals.

Materials that respond to polarized light have different refractive indices along different axes. These axes are always perpendicular to each other, but not necessarily to the crystal edges or material directions (although they typically lie along the long and short edges of a fiber and along the direction of draw and perpendicular in a film.. but more about that later). The refractive index is a measurement of the interaction of the electric field in light with the electric field in matter: the greater the interaction, the higher the value. Mathematically, you can calculate RI by dividing the velocity of light in vacuum (300,000 km/s) with the velocity of light in the material.

I can't send diagrams in this email (will do so, if you contact me off line), but imagine a rectangle as representing your object, with the short side having lower RI and the long side having higher RI (Step 1) . (This is highly simplified, but works). Next, imagine the crossed polars, with direction of vibration transmitted through the first polar as vibrating East-West and the permitted direction of vibration through the second polar (the "analyzer") as North-South (Step 2).

Step 3: Cross the polars and put the object in between. Imagine that you are looking down on this "sandwich".
a. When either the short side or long side of the rectangle is parallel to the polarizer, only that direction receives energy from the polarized light. Since that light is vibrating E-W, it will not pass the analyzer. Rather, it will be absorbed, so the crystal appears dark in these two orientations.
b. Imagine rotating the crystal into any 45 degree position. In this orientation, both RI directions receive energy from the polarizer and, if you complete the rectangle so that you have a resultant vector, you will see that the vector has components that will pass through the analyzer. In this position, the crystal will appear bright between crossed polars.

Now for your multi-lamellar vesicle.
The fact that it has areas that appear bright indicates that its internal structure generates differences in electrical field with direction. In other words, it is anisotropic. If you consider how the lamellae are laid down in the vesicle, this makes perfect sense. The maltese cross that you are seeing indicate the two directions of the primary RI's. In each of these positions, only one RI is visible to the incoming polarized light; the same situation as described above in which the short or long side of the crystal is parallel to the direction of the polarizer or analyzer.

Actually, if you removed the analyzer and did RI tests with just the polarizer in place, then rotated the polarizer 90 degrees and did another RI test, you could actually determine the RI in each of those directions.

I don't have direct experience with this type of vesicle, but I would assume that, like starch and polymer spherulites, if you rotate your sample between XPol, the cross will stay in the same orientation indicating that your lamellae are sitting radially within the vesicle.

Hope this was helpful.

Best regards,
Barbara Foster

Microscopy/Microscopy Education
313 S Jupiter Rd, Suite 100
Allen, TX 75002
P: 972-954-8011
W: www.MicroscopyEducation.com

MME is now scheduling customized, on-site courses through September. Call us today for details.

P. S.
Need a good general reference or light microscopy text for the Spring semester? Call us today to learn more about "Optimizing LIght Microscopy". Copies still available through MME... even for class-room lots ... and we give quantity discounts. Call Ken Piel at (972)954-8011.

At 06:59 AM 5/17/2006, milindphd-at-gmail.com wrote:

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Yes, this can be the case. Distilled water can often be more corrosive to piping
than regular tap water, it does depend upon the water chemistry of the tap
water. What you really want is de-oxygenated water.

Beth, if you'd like more info, I can send you a synopsis of a recent discussion
on this subject from a while back.
Or if you prefer, you may email me directly and we could talk more about this,
as you wish.

dj

On Wed, 17 May 2006 YANGA-at-AGR.GC.CA wrote:

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}
} Someone had said that distilled water will cause copper tubing to ionize, and
correction will be faster than tap
} water. Has anybody heard this?
}
} Ann Fook Yang
} EM Unit/ Unite EM
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