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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
dcrippen-at-bucki
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
} America
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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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7, 19 -- From dianavd-at-eye.usyd.edu.au Tue May 2 20:37:34 2006
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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
Dept. Cell Biology, Box 3011 (for U.S. Mail)
458 Alex Sands Bldg, Research Dr (courier)
Durham, NC 27710

Office 919-668-2534
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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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} 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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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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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
++49 - 931 - 7848701 Fax
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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

Index of Advertisers

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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.
URL: http://www.aecom.yu.edu/aif/

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

==============================Original Headers==============================
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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, 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
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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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} 2, 24 -- From: Gordon Vrololjak {gvrdolja-at-nature.berkeley.edu}
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--
Greg Erdos
5410 SE 185th Ave.
Micanopy, FL 32667

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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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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
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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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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
Remember to consider the Grade/Age of the student when considering the Question
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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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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

This e-mail transmission, and any documents, files or previous e-mail
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attachments without reading or saving in any manner. Thank you.

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�

==============================Original Headers==============================
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This Question/Comment was submitted to the Microscopy Listserver
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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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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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:

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

--
Ritchie Sims Ph D Phone : 64 9 3737599 ext 87713
Microanalyst Fax : 64 9 3737435
Department of Geology email : r.sims-at-auckland.ac.nz
The University of Auckland
Private Bag 92019
Auckland
New Zealand

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Dear Greg,
If you could culture and clone these cells. do
karyotyping.
shashi

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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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5, 20 -- From shashis_99-at-yahoo.com Mon May 15 23:00:09 2006
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5, 20 -- Subject: Re: [Microscopy] Mouse -vs-Human
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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
}
} ==============================Original
} Headers==============================
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==============================Original Headers==============================
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==============================
5, 31 -- From opmills-at-mtu.edu Tue May 16 07:48:57 2006
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5, 31 -- From: "Owen P. Mills" {opmills-at-mtu.edu}
5, 31 -- Subject: dbl stick carbon tab blues
5, 31 -- Date: Tue, 16 May 2006 08:49:15 -0400
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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==============================
16, 23 -- From jchalcro-at-neuro.mpg.de Tue May 16 08:26:15 2006
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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

==============================Original Headers==============================
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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

----------------------------------------------------------------
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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)
} PhillipsT-at-missouri.edu
}
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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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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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} Headers==============================
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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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Please reply to both Dstekl-at-frontiernet.net as well as to the Microscopy 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!!!

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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

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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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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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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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___________________________________________________________
$0 Web Hosting with up to 200MB web space, 1000 MB Transfer
10 Personalized POP and Web E-mail Accounts, and much more.
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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-----
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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
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------------------------------------------------------------------------
---
Please reply to both Dstekl-at-frontiernet.net as well as to the
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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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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
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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==============================
8, 18 -- From beth-at-plantbio.uga.edu Tue May 16 17:45:55 2006
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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)

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

==============================Original Headers==============================
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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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} 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
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I am looking for an older Oxford Pulse Processor so I can do some testing of
EDX detectors. The model number can go back to #2020 and forward.
I don't need the complete analyzer system. If anyone has one of these pulse
processors that is available for sale, please contact me directly at my
email below.
Kind Regards,
Jim Nicolino

PulseTor/AAT
1816 St. Johns Bluff Road
Suite 305
Jacksonville, Florida 32246
904.646.3069
FAX 904.646.3131
JNicolino-at-comcast.net

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Dear group - has anyone used gold grids for cell culture and what luck
have you had with these grids?
Thanks
Barbara

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As the President, Officers, and only member of the Half-Norwegian (on my
Mother's side) Electron Microscopy Society, I would like to wish all you
listers a Happy Norwegian Independence Day!!

Also, thank you for all your recent help on nano-particles in agarose
and the fascinating string on ethics.

Uff Da!

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

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As long as lutefisk isn't involved ...
Phil

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On May 17, 2006, at 8:38 AM, YANGA-at-AGR.GC.CA wrote:

} Someone had said that distilled water will cause copper tubing to
} ionize, and correction will be faster than tap water. Has anybody
} heard this?
}
Dear Ann Fook,
I also have heard this. I have not done the experiment, but if [Cu+]
= [Cu++] = 0, then any dissolved oxidizing agent will cause Cu to be
ionized, so there is good reason to believe that distilled H2O will be
more corrosive. I would not advise using it in an EM.
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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On May 17, 2006, at 9:20 AM, ZZhang-at-uwyo.edu wrote:

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

Dear Zhaojie,
Distillation will remove non-volatile ions, so unless the tap water
has something like NH4HCO3 in it, it will not have many ions after
distillation.
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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This Question was submitted to Ask-A-Microscopist by (Mayya_Saab-at-dps-consulting.com)
from http://microscopy.org/Ask-A-Microscopist/Ask-A-Microscopist.html on Wednesday, May 17, 2006 at 13:52:20
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Email: Mayya_Saab-at-dps-consulting.com
Name: Mayya Saab

Education: Undergraduate College

Location: Herndon, VA, Fairfax

Title: darkfield microscopy

Question: What is this field? How can it help an individual?

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Email: kempfsc-at-auburn.edu
Name: Steve Kempf

Organization: Auburn University

Title-Subject: [Filtered] Running away sections

Question: I've had the strangest thing happen this past weekend and since and no one who I've talked to (some very experienced with thin sectioning) has been able to help.

I had been sectioning a block of tissue embedded in Poly/Bed 812 at 50 nm without difficulty using a Diatome diamond knife on a Leica Ultracut microtome, getting beautiful silver sections, spredding then by wafting a toothpick dipped in chloroform over them, and picking them up on formvar coated slot grids. All has been well until this weekend. Everything was set-up as it had been the previous day. Same knife, same boat water, same toothpicks, same choroform, same block, etc. As was the case previously, I got beautiful silver sections, but when I when to spread them with chloroform on a toothpick, they ran away as I moved the toothpick close to them, lickety-split. I could chase them around the boat with the toothpick. When I tried to pick them up on a grid, I couldn't. They would just slide off, back onto the water surface in the boat. Arrrrgh! Naturally, this is happening just as I reach the critical spot in the tissue I'm sectioning.

Not to be deterred, I cleaned the boat by washing it with clean water, got new beakers to hold the water I use, and tried again. The same thing happened. At that point I called it quits for the day and hoped it was some strange environmental effect that would disappear the following week. No Joy! When I sat down to section today, the same thing happened. I tried cleaning the knife boat with 0.2% ethanol in water, got new glassware, new chloroform, new toothpicks, etc. Still no joy?

So, my question is, can anyone give me some guidance as to what might be going on? I'm at my wits end.

Oh, one other thing, if I let the chloroform evaporate off the dipped toothpick, the sections don't run away from it.

Thanks for any advice,

Steve

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It sounds to me like a static electricity problem. Is your air
suddenly much drier? i.e. did they turn on the air conditioner? We
found many years ago that we could block this with those old darkroom
"dustfree" brushes with polonium strips in them. I don't know if
those are still available.

Joel

Date sent: Wed, 17 May 2006 19:39:32 -0500
To: jbs-at-temple.edu
X-from: kempfsc-at-auburn.edu
Send reply to: kempfsc-at-auburn.edu

I would like to test 3D reconstruction from tilted images (jpeg, tiff or
bmp) from an Hitachi S4000 SEM.
Does anyone knows a software that could do such thing ?
It has to be free, because it is just a test (at the moment) and running
under MS-Windows.
I know the easiest way yo do that is to paint 2 tilted images (with a
tilt difference of about 8 degrees) in red and green with a
photoshop-style software and use special glasses, but I would like to
display something that looks like an AFM map on the screen.
If anyone has experienced that type of job, I would be glad to have
advices.

Ludovic Pinier

==============================Original Headers==============================
3, 28 -- From ludovic.pinier-at-thalesgroup.com Thu May 18 02:10:14 2006
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For a free program for 3D surface topography have a look at

Measuring Surface Topography with Scanning Electron
Microscopy. I. EZEImage: A Program
to Obtain 3D Surface Data
Ezequiel Ponz,1 Juan Luis Ladaga,2 and Rita Dominga Bonetto1*

which appeared in Microscopy and Microanalysis 12 (2006) 170-177.

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: ludovic.pinier-at-thalesgroup.com [mailto:ludovic.pinier-at-thalesgroup.com]
Sent: Thursday, May 18, 2006 9:12 AM
To: j.bilde-at-risoe.dk

I would like to test 3D reconstruction from tilted images (jpeg, tiff or
bmp) from an Hitachi S4000 SEM.
Does anyone knows a software that could do such thing ?
It has to be free, because it is just a test (at the moment) and running
under MS-Windows.
I know the easiest way yo do that is to paint 2 tilted images (with a
tilt difference of about 8 degrees) in red and green with a
photoshop-style software and use special glasses, but I would like to
display something that looks like an AFM map on the screen.
If anyone has experienced that type of job, I would be glad to have
advices.

Ludovic Pinier

==============================Original Headers==============================
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I'll second the notion that your problem is static. In my lab here in
Houston, I have the same problem when there is low humidity. On days of
high humidity, no "run around" sections.

Mannie Steglich
UTMD Anderson Cancer Center

kempfsc-at-auburn.edu

05/17/2006 07:41 PM
Please respond to kempfsc

To:
msteglic-at-mdanderson.org
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Email: kempfsc-at-auburn.edu
Name: Steve Kempf

Organization: Auburn University

Title-Subject: [Filtered] Running away sections

Question: I've had the strangest thing happen this past weekend and since
and no one who I've talked to (some very experienced with thin sectioning)
has been able to help.

I had been sectioning a block of tissue embedded in Poly/Bed 812 at 50 nm
without difficulty using a Diatome diamond knife on a Leica Ultracut
microtome, getting beautiful silver sections, spredding then by wafting a
toothpick dipped in chloroform over them, and picking them up on formvar
coated slot grids. All has been well until this weekend. Everything was
set-up as it had been the previous day. Same knife, same boat water, same
toothpicks, same choroform, same block, etc. As was the case previously, I
got beautiful silver sections, but when I when to spread them with
chloroform on a toothpick, they ran away as I moved the toothpick close to
them, lickety-split. I could chase them around the boat with the
toothpick. When I tried to pick them up on a grid, I couldn't. They would
just slide off, back onto the water surface in the boat. Arrrrgh!
Naturally, this is happening just as I reach the critical spot in the
tissue I'm sectioning.

Not to be deterred, I cleaned the boat by washing it with clean water, got
new beakers to hold the water I use, and tried again. The same thing
happened. At that point I called it quits for the day and hoped it was
some strange environmental effect that would disappear the following week.
No Joy! When I sat down to section today, the same thing happened. I tried
cleaning the knife boat with 0.2% ethanol in water, got new glassware, new
chloroform, new toothpicks, etc. Still no joy?

So, my question is, can anyone give me some guidance as to what might be
going on? I'm at my wits end.

Oh, one other thing, if I let the chloroform evaporate off the dipped
toothpick, the sections don't run away from it.

Thanks for any advice,

Steve

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I would like to bring to your attention a vacancy in the EM Technical Support Group at the Materials Science Department, University of Oxford.

Information can be found at
http://www.materials.ox.ac.uk/vacancies/index.htm

Gabriella Chapman
Senior EM Support Technician
Department of Materials
University of Oxford
Begbroke

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Yesterday was the first time that I tried to cut a "turbinate" nose bone,
and I found it extremely hard, and even the poor razor blades were instantly
dull when I was trying to trim the block.

Does anyone have any good decalcification schemes for processing of
specimens which contain bone for electron microscopy?

This e-mail and/or any documents in this transmission is intended for the address(s) only and may contain legally privileged or confidential information. Any unauthorized use, disclosure, distribution, copying or dissemination is strictly prohibited. If you receive this transmission in error, please notify the sender immediately and return the original.

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Steve,
We gave up smelling chloroform a while ago and switched to
using a hot wire. The supply houses sell these, very nice units, but
with a little care you could probably make one. They deliver a
controlled amount of heat to a thin wire loop. You wave this near the
sections and just like chloroform, the heat flattens the sections
right out. I realize this isn't a fix for your problem right now, but
something to consider for the future. Your liver may thank you!

As ever,
Tobias Baskin
}
} Email: kempfsc-at-auburn.edu
} Name: Steve Kempf
}
} Organization: Auburn University
}
} Title-Subject: [Filtered] Running away sections
}
} Question: I've had the strangest thing happen this past weekend and
} since and no one who I've talked to (some very experienced with thin
} sectioning) has been able to help.
}
} I had been sectioning a block of tissue embedded in Poly/Bed 812 at
} 50 nm without difficulty using a Diatome diamond knife on a Leica
} Ultracut microtome, getting beautiful silver sections, spredding
} then by wafting a toothpick dipped in chloroform over them, and
} picking them up on formvar coated slot grids. All has been well
} until this weekend. Everything was set-up as it had been the
} previous day. Same knife, same boat water, same toothpicks, same
} choroform, same block, etc. As was the case previously, I got
} beautiful silver sections, but when I when to spread them with
} chloroform on a toothpick, they ran away as I moved the toothpick
} close to them, lickety-split. I could chase them around the boat
} with the toothpick. When I tried to pick them up on a grid, I
} couldn't. They would just slide off, back onto the water surface in
} the boat. Arrrrgh! Naturally, this is happening just as I reach the
} critical spot in the tissue I'm sectioning.
}
} Not to be deterred, I cleaned the boat by washing it with clean
} water, got new beakers to hold the water I use, and tried again. The
} same thing happened. At that point I called it quits for the day and
} hoped it was some strange environmental effect that would disappear
} the following week. No Joy! When I sat down to section today, the
} same thing happened. I tried cleaning the knife boat with 0.2%
} ethanol in water, got new glassware, new chloroform, new toothpicks,
} etc. Still no joy?
}
} So, my question is, can anyone give me some guidance as to what
} might be going on? I'm at my wits end.
}
} Oh, one other thing, if I let the chloroform evaporate off the
} dipped toothpick, the sections don't run away from it.
}
} Thanks for any advice,
}
} Steve
}
} -

--
_ ____ __ ____
/ \ / / \ / \ \ 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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Guy,

This is a perfect question for Histonet: www.histonet.org to
subscribe to the mailing list. There are several "boneheads" on the
list who can help you.

Phil

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Although Histo folks use formic acid, it is murder on ultrastructure.

I have better luck with small bone by 1st fixing for an hour or two in
regular fixative, then soaking in a buffered EDTA solution overnight,
then refixing in Glut, then embed as normal or in a hard epoxy.

Still... Don't use your BEST knife

Lou Ann

On May 18, 2006, at 8:18 AM, GBurgess-at-exchange.hsc.mb.ca wrote:

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}
} Yesterday was the first time that I tried to cut a "turbinate" nose
} bone,
} and I found it extremely hard, and even the poor razor blades were
} instantly
} dull when I was trying to trim the block.
}
} Does anyone have any good decalcification schemes for processing of
} specimens which contain bone for electron microscopy?
}
} This e-mail and/or any documents in this transmission is intended for
} the address(s) only and may contain legally privileged or confidential
} information. Any unauthorized use, disclosure, distribution, copying
} or dissemination is strictly prohibited. If you receive this
} transmission in error, please notify the sender immediately and return
} the original.
}
} ==============================Original
} Headers==============================
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} 4, 20 -- From: Garry Burgess {GBurgess-at-exchange.hsc.mb.ca}
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}
~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Lou Ann Miller, MT(ASCP)
Service Supervisor
Center for Microscopic Imaging
College of Veterinary Medicine
Rm 1204 VMBSB
2001 S Lincoln Ave
Urbana, IL 61802

217/244-1567

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

I'm seeking advice from those who have had success corralling small
specimens into dialysis tubing for EM processing. We recently tried
some samples in 2 different types of tubing that were convenient just
because we had them around. The samples were high pressure frozen and
freeze substituted but in the end one type of tube just dissolved in
the acetone and the other shrunk greatly, crushing the contents. Does
anyone have a recommendation for a brand of tubing that is robust in
this kind of treatment?

Our thought has been to embed the entire tube and contents for
sectioning. Is there any reason that might not work?

Thanks in advance,
Jay

Jay Campbell
Research Specialist
University of Wisconsin
Laboratory of Molecular Biology
R.M. Bock Labs
1525 Linden Drive
Madison, WI 53706
jmcampbe-at-wisc.edu
608 263 8481

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Dear listers,

Following 3 days of incubation with a substance, I can
observe an important decrease in cell viability by
ELISA (resazurin assay, BTW I recommand this test
since it is very good and costs nothing). Now I would
like to know if the cells die by apopotosis.
Unfortunately the commercial kits are very expensive
and there seems to be no "home made" solution. I
thought about observing the cells in fluorescence
using DAPI. I know DNA compaction is a late event of
apoptosis, and that microcroscopy does not give
quantitative measurements (easily) but it could at
least show a clear effect, like Y/N apoptosis is
involved in the reduction of cell viability. If I see
no sign of apoptosis by fluorescence I can search in
another direction and save some money in the process.

I would like to read your comments about my idea.

Regards,

St�phane (without-the-i)

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

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We use a Diatome Static LineII (which is an anti static device) with the tip
hanging close to where the sections are cut. I use it for room temperature
slicing even though it was designed for Cryo slicing. We bought ours through
EMS at www.emsdiasum.com. Hope this is helpful.
shawn

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On May 17, 2006, at 5:38 PM, Mayya_Saab-at-dps-consulting.com wrote:

} Question: What is this field? How can it help an individual?
}
Dear Mayya,
Dark field imaging is a technique where the unscattered beam (of
either light or electrons) is removed from the image, so that the image
consists only of the scattered particles. With this technique the most
strongly scattering parts of the specimen appear brightest. It is a
help for specimens of low contrast, and it can be used to highlight
those parts of a specimen that scatter into well-defined areas; e.g.,
crystals having a particular orientation or heavy atoms.
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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Here in Britain we have a gov't TV advertisement aimed at recruiting
teachers, showing young people with "active minds" asking lots of awkward
questions. One is:

"What's the difference between dust and fluff?"

WE know the answer - there should be one in every classroom!

*** from the neutrino who always pulls his weight ***,

-----------------------------------
Robert H. Olley
Reply to: R.H.Olley-at-reading.ac.uk
URL: http://www.rdg.ac.uk/~spsolley
-----------------------------------

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} --------------
}
} For a free program for 3D surface topography have a look at
}
} Measuring Surface Topography with Scanning Electron
} Microscopy. I. EZEImage: A Program to Obtain 3D Surface Data
} Ezequiel Ponz,1 Juan Luis Ladaga,2 and Rita Dominga Bonetto1*
}
} which appeared in Microscopy and Microanalysis 12 (2006) 170-177.
}
} 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

Did anybody tried to use this program?
I would greatly appreciate any comments.
Examples from the paper were not convincing.
I still do not believe that gray level alone
could be used for successful 3D reconstruction.
I would be glad to change my opinion.

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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10% EDTA adjusted with NaOH to PH 7.5 will do the work.
Change solution every other day for 1-2 weeks
(for small pieces of bone, about 0.5 mm square).
Better on a rotator.

As for calcified bone, I trim it with a small
file in a fume hood.

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

} --------------
}
}
} Yesterday was the first time that I tried to cut a
} "turbinate" nose bone, and I found it extremely hard, and
} even the poor razor blades were instantly dull when I was
} trying to trim the block.
}
} Does anyone have any good decalcification schemes for
} processing of specimens which contain bone for electron microscopy?
}
} This e-mail and/or any documents in this transmission is
} intended for the address(s) only and may contain legally
} privileged or confidential information. Any unauthorized use,
} disclosure, distribution, copying or dissemination is
} strictly prohibited. If you receive this transmission in
} error, please notify the sender immediately and return the original.

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Hi all,

Distilled water does have a slightly acidic pH. While we use
distilled water in our cooling systems, we add a bit of sodium
bicarbonate to bring the pH up to neutral. I just pull out the pH
paper 3-4 times a year and check the cooling systems.

Cheers,
Henk

At 08:02 PM 05/17/06, tivol-at-caltech.edu wrote:

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Hendrik O. Colijn colijn.1-at-osu.edu
Campus Electron Optics Facility Ohio State University
(614) 292-0674 http://www.ceof.ohio-state.edu
Time is that quality of nature which keeps events from happening all
at once. Lately it doesn't seem to be working.

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Dear Listers,

We are doing a rush job for a client who requires 4.0 um sections from
JB 4 blocks. Our ultramicrotomists extraordinaire, Cheryl, is having a
dickens of a time getting the sections to remain flat when removing them
from the knife. She is cutting on glass and taking sections from the
dry edge with a fine forceps. As soon as the sections leave the knife,
they curl and won't uncurl when placed on a drop of water on a slide.

Not only is this a rush job in support of a grant proposal, but it
requires serial sectioning with no missing sections, and we have, like,
no real experience with this resin. Cheryl has tried various sized
block faces and different thicknesses for the sections, but nothing is
helping.

The thumping sound you hear is a head hitting a wall----repeatedly. Can
anyone HEEEELLLLP??

Thanks!

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

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The problem of dealing with algal growth in water chillers is
discussed in some detail on p. 216 of my book, Vacuum Methods in
Electron Microscopy (ISBN 1-85578-052-6, available from SPi, Ladd,
Pella, etc.). One basic fact to remember is that algae require light
to grow, and so you can go a long way toward reducing algal growth by
excluding light from the chiller system (a light-tight cover for the
reservoir and opaque tubing leading to and from the instrument).
Beyond that, it is simple to add a bit of an algacide such as
Chloroamine-T or dichlorophene to the system. These chemicals are
available from specialty chemical companies such as Polysciences,
Sigma, etc., and perhaps from your local air conditioning or swimming
pool service company. For Chloramine-T the recommended concentration
is about one gram per gallon of water. It is, of course, advisable to
use distilled water to avoid the build-up of scale in the system.
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-engin.umich.edu;
Fx:734-763-4788; Ph:734-662-5237
Address mail to: 1136 Mixtwood Rd.
Ann Arbor, MI 48103-3035

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ah yes, the joys of bmma. i have spent many a pleasant hour cursing the
curling sections. in my experience, no change in shape, speed, thickness
makes a difference. I learned to be waiting for the section to start
cutting and then either grabbing a corner of it with a fine forceps or
using the forceps' tines to hold the corner down on to the surface of the
knife while it cut. then stopping the microtome, removing the section, and
re-starting the cutting motor. very tedious and time-consuming. it does
help to listen to NPR while doing this. I know you are stuck to the whims
of your client whose blocks are already embedded but I strongly recommend
any fans of JB-4 consider switching to the generic (and therefore less
expensive) butylmethyl methacrylate resin mix of Tobias Baskin. you can
cut on water filled boats and use acetone to extract the resin so the
sensitivity is better and the sectioning is trivial. My condolences to
Cheryl. Tom

At 02:16 PM 05/18/06, you wrote:

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

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

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Whoops - I mistakenly typed the wrong resin name in the first sentence of
my last posting - the correct version is below:

} ah yes, the joys of JB-4. i have spent many a pleasant hour cursing the
} curling sections. in my experience, no change in shape, speed, thickness
} makes a difference. I learned to be waiting for the section to start
} cutting and then either grabbing a corner of it with a fine forceps or
} using the forceps' tines to hold the corner down on to the surface of the
} knife while it cut. then stopping the microtome, removing the section, and
} re-starting the cutting motor. very tedious and time-consuming. it does
} help to listen to NPR while doing this. I know you are stuck to the whims
} of your client whose blocks are already embedded but I strongly recommend
} any fans of JB-4 consider switching to the generic (and therefore less
} expensive) butylmethyl methacrylate resin mix of Tobias Baskin. you can
} cut on water filled boats and use acetone to extract the resin so the
} sensitivity is better and the sectioning is trivial. My condolences to
} Cheryl. Tom
}
} At 02:16 PM 05/18/06, 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
} } ----------------------------------------------------------------------------
} }
} } Dear Listers,
} }
} } We are doing a rush job for a client who requires 4.0 um sections from
} } JB 4 blocks. Our ultramicrotomists extraordinaire, Cheryl, is having a
} } dickens of a time getting the sections to remain flat when removing them
} } from the knife. She is cutting on glass and taking sections from the
} } dry edge with a fine forceps. As soon as the sections leave the knife,
} } they curl and won't uncurl when placed on a drop of water on a slide.
} }
} } Not only is this a rush job in support of a grant proposal, but it
} } requires serial sectioning with no missing sections, and we have, like,
} } no real experience with this resin. Cheryl has tried various sized
} } block faces and different thicknesses for the sections, but nothing is
} } helping.
} }
} } The thumping sound you hear is a head hitting a wall----repeatedly. Can
} } anyone HEEEELLLLP??
} }
} } Thanks!
} }
} } 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 Headers==============================
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} } 11, 23 -- Date: Thu, 18 May 2006 14:15:26 -0500
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} } 11, 23 -- From: "Tindall, Randy D." {TindallR-at-missouri.edu}
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}
} Thomas E. Phillips, PhD
} Professor of Biological Sciences
} Director, Molecular Cytology Core
} 2 Tucker Hall
} University of Missouri
} Columbia, MO 65211-7400
}
} 573-882-4712 (office)
} 573-882-0123 (fax)
} PhillipsT-at-missouri.edu
}
}
}
} ==============================Original Headers==============================
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} 9, 20 -- To: Microscopy-at-msa.microscopy.com
} 9, 20 -- From: Tom Phillips {phillipst-at-missouri.edu}
} 9, 20 -- Subject: Re: [Microscopy] Sectioning JB 4 resin
} 9, 20 -- In-Reply-To: {200605181916.k4IJG7it023109-at-ns.microscopy.com}
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Thomas E. Phillips, PhD
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400

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

==============================Original Headers==============================
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}
} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
} Postdoctoral Position in Scanning Transmission Electron Microscopy -
} University of California, Santa Barbara
}
} Applicants should have demonstrated experience in TEM and a strong
} background in materials science and diffraction. Preference will be
} given to applicants with expertise in STEM techniques, such as
} atomic resolution HAADF imaging. Facilities at UCSB include a
} Tecnai F30U TEM/STEM and other state-of-the-art imaging,
} spectroscopy and diffraction facilities. Research projects include
} the characterization of interfaces and defects in high-permittivity
} oxide thin films, including gate dielectrics and ferroelectrics.
}
} The position is available summer/fall 2006. Duration about 1-2
} years, salary is commensurate with qualifications. Candidates must
} have a Ph.D. in Materials Science or Physics. Interested candidates
} should send a curriculum vitae, publication list and the names of
} three references with their contact information to:
}
} Prof. Susanne Stemmer
} Materials Department
} University of California
} Santa Barbara, CA 93106-5050
} Email: stemmer-at-mrl.ucsb.edu
} http://www.mrl.ucsb.edu/~stemmer
} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

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Email: kempfsc-at-auburn.edu
Name: Steve Kempf

Organization: Auburn University

Title-Subject: [Filtered] RE: Running away sections

Question: I just wanted to thank everyone who replied to my question yesterday. I too thought static charge might be the problem; however, it seemed strange that if that was the case, the sections didn't "run away" when I put the chloroform dipped toothpick next to them after allowing the chloroform to evaporate. At any rate, late yesterday I dismounted the block I had been having the problems with and mounted new one and, guess what, no problems. Everything worked fine. I didn't have time to re-try the block that I had been having the problem with again followng that, so I put it in a 60 degree oven overnight and mounted it this morning. And, as the Gods of sectioning would have it, everything was back to normal with no "running away" problems.

Go figure?

So, if it was static, did just changing the block releave the charge?

At any rate, thanks again for all the suggestions.

Steve

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Hi, All-

I have a client who would like to figure out the composition (in terms of
protein, carbohydrates, and nucleic acids) of small (0.2um - 1um) abiotic
particles. They have already tried to stain with DAPI and a couple of
other things (I don't know what) and look with fluorescence and flow
cytometry, but the size/signal is too low.

The only approach I know is microscopy (I'm a one-trick pony). Should they
be pursuing other methods, talking to biochemists, geochemists, food
nutritionists? Their sample size is very tiny; microscopic.

For microscopy all I can think of is things like antibodies to nucleic
acids, hitting it with all kinds of lectins, and other shotgun approaches.
They have been able to get some of these on a coated grid with an airfuge;
they are pretty electron dense. I shudder to think that I will need to
embed and section them, but I can give it a shot. No idea if they also
contain metallic particles, but I would not be surprised.

Does anyone have any ideas how to approach this challenge?

Mahalo (thanks),
Tina

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

==============================Original Headers==============================
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Randy,

Similar to what Tom suggested below, that is,
mechanical restriction to reduce the roll up of
the section, is to use a fine tipped artists
brush to sort of tamp the section lightly to the
surface of the knife as it comes of the knife
edge, or to use light pressure on top of section
to restrict the roll up. A slow cutting speed
would give you more time to do that. Then use
the brush to lift it off the knife surface to
the slide. I had a microtomist here a year ago
who had to do that and I think it was JB-4 or
GMA resin that she was cutting. Best of luck to
Cheryl!

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

phillipst-at-missouri.edu wrote:
} ----------------------------------------------------------------------------
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} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------------
}
} ah yes, the joys of bmma [JB-4]. i have spent many a pleasant hour cursing the
} curling sections. in my experience, no change in shape, speed, thickness
} makes a difference. I learned to be waiting for the section to start
} cutting and then either grabbing a corner of it with a fine forceps or
} using the forceps' tines to hold the corner down on to the surface of the
} knife while it cut. then stopping the microtome, removing the section, and
} re-starting the cutting motor. very tedious and time-consuming. it does
} help to listen to NPR while doing this. I know you are stuck to the whims
} of your client whose blocks are already embedded but I strongly recommend
} any fans of JB-4 consider switching to the generic (and therefore less
} expensive) butylmethyl methacrylate resin mix of Tobias Baskin. you can
} cut on water filled boats and use acetone to extract the resin so the
} sensitivity is better and the sectioning is trivial. My condolences to
} Cheryl. Tom
}
} At 02:16 PM 05/18/06, 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
} } ----------------------------------------------------------------------------
} }
} } Dear Listers,
} }
} } We are doing a rush job for a client who requires 4.0 um sections from
} } JB 4 blocks. Our ultramicrotomists extraordinaire, Cheryl, is having a
} } dickens of a time getting the sections to remain flat when removing them
} } from the knife. She is cutting on glass and taking sections from the
} } dry edge with a fine forceps. As soon as the sections leave the knife,
} } they curl and won't uncurl when placed on a drop of water on a slide.
} }
} } Not only is this a rush job in support of a grant proposal, but it
} } requires serial sectioning with no missing sections, and we have, like,
} } no real experience with this resin. Cheryl has tried various sized
} } block faces and different thicknesses for the sections, but nothing is
} } helping.
} }
} } The thumping sound you hear is a head hitting a wall----repeatedly. Can
} } anyone HEEEELLLLP??
} }
} } Thanks!
} }
} } 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
} }
}
} 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

==============================Original Headers==============================
5, 19 -- From ahlst007-at-umn.edu Thu May 18 17:53:24 2006
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_____________________________________________
X-from: Randi Olsen
Sent: 18. mai 2006 16:04
To: 'Microscopy-at-microscopy.com'

Hi listers,

Our lab wishes to print out b/w images from scanned TEM negatives and
bypass completely darkroom enlargements.
Printouts should not be expensive but should have a good dynamic range
to allow researchers to evaluate fine structures etc.
Can anyone suggest (or even recommend) the most suitable monochrome
printers for this purpose.
We have already a Fujix pictrography system so final (expensive)
printouts are no problem here.
TIA,

Jim

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Randi

I must admit that I have never attempted growing bacteria on grids.
But I wonder if the copper of the grids is too reactive. It might both
inhibit bacteria and encourage precipitation/reaction products.

You could
try a couple of gold grids as a control to see if matters improve.

Good
luck

Malcolm

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

----- Original Message -----
X-from: Randi.Olsen-at-fagmed.uit.no

Dear Randi,

A possible way out of your problem might be the technique I used years
ago in the study of thermophilic bacteria in nature.
I deposited carbon films in vacuo on small freshly-cleaved mica squares
and let them mature for a few days. The films were carefully floated off
on the water surface and the bacteria were allowed to settle on and
attach to the underside of the carbon. The films were then transferred
to other solutions with a fine Pt loop (or even the original mica piece
- submerged then lifted up below the film).
Staining with aqueous solutions of UA, AM or PTA can be done either
before (preferably) or after the films are picked up with TEM grids.
Good luck.

Jim

-----Original Message-----
X-from: Randi.Olsen-at-fagmed.uit.no [mailto:Randi.Olsen-at-fagmed.uit.no]
Sent: Friday, May 19, 2006 9:49 AM
To: James Chalcroft

_____________________________________________
X-from: Randi Olsen
Sent: 18. mai 2006 16:04
To: 'Microscopy-at-microscopy.com'

Tina,

If they just need things like "protein" or "carbohydrate", how about
cytochemical staining methods for light microscopy? Deposit the
particles on slides, and pretend they're bacteria.
SEM/EDX ought to pick up metals.
Mind, there are things like ninhydrin for protein, and the
protein/carbohydrate/fat methods physiological ecologists (and in the
Old Days, biochemists) use, if there is enough sample.

Phil

} Hi, All-
}
} I have a client who would like to figure out the composition (in terms of
} protein, carbohydrates, and nucleic acids) of small (0.2um - 1um) abiotic
} particles. They have already tried to stain with DAPI and a couple of
} other things (I don't know what) and look with fluorescence and flow
} cytometry, but the size/signal is too low.
}
} The only approach I know is microscopy (I'm a one-trick pony). Should they
} be pursuing other methods, talking to biochemists, geochemists, food
} nutritionists? Their sample size is very tiny; microscopic.
}
} For microscopy all I can think of is things like antibodies to nucleic
} acids, hitting it with all kinds of lectins, and other shotgun approaches.
} They have been able to get some of these on a coated grid with an airfuge;
} they are pretty electron dense. I shudder to think that I will need to
} embed and section them, but I can give it a shot. No idea if they also
} contain metallic particles, but I would not be surprised.
}
} Does anyone have any ideas how to approach this challenge?
}
} Mahalo (thanks),
} 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*
} ****************************************************************************
--
Philip Oshel
Microscopy Facility Supervisor
Department of Biology
Central Michigan University
024C Brooks Hall
Mt. Pleasant, MI 48859
(989) 774-3576

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Hi Jim,
We were apprehensive 3-4 years ago when we made this switch, but were
quickly satisfied with the plain-paper work-print quality and speed
of an HP Laserjet 4100, cost around $2000 back then. You can always
treat yourself to a laser paper of higher brightness, like 98-105,
than the noticeably dingy photo-copier paper we find ourselves
settling for most of the time because "that's what's loaded in the
printer". We "never" make glossies any more; a rare inkjet
grayscale run of printing is the nearest thing. Publications are all
done via digital files now. A local company that is quoting us on a
CCD system for Philips EM 420 says their customers are now very
satisfied with

$625 Hewlett Packard LaserJet 2420 MonoChrome Laser Printer (optional
purchase) * Up to 30 ppm speed, 1200 dpi. Add $375 for
network/duplexing capability.

and they advise: "The printer listed is the printer most recommended
for gray-scale images. I can quote whatever printer you specify.
However, once you are using digital images, you may find that you
don't have a need for printing images. Before deciding that you want
a high-quality printer, it would be good to speak with a few
colleagues using CCD systems and ask how much printing they do.".

There is a frequent rumor among some experts that HP Laser printers
have undocumented capability to produce at 1800 dpi. At any event,
we print at something like 120-150 lines per inch; 120 would leave
each 1200 dpi pixel sized 10 x 10 dots, giving a range of 10x10 gray
levels, but we feel no constraints or shortcomings-- output may not
be 256 gray levels, but seems very adequate. We scan negatives
routinely at 800 ppi, sometimes 1200 ppi using Epson flatbeds via
Photoshop plug-in, and we set up our output-screen properties/
preferences using the interface options between Photoshop and the HP
printer that show up under Page Setup and Print windows (Macintosh);
this and the grayscale display preferences make differences you
probably want to explore and choose among. I'm too preoccupied to
dig for our exact numerical settings; they have been unchanged for
too long to remember the decisions. we settled on.

-mike reedy-

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

--
-mike reedy-

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

Office 919-668-2534
Lab 919-684-5674
Fax 919-681-9929
mike.reedy-at-cellbio.duke.edu
http://note.cellbio.duke.edu/Faculty/Research/Reedy.html

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On May 18, 2006, at 3:07 PM, tina-at-pbrc.hawaii.edu wrote:

} I have a client who would like to figure out the composition (in terms
} of
} protein, carbohydrates, and nucleic acids) of small (0.2um - 1um)
} abiotic
} particles. They have already tried to stain with DAPI and a couple of
} other things (I don't know what) and look with fluorescence and flow
} cytometry, but the size/signal is too low.
}
} The only approach I know is microscopy (I'm a one-trick pony). Should
} they
} be pursuing other methods, talking to biochemists, geochemists, food
} nutritionists? Their sample size is very tiny; microscopic.
}
} For microscopy all I can think of is things like antibodies to nucleic
} acids, hitting it with all kinds of lectins, and other shotgun
} approaches.
} They have been able to get some of these on a coated grid with an
} airfuge;
} they are pretty electron dense. I shudder to think that I will need to
} embed and section them, but I can give it a shot. No idea if they also
} contain metallic particles, but I would not be surprised.
}
} Does anyone have any ideas how to approach this challenge?
}
Dear Tina,
There are some very sensitive methods to measure protein,
carbohydrate, and nucleic acids that give quantitative information
about composition. If the particles can be dissolved, the protein can
be separated from the carbs and NAs by centrifugation, then it can be
hydrolysed into amino acids and run through a column to get the
composition, or it can be subjected to a Lowry determination (or,
perhaps, more up-to-date procedures). Similarly, NA can be amplified
by quantitative PCR, then quantitated or sequenced. I'm pretty sure
that carbs can also be quantitated biochemically, but I am not at all
familiar with those methods. Of course, none of these methods will
give any structural info, so if that is needed, EM would be indicated.
Talking to several members of a biology department should lead to the
experts in dealing with the components of the particles, who would be
able to give details on how best to proceed. Metals could be
determined and quantitated by atomic absorption spectroscopy, but again
this gives only overall composition, not structural info. The small
sample size makes it essential that the best methods be identified
before proceeding; i.e., doing it right is better than doing it fast.
Good luck.
Yours,
Bill Tivol, PhD
EM Scientist and Manager
Cryo-Electron Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu

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Hi Tina,
The University of Georgia Complex Carbohydrate Research Center (CCRC)
may be able to help. They have an amazing state-of-the-art facility - I
call it "The Palace";-) Check out www.ccrc.uga.edu to see if someone
there can help with advice.
best,
Beth

On Thursday, May 18, 2006, at 06:08 PM, tina-at-pbrc.hawaii.edu wrote:

}
}
}
} -----------------------------------------------------------------------
} -----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
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}
} Hi, All-
}
} I have a client who would like to figure out the composition (in terms
} of
} protein, carbohydrates, and nucleic acids) of small (0.2um - 1um)
} abiotic
} particles. They have already tried to stain with DAPI and a couple of
} other things (I don't know what) and look with fluorescence and flow
} cytometry, but the size/signal is too low.
}
} The only approach I know is microscopy (I'm a one-trick pony). Should
} they
} be pursuing other methods, talking to biochemists, geochemists, food
} nutritionists? Their sample size is very tiny; microscopic.
}
} For microscopy all I can think of is things like antibodies to nucleic
} acids, hitting it with all kinds of lectins, and other shotgun
} approaches.
} They have been able to get some of these on a coated grid with an
} airfuge;
} they are pretty electron dense. I shudder to think that I will need to
} embed and section them, but I can give it a shot. No idea if they also
} contain metallic particles, but I would not be surprised.
}
} Does anyone have any ideas how to approach this challenge?
}
} Mahalo (thanks),
} 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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} Headers==============================
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}
**********************************************************************
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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Hi

We are looking to purchase some software that will log use of the
computers that are interface to our microscopes.

Basically on a per user basis the software would record when the user
log on and off, and would record uses of the imaging packages installed
on the computer.

Currently we do this using the built in Windows 2000/ XP security event
logs. This does the job but is not very user friendly.

All systems in question are PCs (no Mac systems) and the users log on
with Windows Active Directory accounts.

Does any one out there know of any package that does this? It would be
particularly useful if the software would not only record the data but
collate it too, i.e. provide tables of total usage per user over a given
time period.

Thanks

Lloyd Williams

Manager of Bio-Imaging Core Facility

Hunter College

695 Park Ave

New York NY 10021

==============================Original Headers==============================
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Hi Randi,

There are two potential problems with the approach you describe. One is that
the copper grids, as pointed out in another reply, may be reacting to the
salts in the growth medium. He other problem is that the bacteria are in th
eprocess of forming a biofilm on the support film surface during the
incubation time.

If you only want to examine the bacteria by negative stain, you can place a
drop of the culture on the support film surface, leave for about 30 sec and
then replace the drop with a drop of negative stain. You have to be careful
to make sure there are no salts in the growth medium that could cause the
negative stain, and in particular the uranyl acetate, to precipitate. There
is no need to centrifuge the bacteria because there are usually enough in
the drop to fall to the surface.

As with most techniques, there are many variations on this simple method,
but it is always a good policy to try the easier way first to make sure it
is possible to obtain results. The next easiest way is to deposit a carbon
film onto a freshly-cleaved mica surface and then allow a drop of bacterial
suspension to infiltrate between the carbon film and the mica. If you then
place a clean grid on the carbon film you should be able to pick it up onto
the support grid and then negative stain. The advantage of this method is
that you don't have to worry about the support film being hydrophilic - it
already is. The disadvantage is that the carbon film is very fragile so
needs a small mesh grid to support it. Even better support comes form holey
grids.

If you are interested in examining the bacteria as they grow on the grids,
first switch to gold or nickel grids. You may then want to extensively wash
the bacteria before you stain them to remove all the extracellular material
they secrete in the short time they have been incubated. You can check to
see if what I am saying is correct by taking a grid with bacteria on
directly from the culture medium (after incubation) and plunge-freezing it
in liquid propane. From there you freeze substitute it in ethanol (with a
little osmium tetroxide if you wish), warm it slowly to 4 degrees and then
critical point dry it and look at it in the SEM. My guess is that the
bacteria will be almost invisible because they will be covered in a large
amount of slimy looking stuff.

I hope this helps.

Paul Webster.

Paul Webster, Ph.D
House Ear Institute
2100 West Third Street
Los Angeles, CA 90057
(213) 273 8026
pwebster-at-hei.org

} From: {Randi.Olsen-at-fagmed.uit.no}
} Reply-To: {Randi.Olsen-at-fagmed.uit.no}
} Date: Fri, 19 May 2006 02:47:23 -0500
} To: {pwebster-at-hei.org}
} Subject: [Microscopy] FW: NEGATIVE STAINING ENTEROCOCCI
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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}
} _____________________________________________
} X-from: Randi Olsen
} Sent: 18. mai 2006 16:04
} To: 'Microscopy-at-microscopy.com'
} Subject: NEGATIVE STAINING ENTEROCOCCI
}
} Dear all,
} We have problems with negative staining of E. Coli, and have tried two
} different approaches with both Uranyl acetate and PTA.
} Without being able to detect the flagellas, we see partly collapsed
} bacterias and lots of dirt (growing bacterias directly on the grids)
} We've also tried embed the E.coli in a mixture of methylcellulose and
} uranyl acetate without any luck.
} We've used carboncoated formvar films on copper grids.
} I enclose the procedure for preparing cells grown on grids, observations
} and some questions form the scientist in charge of this project.
} We might be wrong trying to grow cells directly on grids?
}
}
} Our aim is to prepare grids for TEM with a suitable cell number for
} reliable anaysis without damaging the cells by centrifugation.
} We plan to grow the cells directly onto the grids and have performed the
} following:
}
} Inoculate cultures of enterococci in TSB+0.25%glucose and grow overnight
} at 37*C without shaking. (TSB is a bacterial growth medium made from
} casein and soya peptone).
} Dilute cultures 100-fold in 10 ml growth medium in microtiterplate wells
} containing TEM grids (should contain ca.107 cells/ml).
} Incubate at 37*C without shaking for 1-4 hours.
} Pick up grids at different time-points (i.e. 1h, 2h, 3h and 4h) from the
} culture.
} Dip the grid in particle free water.
} Stain the cells with 0.5% uranyl acetate for 30".
} Rinse the grids gently in particle free water before air-drying them for
} 10-15'.
}
} Observations made:
} 1. there are low cell-numbers on the grids
} 2. there's a lot of stained debris or precipitate on the grids and it
} seems to increase during the incubation.
}
} Questions:
} 1. What cell density is required to obtain one layer of cells on the
} grid but not cells on top of each other?
} 2. Are there ways of increasing the bacterial affinity for the grids?
} 3. Are there ways to reduce the amount of debris on the grids?
} 4. Can grids be incubated on top of a nitrocellulose filter without
} beeing damaged?
}
}
}
}
} Best regards
}
} Randi Olsen
} Department of Electron Microscopy
} University of Tromso
} Norway
}
}
} ==============================Original Headers==============================
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==============================Original Headers==============================
12, 19 -- From PWebster-at-hei.org Fri May 19 13:54:08 2006
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12, 19 -- From: "Webster, Paul" {PWebster-at-hei.org}
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On May 11, 2006, at 1:13 PM, dlowry-at-asu.edu wrote:

} 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
}
Dear David,
Our lab web page has links to our work. We do not have links to other
labs, AFAIK. The URL is

http://www.jensenlab.caltech.edu/

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

==============================Original Headers==============================
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Hi all:

Does anyone have an archived copy of the PDF brochure
for the LEO 1550 FESEM that could be emailed?

I would appreciate that and any other reference material
on the 1550.

gary g.

==============================Original Headers==============================
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Hello All,

Bit of a microscopy fest happening in the UK soon. See below for details.
Hope to see some of you there!

Best Wishes,

Debbie.

Dr Debbie Stokes
University of Cambridge
Dept of Physics
Cavendish Laboratory
Cambridge, UK

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

"Festival of Microscopy" in the UK - 23rd - 30th June 2006

23rd-25th June 2006 - SuperSTEM Summer School, CLRC Daresbury
Laboratory, UK
http://www.superstem.dl.ac.uk/

26th June 2006 - ESEM VII, 7th Annual European ESEM User club meeting,
Novotel ExCel, London, UK
http://www.rms.org.uk/event_esem.shtml

*** EMS sponsored event ***

27th - 29th June 2006 - MICROSCIENCE 2006, ExCel Centre, London, UK
http://www.microscience2006.org.uk

Note: MICROSCIENCE 2006 includes UK SPM 2006

30th June 2006 - In Situ Electron Microscopy and Analysis, Institute of
Physics, London, Portland Place.
http://conferences.iop.org/ISEM/

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

==============================Original Headers==============================
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Hello Randi,
I do not have answers for all the questions but this is what we have
done in the past while negatively staining Gram negative bacteria
Questions:
} } 1. What cell density is required to obtain one layer of cells on the
} } grid but not cells on top of each other?
2. Are there ways of increasing the bacterial affinity for the grids?
} } 3. Are there ways to reduce the amount of debris on the grids?
} } 4. Can grids be incubated on top of a nitrocellulose filter without
} } beeing damaged?

Answer: It might be challenging to achieve a monolayer of bacterial
cells on the grid as every bacteria behaves a little differently when
it comes to attaching to any surface.
In order to view flagella or pili, we grew the bacteria in stationary
cultures to prevent mechanical loss of flagella or pili. Once the
bacteria were in in their log phase we pipetted out around 100
microliters of the culture and floated a fomvar coated nickle grid on
it for about 5 mins and then let the grids dry. We found the
incubation time to vary with different strains.
These grids were rinsed in distilled water for 30 sec to a min. I have
found that rinsing the grids was very important to remove all the
debris and residual media from the grid leaving a clean prep.
If you find low number of bacterial cells you can try increasing the
incubation time and decreasing the rinse time.
Staining:
The grids were stained for 10 mins with 0.2% UA for 10 mins followed
by a 1 min rinse with distilled water. The grids were then air dried
before examination.

I found rinsing steps were key to a good prep. You have to play with
the innoculation and rinsing steps to obtain desirable results.

Hope that helps. Good luck with your experiments.

regards,
Vinod Nair
Electron microscopy lab
New Mexico State University
Las Cruces
NM 88003

} } 2. Are there ways of increasing the bacterial affinity for the grids?
} } 3. Are there ways to reduce the amount of debris on the grids?
} } 4. Can grids be incubated on top of a nitrocellulose filter without
} } beeing damaged?

On 5/19/06, PWebster-at-hei.org {PWebster-at-hei.org} 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 Randi,
}
} There are two potential problems with the approach you describe. One is that
} the copper grids, as pointed out in another reply, may be reacting to the
} salts in the growth medium. He other problem is that the bacteria are in th
} eprocess of forming a biofilm on the support film surface during the
} incubation time.
}
} If you only want to examine the bacteria by negative stain, you can place a
} drop of the culture on the support film surface, leave for about 30 sec and
} then replace the drop with a drop of negative stain. You have to be careful
} to make sure there are no salts in the growth medium that could cause the
} negative stain, and in particular the uranyl acetate, to precipitate. There
} is no need to centrifuge the bacteria because there are usually enough in
} the drop to fall to the surface.
}
} As with most techniques, there are many variations on this simple method,
} but it is always a good policy to try the easier way first to make sure it
} is possible to obtain results. The next easiest way is to deposit a carbon
} film onto a freshly-cleaved mica surface and then allow a drop of bacterial
} suspension to infiltrate between the carbon film and the mica. If you then
} place a clean grid on the carbon film you should be able to pick it up onto
} the support grid and then negative stain. The advantage of this method is
} that you don't have to worry about the support film being hydrophilic - it
} already is. The disadvantage is that the carbon film is very fragile so
} needs a small mesh grid to support it. Even better support comes form holey
} grids.
}
} If you are interested in examining the bacteria as they grow on the grids,
} first switch to gold or nickel grids. You may then want to extensively wash
} the bacteria before you stain them to remove all the extracellular material
} they secrete in the short time they have been incubated. You can check to
} see if what I am saying is correct by taking a grid with bacteria on
} directly from the culture medium (after incubation) and plunge-freezing it
} in liquid propane. From there you freeze substitute it in ethanol (with a
} little osmium tetroxide if you wish), warm it slowly to 4 degrees and then
} critical point dry it and look at it in the SEM. My guess is that the
} bacteria will be almost invisible because they will be covered in a large
} amount of slimy looking stuff.
}
} I hope this helps.
}
} Paul Webster.
}
}
} Paul Webster, Ph.D
} House Ear Institute
} 2100 West Third Street
} Los Angeles, CA 90057
} (213) 273 8026
} pwebster-at-hei.org
}
}
} } From: {Randi.Olsen-at-fagmed.uit.no}
} } Reply-To: {Randi.Olsen-at-fagmed.uit.no}
} } Date: Fri, 19 May 2006 02:47:23 -0500
} } To: {pwebster-at-hei.org}
} } Subject: [Microscopy] FW: NEGATIVE STAINING ENTEROCOCCI
} }
} }
} }
} }
} } ----------------------------------------------------------------------------
} } 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
} } ----------------------------------------------------------------------------
} }
} }
} }
} } _____________________________________________
} } X-from: Randi Olsen
} } Sent: 18. mai 2006 16:04
} } To: 'Microscopy-at-microscopy.com'
} } Subject: NEGATIVE STAINING ENTEROCOCCI
} }
} } Dear all,
} } We have problems with negative staining of E. Coli, and have tried two
} } different approaches with both Uranyl acetate and PTA.
} } Without being able to detect the flagellas, we see partly collapsed
} } bacterias and lots of dirt (growing bacterias directly on the grids)
} } We've also tried embed the E.coli in a mixture of methylcellulose and
} } uranyl acetate without any luck.
} } We've used carboncoated formvar films on copper grids.
} } I enclose the procedure for preparing cells grown on grids, observations
} } and some questions form the scientist in charge of this project.
} } We might be wrong trying to grow cells directly on grids?
} }
} }
} } Our aim is to prepare grids for TEM with a suitable cell number for
} } reliable anaysis without damaging the cells by centrifugation.
} } We plan to grow the cells directly onto the grids and have performed the
} } following:
} }
} } Inoculate cultures of enterococci in TSB+0.25%glucose and grow overnight
} } at 37*C without shaking. (TSB is a bacterial growth medium made from
} } casein and soya peptone).
} } Dilute cultures 100-fold in 10 ml growth medium in microtiterplate wells
} } containing TEM grids (should contain ca.107 cells/ml).
} } Incubate at 37*C without shaking for 1-4 hours.
} } Pick up grids at different time-points (i.e. 1h, 2h, 3h and 4h) from the
} } culture.
} } Dip the grid in particle free water.
} } Stain the cells with 0.5% uranyl acetate for 30".
} } Rinse the grids gently in particle free water before air-drying them for
} } 10-15'.
} }
} } Observations made:
} } 1. there are low cell-numbers on the grids
} } 2. there's a lot of stained debris or precipitate on the grids and it
} } seems to increase during the incubation.
} }
} } Questions:
} } 1. What cell density is required to obtain one layer of cells on the
} } grid but not cells on top of each other?
} } 2. Are there ways of increasing the bacterial affinity for the grids?
} } 3. Are there ways to reduce the amount of debris on the grids?
} } 4. Can grids be incubated on top of a nitrocellulose filter without
} } beeing damaged?
} }
} }
} }
} }
} } Best regards
} }
} } Randi Olsen
} } Department of Electron Microscopy
} } University of Tromso
} } Norway
} }
} }
} } ==============================Original Headers==============================
} } 15, 25 -- From Randi.Olsen-at-fagmed.uit.no Fri May 19 02:42:19 2006
} } 15, 25 -- Received: from mux2.uit.no (mux2.uit.no [129.242.5.252])
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} 12, 19 -- From PWebster-at-hei.org Fri May 19 13:54:08 2006
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Thanks to all for the advice on sectioning JB 4 resin, with its tendency
to fold and curl. I am summarizing the replies below:

1) From Glen McDonald: "For serial sections, which thankfully I
haven't had to do with JB-4 in many, many years, I got a tackle box,
found in any small parts supplier or fishing supply shop - a clear
plastic box with 2x6 or 4x6 array of compartments about 2 inches square.
Fill with deionized water or 3% ethanol. As the section comes off of
the knife edge, either lay an eyelash across the bottom edge to prevent
the curling, or grab with a pair of Dumont #55 forceps. move the
eyelash or forceps along with the motion of the section, then lift the
section and drop onto the liquid. Place one section in each compartment
to fill the tackle box, then mount them by immersing the slide and
bringing it up under the section. *Gently* touch one corner of the
section to guide into position. If not gently enough, the section will
cling to the eyelash and wad up like a used tissue during a bad cold."
There were a couple other variations on this technique of lifting slides
up underneath the sections. Off to the Bass Pro Shops electron
microscopy department for me!

2) Several people described helping the section come off the block by
pulling with a forceps on one corner during the cutting stroke or
holding it flat with an eyelash or brush, then flicking the section
quickly onto a drop of water or water/ethanol mixture. Dexterity
required, methinks.

3) Another repeated suggestion was to put the sections onto drops of
water with a little ammonium hydroxide in it.

4) As mentioned above, putting sections into water with ethanol, up to
50%, was mentioned several times, sometimes followed by transferring
sections to distilled water afterwards.

5) Don't use JB 4. (My favorite.)

6) Tobias Baskin has published his own formulation of BMM resin which
apparently sections much better. He uses DTT in the mix and says he is
happy to help anyone with this resin. We haven't tried this, yet, but
we intend to. Tobias posts to the list often, but if you can't find
his email, let me know and I'll forward your questions to him.

7) Humidity should be in the 40-50% range and the block should neither
be too wet or too dry or "sectioning is nearly impossible". From Ralph
Common. Humidity? In Missouri? Who could have guessed?

8) Related to 7, if the block is too soft, it won't cut well. This one
did it for us! Four more hours in the oven solved the worst of the
problem. Sections coming off flat and staying flat. These sections did
fold when placed on 50% ethanol/water, but did not fold when placed on
distilled water. Yaaaaaayy! Thanks, Teri Johnson!!

This was a crash course in JB 4 emergency microtomy. Our client is
happy, and Cheryl is tired. Cheryl gets a free lunch at the restaurant
of her choice (in Columbia, anyway).

Thanks again to all who responded. You guys are great!

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

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Head of Bioimaging Facility

Rothamsted Research, UK

We are seeking a biological scientist with experience in electron
microscopy and other imaging techniques to direct the Rothamsted Centre
for Bioimaging. This state of the art facility opened in 2003 and houses
3 electron microscopes, a confocal microscope, digital imaging and
associated image analysis software, a laser capture system and a fully
equipped cryo-preparation laboratory. The centre provides a high quality
service for a wide range of users both inside and outside the research
Institute. The Institute is the largest agricultural research centre in
the United Kingdom. The scientific research ranges from studies of
genetics, biochemistry, cell biology and soil processes to
investigations at the ecosystem and landscape scale.

You will be responsible for developing innovative research projects in
collaboration with other staff and for providing scientific and
technical advice on bioimaging within institute research programmes. You
will need to keep abreast of technologies and instrumentation to ensure
that the Centre develops new science and imaging applications and
remains at the cutting edge. The post also entails management of two
support staff.

Applicants should have a PhD or other higher qualification in a relevant
biological science, plus at least 5-7 years' postdoctoral experience in
microscopy, imaging or structural biology. You must be well versed in
both conventional and cryo-preparative techniques and their application
to biological samples. Experience in running a microscopy facility would
be an advantage.

The appointment will be at Band 5 with a starting salary normally within
the range �27,584 - �32,122 per annum.

For further information please contact Professor John Lucas ++ 44(0)1582
763133 x 2779 or john.lucas-at-bbsrc.ac.uk).

Apply by application form only, please visit our website
http://www.rothamsted.bbsrc.ac.uk/careers/vacancies/Vacancies.html for
further information and details regarding the application process.

Closing date: 8 June 2006

**
Dr Smita Kurup
CPI Division
Rothamsted Research
Harpenden
Herts AL5 2JQ

Tel No. 01582 763133 ext 2589
Fax No. 01582 763010

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Hello,
Does anyone have some experience with the:
Balzers RES 010 Rapid Etching System
I was wondering if it could be used for coating of samples for SEM as well
as etching?

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\
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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Email: roderi_co-at-yahoo.com
Name: Roman Derico

Organization: Faculty of food technology and biotechnology

Title-Subject: [Filtered] TEM - Problem with heating specimen holder

Question: Hi!
I have a weird problem. I have two heating specimen holders and recently I tried to use them on Jeol JEM200CX TEM (both holders are original parts from Jeol; last time they have been used on this microscope several years ago). When I inserted the holder in microscope, the electron beam disappeared from screen. I moved the holder step-by-step from position I to position II and changed the x and y position systematically (with specimen shifting knobs), but no beam appeared on screen. The beam doesn't appear on screen even when heating holder is empty (with specimen grid removed from it). The strange thing is that hole on heating holder matches exactly in position with holes on specimen holders of other types (it corresponds to specimen position I hole), and all other types of specimen holders work well.
Does anyone have idea why beam doesn't pass through heating holder?

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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dfine-at-seton.org) from http://microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Monday, May 22, 2006 at 16:28:41
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Email: dfine-at-seton.org
Name: Belinda Torres

Organization: University of Texas

Education: Graduate College

Location: Austin,Texas

Question: I am doing some experiments in electron microscopy and I am having a hard time removing wrinkles from my resin(thicks) sections. Can you help.

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Email: jacqui.ross-at-auckland.ac.nz
Name: Jacqueline Ross

Organization: The University of Auckland

Title-Subject: [Filtered] Job posting: Lecturer position in Auckland, New Zealand

Question: The Department of Anatomy with Radiology has a vacancy as detailed below. This person will also be associated with the Biomedical Imaging Research Unit. Please see the link to the job description if you are interested.

LECTURER

Anatomy with Radiology

School of Medical Sciences

Faculty of Medical and Health Sciences

The University of Auckland

Vacancy Number A304-06

We require a Lecturer to facilitate imaging research in cellular and molecular imaging and assist in providing for the ongoing development of imaging technologies within the Biomedical Imaging Research Unit and the wider faculty. The appointee will also be expected to provide leadership to both undergraduate and postgraduate teaching programmes, to assist in the development, planning and management of courses and programmes relevant to their field of expertise and to act in a supervisory role to graduate students.

Applications are invited from PhD qualified candidates with a track record of research and teaching in the areas of cellular/molecular imaging and cell and tissue biology.

For further information and to apply online, please visit http://www.vacancies.auckland.ac.nz/ or alternatively call +64 9-373 7599 ext 83000.

Please quote the vacancy number. Applications close 23 June 2006.

The University has an equal opportunities policy and welcomes opportunities policy and welcomes applications from all qualified persons.

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19, 12 -- From zaluzec-at-microscopy.com Mon May 22 21:22:28 2006
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I would suggest that you post the details for "foreign" applicants
as they are not likely to be acceptable to your solicitation.

There are restrictions on US jobs as well on non-US jobs.
I think that each solicitation should identify the nuances
of their particular geopolitical/geographic location.

That might save many folks a lot of wasted time. Just MO.

gary g.

At 07:24 PM 5/22/2006, you wrote:

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Hi all,

Okay, I'm a biologist and have absolutely no idea of how to go about setting
up a Philips CM-10 for dark field imaging. Now I have a user who needs it
and we need some help.

I would appreciate some suggestions for basic settings to get us started
such as aperture sizes, spot size, tilt angles, etc.

Thanks in advance,
Debby

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

==============================Original Headers==============================
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Hi,
I think that operation in dark-field (DF) mode on CM10 is similar to CM12.

Simply, we are using following procedure for beginners in DF on our CM12:
-start with well aligned microscope, C2 aperture 100-200 µm, spot
size 2, 80kV
-insert suitable sample (it should be thin!! e.g. DNA on carbon support film)
-adjust eucentric height of the sample
-check rotation center and pivot points in bright-field mode
-check stigmators
-switch to diffraction mode and center diffraction spot
-insert and center OBJ aperture
-press DF button on CM10 panel
-press RESET button under OPCON to center diffraction spot in DF mode
-move the diff. spot with Multifunction knobs just behind the rim of the
aperture disk on main screen
-switch to imaging mode and you should see dark-field image of your sample
-focus the sample and adjust illumination with Intensity button

You should try different OBJ apertures to get optimal image in DF with
your sample. I hope the procedure will work for you.

Best regards from Prague Oldrich

-----------------------------------
Oldrich Benada
Institute of Microbiology
Acad. Sci CR
Videnska 1083
142 20 Prague 4
Czech Republic

}
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} Hi all,
}
} Okay, I'm a biologist and have absolutely no idea of how to go about
} setting
} up a Philips CM-10 for dark field imaging. Now I have a user who needs it
} and we need some help.
}
} I would appreciate some suggestions for basic settings to get us started
} such as aperture sizes, spot size, tilt angles, etc.
}
} Thanks in advance,
} Debby
}
} Debby Sherman, Manager Phone: 765-494-6666
} Life Science Microscopy Facility FAX: 765-494-5896
} Purdue University E-mail: dsherman-at-purdue.edu
} S-052 Whistler Building
} 170 S. University Street
} West Lafayette, IN 47907
} http://www.agriculture.purdue.edu/microscopy
}
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jeger-at-bgu.ac.il) from http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on Tuesday, May 23, 2006 at 05:05:26
---------------------------------------------------------------------------

Email: jeger-at-bgu.ac.il
Name: Rina Jeger

Organization: Ben Gurion University of the negev

Education: Graduate College

Location: City, State, Country

Question: We have a problem with blocks damaging the diamond knife. Cell cultures grown in flasks and processed without any glass cause us much damage. We don't use molecular sieves. We use Araldite and just the blocks of cells from the flasks cause the damage. Please help!

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

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7, 12 -- From zaluzec-at-ultra5.microscopy.com Tue May 23 08:12:22 2006
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Rina,

We had that problem and it turned out to be the microtome rather than the
blocks. If bearings are worn out or advance motors are not working properly
you can get enough instability in the cutting stroke to cause fine damage to
the knife.

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: {jeger-at-bgu.ac.il}
} Reply-To: {jeger-at-bgu.ac.il}
} Date: Tue, 23 May 2006 08:15:46 -0500
} To: {dsherman-at-purdue.edu}
} Subject: [Microscopy] AskAMicroscopist: Help, blocks damaging the diamond
} knife
}
}
}
}
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} by (jeger-at-bgu.ac.il) from
} http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
} Tuesday, May 23, 2006 at 05:05:26
} ---------------------------------------------------------------------------
}
} Email: jeger-at-bgu.ac.il
} Name: Rina Jeger
}
} Organization: Ben Gurion University of the negev
}
} Education: Graduate College
}
} Location: City, State, Country
}
} Question: We have a problem with blocks damaging the diamond knife. Cell
} cultures grown in flasks and processed without any glass cause us much damage.
} We don't use molecular sieves. We use Araldite and just the blocks of cells
} from the flasks cause the damage. Please help!
}
} ---------------------------------------------------------------------------
}
} ==============================Original Headers==============================
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7, 24 -- From dsherman-at-purdue.edu Tue May 23 08:43:36 2006
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7, 24 -- diamond knife
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Hi,
I always request that tissue culture flasks/dishes are rinsed with a
sterile PBS prior to growing any cell lines to remove any debris that
is in there. I had problems like that and I rinsed the flask, spun the
content and checked under light microscope. There was plenty of
debris, probably of plastic origin. It might ease the problem but it
will not eliminate it 100%. If it is really bad I'd use a glass knife.
Dorota

==============================Original Headers==============================
1, 22 -- From wadowska-at-avcn1.novell.upei.ca Tue May 23 08:46:13 2006
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I had this problem. The problem diminished when I switched from
smashing osmium ampoules to dissolve the crystals to buying made up
solutions.

Dave

-----Original Message-----
X-from: jeger-at-bgu.ac.il [mailto:jeger-at-bgu.ac.il]
Sent: 23 May 2006 14:14
To: David Patton

Below is the result of your feedback form (NJZFM-ultra-55). It was
submitted by (jeger-at-bgu.ac.il) from
http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
Tuesday, May 23, 2006 at 05:05:26
------------------------------------------------------------------------
---

Email: jeger-at-bgu.ac.il
Name: Rina Jeger

Organization: Ben Gurion University of the negev

Education: Graduate College

Location: City, State, Country

Question: We have a problem with blocks damaging the diamond knife. Cell
cultures grown in flasks and processed without any glass cause us much
damage. We don't use molecular sieves. We use Araldite and just the
blocks of cells from the flasks cause the damage. Please help!

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

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24, 34 -- From David.Patton-at-uwe.ac.uk Tue May 23 09:07:12 2006
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Dave,

A simple trick to eliminate smashing osmium vials is to dip the sealed
ampoule into liquid nitrogen. This releases the osmium crystals from the
glass walls. Then simply break the vial using an ampoule cracker (available
through EM supply houses) and pour the osmium crystals into your bottle
containing the water.

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: {David.Patton-at-uwe.ac.uk}
} Reply-To: {David.Patton-at-uwe.ac.uk}
} Date: Tue, 23 May 2006 09:10:02 -0500
} To: {dsherman-at-purdue.edu}
} Subject: [Microscopy] RE: AskAMicroscopist: Help,
}
}
}
}
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}
} I had this problem. The problem diminished when I switched from
} smashing osmium ampoules to dissolve the crystals to buying made up
} solutions.
}
} Dave
}
}
}
} -----Original Message-----
} X-from: jeger-at-bgu.ac.il [mailto:jeger-at-bgu.ac.il]
} Sent: 23 May 2006 14:14
} To: David Patton
} Subject: [Microscopy] AskAMicroscopist: Help, blocks damaging the
} diamond knife
}
}
}
}
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} Below is the result of your feedback form (NJZFM-ultra-55). It was
} submitted by (jeger-at-bgu.ac.il) from
} http://www.microscopy.com/Ask-A-Microscopist/Ask-A-Microscopist.html on
} Tuesday, May 23, 2006 at 05:05:26
} ------------------------------------------------------------------------
} ---
}
} Email: jeger-at-bgu.ac.il
} Name: Rina Jeger
}
} Organization: Ben Gurion University of the negev
}
} Education: Graduate College
}
} Location: City, State, Country
}
} Question: We have a problem with blocks damaging the diamond knife. Cell
} cultures grown in flasks and processed without any glass cause us much
} damage. We don't use molecular sieves. We use Araldite and just the
} blocks of cells from the flasks cause the damage. Please help!
}
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8, 23 -- From dsherman-at-purdue.edu Tue May 23 09:24:30 2006
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On May 22, 2006, at 8:07 PM, dsherman-at-purdue.edu wrote:

} Okay, I'm a biologist and have absolutely no idea of how to go about
} setting
} up a Philips CM-10 for dark field imaging. Now I have a user who
} needs it
} and we need some help.
}
} I would appreciate some suggestions for basic settings to get us
} started
} such as aperture sizes, spot size, tilt angles, etc.
}
Dear Debby,
I have only used dark field on our Tecnai scopes, which have a
push-button to select dark field, but the process is likely to be the
same on the CM10. The settings used will depend on what the user is
trying to see. To get an idea, take a typical area of the specimen--or
a positive control--and look in diffraction mode with no objective
aperture to see where the signal you want is. When dark field is
activated, the diffraction pattern shifts by an amount that varies with
the tilt angle, so if you increase the tilt from zero, there will be a
value that puts the signal you want to see at the position that the
unscattered beam occupies for zero tilt, and this will be the tilt
angle you want. If you now turn off dark field, insert and center an
objective aperture, and turn on dark field, you will see what part of
the diffracted beam will constitute the dark field image. The smaller
the aperture, the more selective it will be, but the smaller the signal
that will get through. In any case, you want an aperture that blocks
the unscattered beam, and its maximum size will depend on the dark
field tilt angle you chose. If you have crystalline material, the
aperture will select signal from only a subset of possible
orientations, which could either be what you want or just a nuisance.
If the latter, dynamical dark field, AKA hollow-cone illumination, will
solve it, but on the Tecnai scopes, it is only available with STEM.
Spot size should be selected to give you sufficient signal, and other
than that it will have little effect on the DF image unless you are
using an extremely small objective aperture, in which case, a smaller
spot size number will smear the diffraction rings or spots and lessen
the amount and selectivity of the signal. For taking DF images of
terbium, a 1 deg tilt and 70 um objective aperture allowed me to see
the terbium as bright areas with faint gray corresponding to places
where lower-Z material was and very few counts corresponding to blank
areas of the grid. Good luck.
Yours,
Bill Tivol, PhD
EM Scientist and Manager
Cryo-Electron Microscopy Facility
Broad Center, Mail Code 114-96
California Institute of Technology
Pasadena CA 91125
(626) 395-8833
tivol-at-caltech.edu

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Hello,
I'm seeking reccomendations for a small CO2 incubator to use in an
imaging facility to hold samples waiting for their turn on the
microscope. This period could be as long as several hours. Any
suggestions for 1 cubic foot (0.028 m^3) to 1.4 cubic foot (0.039
m^3) incubator? Have air jacketed been sufficient or does anyone
wish they'd purchased a water jacketed incubator instead?

thanks,
glen

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

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

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Have you got the holder connected to the power supply, temperature sensor, etc?

It's possible the outer part of central furnace is grounded via the
the electrical connections - if you don't have it connected up, it
may be charging up and deflecting the beam. Even if you have it
plugged in, if the holder hasn't been used for several years, there
may be a broken connection.

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

PLEASE NOTE - Agressive SPAM filtering is employed. If you don't get
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e-mail will never be seen.

==============================Original Headers==============================
5, 17 -- From larry-at-cymru.freewire.co.uk Tue May 23 14:59:58 2006
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5, 17 -- holder
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Email: istillma-at-bidmc.harvard.edu
Name: Isaac E Stillman

Organization: BIDMC/Harvard Medical School

Title-Subject: [Filtered] Advice

Question: Hi-

I am a renal pathologist who is purchasing a new instrument for our diagnostic EM unit. The choice has come down to FEI (Phillips) Morgagni vs JEOL JEM-1011. Of course, we would be adding a 2K digital camera. I would be most grateful to hear your thoughts on this choice, off list if you prefer.

Thanks!
Isaac E Stillman MD
Dept. of Pathology - BIDMC/HMS

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

==============================Original Headers==============================
11, 12 -- From zaluzec-at-microscopy.com Tue May 23 18:23:52 2006
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Email: huisheng.jiao-at-gmail.com
Name: Ding

Organization: The University of Birmingham

Title-Subject: [Filtered] Suva 124 (R124)?

Question: Does anybody have experience in Suva 124 (R124) as cryogen for rapid freezing?

Huisheng

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8, 12 -- From zaluzec-at-microscopy.com Wed May 24 07:42:09 2006
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Hi:

In the late 80's early 90's M. Parthasarathy, Cornell University, & Theo
Muller, BAL-TEC AG published a paper examining using SUVA 124 for
cryofixation.

Best,

Al Coritz, Technical Engineer
Electron Microscopy Sciences
www.emsdiasum.com

----- Original Message -----
X-from: {huisheng.jiao-at-gmail.com}
To: {Sampleprep-at-earthlink.net}
Sent: Wednesday, May 24, 2006 8:43 AM

Dear Al,
dear Huisheng,

I'v found in the MSA listserver-Archives
cf:
http://www.microscopy.com/cgi-bin/ReadPrintEmailHTML.pl?filename=9807.txt

a short thread on the possible use of SUVA, in which perhaps also the
Article mentioned below has been referenced:
It says (for the whole thread use "FREON" or "replacement" as a search
phrase):
} } } } } } } } } } } } } } } } } } } } { { { { { { { { { { { { { { { { { { { { { { { {
Re: Freon replacements for cryo

Try Muller T, Moser S, Vogt M, Daugherty C & Parasarathy MV (1993).
Optimisation and application of jet freezing. Scanning Microscopy 7,
1295-1310.

Using HCFC 124 (SUVA 124-CHClFCF3) with thin titanium
supports, cooling rates were obtained similar to those from propane and
standard copper supports. This was suggested (I hesitate to plug this !!)
in
Ryan KP (1992) Cryofixation of tissues for electron microsocpy: a
review of plunge cooling methods.
Scanning Microsc. 6, 715-743. See
next-to-last page , p. 742 in Discussion with Reviewers section..

Keith Ryan
Plymouth Marine Lab., UK
{ { { { { { { { { { { { { { { { { { { { { { { {} } } } } } } } } } } } } } } } } } } }

Best regards and wishes to solve the problem,

Wolfgang MUSS
Salzburg, Austria

----------
Von: sampleprep-at-earthlink.net[SMTP:sampleprep-at-earthlink.net]
Antwort an: sampleprep-at-earthlink.net
Gesendet: Mittwoch, 24. Mai 2006 15:41
An: W.Muss-at-salk.at
Betreff: [Microscopy] Re: cryogen for rapid freezing

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Hi:

In the late 80's early 90's M. Parthasarathy, Cornell University, & Theo
Muller, BAL-TEC AG published a paper examining using SUVA 124 for
cryofixation.

Best,

Al Coritz, Technical Engineer
Electron Microscopy Sciences
www.emsdiasum.com

----- Original Message -----
X-from: {huisheng.jiao-at-gmail.com}
To: {Sampleprep-at-earthlink.net}
Sent: Wednesday, May 24, 2006 8:43 AM

Micromavens,

Has anyone on the list had recent experience shipping small flowers
fixed for EM? Or bringing such samples along as baggage on an
airplane? I've done similar things in the past, but not recently, and
times and regulations have changed.
What's allowed? What's not? This is specifically for flowers (small
ones) and flights or courier shipments originating and ending in the
USA.
Thanks.

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

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I've been asked to seek opinions from folks who have used 2 or more
of the currently available knife-makers:
The GMK triangular knife maker sold by Energy Beam Sciences, and the
two "balanced break" knife-makers, the Leica EM KMR 2 and the RMC GKM
(an unfortunately confusing model designation).
What do folks think of them, and how easily can novices be taught to
consistently make good knives usable for thin sectioning?
Of these, I've only used the GMK, formally known as the Sorvall brick.
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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Dear colleagues...

We're plannig to build a sterotaxic instrument for animal brain studies.
It must be adaptable for mouse, rat and rabbit. We're looking for plans,
drawings etc. anything beneficial. Does anybody know this kind of
materials reachable on internet or another source?

Thanks in advance...

Dr. Necat Yilmaz
Mersin University School of Medicine
Histology and Embryology Dept.
Mersin/TURKEY

==============================Original Headers==============================
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Dear friends
I am intersted in the following pdf reprint or xerox if possible (fax
is in the signature).
As well I use this occasion to remind that we have two schools in
Genoa, namely: Fluorescence 19-22 June; Confocal and Multiphoton
microscopy 4-7 July. SInce the number of partecipants is limited,
please have a check at www.lambs.it or send an e.-mail to me for
reservations.
All my best
Alby

paper ref
Squeezing in two photon absorption from a strong coherent beam
G. S. Agarwal
Optics Communications
Pages 190-192, Volume 62, Issue 3, 1 May, 1987.

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

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We have been using a flatbed scanner for documenting rock core before
processing it for other analytical techniques. Now, we want to explore the
technique a bit further, possibly towards applied image analysis, but need
one last ingredient for better quality scans. We want to use some akin to
immersion oil for the interface between rock and flatbed, but want to find
something inexpensive, non-toxic and easy to clean. Any thoughts?

TIA & genuinely :o)
michael shaffer

SEM/MLA Research Coordinator
(709) 737-6799 (ofc)
(709) 737-6790 (lab)
(709) 737-6193 (FAX)
{http://www.mun.ca/creait/maf/}
{http://www.esd.mun.ca/epma/}

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

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7, 19 -- Subject: imaging rock core with flatbed scanner
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Hi,
Our lab has been using Leica EM KMR 2 for a few years now. So
far we have no problem with the machine. It is reliable and makes
consistently good knives.
Dorota

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Hi Michael,

If you feel you would like to use immersion oil, why don't you - non
fluorescent immersion oil is pretty cheap from the likes of Zeiss and
Cargille, and isn't particularly toxic (the anti-fluorescent variety is a
little more toxic). As long as you keep the oil in the centre of the glass
platen and don't shut the platen lid down, it's not going anywhere. In fact
you can get quite good scans of things like fruit, seeds, leaves,
calculators, small toys etc.. using film scanning flatbeds in reflective
mode at high resolutions (certainly better than the likes of that from an
$80 QX-5 toy microscope). You may need to use Silverfast.com twain software
to overcome the scanners reluctance to go above 1,200 dpi with a reflective
A4 scan though. Also many scanners are fine focussed a little above the
glass platen, as it is assumed you are using film in holders. The superb
Epson V750 pro flatbed scanner even has optional little legs on the film
holders to help with this. I'm pretty sure that the scanner optics aren't
optimised for oil immersion though, it being an air interface on the
underneath of the scanner. You won't get good scan results with a cheap LiDE
type scanner (no depth of field), you really need a film scanner type
flatbed.

You can easily remove immersion oil with 50:50 alcohol water and tissues, in
fact I use 50:50 propanol and water regularly on my Canon 9950F and Epson
4990 photo flatbeds (naturally avoiding the platen edges when spraying).
This mixture also cleans all our imaging workstation's VDU CRT screens
(removing all finger marks). Also try 100% ether for stubborn stains.
Fingermarks are a similar oily mess to immersion oil, but it's only non
removable scratches from sharp rocks that are the real pain (try using the
sample on glass cover slips or slides perhaps ?). You can try using things
like thin plastic strips (or thin card with film) to scoop under the sample
to remove it, without touching the platen glass.

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: michael-at-Shaffer.net [mailto:michael-at-Shaffer.net]
Sent: 25 May 2006 11:57
To: keith.morris-at-ucl.ac.uk

We have been using a flatbed scanner for documenting rock core before
processing it for other analytical techniques. Now, we want to explore the
technique a bit further, possibly towards applied image analysis, but need
one last ingredient for better quality scans. We want to use some akin to
immersion oil for the interface between rock and flatbed, but want to find
something inexpensive, non-toxic and easy to clean. Any thoughts?

TIA & genuinely :o)
michael shaffer

SEM/MLA Research Coordinator
(709) 737-6799 (ofc)
(709) 737-6790 (lab)
(709) 737-6193 (FAX)
{http://www.mun.ca/creait/maf/}
{http://www.esd.mun.ca/epma/}

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

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Michael,

Glycerin comes to mind as being cheap, non-toxic, easier to clean,
clear, refractive index ~1.47 and just all around more "friendly" than a
traditional refractive index liquid. I've never used it, but it comes
to mind as something I have seen mentioned as usable for immersion
media. There are lots of household liquids that you might also
consider, vegetable oil, olive oil, and mineral oil come to mind.

Good Luck,
Bryan Bandli

michael-at-Shaffer.net wrote:

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You could try water as well. Glycerol can be diluted in it. We regularly use
water or glycerol immersion and it works great with special water/glycerol
immersion objectives.

On a scanner in reflected mode, oil can really shine sometimes though
(particularly at oil/air interfaces). Some vegetable oils such as corn oil
also evaporate to leave a very hard resin residue that can't easily be
removed (naturally immersion oil doesn't) - so the glass must be cleaned
well afterwards. If your sample is huge (6 inches or more) I would have
thought any cheap clear mineral oil would be ideal (there are plenty to
chose from, even baby oil). On a flat polished sample you shouldn't need
much oil.

If a sample is very very thin (we used annular saws with our hard human bone
samples and ground the surfaces flat - so I'm thinking along those lines) -
and you can get some light through it (or you are looking at something like
loose soil particles), running the scanner in transmission film scan mode
might work. You have more than half a centimetre or so between the platen
glass and the film scanner lid with an Epson 4990 photo, so you can get a
thin sheet of glass (e.g. slide thickness) and a thin sample in there.
Samples in oil appear to scan better in this film scan mode than in
reflected mode, particularly at air/oil surfaces.

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: bbandli-at-mvainc.com [mailto:bbandli-at-mvainc.com]
Sent: 25 May 2006 14:08
To: keith.morris-at-ucl.ac.uk

Michael,

Glycerin comes to mind as being cheap, non-toxic, easier to clean,
clear, refractive index ~1.47 and just all around more "friendly" than a
traditional refractive index liquid. I've never used it, but it comes
to mind as something I have seen mentioned as usable for immersion
media. There are lots of household liquids that you might also
consider, vegetable oil, olive oil, and mineral oil come to mind.

Good Luck,
Bryan Bandli

michael-at-Shaffer.net wrote:

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HI, Michael

Normal mineral oil from the drug store has RI 1.51. I would try it on a glass surface other than the scanner bed first, to make sure that you don't get moire patterns. Also, try various cleaning methods. I think that wiping it dry first then using either Windex or rubbing alcohol should do the trick.

Good hunting!
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 05:55 AM 5/25/2006, michael-at-Shaffer.net wrote:

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

All the best

Jacob

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

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

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

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

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

michael-at-Shaffer.net wrote:
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} We have been using a flatbed scanner for documenting rock core before
} processing it for other analytical techniques. Now, we want to explore the
} technique a bit further, possibly towards applied image analysis, but need
} one last ingredient for better quality scans. We want to use some akin to
} immersion oil for the interface between rock and flatbed, but want to find
} something inexpensive, non-toxic and easy to clean. Any thoughts?
}
} TIA & genuinely :o)
} michael shaffer
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--
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~

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

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

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

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

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Email: mccaulak-at-wfu.edu
Name: Anita McCauley

Organization: Wake Forest University

Title-Subject: [Filtered] procedure for quantitative fluorescence

Question: I am looking for advice for doing quantitative fluorescence microscopy. My focus is on the microscope, camera, and software end of it, rather than the sample prep. Last year, a question about quantitative fluorescence on this list resulted in a number of very helpful posts about things to do, like controlling camera settings and using fluorescence reference slides. However, what is not clear is how to combine all these things together into an appropriate procedure. If anybody could help with any of the following, I would greatly appreciate it:

1. I purchased a set of fluorescent reference slides but have been unable to obtain any documentation or instructions from the supplier for how to use them correctly for fluorescent imaging.

2. When comparing a signal between two samples, is it best to subtract the background from the signal or just compare the two signals?

3. To determine the signal, is it best to use irregular AOIs and get a mean or summed value or to use multiple line profiles, or something else?

4. If anyone would be willing to share a step-by-step protocol that they have developed to help users do image capture and analysis of fluorescence correctly, I would greatly appreciate it.

Thanks.

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Hello,
I've been processing some small plant roots for scanning electron
microscopy. I use Glutaraldehyde fixation followed by 0.8% KFerrocyanide
and 1% OsO4 - otherwise standard fixation. I get great root preservation,
but I've found that the root hairs all look very collapsed. I can send a
picture if you like, but was wondering if anyone had experience in this
area.

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\
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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Hello Philip,

We are using the Leica EM KMR2 without problems. It is
easy to learn and to use and is a reliable instrument.

Stephane

--- oshel1pe-at-cmich.edu wrote:

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} I've been asked to seek opinions from folks who have
} used 2 or more
} of the currently available knife-makers:
} The GMK triangular knife maker sold by Energy Beam
} Sciences, and the
} two "balanced break" knife-makers, the Leica EM KMR
} 2 and the RMC GKM
} (an unfortunately confusing model designation).
} What do folks think of them, and how easily can
} novices be taught to
} consistently make good knives usable for thin
} sectioning?
} Of these, I've only used the GMK, formally known as
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}
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Hi,

My answer will deal with confocal microscopy: I use to
insert my negative control and choose the lasers and
detection settings so that I get no signal. Then I
insert my samples and won't change the settings.
To compare fluorescence intensities, I use to draw a
profile line and compare the peaks (maximum
intensities). I do it because I observe compact
homogeneous bodies, but I suppose I would draw a ROI
in the case I had to compare the total intensity of
cell cytoplasm for example. It all depends on what and
how you measure, there is no general rule I think, it
must be adapted to be as "ethical" (yes again ;-)) as
possible.

regards,

St�phane

--- mccaulak-at-wfu.edu wrote:

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} Email: mccaulak-at-wfu.edu
} Name: Anita McCauley
}
} Organization: Wake Forest University
}
} Title-Subject: [Filtered] procedure for quantitative
} fluorescence
}
} Question: I am looking for advice for doing
} quantitative fluorescence microscopy. My focus is
} on the microscope, camera, and software end of it,
} rather than the sample prep. Last year, a question
} about quantitative fluorescence on this list
} resulted in a number of very helpful posts about
} things to do, like controlling camera settings and
} using fluorescence reference slides. However, what
} is not clear is how to combine all these things
} together into an appropriate procedure. If anybody
} could help with any of the following, I would
} greatly appreciate it:
}
} 1. I purchased a set of fluorescent reference slides
} but have been unable to obtain any documentation or
} instructions from the supplier for how to use them
} correctly for fluorescent imaging.
}
} 2. When comparing a signal between two samples, is
} it best to subtract the background from the signal
} or just compare the two signals?
}
} 3. To determine the signal, is it best to use
} irregular AOIs and get a mean or summed value or to
} use multiple line profiles, or something else?
}
} 4. If anyone would be willing to share a
} step-by-step protocol that they have developed to
} help users do image capture and analysis of
} fluorescence correctly, I would greatly appreciate
} it.
}
} Thanks.
}
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} Headers==============================
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} 11, 12 -- Subject: viaWWW: procedure for
} quantitative fluorescence
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Dear listers,

I have a triple labeling in fluorescence
(green-red-blue), but I take pictures in
epifluorescence with a B/W camera, which is more
sensitive than the colour camera. Now for presentation
purposes I would like to give colours to my B/W
pictures in the 3 colours originally used and
(perhaps) merge them in the end.
What is the procedure in Photoshop?

St�phane

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4, 18 -- From: Stephane Nizet {nizets2-at-yahoo.com}
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Hi Stephanie,

It's a bit of a fag in Photoshop - much easier in MetaMorph or ImagePro Plus
(and probably in ImageJ/NIHImage for poor people). Photoshop also gets cross
with 12-bit B&W images (they go black and it's not worth bringing the image
back with contrast adjustment) so export at 8-bit (256 grey) TIF.

I'll email the complete pdf on 'how to do it' that includes VDU screen
dumps. For the listserver the basic text is appended below. I have the
Bio-Rad PIC plug-ins for Photoshop if anyone would like them.

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

Text from the pdf file 'Colouring B&W pictures in photoshop'

Creating Combined RGB Bio-Rad images in Photoshop

(You may need the Bio-Rad Plugin for *.PIC files otherwise use *.TIF)
Load images (via import, Bio-Rad PIC Import)
Convert both to RGB (Image, Mode,RGB Color)
Find the image size (Image, Image Size � 1024x1024 in this case)

Create a new Image for the blue channel (1024x1024 � black)
File, New (White, 1024x1024 � it must be the same size as the red and green
images)
Select Color Picker (left menu � black/white boxes) and change the
foreground colour to black (click OK). Right Click over the new image and
select all.

Select Edit,Fill and click OK. Image will now be black (the unused blue
channel).

Select Windows, Show Channels (to see RGB channels for all images)

Go to the black (blue) image (right click to select all if necessary, and
Edit, Copy
Select the Green Channel Image and highlight its blue channel

Select Edit, Paste and the blue channel is �removed� on the target image.

Repeat this pasting the black into the red channel (leaving the Green
channel left)

Now right click select all over the �red channel� B&W image.
Select Edit, Copy

Reselect the �green channel� image (that has black over its red and blue
channels).

Select the blackened red channel in the �green� image.
Now Edit, Paste (and the red channel is pasted into the red channel of the
green image creating a Red and Green combined RGB image (black for blue).

I also adjusted the Image, Adjust, brightness/Contrast for both the red and
green images prior to combining them. You must select the red or green
channel when adjusting. Once finished select and copy the red image into the
red channel of the green image to combine as before. This time the green
brightness/contrast has been reduce and the red increased. See resulting
image below and compare with the unadjusted original intensities above.

Author: Dr Keith J Morris, Imaging Facility Manager, Cell Biology Division,
the institute of Ophthalmology,
11-43 Bath Street, London EC1V 9EL.. Telephone: 020 7608 4050.
Email: keith.morris-at-ucl.ac.uk.

Please advise me of any errors found and/or difficulties encountered when
using this help document.

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: 26 May 2006 09:33
To: keith.morris-at-ucl.ac.uk

Dear listers,

I have a triple labeling in fluorescence
(green-red-blue), but I take pictures in
epifluorescence with a B/W camera, which is more
sensitive than the colour camera. Now for presentation
purposes I would like to give colours to my B/W
pictures in the 3 colours originally used and
(perhaps) merge them in the end.
What is the procedure in Photoshop?

St�phane

==============================Original Headers==============================
30, 27 -- From keith.morris-at-ucl.ac.uk Fri May 26 03:59:23 2006
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30, 27 -- Subject: RE: [Microscopy] colouring B/W pictures with Photoshop
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Dear Anita,

It's not really possible to get quantitative measurements of
epi-fluorescence. The gel fluorescence standards commonly used are simply to
check that your camera/detector and electronics are genuinely linear in
response (something quite important to know) - they are no use as standards
for calibrating fluorescence within the cell (although you can use things
like BCECF in solution at known pH's to roughly calibrate intracellular pH).
To properly calibrate cellular fluorescence you need standards that are
essentially known concentrations and masses of labelled protein etc.. within
cells, not something that's easy to get. You will always have problems with
laser power variation, differential bleaching, uneven inconsistent
labelling, internal quenching etc.. so you will probably never be able to
say that one cell has exactly twice the mass (total pixel brightness) or
concentration (mean pixel brightness) of a given protein compared to
another. However it is reasonable to assume that a very brightly labelled
cell has definitely more labelled fluorochrome in it than a poorly
fluorescent cell or intracellular region (and you could say something like
'suggesting' x times concentration etc.). Also try measuring something
simple like concentrations of fluorescent beads to see how (badly) the image
analysis results work out (these do have the advantage that you can actually
count them as well).

You will probably have to use regions of interest to include darker as well
as bright areas if comparing regions, plus you can use peak values as
described by Stephane (but often the bright sample fluorescence area has to
be very clearly defined to one region to get this to work). You can also try
things like what % of the cell is brighter than a set grey level.
Co-localisation (i.e. the % of bright pixels in both the red and green
channel across the cell) is also useful. Generally each set of samples
require their own image analysis protocols depending on what you want to
find out, plus you need simple stats to say if the difference is significant
to 95% level.

When looking at optical density, this can be more easily calibrated - here
you are measuring optical white light transmission though a sample. So you
can create black (metal disk), white (free space) and have a selection of
known density materials like polythene (grey) etc. in the image. You can
therefore assign a grey level to a particular density. We have used this to
estimate things like bone density, assuming your can get very evenly sliced
samples.

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: mccaulak-at-wfu.edu [mailto:mccaulak-at-wfu.edu]
Sent: 25 May 2006 23:57
To: keith.morris-at-ucl.ac.uk

I know this does not answer the question...

I have found that live wheat seedling root hairs collapse almost
immediately even in an ESEM.

Dave

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

Sent: 26 May 2006 00:16
To: David Patton

Hello,
I've been processing some small plant roots for scanning electron
microscopy. I use Glutaraldehyde fixation followed by 0.8%
KFerrocyanide
and 1% OsO4 - otherwise standard fixation. I get great root
preservation,
but I've found that the root hairs all look very collapsed. I can send
a
picture if you like, but was wondering if anyone had experience in this
area.

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/
\/\/\
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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Dear Gordon,

Have you considered cryo-SEM? This would seem to the ideal method -
maintaining the sample in its hydrated state and avoiding mechanical
damage that critical point drying can cause some delicate samples.

See:

http://www.quorumtech.com/Applications/Cryo_Apps_Library/Frozen-Hydrated
-Root-Hairs.htm

This is quite an old image and at one time (now lost) there was a
companion image showing the clean surface after controlled etching
(sublimation) of the ice.

Declaration of commercial interest: Quorum Technologies are
manufacturers of the PP2000 and PP2000T cryo-SEM systems.

Best regards,

Mike Wombwell
Quorum Technologies
Newhaven, East Sussex, UK
Tel: +44(0)1273 510535
Fax: +44(0)1273 510536
mike.wombwell-at-quorumtech.com
http://www.quorumtech.com
E & O E

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Hello,
I've been processing some small plant roots for scanning electron
microscopy. I use Glutaraldehyde fixation followed by 0.8%
KFerrocyanide
and 1% OsO4 - otherwise standard fixation. I get great root
preservation,
but I've found that the root hairs all look very collapsed. I can send
a
picture if you like, but was wondering if anyone had experience in this
area.

\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/
\/\/\
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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Email: lenoska1-at-seznam.cz
Name: Zuzana Lenochova

Organization: Charles UNiversity in Prague, Czech Republic

Title-Subject: [Filtered] FDA

Question: Dear listers,
some time ago I decided to distinguish between dead and living cells in plant roots using FDA (fluorescein diacetate) combined with PI (propidium iodide). But it seems cells showing the brigtest FDA fluorescence are those that are prone to die, forming strange fluorescing vesicles or even their whole cell content fluoresces. But haven't found enough supporting information in literature. Does any of you have similar experience (or any citations) showing the activity of esterases would more refer to physiological changes (let's say connected with cell death) of the cell than just to its viability?
Thanks in advance for your help,

Zuzana

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Hi Zuzana,

I don't know the real answer to your question but I suspect it's
something to do with the toxicity of propidium iodide. Do you use PI
and FDA at the same time? PI on it's own is a fairly reliable live/dead
marker in Arabidopsis roots when used at 5 micrograms/ml. Living cells
exclude it and so remain outlined while dead cells fill with the stain
and fluoresce brightly. The problem with PI is that it will kill the
cells if they are left too long in it (greater than 30-45 minutes).
Maybe that is why the wierd FDA effect. What happens when you use the
stains on their own?

Regards, John.

lenoska1-at-seznam.cz wrote:

} Email: lenoska1-at-seznam.cz
} Name: Zuzana Lenochova
}
} Organization: Charles UNiversity in Prague, Czech Republic
}
} Title-Subject: [Filtered] FDA
}
} Question: Dear listers,
} some time ago I decided to distinguish between dead and living cells in plant roots using FDA (fluorescein diacetate) combined with PI (propidium iodide). But it seems cells showing the brigtest FDA fluorescence are those that are prone to die, forming strange fluorescing vesicles or even their whole cell content fluoresces. But haven't found enough supporting information in literature. Does any of you have similar experience (or any citations) showing the activity of esterases would more refer to physiological changes (let's say connected with cell death) of the cell than just to its viability?
} Thanks in advance for your help,
}
} Zuzana
}
}
}

--

*********************************
C. John Runions, Ph.D.
School of Biological and Molecular Sciences
Oxford Brookes University
Oxford, UK
OX3 0BP

email: jrunions-at-brookes.ac.uk
phone: +44 (0) 1865 483 964

==============================Original Headers==============================
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Stephane,

As Keith implies, it is quite easy to do the kind of work you want
with ImageJ. There is a function under the Image Menu} Colors, called
RGB merge. Using that, you can assign any b/w image to one of the
three color channels, or to none. This does, however, presuppose
that your three images are properly aligned. There is also a plugin
for ImageJ that will allow you to align the images.

To replace a single b/w image with a colored one, you can use the
ImageJ LookUpTables, which will remap the 8-bit gray scale to a color
of your choosing.

Check with me if you need more information.

Joel

}
}
}
} ----------------------------------------------------------------------
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} of America To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver On-Line Help
} http://www.microscopy.com/MicroscopyListserver/FAQ.html
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}
} Hi Stephanie,
}
} It's a bit of a fag in Photoshop - much easier in MetaMorph or
} ImagePro Plus (and probably in ImageJ/NIHImage for poor people).
} Photoshop also gets cross with 12-bit B&W images (they go black and
} it's not worth bringing the image back with contrast adjustment) so
} export at 8-bit (256 grey) TIF.
}
} I'll email the complete pdf on 'how to do it' that includes VDU screen
} dumps. For the listserver the basic text is appended below. I have the
} Bio-Rad PIC plug-ins for Photoshop if anyone would like them.
}
} 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
}
}
} Text from the pdf file 'Colouring B&W pictures in photoshop'
}
} Creating Combined RGB Bio-Rad images in Photoshop
}
} (You may need the Bio-Rad Plugin for *.PIC files otherwise use *.TIF)
} Load images (via import, Bio-Rad PIC Import) Convert both to RGB
} (Image, Mode,RGB Color) Find the image size (Image, Image Size �
} 1024x1024 in this case)
}
} Create a new Image for the blue channel (1024x1024 � black)
} File, New (White, 1024x1024 � it must be the same size as the red and
} green images) Select Color Picker (left menu � black/white boxes) and
} change the foreground colour to black (click OK). Right Click over the
} new image and select all.
}
} Select Edit,Fill and click OK. Image will now be black (the unused
} blue channel).
}
} Select Windows, Show Channels (to see RGB channels for all images)
}
} Go to the black (blue) image (right click to select all if necessary,
} and Edit, Copy Select the Green Channel Image and highlight its blue
} channel
}
} Select Edit, Paste and the blue channel is �removed� on the target
} image.
}
} Repeat this pasting the black into the red channel (leaving the Green
} channel left)
}
} Now right click select all over the �red channel� B&W image.
} Select Edit, Copy
}
} Reselect the �green channel� image (that has black over its red and
} blue channels).
}
} Select the blackened red channel in the �green� image.
} Now Edit, Paste (and the red channel is pasted into the red channel of
} the green image creating a Red and Green combined RGB image (black
} for blue).
}
} I also adjusted the Image, Adjust, brightness/Contrast for both the
} red and green images prior to combining them. You must select the red
} or green channel when adjusting. Once finished select and copy the red
} image into the red channel of the green image to combine as before.
} This time the green brightness/contrast has been reduce and the red
} increased. See resulting image below and compare with the unadjusted
} original intensities above.
}
} Author: Dr Keith J Morris, Imaging Facility Manager, Cell Biology
} Division, the institute of Ophthalmology, 11-43 Bath Street, London
} EC1V 9EL.. Telephone: 020 7608 4050. Email:
} keith.morris-at-ucl.ac.uk.
}
} Please advise me of any errors found and/or difficulties encountered
} when using this help document.
}
}
}
}
}
} -----Original Message-----
} X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: 26 May 2006 09:33
} To: keith.morris-at-ucl.ac.uk
} Subject: [Microscopy] coloring B/W pictures with photoshop
}
}
}
}
} ----------------------------------------------------------------------
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} ----------------------------------------------------------------------
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}
} Dear listers,
}
} I have a triple labeling in fluorescence
} (green-red-blue), but I take pictures in
} epifluorescence with a B/W camera, which is more
} sensitive than the colour camera. Now for presentation
} purposes I would like to give colours to my B/W
} pictures in the 3 colours originally used and
} (perhaps) merge them in the end.
} What is the procedure in Photoshop?
}
} St�phane
}
}
}
} ==============================Original
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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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Dear folks,

I would like to transfer 16 bit sun raster files into TIF format
without loosing the resolution !. Anybody aware of a
technique/software which can do that?

Regards,

Jafar

Rutgers University,
Piscataway, NJ 08854
Jafar F. Al-Sharab, Ph.D.
Rutgers, The State University of New Jersey
Department of Materials Science and Engineering
607 Taylor Road, Room 237
Piscataway, NJ 08854-8065
Tel. 732-445-5615 Work
732-688-1955 Cell
Fax 732-445-3258

E-mail: jafarhan-at-rci.rutgers.edu

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Folks-
Can anyone advise us who can service older ultramicrotomes in the
Midwest? We have Sorval MT-2 and MT-2B and LKB Ultrotome III
ultramicrotomes.
Carol Heckman
--
__
Carol A. Heckman, Ph.D.
Professor of Biological Sciences
Director, Center for Microscopy & Microanalysis
Bowling Green State University, Bowling Green, OH 43403
fax: (419) 372-2024 email: heckman-at-bgsu.edu
http://www.bgsu.edu/departments/biology/facilities/MnM
___________________________________________________________________________

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This is a great email to bring back up the ethics of imaging!

What *color* was really collected? For a merged image the interpretation is great using color as described in the follow up emai. BUT, for single image display on a computer (PDF or Powerpoint) the information content is much more accessible to the viewer as grayscale than when displayed in color.

Why? RGB. When you have a grayscale image on the monitor it uses all three equally to make the image. When you have only the R channel on, the intensity is cut by 1/3. The issue there becomes detectable differences. Its a simple exercise. And I encourage everyone to try it: Take a grayscale image (any will do, assuming it has a close to normal distributed histogram) and then make it one color (just Red for example). For many of you this is a normal and obvious statement.

If you are overlaying color it makes sense to convert to color. If you are collecting an image with a color camera, and it is looking at the color (is it a real color or just the filtered spectrum defined by the bandpass filters), okay. But why reduce the information displayed in a single channel.

Maybe my personal bias against such practices as displaying the single chanel GFP label as green is unjust, but I ask:
If displaying as a color makes it more difficult to see the information in the micrograph, Why do it? (suggesting that it is less confusing for the viewer doesn't cut it - that's what figure legends and labels do very successfully).

-Geoff

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Fri 5/26/2006 4:32 AM
To: Williams, Geoffrey

Dear listers,

I have a triple labeling in fluorescence
(green-red-blue), but I take pictures in
epifluorescence with a B/W camera, which is more
sensitive than the colour camera. Now for presentation
purposes I would like to give colours to my B/W
pictures in the 3 colours originally used and
(perhaps) merge them in the end.
What is the procedure in Photoshop?

St�phane

__________________________________________________
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Dear Stephane,
the simplest manner in Photoshop is to open all 3 images then convert
1 to RGB and paste in the other channels. Some acquisition software
automatically exports monochrome images as 24-bit RGB, saving you
this step. Different steps may be needed for the various formats in
which captured images are saved.

In Photoshop, view the Channels window, If you have the "red" image
converted to RGB, select its "green" channel, then go to the "green"
image, 'Select All', 'Copy', return to the green channel in the
target image and 'Paste'. Repeat with the "Blue" channel and you are
done. Now, select the "RGB" channel at the top of the Channels
window to observe the merged image. Open the Levels command to
adjust histogram stretch and gamma for each channel in the merged
image (be certain to note gamma changes in methods or caption). If
you have 16-bit images, John and Christian Russ have available a
plugin to scale from 16-bit to 8-bit.

Regards
Glen

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

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

On May 26, 2006, at 1:30 AM, nizets2-at-yahoo.com wrote:

} Dear listers,
}
} I have a triple labeling in fluorescence
} (green-red-blue), but I take pictures in
} epifluorescence with a B/W camera, which is more
} sensitive than the colour camera. Now for presentation
} purposes I would like to give colours to my B/W
} pictures in the 3 colours originally used and
} (perhaps) merge them in the end.
} What is the procedure in Photoshop?
}
} St�phane

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Jafar F. Al-Sharab asked: "I would like to transfer 16 bit sun raster files
into TIF format without loosing the resolution!"

Jafar, what application generated your 16 bit Sun raster files?

Everything I have seen concerning '.ras' files specifies a header structure
that does NOT include a 16 bit per pixel option. For example, the excellent
site at the University of Rochester's EE site
(http://www.earth.rochester.edu/ees254/gmt/doc/html/GMT_Docs/node119.html)
lists the ras_depth member of the header structure as having values "Depth
(1, 8, 24, 32 bits) of pixel". Likewise, the SunRasterDecoder at the Code
Project site (http://www.codeproject.com/bitmap/SUN_Raster_File_Decoder.asp)
has also notes that ras-depth is "Depth (1, 8, 24 or 32 bits) of each
pixel".

If there is an updated specification or your application "cheats" in
handling the format (for instance using 16 bits of a 24 bit pixel, I suspect
most of the free converters will not work.

Jafar, if you send me directly (not to the list, please) a small (e.g.
512x512) 16 bit sun raster file, I'll try the tools I have and let you know
if any work...

Best regards,
John Minter
jrminter_at_rochester.rr.com
(replace the _at_ with -at-; I'm hoping to avoid the address harvesting bots
and minimize spam. My apologies for the inconvenience)

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There is a very good FREE Image converter program called
Imagemagick which I use on Unix, Linux, & Mac's. It
has also been ported to Windoze.

http://www.imagemagick.org/script/index.php

If you go to the above site you will be able to download
a version for SUN which should let you do the job.

Nestor
Your Friendly Neighborhood SysOp

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

--
===========================================
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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9, 16 -- To: microscopy-at-microscopy.com
9, 16 -- From: "Nestor J. Zaluzec" {zaluzec-at-aaem.amc.anl.gov}
9, 16 -- Subject: Converting sun raster into tif format
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On Fri, 26 May 2006 11:31:37 -0600, {zaluzec-at-aaem.amc.anl.gov} wrote:

} There is a very good FREE Image converter program called
} Imagemagick which I use on Unix, Linux, & Mac's. It
} has also been ported to Windoze.
}
} http://www.imagemagick.org/script/index.php
}
} If you go to the above site you will be able to download
} a version for SUN which should let you do the job.

In addition to the excellent ImageMagick toolset Mr. Zaluzec has just
recommended, I'd like to encourage you to investigate the Gimp
( http://www.gimp.org ). It is an open-source project intended to provide
functionality very similar to Photoshop. It has been ported to just about
everything: I use it daily on Mac OS X (under Apple's X11), and prior to
that I used it on both Solaris and Windows 98 and XP.

It is a downright invaluable tool, and would still be invaluable at twice
the price. Having said that, it's also free, and free is a property that
excuses many faults (even though there aren't many faults to object to
here!). Precompiled binaries are available at the Gimp web site as well.

It is well worth checking out, especially for those in academia with
limited budgets for image manipulation software. Between ImageMagick and
the Gimp, you can do an astonishing amount of work.

Hope that helps!

-skod

--
Scott Griffith
ISES-LLC
9745 Steeplechase Drive
Franktown, CO 80116
303-660-2541 voice
303-660-2542 fax
skod-at-ises-llc.com

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A postdoctoral position is available in Surface Transmission Electron
Microscopy at Northwestern University. Work would involve collaborative
work on a number of different projects including charge density at
surfaces, oxide surface structures, surface plasmonics and the surface
structure of fuel cell materials. A strong background in TEM is necessary,
including basic knowledge of dynamical diffraction and imaging theory.
Experience in plan or profile imaging of surfaces would be a plus, but
not a requirement.

To apply, send a short CV with a list of publications (no papers) as well
as the names of three referees to L-marks-at-northwestern.edu. Include a
brief (one page maximum) description of your strengths as relevant to
transmission electron microscopy of surfaces.

-----------------------------------------------
Professor Laurence Marks
Department of Materials Science and Engineering
MSE Rm 2036 Cook Hall
2220 N Campus Drive
Northwestern University
Evanston, IL 60201, USA
Tel: (847) 491-3996 Fax: (847) 491-7820
email: L - marks -at- northwestern . edu
http://www.numis.northwestern.edu
-----------------------------------------------

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5, 17 -- From: LDM Microscopy {ldmm-at-risc4.numis.northwestern.edu}
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5, 17 -- Subject: Postdoctoral Opening in Surface Microscopy
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I look for etchants for Ni3Al (grain size) ?

Best regards

Krzysztof Jan H�bner

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Thanks Geoffrey for this remark. In the present case
the merging of the 3 colors is meaningful because I
look for colocalisation (I will add "or not", to
remain ethically objective ;-)), but your remark makes
sense.
Please could you answer to the following
considerations?

- What if you make color photo films of TIFF images to
project them (classical slide projectors)?
- What if you project a TIFF image with a projector
directly coupled to the computer instead of a monitor?
- Would it be ethically correct to increase the
intensity of one color 3 fold in order to compensate
the decrease due to RGB pictures displayed on a
monitor?

St�phane

--- Geoffrey_Williams-at-brown.edu wrote:

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}
} This is a great email to bring back up the ethics of
} imaging!
}
} What *color* was really collected? For a merged
} image the interpretation is great using color as
} described in the follow up emai. BUT, for single
} image display on a computer (PDF or Powerpoint) the
} information content is much more accessible to the
} viewer as grayscale than when displayed in color.
}
} Why? RGB. When you have a grayscale image on the
} monitor it uses all three equally to make the image.
} When you have only the R channel on, the intensity
} is cut by 1/3. The issue there becomes detectable
} differences. Its a simple exercise. And I
} encourage everyone to try it: Take a grayscale image
} (any will do, assuming it has a close to normal
} distributed histogram) and then make it one color
} (just Red for example). For many of you this is a
} normal and obvious statement.
}
} If you are overlaying color it makes sense to
} convert to color. If you are collecting an image
} with a color camera, and it is looking at the color
} (is it a real color or just the filtered spectrum
} defined by the bandpass filters), okay. But why
} reduce the information displayed in a single
} channel.
}
} Maybe my personal bias against such practices as
} displaying the single chanel GFP label as green is
} unjust, but I ask:
} If displaying as a color makes it more difficult to
} see the information in the micrograph, Why do it?
} (suggesting that it is less confusing for the viewer
} doesn't cut it - that's what figure legends and
} labels do very successfully).
}
} -Geoff
}
}
} -----Original Message-----
} X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Fri 5/26/2006 4:32 AM
} To: Williams, Geoffrey
} Subject: [Microscopy] coloring B/W pictures with
} photoshop
}
}
}
}
}
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} Dear listers,
}
} I have a triple labeling in fluorescence
} (green-red-blue), but I take pictures in
} epifluorescence with a B/W camera, which is more
} sensitive than the colour camera. Now for
} presentation
} purposes I would like to give colours to my B/W
} pictures in the 3 colours originally used and
} (perhaps) merge them in the end.
} What is the procedure in Photoshop?
}
} St�phane
}
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} 4, 18 -- Subject: coloring B/W pictures with
} photoshop
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} 18, 30 -- From Geoffrey_Williams-at-brown.edu Fri May
} 26 11:21:21 2006
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} pictures with photoshop
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9, 20 -- From: Stephane Nizet {nizets2-at-yahoo.com}
9, 20 -- Subject: Re: [Microscopy] RE: coloring B/W pictures with photoshop
9, 20 -- To: Geoffrey_Williams-at-brown.edu
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Thanks to all for the easy methods to create colourful
B/W pictures! It is so simple, when you know what to
do!

St�phane

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3, 19 -- Subject: THANKS ! coloring B/W pictures with photoshop
3, 19 -- To: microscopy-at-microscopy.com
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Email: DrJohnRuss-at-AOL.com
Name: John Russ

Organization: North Carolina State University

Title-Subject: [Filtered] RE: colouring B/W pictures with Photoshop

Question:

-----Original Message-----
} From: keith.morris-at-ucl.ac.uk
To: DrJohnRuss-at-AOL.com
Sent: Fri, 26 May 2006 03:59:50 -0500

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Email: Amr_elsirafy-at-hotmail.com
Name: afaf Ibrahim Mahdi

Organization: Cairo university

Title-Subject: [Filtered] ESEM-detectors

Question: Hello
I' d like to know is GSED is used in ESEM with tungesten filament and if it can be used in both low and high vacuum mode because the BSD in my XL30 TMP instrument is damaged and I want to know iif it is best to buy back scatter detectors or GSED and is it need a new software.

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6, 12 -- From zaluzec-at-microscopy.com Tue May 30 08:08:18 2006
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Email: eric.leroy-at-glvt-cnrs.fr
Name: Eric Leroy

Organization: CNRS - LCMTR

Title-Subject: [Filtered] AnalySIS TIFF images

Question: Hi,

We have a SIS Megaview III camera attached to our Tecnai and we use AnalySIS to record the images. Since we only have a full version of AnalySIS installed on the microscope's PC we usually treat ours images with ImageJ. ImageJ can perfectly read the TIFF images but is unable to read the spatial calibration of the image. Do you know how this spatial calibration is written in the image ?

Thanks

Eric

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I Googled GSED and couldn't find anything related to SEM, so for those
of us who are acronym-challenged, could you (or someone else on the
list) please define GSED?

Thanks,
Bryan Bandli
MVA Scientific Consultants

Amr_elsirafy-at-hotmail.com wrote:

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==============================Original Headers==============================
6, 19 -- From bbandli-at-mvainc.com Tue May 30 08:52:33 2006
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GSED = Gaseous Secondary Electron Detector. It is the secondary
electron detector of the ESEM which works at poor vacuum unlike a
conventional SE detector.

Dave

-----Original Message-----
X-from: bbandli-at-mvainc.com [mailto:bbandli-at-mvainc.com]
Sent: 30 May 2006 14:54
To: David Patton

I Googled GSED and couldn't find anything related to SEM, so for those
of us who are acronym-challenged, could you (or someone else on the
list) please define GSED?

Thanks,
Bryan Bandli
MVA Scientific Consultants

Amr_elsirafy-at-hotmail.com wrote:

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You cannot use GSED in high vac; it is only for low vac. I have a XL30 ESEM and never use the backscatter detector (BSD) since installation six years ago for morphological study, except using it to compare with GSED at the very beginning. A GSED produces images a lot better than a BSD. If I were you, the pick would have been a GSED. If you need a BSD for another reason, then that is another matter to consider.
As for software, you don't have to worry; just put in a new GSED and off you go. I have checked with my service engineer.

Ann Fook Yang
EM Unit/ Unite EM
Bldg 20, AAFC/AAC
960 Carling Ave,
Ottawa,Ontario
Canada K1A 0C6
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701
yanga-at-agr.gc.ca

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada

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Email: Amr_elsirafy-at-hotmail.com
Name: afaf Ibrahim Mahdi

Organization: Cairo university

Title-Subject: [Filtered] ESEM-detectors

Question: Hello
I' d like to know is GSED is used in ESEM with tungesten filament and if it can be used in both low and high vacuum mode because the BSD in my XL30 TMP instrument is damaged and I want to know iif it is best to buy back scatter detectors or GSED and is it need a new software.

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Eric,

The TIFF standard allows for spatial resolution data to be saved in the
fields XResolution (no of pixels per measurement unit in X dir) and
YResolution (no of pixels per measurement unit in Y dir). The measurement
unit data is contained in the ResolutionUnit field (usually inch or cm).
From these you should be able to calculate the pixel size, magn or image
width. There are a number of free TIFF tag viewers available, for example,
http://www.awaresystems.be/imaging/tiff/astifftagviewer.html, or just
Google 'tiff tag viewer' .
Hope this helps.

David Vowles
Electron Microscope Unit
Dept of Materials Science and Metallurgy
University of Cambridge
Pembroke St Cambridge
UK CB2 3QZ
Tel: +44 (0)1223 334325
Fax: +44 (0)1223 334567
Email: djv23-at-cam.ac.uk

} Name: Eric Leroy
}
} Organization: CNRS - LCMTR
}
} Title-Subject: [Filtered] AnalySIS TIFF images
}
} Question: Hi,
}
} We have a SIS Megaview III camera attached to our Tecnai and we use
} AnalySIS to record the images. Since we only have a full version of
} AnalySIS installed on the microscope's PC we usually treat ours images
} with ImageJ. ImageJ can perfectly read the TIFF images but is unable to
} read the spatial calibration of the image. Do you know how this spatial
} calibration is written in the image ?
}
} Thanks
}
} Eric
}
} ---------------------------------------------------------------------------
}
} ==============================Original Headers==============================
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Salut Eric,

We have the same system. The easiest way to keep the
magnification in my pictures I found is to "burn" the
scale bar into the image (on your screen it is just in
overlay) before saving the picture in jpg or TIFF
format. It is a little bit a pain, I would appreciate
a method to automatize it, but I don't know if it is
possible (I work only for 8 months with this system).
It is clear that the pixels "behind" the scale bar
will be burned and the information lost.

- Choose "Image" in the Menu bar
--} Scale bar...
--} draw into overlay
Then save your image

Sorry if I misunderstood your question and did not
answer it (ask it in french it will be clearer ;-)).

St�phane

--- eric.leroy-at-glvt-cnrs.fr wrote:

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} Email: eric.leroy-at-glvt-cnrs.fr
} Name: Eric Leroy
}
} Organization: CNRS - LCMTR
}
} Title-Subject: [Filtered] AnalySIS TIFF images
}
} Question: Hi,
}
} We have a SIS Megaview III camera attached to our
} Tecnai and we use AnalySIS to record the images.
} Since we only have a full version of AnalySIS
} installed on the microscope's PC we usually treat
} ours images with ImageJ. ImageJ can perfectly read
} the TIFF images but is unable to read the spatial
} calibration of the image. Do you know how this
} spatial calibration is written in the image ?
}
} Thanks
}
} Eric
}
}
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}
} ==============================Original
} Headers==============================
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} 08:08:56 2006
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11, 20 -- From nizets2-at-yahoo.com Wed May 31 01:51:14 2006
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nizets2-at-yahoo.com wrote:
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}
} Salut Eric,
}
} We have the same system. The easiest way to keep the
} magnification in my pictures I found is to "burn" the
} scale bar into the image (on your screen it is just in
} overlay) before saving the picture in jpg or TIFF
} format. It is a little bit a pain, I would appreciate
} a method to automatize it, but I don't know if it is
} possible (I work only for 8 months with this system).
}
If you save to TIFF file format there is an option you can select in the
"Save..." dialog to burn the overlay into the image data. (The other
option is to convert to 8bit TIFF file, which sometimes may be helpful,
too.)

-at-Eric: There is a viewer that can be downloaded from the SIS webpage[1],
which can handle the image files (and databases) created with the full
version. This viewer also shows you the data listed in the TIFF header.

Hope that helps...
Hanno

[1] http://www.soft-imaging.net/rd/english/419.htm

==============================Original Headers==============================
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Dear all,

Is it possible to use a immunogold purpose silver enhancement kit for
revealing other heavy metals, such as cadmium, on paraffin or resin sections?

Best wishes

Fabio

Dr. Fabio D'Amico
Dept. Biomedical Sciences
University of Catania
Via Androne 87/a
I-95124 Catania
ITALY
tel/fax +39 095312017
email: f.damico-at-unict.it

==============================Original Headers==============================
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David,

Thank you for the information. I downloaded the software you told me but
unfortunately, the information written int Xresolution tag is allways 200
and the ResolutionUnit is 0. So it seems that the saptial calibration is
stored elsewhere.

Best regards

Eric

Le 30/05/06 17:58, ��David Vowles�� {djv23-at-cam.ac.uk} a �crit�:

} Eric,
}
} The TIFF standard allows for spatial resolution data to be saved in the
} fields XResolution (no of pixels per measurement unit in X dir) and
} YResolution (no of pixels per measurement unit in Y dir). The measurement
} unit data is contained in the ResolutionUnit field (usually inch or cm).
} From these you should be able to calculate the pixel size, magn or image
} width. There are a number of free TIFF tag viewers available, for example,
} http://www.awaresystems.be/imaging/tiff/astifftagviewer.html, or just
} Google 'tiff tag viewer' .
} Hope this helps.
}
} David Vowles
} Electron Microscope Unit
} Dept of Materials Science and Metallurgy
} University of Cambridge
} Pembroke St Cambridge
} UK CB2 3QZ
} Tel: +44 (0)1223 334325
} Fax: +44 (0)1223 334567
} Email: djv23-at-cam.ac.uk
}
}
}
} } Name: Eric Leroy
} }
} } Organization: CNRS - LCMTR
} }
} } Title-Subject: [Filtered] AnalySIS TIFF images
} }
} } Question: Hi,
} }
} } We have a SIS Megaview III camera attached to our Tecnai and we use
} } AnalySIS to record the images. Since we only have a full version of
} } AnalySIS installed on the microscope's PC we usually treat ours images
} } with ImageJ. ImageJ can perfectly read the TIFF images but is unable to
} } read the spatial calibration of the image. Do you know how this spatial
} } calibration is written in the image ?
} }
} } Thanks
} }
} } Eric
} }
} } ---------------------------------------------------------------------------
} }
} } ==============================Original Headers==============================
} } 9, 12 -- From zaluzec-at-microscopy.com Tue May 30 08:08:56 2006
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}

\\_//
-(-at- -at-)-
----------------------oOO--(_)--Ooo-------------------------

Eric LEROY Dr.
Laboratoire de Chimie Metallurgique des Terres Rares
UPR 209 - CNRS
Groupe des Laboratoires de Thiais
2-8, rue Henri Dunant
94320 THIAIS cedex

Tel : (33) (0)1 49 78 12 09 ou 13 24
Fax : (33) (0)1 49 78 12 03
email : eric.leroy-at-glvt-cnrs.fr
http://www.glvt-cnrs.fr/microscope
------------------------------Oooo.-------------------------
.oooO ( )
( ) ) /
\ ( (_/
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Hello,

The problem of Eric brings a new question to me:
The TEM pictures may be saved in JPG or TIFF formats.
I know that TIFF format keeps all the information
without compression. But does JPG without compression
(quality 12) deteriorates the quality of the picture?
I thought that JPG without compression was as good as
TIFF.

Stephane

__________________________________________________
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Title-Subject: [Filtered] perforated paper labels for embedding molds

Question: Where can I buy some? A full page of 5 1/4" x 7 3/4" was perforated into a total of 7x31 labels, each label is 3/4" x 1/8" and fits one BEEM or gelatin capsules. The full pages are bound like a notepad.

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Stephanie:

Please be advised that there is no implementation of JPG with lossless
compression anywhere in the world that I know of. The last time I saw a
program that actually was lossless was in DOS 3.3 almost twenty years ago.

Saving with JPG at a setting of 12 removes data. period

If you load an image and make adjustments and then resave it, the image
file will have compression on top of compression. The situation gets
worse and worse.
JPG is an evil format that is not supported for scientific imaging by
the Society for really good reasons. If a system produces a different
format, the first time you save it should ALWAYS be to tiff if you want
to save it.

JPG is for email and web pages and is good for nothing else

Luckily I don't have strong opinions

John

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At 02:25 AM 5/31/2006, h.dierke-at-tu-braunschweig.de wrote:

} If you save to TIFF file format there is an option you can select in the
} "Save..." dialog to burn the overlay into the image data. (The other
} option is to convert to 8bit TIFF file, which sometimes may be helpful,
} too.)

On our AnalySIS-based systems, "burning in" the overlay also automatically
converts the file to 8 bit. The original quantitative image data is lost
in the conversion. This gave us fits a few years ago until we found out
what was gong on. If I may personify a little, it seems like AnalySIS
desperately wants to convert images to 8 bit - the only sure strategy we
have found for preserving the 16 bit images is to save them immediately
after acquisition, then save again under a different name after any
manipulation in AnalySIS.

I should also mention that we are several versions out of date on the
software; newer versions of AnalySIS / iTEM / whatever may have changed
this behavior.

Best wishes,
Paul Voyles

Paul Voyles
Assistant Professor
Materials Science and Engineering Department
University of Wisconsin - Madison
1509 University Ave.
Madison, WI 53706-1595
Voice: (608) 265-6740
Fax: (608) 262-8353
voyles-at-engr.wisc.edu
www.engr.wisc.edu/mse/faculty/voyles_paul.html

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Good one John! Now I have an image of Buffy the JPEG Slayer working to
save us all from the 'evil JPG.' John.

john_mackenzie-at-ncsu.edu wrote:

}
} Stephanie:
}
} Please be advised that there is no implementation of JPG with lossless
} compression anywhere in the world that I know of. The last time I saw a
} program that actually was lossless was in DOS 3.3 almost twenty years ago.
}
} Saving with JPG at a setting of 12 removes data. period
}
} If you load an image and make adjustments and then resave it, the image
} file will have compression on top of compression. The situation gets
} worse and worse.
} JPG is an evil format that is not supported for scientific imaging by
} the Society for really good reasons. If a system produces a different
} format, the first time you save it should ALWAYS be to tiff if you want
} to save it.
}
} JPG is for email and web pages and is good for nothing else
}
} Luckily I don't have strong opinions
}
} John
}
}
}
}

--

*********************************
C. John Runions, Ph.D.
School of Biological and Molecular Sciences
Oxford Brookes University
Oxford, UK
OX3 0BP

email: jrunions-at-brookes.ac.uk
phone: +44 (0) 1865 483 964

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Just a reminder that TIFF is the officially recognized format by the Microscopy Society. Basic rule of thumb: save the original in TIFF then save a copy in JPEG for easier email communication etc.

The big problem with JPEG used to be degradation of the image on resizing, etc. I don't know if this is still the case with their new algorithms.

Hope this was 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 07:34 AM 5/31/2006, nizets2-at-yahoo.com wrote:

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Good evening,
dear Fabio,
in addition to the helpful message of Rick Powell on Silver Amplifying /
Autometallographic localization of metals / Gorm Danscher papers etc. I
would like to comment on your problem (since silver precipitation alone is
more/less unspecific and will label a lot of metals in your specimen)....

Perhaps my message does not help you really, but if your } silver enhance
kit { alone is not able to localize Cd qualitatively / quantitatively (and
your Cd-concentration in the probe likely is very small) you could use also
perhaps a precipitating method

}
By means of crystalline precipitation as Cd (TH)2 [Cr (NH3)2 (CNS)4]2,1
0,1% of cadmium can be detected, without previous separation, in the
presence of the metals of the ammonium sulphide group and the alkaline
earths. Moreover, about 0,2% of cadmium can be detected by aid of a simple
separation, in the presence of whatsoever metals of the hydrochloric acid
and hydrogen sulphide groups. {
source:
Microchimica Acta Publisher: Springer Wien ISSN: 0026-3672 (Paper)
1436-5073 (Online) DOI: 10.1007/BF01471844 Issue: Volume 3, Number 4

Assuming that precipitation of Cd as CdS (with ammoniumsulfide, and
-perhaps -using also KCN for masking copper) is not the method of your
choice, I have had a look into PubMed and found the following reference:
Biomarkers. 2002 Nov-Dec;7(6):491-500.
Autometallography and metallothionein immunohistochemistry in hepatocytes
of turbot (Scophthalmus maximus L.) after exposure to cadmium and
depuration treatment.
Amaral AF, Alvarado N, Marigomez I, Cunha R, Hylland K, Soto M.
Section of Ecology, Department of Biology, University of the Azores, R Mae
de Deus, 9500 Ponta Delgada, Sao Miguel, Azores, Portugal.
Abstract: In this study, autometallography and immunohistochemistry were
used to localize and quantify cadmium and metallothionein (MT) levels,
respectively, in cellular compartments of turbot liver on exposure to
cadmium for 7 days and further depuration treatment for 14 days. Metals
weakly bound to proteins (i.e. MTs) in hepatocyte lysosomes were visualized
as black silver deposits (BSDs) using a light microscope. With the aid of a
newly developed immunohistochemical procedure, MTs were localized and
semi-quantified in both the cytosolic and the lysosomal compartments of
hepatocytes. The BSD extent in the lysosomes of hepatocytes increased
significantly as a result of cadmium exposure. This response was evidenced
after 1 h. Further, a progressive increase in the volume density of BSDs
occurred up to the seventh day. Total MT immunohistochemical levels
increased at a lower rate, starting after 1 day of cadmium exposure. BSD
extent values recovered after depuration, whilst MT levels remain
unchanged. It is possible that the detoxification rate of metals via
lysosomes was diminished, whilst MT levels remained unchanged, at least
after 14 days of depuration. It can be concluded that autometallography and
MT immunohistochemistry are good tools for clarifying metal and metal-MT
trafficking routes in hepatocytes, and also that BSD extent and MT
immunohistochemical levels in the lysosomes and cytosol of fish hepatocytes
can be considered to be useful biomarkers of metal exposure.

Unfortunately this is an article published in a Taylor and Francis Journal,
so I was not able to retrieve the article as pdf.......but you can find the
Abstract and Journal reference at:
http://taylorandfrancis.metapress.com/link.asp?id=efeg34kqvvyn0vtg

The search in Pubmed for "cadmium localiz" resulted only with 17 hits.

Also I have found (for: "cadmium" and "EM" only 14 results) at:
http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B6TCP-4HHGNPH-1
&_coverDate=06%2F30%2F2006&_alid=408694459&_rdoc=1&_fmt=&_orig=search&_q
d=1&_cdi=5176&_sort=d&view=c&_acct=C000057295&_version=1&_urlVersion=0&_
userid=2450241&md5=5a4679d3894972bb9ba28a4ab1f88de4

an article dealing with Cd-localization perhaps), also there only the
abstract is available.

When using: } "cadmium" and "TEM" { as search phrase, I found a paper,
entitled
} Bioaccumulation and localization of exogenous cadmium in a teleost by
electron microscopy (TEM) and its specific quantitation by electron probe
X-ray microanalysis (EPMA) {.
Bioorg Med Chem. 2000 Mar;8(3):475-82.
by Tayal AK, Kaur I, Mathur RP.

the abstract of which tells us:
} A cadmium bioconcentration study was carried out in a fresh water teleost,
Colisa fasciatus, to study the bioaccumulation kinetics and fate of
exogenous cadmium (Cd) in biological tissues. Study shows that on exposure
of the fish to a sublethal concentration of cadmium in test water, Cd
uptake results in its bioconcentration in gills, liver and muscle tissues.
To explore whether the accumulated Cd reaches the membranes or inside the
cells, transmission electron microscopy (TEM) of the thin sections of
tissues was done after
histochemical localization of Cd in cells by modified SST method.

TEM studies of sections of gills, liver and muscle tissues showed the
deposits of exogenous Cd (visualized as dense clouds) in biological cells.
This suggests the presence of free or loosely bound Cd on the membranes and
inside the cells, which in the presence of Na2S is converted into insoluble
metal sulfides. Electron probe X-ray microanalysis (EPMA) studies confirmed
the presence of Cd on the membrane surface as well as inside the cells of
bioindicator organs suggesting involvement of membrane transport of
exogenous Cd inside the cells and its deposition as loosely bound insoluble
metal complexes.

SST-Method: excerpt out of the pdf (which I am able to send to you, if you
allow):
} } }
The SST process employs a developer with a low pH, silver nitrate as silver
ion donator and a protecting colloid, gum arabic. The developer involves
the use of small amounts of hydroquinone as reduction molecules.
The SST results in reduced silver ion magnfications at specfic sites with
gum arabic reducing the autocatalytic activity in the developer itself and
the catalytic activity
of the zone between the developer and the surface of the section [6,8].
However, SST visualizes only a certain fraction of heavy metals represented
by free or loosely bound metal that in the presence of Na2S is converted
into insoluble metal sufdes. The metal that is strongly bound to
organic/inorganic ligands may not be sulfidated and remains histochemically
invisible.
The visualization of the fraction of Cd in fish tissue presented herein
support Danscher and Norgaard [18] in that SST could be used for the
demonstration of trace amounts of certain heavy metals at ultra-structural
level and that the metal sufides can be shown by the technique. In their
investigation, George et al.,[9] using the precipitation reactions (sufide
for heavy metals), reported precipitation of Cd as a sufide in bivalve
tissues.
They also reported that precipitation reactions result in a better
localization with a loss of 14% as compared to .....

Also, with this search phrase, you will discover some older references,
like
} Cellular alterations in collembolan midgut cells as a marker of heavy
metal exposure: ultrastructure and intracellular metal distribution. {
Sci Total Environ. 1996 Mar 29;181(3):187-200.
by Pawert M, Triebskorn R, Graff S, Berkus M, Schulz J, Kohler HR.
(which you can find at:
http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B6V78-3VWF92S-1
P&_coverDate=03%2F29%2F1996&_alid=408705808&_rdoc=1&_fmt=&_orig=search&_
qd=1&_cdi=5836&_sort=d&view=c&_acct=C000057295&_version=1&_urlVersion=0&
_userid=2450241&md5=77742f5cebfe3a73991ef90c3973adff )

the .pdf of which I also could send to you (please send } allowance { mail)

Hope this helps

best regards and wishes,

Wolfgang MUSS
Salzburg, Austria

----------
Von: fab-at-tariffenet.it[SMTP:fab-at-tariffenet.it]
Antwort an: fab-at-tariffenet.it
Gesendet: Mittwoch, 31. Mai 2006 11:21
An: W.Muss-at-salk.at
Betreff: [Microscopy] Silver enhancement kit

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Is it possible to use a immunogold purpose silver enhancement kit for
revealing other heavy metals, such as cadmium, on paraffin or resin
sections?

Best wishes

Fabio

Dr. Fabio D'Amico
Dept. Biomedical Sciences
University of Catania
Via Androne 87/a
I-95124 Catania
ITALY
tel/fax +39 095312017
email: f.damico-at-unict.it

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39, 28 -- From W.Muss-at-salk.at Wed May 31 11:28:09 2006
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39, 28 -- Cc: "'Microscopy-at-Microscopy.Com'" {Microscopy-at-Microscopy.Com}
39, 28 -- Subject: [Microscopy]Re: (LONG) Silver enhancement kit
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A user wants to remove the data bar from a stored image. Can anyone
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Dave

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Listers,

My recent response to ESEM detectors was subjective and incomplete. My errors have brought Daniel Phifer, the application specialist of FEI to my rescue. Please read below. Daniel, thank you.

Ann Fook Yang
EM Unit/ Unite EM
Bldg 20, AAFC/AAC
960 Carling Ave,
Ottawa, Ontario
Canada K1A 0C6
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701
yanga-at-agr.gc.ca

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada
============================================================================

AnnFook,
I was forwarded your list server response and wanted to give you some info to better respond.

For your samples which have little to no atomic contrast, there is likely no need for the BSE except when looking for gold labels, etc. This is usually on dried prepared biological samples. With the majority of your samples at Agri-Food Canada, the BSE and SE images may be similar as the atomic composition is very similar.

The query could be from a customer who has lots of atomic differences and thus may need a BSE detector to show the atomic differences. Maybe not, but this should be qualified to give the best response.

FEI has two detectors for picking up BSE signal. The solid state BSE crystal can be used in high and low vacuum but not in ESEM (as it requires the gas to amplify the signal. As the pressure is increased above 2-3 Torr in the chamber, the BSE crystal will get less of the signal reaching the detector as the BSE collide with the gas and scatter. For this situation, there is an optional Gaseous Backscattered Detector (GBSD) which can be used in ESEM mode (4-7Torr optimum). The GBSD will give atomic contrast (even in fully hydrated samples) using a patented detection mechanism. If BSE is desired, these two detectors are the best options. The GSED (Gaseous Secondary Electron Detector) only picks up secondary electrons which yield surface information, not composition.

It is my impression from the question that the terminology is getting confused in translation. Hopefully this information will allow you to clarify your response. Please feel free to paraphrase or plagiarize my words and make them your own if you like. I just want you to have the information. If you have questions about the technology and would like me to help you answer a query, please do not hesitate to contact me directly.

Hope all is well in Ottawa.

Daniel

Daniel Phifer

FEI Company

-----Original Message-----
X-from: Yang, Ann-Fook
Sent: Tuesday, May 30, 2006 11:38 AM
To: Yang, Ann-Fook

You cannot use GSED in high vac; it is only for low vac. I have a XL30 ESEM and never use the backscatter detector (BSD) since installation six years ago for morphological study, except using it to compare with GSED at the very beginning. A GSED produces images a lot better than a BSD. If I were you, the pick would have been a GSED. If you need a BSD for another reason, then that is another matter to consider.
As for software, you don't have to worry; just put in a new GSED and off you go. I have checked with my service engineer.

Ann Fook Yang
EM Unit/ Unite EM
Bldg 20, AAFC/AAC
960 Carling Ave,
Ottawa,Ontario
Canada K1A 0C6
Telephone/T�l�phone: 613-759-1638
Facsimile/T�l�copieur: 613-759-1701
yanga-at-agr.gc.ca

Agriculture and Agri-Food Canada - Agriculture et Agroalimentaire Canada

-----Original Message-----
X-from: Amr_elsirafy-at-hotmail.com [mailto:Amr_elsirafy-at-hotmail.com]
Sent: Tuesday, May 30, 2006 9:21 AM
To: Yang, Ann-Fook

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Email: Amr_elsirafy-at-hotmail.com
Name: afaf Ibrahim Mahdi

Organization: Cairo university

Title-Subject: [Filtered] ESEM-detectors

Question: Hello
I' d like to know is GSED is used in ESEM with tungesten filament and if it can be used in both low and high vacuum mode because the BSD in my XL30 TMP instrument is damaged and I want to know iif it is best to buy back scatter detectors or GSED and is it need a new software.

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

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Another informative listserver for FEI/Philips XL30 ESEM's is maintained by Dr. Cameron Begg at Ohio State University. See instructions below. I hope this paste is still valid.

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} } } {David.Patton-at-uwe.ac.uk} 05/31/06 11:43AM } } }

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A user wants to remove the data bar from a stored image. Can anyone
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Barbara writes ...

} Just a reminder that TIFF is the officially recognized format
} by the Microscopy Society.

More exactly ... It is the ^uncompressed^ TIFF that is the MSA standard
format ... Presumably because the bitmap is intact, while compression
algorithms come and go.

cheerios :o)
michael shaffer

SEM/MLA Research Coordinator
{http://www.mun.ca/creait/maf/}
{http://www.esd.mun.ca/epma/}
Inco Innovation Centre
c/o Memorial University
St. John's, NL A1C 5S7

}
} At 07:34 AM 5/31/2006, nizets2-at-yahoo.com wrote:
}
}
}
} } -------------------------------------------------------------
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} } Hello,
} }
} } The problem of Eric brings a new question to me:
} } The TEM pictures may be saved in JPG or TIFF formats.
} } I know that TIFF format keeps all the information without
} compression.
} } But does JPG without compression (quality 12) deteriorates
} the quality
} } of the picture?
} } I thought that JPG without compression was as good as TIFF.
} }
} } Stephane
} }
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Sorry, I was incommunicado for a few days and just saw this posting.

The resolution information is not stored in the tags mentioned below for the following reason: Many word processing and page layout programs use that tag to print something at the "right" size (for example MS Word). If you enter the calibration value in this field, the software will try to print the image with a width of a few microns, resulting in a single dot on the paper. We ran into this behavior a few years ago and were forced to store the calibration values in a private tag. If you want to know what the tag is, let me know, I can find out for you.

ananlySIS does not automatically convert images to 8 bit, unless instructed to do so. One option is, as already mentioned, the possibility to automatically convert to 8-bit when saving the image. Of course, if that is selected, the software will do that.

Michael Bode

General Manager

OLYMPUS SOFT IMAGING SOLUTIONS CORP.
12596 West Bayaud Ave #300
Lakewood, CO 80228
USA
Tel.: +1 (303) 234-9270
Fax.: +1 (303) 234-9271
E-Mail: Mike.Bode-at-olympus-sis.net
www.olympus-sis.net
-----Original Message-----
X-from: eric.leroy-at-glvt-cnrs.fr [mailto:eric.leroy-at-glvt-cnrs.fr]
Sent: Wednesday, May 31, 2006 3:45
To: Mike Bode

David,

Thank you for the information. I downloaded the software you told me but
unfortunately, the information written int Xresolution tag is allways 200
and the ResolutionUnit is 0. So it seems that the saptial calibration is
stored elsewhere.

Best regards

Eric

Le 30/05/06 17:58, ��David Vowles�� {djv23-at-cam.ac.uk} a �crit�:

} Eric,
}
} The TIFF standard allows for spatial resolution data to be saved in the
} fields XResolution (no of pixels per measurement unit in X dir) and
} YResolution (no of pixels per measurement unit in Y dir). The measurement
} unit data is contained in the ResolutionUnit field (usually inch or cm).
} From these you should be able to calculate the pixel size, magn or image
} width. There are a number of free TIFF tag viewers available, for example,
} http://www.awaresystems.be/imaging/tiff/astifftagviewer.html, or just
} Google 'tiff tag viewer' .
} Hope this helps.
}
} David Vowles
} Electron Microscope Unit
} Dept of Materials Science and Metallurgy
} University of Cambridge
} Pembroke St Cambridge
} UK CB2 3QZ
} Tel: +44 (0)1223 334325
} Fax: +44 (0)1223 334567
} Email: djv23-at-cam.ac.uk
}
}
}
} } Name: Eric Leroy
} }
} } Organization: CNRS - LCMTR
} }
} } Title-Subject: [Filtered] AnalySIS TIFF images
} }
} } Question: Hi,
} }
} } We have a SIS Megaview III camera attached to our Tecnai and we use
} } AnalySIS to record the images. Since we only have a full version of
} } AnalySIS installed on the microscope's PC we usually treat ours images
} } with ImageJ. ImageJ can perfectly read the TIFF images but is unable to
} } read the spatial calibration of the image. Do you know how this spatial
} } calibration is written in the image ?
} }
} } Thanks
} }
} } Eric
} }
} } ---------------------------------------------------------------------------
} }
} } ==============================Original Headers==============================
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}

\\_//
-(-at- -at-)-
----------------------oOO--(_)--Ooo-------------------------

Eric LEROY Dr.
Laboratoire de Chimie Metallurgique des Terres Rares
UPR 209 - CNRS
Groupe des Laboratoires de Thiais
2-8, rue Henri Dunant
94320 THIAIS cedex

Tel : (33) (0)1 49 78 12 09 ou 13 24
Fax : (33) (0)1 49 78 12 03
email : eric.leroy-at-glvt-cnrs.fr
http://www.glvt-cnrs.fr/microscope
------------------------------Oooo.-------------------------
.oooO ( )
( ) ) /
\ ( (_/
\_)

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Email: afaf_mahdi-at-hotmail.com
Name: Afaf Ibrahim Mahdi

Organization: Cairo Univ

Title-Subject: [Filtered] ESEM detectors

Question: Hello all
I have already recieved answers from some scientists for my question about GSED and it was valuable. I'am a mieralogist and I used ESEM XL30 TMP in examining geological samples and also biological samples. I used BSD fo about 90% of my work. For about 6 years I used BSD in both high vacuum and low vacuum modes by converting Key no 4 and I used only carbon sticker tofix samples. it gives satisfied EDX analyses but images some time satisfactory and sometimes it is very bad. The magnification dont exceed 8000X. Now the back scatter detector is damage and I want to buy one suitable for both modes because I don' have enough moner to buy 2 detectors. Is there one detector can used in both modes taking into account that i still have GSD or it is better to buy BSD. In the last monty I noticed that the two pumps are begging to heat severly, could be the detector is the reason.
thank you

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(5/31/06 10:47) bfoster-at-mme1.com {bfoster-at-mme1.com} wrote:

} Just a reminder that TIFF is the officially recognized format by the Microscopy Society. Basic rule of thumb: save the original in
} TIFF then save a copy in JPEG for easier email communication etc.
}
} The big problem with JPEG used to be degradation of the image on resizing, etc. I don't know if this is still the case with their new
} algorithms.

JPEG2000 is no more lossless than the original JPEG - only the method of compression has changed.

Also - I warn people to beware of TIFFs from unknown programs. The TIFF 6 specification does allow for storing JPEG compressed data in the TIFF wrapper - there is a flag that indicates the compression level.

Brent
--
Brent Neal, Ph.D.
Asheville, NC
{brentn-at-gmail.com}

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