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1. Standard
Procedure to Prepare Specimens for Transmission Electron Microscopy,
EM Center, Indiana University School of Medicine:
Fix the specimen with an appropriate aldehyde fixative for a minimum
of one hour, depending on the size of the specimen. Ideal size for
the specimen should be a cube less than 3mm square. Cut the specimen
and place in the fixative as quickly as possible. The fixative routinely
used in this lab is a modified Karnovsky's, 2% Paraformaldehye/2%
Glutaraldehyde in 0.1M Phosphate Buffer. After fixation the specimens
are rinsed several times with phosphate buffered saline (PBS) followed
by post fixation with 1% osmium tetroxide in phosphate buffer for
one hour. After rinsing again with PBS for 15 minutes, the tissue
specimens are dehydrated through a series of graded ethyl alcohols
from 70 to 100%. The schedule is as follows: 70% for 10 min., 95%
for 10 min. and three changes of 100% for 5 minutes each. After
dehydration the infiltration process requires steps through an intermediate
solvent, 2 changes of 100% propylene oxide (P.O.) for 15 minutes
each and finally into a 50:50 mixture of P.O. and the embedding
resin (Embed 812, Electron Microscopy Sciences, Fort Washington,
PA) for 12-18 hours. The specimen is transferred to fresh 100% embedding
media for at least one hour. The tissue is then embedded in a fresh
change of 100% embedding media. Following 12-18 hours in the oven
at 60°C. for polymerization, the blocks are then ready to section.
Protocol for sectioning is on a separate page. |
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2.
Standard Procedure To Section for Transmission Electron Microscopy,
EM Center, Indiana University School of Medicine:
The resin blocks are first thick sectioned at 1-2 microns with glass
knives using an Ultracut UCT (Leica, Bannockburn, IL) or a MT 5000
(RMC, Tucson, AZ) and stained with Toluidine Blue, these sections
are used as a reference to trim blocks for thin sectioning. The
appropriate blocks are then thin sectioned using a diamond knife
(Diatome, Electron Microscopy Sciences, Fort Washington, PA)) at
70-90nm (silver to pale gold using color interference) and sections
are then placed on either copper or nickel mesh grids. After drying
on filter paper for a minimum of 1 hour, the sections are stained
with the heavy metals, uranyl acetate and lead citrate for contrast.
After drying the grids are then viewed on a Tecnai BioTwin (FEI,
Hillsboro, OR). Digital images are taken with an AMT CCD camera
and saved on a CD for the researcher. |
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3.
Standard Procedure to Process and Embed Tissue Cultures on Coverslips,
EM Center, Indiana University School of Medicine:
Cells can be grown on Thermanox Coverslips (Nalge Nunc International,
supplied through Fisher Scientific). After the appropriate amount
of time in culture, the coverslips are drained of their media and
immediately placed in the appropriate fixative for a minimum of
an hour. The fixative routinely used in this lab is a modified Karnovsky's,
2% Paraformaldehye/2% Glutaraldehyde in 0.1M Phosphate Buffer. After
fixation the specimens are rinsed several times with phosphate buffered
saline (PBS) followed by post fixation with 1% osmium tetroxide
in phosphate buffer for one hour. After rinsing again with PBS for
15 minutes, the tissue specimens are dehydrated through a series
of graded ethyl alcohols from 70 to 100%. The schedule is as follows:
70% for 10 min., 95% for 10 min. and three changes of 100% for 5
minutes each. Make sure the specimen (coverslip) is not allowed
to dry out. After dehydration the infiltration process requires
a 12-18 hour stay in a 50:50 mixture of 100% ethyl alcohol and embedding
resin (Embed 812, Electron Microscopy Sciences, Fort Washington,
PA). Do not use propylene oxide (P.O.) as an intermediate solvent
as it will dissolve the coverslip and petri dish. The following
day the coverslips are placed in a change of 100% embedding media
for a minimum of 1 hour. Coverslips can then be embedded by two
different methods. The first method is for cross sections through
the sample. Cut the coverslips to an appropriate size to fit into
a flat embedding mold and place in a mold with fresh resin. The
second method is for sectioning in the plane of a monolayer. Overfill
Easy Molds (Ted Pella, Inc, Redding, CA) with fresh resin and place
coverslip on top with cell side down. Following a 12-18 hour stay
in a 60°C. oven for polymerization the both types of samples
are then ready for sectioning. After polymerization of the monolayer
blocks the coverslip is peeled off using liguid nitrogen leaving
the cell layer behind. |
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4.
Standard Procedure for Post-Embedding Immunostaining on Grids, EM
Center, Indiana University School of Medicine:
Tissue samples are fixed with 2-4% Paraformaldehyde in 0.1M phosphate
buffer, dehydrated through a graded series of ethyl alcohols and
embedded in Unicryl (Vector Labs). Thin sections (70-90nm) are mounted
on Formvar/carbon coated nickel grids. After drying the grids are
floated on drops of 0.05M glycine for 15 minutes to quench the aldehydes.
After rinsing* with 0.1M Phosphate buffer or PBS, the grids are
then placed into the Blocking buffer for a block/permeablization
step of 30-45 minutes. The grids are then placed in the primary
antibody overnight at 4°C. During the immuno-labeling process,
we do not let the grids dry out. The grids are then rinsed* with
the incubation buffer and then floated on drops of the secondary
antibody with attached 1.4 nm gold particles (AURION) for 2 hours
at room temperature or overnight at 4°C. After rinsing* in a
rinse buffer (incubation buffer without Tween) the grids are placed
in 2.5% Glutaraldehyde in 0.1M Phosphate buffer for 5 minutes. After
rinsing* in PBS (Phosphate Buffered Saline) and distilled water,
the grids are then ready for Silver Enhancement (AURION) for 45
minutes or GoldEnhance (Nanoprobes, Inc.) for 2 minutes. After rinses*
in distilled water, the grids are allowed to dry and then are stained
for contrast with uranyl acetate and lead citrate. The samples are
viewed with a Tecnai Bio Twin transmission electron microscope (FEI,
Hillsboro, OR). The block/perm buffer consists of 2% BSA, 0.1% Cold
Water Fish Gelatin and 0.1% Tween in PBS. The primary and secondary
antibodies are diluted in an incubation buffer containing 0.1% BSA-c
(AURION), 0.05% Tween in PBS. Times and dilutions are determined
for each particular primary antibody being used. *All rinses are
3 times for 5 minutes each. Other types and sizes of gold can be
used, such as Protein-A-Gold at 10nm. This size of gold does not
require enhancement. |
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| 5. Standard Procedure to Embed Tissue in LR Gold for Immunoelectron Microscopy. Tissue is fixed or perfused with 4% paraformaldehyde in 0.1M sodium cacodylate buffer to which 0.05 or 0.2% glutaraldehyde may be added. If phosphate buffer is used, it must be rinsed out of the tissue with sodium cacodylate buffer before an en bloc stain with uranyl acetate is used. The tissue is cut down to 0.5 x 0.5mm or less before processing. The tissue is rinsed three times in 0.1M sodium cacodylate buffer with 3.5% sucrose at 4 ° C, followed by 1 hour in 50mM ammonium chloride in the same buffer. The tissue is then rinsed 3x in 0.1M maleate buffer with 3.5% sucrose, pH 6. The tissue is then placed in 2% uranyl acetate in maleate/sucrose buffer for 2 hrs. At 4 ° C. After en bloc staining the tissue is rinsed three times in maleate/sucrose buffer. The tissue is dehydrated up to 90% acetone as follows: 1 hour in 50% acetone at 4 ° C, 1.5 hours in 70% acetone at -20 ° C, 1.5 hours in 90% acetone at -20 ° C. Infiltration with LR Gold is done at -20 ° C on a rotator as follows: 1:1, 90% acetone: LR Gold with no bensoin methyl ether (BME) for 2 hours; 100% LR Gold with no BME overnight; two changes of LR Gold with 0.5% BME two times during the day; two changes the next day; fresh LR Gold with BME. The tissue is embedded and polymerized at -20 ° C in a chest freezer.
The blocks with tissue are sectioned and placed on formvar, carbon coated nickel grids. After immunolabeling the sections may be post stained with saturated uranyl acetate only or uranyl acetate and lead citrate. For increased membrane staining, after immunolabeling grids may be place on a drop of 2% aqueous osmium tetroxide for 10 to 15 minutes and then post stained with UA and Pb. |
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6.
Standard Procedure for Pre-Embedding Immunostaining on Vibratome
Sections, EM Center, Indiana University School of Medicine:
After fixation in 2-4% Paraformaldehyde, tissues samples are vibratomed
(50 microns). After rinses* in PBS, the sections are placed in
0.1% sodium borohydride for 15 minutes to quench the aldehydes.
After rinsing in 0.1M phosphate buffer follows until all the bubbles
are gone, do not use Phosphate buffered saline (PBS) the samples
are placed into a Blocking buffer for the block/permeablization
step for 45 minutes. The samples are then ready for incubation
in the primary antibody overnight at 4°C. The sections are
rinsed* with the incubation buffer and placed into the secondary
antibody attached to 1.4nm gold particles (AURION) for 2 hours
at room temperature or overnight at 4°C. After rinsing* in
the rinse buffer (incubation buffer without Tween) the sections
are placed in 2.5% Glutaraldehyde in 01.M phosphate buffer for
30 minutes. After rinsing* in PBS (phosphate buffer saline) and
distilled water the samples are then ready for silver enhancement
(AURION) for 45 minutes. After rinses* in distilled water and
PBS, the sections are post fixed with 0.5% osmium and processed
for standard embedment using Embed 812 (Electron Microscopy Sciences).
The block/perm buffer consists of 2% BSA, 0.1% Cold Water Fish
Gelatin and 0.1% Tween in PBS. The primary and secondary antibodies
are diluted in an incubation buffer containing 0.1% BSA-c (AURION)
and 0.05% Tween in PBS. |
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7.Standard Procedure for Cryosectioning
Tissue or cell culture may be processed for immunohistochemistry using semi-thin (0.5-2 µm) or immunocytochemistry using ultra thin (50- 100nm) cryosections using the Leica Ultracut UCT with the FCS cryo attachment. The tissue is fixed with 4% paraformaldehyde in phosphate buffer, cryoprotected in 2.3M sucrose and kept frozen until sectioning. Semi-thin sections are picked up on glass slides for processing using a fluorescent label for light or confocal microscopy or they may be immunogold labeled and silver enhanced. For unltra-thin sections the frozen tissue is trimmed using a Diatome cryo trim knife and then thin sectioned using a Diatome 35° dry cryo diamond knife. The sections are picked up in a sucrose/methycellulose mixture and put on formvar carbon coated grids. The sections are immunolabled, stained with uranyl acetate and embedded in a 2% methycellulose/uranyl acetate mixture. This procedure is good for labeling membranes and also retains antigens that may be lost when tissue is dehydrated and embedded in a resin.
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8. High Pressure Freezing-Automated Freeze Substitution
High Pressure Freezing (HPF) is a method of fixation that uses ultra rapid freezing at high pressure to preserve cells and tissue that shows superior ultrastructural preservation and retention of antigens using a Leica, EM PACT high pressure freezer, Microbiopsy system, and Automatic Freeze Substitution Unit (AFS).
Tissue must be quickly excised from a live anesthetized animal or specimens taken with the Microbiopsy system. Tissue cultures cells may be brought in buffer as a cell suspension or grown on sapphire discs. Once the tissue or cells have been HPF they may be stored in liquid nitrogen indefinitely. Experiments with the proper application may require moving the EM Pact next to a confocal or light microscope for correlative studies.
Once the samples are frozen they may be freeze substituted. Substitution protocols must be worked out for each tissue. A basic freeze substitution protocol for morphology would be 3 days at -90°C in 1% OsO4 in acetone followed by a slow warm up to room temperature and embedded in Epon. Substitution media for immunocytochemistry will vary but may include small percentages of glutaraldehyde (0.01%) or uranyl acetate (0.25 – 0.5%) in acetone at -90 for 3 days followed a slow warm up to a temperature compatible with the resin to be used, Unicryl, LR Gold -20°C, Lowicryl -35° or HM20 -50°C.
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9.
Standard Procedure to Prepare Specimens for Scanning Electron Microscopy,
EM Center, Indiana University School of Medicine:
Fix the specimen with an appropriate aldehyde fixative for a minimum
of 1 hour, depending on the size of the specimen. This laboratory
routinely uses a modified Karnovsky's fixative, 2% Paraformaldehye/2%
Glutaraldehyde in 0.1M Phosphate Buffer. After initial fixation,
the specimens are rinsed several times with PBS (phosphate buffered
saline) for a minimum of 15 minutes, followed by post fixation with
1% Osmium tetroxide in 0.1M Phosphate Buffer for 1 hour. After rinsing
with PBS for a minimum of 15 minutes the specimens are dehydrated
using a series of graded ethyl alcohols (70% for 15 min, 95% for
15 min. and 3 changes of 100% for 10 min. each). From this point
on there are two methods this lab uses routinely for drying the
tissue, either critical point drying using a Samdri-790 (Tousimi,
Rockville, MD) or chemically drying using HMDS (hexamethyldisilazane).
A protocol for this procedure is on a separate page. After drying,
the specimens are mounted on aluminum stubs with adhesive tabs and
sputter coated for 3 minutes using a Polaron (Energy Beam Sciences,
Agawam, MA). The specimen is then ready to view on the AMRAY 1000A
(Bedford, MA) scanning electron microscope. |
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10.
Specimen Preparation for Scanning Electron Microscopy Using Chemical
Drying, EM Center, Indiana University School of Medicine:
After the appropriate primary fixation, post fixation and dehydration
through 100% ethyl alcohol the specimen is then ready for chemical
drying. The schedule is as follows: 2 parts 100% ethyl alcohol/1
part HMDS (hexamethyldisilazane, Electron Microscopy Sciences,
Fort Washington, PA) for 15 minutes, 1 part 100% ethyl alcohol/2
parts HDMS for 15 minutes, then 2 changes for 15 minutes each
with 100% HDMS. Finally remove as much of the HDMS as you can
and allow the specimen to air-dry in a hood over night. The samples
are then ready to mount and sputter coat. |
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11. Negative Staining of Particles on Grids (for virus, bacteria, small particles, etc.) for Transmission Electron Microscopy, EM Center, Indiana University School of Medicine:
Fix the specimen with the appropriate fixative. An Optimal concentration and clean specimen is needed for the best negative staining. The specimen is dropped onto a 200-400 mesh carbon/formvar coated grid and allowed to absorb to the formvar for a minimum of 1 minute. The excess liquid does not need to be wicked off. A drop of the negative stain is placed on the grid for the appropriate amount of time for the stain you are using and type of specimen. The EM Center uses either 1% aqueous uranyl actetate made up fresh or Nanovan (Nanoprobes, Inc. Yaphank, NY). For both stains the time in the stain is very short, generally less than 1 minute. The time needs to be worked out for the specimen you are using. The excess liquid is then wicked off and the grids are allowed to dry. After the specimen is placed into the TEM, allow the specimen to sit for a few minutes so the sample can be vacuumed dried before being irradiated. The sample is then ready for viewing and images. |
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