List of standard EM Center procedures/protocols:
- Standard Procedure to Prepare Specimens for Transmission Electron Microscopy
- Standard Procedure to Section for Transmission Electron Microscopy
- Standard Procedure to Process and Embed Tissue Cultures on Coverslips
- Standard Procedure for Post-Embedding Immunostaining on Grids
- Standard Procedure for Pre-Embedding Immunostaining on Vibratome Sections
- Standard Procedure for Cryosectioning
- Standard Procedure for High Pressure Freezing-Automated Freeze Substitution
- Standard Procedure to Prepare Specimen for Scanning Electron Microscopy
- Specimen Preparation for Scanning Electron Microscopy Using Chemical Drying
- Negative Staining of Particles on Grids (for virus, bacteria, small particles, etc)
*Note: With all specimens brought to the EM Center for immunostaining (non-standard processing ~#s 4 & 5 above):
- The EM Center does not recommend having any Glutaraldehyde in the fixative
- The EM Center recommends 4% Paraformaldehyde in a 0.1M buffer, either phosphate or cacodylate
- ALWAYS bring your specimen in fixative, do not rinse in a buffer
- Paraformaldehyde does not cross link completely and will leach out of the tissue if left in buffer, therefore the tissue will become unfixed and ruined
**Note: With all specimens brouth to the EM Center for standard processing (#1 above):
- Biohazardous specimens must always be brought in fixative
- The EM Center recommends using the fixative provided by the lab. Fixation for electron microscopy is extremely important, if not done right, there is no way to correct it. if you do want to make up your own fixative, please clear it with the EM Center first.
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 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, Hatfield, PA) for 12-18 hours. The specimen is transferred to fresh 100% embedding media under vacuum overnight. The following day the specimen 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.
The resin blocks are first thick sectioned at 1-2 microns with glass knives using an Ultracut UCT (Leica, Bannockburn, IL) 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, Hatfield, 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 metal uranyl acetate 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.
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 coverslips 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 coveslips 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 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, Hatfield, 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 under vacuum for a minimum of 4 hours. 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 liquid nitrogen leaving the cell layer behind.
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 (Electron Microscopy Sciences, Hatfield,PA). Thin sections (70-90nm) are mounted on Formvar/carbon coated nickel grids. 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 phosphate buffered saline (PBS) and then floated on drops of the appropriate secondary antibody attached with 10nm gold particles (AURION, Hatfield, PA) for 2 hours at room temperature. After rinsing with PBS the grids are placed in 2.5% Glutaraldehyde in 0.1M Phosphate buffer for 15 minutes. After rinsing in PBS and distilled water. After rinses in distilled water, the grids are allowed to dry and then are stained for contrast with uranyl acetate. 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. Other types and sizes of gold can be used from 1.4nm-25nm. 1.4nm sized gold requires silver enhancement to visualize at the TEM level.
After fixation in 2-4% Paraformaldehyde, tissues samples are vibratomed (50 microns). After rinses in phosphate buffered saline (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 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 PBS and placed into the secondary antibody attached to 10nm gold particles (AURION, Hatfield, PA) for 2 hours at room temperature. After rinsing in PBS the sections are placed in 2.5% Glutaraldehyde in 01.M phosphate buffer for 30 minutes. After rinsing in PBS the sections are post fixed with 0.5% osmium and processed for standard embedment using Embed 812 (Electron Microscopy Sciences, Hatfield, PA). 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.
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 ultra-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.
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.
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 JEOL 6390LV (Peabody, MA) scanning electron microscope.
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.
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 Nanovan (Nanoprobes, Inc. Yaphank, NY). The time in the stain is very short, generally less than 1 minute and this 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.