The standard procedures and protocols for the IU School of Medicine Electron Microscopy Center are provided here.
For all specimens brought for immunostaining (non-standard processing), the Center does not recommend having any Glutaraldehyde in the fixative. The Electron Microscopy Center recommends 4% Paraformaldehyde in a 0.1M buffer, either phosphate or cacodylate. The specimen should always be submitted in fixative—and not rinsed in a buffer. Paraformaldehyde does not cross link completely and will leach out of the tissue if left in buffer; therefore the tissue becomes unfixed and ruined.
Biohazardous specimens must always be brought in fixative.
For all specimens brought to the Electron Microscopy Center for standard processing, the Center recommends using the fixative provided by the lab. Fixation for electron microscopy is extremely important; if not done correctly, there is no way to correct it. Investigators who want to make up their own fixative must clear it with the Electron Microscopy Center first.
Fix specimens with an appropriate aldehyde fixative for a minimum of an hour, place specimens into the fixative immediately, do not allow to air dry. Ideally, specimens should be no larger than 3mm in width. Fixative routinely used by the EM Center and provided, 3% Glutaraldehyde in 0.1M Phosphate buffer, cacodylate buffer is used for muscle and nerve specimens. After primary fixation, specimens are rinsed in the appropriate buffer and post fixed in 1% Osmium tetroxide in buffer for at least 1 hour. After rinsing in buffer the specimens are dehydrated through a series of graded alcohols from 70-100%, after 2 changes with an intermediate solvent, 100% acetone, specimens are placed in a 50:50 mixture of acetone and embedding media (Embed 812, Electron Microscopy Sciences) for a minimum of overnight. The following day specimens are placed into a fresh change of pure resin for at least 4 hours, then embedded in another fresh change of 100% resin, and placed in a 60°C oven for 12-18 hrs for polymerization. The blocks are then ready to section.
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 placed on either copper or nickel mesh grids. After drying on filter paper for a minimum of one hour, the sections are stained with the heavy metal Electron Microscope Sciences, UA replacement stain for contrast. After drying, the grids are viewed on a Spirit (FEI, Hillsboro, OR). Digital images are taken with an AMT CCD camera and are uploaded to a Box folder for the researcher.
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 (see note above) with 0.1M Phosphate buffer or PBS, the grids are 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, the grids are not allowed to dry out. The grids are 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 two 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 rinsed 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 rinsed 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 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 two 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.
Fix specimens with an appropriate aldehyde fixative for a minimum of an hour. Fixative routinely used by the EM Center and provided, 3% Glutaraldehyde in 0.1M Phosphate buffer. For larger specimens can use 2% Paraformaldehyde/2% Glutaraldehyde in 0.1M Phosphate buffer. After primary fixation, specimens are rinsed in buffer and post fixed in 1% Osmium tetroxide for at least 1 hour. After rinsing in buffer the specimens are dehydrated through a series of graded alcohols from 70-100%. From this point on the lab routinely used chemical drying, HMDS. After drying, the specimens are mounted on aluminum stubs with adhesive or carbon tabs, sputter coated and then ready to view on JEOL 6390LV (Peabody, MA) scanning electron microscope.
After the appropriate primary fixation, post fixation and dehydration through 100% ethyl alcohol, the specimen is ready for chemical drying. The schedule is as follows: two parts 100% ethyl alcohol/one part HMDS (hexamethyldisilazane, Electron Microscopy Sciences, Fort Washington, PA) for 15 minutes, one part 100% ethyl alcohol/two parts HDMS for 15 minutes, then two changes for 15 minutes each with 100% HDMS. Finally as much of the HDMS as possible is removed and the specimen is allowed to air-dry in a hood overnight. 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 one 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 being used and type of specimen. The Center uses Nanovan (Nanoprobes, Inc. Yaphank, NY). The time in the stain is very short, generally less than one minute, and this time needs to be worked out for the specimen being used. The excess liquid is then wicked off and the grids are allowed to dry. After the specimen is placed into the TEM, investigators should 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.