Transmission Electron Microscopy Tem Biology Essay

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This unit describes TEM preparation techniques for examining particulate samples as well as samples presenting more complex ultrastructural considerations that require analysis in thin sections. Negative staining is a simple but valuable technique that is useful for routine examination of particulate samples in suspension ranging from bacteria to viruses to purified macromolecules. .

Safety

Aside from the biohazards posed by handling and disposing of pathogenic microbes (refer to unit 1A.1), many of the chemicals used in the processing of samples for electron microscopy are either volatile, inflammatory, toxic, carcinogenic, and/or radioactive. It is important to understand the nature of health and environmental hazards posed by these chemicals and to handle and dispose of them according to appropriate safety regulations (see units 1A.1, 1A.3 & 1A.4). Expose electron microscopy grids containing negatively stained pathogenic specimens to UV radiation or chemical fixation before examination and store in a suitable container for safe temporary storage. Discard grids into a small wide-based container with appropriate decontamination reagents upon completion of the analysis. Rinse forceps used to handle grids with pathogenic samples in alcohol and flame before and after use. Specimens processed for embedding and thin sectioning are typically chemically stabilized in a fresh aldehyde-containing fixative, which will inactivate/kill the majority of viruses and bacteria. Autoclave other disposable preparative materials that come into contact with pathogens (e.g., Parafilm, filter paper) in appropriate containers prior to disposal.

Fix cells (Standard protocol)

1.

In a fume hood, add an equal volume of TEM primary fixative 2 to bacterial cells suspended in culture medium or buffer in a 15-ml conical centrifuge tube.

This will result in a final fixative concentration of 2% glutaraldehyde.

2.

Immediately invert mixture and place on a horizontal mixer 60 min at room temperature.

3.

After initial room temperature fixation, transfer cell suspension to a 4°C ice bath for an additional 1 hr of fixation.

4.

Centrifuge cell suspension 10 min at 1000 Ã- g, 4°C, and discard supernatant.

5.

Resuspend cells in phosphate/sucrose rinse buffer and incubate at 4°C for 10 min.

6.

Centrifuge cells 10 min at 1000 Ã- g, 4°C, and discard supernatant.

Encase cells in agar and fix again

7.

Resuspend cells in enough 2.5% molten agar to cover pellet, mix with a warmed transfer pipet, and transfer to polyethylene microcentrifuge tubes.

8.

Centrifuge the agar and cell suspension 5 min at 5000 Ã- g, room temperature, to pellet cells.

9.

Chill tubes briefly on ice to solidify agar.

10.

Using a single-edged razor blade, cut through the microcentrifuge tube just above the cell pellet in the agar to remove the pellet. Slice the removed agar encased pellet into thin slices, ~0.5 mm wide, and place in fixation vials containing chilled phosphate/sucrose rinse buffer.

Perform post-fixation and en bloc staining

11.

Replace buffer with osmium tetroxide post-fixative and incubate 90 min at 4°C. Rock pellets frequently during post-fixation or place vials on a platform rocker and rock at low speed to facilitate penetration of osmium into the tissue.

12.

Rinse pellets twice in chilled phosphate/sucrose rinse buffer.

13.

Rinse pellets six times over the course of 1 hr with chilled distilled water to remove phosphate buffer residue.

14.

Stain pellets en bloc in uranyl acetate staining solution for 90 to 120 min at 4°C with periodic agitation.

Alternatively, stain in 0.5% aqueous uranyl acetate overnight at 4°C.

15.

Wash three times in cold distilled water to remove uranyl acetate.

Dehydrate and embed pellet

16.

Dehydrate through ethanol series by immersing successively for 10 min each in the following solutions:

50% ethanol, 4°C

75% ethanol, 4°C

95% ethanol, room temperature.

Transfer to 100% ethanol.

17.

Rinse tissue three times, each time for 10 min, in acetone or propylene oxide.

18.

Embed in epoxy resin (see Basic Protocol 4).

Basic Protocol 4: Embedding of Tissues and Cell Pellets for Thin Sectioning

The following basic protocol is a continuation of Basic Protocol 3 and Alternate Protocols 4 and 5. All material to be thin sectioned for TEM examination must be embedded in a hardened plastic matrix. The unpolymerized plastic matrix must be fluid enough to completely infiltrate the specimen. Once polymerized, the plastic must not alter the ultrastructure of the specimen (i.e., as a result of specimen shrinkage, swelling, or extraction of cellular components) while being able to withstand the stress of thin sectioning. The resulting sections should remain stable in the electron beam and impart adequate contrast to the sample to allow for the visualization of ultrastructural detail. An additional consideration is the preservation of antigenicity in the case of thin-section immunogold labeling; a separate procedure is provided to deal with this issue (see Alternate Protocol 9).

In general, the goal is to introduce a plastic resin into the sample over time by the gradual replacement of dehydrating agent and resin solvent (usually ethanol and acetone, or ethanol and propylene oxide) with increasing concentrations of resin and solvent over time. Tissue pieces and cell pellets are eventually transferred to resin-filled embedding molds for polymerization. Cells in polymerized resin can then be prepared for thin sectioning.

CAUTION: Some resin components are possible carcinogens and others can produce allergic reactions in certain individual. Their preparation may therefore require a fume hood, a suitably vented oven, and other adequate measures for personal protection (also see unit 1A.3).

Materials

Fixed and processed tissue (Basic Protocol 3), cell pellet (Alternate Protocol 4), or bacterial sample (Alternate Protocol 5)

Luft's epoxy mixture (see recipe)

Acetone or propylene oxide

Orbital shaker

Transfer forceps or applicator stick

Embedding molds or capsules (available from Ted Pella; BEEM flat embedding molds, cat. no. 111-2, PTFE flat embedding molds, cat. no. 10509, or BEEM embedding capsules, size 00, cat. no. 13)

60°C oven

1.

Pipet out the last rinse of acetone or propylene oxide from the fixation vial and replace with a mixture of 50% (v/v) Luft's epoxy mixture and 50% acetone or propylene oxide. Place the vial on an orbital shaker and rotate sufficiently to suspend the tissue pieces or pellet in the resin.

Upon initial introduction of the resin, the tissue pieces or pellets will float to the top. With time, they will sink as they become infiltrated.

2.

Incubate at room temperature for 30 min with rotation on the orbital shaker to aid infiltration.

3.

Remove all of the 50-50 resin mixture and replace with a mixture of 75% (v/v) Luft's epoxy mixture and 25% acetone or propylene oxide. Incubate at room temperature for 30 min with rotation to aid infiltration.

The pellets will again float to the top of the suspension and will slowly sink as the resin infiltrates the pellets.

4.

Remove the resin from step 3 and replace it with 100% Luft's epoxy mixture. Rotate on an orbital shaker for 1 hr at room temperature, then remove the epoxy mixture and replace with freshly prepared 100% Luft's epoxy mixture.

5.

Remove the resin and, using transfer forceps or an applicator stick, carefully transfer pellets to suitable embedding molds containing freshly prepared 100% Luft's epoxy mixture. Transfer molds to a vented 60°C oven and incubate overnight.

A variety of embedding molds and capsules are commercially available that allow tissue to be optimally positioned for sectioning once released from the mold. For pelleted material, polyethylene caps for snap-cap sample vials (Electron Microscopy Sciences #64257-30) work well for this purpose. They come in a variety of sizes, allow for easy placement of the sample, have plenty of surface area for outgassing of residual solvent if necessary, and are reusable; in addition, the polymerized resin is easily removed. Pellets or tissue can be excised from the resin using a jeweler's saw or a single-edged razor blade if the resin is slightly heated and mounted on a blank BEEM capsule or Plexiglas pegs (Ladd Research #21830) using superglue. Small pieces of tissue can be directly placed in resin-filled BEEM capsules for polymerization. However, when BEEM capsules are used for direct embedding of tissue, it is often advantageous to employ flat embedding molds for embedding extra pieces of tissue if the need arises.

Fixation and Initial Processing of Samples for Immunogold Labeling of Thin Sections

The fixation and processing of samples that will permit labeling of antigens in thin-sectioned materials pose substantial challenges. Methods commonly used for optimal ultrastructural morphology must usually be modified to ensure that antigens are not only retained but available for labeling. This often involves optimization, with compromises between the fixation, processing, and embedding steps; however, careful selection of preparation methods can yield high-quality combinations of structural detail and immunochemical labeling.

Additional Materials (also see Basic Protocol 3)

Tissue or cell sample

TEM primary fixative 3 (see recipe)

Aldehyde quenching solution (see recipe)

1a.

For tissue specimens: In a fume hood, fix tissue specimens by incubating with TEM primary fixative 3 for 12 to 24 hr at 4°C.

1b.

For free cells or cell suspension: In a fume hood, fix cells or pellet by incubating with TEM primary fixative 3 for 2 to 4 hr at 4°C.

2.

Rinse in phosphate/sucrose rinse buffer three times for at least 10 min each at 4°C.

3.

Incubate in aldehyde quenching solution for 30 min at 4°C to remove residual free aldehydes.

4.

Wash in phosphate/sucrose rinse buffer two times for 15 min each at 4°C.

5.

Dehydrate through ethanol series by immersing successively in the following solutions for the indicated amounts of time:

50% ethanol, 4°C

10 min

75% ethanol, 4°C

10 min

95% ethanol, room temperature

10 min

100% ethanol, room temperature

30 min

100% ethanol, room temperature

30 min.

6.

Directly embed in resin (see Alternate Protocol 9).

Note that this procedure omits the acetone or propylene oxide infiltration step included in the previous tissue fixation and processing protocols.

. Resin Embedding for Thin Section Immunogold Labeling

Preservation of sample antigenicity is of paramount importance when choosing an embedding resin for thin section immunogold labeling. Dehydration and embedding protocols must minimize loss of potential antigenic sites. This generally requires the use of ethanol alone as the dehydrating agent, an altered infiltration schedule, and the use of a more hydrophilic embedding resin coupled with room-temperature or cold polymerization. Several specific resins are available, all acrylic based, that offer good antigen protection, although some require elaborate cooling chambers in order to optimize their superior antigenic preservation qualities during polymerization. The resin protocol described below utilizes a one-part resin, which only needs an accelerator for room temperature polymerization and yields consistent results.

Materials

Fixed and processed tissue or cell sample (Alternate Protocol 7)

LR White resin (Polysciences)

LR White accelerator (Polysciences)

Embedding molds (available from Ted Pella; BEEM flat embedding molds, cat. no. 111-2, PTFE flat embedding molds, cat. no. 10509, or BEEM embedding capsules, size 00, cat. no. 13)

Cooled water bath or cooling block (optional)

Jeweler's saw

Plexiglas pegs (Ladd Research, cat. no. 21830; http://www.laddresearch.com/)

Cyanoacrylate glue (e.g., Superglue, Krazy Glue)

1.

Following the last two washes of tissue in 100% ethanol described Alternate Protocol 7, infiltrate the sample with LR White resin as follows. Remove the ethanol and replace with 100% LR White resin, incubate 1 hr at room temperature, then replace with fresh 100% LR White resin and incubate an additional 1 hr at room temperature. Finally, replace again with fresh 100% LR White resin and incubate an additional 1 hr to overnight at room temperature.

See technical data sheet for LR White resins at http://www.polysciences.com/shop/assets/datasheets/305A.pdf.

2.

To 10 ml LR White resin, add 20 µl LR White accelerator and mix well. Partially fill embedding mold(s) with the resin/accelerator mixture. Remove LR White resin-infiltrated tissue pieces or pellets from their vials using fine-tipped forceps or a wooden stick with a fine tip. Deposit the tissue pieces into the resin-containing embedding mold(s). Allow molds to polymerize for 48 hr at room temperature.

If flat embedding molds are used, it may be necessary to exclude air from the surface of the resin and overlay the mold with a piece of ACLAR film (Ted Pella, cat. no. 10501-10) to exclude as much air as possible from the surface of the mold. Embedding in BEEM or gelatin capsules is preferred over the use of flat embedding molds, as the surface area exposed to air is minimized and the capsules do not require the film overlay.

3.

Remove polymerized block(s) from embedding mold(s) or capsule(s). If the tissue has been flat-embedded, cut out the desired area with a jeweler's saw and mount on Plexiglas specimen-mounting pegs (that fit into the specimen chucks of the ultramicrotome) using cyanoacrylate glue.

Basic Protocol 5: Overview of Ultramicrotomy

As noted previously, the goal of ultramicrotomy is to prepare thin sections between 50 and 90 nm thick. The generation of sections that adhere to one another as straight ribbons using an ultramicrotome is an extremely challenging manual skill that requires a high level of dexterity. Ultramicrotomy is not a technique that can be effectively learned using a protocols manual and is therefore given as an overview rather than a method in this unit.

While it is assumed that the reader has access to ultramicrotomy support service or hands-on training in this key element of specimen preparation for analysis of thin sections, a brief description of ultramicrotomy follows. In order to thin-section embedded tissue that has been placed in an embedding mold, it must be mounted in a holder for trimming. Some ultramicrotomes have different types of holders that can grip flat embedded tissue or hold a cylinder of plastic with a tapered end (pyramid shape) containing the tissue. The tapered end must be trimmed further to form a small trapezoid of ~200 µm2. This is usually done by hand with a single-edged razor blade under a stereo microscope and requires considerable dexterity and practice. Some microtomes have specimen holders, which permit rapid trimming of the block to form the trapezoid with a highly polished face.

Thin sections are cut with either a glass or diamond knife. Glass knives are normally made freshly with a knife-maker instrument, which breaks ~1-in. (~2.5-cm) squares of glass from high-quality glass strips. The squares of glass are scored diagonally and carefully broken to form triangular pieces, one edge of which contains the knife edge. A commercially made plastic trough is sealed with dental wax. Alternatively, a trough or "boat" is fashioned around the knife edge with Mylar or other tape that will provide a distilled water-filled reservoir that reaches the edge of the knife and that will allow sections to float away from the edge of the knife from the block during the cutting stroke of the microtome arm. The wide side of the trapezoid-shaped block face makes first contact with the knife edge during the cutting stroke of the microtome. Prior to collecting sections, the block face must be "polished" in preparation for thin sectioning by cutting initial sections until the block face appears to have a mirrored surface. Thin sections are then cut, and individual trapezoid-shaped sections adhere to the knife edge until the subsequent section displaces the previous section during the cutting stroke of the microtome arm. With each consecutive section, a ribbon resembling the shape of a tapeworm floats into the boat (Fig. 2B.1.2). These ribbons are typically manipulated into parallel rows with a fine eyelash attached to the end of a small applicator stick just prior to picking up the sections on a 3-mm-diameter electron microscope grid. Sections are adhered to the grid by lowering it with a pair of forceps above the sections, gently touching the grid to the surface of the water, and then lifting the grid off the water surface in a smooth motion. Residual water is removed from the edge of the grid with a wedge of filter paper as is done for negative staining.

Figure

Figure 2B.1.2 Thin sections (~70 to 80 nm) prepared on a diamond knife as seen through a stereo microscope. Trapezoid-shaped sections are cut sequentially, adhering to the knife edge and forming ribbons during sectioning (black arrow). They are visible as the result of reflected light from above, which generates an interference pattern that can be used to estimate section thickness. As sections accumulate, they can be separated into smaller ribbons, which float on the water surface and can be aligned for collection onto a grid. The width of the trough of the diamond knife shown is ~10 mm.

Copper grids are routinely used for most biological samples; nickel grids are usually used with samples prepared for immunogold electron microscopy. The 50- to 90-nm thickness of each section can be estimated by adjusting the microtome stereomicroscope viewing and illumination system to detect interference colors reflecting from the sections. Generally, gray sections indicate a thickness of less than 60 nm, while silver sections range from 60 to 90 nm (the typical section thickness for most biological samples), and gold sections range from about 90 to 150 nm. The diamond knife, which contains a high-quality diamond with a cleaved knife edge mounted in an aluminum holder fashioned into a trough for floating sections, is a considerably more expensive and longer-lasting alternative to disposable, freshly made glass knives.

Alternate Protocol 10: Immunogold Post-Embedding Staining of Thin Sections

Localization of antigens in thin sectioned material usually requires special fixation procedures to limit the extent of macromolecular cross-linking, as well as embedment in a plastic resin which preserves antigenic sites. The following is one of many alternative procedures that can be used to localize antigens with immunogold reagents.

Materials

Thin sections on 300-mesh Formvar-coated nickel grids (Basic Protocol 5)

Immunogold blocking buffer (see recipe)

Primary antibody against antigen of interest and control (irrelevant) primary antibody of same Ig class

TBS-Tween (see recipe)

Reagents for colloidal gold labeling-one of the following:

Colloidal gold-labeled secondary antibody against species from which primary antibody was obtained

Biotinylated secondary antibody against species from which primary antibody was raised, and streptavidin-conjugated colloidal gold

Colloidal gold-labeled protein A and/or protein G

TEM primary fixative 2 (optional; see recipe)

Whatman no. 4 filter paper

Spray bottle

Grid box

Additional reagents and equipment for etching of epoxy embedding resin (optional; see Support Protocol 2) and uranyl acetate/lead citrate staining of thin sections (see Basic Protocol 6)

1.

If the sample is not embedded in LR White resin, perform Support Protocol 2.

Block sample

2.

Place an ~50-µl drop of immunogold blocking buffer onto Parafilm and float sample-containing grid on top of it for 30 min as a blocking step.

Treat with primary antibody

3.

Prepare appropriate dilution of primary antibody in TBS-Tween, place 50-µl droplets of the diluted antibody onto Parafilm, then place grids on droplets and incubate at room temperature for 2 hr.

Controls can be processed at the same time and should include both an irrelevant primary antibody of the same class and another preparation without primary antibody.

4.

Remove grid and draw off excess solution with a filter paper wedge (see Basic Protocol 1) and gently rinse with a stream of TBS-Tween from a spray bottle.

Label with colloidal gold

5a.

To use colloidal gold-conjugated secondary antibody: Place appropriate dilution of colloidal gold-conjugated secondary antibody onto Parafilm in 50-µl droplets, then transfer grids onto droplets and incubate at room temperature for 60 min.

5b.

To use biotinylated secondary antibody: Place appropriate dilution of biotinylated secondary antibody onto Parafilm in 50-µl droplets, then transfer grids onto droplets and incubate at room temperature for 1 hr. Wash grids briefly in 50-µl droplets of distilled water, then transfer to 50-µl droplets of streptavidin-conjugated colloidal gold and incubate at room temperature 60 min.

5c.

To use protein A and/or protein G: Place appropriate dilution of colloidal gold-conjugated protein A, protein G, or protein A/G onto Parafilm in 50-µl droplets, then transfer grids onto droplets and incubate at room temperature for 60 min.

6.

Rinse grids 1 min in a succession of four or five 50-µl droplets of TBS-Tween on Parafilm.

Post-fix (optional) and stain with heavy metals

7.

Optional: Post-fix by transferring grid to a 50-µl droplet of TEM primary fixative 2 for 10 min, then rinse in a 50-µl droplet of TBS-Tween for 1 min.

8.

Wash grid with a stream of Milli-Q or deionized distilled water from a spray bottle.

9.

Stain grids with uranyl acetate and lead citrate as described in Basic Protocol 6.

10.

Rinse grids in a gentle stream of distilled water and remove excess water with a filter paper wedge.

11.

Ensure that grids are completely dry and place them into a grid box. Store in a desiccator until ready to image in the electron microscope.

Basic Protocol 6: Heavy Metal (and Immunogold) Staining of Thin Sections

Heavy metal staining of thin sections is normally required to impart contrast in thin-sectioned biological materials. Two of the most widely used heavy metal post-staining solutions (i.e., uranyl acetate and lead citrate) are described. It is also possible to immunolocalize antigens in thin sections using colloidal gold particles of various sizes (5 to 20 nm) that effectively scatter electrons, which can be easily seen even when sections are post-stained with uranyl acetate and lead citrate.

Materials

Thin sections collected on copper grids (Basic Protocol 5)

Uranyl acetate staining solution (see recipe)

Lead citrate staining solution (see recipe)

Whatman no. 4 filter paper

Spray bottle with distilled H2O

Grid box

1.

Place 50-µl drops of uranyl acetate staining solution onto a clean Parafilm surface using the technique described in Basic Protocol 1.

2.

Using forceps, place grids containing sections face-down onto uranyl acetate drops using the technique described in Basic Protocol 1, and incubate for 3 to 10 min at room temperature.

3.

Remove grids from uranyl acetate drops and carefully direct a gentle stream of Milli-Q or deionized distilled water from a spray bottle onto the section side of the grid for 10 to 15 sec, slowly wetting the back side of the grid (i.e., the side without the section).

4.

Remove excess water with a wedge of Whatman no. 4 filter paper (see Basic Protocol 1); do not let section dry.

5.

Place 50-µl drops of lead citrate staining solution onto a clean sheet of Parafilm and deposit the grid, section-side-up, into the drop. Incubate 2 to 5 min at room temperature. Use a new drop of stain for each grid.

6.

Remove grids and gently run a stream of Milli-Q or deionized distilled water from a spray bottle down the tips of the forceps onto the grid to remove residual stain.

7.

Remove excess water with a filter paper wedge and ensure that no water remains between the tips of the forceps by sliding a piece of filter paper down between them while releasing the grid from the forceps onto a clean piece of filter paper.

8.

Ensure that grids are completely dry and place them into a grid box. Store in a desiccator until ready to image in the electron microscope.

9.

Place lead and uranyl waste in appropriate containers for disposal.

Reagents and Solutions

Use Milli-Q-purified or deionized, distilled water in all recipes and protocol steps. For common stock solutions, see appendix 2A; for suppliers, see suppliers appendix.

Aldehyde quenching solution

Prepare 100 mM glycine by dissolving 0.75 g in 100 ml phosphate/sucrose rinse buffer (see recipe). Alternatively, prepare 100 mM ammonium chloride by dissolving 0.53 g ammonium chloride in 100 ml phosphate/sucrose rinse buffer. Store up to 1 month at 4°C.

Immunogold antibody dilution buffer

To 180 ml Milli-Q-purified water or distilled H2O add:

1.04 g sodium phosphate, monobasic (Na2PO4Ã-H2O)

8.70 g sodium phosphate, dibasic, heptahydrate (Na2HPO4Ã-7H2O)

4 g bovine serum albumin (BSA), fraction V

0.6 ml Tween 20

Milli-Q-purified or deionized distilled water to 200 ml

Store up to 2 months at 4°C

Final concentrations 1 mM PBS containing 2% BSA and 0.3% Tween 20.

Immunogold blocking buffer

To 100 ml TBS-Tween (see recipe) add:

1 g bovine serum albumin (BSA), fraction V

3 ml normal serum from species in which secondary antibody was generated

Stir to dissolve

Store up to 6 months at 4°C

Final concentrations: 1% (w/v) BSA, 3% (v/v) serum in TBS-Tween.

TBS-Tween

To 1 liter of distilled water add:

6.1 g Trizma base

9 g NaCl

Mix to dissolve

Adjust pH to 7.6 using 1 N HCl

Add 0.5 ml Tween 20

Store up to 6 months at 4°C

Final concentrations: 0.5 M Tris-buffered saline, 0.05% Tween 20.

TEM fixative stock solutions

The most convenient source for EM-grade fixative stocks is from commercial electron microcopy suppliers (e.g., Electron Microscopy Sciences, Energy Beam Sciences, Structure Probe/SPI Supplies, or Ted Pella). For example, stock solutions of 16% paraformaldehyde, 8% or 25% glutaraldehyde, and 4% aqueous osmium tetroxide can be purchased in 10-ml quantities in sealed ampules individually or in boxes of 10 or more units. These stocks typically have a shelf life of 12 months if unopened, but once opened must be used in fixative solutions on the same day, preferably within hours of preparation.

Uranyl acetate staining solution

Immunoelectron microscopy

In general, immunoelectron microscopy applications involve the direct electron microscopy imaging of antigen-antibody complexes. A wide range of applications have been developed to improve the sensitivity of microbe detection and to reveal the ultrastructure of elusive organisms. When used to increase detection sensitivity, antibodies serve to aggregate viruses or bacteria in solution or onto a grid for negative staining. Immunogold staining can be combined with negative staining to provide a sensitive technique that enables the identification and visualization of individual antigens by TEM, and which can be useful in the identification of the biologic agent itself.

Fixation techniques for TEM

Ultrathin sections of samples provide insights regarding the internal structure of chemically stabilized samples. Individual sections that are used to provide images in the TEM represent two-dimensional views of three-dimensional objects; therefore, considerable efforts must be made to understand the three-dimensional ultrastructure of a sample. It is also important to appreciate that the appearance of the thin-sectioned sample is the result of the multiple steps in the protocol, including isolation of the sample, one or more chemical fixation steps, dehydration and embedding, sectioning, and staining.

Immunoelectron microscopy

Adequate controls must be incorporated into the procedure for the proper analysis of experimental results. Controls should include a grid in which the primary antibody incubation is omitted and a grid incubated with a nonspecific IgG of the same species as the primary antibody to identify nonspecific binding. Adjustments to increase labeling and decrease nonspecific binding may include, for example, use of different or additional blocking agents and/or concentrations and blocking times, changing concentration or incubation time of primary antibody, or addition of more rinses (see Table 2B.1.2).

Table 2B.1.2 Troubleshooting Guide for Negative Stain Immunoelectron Microscopy

Problem

Possible cause

Solution

No gold labeling

The antigen may be present in very low amounts

Use longer incubation times and more concentrated primary antibody

The primary antibody may be bad, e.g., due to poor titer, age, improper storage, improper dilution, or excessive freezing and thawing

If available, run a positive control to check

The pH of solutions may be excessively acidic or alkaline

Adjust the pH

Heavy negative staining may mask gold particles

Reduce the concentration of negative stain and/or use larger gold particles. Use higher magnification to visualize gold particles (e.g., 200,000Ã- for 5 nm; 100,000Ã- for 10 nm; 80,000 for 15 nm; 50,000 for 20 nm).

The section may not have been exposed to solutions (as a result of being wrong-side-up) if on a plastic film

Be careful when transferring and washing grids that the side of the grid containing the sample is kept facing up

The antigen may be destroyed by preparative procedures

Use a different procedure. Consider brief fixation in 1% paraformaldehyde.

Excessive background gold particles

Ionic concentration of solutions may be too low

Use increased salt concentration (up to 2.5%). Add ovalbumin, BSA, or normal goat serum (not for protein A) to ~1% in incubation solutions.

Sections may have been inadequately washed between incubations

Increase washing steps

Nonspecific charge attraction of antibody can cause background

Use 1% detergent (e.g., Tween 20) in all solutions. Include normal goat serum (not for protein A) in all solutions. Increase concentration of normal goat serum before primary antibody incubation.

Free aldehyde groups in fixed tissue may be a source of background

Reduce by exposing sections to 0.5 M ammonium chloride for 1 hr before incubations.

The primary antibody concentration may be too high

Dilute by orders of magnitude

The gold conjugate concentration may be too high

Dilute further

Clustering of gold particles

Clumped primary antibody.

Use fresh antiserum

Clustering can be caused by the natural amplification factor of the gold conjugate. For IgG-gold conjugates, up to 10 conjugated gold particles may attach to the Fc component of the primary antibody, producing the appearance of clusters on the section. This does not occur with protein A conjugates.

Use higher dilution of gold conjugates if desired

Gold particles over surface of support film

Nonspecific binding

Incorporate additional blocking steps prior to delivery of primary antibody

When considering the use of the protein A-, protein G-, or protein A/G-gold procedure, it is important to understand that, while these conjugates interact with mammalian immunoglobulins, their individual affinity for immunoglobulins is not equivalent for all species or for all antibody subclasses. For example, in the case of polyclonal antibodies, protein A or protein G can be used for human, pig, rabbit, or mouse. Horse and cow immunoglobulins have relatively high affinity for protein G, while sheep, goat, chicken, hamster and rat antibodies will bind protein G weakly. Guinea pig immunoglobulins will bind protein A. For use with monoclonal antibodies, human isotypes IgG1, IgG2, and IgG4 have good affinity for protein A or protein G, while human IgG3 has good affinity for protein G only. Protein G is used with rat isotypes, whereas mouse isotypes IgG1, IgG2a, IgG2b, and IgG3 all bind protein G, and mouse IgG2a and IgG2b also bind protein A. The use of protein A/G can therefore be useful if the antibody isotype is unknown.

Another important consideration is size and affinity of the conjugated gold particles for the primary antibody. In general, the smaller the gold particle, the lower the steric hindrance to antigen detection; however, a confounding consideration is that the smaller the gold particle, the fewer the molecules of protein A bound per gold particle. For example, it has been estimated that a 5-nm gold particle can bind about four protein A molecules, whereas a 20-nm particle can bind ~48 protein A molecules. Similarly, a 5-nm gold particle can bind about three secondary antibody molecules, while 10-nm gold particles can bind ~12 antibody molecules and 20-nm gold particles can bind ~ 48 antibody molecules. Therefore, the affinity of the gold particle can increase with increasing size. An additional consideration when immunogold staining is combined with negative staining is that the smallest gold particles (e.g., 5 nm) are much more difficult to detect in negatively stained preparations; therefore, the recommended range of gold particle sizes is from 10 to 30 nm.

The use of colloidal gold-tagged secondary antibodies described above is an alternative to protein A/G and involves a secondary antibody generated in a different species against the Fc fragment of the primary antibody. A third alternative involves the use of biotinylated secondary antibodies that are subsequently rendered visible in the TEM by binding of colloidal gold-conjugated streptavidin. Immunolocalization of two antigens in the same preparation is also possible if compatible primary antibodies are available (e.g., primary antibodies generated in different species). This procedure involves the sequential indirect labeling of two primary antibodies with secondary antibodies conjugated to gold particles of different size.

Fixation techniques for TEM

Preparation of the sample and the choice of fixative, stains, and plastic resin can affect the quality of the preparation. Unfortunately, there is no optimal fixative for all biological samples, although freshly prepared fixatives containing glutaraldehyde are considered among the best primary fixatives currently available. The useful property of glutaraldehyde, a five-carbon dialdehyde, is its ability to cross-link proteins. Freshly prepared glutaraldehyde at neutral pH will also polymerize into longer chain dialdehydes, which provide cross-links of variable size that efficiently stabilize protein structure. Paraformaldehyde is commonly combined with glutaraldehyde to better stabilize biological samples. In general, when fixatives are identified as containing paraformaldehyde, it usually means that they contain formaldehyde generated from the paraformaldehyde polymer of formaldehyde. The advantages of this freshly prepared one-carbon monoaldehyde, formaldehyde, which reacts in an aqueous solution as methylene glycol, are its rapid penetration properties and ability to polymerize and cross-link proteins and nucleic acids, and also modify the chemical properties of lipids. While not included here, acrolein is another highly reactive fixative that is often used in combination with other aldehydes such as glutaraldehyde and/or paraformaldehyde for fixing very dense specimens.

The primary aldehyde-fixation step is frequently combined with a post-fixation step involving osmium tetroxide, which acts as a fixative and electron stain and also as a mordant to enhance staining of the sections with lead. The primary fixatives and post-fixatives described in these protocols are generally considered to be good starting points for the preservation of solid tissues and cells in either suspension or pellets. More specific protocols are available in the literature that address specific fixation requirements of specific organisms and cells (Fassel et al., 1997; Karlyshev et al., 2001). Variations on fixative concentrations, fixation times, buffer types and osmolarity are all important considerations for optimal preservation of any particular sample.

The preparation of individual cells or aggregates of cells poses special challenges. Generating a sample of adequate size for processing usually requires concentration of cells by centrifugation into pellets. Maintenance of pellet integrity is also an important consideration during primary fixation, post-fixation, post-staining, dehydration, and embedding. If cells are not sufficiently concentrated during centrifugation before addition of fixative, as described in Alternate Protocol 5, there can be a tendency for the pellet to disintegrate during subsequent processing. If the pellet is of sufficient size to yield multiple slices for processing, some cell loss can be tolerated. If not, then it is important to encase the slices of pellet in a thin layer of agar to preserve the integrity of the pellet. Proper temperature management of the agar used to encase suspension fixed cells is also important during the preparative procedure. The agar must be warm enough to remain molten when the cells are introduced while at the same time not being hot enough to damage tissue and/or to allow the cells to effectively spin out of the agar during centrifugation. It is advisable, if the sample size permits, to subdivide the sample and perform multiple runs to ensure preservation of the pellet.

Preparation of tissues and cells for immunogold labeling requires a balance between preservation of tissue/cell ultrastructure and the preservation of antigens. The goal is to optimize these compromises in order to combine adequate structural detail and adequate immunochemical labeling. Concentrations of glutaraldehyde that are normally used for ultrastructure most often destroy or cross-link antigens. Therefore paraformaldehyde alone or in combination with dilute glutaraldehyde (0.1% to 1%) is a useful starting point for fixation. Tissues processed for immunolabeling in thin sections require special embedding procedures, as discussed below.

The protocols presented in this section can be classified as representing classical approaches to the preservation of tissue for examination by thin-section electron microscopy. Other techniques are available that offer the advantages of potentially better preservation of specimen ultrastructure and possibly shorter processing times at the expense of having specialized processing equipment, additional safety concerns, and special modifications of the electron microscope itself. Freeze substitution, accomplished by the formation of vitreous ice within the sample by either plunging the sample into cooled liquid propane or ethane or exposing samples to a jet of liquid propane or to high-pressure freezing, offers the advantage of better preservation of ultrastructure, and usually better antigen preservation as well. Microwave processing techniques offer the advantage of rapid specimen fixation and embedding times. All of these techniques require specialized preparation protocols adapted to suit the specific equipment on hand, and are beyond the scope of this chapter.

Embedding techniques for TEM

By their nature, epoxy resins and their components are viscous, and this viscosity tends to increase with time after the individual components of the resin are mixed. Because of this, it is advisable to prepare resins in small batches just prior to use. Dilutions of complete resin mixtures in dehydration agents are prepared by dispensing the resin components into new disposable beakers. Preparations of resin and dehydration agent are kept capped to prevent evaporation of the agent over time, an especially important consideration when working in a fume hood. In the case of epoxy resin, accurate weighing of all components and dilution of resins is of critical importance in their preparation to ensure a resin mixture that will give the proper characteristics. It is also important that the individual components of the resin be completely mixed. Inadequate mixing of resin components can result in poorly infiltrated samples that section poorly, that have poor specimen contrast, and that do not hold up in the electron beam. Shaking or vigorous stirring results in well mixed resin. Any resulting entrapped air is liberated upon standing or can be removed by use of a vacuum chamber for 5 to 10 min or until the generation of gas bubbles stops (the vacuum provided by a one-stage rotary pump is sufficient). Caution must be used when degassing the resin mixture, as it can boil out of the disposable beaker; careful monitoring of the mixture in the vacuum chamber is necessary. Furthermore, the ability to introduce air into the chamber to minimize initial boiling of the resin during degassing is also necessary.

TEM of thin sections

Staining with uranyl acetate and lead are relatively straightforward procedures. Occasionally, electron-dense precipitates occur that are the result of a staining problem. Care is taken with lead citrate to minimize the exposure to CO2, which can come from one's breath and result in the precipitation of insoluble lead carbonate. Precipitates can be formed from contaminants in glutaraldehyde and osmium tetroxide, and, as noted previously, uranyl phosphate forms an insoluble precipitate. For valuable samples, post-staining precipitates can sometimes be removed by using the plastic etching solution for 10 min, washing in a stream of distilled water, and then restaining (see Aldrich and Mollenhauer, 1986).

Troubleshooting suggestions for immunogold staining of negatively stained preparations are summarized in Table 2B.1.2 and are generally relevant to immunogold staining of thin sections. It is important that a suitable nonspecific binding control be included for all samples. This should at least consist of a section blocked according to the standard procedure described above and then incubated on the colloidal gold suspension, eliminating the primary antibody incubation step. The addition of a grid incubated with a purified IgG from the same species as that in which the specific primary antibody was raised and used at the same concentration as the primary antibody is also appropriate. Before any meaningful assessment of labeling is performed, these controls must be evaluated for labeling. In addition, the degree of labeling in the portions of the grid containing only support film and, if present, plastic without sample, provides an index of background labeling. While it is not absolutely necessary to use Formvar-coated grids in this procedure, they greatly aid the retention of sections on the grid, as the numerous wash steps in this procedure increase the chances of losing sections from uncoated grid surfaces.

LR White embedding resin was chosen for the above-referenced protocol due to its good antigen-preservation characteristics. Tissues embedded in epoxy resins may need to be etched to expose antigenic sites. Etching involves basic attack on the end-linked epoxide rings. Some proteins and peptides and many amino acids have been successfully deplasticized by etching methods to restore immunoreactivity lost in the embedding process.

Anticipated Results

Immunoelectron microscopy

Fixation techniques for TEM

At the conclusion of the fixation and initial processing of samples for thin-sectioning procedures, samples sufficiently stabilized to preserve ultrastructural organization are now ready for the step of embedding in plastic. This step is necessary to stabilize tissues in a matrix that is uniformly hardened to allow ultrathin sectioning with glass or diamond knives.

Embedding techniques for TEM

At the conclusion of the embedding process, tissues are infiltrated with a hardened matrix which is able to withstand the stress of ultrathin sectioning (i.e., ultramicrotomy) with glass or diamond knives.

Time Considerations

TEM of thin sections

Successful heavy metal staining of thin sections results in sections that are relatively easy to visualize on the phosphor screen of the electron microscope. The deposition of sufficient stain is required to record images, as identification of optimal focus in a TEM is another procedure that requires considerable care and practice. Figure 2B.1.3 and Figure 2B.1.4 show examples of samples suitably stained with uranyl acetate and lead. Figure 2B.1.8 provides an example of the use of immunogold localization of antigens followed by heavy metal staining. In this figure, differences between the electron scattering of 10-nm gold particles and heavy metal stains that generate contrast in the TEM can be appreciated.

Figure

Figure 2B.1.8 Propionibacterium fixed and initially processed for immunogold labeling and embedded in LR White resin. In panels (A) and (B) shown at the same magnification, the LR White-embedded sections were labeled first with a primary monoclonal antibody specific for this particular isolate of Propionibacterium; this was followed by secondary labeling with 10-nm protein G-gold (white arrows). Panel A shows inherent specimen contrast without staining uranyl acetate and lead staining of the section, whereas panel B shows the added contrast obtained by employing those stains. Note that while the ultrastructural detail of cells is improved with heavy metal staining, the visualization of gold particles can be more difficult and may require examination of sections at higher magnification.

Immunoelectron microscopy

These procedures require extensive manipulation of grids and several incubations, which will require several hours to complete. It is therefore desirable to process several girds simultaneously, not only to speed up the process of evaluating samples, but to ensure that a sufficient number of samples are observed along with appropriate controls. It is also possible to run several dilutions of antibody and gold conjugate simultaneously to expedite the procedure.

Fixation techniques for TEM

Preservation of cells for examination by thin sections is an extensive protocol that requires multiple steps to accomplish. If a sample is presented early in the work day, it is possible to have the infiltrated sample polymerizing in the oven by the end of the day. However, it is more likely that these procedures will be performed over several days. Fortunately, there are several points at which the processing can be arrested until such time that it can be resumed without doing damage to the sample, all of which involve storage at 4°C. Some potential points at which it is possible to halt the procedure include the primary fixative stage (if the tissue is being fixed overnight at 4°C), after the first buffer rinse following primary fixation, after the buffer rinse following post-fixation, or when the sample is in the 100% resin mixture.

Embedding techniques for TEM

In the embedding process, it typically takes several hours to replace the resin/solvent mixture with unpolymerized resin, which is then usually hardened in an oven overnight. On the following day, blocks are removed from the oven and either trimmed in preparation for ultramicrotomy or stored in appropriate containers until samples are needed for preparation of thin sections. Embedded tissue will last many years when kept at normal room temperature and humidity.

TEM of thin sections

Staining with uranyl acetate and lead usually takes about 30 min. Immunogold staining of antigens on thin sections typically takes 4 to 6 hr to complete.

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