22 Transmission electron microscopy
Tissue preparation for transmission electron microscopy
The basic preparation methods for routine TEM are provided in this chapter. More detailed discussions of these, plus alternative and specialized procedures, can be found elsewhere (Glauert 1972; Robards & Wilson 1993; Allen & Lawrence 1994; Glauert & Lewis 1998; Hayat 2000). A flow chart summarizing the steps required for preparing the basic range of diagnostic TEM specimens is given in Figure 22.1.
Fixation
Buffers
Fixatives are normally buffered within the range of pH 7.2–7.6 (Robinson & Gray 1996). Ideally the osmolarity and ionic composition of the buffer should mimic that of the tissue being fixed. In general practice this is not a major requirement but, if required, 300–330 mOsm (the osmolarity equivalent to that of plasma or slightly hypertonic) is suitable for most circumstances. Non-ionic molecules such as glucose, sucrose or dextran are used to adjust tonicity as these will not influence the ionic constitution of the buffer. The addition of various salts, particularly calcium and magnesium, is thought to improve tissue preservation, possibly by stabilizing membranes (Hayat 1981). This is unlikely to have a major effect in routine diagnostic applications.
Phosphate buffers
Alternative buffers
Other buffers that have been recommended for use in TEM include cacodylate (Plumel 1948; Sabatini et al. 1963), HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), MOPS (3-(N-morpholino) propanesulfonic acid) and PIPES (piperazine-N,N′– bis2-ethanesulfonic acid) (Good et al. 1966; Good & Izawa 1972; Massie et al. 1972; Salema & Brandão 1973; Ferguson et al. 1980).
Aldehyde fixatives
Aldehyde combinations
Paraformaldehyde (2%) and glutaraldehyde (2.5%) fixative (buffered) (based on Glauert 1972; Karnovsky 1965)
Stock reagents
0.2 M buffer, pH 7.4 (phosphate, cacodylate) | 50 ml |
Paraformaldehyde | 2.0 g |
25% aqueous glutaraldehyde | 10 ml |
Distilled/deionized water to | 100 ml |
Method
1. Completely dissolve paraformaldehyde in buffer using heat and with continuous stirring. It may be necessary to add a few drops of 1.0 M sodium hydroxide to clarify the solution.
2. Cool the solution rapidly under running water.
3. Add aqueous glutaraldehyde. Check the pH of the mixture and adjust if necessary to pH 7.4.
Osmium tetroxide
The use of osmium tetroxide fixation to preserve lipids is fundamental to electron microscopy (Palade 1952; Millonig & Marinozzi 1968). While primary fixation in osmium tetroxide is effective, its extremely slow penetration rate can give rise to autolytic changes. For this reason osmium tetroxide is almost always used as a secondary fixative (termed ‘post-fixation’) after primary fixation in aldehyde. The penetration rate of osmium tetroxide is also higher in stabilized tissue, such that immersion for 60–90 minutes is usually sufficient for most specimens.
Osmium tetroxide fixative (2% aqueous)
Stock reagent
Osmium tetroxide | 1.0 g |
Distilled/deionized water | 50 ml |
Method
1. Clean, then score the glass ampoule with a diamond pencil and place in a dark-glass storage bottle.
2. Break the ampoule with a glass rod and add water. It may take 24 hours or longer for the osmium tetroxide to dissolve completely.
3. Prepared solutions may be stored for short periods at room temperature in the dark in a well-sealed bottle (double wrap the bottle in aluminum foil); for long periods store at 4°C. All osmium solutions should be stored inside a second closed container to prevent the leakage of osmium fumes. Aqueous osmium solutions that are prepared and stored in a clean container should last for ~1 year; solutions in buffer may only last a few days before they deteriorate.
4. For a 1% working solution combine 1:1 with water or buffer.
Epoxy resins
Examples of widely used epoxy resin composites include Araldite (Glauert & Glauert 1958), Epon (Luft 1961) and Spurr’s resin (Spurr 1969). Although the original product names Araldite and Epon refer to epoxy resins developed by the CIBA Chemical Company and Shell Chemical Company, respectively, these terms are now in general use. Araldite polymers are preferred as these react with a higher degree of cross-linking and are the most stable.
Occupational exposure to epoxy resins is a common cause of allergic contact dermatitis (Kanerva et al. 1989; Jolanki et al. 1990). These agents are also probable carcinogens, primary irritants and systemically toxic (Causton 1981). Spurr’s resin in particular is toxic and should be handled with great care (Ringo et al. 1982).
Acrylic resins
The main commercial acrylic resins are LR White and LR Gold and the Lowicryl series (K4M, K11M, HM20 and HM23). Each of these can be used for low-temperature dehydration and embedding to reduce the heat damage from exothermic polymerization and extraction by solvents and resin components (Acetarin et al. 1986; Newman and Hobot 1987, 1993). These characteristics make several forms of acrylic resin ideally suited to electron immunogold labeling (Stirling 1994) and enzyme cytochemical studies.
Tissue processing schedules
Table 22.1 Standard processing schedule for solid tissue cut into 1 mm3 blocks (each step is performed at room temperature unless stated otherwise)
Primary fixation | 2.5% glutaraldehyde in 0.1 M phosphate buffer | 2–24 hours (room temperature or 4°C) |
Wash | 0.1 M phosphate buffer | 2 × 10 minutes on rotator |
Wash | Distilled water | 2 × 10 minutes |
En bloc staining (optional) | 2% aqueous uranyl acetate | 20 minutes |
Dehydration | 70% ethanol | 10 minutes on rotator* |
90% ethanol | 10 minutes on rotator | |
95% ethanol | 10 minutes on rotator | |
100% ethanol | 15 minutes on rotator | |
Dry absolute ethanol | 2 × 20 minutes on rotator | |
Transition solvent (clearing) | 1,2-epoxypropane | 2 × 15 minutes on rotator |
Infiltration | 50 : 50, clearant : resin# | 1 hour |
25 : 75, clearant : resin | 1 hour | |
Resin only | 1–24 hours (with vacuum to remove bubbles) | |
Embedding | Fresh resin in embedding capsules | 12–24 hours at 60–70°C |
* Tissues may be stored at this stage.
# As batches may vary, resin should be prepared in accordance with manufacturer’s instructions.
Procedures for other tissue samples
Cell suspensions or particulate matter
Cell suspensions (such as fine needle biopsy aspirates, bone marrow specimens or cytology samples) or particulate materials (including fluid aspirates, tissue fragments or products and specimens for the assessment of ciliary structures) are best embedded in a protein support medium before processing. Plasma, agar or bovine serum albumen (BSA) can be used. The addition of tannic acid (Hayat 1993) during the preparation of ciliary specimens gives improved visualization of axonemal components (Sturgess & Turner 1984; Glauert & Lewis 1998). The tannic acid is thought to act as a fixative and also a mordant, facilitating the binding of heavy metal stains (Hayat 2000). Double en bloc staining with uranyl acetate and lead aspartate may also improve the visibility of dynein arms (Rippstein et al. 1987).
Preparing particulate specimens
Method
1. Centrifuge the material in buffer in a plastic centrifuge tube to form a loose pellet.
2. Discard supernatant and resuspend the material in glutaraldehyde fixative at room temperature for a minimum of 1 hour.
3. Centrifuge the material and carefully discard the supernatant.
4. Wash the specimen by resuspending it in buffer for 10–15 minutes.
5. Centrifuge the material to form a loose pellet.
6. Discard supernatant and introduce 0.5 ml of 15% aqueous BSA. Resuspend the specimen and allow it to infiltrate for a minimum of 1 hour.
7. Centrifuge the material and discard most of the supernatant, leaving sufficient to cover the pellet to a depth of approximately 1 mm.
8. Introduce an equal volume of glutaraldehyde fixative to form a layer above the BSA. Allow material to solidify for 2–24 hours.
9. Remove the material (this is most easily achieved by cutting away the plastic centrifuge tube) and divide into small portions.
10. Wash in four changes of buffer, each for 5 minutes (for ciliary biopsies only, incubate for 15 minutes in buffered tannic acid solution, then wash in four changes of buffer, each for 5 minutes, before proceeding to step 11).
11. Postfix in 1% aqueous osmium tetroxide and process as normal.
Material embedded in paraffin/cell smears
Reprocessing paraffin-embedded material
Method
1. Remove the area of interest from the block, taking care not to damage the tissue.
2. Dewax the specimen by passing through several changes of xylene. The time required depends on the size of the sample but should be at least 1 hour. A minimum of three changes is recommended.
3. Rehydrate the material in a graded ethanol series.
4. Wash in water, postfix in osmium tetroxide and process as routine specimen (see above).
Pop-off technique for slide-mounted sections (after Bretschneider et al.1981)
If additional fixation is required
2. Rehydrate the tissue in a graded ethanol series.
3. Wash in buffer and fix the tissue in glutaraldehyde fixative for 15–20 minutes.
4. Wash in buffer and postfix in 1% osmium tetroxide for 20–30 minutes.
5. Wash in buffer or distilled water, and then cover with 2% uranyl acetate for 15 minutes.
6. Dehydrate the tissue by passing the slide through 70%, 90%, 95%, 100% and super dry ethanol for 5 minutes in each stage.
7. Dip the slide into propylene oxide for 5 minutes. The tissue should not be allowed to dry.
8. Cover the tissue with a 2 : 1 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.
9. Cover the tissue with a 1 : 2 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.
10. Cover the tissue with neat epoxy resin for 5–15 minutes.
11. Drain off surplus resin mixture. Invert a freshly filled (to overflowing) embedding capsule over the section and press onto the slide.
12. Incubate the slide and capsule at 60°C for 24 hours for polymerization to occur.
13. Remove the slide and, while still warm, separate the capsule and the newly embedded tissue from the glass slide.
If additional fixation is not required
2. Dip the slide in equal parts of propylene oxide and xylene, then into propylene oxide for 5–10 minutes. The tissue should not be allowed to dry.
3. Cover the tissue with a 2 : 1 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.
4. Cover the tissue with a 1 : 2 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.
5. Cover the tissue with neat epoxy resin for 5–15 minutes.
6. Drain off surplus resin mixture. Invert a freshly filled (to overflowing) embedding capsule over the section and press onto the slide.
7. Incubate the slide and capsule at 60°C for 24 hours for polymerization to occur.
8. Remove the slide and, while still warm, separate the capsule and the newly embedded tissue from the glass slide.