It is essential to be familiar with the operation of the cryostat. It is highly recommended to read the manual, which contains a great deal of information of practical value, and will save many hours of frustration. Thin cryosections (about 4 microns) give the best morphology. Choice of section thickness depends on the sensitivity required. Thicker sections have more antigen, so they may give stronger signals but worse resolution.
(1) The knife must be sharp and with no nicks or other damage. Nicks will cause tears in the sections. A blunt knife will cause multiple small tears parallel to the knife edge and/or alternate thick and thin sections.
(2) The knife and tissue block holder must be firmly screwed in place. This seemingly trivial point is not an uncommon source of problems.
(3) The knife must be at the correct angle (consult the Cryostat manual). If the angle is too steep, multiple small tears parallel to the knife edge may develop. If the angle is too shallow, it may result in the blade leaving the block surface before the whole of the block is sectioned.
(4) Operation should be smooth—a consistent cutting speed will provide consistent thickness of serial sections.
(5) It is essential that the cryostat mechanism be functioning well. Lubrication is important and takes only seconds to perform. The cryostat mechanism should be kept ice-free because the presence of ice may cause irregular advancement of the block.
(6) Keep the glass door of the cryostat closed when not in use to minimize condensation and dust.
(7) Do not allow the cryostat and tissue to warm up excessively. Individual tissues cut best at different temperatures; consult the cryostat manual. Warming of the block surface can cause curling of sections.
(8) Frost and/or tissue debris on the knife and Perspex anti-roll shield may cause sticking and folding of sections. Wipe the knife clean with a thick wad of paper tissues, away from blade edge. The use of a brush will almost inevitably result in damage to the blade edge. The blade edge will also be damaged by trimming the tissue block while it is mounted adjacent to the knife. It is a good idea to wear a face-mask while cutting sections, to avoid breath causing frosting of the knife and Perspex shield.
(9) Curling of sections is usually due to warming of either the tissue block surface or the Perspex shield. Close the door of the cryostat and wait for the temperature to equilibrate.
(10) Transfer the sections to a microscope slide (preferably gelatinized to facilitate adhesion) kept in a slide rack at room temperature.
(11) If the block of tissue is uneven, a sloping side can cause twisting or tilting of one side of the section. This can be avoided by embedding in OCT and trimming the excess OCT to give square sides.
(12) Trimming the upper edge of the tissue block to have a pointed shape (vertex upwards) may make it easier to detach the sections for manoeuvring into position onto the glass slide.
(13) Grit and other hard debris in blocks will result in tearing of sections and may cause nicks in the knife.
(14) Check the quality every 10-12 sections using the rapid 'Paragon' multi-stain. The macroscopic appearance of the cut, unstained section on the slide can give much information as to the quality of the section. Comparing it with a satisfactory Paragon-stained section is useful, looking at the outline as well as for easily recognized shapes within the section.
(15) The Perspex antiroll shield must be parallel to and slightly ahead of knife edge. Begin cutting with the Perspex just behind knife edge, then advance it forward until perfect sections are cut. Care must be taken to adjust the Perspex shield accurately over the knife, otherwise the section may roll over it or fold up on itself under it.
(16) Electrostatic sticking of sections to the Perspex shield may occur if there is low humidity in the air. The remedy may be to discharge the static by 'earthing'.
Cutting cryostat sections of fixed material such as PLP-fixed blocks requires more skill and perseverance than for straight unfixed, snap-frozen tissues.
It is a good idea to coat the glass slides with gelatin prior to cutting the sections. Gelatin helps the section to stick firmly on the slide, and stops them lifting off or rolling back on themselves. The coating is done by dipping the slides in a solution consisting of 2% gelatin in water for 30 seconds at room temperature. Gelatin solution is kept at 4°C and is solid at this temperature. Liquify by heating at 37°C for about 30 min. After dipping, the gelatin slides may be fixed in 10% formalin in water for 30 s (this is optional; residual formalin may decrease antigenicity of the specimen). Coated slides are dried overnight in a staining rack at room temperature to allow adequate ventilation, and stored in a box at room temperature.
If the sections detach from the gelatin-coated slides during washing, trials of other slide adhesives such as poly-L-lysine, starch, or proprietary glue preparations may be required. Gelatin coating should not be used if sections are to be protease-digested, the sections will lift off almost immediately; adherence of sections to the slide in this case may be achieved by drying at 37°C or use of a glue solution.
Opinions vary about how to dry the sections. If the sections are to be dried, this should probably be done before fixing and staining. They may be dried at room temperature for at least 1 hour or overnight, but it is probably best to fix and stain as soon as possible after cutting, as time-dependent loss of antigenicity has sometimes been observed. Alternatively, the slides may be air-dried in the cold room for 30 min to 2 h. If slides are dried in the cold room, they should be placed in a position where the air circulates freely, to avoid the accumulation of condensation.
After sections are prepared, the key factors that determine the extent and rate of deterioration in antigenicity are moisture and ambient temperature. Once dried, sections that are placed in slide boxes with dessicant and stored at -70°C are stable for years.
13.5.5 Should Frozen Sections be Fixed Before Antibody Staining?
This is a vexed question. In some cases, fixation is essential to prevent the relevant antigen from washing out during staining. However, as discussed at the beginning of this chapter, all fixation procedures, by their very nature, will cause some damage to antigenicity. In most cases, the fixation procedure will have to be determined empirically for each antigen and monoclonal antibody.
If the sections are to be fixed prior to staining, this should be done as soon as possible after the section is dry. Store the fixed section sealed in a plastic bag containing dessicant (silica gel) at 4°C or -20°C.
The most popular fixation procedure is probably a brief dip in acetone, usually pre-cooled in the -20°C freezer in a Coplin jar. Time may be 1-30 s, followed by aqueous wash in isotonic saline or drying in air for 5-30 m at room temperature. Fixation with organic solvents such as acetone will remove most of the membrane lipids and precipitate membrane proteins in situ. Fixation with organic solvents such as acetone also permeabilises cells, allowing antibodies access to the cytoplasm.
A brief dip in acetone is compatible with retention of many antigenic determinants. There is a greater chance of loss of antigenicity using ethanol, and even greater chance if the sections are fixed in methanol. PLP-fixed cryostat sections often give better morphology than fixation in organic solvents (see Hancock and Atkins, 1986).
There are many choices of strategy for immunological staining of tissue sections. In recent years, there has been a marked trend towards the use of enzyme-labelled antibodies rather than immunofluorescence, because antigens can be localized at the same time as viewing the architecture of the tissues, and a permanent record is obtained. However, the recent development of highly fluorescent substrates for alkaline phosphatase (see Sections 12.10 and 13.8.6) has provided a powerful new option.
Multilayer systems give greatly increased sensitivity due to the fact that each first antibody can bind several second antibodies. If a third antibody is added, the sensitivity increases still further, but at the risk of more 'background' staining and experimental artefacts (see Hancock and Atkins, 1986 for discussion).
13.6.1 Simultaneous Detection of Two or More Antigens
If two or more antigens are to be detected simultaneously, careful consideration will need to be given to the choice of enzymes and substrates, or to the use of fluorochromes that have easily separable emission spectra. The possibility of a combination of conventional substrates and the new fluorescent substrates may be particularly useful. Great care must be taken to ensure that the antibody systems used in multicolour work do not interact in unplanned ways. Many of these considerations have been discussed in Section 12.7.1 and other parts of Chapter 12 (see also Mason and Woolston, 1982).
The biotin-avidin system is particularly helpful in the staining for several different antigens because it allows one antigen to be detected with anti-immunoglobulin and a second to be detected with labelled avidin or streptavidin (see Sections 10.3 and 12.9). The tetrameric structure of avidin and streptavidin allows the use of a 'bridging' technique in which the tissue is stained with biotinylated antibody, washed, incubated with avidin or streptavidin, washed again, and incubated with a biotinylated enzyme or other biotinylated reagent which can bind to unoccupied biotin-binding sites.
As discussed elsewhere (see Section 12.9.3), avidin is very basic and may bind nonspecifically to tissues. It binds particularly strongly to DNA and other acidic structures. This problem may be particularly severe in immunohistology, but may sometimes be overcome by raising the salt concentration to 0.3-0.5 M and/or adding cytochrome c as a competitor. Alternatively, one may use a modified form of avidin that has been chemically treated to raise its isoelectric point (e.g. Neutravidin from Pierce Chemicals). The use of streptavidin overcomes many of these problems, but can create new problems because it can bind to certain receptors which have specificity for the tripeptide Arg-GIy-ASP RGD and related motifs (Section 12.9.3).
If the tissue sections are to be incubated with hybridoma supernatants, it must be remembered that most tissue culture medium contains biotin, which could inhibit binding to avidin if it is not removed by washing. It should also be noted that some proteins in cells may contain attached biotin, and this could cause nonspecific staining with avidin or streptavidin.
412 Monoclonal Antibodies: Principles and Practice
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