Speciment Preparation

Fixation

There are three main ways in which cells and tissues may be processed to retain their structural organization for subsequent staining. These are fixation by cross-linking, fixation by precipitation, and fixation by freezing (cryofixation). The last method, cryofixation, is probably the best technique for cellular preservation, and is often employed for electron microscopy for this reason. It involves rapidly freezing the cells or tissues on a cooled block of heat-conductive metal or rapid plunging into a cold medium, such as liquid nitrogen or freon. Following freezing, the samples may then be treated with a cross-linking reagent, discussed below, in a process called 'freeze substitution'. The disadvantages of cryofixation are that it typically requires specialized equipment usually unavailable in most laboratories, as well as experience. I do not have experience with these techniques, and will not discuss them further.

The selection of a specific fixation protocol will be dictated by several factors. Firstly, the fluorescent probe to be used may place restrictions on which treatment may be used (i.e. some fixations prevent binding of certain dyes). Secondly, the size or thickness of a given sample may preclude the use of certain fixatives due to permeability (i.e. a fixative that is unable to penetrate into thick samples will only preserve the outer layers).

Cross-linking.

Fixation by cross-linking is a method commonly used for fluorescence microscopy. It involves treating specimens with reagents that penetrate into the cells and tissues and form covalent cross-links between intracellular components. The most commonly used cross-linking agents are aldehydes, which form covalent bonds between adjacent amine-containing groups through a Schiff acid-base reaction. These bonds form both inter- and intra-molecularly and are, therefore, very effective fixatives for proteins and nucleic acids. The two most frequently used aldehydes are formaldehyde and glutaraldehyde. Both fixatives have advantages and disadvantages which will be discussed below. Other aldehydes, such as acrolein, have been used historically, but do not preserve samples as well as the other two I have listed and will not be discussed here. Aldehydes are suspected carcinogens, they should be used only in well-ventilated areas or fume hoods and contact with skin or eyes should be carefully guarded against.

Glutaraldehyde.

Glutaraldehyde is a four carbon molecule terminated at both ends by aldehyde groups. It is an extremely efficient fixative, and is widely used by light and electron microscopists for its efficacy in preserving cellular structure. Glutaraldehyde is commercially available as an aqueous stock from many vendors, and may be purchased in stock concentrations ranging from 2 to 50%. Electron microscopy suppliers often offer glutaraldehyde in glass ampules sealed under nitrogen. This is because upon exposure to oxygen, glutaraldehyde will polymerize with time, diminishing its effectiveness. Hard-core electron microscopists will typically use one vial per experiment, and discard any unused fixative. I have used glutaraldehyde in vials and in large stock bottles which we have stored on a shelf for a year or so without any observable difference in its potency, at least at the light microscope level. Use of glutaraldehyde does have certain disadvantages, however. Firstly, its comparatively high molecular weight limits its ability to diffuse into thick specimens, such as tissue sections or embryos. This is further exacerbated by the fact that as the tissue is cross-linked by the fix, its ability to penetrate over time diminishes. For such samples, formaldehyde may be a better option. Secondly, unreacted aldehyde groups fluoresce efficiently at the same wavelengths as many of the fluorescent probes employed by biologists. As glutaraldehyde possesses two functional groups per molecule, background autofluorescence may be a significant problem in fixed tissues, effectively lowering the probe's signal to noise. This problem may be circumvented by using relatively low concentrations of glutaraldehyde (i.e. less than 1%). Unreacted aldehydes may also be quenched by treating fixed samples with reducing agents, such as sodium borohydride, to reduce free aldehyde groups to alcohols, or by reacting them with exogenous amine-containing reagents, such as ammonium chloride or glycine. Thirdly, the 4 carbon chain of glutaraldehyde may mask amine-containing epitopes, making immunostaining impossible. Although I have never used an antibody which did not stain glutaraldehyde-fixed cells, this should be taken into consideration. Despite these problems, it has been my experience that glutaraldehyde is an excellent fixative, and the drawbacks to its use may be circumvented effectively with a little extra effort.

Formaldehyde.

Formaldehyde is probably the most commonly used cross-linking fixative for biological samples. It has a single aldehyde-containing carbon and exists as a gas. Formaldehyde does not cross-link as effectively as glutaraldehyde, and for this reason is rarely used by-itself for electron microscopy. However, its small molecular weight allows it to penetrate cells and tissues rapidly, making it a choice fixative for thicker samples and autofluorescence of unreacted aldehyde groups is not usually a problem. Formaldehyde is commercially available in solution, termed 'formalin', which is typically provided as 10% or 37% stocks. These stock solutions are convenient as preparation of the fixative simply entails diluting them into buffer to the desired concentration. However, in order to stabilize the solubility of formaldehyde, manufacturers will usually include additives such as methanol to formalin, which may be undesirable (see fixation by precipitation, below). It is preferable, therefore, to freshly prepare formaldehyde in the lab just prior to its use from its polymeric form, paraformaldehyde (see below). Unlike glutaraldehyde, formaldehyde has a relatively rapid reverse rate, on the order of weeks, so specimens fixed in this manner should not be stored for extended periods of time prior to staining and analysis. Formaldehyde is typically used at final concentrations of between 1 to 4%. Unlike glutaraldehyde, formaldehyde exhibits a pH-dependence in its ability to cross-link amine groups, the reaction occurring 100-fold more rapidly with protonated amines. Greater cross linking may, therefore, be induced by fixing samples in buffers of high pH. The greater extent of cross-linking under basic conditions can inhibit permeability of the fixative throughout specimens, however. This problem has been solved by the development of a two-phase fixation, where samples are first treated with formaldehyde at a pH of 6.5 for a short time to infuse them with fixative, followed by a shift to basic pH to induce rapid cross-linking¹. An adaptation of this protocol is given later.

Precipitation.

Another often-used fixation method entails the simultaneous denaturation and precipitation of biological samples. This is typically accomplished by immersion in cooled organic solvents, such as methanol or acetone or acids. Acidic precipitation does not preserve cellular structures well, and is rarely used except for specific protocols, such as mitotic chromosome spreads. Fixation by precipitation does not preserve the three-dimensional organization of specimens, and is therefore not recommended for confocal microscopy. Quantitative comparisons of different fixation protocols have shown that cultured cells fixed with cold methanol shrink by as much as 50%². With those caveats, precipitation does have several advantages over cross-linking protocols. The first is speed - this type of fixation is relatively rapid, usually taking a few minutes. Secondly, precipitation rarely destroys antibody binding sites, as epitopes are not covalently modified as they might be with aldehyde fixation. Thirdly, fixation with organic solvents simultaneously permeabilizes cellular membranes, circumventing the necessity for detergent-treatment. And fourth, precipitation will not introduce autofluorescence into a specimen (as glutaraldehyde sometimes does). For conventional, wide-field fluorescence microscopy, the solvent-induced shrinkage may actually be beneficial as thicker samples may more readily fit within the depth of field of high mag, high numerical aperture objective lenses.

Buffer selection.

One other consideration when designing a fixation protocol is the choice of buffer in which the fixative will be dissolved. Ideally, it should be isotonic with the cell so as not to perturb cellular structure, and it should maintain the cell is as close to its physiological state as possible. The most frequently used buffer is probably phosphate-buffered saline (PBS) as it is inexpensive and approximates the physiological, extracellular environment fairly well. When using aldehydes as fixatives, amine-containing buffers such as Tris should never be used, as they will react with the fixative. In certain cases, specific buffers may be chosen for their efficacy at preserving specific cellular structures (i.e. microtubules, see below).

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