Application of immunohistochemical techniques
to the study of the nervous system

Immunohistochemical visualization of the nerve cells and their fibers is based on selective labeling of the chemical substances specifically present in these cells or fibers using antibodies targeted against these chemical substances. One of the substances characteristic of the nervous system is tubulin, which is present in most fibers and often in the cytoplasm of neurons of both vertebrate and invertebrate animals. Tubulin forms the microskeleton of the neuronal microtubules and also is a component of the ciliary apparatus of epithelial and receptor cells. The nerve cells are also characterized by the presence of regulatory substances functioning as neurotransmitters, hormones, and neuromodulators; these substances are secreted by the neurons and can be present in their cell bodies and fibers. The neuroactive substances include acetylcholine, some amino acids (glutamic acid, gamma-aminobutyric acid, etc.), monoamines (serotonin and catecholamines such as dopamine, adrenalin, noradrenaline, etc.), enkephalins and a great variety of as yet little explored neuropeptides (neurotensin, substance Р, FMRFamide, etc.). The antibodies that have now been produced against some of the neuroactive substances allow neuroscientists to localize these substances in the neurons and study the arrangement of these cells in various regions of the nervous system. The primary antibodies are labeled by the secondary antibodies linked to fluorescent or non-fluorescent probes.

The advantages of all immunohistochemical techniques are:

1. high specificity towards targeted chemical substances and the ability to determine their precise localization;
2. the ability to use fluorescent stains in conjunction with confocal and multiphoton microscopy to make precise three-dimensional reconstructions of the architecture of the nerve nets containing specific neuroactive substances, with an additional benefit of studying the nervous system as a whole on the whole-mount preparations of small-sized animals;
3. the ability to combine immunohistochemistry of the nervous elements with visualization of musculature using phalloidin, which allows for studying spatial relationships between the nervous elements and muscles.

The disadvantages (limitations) of immunohistochemical techniques are:

1. the inability to reveal the general cytoarchitecture of the nervous system and its regions (including localization of cell bodies);
2. the inability to reveal the shapes of the cells and their fibers and relations between the neurons containing different neuroactive substances (except with 2–3 substances simultaneously), because the immunohistochemical staining detects only the sites where the targeted substances are localized inside the neurons, but the plasma membranes are not visualized and the shapes of the cells cannot be determined.

Immunohistochemistry is a set of techniques for studying tissues based on the ability of the immune system of vertebrates (primarily, mammals and birds) to produce specific proteins (antibodies) that can selectively bind to certain substances (antigens). Inside the body of a vertebrate, the antibodies are used primarily to identify and neutralize pathogens. The immune system functions in such a way that theoretically the antibodies can be produced against almost any chemical substance. The larger the molecule of an antigen, the faster and better is the production of antibodies. These unique properties of the immune system are employed in immunohistochemistry: the antibodies are raised against the substance of interest, then isolated from the serum and used to detect this substance in the cells and tissues of different organisms.

The immunohistochemical techniques are not new. The first attempts to use these techniques were made in the 1930s, when scientists have successfully produced the dye-labeled antibodies that were able to react with certain antigens and stain them. The antibodies conjugated with a fluorescent probe have been produced for the first time at the beginning of the 1940s. These first studies were conducted using conventional fluorescent microscopes. The advent of the confocal microscope has greatly expanded the utility and the scope of these techniques giving them the ability to analyze the three-dimensional structure of specimens.

The antibodies are glycoproteins that belong to the group of immunoglobulins. The molecule of an antibody is composed of two identical heavy and two identical light polypeptide chains. Consequently, each antibody molecule has two identical antigen-binding regions that can recognize the target antigen and bind to it.

The antibodies are secreted by B cells; each line of В cell produces only one type of antibodies that can recognize one specific epitope. The antibodies thus are classified into monoclonal (those that are produced by a single cell line and recognize only one epitope) and polyclonal (those that are secreted by cells of different lines and probably recognize different epitopes of presumably the same antigen). Monoclonal antibodies are more specific than polyclonal antibodies, but their production is more expensive, and for this reason the polyclonal antibodies are more often used for research purposes.

The antibodies are themselves colorless and unable to fluoresce. There are several different visualization methods in immunohistochemistry based on linking a fluorescent or chromogenic probe to an antibody molecule. The probe can be linked directly to the antibodies produced against the target antigen, but there is also another technique, the so-called indirect immunohistochemical method. This method is more sensitive and versatile. It involves two different, primary and secondary, antibodies. The primary antibodies are produced against a certain target antigen. These antibodies are pure, not linked to any molecule. By contrast, the secondary antibodies are raised against proteins of the organism in which the primary antibodies have been produced. These antibodies are conjugated to the probe molecules. Over the course of the reaction, the secondary antibodies find the primary antibodies bound to the antigen and attach to them. For this method to work, the specimen must first be incubated with the primary antibodies and then with the secondary antibodies. This technique is not only more sensitive, but also allows for different markers to be used with the same primary antibodies in different situations. The secondary antibodies linked to the same type of probe can be used with different primary antibodies. One should only make sure that the primary antibodies be raised in the same animal species, against which the secondary antibodies have been produced.

The same specimen can be labeled with antibodies raised against several different antigens. Their number is theoretically limited only by spectral characteristics of the probes and by the number of host animals against which the secondary antibodies have been raised. It should be kept in mind that if the primary antibodies against different substances were produced in the animals of the same species (for instance, in rabbits), they cannot be used together in the same experiment. The reason for this is that the same secondary antibodies will bind to all primary antibodies making it impossible to know, which substances were eventually identified. Likewise, different secondary antibodies with fluorescent labels having similar or identical absorption/emission spectra should not be used together in the same experiment, because fluorescent signals will be difficult or impossible to differentiate.

Fluorescent-antibody staining works well with the majority of the common fluorescent methods, such as staining of filamentous actin with phalloidin or staining of nuclei with DAPI. Furthermore, fluorescent immunohistochemistry can be combined with chromogenic methods of detection used in in situ RNA-hybridization.

Since antibodies are relatively large molecules (the molecular weight of IgG is about 150 kDa), their delivery into the cell (or even into the body) may pose a challenge. Two approaches have been devised to resolve this problem. The first approach involves a slight damage to the plasma membranes of the specimen using detergents. The most widely used detergent is Triton X-100 at a concentration of 0.05% to 5% (the most common concentration is 0.1%). In some cases, if the organism has a dense cuticle (e.g. nematodes or arthropods), the mechanical force is applied to damage the cuticle. The damage can be inflicted mechanically, for instance, by a scalpel, or by a brief application of ultrasound. Also, in some cases, the specimen is pre-incubated with enzymes such as protease, chitinase or collagenase.

An alternative set of approaches is to reduce the size of antibody molecules. The heavy chains include the so-called hinged region; the antibody can be cut across this region by the enzyme papain into three fragments: two Fab-fragments (fragment antigen-binding) and one Fc-fragment (fragment crystallizable). The Fab-fragment can penetrate the cells faster and more efficiently than the uncut molecules. This approach is most often used in non-fluorescent detection, when the enzyme alkaline phosphatase is linked to the antibody fragment instead of a fluorophore molecule. This type of detection is employed primarily in in situ RNA-hybridization.

Immunohistochemical staining is a complex, multi-step process, but as long as the procedure is performed with reasonable care and accuracy, it will consistently yield reproducible and high-quality results. In some cases, antibodies can bind non-specifically to certain substances or the specimen can fluoresce all by itself. For this reason, the antibody-staining procedures should always be performed with a set of controls to detect possible non-specific staining. The most important is the negative control for both the primary and secondary antibodies, which is used to reveal potential autofluorescence and non-specific antigen binding. The quality of the reaction should also be checked with the negative control. This control can be performed using the immunohistochemical reaction with an organism, for which the antibody-binding pattern is already known. In some cases, the positive control can be replaced with a simultaneous application of several different primary and secondary antibodies. If the reaction is successful, one antibody will serve as a positive control for another and vice versa.

In immunohistochemistry, the quality of antibody binding is strongly dependent on the quality of fixation. The classical and most commonly used fixative is 4% formaldehyde solution in phosphate-buffered saline. The quality of the fixative is of the utmost importance for good staining. In water solutions, formaldehyde is gradually degraded into formic acid, methanol and acetone, which have a negative effect on fixation, and therefore the fixation must be as fresh as possible. The best option is to use the fixative immediately after it was prepared. It can be stored in the refrigerator for one week. Longer storage is possible, but not recommended. Sometimes, for a more prolonged storage, the fixative is prepared as 16% stock solution in water and stabilized with methanol (about 1 mL per 500 mL of fixative). This stock solution can be stored for several months.

Under normal conditions, formaldehyde is a gas. In laboratory practice, 4% formaldehyde solution is made from paraformaldehyde, which is a white powder produced by polymerization of formaldehyde. To prepare a good fixative, paraformaldehyde should be as chemically pure as possible. The best practice is to use paraformaldehyde specially designated for these purposes.

When heated, paraformaldehyde breaks down into monomers. Heat the mixture gradually, under constant stirring (preferably using a magnetic stirrer), while trying to avoid boiling. The good fixative should not contain any residue and should have a neutral рН. When the solution is being prepared, its pH goes down, which slows down the depolymerization reaction. If the fixative is prepared in buffer or in seawater, the buffer capacity of the system is usually enough to maintain the neutral pH and paraformaldehyde should dissolve completely, without leaving a residue. If the fixative is prepared in distilled water (especially if high-concentration solutions are used), 1N NaОН solution is usually added by drops to neutralize the fixative. Control the pH of the solution using an indicator paper to keep it close to the neutral pH. Cool the prepared fixative down and store in the refrigerator.

The anesthetized animals are fixed at the temperature, which is ambient for these animals, or in a cold fixative (cooled on ice). As with the histological fixation, it is recommended to use the volume of fixative, which is at least 10 times that of the specimen (for aquatic organisms the volume of the specimen must also include the volume of water around the organisms during fixation). Optimal are the results of fixation with the ratios of specimen-to-fixative volume of 1:20 to 1:40. The fixation time and temperature regime vary depending on the species, but the fixation time usually does not exceed 24 hours. The larger the specimen, the longer must be the period of fixation, and therefore the fixation of overly large specimens (over 1 cm in diameter) should be avoided. Small-sized specimens such as larvae of aquatic organisms with a length of up to 500–1000 µm can be fixed for 2 hours at room temperature or at +4°С. Larger organisms should be fixed for 4–6 hours at room temperature or for 8–12 hours at +4°С. An unreasonably long fixation may have a negative effect on staining.

Once fixation is over, the specimens should be rinsed in a buffer, usually in the same buffer that was used to prepare the fixative, or in 0.1 М phosphate-buffered saline (рН 7.2–7.4). The duration of rinsing may somewhat vary depending on sizes and properties of the specimens. It is recommended to wash the specimen at least in three changes of the rinsing solution, about 30 min each.

After the fixative is washed away, the specimen can be stored in the rinsing solution for a short period of time (in some cases for up to a week) at +4°С. If a longer storage is needed, add 0.01–0.1% of sodium azide to the rinsing solution. Some specimens, however, macerate quickly even if sodium azide is present. In this case, continue with the next steps of the procedure as soon as possible or transfer the specimens to 70% ethanol or absolute methanol and store at -20°С or lower.

Once the specimen is thoroughly washed and contains no fixative, it can be used for further staining. The first important step in immunohistochemical staining is preincubation in a solution of serum or bovine albumen. The purpose of preincubation is to block non-specific binding sites in the tissues of the specimen, which reduces non-specific binding of the primary and secondary antibodies. The best serum is the one that was obtained from an animal of the same species, in which the secondary antibodies had been produced. This serum would be most effective in blocking the potential non-specific activity and at the same time would not cross-react with the secondary antibodies. The usual concentration of the serum is 5–20% solution in buffer. An alternative to the serum is a buffered solution of bovine serum albumin (BSA) at a concentration of 0.25 to 2.5%. The specimens are recommended to be incubated in the refrigerator for 1.5 hours to 1 day. The stock serum should be aliquoted to smaller volumes and then frozen. It is recommended to prepare the bovine albumen solution immediately before preincubation or add 0.05–0.1% sodium azide to the prepared solution and store it in the refrigerator.

Good results can be achieved if another detergent, Tween 20, is used instead of Triton X-100. This detergent has a less detrimental effect on membranes and allows a more specific binding of antibodies.

Preincubation is followed by incubation with antibodies. For incubation, use the same buffer as for fixative rinsing. The dilution buffer sometimes contains bovine serum albumen or the serum used in preincubation. In my opinion, for most invertebrate animals the use of the serum during incubation with antibodies is unnecessary.

The optimal concentration of antibodies should be determined individually for each batch. For this, prepare a series of successive dilutions from 1:100 to 1:10000 (for instance, 1:100, 1:200, 1:500, 1:1000, 1:2000 1:5000, and 1:10000) and use the concentration that gave the best results. The manufacturer of the antibody may sometimes indicate the optimal dilution, but even in this case I recommend first to prepare a test series of dilutions, because in some cases the activity of the antibody may vary significantly from one batch to another.

The incubation times and temperature regime can also vary depending on the properties of the specimen under study. The incubation normally takes at least 16 hours (overnight) and is carried out at +4°С. When necessary, the incubation time can be extended for up to three days. As an alternative, the incubation for 16 hours at room temperature can also give good results.

After incubation, the unbound antibodies should be removed. Three successive rinses in a buffer, 15-20 minutes each, are usually enough. The rinses are followed by incubation with the secondary antibodies. The optimal dilution of the secondary antibodies is usually indicated by the manufacturer. As a rule, the incubation time for the secondary antibodies is the same as for the primary antibodies or slightly shorter. The secondary antibodies should be washed away in the same way as the primary antibodies.

After incubation with secondary antibodies, the specimens can be additionally stained with phalloidin or a nuclear stain (for instance, DAPI) and then mounted in glycerol, Mowiol or other mounting medium for fluorescent microscopy.

This protocol is suitable for most marine Lophotrochozoa.

1. Fix specimens with 4% solution of paraformaldehyde in 0.1 М phosphate-buffered saline (PBS) for 4–6 hours at room temperature or for 8-12 hours at +4°С. The specimens can be stored overnight in a refrigerator.
2. Rinse 3x30 min in PBS+0.1% Triton X-100 (PBT) at room temperature. It is recommended that the remaining fixative be removed as thoroughly as possible with the specimen exposed to the air.
3. Rinse in PBT.
4. Preincubate in PBT+1% BSA for 2 hours at +4°C. The specimens can be kept overnight in the refrigerator.
5. Incubate with primary antibodies diluted in PBT (1:1000) for one day/overnight at +4°С.
6. Rinse 3x20 min in PBS+0.1% Triton X-100 (PBT).
7. Incubate with secondary antibodies diluted in PBT for one day/overnight at +4°С.
8. Rinse 3х15 min in PBT at room temperature.
9. Stain the musculature with phalloidin and nuclei with DAPI (2 hours at +4°С).
10. Rinse in PBT.
11. Mount in Mowiol.