21 Molecular pathology
Molecular pathology seeks to apply gene expression against morphology and use gene expression analysis to validate large numbers of targets. A glossary of common definitions and terminology can be found at the end of this chapter.
Molecular pathology techniques have been used in the clinical laboratory to aid in the diagnosis and monitoring of treatment regimens of many infectious diseases such as HIV, hepatitis B, and tuberculosis (Netterwald 2006). These tests are usually performed on serological or other body fluids, such as sputum and seminal fluid. Currently the most well-known and most advertised molecular testing is for human papillomavirus (HPV) and human epidermal growth factor receptor 2 (HER2).
Clinical and research laboratories may use additional molecular pathology techniques, such as blotting methods which are used to study extracted ribonucleic acid (RNA) and deoxyribonucleic acid (DNA). Blotting methods consist of extracting DNA and/or RNA from homogenized tissues and then analyzing them using dot, Southern and Northern blotting filter hybridization methods (Sambrook et al. 1989). Blotting techniques such as these are powerful tools for the qualitative analysis of extracted nucleic acid from fresh or frozen cells and frozen tissues.
The polymerase chain reaction (PCR) is included in molecular pathology methods. PCR is a common method of creating copies of specific fragments of DNA. It rapidly amplifies a single DNA molecule into many billions of copies. In one application of the technology, small samples of DNA, such as those found in a strand of hair at a crime scene, can produce sufficient material to carry out forensic tests. PCR may also be used in addition to in situ hybridization (ISH) to study a specific genome of a tissue (Innis et al. 1990).
All of this leads to the role the histology laboratory plays in molecular pathology. In the histology laboratory the main method used in molecular pathology is ISH. John et al. (1969) and Gall and Pardue (1969) described the technique of ISH almost simultaneously.
ISH is a method of localizing and detecting specific mRNA sequences in preserved tissue sections or cell preparations by hybridizing the complementary strand of a nucleotide probe to the sequence of interest.
The method consists of denaturing (breaking apart) DNA and RNA strands using heat. A probe (a labeled complementary single strand) is incorporated with the DNA/RNA strands of interest. The strands will anneal with complementary nucleotides bonding back together with their homologous partners when cooled (Fig. 21.1). Some will anneal with the original complementary strands, but some will also anneal or hybridize with the probe. As probes increase in length, they become more specific. The chances of a probe finding a homologous sequence other than the target sequence decreases as the number of nucleotides in the probe increases. A longer probe can hybridize less specifically than shorter probes. Optimal probe size for ISH is small fragments of about 200–300 nucleotides. However, probes may be as small as 20–40 base pairs (bp) or as large as 1000 bp.
Figure 21.1 The genetic information for humans is encoded in billions of nucleotides (the building blocks of the DNA code) arranged in a double-helix molecule. Nucleotides consist of a base, a sugar (S), and a phosphate (P). The DNA code is written in an alphabet that uses four letters to represent each of the bases: (A) = adenine, (T) = thymine, (C) = cytosine, (G) = guanine. These bases will form pairs. (A) will only pair with (T). (G) will only pair with (C). Therefore, double-stranded DNA consists of two strands of homologous nucleotides. The genetic code in DNA is in triplets such as ATG. The base sequence of that triplet in the partner strand is therefore TAC.
The detection of specific nucleic acid sequences (RNA, viral DNA or chromosomal DNA) in cells, tissues or whole organisms by ISH has numerous applications in biology, clinical and anatomical pathology, as well as in research.
ISH methods may employ radiolabeled probes that are visualized on a photographic film or photographic emulsion. However, most of these probes do not work well on routinely fixed, processed tissues and require the use of frozen sections. They may need 20–50 days of exposure before the results are visible. The development of non-radiolabeled probes that perform well on routine surgical and autopsy specimens has extended the field of anatomic pathology.
Detection of mRNA by using ISH is particularly useful if the protein product is quickly degraded or rapidly transported out of the target cell.
In ISH detection, immunohistochemistry (IHC)-like methods may be incorporated to detect the labeled (biotin, digoxigenin (DIG)) probe. So, the question arises, why not just do IHC? After all IHC is well-established, reliable, and less time consuming than ISH. IHC has been employed in the clinical and research arenas for several decades and has become a routine procedure in the histology laboratory. Furthermore, IHC has provided diagnostic procedures and a close look at the proteins within and on the cell membranes. The advantages of ISH over IHC include:
• DNA and mRNA are not as sensitive to formalin fixatives.
• Probe-target hybrid is stronger than antibody-antigen complex.
• Provides an alternative means of detection when reliable antibodies are not available.
It is important to understand the ‘how and why’ of the different stages in the ISH process in order for the testing to result in a functional outcome. This revised chapter continues to focus on the ‘how and why’ of ISH, and includes a review of automated ISH versus the manual processes (Sterchi 2008). It also includes a revised comprehensive review of fluorescence in situ hybridization (FISH) staining procedures and analysis.
There are many modifications of ISH methods that relate to application needs. Although the demonstration of DNA and RNA sequences by ISH is a valuable research tool, according to Warford and Lauder (1991) and Mitchell et al. (1992), it is also used diagnostically in:
ISH comes in many forms and methods, and over the past 10 to 20 years the methodology has expanded significantly. At one time only FISH was the ‘standard’ method for ISH. Now there are methods that allow visualization of the stain using a bright field microscope, reducing the need for a fluorescence microscope. However, FISH still has an advantage over chromogenic methods for labeling specific nucleic acid sequences in cells and tissues. It is a ‘direct’ technique, so it is faster and in some cases it does not require IHC-like detection. These probes employ fluorescent (fluorescein) tags that glow under ultraviolet light to detect the hybridization. FISH allows the use of multiple probes on the same tissue that may spatially or spectrally overlap. The literature suggests that it is possible to distinguish at least four or five different fluorescent signals in a single sample (Haugland & Spence 2005), whereas chromagens are often limited to one or two color options per slide. FISH, also known as molecular cytogenetics, has enabled a huge advance in the diagnostic and prognostic capability of the clinical cytogenetic laboratory. FISH can also vividly paint chromosomes or portions of chromosomes with fluorescent molecules (thus, the term ‘chromosome painting’).
ISH can provide cytological information on the location and alteration of genomic sequences in chromosomes. Traditionally, the technique has been applied to metaphase chromosome spreads (Davis et al. 1984; Lux et al. 1990), but it has been shown to be applicable to interphase nuclei (Hopman et al. 1988; Poddighe et al. 1992). Routine paraffin wax preparations of tissues can be used and ‘interphase cytogenetics’, as the method is termed, can provide direct information on chromosomal abnormalities in unselected tumor cell populations.
Viral identification can be undertaken using a variety of methods, of which only immunohistochemistry and ISH provide simultaneous morphological information. The sensitivity of immunohistochemistry for the visualization of viral antigens, and ISH for the demonstration of cytomegalovirus, correlate well (Van den Berg et al. 1989). Most viral ISH methods use probes for DNA. Others, such as in the demonstration of the Epstein-Barr virus (Fig. 21.2), the detection of a virally encoded RNA transcript, provide results that are more sensitive than the use of antibodies and may even approach that of the PCR (Pringle et al. 1992).
Figure 21.2 Example of automatic chromogenic in situ hybridization (CISH) staining.
(a) Epstein-Barr virus-encoded RNA (EBER) and (b) cytomegalovirus (CMV).
(Photographs courtesy of Leica Microsystems, Inc.)
The light chain portion kappa and lambda mRNA may be detected in normal and neoplastic B cells in human lymphoid tissue. Restriction of either kappa or lambda mRNA denotes monoclonality of lymphoid neoplasms and is useful in distinguishing between neoplastic and reactive lymphoid proliferations. Due to the destruction of RNAases by formalin fixation, kappa and lambda sequences are conserved in routine surgical tissues. (See Data Sheet Kappa and Lambda Probe ISH Kit, Novacastra.) (Fig. 21.3.)
Figure 21.3 Example of automatic chromogenic in situ hybridization (CISH) staining (a) kappa (b) lambda.
(Photographs courtesy of Leica Microsystems, Inc.)
Chromogenic in situ hybridization (CISH) is a method ‘that enables the detection of gene expression in the nucleus using a conventional histochemical reaction’ (White 2005); it is used for the detection of abnormal genes and to identify a gene therapy treatment direction. CISH can be used as an alternative in screening archived breast cancer tissue samples for HER2/neu (type 1 growth factor receptor gene) (Madrid & Lo 2004). Automated CISH techniques were used for detecting light chain expression in paraffin sections on plasma cell dyscrasias and B-cell non-Hodgkin lymphomas ‘appeared superior to IHC’ in that the ISH resulted with no background staining (Beck et al. 2003).
Silver precipitation in situ hybridization (SISH) is an emerging ISH method that works well with formalin fixed paraffin embedded (FFPE) tissues. It is also similar to FISH performance in detecting the location of genomic targets using probes. The major advantage of CISH and SISH is the possibility of long term storage of the stained slides. The chromagens or silver signals do not quench over time, unlike FISH signals (Fig. 21.4).
Figure 21.4 Example of automatic silver (silver deposition technology) in situ hybridization (SISH) staining.
(a) HER2 and Chr17. (b) HER2 and Chr17.
(Photographs courtesy of Ventana Medical Systems, Inc.)
In situ zymography (ISZ) is a method that uses specific protease substrates to detect and localize protease activities in tissue sections. In the regulation of biological processes, proteases modulate several cellular functions. Several molecular techniques identify and characterize proteases in cells and tissue, such as a Northern blot and reverse transcription-polymerase chain reaction (RT-PCR) but ISZ works as well. One of its drawbacks is that unfixed fresh frozen tissues must be used. In contrast, its advantages are that it costs less than conventional ISH methods, there are two approaches (one uses a photographic emulsion, the other uses a fluorescent-labeled substrate) and it is applicable to almost any protease (Yan & Blomme 2003).
Immunolabeling electron microscopy (IEM) in combination with ISH has been used in detecting severe acute respiratory syndrome (SARS). Viral immunogold labeling and ultrastructural ISH were used to analyze the morphogenesis of this recently emergent virus. A negative-sense riboprobe was used for the ultrastructural ISH (Goldsmith et al. 2004).
Polymerase chain reaction-ISH (PISH) is another form of ISH. Viral RNA is detected by RT-PCR, using formalin-fixed paraffin-embedded tissue (FFPE). PISH results have been compared to IHC on staining for Newcastle disease in veterinary medicine. Newcastle disease is an avian viral infection that has a potential for rapid spread and may cause serious economic impact and international trade restrictions in the poultry industry (Wakamatsu et al. 2005). PISH is also used in the detection of human papillomavirus in uterine cervical neoplasia (Xiao et al. 2001).
The identification of mRNA sequences by in situ hybridization may be the technique of choice for the rapid and sensitive identification of viral infection. Another advantage of in situ hybridization for viral detection is that some viral coat antigens are not expressed at certain stages of the viral replication cycle, thus negating the use of immunohistochemical methods.
ISH methods have been developed over the years so that most FFPE tissues, including decalcified tissues, can be used (Janneke et al. 1999).
Another area in which in situ hybridization and immunochemistry can be viewed as complementary techniques is in the phenotyping of tumors. Many monoclonal and polyclonal antibodies are available for phenotyping and these may be employed in sensitive and rapid techniques. When problems arise in the interpretation of immunohistochemical results, mRNA phenotyping by in situ hybridization can be helpful (Pringle et al. 1990, 1993; Kendall et al. 1991; Ruprai et al. 1991).
Listed here are names of reagents that are used in ISH techniques. The formulae for preparing these reagents are located in Appendix VIII. The majority of these reagents can be purchased pre-mixed or in a kit for easy mixing. Keep in mind that different reagents may be suggested with some ISH methods and automatic IHC equipment often provide pre-package reagents as ready to use (RTU) with their equipment. Purchasing the reagents pre-mixed or in kits is convenient and safer, and it provides some reassurance that they are mixed according to manufacturer’s specifications and guaranteed by the vendor. This may cut down the possibility of human error.
1. Diethylpyrocarbonate (DEPC) treated water
2. 2% aminoalkylsilane (positively charged slides)
These slides may be purchased pre-coated
Make sure they are RNA/DNA free
Aliquot and freeze below −20°C
5. 0.1 M triethanolamine (TEA), freshly made
6. 1 M Tris (this is to make buffers that vary in pH: buffer #1, pH 7.5; buffer #2, pH 9.5)
9. Maleic acid buffer a washing buffer
10. 20× Saline sodium citrate (SSC) buffer (this is also used to make 2× SCC and 1× SCC buffers)
(may cause increase in background)
14. Detection method reagents:
15. Colorimetric detection reagents:
Probes and their choice
Probe choice is based on the type of sequence you are trying to detect. The technologist needs to optimize the conditions used as much as possible. The strength of the bonds between the probe and the target plays an important role. The strength decreases in the order RNA-RNA to DNA-RNA. Various hybridization conditions such as concentration of formamide, salt concentration, hybridization temperature, and pH influence this stability.
A probe is a labeled fragment of DNA or RNA used to find its complementary sequence or locate a particular clone. The choice of probes will depend on availability, sensitivity, and resolution required. The sensitivity of the probe will also depend on the degree of substitution and the size of the labeled fragments. Degree of substitution refers to the original nucleotide substituted by the labeled analogues. The sensitivity of detection correlates with the amount of label substituted. In general, probes with 25–32% substitution yield the highest sensitivity. There are several different types of probe. Each has unique characteristics that must be considered for each application.
Probe type and means of synthesis
There are essentially four types of probe that can be used in performing in situ hybridization. Oligonucleotide probes are usually 20–50 bases in length. They are produced synthetically by an automated chemical synthesis employing a specific DNA nucleotide sequence (of your choice). These probes are resistant to RNases and are small, thus allowing easy penetration into the cells or tissue of interest. However, the small size has a disadvantage in that it covers fewer targets. The label should be positioned at the 3′ or the 5′ end. To increase sensitivity one can use a mixture of oligonucleotides that are complementary to different regions of the target molecule. Oligonucleotide protocols can be standardized for many different probes regardless of the target genes being sought. Another advantage of oligonucleotide probes is that they are single stranded, therefore excluding the possibility of renaturation.
Single-stranded DNA probes cover a much larger size range (200–500 bp) than oligonucleotide probes. They can be prepared by a primer extension on single-stranded templates by RT-PCR of RNA, or by an amplified primer extension of a PCR-generated fragment in the presence of a single antisense primer, or by the chemical synthesis of oligonucleotides. PCR-based methods are much easier and probes can be synthesized from small amounts of starting material. Moreover, PCR allows great flexibility in the choice of probe sequences by the use of appropriate primers.
Double-stranded DNA probes can be prepared by nick-translation, random primer, or PCR in the presence of a labeled nucleotide, and denatured prior to hybridization in order for one strand to hybridize with the mRNA of interest. They can also be produced by the inclusion of the sequence of interest in bacteria, which is replicated, lysed, and then the DNA is extracted and purified. The sequence of interest is removed with restriction enzymes. Random priming and PCR give the highest specific activities. These probes are less sensitive than single-stranded probes, since the two strands have a tendency to rehybridize to each other, thus reducing the concentration of probe available for hybridization to the target. Nevertheless, the sensitivity obtained with double-stranded probes is sufficient for many purposes, although they are not widely used today.
RNA probes (cRNA probes or riboprobes) are thermostable and are resistant to digestion by RNases. These probes are single stranded and are the most widely used in ISH. RNA probes are generated by in vitro transcription from a linearized template using a promoter for RNA polymerase that must be available on the vector DNA containing the template (SP6, T7, or T3). RNA polymerase is used to synthesize RNA complementary to the DNA substrate. Most commonly, the probe sequence is cloned into a plasmid vector so that it is flanked by two different RNA polymerase initiation sites enabling either sense-strand (control) or antisense (probe) RNA to be synthesized. The plasmid is linearized with a restriction enzyme so that plasmid sequences are not transcribed, since these may cause high backgrounds. Single-stranded probes provide advantages over double-stranded probes such as:
• The probe does not self-anneal in solution, so the probe is not exhausted.
• Large probe chains are not formed in solution; thus, probe penetration is not affected.
If high sensitivity is required, single-stranded probes should be used (Table 21.1).
Probe preparation and labeling
To visualize where the probe has bound within your tissue section or within your cells, you must attach a detectable label to your probe before hybridization. Two major choices must be made for the preparation of a probe:
• What type of nucleic acid is to be used (DNA or RNA, single or double stranded)?
A vital consideration is the length of the probe, and the means by which this is controlled depends on the type and the method of synthesis. There are two methods of probe labeling. They are:
• Direct: the reporter molecules (enzyme, radioisotope or fluorescent marker) are directly attached to the DNA or RNA.
• Indirect: a hapten (biotin, digoxigenin, or fluorescein) is attached to the probe and detected by a labeled binding protein (typically an antibody).
Methods for incorporating labels into DNA are nick translation and random primer methods.
Oligonucleotide probe labeling
The 5′ end of DNA or RNA undergoes direct phosphorylation of the free 5′-terminal OH groups. The free 5′-OH substrates can be labeled using T4 polynucleotide kinase. This method is usually used for radiolabeling. Non-radiolabels use a covalent linker.
Terminal dexoxynucleotidyl transferase (TdT) is used to add a labeled residue to the 3′ end of a synthetic oligonucleotide that is approximately 14–100 nucleotides in length. These probes provide excellent specificity but only moderate sensitivity. See the oligonucleotide 3′-end labeling procedure on page 555 of sixth edition.
A tail containing labeled nucleotides is added to the free 3′ end of double- or single-stranded DNA using TdT. These probes are more sensitive than the 3′-end labeled versions, but can produce more non-specific background. Oligonucleotide tailing kits are commercially available.
It should be noted that the use of commercially available labeling kits can greatly assist in making methods simpler to undertake while providing results of an assured standard.
Purification of labeled probes
There are several methods that can be used to test the purification. If the probe is homemade or purchased it must be tested for its sensitivity to the selected target. Here is a list of methods that can be used, but it is advisable to follow the manufacturers’ recommendation on their use:
Estimating the labeling efficiency and testing the probe
It is always good practice to estimate the yield of labeled nucleic acids. This confirms a successful labeling reaction before performing the staining.
Before using a labeled probe, it is useful to prepare and demonstrate test strips to gauge the degree of label incorporation. This may be done by using the normal detection procedure for the ISH method on dots of labeled nucleic acid sequence and a labeled control applied at matching descending concentrations on a positively charged nylon membrane. Some technicians will prepare several test strips from one labeled sequence to compare the sensitivity of different detection systems. The nylon membrane is subjected to an immunological detection which can be either a colorimetric or chemiluminescent method depending on the protocol used. Direct comparison of the signal intensities of sample and control allows estimation of labeling yield. Kits for this technique are commercially available in which the labeled control already exists on a test strip. A much quicker method for estimating the yield of labeled nucleic acids is to use a bioanalyzer, since this can give quantitative results in as little as 30 minutes.
A method for estimating labeling efficiency of the nucleic acid using a dilution series followed by a spot test is described below.
Preparation of the dilution series
1. Dilute the labeling probe using dilution buffer to a starting concentration of 2.5 pmol/µl.
2. Make a dilution series of purified probe in Eppendorf tubes to give nucleic acid concentrations of 300 pg/µl, 100 pg/µl, 30 pg/µl, 10 pg/µl, 3 pg/µl, and one tube containing diluent only. Ensure that all tube volumes are equal. Repeat the same dilution series with your control or used pre-labeled (with control) test strips.
3. Apply 1 µl drops from each tube onto the nylon membrane (Roche). The control dilutions should be lined up with the test sample dilution concentration. For an example of placing spots, see Figure 21.5.
4. Label the position of each application with a pencil on the side of the strip (not on the strip).
5. Fix the nucleic acid to the membrane by either baking the membrane for 30 minutes at 120°C or using a UV light.
6. Wash the membrane briefly in washing buffer.
7. Immerse in blocking solution for 10 minutes.
8. Incubate with reagents used in ISH detection technique.
Note: Dilute reagents in blocking solution and use this solution for washing.
9. Detect enzyme using the same solutions and procedure as for ISH method.
Commercially made probes
Custom designed, pre-made cloned DNA and oligonucleotide sequences and labeling kits are commercially available. Their use can make methods simpler to undertake while providing results of an assured standard. Depending on the type of laboratory you have and how many ISH requests you receive, premixed reagents and pre-labeled probes can be cost-effective. The kits, reagents, or ordering the labeled probes can cut down on precious technologist time, and they come with instructions – a benefit for the novice technologist. However, this does not mean that the theory behind ISH can be ignored. The technologist must understand ISH to order labeled probes and the ISH kits.
For DNA probes the concentration of the probe will be ~0.5–2 µg/ml. Oligonucleotide probes can be used with, or without, acetylation. Probes without acetylation pretreatment of the sample will have a concentration of ~50–200 ng/ml and may provide more intense results with minimal background. For probes with acetylation pretreatment, a higher concentration of oligonucleotide probe may be used without incurring non-specific background staining.
Length of probe
As mentioned previously, one must consider the length of the probe. Longer probes give weaker signals and they penetrate less effectively into the cross-linked (fixed) tissue. The extent of weaker signals and penetration depends also on the nature of the tissue, the choice of fixative and whether a pretreatment has been carried out.
The length of probe can be controlled in either the synthesis reaction or the subsequent partial cleavage. In nick-translated probes, the DNA length is determined by the amount of DNase in the reaction, whereas in random priming the length is determined by concentration of the primer. Long RNA probes may show poor tissue penetration, whereas chemical shortening (hydrolysis) enhances tissue penetration, and may also increases the likelihood of non-selective binding to other non-targeted gene sequences.
Once a probe is prepared, its size should be checked. If the probe is too small, it may yield low signals with high background. It is necessary to know whether a reduction in probe size (length) will improve both the signal for the tissue and the preparation method used.
The choice of detection system will be principally determined by the probe label used and secondly by the ISH procedure type. One must consider the sensitivity and resolution required.
Colorimetric detection substrate systems include horseradish peroxidase with either 3-amino-9-ethylcarbazole (AEC) or 3,3′-diaminobenzidine tetra-hydrochloride (DAB) substrates. AEC forms a red-brown product which is alcohol soluble; therefore aqueous mounting media are required. Methyl green/blue has been the most often used counterstain in earlier publications but is losing popularity. DAB forms a permanent, insoluble, brown product that is compatible with solvent-based mounting media. Alkaline phosphatase systems can use 5-bromo-4-chloro-3-indolyl phosphate/nitro-blue tetrazolium (BCIP/NBT) or fast red. BCIP/NBT forms a purple/blue alcohol-insoluble stain. Eosin is a compatible counterstain if a nuclear target is expected, or nuclear fast red if the target is cytoplasmic. Fast red forms an intense red product which is alcohol soluble and an aqueous mounting media is required. Methyl green or blue is compatible if a nuclear target is expected, or light hematoxylin if the target is cytoplasmic.
Detection methods can be either direct or indirect. Incorporation of a stable hapten to a probe is the cornerstone of non-radiographic detection. Hybridized probes can be detected by enzymatic reactions that produce a colored precipitate at the site of hybridization. The most commonly used enzymes for this application are alkaline phosphatase (AP) and horseradish peroxidase (HRP). Although these enzymes can be conjugated directly to nucleic acid probes, such enzyme-coupled probes are often inappropriate for ISH to tissue preparations because probe penetration is hampered by the presence of the conjugated enzyme. Therefore, indirect methods are preferred (Knoll & Lichter 1995).
Biotin, fluorochromes (fluorescein), and digoxigenin (DIG) are the most common labels used. Probes that are labeled with these reporter molecules are usually detected by an AP or HRP conjugate to avidin (for biotin) or antibodies (for DIG). Fluorescein and DIG have an advantage over biotin in that they produce lower levels of background signals in tissues that contain high amounts of endogenous biotin.
Direct detection of a fluorescent label is often employed for the demonstration of multiple chromosome targets (Nederlof et al. 1989), but when single target sequences are to be identified, indirect methods may be used.
To reduce non-specific staining (particularly of collagen) by indirect detection reagents it is advisable to pre-incubate preparations in a Tris (Triton X-100 or Tween) buffer solution containing bovine serum albumin. It may also prove beneficial to use this solution as the diluent for the primary detection reagent. However, at this step it is more important to use a Fab fragment of an antibody as this may reduce background staining.
Indirect detection procedures offer increased sensitivity. The selection of the enzyme and substrate (De Jong et al. 1985) should be included in weighing the benefits of different detection systems. A substrate system that employs conjugated antibodies such as anti-DIG or anti-FITC that are conjugated with AP together with the application of a colorimetric BCIP/NBT that can be cycled to produce an insoluble blue/black precipitate over a period of 24 hours is recommended. Another substrate system one could use for a more intense fluorescent signal is a fluorescent 2-hydroxy-3-naphthoic acid-2′-phenylanilide phosphate (HNPP) with fast red TR.
The main advantages of this procedure are low levels of non-specific staining, simplicity, and the use of an enzyme substrate system that can produce an insoluble blue/black precipitate.
Many commercially available probes for ISH are labeled with biotin. When used in combination with streptavidin detection systems, high sensitivity can be achieved. A disadvantage of this combination is having a widespread endogenous tissue distribution. Substantial quantities of endogenous biotin are, for example, present in the liver and kidney (Wood & Warnke 1981), as well as in other tissues, such as pituitary, submandibular gland, thyroid, and parathyroid. Furthermore, proliferating cells may often produce enough biotin to make the discrimination between true and false-positive results difficult. However, methods of blocking endogenous biotin have greatly improved and work well to prevent false positives.
Digoxigenin (Herrington et al. 1989) in combination with a Fab fragment-enzyme conjugate detection system currently provides results of equal or superior sensitivity to biotin, with extremely low non-specific background staining. Another label that may be used in conjunction with a single-step detection method is fluorescein. Using this label it is possible to undertake rapid ISH methods in which target sequences of moderate to high copy number can be demonstrated in a working day.
Fixation is an initial step in specimen preparation or can be an intermediate step in a protocol, as in methods using cryostat sections. The duration, type, and temperature of fixation may also differ according to preparation. Together, these factors will have an effect, not only on the preservation of the tissue but also on the retention of nucleic acid and the resistance of DNA and RNA to nuclease digestion. The choice of fixative will have an influence on the conservation of nucleic acids and their availability for hybridization. Specimens that are immersion-fixed prior to paraffin embedding appear to be unaffected by ‘normal’ contamination levels of nucleases, thus indicating that only the hybridization solutions need to be scrupulously free of the enzymes.
The functional groups involved in base pairing are protected in the double-helix structure of duplex DNA. RNA is fairly unreactive to cross-linking agents.
Methanol/acetic acid fixation is recommended for metaphase chromosome spreads. Cryostat sections may be fixed with 4% formaldehyde (~30 minutes), Bouin’s fixative, or paraformaldehyde vapor fixation. This fixation also helps to secure the tissue to the slide.
Proteins surround DNA and RNA target sequences and the extensive cross-linking of these proteins may mask the target nucleic acid. Therefore, ‘permeabilization’ procedures may be required.
After the tissue is removed from the patient or animal, it must be fixed to prevent autolysis, inhibit bacterial/fungal growth, and make it resistant to damage from subsequent processing. There are two main groups of fixatives, coagulant and non-coagulant, classified by their reaction with soluble proteins. Ethanol and mercuric chloride are coagulant fixatives. They are not the preferred fixative for use with ISH, since ethanol dehydrates, coagulates, and precipitates cellular proteins, nucleic acids, and carbohydrates. Covalent bonding does not occur with ethanol fixatives and the tissue components, so mRNA is not anchored within the tissue and is likely to be lost during post-fixation processing procedures. For ISH, non-coagulant, cross-linking aldehydes (formaldehyde, paraformaldehyde, and glutaraldehyde) are recommended.
Tissues fixed for ISH should retain mRNA within the tissue but not raise background. Both background and signal are generally higher on perfused-fixed paraffin tissue sections than on frozen sections. The signal-to-noise (S/N) ratio on perfused-fixed tissue sections is better.
Most commonly, tissue specimens are routinely fixed in 10% buffered formalin, processed overnight in an automatic tissue processor, and embedded in paraffin wax. Fixation time of 8–12 hours is optimal. Keep in mind that the longer the fixation, the more rigorous the enzyme digestion is required to optimize the signal. Alcohol-fixed tissues should be post-fixed with an aldehyde fixative to prevent the diffusion of mRNA (if you are looking at RNA).
Sections are cut at 4–6 µm on an alcohol-cleaned microtome using positively charged or hand-coated slides. Sections are drained well and then air-dried at room temperature. After deparaffinization, slides are placed in an alcohol-cleaned staining container of DEPC water. The staining container is then placed in the heated water bath at 23–37°C and held until the start of ISH. Gloves must be worn to prevent contamination, and all utensils, such as brushes and forceps, should be cleaned with alcohol and kept within the cleaned area designated for ISH.
The use of formaldehyde-based fixatives prior to paraffin embedding of specimens will mask nucleic acid sequences. Digestion is a important step when performing ISH. Digestion improves probe penetration by increasing cell permeability with minimal tissue degradation. Although the nucleic acid is not directly affected by proteolytic digestion, it is important to control this step carefully. Under-digestion will result in insufficient exposure of the nucleic acid, while over-digestion can sufficiently weaken the protein structure surrounding the sequence, bringing about its loss into subsequent solutions. mRNA sequences tend to be loosely associated with proteins, while DNA targets are intimately associated with histone and other nuclear proteins. Due to these differences, the concentration of proteolytic enzyme required to unmask mRNA will be less than that necessary to expose DNA.
The selection of the proteolytic enzyme is important. This should be of molecular biology grade to ensure the absence of nuclease activity. Proteinase K and pepsin are two enzymes commonly used for digestion. Proteinase K has an advantage over other proteolytic enzymes because during incubation it digests any nucleases that might be present. However, higher concentrations of the enzyme may be required, depending on the tissue fixation time.
Nuclease digestion is used as a negative control. Treatment of same tissue/patient tissue sections with RNase A will demonstrate that ISH signals are due to RNA hybridization.
Proteoglycan digestion is required for bone and cartilage. Kidney and brain may also need proteoglycan digestion, if the signal is weak. Post-fixation in 4% paraformaldehyde is necessary after digestion to prevent tissue loss. In addition, post-fixation after digestion (for all digestions) prevents leaching and will inhibit RNase activity.
When ISH methods are used to demonstrate mRNA sequences, non-specific attachment of digoxigenin and fluorescein-labeled oligonucleotides to epithelial tissues can create non-specific results. To minimize this interaction, preparations can be acetylated after proteolytic digestion and before post-fixation. Acetylation decreases non-specific binding of the probe to the tissue. Positive charges on the tissue are neutralized by reducing electrostatic binding of the probe.
Prehybridization is intended to reduce non-specific binding. Sites in the tissue become saturated with the components of the prehybridization solution preventing non-specific binding. The purpose of the prehybridization solution is to equilibrate the specimen with the hybridization solution prior to the addition of the probe, and to allow anionic macromolecules to block sites of potential non-specific probe interaction. The prehybridization solution contains all the ingredients of the hybridization mixture except the probe. Non-complementary sequences such as bovine serum albumin (BSA at 1 mg/ml), Denhardt solution (Ficoll, BSA, and polyvinylpyrrolidone all at 0.02%), and tRNA are used to reduce non-specific binding. Most data indicate that blocking is required, but a separate prehybridization step may not be necessary. In some cases, adequate blocking may be accomplished during the hybridization step. Electrostatic binding of probes to the tissue and slides can be neutralized by treatment in TEA buffer containing 0.1 M triethanolamine.
Hybridization occurs after denaturization, during cooling, in the presence of a complementary probe, and permits hydrogen bonding of the two strands of nucleic acids. The probe must form stable hydrogen bonds with the target with minimal hybridization with non-target sequences. The probe and target sequences must be single stranded. Simultaneously heating the probe and target to high temperatures may increase the consistency and sensitivity of detection. This can only be met if care is taken to precisely control this step of the ISH procedure. Control is achieved through balancing the various components of the hybridization solution and hybridizing at an optimal temperature for the correct length of time.
If a DNA probe is employed or a DNA target demonstrated, then it is essential that these are rendered single stranded. This is achieved by using dry heat with the hybridization solution placed over the specimen, which is then covered with a coverslip, plastic sheet, or cap. The denaturation will differ according to the percentage of guanine-cytosine base pairs within the target sequence of interest. When there is a high percentage of guanine-cytosine base pairs present, the third hydrogen bond associated with the base pair will occur at a higher melting temperature (Tm) than in the sequences in which adenosine and thymidine pairing predominates. Overheating to temperatures greater than 100°C at this stage may compromise specimen preservation.
At the molecular level, hybridization involves an initial nucleation reaction between a few bases, followed by hydrogen bonding of the remaining sequences. The control of temperature during hybridization is crucial, as variations will influence the specificity (stringency) of annealing. RNA and DNA hybrids are formed optimally at about 25°C, below their Tm, but when lower temperatures are used some partial homologous annealing may occur. Although this situation should be avoided, it can be usefully employed to screen for sequences with partial homology (e.g. human papillomavirus subtypes). By incorporating formamide, a helix-destabilizing reagent, the annealing may be maintained using lower temperatures (e.g. at 37°C, at which the tissue preservation is not affected).
Stringency can also be altered by adjusting the availability of monovalent cations in the hybridization solution. These cations are usually supplied by sodium chloride and they regulate the degree of natural electrostatic repulsion between the probe and target sequences. When used at high concentration their effect is to produce conditions of low stringency, while at low concentrations only sequences with complete homology can hybridize.
Anionic macromolecules are often included in the hybridization solution to reduce non-specific interactions of the probe. Sonicated and denatured salmon sperm DNA can shield non-homologous nucleic acid sequences from the probe and reduce the opportunity for cellular electrostatic interactions. Dextran sulfate will also reduce the possibility of cellular electrostatic interactions and locally concentrate the probe, enhancing the rate of hybridization. Particular attention should be taken to ensure that hybridization solutions are prepared using reagents free from nuclease contamination.
The rate of annealing will be influenced by time and temperature as well as by the composition of the hybridization solution as discussed above. Due to steric constraints, ISH proceeds at a slower rate than in blotting methods. However, high probe concentrations can be used to compensate for this factor. Hybridization times of 1–2 hours for biotinylated and fluorescein-labeled probes are often effective. However, with digoxigenin-labeled probes, overnight hybridization may be required to provide high sensitivity.
Post-hybridization washes are used to adjust the stringency of hybridization. The sections must be rinsed with solutions that contain high concentrations of salt to remove the unbound probe. Subsequent washing with solutions containing decreasing salt concentrations and increasing temperature reduces mismatching of base pairs. Longer probes and those with higher G + C content are more stable. Increases in temperature and formamide concentration are the destabilizing factors. By reducing the concentration of formamide in the hybridization solution, while maintaining a constant temperature, the annealing conditions will become less stringent, thereby increasing the sensitivity of mRNA detection when using fluorescein-labeled oligonucleotide probes.