During the in situ hybridization procedure it is important to work accurately and cleanly to ensure the results are reproducible, to reduce the accumulation of dirt that causes background, and to reduce loss or damage to the material. All tools and containers must be clean/ sterile, and solutions without fungal/ microbial growth, although aseptic conditions are not necessary. The preparations on slides must not touch other slides (a frequent cause of scratching material), nor dry out or accumulate water when incubating and between steps. Wash solutions should cover slides totally and should be changed carefully to avoid strong turbulence.

The in situ hybridization procedure involves following steps:

Following solutions are required for FISH procedures. The quantity of chemicals and the method of preparing the solutions we follow in our laboratory are as follows:

1) 0.005 % Pepsin: dissolve 0.25 mg pepsin in 5 ml of protease buffer (0.001 N HCl pH 2-3→50 μl of 1N HCl in 49.95 ml of DDW).

2) 20X SSC: 3M NaCl, 0.3M Sodium citrate, pH 7.

3) PBS

4) 1% Formaldehyde

5) Denaturation Buffer: 35 ml Deionised formamide, 5 ml of 20X SSC, 10 μl of 0.5 EDTA pH 7, make final volume of 50 ml.

6) Hybridization Buffer: 5.5 ml formamide, 1 g dextran sulphate, 0.5 ml 20X SSC pH 7, make final volume 7 ml.

7) Hybridization mixture: 7 μl hybridization buffer, 1μl (2.5 μg) salmon sperm DNA and 2 μl labeled probe (100 ng).

8) Washing solution 1: 50% formamide, 10% 20X SSC and 40% distilled water.

9) Washing solution 2: 4X SSC and 0.05% NP-40.

9) Washing solution 3: 4X SSC.

High quality chromosome preparations are required for the best DNA:DNA in situ hybridization. An ideal method of tissue preparation ensures both good specimen morphology and that the target molecules are in the optimum state for probe access and hybridization. DNA:DNA in situ hybridization is usually carried out on chromosome spread preparations where chromosome and nuclei are released from cells and spread on a glass microscope slide. This method yields well-separated and enlarged chromosomes with good morphology, which can be analyzed in transmitted light or fluorescence microscopes. As precipitating fixatives including methanol: acetic acid are used, many proteins are destroyed and access of probes to the DNA within the spread chromosomes is very good. Spread preparations are used for mapping and analyzing the long-range organization of DNA sequences, and can be made from mitotic or meiotic material or DNA depending on the resolution required.

The protocols about making chromosome preparations from different cells and for different purposes including karyotype analysis and chromosome banding have been discussed in previous chapters.

The first requirement for chromosome preparation is the glass slide. Different batches and suppliers of glass clearly give different qualities of in situ hybridization results. With the acid treatment below, all slides seem to work successfully. Omitting this step often leads to selective loss of nuclei and metaphase spreads and poor in situ hybridization results.

Freshly prepared (1-3 day old) slides are always better option for in situ hybridization results. However, slides stored at –80 °C or –20 °C can also be used. Preparation of fresh slide from the already fixed can be made on acid cleaned slides as given below.

• Take fixed (Methanol: Acetic acid 3: 1) cells in a 1.5 ml eppendorf tube and centrifuge at 3,000 rpm for 5 min.
• Remove the supernatant, add fresh fixative and centrifuge again at 1500 x g for 10 min.
• Remove supernatant and resuspend cell pellet in small volume of fresh fixative (according to cell pellet size).
• Take 10 μl of final cell suspension and drop onto the centre of a slide (ice cold). Spread cells by providing heat by touching it the backside of slide with the back of palm.
• Check cell density under microscope (mark the best area by diamond tip marker).
• Dehydrate the slide in 70%, 90%, and 100% ethanol series for 3 min each at room temp.
• Leave it to air dry at room temp.

Pretreatments of slide preparations are required for following purposes:

1. To remove extraneous RNA and proteins, which will bind to probe and detection reagents increasing background.
2. To enable access of probes to the DNA by permeabilizing the target material, and to fix the preparation so that the chromosomes and nuclei are not lost from the slide during the procedure.

The pretreatment of slides before proceeding to the in situ hybridization is usually done by following the steps given below:

• Age the slide at 90 ?C for 1 h in Thermal cycler/ Thermobrite/ Hot air oven or Chemical aging of slide in 2X SSC for 1 h at 37 ?C, may add 0.1% Tween 20.
• Treat the slide with 0.005 % pepsin at 37 ?C for 10-20 min.
• Wash slide with PBS for 5 min.
• Fix the slide with 1% formaldehyde for 5-7 min.
• Wash slide with PBS for 5 min.
• Perform serial dehydration of slide in 70%, 90% and 100% ethanol for 1-2 min each.

Procedures for in situ hybridization of labeled DNA probes to spread or sectioned chromosomes and nuclei (DNA) are as below:

• Set water bath at 37 ?C and 75 ?C.
• Denature 10 μl hybridization mix at 95 ?C for 10 min.
• Immediately chill the mix on ice for 5 min.
• Transfer the denatured probe at 37 ?C in incubator for 15-60 min for pre-hybridization for binding of salmon sperm DNA (SSD) with small repetitive DNA probes.
• Incubate slide in denaturation buffer for 2-5 min at 75 ?C.
• Wash slide with cold 2X SSC for 2 min, repeat this step two times.
• Perform serial dehydrate of slide with 70%, 90% and 100% chilled ethanol for 1-2 min.
• Dry the slide at 42 ?C for 5 min.
• Apply 10 μl of denatured probe mix on the slide.
• Immediately cover with cover slip and seal with rubber seal. Keep the slide in moist chamber at 37 °C overnight - two days for hybridization.

• Set water bath at 45°C.
• Wash the slide with washing solution 1 at 45° for 5 min.
• Wash slide with washing solution 2 at room temp for 2 min.
• Wash slide with washing solution 3 at room temp for 2 min.
• Perform serial dehydration of slide with 70%, 90% and 100% ethanol for 1-2 min each and allow to air dry.

The location of the hybridized probe on the chromosome is observed under fluorescent microscope using suitable filter. If the probe is made with fluorescent-tagged dUTP by nick translation or PCR (direct labeling), the probe signal can directly be observed. In this case following procedure is adopted:

• Apply 40 μl of DAPI (with anti-fade solution) for 1 h at room temperature and cover with cover slip to counter stain.
• Examine the slides under fluorescent microscope using suitable filters.

If the probe is labeled using Biotinylated (Biotin) dUTP by nick translation (indirect labeling), for signal detection the treatment procedure involves successive rounds of Fluorescein Avidin DCS and Biotinylated Anti-Avidin to detect and amplify in situ hybridization signals. The multiple binding capacities of Biotinylated Anti-Avidin provide the potential for significant amplification. This antibody binds to Avidin through the antigen binding sites or through the biotin residues that are covalently attached to the molecule. Following the first application of Fluorescein Avidin DCS, the signal is amplified by incubation with Biotinylated Anti-Avidin, followed by a second incubation with Fluorescein Avidin DCS. This procedure results in the introduction of several more fluorochromes at the target site. Following steps are followed for observing the probe signals.

1. After hybridization with biotinylated DNA/RNA probes, block tissue sections or chromosome spreads for 30 min in 1x ISH Blocking Solution (5x ISH Blocking Solution, Vector lab Cat. No. MB-1220). The effectiveness of the blocking solution may be enhanced by pre-warming the solution to 37°C and incubating tissue sections/ chromosome spreads for 30 min or longer at 37°C.

Note: 5% nonfat dry milk plus 0.1% Tween 20 in 4X SSC can be used as an alternative blocking solution. (4X SSC is 0.6 M NaCI, 60 mM sodium citrate, pH 7). However, non-fat dry milk can contain variable amounts of biotin which could reduce staining if used as a diluent for (strept)avidin conjugates.

2. Dilute each of the detection reagents, i.e. Fluorescein Avidin DCS (Vector lab Cat. No. A-2011) and Biotinylated Anti-Avidin (Vector lab Cat. No. BA-0300) to 5 μg/ ml in 1x ISH Blocking Solution, approximately 30 min before use to minimize non-specific binding.

Note: This procedure will require twice the volume of Fluorescein Avidin DCS as Biotinylated Anti-Avidin.

3. Flip off the blocking solution and add the Fluorescein Avidin DCS solution (5 μg/ ml). Incubate for 30 min at room temperature.

4. Wash slide for 2 - 3 min in 1X ISH Blocking Solution.

If satisfactory sensitivity has been achieved, skip up to step 7. For increased sensitivity, continue with steps 5 through 7.

5. Incubate with the Biotinylated Anti-Avidin solution (5 μg/ ml) for 30 min at room temperature.

6. Wash slide for 2 - 3 min in 1X ISH Blocking Solution.

7. Follow with a second incubation of the same Fluorescein Avidin DCS solution (5μg/ ml) for 30 min at room temperature.

8. Wash slide 2 - 5 min in 4X SSC + 0.1% Tween 20 before cover slipping with mounting medium.

• Apply 40 μl of DAPI (with antifade solution) to counterstain for 1 h at room temperature and cover with cover slip.
• Examine the slide under fluorescent microscope using suitable filters in dark room.

1. Kumar Ravindra, Kushwaha B, Nagpure NS (2012). Fluorescence in situ hybridization (FISH) in fishes: A Practical Approach. NBFGR Publication.