Robust Detection of Gene Amplification in Formalin-Fixed Paraffin-Embedded Samples by Fluorescence In Situ Hybridization

This research protocol was approved by the Institutional Review Board (IRB) of the University of Texas Southwestern Medical Center. Informed consent was obtained from all patients prior to surgery.

1. Reagents and materials preparation

1. Prepare the 0.2 N sodium chloride (HCl) solution in a fume hood by slowly adding 8.212 mL of HCl (37% w/w or 12.1 N) to 491.788 mL of ddH₂O. Store at room temperature (RT).
   CAUTION: Slowly add acid to water. Do not add water to acid.
2. Prepare 10 mM citric acid solution (pH 6.0) by dissolving 1.47 g of Tri-sodium citrate (dihydrate) in 400 mL of ddH₂O. Use HCl to adjust to pH 6.0, and then bring the final volume to 500 mL with ddH₂O. Store the buffer at RT.
3. Prepare the 10% Tween-20 solution by adding 100 µL of Tween-20 to 900 µL of ddH₂O. Store it at RT.
4. Prepare the 10% IGEPAL solution by adding 5 mL of IGEPAL CA-630 to 45 mL of ddH₂O. Store it at RT.
5. Prepare the 20x SSC (pH 7.0, 3 M NaCl, 0.3 M Sodium Citrate) solution by dissolving 44.1 g of Tri-sodium citrate (dihydrate) and 87.65 g of sodium chloride (NaCl) in 900 mL of ddH₂O. Use HCl to adjust to pH 7.0, and then bring the final volume to 1000 mL with ddH₂O. Store the buffer at RT.
6. Prepare the 2x SSC solution by adding 100 mL of 20x SSC to 900 mL of ddH₂O. If necessary, add 0.5 mL of preservative (Table of Materials). Store it at RT.
7. Prepare the probe hybridization buffer by mixing 910 µL of ddH₂O, 500 µL of 20x SSC, 50 µL of 10% Tween-20, 40 µL of RNase A, 1 mL of 50% dextran sulfate, and 2.5 mL of formamide. Aliquot into 1 mL and store them at -20 ˚C.
8. Prepare the 0.4x SSC with 0.3% IGEPAL solution by mixing 100 mL of 2x SSC, 15 mL of 10% IGEPAL, and 385 mL of ddH₂O. Store it at RT.
9. Prepare the 2x SSC with 0.1% IGEPAL solution by adding 5 mL of 10% IGEPAL to 495 mL of 2x SSC solution. Store it at RT.
10. Freshly prepare the Proteinase K digestion buffer before use by adding 1 µL of Proteinase K to 99 µL of Tris-EDTA buffer.
11. Prepare 1 mg/mL DAPI storage stock by dissolving 1 mg of DAPI into 1 mL of ddH₂O. Store it at -20 °C and away from light. Prepare the DAPI working solution by adding 1 µL of the DAPI storage stock to 999 µL of 2x SSC solution. Keep away from light till use.

2. Sample pretreatment

NOTE: The slide used here contains the specimen.

1. Age the slide at 60-90°C for 20 min or overnight (Figure 1).
   NOTE: This step facilitates paraffin melting. Usually, 20 min of heating is sufficient. It can be extended to overnight to accommodate the schedule.
2. Deparaffinize the slide by immersing it into xylene or its substitutes in a Coplin jar for 10 min. Repeat this step with fresh xylene or its substitutes. Perform all the following pretreatment and washing steps in a Coplin jar.
   NOTE: Xylene substitutes are safe and eco-friendly alternatives to xylene. One of the substitutes (see Table of Materials) performs as well as xylene, if not better. Although xylene substitutes produce less odor than xylene, using it inside a fume hood is recommended. If xylene substitutes are not accessible, all procedures are compatible with xylene-based deparaffinization without any changes.
3. Wash off the xylene substitute with 100% ethanol for 5 min.
4. Rehydrate the slide with 70% ethanol immersion for 5 min.
5. Immerse the slide into 0.2 N hydrochloric acid (HCl) at RT for 20 min.
   NOTE: HCl effectively extracts acid-soluble proteins, such as basic nuclear proteins, to improve DNA accessibility to FISH probes¹¹.
6. Immerse the slide into 10 mM of hot citric acid solution and incubate at 90-95 °C for 20 min.
   NOTE: Citric acid treatment under high temperatures similarly extracts acid-soluble proteins. Both acid treatments are thought to extract extracellular matrix proteins to decrease autofluorescence¹². It is recommended that the citric solution be preheated to the desired temperature range before being applied to the slide. Microwave can be a convenient way to do so. A water bath, such as with a sous vide cooker, is the most effective and economical solution for high-temperature incubation.
7. Rinse the slide briefly in 2x SSC to neutralize pH.
8. Digest the tissue by adding 100-200 µL (enough to completely cover the tissue depending on the size of the section) of Proteinase K digestion buffer and incubate at RT for 1 min.
   NOTE: Proteinase K digestion further increases the accessibility for FISH probes and reduces autofluorescence. The time of digestion should be optimized based on the tissue types. In most cases, 1 min of digestion is sufficient. Over-digestion leads to halo-shaped nuclei morphology, and the digestion time should be reduced.
9. Immediately stop Proteinase K digestion and dehydrate the slide by immersing it into 70% ethanol for 2 min, followed by 85% and 100% ethanol treatment for 2 min each.

3. FISH and imaging

1. Prepare the FISH hybridization mix by diluting 2 µL of FISH probe stock with 8 µL of hybridization buffer, then apply it to the slide. Cover the sample with a coverslip.
   NOTE: The total FISH probe stock used ranges from 0.5-4 µL, depending on the image quality. If the signal is too low, increase the probe input. Reduce the FISH probe input if the background is too high, especially when fluorescent debris outside the nuclei is observed. The hybridization buffer can either be the one provided with the commercially purchased probes or prepared as in the section 1.
2. Place the slides onto a hot plate, such as a slide moat hybridization system, to denature DNA at 75 °C for 2-5 min. Then, transfer the slide onto another hot plate set at 37 °C to hybridize overnight.
   NOTE: If the hot plate has a water tray or reservoir to maintain the humidity during hybridization, sealing the coverslip with rubber cement is unnecessary.
3. After hybridization, dip the slide into 40-60 °C warmed 0.4x SSC with 0.3% IGEPAL CA-630 washing buffer, then carefully remove the coverslip. Continue the washing twice for 5 min each in the dark, with agitation for the first 10-15 s.
   NOTE: Dipping the slide into the washing buffer helps gently release the coverslip.
4. Wash the slide by in SSC with 0.1% IGEPAL CA-630 for 5 min at RT in the dark, with agitation for the first 10-15 s.
5. To quench autofluorescence, treat the slide with the autofluorescence quenching kit (see Table of Materials) by applying 150 µL of reagent (50 µL + 50 µL + 50 µL of reagents A, B, C) for 2-5 min, then wash it with 2x SSC for 5 min.
   NOTE: This is an optional step. Tissue autofluorescence primarily originates from extracellular matrix components, such as collagen and elastin. It is also significantly influenced by lysosomes and mitochondria due to their lipofuscin, NADPH, and flavin coenzyme content. Aldehyde fixation and blood cell presence may also increase autofluorescence well^13,14.
6. Stain the slide with DAPI for 10 min. Rinse the slide with 2x SSC buffer for 5 min.
   NOTE: If the slide is not treated by the autofluorescence quenching kit, DAPI staining can be reduced to 2 min.
7. Quickly dip the slide into deionized water for no more than 1 s, then quickly dry it by absorbing extra moisture with a paper towel.
   NOTE: This is an optional step. Deionized water treatment effectively prevents salt crystal deposition of SSC buffer on the slide and improves imaging quality. However, under such low ion conditions, hydrogen bonds between the FISH probe and targeted DNA are weakened, leading to probe dissociation and signal loss. Therefore, maintaining the water treatment step for a very short time is crucial.
8. Dry the slide, then mount it with antifade mounting media. Seal the coverslip with nail polish before imaging.
   NOTE: If the slide is treated by autofluorescence quenching reagent, mount the slide with an antifade mounting medium per the manufacturer's instruction. Moreover, depending on the type of mounting media, hardening or non-hardening, the sample must be cured for 1-24 h before sealing and imaging. It is recommended that the sample be cured at least overnight to achieve the best refractive index for imaging.
9. Use a 60× oil lens to capture fluorescence signal. Use the DAPI channel to adjust the focus. Ensure to obtain multiple Z-stacks. Typically, 5-10 z-stacks with a 1-µm interval are sufficient. Perform a maximum 3D projection to achieve the best resolution. Apply deconvolution or other background-clearing algorithms to further improve the image quality.