Procedure for Fluorescence In Situ Hybridization (FISH)

    The basic principle of fluorescence in situ hybridization (FISH) is that, based on the principle of complementary base pairing between two nucleic acids, DNA probes are labeled with specially modified nucleotide molecules, which are then directly hybridized in situ to chromosomes or DNA fiber sections. The labeled probes subsequently bind specifically to fluorescein-conjugated monoclonal antibodies, allowing for the localization, identification, and relative quantification of DNA sequences on chromosomes or DNA fibers through a fluorescence detection system and image analysis techniques. The experimental procedure is as follows:

I. Slide Preparation:

(1) Slide washing: Scrub with hot soapy water, soak in 1% hydrochloric acid for 24 hours, then soak in 0.1% diethyl pyrocarbonate (DEPC) for 24 hours.

(2) Silanization: Boil the slides and coverslips in 1% (w/w) hydrochloric acid for 10 minutes, dry them, wrap them in aluminum foil, and store at 4 °C for later use.

(3) Preparation of Gelatin Mounts: Place the dried slides in gelatin for 10 minutes, then dry them at 60 °C overnight for later use.

(4) Treatment of reagent vials, plasticware, and tissue homogenizers: First, thoroughly clean the reagent vials and tissue homogenizers. Soak them in a 1 mL/L DEPC (diethyl pyrocarbonate) aqueous solution (37 °C for 2 hours, then at room temperature overnight). Remove the DEPC by autoclaving, then dry at 250 °C for at least 4 hours or at 200 °C overnight. Weighing spoons should also be dry-heated. It is best to use sterile, disposable plasticware, which should be autoclaved before use. To ensure quality, all pipette tips, test tubes, and other items used must be treated with a 0.5 mL/L DEPC (diethyl pyrocarbonate) aqueous solution for at least 3 hours, followed by autoclaving and drying. Imported pipette tips and test tubes that have already been treated to be free of RNase and DNase may be used directly without further treatment.

II. Sample Preparation and Related Reagents Include:

(1) Buffer Solutions:

1× PBS buffer solution: 8 g NaCl, 2.9 g Na₂HPO₄, 0.2 g KCl, 0.2 g KH₂PO₄, dissolved in 1000 mL ultrapure water;

3× PBS buffer: 24 g NaCl, 8.7 g Na₂HPO₄, 0.6 g KCl, and 0.6 g KH₂PO₄ dissolved in 1000 mL ultrapure water;

Typically, a 10× PBS stock solution is prepared; 3× PBS and 1× PBS can be obtained by diluting the stock solution with DEPC-treated water.

(2) 4% Paraformaldehyde (PFA)

Weigh 2 g of PFA and add it to 30 mL of hot water at approximately 60 °C. Add a few drops of 20% NaOH and stir on a magnetic stirrer until completely dissolved. Then add 16.5 mL of 3× PBS buffer to the mixture, mix thoroughly in an ice bath until cooled, and after cooling, adjust to neutrality (approximately 7.2) with HCl. Remove the stir bar. Add ultrapure water to the PFA solution and make up the volume to 50 mL. Filter through a 0.45-micron membrane before use. (Store at room temperature or 4 °C for later use.)

(3) Prepare 50%, 80%, and 98% anhydrous ethanol solutions, each with a total volume of 20 mL.

(4) DAPI Solution

Dissolve DAPI in 1 mL of ddH₂O to prepare a 2.9 mM DAPI solution (1 mg DAPI/1 mL H₂O). (DAPI cannot be dissolved directly in buffers such as PBS; it must first be dissolved in water.) Add an appropriate amount of the DAPI aqueous solution to PBS to prepare a 10–50 µM DAPI solution.

(5) Preparation of Various Solutions: All water-soluble liquids should be prepared using 1 mL/L DEPC-treated water.

Note:

① When preparing the solution, work in a well-ventilated area and avoid skin contact and inhalation (wear gloves and a mask), as PFA has strong fixing properties and is toxic; it can cause fixation, toxicity, and irritation to mucous membranes and skin.

② When heating, do not exceed 60–65 °C; otherwise, PFA will degrade and lose its effectiveness.

③ Although prepared PFA can be stored for a certain period, the fixing efficacy of the solution decreases over time; therefore, it should be used as soon as possible.

④ 4% PFA is currently the most commonly used fixative in in situ hybridization histochemistry. It effectively preserves RNA within tissues and cells while also maintaining good morphological integrity. The sample fixation time is 2–3 hours.

⑤ The RNA content is relatively constant. Excessively prolonging the fixation time can cause excessive cross-linking of biomolecules within the cells, affecting probe penetration and reducing hybridization efficiency.

III. Sampling and Storage Methods:

(1) Before precipitation in the reactor, transfer 2–5 mL of the solution to a centrifuge tube.

(2) Centrifuge (2000 r/min) for 5 min, then remove the supernatant (repeat twice after adding distilled water).

(3) Add 1 mL of paraformaldehyde to the sample, mix well, and incubate at 4 °C for 3 h.

(4) Centrifuge the sample (12,000 r/min) for 5 min, then remove the supernatant.

(5) Add 1× PBS, mix well, centrifuge (10,000 r/min) for 5 min, and remove the supernatant (repeat 3 times).

(6) Add 0.5 mL of 1× PBS and 0.5 mL of anhydrous ethanol, mix well, and store at –20 °C (can be stored for 6 months).

IV. Sample Fixation:

    Dilute the sample 3–5 times, then subject it to ultrasonic treatment (3 W for 2–3 min, allow to settle for 1 min, remove the sediment, then 5 W for 5 min) to break up the activated sludge flocs into individual cells for microscopic counting. Then, apply 3 μL of the sample to a glass slide coated with embedding gelatin (check the slide background), and fix in a 37 °C oven for 2 h (or air-dry for 2 h or overnight). Successively immerse the slide in 50%, 80%, and 98% (by mass) ethanol for 3 min each to dehydrate the cells, then stand the slide upright and allow it to dry.

V. Sample Hybridization:

    The buffers used in this experiment include Hybridization Buffer (HB) and Washing Buffer (WB), whose components are listed in Table 1-1 and Table 1-2.

(1) Pipette 2 mL of hybridization buffer and spread it evenly over the folded absorbent paper inside the hybridization cassette. Place the slide with the fixed sample into the hybridization tube, then incubate it in a 46 °C hybridization oven for several minutes.

(2) Pipette 10 μL of probe stock solution and mix it with 80 μL of hybridization buffer; wrap the mixture in foil and preheat it in a 46 °C hybridization oven for several minutes. The probe stock solution has a concentration of 25 ng/μL; dilute the purchased probe with sterile water. The probe sequences and hybridization conditions used in the experiment are shown in Tables 1–3.

Note:

(1) Indicates the concentration of deionized formamide in the hybridization buffer (%).

(2) Pipette 9 μL of the preheated probe dilution onto the sample on the slide, then quickly return the slide to the hybridization tube and allow hybridization to proceed at 46 °C (in the dark) for 2–3 hours.

(3) Post-hybridization elution: Turn on the constant-temperature water bath and heat it to 48 °C to preheat the hybridization buffer and wash buffer. Remove the slide from the hybridization kit, rinse the sample with hybridization buffer, then quickly place it in the wash buffer. After 20 minutes in a 48 °C water bath, rinse the sample with 4 °C ice water and air-dry it on a clean work surface.

VI. DAPI Staining:

   Add 9 μL of DAPI to each well, incubate in the dark at 4 °C for 5 minutes, rinse with 4 °C ice water (BPS buffer), and air-dry. Observe the cells using a fluorescence microscope equipped with filters for a 360 nm excitation wavelength and a 460 nm emission wavelength. DAPI (4,6-diamido-2-phenylindole) staining is used for whole-cell counting.

VII. Mounting:

    Apply mounting medium, place a coverslip on top, and seal with nail polish after ensuring there are no air bubbles.

VIII. Fluorescence Microscopy Analysis:

    Acquire images from 20 different regions for each sample and each probe. Each region must contain more than 1,000 cells, and the  results are then averaged to obtain the hybridization results for each probe in each sample. Bacterial abundance or bacterial count (cells/g or cells/L) is calculated as AS1/(S2V), where A is the average number of bacteria in the field of view; S1 is the smear area of the sample; S2 is the smear area of the field of view; and V is the sample volume.