Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

A summary of tissue pre-processing steps may be found in Figure 1. All procedures were carried out in compliance with the Animal Care and Ethics Committee of the University of New South Wales in accordance with the guidelines for the use and care of animals for scientific purposes (Australian National Health and Medical Research Council).

1. Sample preparation of fresh frozen brain tissue

1. Transcardial Perfusion
   1. Prepare heparinized (2500 U/L) 0.1 M phosphate buffer (PB), pH 7.5. Make dry ice ethanol slurry by mixing dry ice with ethanol. This will have a temperature of approximately −72 °C and will be used for immediate freezing of the harvested tissue.
   2. Euthanize adult C57BL/6 and Phox2b-GFP¹¹ (Mouse Genome Informatics database ID MGI:5776545) mice by anesthetizing with sodium pentobarbital (70 mg/kg, i.p.), using a 27.5 inch needle gauge.
      CAUTION: Pentobarbital is a barbiturate. It is acutely toxic in high doses and may cause death by respiratory arrest. Consult local healthcare, legal and material safety guidelines before use.
   3. Expose the heart and cannulate the left ventricle with a drawing-up needle (23 inch gauge). Perform transcardial perfusion with heparinized 0.1 M PB until the blood clears (2-3 minutes) at a flow rate of 11-13 mL/min. Determine blood clearing by monitoring the coloration of the liver and the effusate from the right atrium¹².
   4. Isolate the brain from the skull cavity, immediately embed it in Optimal Cutting Temperature Compound (OCT) in a cryomold or aluminum foil and place it on the dry ice ethanol bath. Store the frozen embedded tissue in an airtight container at – 80 °C for up to 3 months.
2. Sectioning of fresh frozen tissue
   1. Set the cryostat temperature to -20 °C. Leave the OCT-embedded tissue and a cryostat chuck in the cryostat for ~30 minutes to allow for equilibration to the new temperature.
      NOTE: Keep the tissue frozen at all times; transport the tissue from the -80 °C freezer to the cryostat on dry ice.
   2. Secure the tissue to the pre-chilled cryostat chuck using OCT compound. In this protocol, tissue blocks were mounted onto the chuck in the coronal plane.
      NOTE: Trim excess OCT from the tissue, using a razor blade, to minimize the amount of OCT being cut by the cryostat and subsequently transferred onto the glass slide.
   3. Cut 14 µm thick coronal sections and mount them onto charged glass microscopy slides.
      1. Warm the slides to room temperature before mounting the sections. Once the section has been mounted, keep the slides in a slide box in the cryostat.
      2. If more than one section needs to be mounted on one slide, warm the area for the second section by placing a finger on the opposite side of the slide for 5-10 seconds to aid adherence of the section to the slide. A cold tissue section will not attach to a cold slide. The sections should adhere to the slides flat; folding will cause them to fall off the slides during wash steps.
      3. If cracks are noticed in the sections, increase the cryostat temperature by 1-5 °C to avoid this. It is particularly important to place tissue sections in close proximity to one another on the same slide. This will prevent wastage of FISH probes and reagents during the assay.
   4. Store tissue sections mounted onto glass slides in an air-tight container at -80 °C for up to 6 months.
      NOTE: Keep the sections frozen at all times and avoid freeze thaw cycles, to prevent RNA degradation. Transport the slide box from inside the cryostat to the -80 °C freezer on dry ice.
3. Fixation of fresh frozen tissue
   1. On the day the FISH probe assay is to be performed, prepare 4% paraformaldehyde (PFA) in 0.1 M PB, pH 7.5 (4% PFA solution). Filter by passing through filter paper (Grade 1: 11 µm, Table of Materials) in a Buchner funnel or crucible filter.
      CAUTION : PFA is harmful and toxic by skin contact or inhalation. All procedures with PFA solution should be performed in a fume hood cabinet. PFA solution waste should be disposed of carefully following institutional safety protocols.
   2. Cool the 4% PFA solution to 4 °C. Transport the slide-mounted tissue from the -80 °C freezer in dry ice and immediately immerse it in the pre-chilled fixative for 15 minutes.
      NOTE: It is important that this fixation step does not exceed 15 minutes as over-fixation will result in non-specific background labelling.
4. Dehydration of fresh frozen tissue
   1. Dehydrate tissue sections by submerging the slides in graded concentrations of ethanol. In a Coplin jar, first submerge in 50%, then 70% and finally absolute ethanol, for 5 minutes each at room temperature. Repeat the final absolute ethanol incubation a second time.
   2. Air dry slides and, outline the group of sections using a hydrophobic barrier pen, ensuring that the internal area is kept to a minimum.
      NOTE: Make sure that the glass slide is completely dry before drawing the hydrophobic barrier. The hydrophobic barrier should surround the tissue sections completely without gaps and must be dry before further processing.

2. Sample preparation of fixed frozen brain tissue

1. Transcardial perfusion fixation
   1. Euthanize mice by anesthetizing with sodium pentobarbital (70 mg/kg, i.p) followed by transcardial perfusion, first with 0.1 M PB then 4% PFA solution. Fix with 10 minutes of perfusion at 11-13 mL/min.
   2. Isolate the brain from the skull cavity following perfusion-fixation and submerge overnight in 4% PFA solution, at 4 °C.
2. Tissue sectioning of fixed tissue
   1. Rinse the brain in sterile 0.1 M phosphate buffered saline (PBS) before removing the meningeal layers, with the aid of a dissecting microscope, using fine forceps.
   2. Cut the brain precisely into blocks (separate the brainstem from the forebrain prior to vibratome sectioning) using a brain matrix (Table of Materials). Specifically, cut the brainstem caudally at the pyramidal decussation and dissect away the cerebellum. Similarly, cut the forebrain immediately rostral to the optic chiasm.
   3. Secure the tissue onto a vibrating microtome chuck using cyanoacrylate and embed in 2% agar solution.
   4. Cut 30 µm thick tissue sections using a vibrating microtome and store cut sections in cryoprotectant solution (30% RNase free sucrose, 30% ethylene glycol, 1% polyvinylpyrrolidone (PVP-40), in 0.1 M PB, pH 7.4). Tissue sections may be stored in cryoprotectant at -20 °C for up to 6 months.
3. Preparation of fixed sections prior to FISH
   1. On the day of FISH, wash free floating sections three times, for 10 minutes per wash, to remove the cryoprotectant solution. To wash, place sections in 0.1 M PBS in a 12 well cell culture plate and agitate on a rotating platform shaker (90 – 100 rpm).
   2. After washes, use a paintbrush to mount sections on glass microscopy slides and air dry for at least 2 hours.
      NOTE: The sections should adhere flat onto the slides as any pronounced folds will cause them to detach during washes.
   3. Using a hydrophobic barrier pen, draw a barrier around the sections to restrict the FISH reagents to the sections. Once more, it is important to minimize the internal area of the outline drawn with the barrier pen.
      ​POSSIBLE BREAK POINT: The sections could be stored at room temperature, overnight, to continue the assay the next day.

3. FISH assay

NOTE: The rest of the protocol applies to both fresh frozen and fixed frozen tissue.

1. Prepare the reagents and instruments for hybridization and amplification steps.
   1. Set a benchtop incubator and water bath to 40 °C.
   2. Prepare a humidified, light-protected chamber for incubating slides. Humidification prevents drying of tissues – slides are securely located above a moist reservoir. Ideally, the chamber is made of heavy-duty polystyrene, it is light-proof and airtight to maintain a saturated water vapor atmosphere. Closure of the chamber relies on minimal friction to avoid movement. We used a slide box lined with wet laboratory wipes (Table of Materials) at the bottom. Place the slide box inside the incubator to prewarm it to 40 °C.
   3. Warm the 50x Wash Buffer (Table of Materials) and probes to 40 °C for 10 minutes, using the water bath, then cool to room temperature.
   4. Prepare 1 L of 1x Wash Buffer from the 50x stock concentration.
   5. Prepare probe mixture (Table of Materials): the C1 probe is ready to use at stock concentration whereas C2 and C3 probes are shipped as 50x concentration and require dilution with the diluent supplied in the kit.
      NOTE: Probe mixtures can be stored at 4 °C for up to 6 months.
2. Protease treatment
   1. Incubate sections with Protease III (Table of Materials) at room temperature for 30 minutes.
      NOTE: Ensure that Protease III and incubation reagents in downstream processes (probe mixture, amplification solutions, blocking buffer and antibody sera) cover the sections entirely. A pipet tip may be used to spread the reagent onto the section to cover the entire area inside the hydrophobic barrier.
   2. Wash slides twice with 0.1 M PBS, for 2 min each time, in a large plastic square Petri dish. Here a 245 mm x 245 mm square bioassay dish was used (Table of Materials). Hold from one side of the dish and tilt gently 3-5 times. After washes, flick excess 0.1 M PBS from slide and immediately add the next reagent. Do not let tissue sections dry.
      NOTE: During each wash, the slides are immersed in solution at room temperature. This is the workflow for all subsequent wash steps. The fixed 30 µm thick sections dislodge from slides more easily than 14 µm thick sections, be gentle during the washes.
3. Hybridization and amplification
   1. After washing off the protease solution, place the slides in the humidified, prewarmed chamber. Incubate sections with probe mixture (Table of Materials) for 2 hours at 40 °C inside a benchtop incubator.
      ​NOTE: Ensure there are at least 2 sections set aside for positive and negative control probes to assess sample RNA quality and optimal permeabilization. Positive control probes target house-keeping genes; here, these were a cocktail of RNAs targeting ubiquitin C (UBC; high-abundance), peptidylpropyl isomerase B (PPIB; moderate-abundance) and RNA polymerase 2a (POLR2A; low-abundance). Negative control probes target the bacterial 4-hydroxy-tetrahydrodipicolinate reductase (DapB) gene, which is normally absent in mouse brain samples. Positive DapB signal indicates non-specific signal and/or bacterial contamination of the sample.
   2. Following hybridization with the probe mixture, the signal amplification steps consist of incubation with Amp 1-FL (30 minutes), then with Amp 2-FL (15 minutes), followed by Amp 3-FL (30 minutes) and finally Amp 4-FL (15 minutes) – each at 40 °C. Using the dropper bottles provided, cover tissue sections with amplification solution. Proceed to IHC assay following the last amplification step.
   3. Rinse slides with Wash Buffer twice for 2 minutes between probe hybridization and each amplification step.

4. IHC Assay

1. IHC blocking step
   1. To prevent non-specific binding of antibodies, incubate the sections for 1 h at room temperature with blocking solution containing 10% normal horse serum, 0.3% Tween20 in 1x TBSm (50 mM Tris-Cl, pH 7.5, 150 mM NaCl, 0.05% merthiolate) following the FISH assay. Prepare primary antibodies in a dilution buffer containing 1x TBSm, 5% normal horse serum and 0.1% Tween20. Primary antibody suppliers are listed in the Table of Materials.
2. Immunohistochemistry
   1. Remove excess blocking buffer by flicking the slide and incubate sections with primary antibodies overnight at 4 °C.
   2. Wash slides 3 times (5 minutes each) with 1x TBSm and incubate with secondary antibody in diluent containing 1x TBSm, 1% normal horse serum and 0.1% Tween20 for 2 hours at room temperature. Secondary antibodies used in this protocol are listed in the Table of Materials.
   3. Wash slides 3 times with 1x TBSm (5 minutes each) before coverslipping with mounting medium with or without DAPI (Table of Materials).

5. Imaging

1. Examine immunostaining under an epifluorescence microscope equipped with a camera (see Table of Material for details). Acquire representative images at 20x magnification and save as TIFF files.
2. Export representative images into an image processing software (Table of Materials) for brightness/contrast adjustment to increase the clarity and to reflect true rendering.

6. OPTIONAL: Quantitative analysis of target transcripts

NOTE: This is a methods article and quantitative results are not provided. The method of quantification presented here is sourced from Dereli et al.¹⁰.

1. Acquire images from the regions of interest as explained in 5.1 and apply the same microscope and camera settings (such as exposure time and light intensity) to all images of the same fluorophore.
2. Plot the neuronal profiles using an image analysis software (Table of Materials).
3. Align the sections with reference to Bregma level according to a stereotaxic brain atlas¹³.
4. Apply the same brightness and contrast to all the images of the same fluorophore. Only consider the neurons with DAPI-stained nuclei.
5. Manually count the number of mRNA, protein expressing, mRNA/mRNA, protein/protein and mRNA/protein coexpressing cells within the region of interest.
6. To decrease bias in the experimental results, have the person quantifying experimental outcomes blinded to the experimental groups.
7. Apply Abercrombie correction¹⁴ to total cell counts by using the following Abercombie equation:
   Corrected cell count = manual cell count x section thickness / (section thickness + nuclear size)
   For example, for 14 µm thick sections, the average nuclear width is calculated to be 7.7 ± 0.3 µm and average section thickness is 14 ± 1 µm based on 30 cells and 10 sections respectively in 5 animals¹⁰. According to the Abercrombie equation, corrected cell count would be manual cell count x 14/(14+7.7).

Figure 1: Parallel workflow of tissue pre-processing steps for both fresh-frozen and paraformaldehyde fixed tissue. Processing steps for fresh-frozen tissue are displayed in the red outlined boxes, whereas those for paraformaldehyde (PFA) fixed tissue are displayed in the blue outlined boxes. Please click here to view a larger version of this figure.

Figure 2: Summary of combined FISH probe and immunohistochemistry procedure. Following tissue pre-processing, the slide mounted tissue is encircled using a hydrophobic barrier pen, as seen in the first frame, and incubated in a protease solution at room temperature. Following washes, tissue is transferred to a benchtop incubator for hybridization for 2 hours before sequential amplification steps. The in situ hybridization system utilizes a proprietary 'Z probe' design, preamplifiers and amplifiers as seen in frames 3-6⁶. Once tissue has undergone FISH probe processing, it is washed before blocking with normal horse serum. The primary antibody incubation is carried out overnight at 4 °C to maximize antibody-antigen binding. The secondary antibody incubation (2 hours) was carried out at room temperature. Please click here to view a larger version of this figure.