Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

1. Preparation of DNA Templates

1. Download the 16S rRNA sequence specific to each bacterial genera from the NCBI database and design polymerase chain reaction (PCR) compatible forward and reverse primers that will amplify genera-specific regions (~200 – 250 base-pair long) as described earlier¹⁹. Also refer to the open source databases SILVA^20,21 and probeBase²² online resources for rRNA-targeted oligonucleotide probes and primers, as RNA probes for some bacterial genera may be commercially available.
   Note: It is important to select gene regions that are specific to specific genera and ensure that designed primers do not amplify related bacterial genera. This is critical to obtain gene/genera-specific probe hybridization.
2. Feed ticks for 24, 48 or 72 h on mice, dissect tick guts and prepare RNA and cDNA from tick guts as described earlier²³. Use cDNA as template to PCR amplify genera-specific amplicons using a thermocycler. Run an aliquot of the PCR-product/amplicon on a 1.5% agarose gel and confirm that the amplicon corresponds to the expected size by visualization under ultraviolet (UV) light using a gel imaging system.
3. Clone the bacterial 16S ribosomal RNA amplicon of interest into a commercially available TA vector (Table of Materials) containing RNA polymerase promoters suitable for bacteriophage polymerases (SP6, T7 or T3). Sequence the clones to confirm the identity of the amplicon and the directionality of the clone with respect to each of the promoters. Based on this, determine the promoter of choice to generate sense or antisense RNA probes (Figure 1).
4. Linearize 1 mg of the plasmid with an appropriate restriction enzyme in a reaction volume of up to 30 µLfor 2 h at temperatures recommended by the manufacturer. Choose restriction enzymes based on the promoter that will be utilized to generate the sense or antisense probe (Figure 1). Verify linearization by visualization of 2 µL of digest reaction on a 1% agarose gel.
5. Purify the remaining 28 µL of digested DNA using a commercially available PCR product column purification kit and quantify DNA concentration by spectrophotometer.
   Note: Some sense probes may cause background staining much higher than the corresponding antisense probe and other options may need to be explored. Alternative negative controls include plasmids encoding sequences not represented in the sample or another probe that yields a distinctive pattern of hybridization.

2. Construction of Digoxygenin-UTP RNA Probes

1. Prepare the nucleotide mix by combining 7.5 µL of 100 mM UTP, 25 µL of 10 mM digoxygenin-UTP, and 10 µL each of 100 mM ATP, GTP, and CTP in a 1.5 mL centrifuge tube.
2. Perform in vitro transcription using commercially available T7 or Sp6 RNA polymerase kits, using 1 µL of the nucleotide mix (Step 2.1) and 0.25 – 1 µg of template DNA. Bring the volume up to 20 µL with water. Flick the tube to combine and pulse spin. Incubate at 37 °C for 2.5 – 3 h.
   Note: Probe construction has been successful with digested plasmid concentrations as low as 7 ng/µL, but typical concentrations at this step range from 15 – 25 ng/µL. The transcription reaction may also be incubated overnight at 37 °C, but this does not significantly increase the RNA yield.
3. Add 1 µL of DNase (2,000 units/µL) and incubate at 37 °C for 10 min. Do not exceed 10 min, or the enzyme may begin degrading RNA.
4. Add 30 µL of ice-cold 7.5 – 8 M lithium chloride and 30 µL of chilled nuclease-free water to precipitate the RNA. Flick the tube to mix. Allow precipitation of RNA to proceed overnight at -20 °C.
5. Collect RNA by centrifugation at 17,000 x g for 20 min at 4 °C. Wash the pellet once with 1 mL of freshly prepared 75% ethanol in Diethyl pyrocarbonate (DEPC) water, then repeat centrifugation.
6. Remove and discard the supernatant, invert the tube on a clean paper towel, and allow to dry for 10 min. Resuspend the pellet in nuclease-free water. If the pellet is easily visible, resuspend in 25 µL. If the pellet is very small or not visible, resuspend in 15 – 20 µL. Obtain an estimate of RNA concentration by spectrophotometer.
   Note: Typical RNA concentrations range from 200 – 500 ng/µL.

3. Visualization of RNA Probes by RNA Formaldehyde Gel Electrophoresis to Assess RNA Purity

CAUTION: Formaldehyde poses an inhalation hazard; therefore, generate the solutions required for RNA gel analysis in a fume hood. Ethidium bromide is a suspected carcinogen; handle with care.

1. Prepare a 1% formaldehyde agarose gel as previously described²⁴ and cast the gel in a gel box free of RNases. Once cool, fill the gel box with 1x running buffer (1x MOPS at pH 7.0, 5% formaldehyde).
2. Prepare the Sample Buffer (5 µL 400 mg/mL ethidium bromide, 20 µL 10x MOPS, 35 µL formaldehyde, and 100 µL formamide). Combine 2 µL of RNA probe (Step 2.6) with 8 µL of Sample Buffer. Incubate at 70 °C for 10 min.
3. Prepare RNA Loading Buffer by combining 5 mL 50% glycerol, 1 mL 10% formaldehyde, 40 mg bromophenol blue, 40 mg xylene cyanol, and 20 µL of 0.5 M EDTA (pH 7.5). After use, store this buffer at -20 °C.
4. Add 3 µL of Loading Buffer to the RNA sample from Step 3.2 and mix. Keep sample tubes on ice.
5. Load the entire sample volume from Step 3.4 into the gel. Load an aliquot of single-stranded RNA ladder alongside to verify RNA size. Run the gel at 100 V or lower until the blue dye migrates approximately 75% of the way down the gel. Visualize the RNA under UV light using a gel imaging system.  
   Note:  A good quality RNA probe should appear as a discrete band with a mobility corresponding to the expected size of the probe (Figure 2, lane 1). If the probe appears as a diffuse and smeary band (Figure 2, lane 2) this should not be used.

4. Tick Gut Collection and Fixation

1. Collect Ixodes scapularis nymphs that have fed on mice for the time points of interest.
   Note: For the purposes of this technique, ticks fed for 48 or 72 h work best. 
2. Dissect the gut from the tick into MEMFA (Table 1) on a glass slide under a dissection microscope, avoiding damage to the organ as much as possible. Place a tick on a clean microscope slide in 20 – 30 µL of MEMFA. Using a pair of sterile high-carbon steel razor blade #11 slice the head region at the scutum of the tick leaving the body of the tick. Gently press the body to push the gut diverticula and salivary glands out the body cuticle. Separate the salivary glands and the gut and scoop the guts on the razor blade. 
3. Collect guts into a 1.5 mL tube containing 500 µL MEMFA. Incubate the tube at room temperature with gentle rocking for 1 h or overnight at 4 °C.
   Note: Tick salivary glands may also be dissected and similarly fixed and stained.
4. Dehydrate the tissue by washing gently 3 times with 1 mL ice-cold 100% ethanol. Let the guts sink naturally to the bottom of the tube between each wash, as centrifugation may damage the tissue. For long-term storage, keep the samples in 100% ethanol at -20 °C.

5. Construction of Mesh Sample Baskets and 24-well Basket Holder

1. Cut the bottoms off of 2 mL or 5 mL snap-top centrifuge tubes using a heated single-edged razor blade on a scalpel.
   CAUTION: The blade may need to be reheated several times before the cut is completed.
2. Cut squares of a 110 µm nylon mesh slightly larger than the new tube opening. Bond the mesh to the bottom of the tube by pressing them together lightly onto a hot plate on medium-low heat until a good seal is made. Let cool, ensure there are no gaps between the mesh and the tube, and trim excess mesh around the edges. 
3. Cut holes in the cover of a 24-well plate such that the lip of the sample basket made in Step 5.2 will sit at the hole and not fall through, yielding a set-up where the bottoms of the sample baskets will sit all the way in the wells when they are placed in the holes in the cover.
4. Use this fitted basket holder to transfer baskets from one plate to another with relative ease for the entirety of the in situ hybridization protocol. Use two 24-well plates so that while one incubation is occurring, the other plate is prepared for the next wash.

6. In Situ Hybridization: Day 1 (Timing: 4 – 5 h)

1. Transfer guts to labeled sample baskets, separated by experimental condition and intended RNA probe.
   Note: Stain at least 5 guts per condition to account for natural variation among replicates.
2. Rehydrate the samples gently to avoid introducing air bubbles into the tissue by gradually increasing the ratio of phosphate-buffered saline with 0.1 % non-ionic surfactant (PTw; Table 1) to ethanol in the washes.
   Note: Complete all washes in the 24-well basket holder at ambient temperature with gentle agitation unless otherwise noted. Aliquot 500 – 600 µL of wash solutions into each well of the holder such that the guts in each basket are completely submerged. Prepare all solutions with DEPC-treated water to prevent RNA degradation.
   1. Wash the samples for 5 min in 500 µL 100% ethanol.
   2. Prepare at least 500 µL per sample basket of 75%, 50%, and 25% ethanol in PTw. Rehydrate the samples by washing for 5 min in each ethanol solution, moving from the highest ethanol concentration to the lowest.
   3. Wash the samples 4 times for 5 min each in PTw.
3. Permeabilize the samples with Proteinase K (10 µg/mL) in PTw for 5 min.
4. Prepare the tissue for hybridization with the RNA probes.
   1. Wash the samples twice as described in 6.2 in the 24-well basket holder at ambient temperature with gentle agitation for 5 min each in 500 µL triethanolamine buffer (0.1 M, pH 7.0 – 8.0) (Table 1) to maintain the samples in an amine-free environment.
   2. Treat the samples twice with 500 µL 0.25% acetic anhydride in triethanolamine buffer (0.1 M, pH 7.0 – 8.0) for 5 min each, then wash the samples twice in PTw for 5 min each.
      Note: The acetic anhydride acetylates free amines to neutralize positive charge, which is critical to prevent non-specific electrostatic interactions and ensure specific binding of the probe to mRNA.
   3. Fix the guts for 20 min with 500 µL 4% paraformaldehyde in PTw, then wash 5 times in PTw for 5 min each.
   4. Prehybridize by incubating the samples in 500 µL hybridization buffer /in the 24-well basket holder in the 24 well plate (Table 1) for 1.5 – 2.5 h at 60 °C with gentle agitation in a shaking water bath. Save the hybridization buffer used for the prehybridization step to reuse in the Day 2 protocol (Step 7.1).
   5. Prepare probe hybridization solutions by diluting RNA probes in hybridization buffer to a final concentration of 1 µg/mL.  Incubate samples with probe hybridization solutions in the 24-well basket holder at 60 °C with gentle agitation overnight in a shaking water bath (Table 2). Cover the plate carrying the basket holder snugly with aluminum foil to prevent evaporation of the hybridization solution.

7. In Situ Hybridization: Day 2 (Timing: 4.5 – 5.5 h)

1. Remove the samples from the probe hybridization solutions and place them into the reserved hybridization buffer from the previous day. Incubate at 60 °C with gentle agitation for 3 min.
   Note: Probe hybridization solutions may be stored at -20 °C for reuse up to 3 times.
2. Prepare 2x saline-sodium citrate buffer (SSC) by diluting 20x SSC in DEPC-treated water (Table 1). Wash the samples twice for 3 min with 2x SSC, then 3 times for 20 min with 2x SSC at 60 °C to remove all traces of probe and hybridization buffer.
3. Treat with RNase A (20 µg/mL) and RNase T₁ (10 µg/mL) in 2x SSC for 30 min at 37 °C to remove single stranded RNA probe representing free non-hybridized RNA.
4. Wash once with 2x SSC for 10 min at room temperature. Prepare 0.2x SSC by diluting 2x SSC in DEPC-treated water, then wash twice with 0.2x SSC for 30 min at 60 °C to remove excess RNases.
5. Wash twice with 500 µL of maleic acid buffer (MAB; Table 1) for 10 min each.
6. Incubate samples with blocking solution (2% blocking reagent in MAB) for 1.5 – 2 h.
   Note: The blocking reagent can be difficult to dissolve; gentle heating aids dissolution.
7. Replace the blocking solution with anti-digoxigenin alkaline phosphatase-conjugated 500 µL of antibody at a 1:3,000 dilution in blocking solution and incubate at room temperature for about 4 h or at 4 °C overnight.

8. In Situ Hybridization: Day 3 (Timing: 6 h to overnight)

1. Wash the samples at least 5 times for 30 min each with 500 µL MAB to remove excess antibody.
   Note: These washes are flexible and may be extended to 1 – 2 h, or 3 – 4 washes may be substituted for one overnight wash at 4 °C with minimal effect on efficacy.
2. Wash twice for 5 min with alkaline phosphatase buffer, pH 9.5 (AP buffer; Table 1).
3. Begin the color reaction by replacing the last wash with 500 µL of the chromogenic substrate for alkaline phosphatase (Table of Materials). Cover the sample plate with aluminum foil and let the staining proceed at room temperature for 3-  5 h or overnight at 4 °C. Monitor the staining reaction periodically by viewing the tissue under a dissecting microscope to avoid overstaining.
   Note: When the staining is complete, a purple tint is detectable by eye in the antisense-probed samples under the microscope, but the tissue should not be allowed to become very dark. Staining proceeds very slowly at 4 °C and may take up to 20 – 24 h.
4. Stop the color reaction by washing for 5 min in 500 µL MAB, then fix the tissue overnight in Bouin’s solution.
   CAUTION: Handle the Bouin’s solution in a fume hood; it is a corrosive carcinogen.

9. In Situ Hybridization: Day 4 (Timing: 3 – 3.5 h)

1. Prepare at least 7 mL per sample basket of 1x SSC by diluting 20x SSC in DEPC-treated water. Remove the Bouin’s solution by washing the samples 4 to 6 times with 70% ethanol in 1x SSC until the tissue is no longer yellow.
2. Prepare 500 µL per sample basket of 75%, 50%, and 25% ethanol solutions in 1x SSC. Wash the samples for 5 min in each solution, moving from highest ethanol concentration to lowest to rehydrate the tissue. Wash twice for 5 min each in 1x SSC.
3. Incubate the samples in the bleach solution (Table 1) for 90 min on a lightbox in a fume hood.
   Note: This step allows any pigmentation in the samples or coloration from Bouins solution to be removed and enhances the probe signal for clear visualization.
4. Wash the samples twice with 500 µL of 1x SSC for 5 min.
5. Relocate the guts with minimal damage by inverting the sample basket into a well of a new 24-well plate and wash with 70% ethanol through the mesh bottom using a transfer pipette. Store at -20 °C if needed.
6. To obtain an overview of the staining patterns of multiple samples as shown in Figure 3, maintain the guts in 70% ethanol in the 24 well-plate and view with any bright-field microscope. To examine individual guts under a bright-field microscope as shown in Figure 4, Figure 5, and Figure 6, mount the guts in 100% glycerol under coverslips. Use a plastic spatula to gently manipulate the tissue with minimal damage.
   Note: The high viscosity of glycerol helps protect the tissue from damage by compression and allows the careful positioning of diverticula such that the tissue can be viewed optimally at higher magnifications.
7. Capture images on the microscope using microscope specific image capture software (Table of Materials) and export as Tiff images to a generic image editing software. To quantify the relative staining differences between experimental treatments, download a generic image analysis software (Table of Materials). Open the image within the analysis software and use the instructions within the software to measure pixel intensity of the staining and compute histogram plots of the staining.