Spatial High-resolution Analysis of Gene Expression Levels in Tendons

All animal experiments were performed in accordance with the Institutional Animal Care and Use Committee (IACUC) and AAALAC guidelines. Experiments were performed under approved protocol #2013N000062 at Massachusetts General Hospital. In this study, C57BL/J6 mice (5 weeks of age and P0) were used. See the Table of Materials for details related to all materials, reagents, and instruments used in this protocol.

1. Sample preparation and fixation

1. Euthanize mice in a CO₂ chamber followed by cervical dislocation as a secondary method of euthanasia. Then, use scissors to cut the mouse hindlimbs along the hip joint¹². In a scintillation vial, immerse the mouse hindlimb in enough 4% paraformaldehyde (PFA) solution to completely cover the sample, and fix the tissue for 24 h.
   NOTE: Instead of 4% PFA, 10% formalin may also be used for fixation. After fixation, the sample may be stored in phosphate-buffered saline (PBS) until further processing. By fixing the whole hindlimb, we are able to preserve tension on the tendon and have the opportunity to examine the enthesis, tendon midbody, or myotendinous junction.
2. After removing the PFA from the scintillation vial, wash the sample for 3 x 10 min with 1x PBS by adding enough PBS to cover the sample. Then, immerse the hindlimbs in 0.5 M EDTA at 4 °C with agitation for 1-2 weeks, replacing with fresh EDTA every 2-3 days.
3. After removing the EDTA, wash the sample for 3 x 10 min with 1x PBS, dehydrate the samples in 70% EtOH, and embed them in paraffin in the desired orientation. Use conventional FFPE sectioning techniques to section the tissue with a thickness of 3 µm and transfer the sections onto a treated, charged microscope slide (see Table of Materials)¹³.
   NOTE: Sections can be stored in 4 °C prior to proceeding to the next steps.
4. Deparaffinization
   1. Bake the slides on a hot plate at 60 °C for 1 h and then allow the slides to come to room temperature (RT). To remove the paraffin wax from the slides, place the slides on a slide rack and lower them into a staining dish filled with xylene. Repeat in fresh xylene for 3 x 5 min.
      CAUTION: Xylene is hazardous; handle with care.
   2. To rehydrate the sections, immerse the slides in the following for 5 min per wash, all at RT: wash 2x in 100% EtOH, 1x in 75% EtOH diluted in dH₂O, 1x in 50% EtOH diluted in PBS, 1x in 25% EtOH diluted in PBS, and 2x in PBS.
      NOTE: All the rehydration steps are done with the slides on a slide rack and with the various EtOH solutions in staining dishes.

2. RNAscope (commercialized ISH) protocol¹⁴ adaptation

1. Preparing solutions and probes
   1. Prepare TEG solution for the pretreatment of sectioned samples by combining 25 mM of Tris-HCl at pH 8, 10 mM of EDTA, and 50 mM of glucose. Make a sufficient volume for all samples to be treated or prepare excess; filter-sterilize and store at RT; mix the solution properly before using it.
   2. Prepare enough 1x Wash Buffer (see the Table of Materials) for all washes and store it at RT.
   3. In a 40 °C water bath or incubator, warm the probes of interest for 10 min. Then, prepare the probes according to the ISH protocol¹⁴.
2. Pretreatment of samples
   1. Rather than using the recommended pretreatment solution, immerse the slides in a staining dish containing TEG buffer for 4 h at 60 °C.
      NOTE: TEG buffer incubation can be extended for up to 6 h if the tissue is particularly dense or if antigen retrieval of the RNA binding motif of interest is challenging. In that case, a stop point can be added here, the slides dried and kept in a 4 °C fridge overnight.
   2. Remove the samples from the buffer and let them dry. Then, draw around the samples with a hydrophobic barrier pen, immerse them in another staining dish containing protease IV, and incubate them in an oven for 45 min at 40 °C.
      NOTE: We recommend protease IV for adult mice. If using younger mice, such as P0, we recommend using a milder enzyme, such as protease III, for 30 min. When troubleshooting for different ages, observe the integrity of the tendon and check for degradation. If the tissue is dissociating or degrading, decrease the incubation time or change the protease used.
   3. Wash samples again in TEG buffer for 30 min and then hybridize the probes according to the ISH protocol¹⁴.
   4. Add 50-100 µL of mounting reagent on top of the sample. Place a coverslip on top of the samples for microscopy. When imaging, use a high magnification (40x objective or higher) to view the signal.

3. HCR ISH protocol¹⁵ adaptation

1. Postfixation
   1. Prepare 4% PFA (in PBS), PBT solution (0.1% Tween-20 in PBS), and Proteinase K in PBS (stock concentration of 10 mg/mL, final concentration of 5 µg/mL).
   2. To begin the first postfix step, immerse the slides in a staining dish containing 4% PFA for 5 min; then, remove the 4% PFA solution and store it for use in later steps. Rinse the slides for 2 x 5 min in PBT solution and remove the PBT solution. Place the slides in a staining dish containing Proteinase K solution (5 µg/mL) for 5 min and rinse them in the PBT solution 2 x 5 min.
   3. Begin the second postfix step and immerse the slides in the 4% PFA for 5 min.
      NOTE: This is the same 4% PFA saved from step 3.1.2.
   4. Rinse the slides for 3 x 5 min in the PBT solution.
      NOTE: All the postfixation steps are done with the slides on a slide rack and in staining dishes containing the respective solutions.
2. Acetylation
   1. Prepare the acetylation solution containing 625 µL of acetic anhydride, 3.3 mL of 1 M triethanolamine (TEA) buffer, and 246 mL of ddH₂O. Be sure to mix well and use immediately after preparation.
   2. Place the slides in the acetylation solution for 10 minutes and then rinse 3 x 5 min in PBT solution. Rinse the slides in ddH₂O and let them air dry for 30 min. Then, draw around the samples with a hydrophobic barrier pen.
      NOTE: All the acetylation steps are done with the slides on a slide rack and in staining dishes containing the respective solutions
3. Hybridization
   1. Preheat the hybridization buffer to 37 °C and prewarm a humidified chamber to 37 °C.
      CAUTION: The buffer contains formamide, a hazardous material. If the slides are not sufficiently dry, blot the edges with a laboratory wipe to remove excess dH₂O.
   2. Prepare probe solutions by adding 0.4 pmol of each probe mixture to 100 µL of hybridization buffer.
      NOTE: If using dHCR imaging, use a higher concentration of probe to improve probe hybridization efficiency.
   3. Prepare 4 L of Sodium Chloride Sodium Citrate Buffer (20x SSC) by combining 3 M NaCl (701.1 g of NaCl in a final volume of 4 L) and 0.3 M of Na₃CH₆H₅O₇•2H₂O (352.8 g) and then adjusting the final pH to 7 by adding HCl or 10 N NaOH. Add enough H₂O to reach 4 L. Prepare 5x SSCT by diluting 20x SSC to 5x and adding enough 10% Tween 20 so that it is 0.1% of the total volume.
   4. Add 200 µL of hybridization buffer to the sample, place the slides in a humidified chamber, and let the slides incubate for 10 min. Remove the hybridization solution and drain the excess buffer on the slide by blotting the edges with a laboratory wipe.
   5. Add 100 µL of the prepared probe solution on top of the sample and place a coverslip on the sample. Incubate in an oven overnight or for around 12-16 h in a chamber humidified with a solution containing 5x SSC and 50% formamide. Set the temperature to 37 °C.
      NOTE: Since the incubation is overnight, the addition of the coverslip is to minimize evaporation.
   6. Leave an aliquot of 5x SSCT solution in a 37 °C water bath and a second aliquot at RT. Anticipate the volumes of these aliquots needed based on the sample number and the number of washes in the next step.
4. Washing
   1. Using the previously preheated 5x SSCT solution, prepare 75% wash buffer/25% 5x SSCT, 50% wash buffer/50% 5x SSCT, and 25% wash buffer/75% 5x SSCT solutions.
   2. To remove excess probes, immerse the slides in series for 15 min per wash at 37 °C in 75% wash buffer/25% 5x SSCT, 50% wash buffer/50% 5x SSCT, and 25% wash buffer/75% 5x SSCT. Allow the coverslips to float off from the samples.
   3. Incubate the slides 2 x 15 min with 100% 5x SSCT and then immerse them in 5x SSCT for 5 min at RT.
      NOTE: The washing steps can be done with the slides on a slide rack and in staining dishes containing the respective solutions or with the solutions added as droplets.
5. Amplification
   1. Prepare 6 pmol of hairpin h1 and 6 pmol of hairpin h2 by snap cooling 2 µL of a 3 µM stock (heat at 95 °C for 90 s and cool to RT). Protect the hairpins from light during this process.
      NOTE: HCR hairpins h1 and h2 are provided in a hairpin storage buffer and are ready for snap cooling. Snap cool h1 and h2 hairpins in separate tubes.
   2. Dry the slides by blotting their edges with a laboratory wipe. Then, add 200 µL of amplification buffer on top of the sample and place it in a humidified chamber for 30 min at RT. Prepare the hairpin mixture by adding snapcooled h1 hairpins and snapcooled h2 hairpins to 100 µL of amplification buffer at RT.
   3. Remove the amplification buffer and drain the excess liquid on the slide by blotting the edges with a laboratory wipe. Add 100 µL of the previously prepared hairpin mixture and place parafilm on top of the sample. Incubate the slides for a minimum of 4 h or overnight in a dark, humidified chamber at RT.
   4. To remove excess hairpins, immerse the slides in 5x SSCT in a staining dish and incubate at RT for 30 min. Repeat the wash in fresh 5x SSCT for 30 min, and a third time for 5 min.
   5. Dry the slides by blotting the edges with a laboratory wipe and add 50-100 µL of mounting reagent on top of the sample. Place a coverslip on top of the samples for microscopy. When imaging, use a high magnification (40x objective or higher) to view the signal.