In Vivo Calcium Imaging of Dorsal Root Ganglia Neurons’ Response to Somatic and Visceral Stimuli

This animal protocol was approved by the Animal Care and Use Ethics Committees of the Institute of Acupuncture and Moxibustion, China Academy of Chinese Medical Sciences, and complied with the National Institutes of Health Guide for the Care and Use of Experimental Animals to ensure minimal animal use and discomfort. Pirt-cre mice were kindly donated by Dr. Xinzhong Dong from Johns Hopkins University (Baltimore, MD). Rosa26-loxP-STOP-loxP-GCaMP6s mice were obtained from a commercial source (see Table of Materials). The following procedure is optimized for either gender of mice weighing 20-35 g. The recommended age for mice undergoing the operation was between 8 and 14 weeks. Either genetically encoded calcium indicators (GECIs), such as Pirt-GCaMP6s mice, or normal C57 mice intrathecally injected with viral GECIs, were adopted to show the main populations of DRG neurons. Genetically engineered mice intrathecally injected with viral GECIs or crossed with Rosa26-GCaMP mice would show the specific population of DRG neurons. Laboratory gowns, gloves, and masks were worn throughout the protocol. After the experiment, the mice were euthanized with CO₂ inhalation followed by cervical dislocation.

1. Pre-operative setup

1. Verify that the anesthesia ventilation device (see Table of Materials) has been properly installed.
2. Ensure that the imaging system (see Table of Materials) has been pre-opened and is functioning correctly.
   ​NOTE: The operation and imaging procedures were conducted sequentially, while the analysis process can be completed offline later.

2. Anesthetization

1. Intraperitoneally inject the mouse with a tribromoethanol solution (1.25%, 250 mg/kg, see Table of Materials)^13,14,15. Assess the depth of anesthesia via toe pinch before proceeding.
2. Shave the hair in the lower back area and the ventral cervical part.
3. Place the mouse in a supine position on the heating pad (see Table of Materials).
4. Apply ointment to the animal's eyes to prevent them from drying out.
   ​NOTE: The depth of anesthesia was maintained as long as the mouse showed no response to a hindpaw pinch during all surgical and recording procedures. If the animal exhibited any signs of pain, an additional 0.2 mL of tribromoethanol solution was administered intraperitoneally, or 0.5% isoflurane was given intratracheally after the ventilator was applied to prevent muscle twitching and ensure the depth of anesthesia.

3. Tracheotomy

1. Make a midline incision of approximately 2 cm in the skin of the throat, using a surgical blade, from the sternum to the chin.
2. Extend the incision and gently push aside the sub-maxillary glands, ensuring full exposure of the digastric muscles located beneath the glands, running longitudinally along the middle of the trachea.
3. Once the seam of the paratracheal musculature in the middle of the muscles is seen, expose the trachea by gently separating the musculature and retracting it open.
4. Create a transverse incision on the trachea, large enough to accommodate the trachea cannula (outer diameter 1.3 mm, length 13 mm, with a Y-adapter, see Table of Materials) directed toward the lung (Figure 1A).
5. Cover the trachea with the paratracheal muscles and sub-maxillary glands, then suture the surrounding tissues to the skin layer by layer using 5-0 surgical sutures.

4. Exposure of lumbar vertebrae

1. Place the animal in a prone position on the heating pad.
2. Create a midline incision of approximately 7 cm in the lower back, extending from lumbar-5 to sacral-1 vertebra.
3. Carefully expose the spinous processes of the three vertebrae by moving aside the longissimus lumborum muscles.
4. Expose the successive three articular processes and mamillary processes, with the recording segment in the middle of the vertebrae, by moving aside the inner parts of the bilateral longissimus lumborum, dorsal cervical spine muscles, and dorsal cervical semispine muscles (Figure 1B).
   ​NOTE: The articular processes and mamillary processes of the three vertebrae were exposed to allow the spinal clamp to secure the animal without displacement during recording.

5. Exposure of dorsal root ganglion

1. Securely immobilize the animal with a toothed forceps (see Table of Materials) on the bilateral articular processes of the neighboring vertebrae. Then, use rongeur forceps to remove either the left or right articular and mamillary processes of the lumbar-6 vertebrae.
2. Carefully expose the left or right lumbar-6 DRG after clearing away the connective tissues above it, keeping the intact epineurium on the left or right side of the lumbar-6 vertebrae.
3. Ensure the active conditions of the exposed DRG by covering the exposed areas with small cotton balls soaked in saline.
   ​NOTE: Change the cotton balls until there is no subtle leakage of blood near the exposed DRG and the surrounding tissue, as this could affect the clear imaging of the DRG neurons. Replace the saline-wetted small cotton balls until there is no red or yellow leakage of tissue fluids.

6. Immobilization and ventilation

1. Place the animal with the exposed lumbar-6 DRG onto a heating pad on the stage of a customized spinal clamp^16,17 (Figure 1C).
2. Secure the animal by using two clamps on the spinal columns of the neighboring vertebrae to minimize movement during testing (Figure 1D).
3. Continue to change the saline-wetted cotton balls until the surgical area is sufficiently clean (Figure 1E).
4. Monitor ventilation and pulse oximetry throughout the experiment.
   ​NOTE: Immobilize the spinal columns using the customized spinal clamp and then adjust the pad's angle so that the microscope's objective is perpendicular to the DRG. Make necessary adjustments to align the objective and the DRG using angle adjusters.

7. Hardware and software setup for imaging

1. Insert the customized spinal clamp into the custom-designed microscope stage (Figure 1F,G).
2. Position a 10x/0.32 long-working distance air objective lens of a confocal microscope (see Table of Materials) over the exposed DRG for imaging (Figure 1G).
3. Capture time-lapse z-stacks of the intact L-6 DRG with a 25 µm/step and either 512 x 512 or 1024 x 1024 pixel resolution. The duration of one stack scan of the entire DRG varies from 5-15 s based on factors such as the perpendicularity of the DRG's position to the objective lens, the depth and speed of each scanning step, and the imaging resolution.
4. Conduct XYZT scanning of the DRG with 4-8 stacks, which include 1-3 baseline states, 2-3 acupuncture or visceral colorectal distension (CRD) stimuli, and 1-2 post-stimuli states.

8. Acupuncture/visceral stimuli and imaging

1. Apply brushing or pinching stimuli to the lower back, hindlimb, or hindpaw areas of the animal to assess the most sensitive receptive field of the imaged DRG neurons (Figure 1H).
2. Administer acupuncture stimuli manually or with a commercially available acupoint and nerve stimulator (Figure 1F) (see Table of Materials).
3. Induce visceral stimuli by using colorectal distension (CRD) with a self-made air pressure gauge (Figure 1H).
   ​NOTE: Test the most sensitive receptive field of the imaged DRG neurons by brushing or pinching the receptive areas of the lumbar-6 DRG. If these stimuli cause fluorescence changes during XYZT stack recording, select acupoints in these areas, such as the lower back acupoint BL25 (Dachangshu)¹⁸. Choose acupoints (as shown in Figure 2) that have the skin and muscle within the receptive field of lumbar-6 innervation.

9. Data analysis and processing

1. Assess evoked calcium responses using a confocal microscope by measuring the increase in green fluorescence of the neuronal GCaMP upon binding to intracellular calcium following somatic or visceral stimuli.
2. Manually trace visible cells and determine cell size and relative fluorescence intensity using the imaging software (Fiji, see Table of Materials). Define small, medium, and large neurons as having soma diameters of <20 µm, 20 to 30 µm, and >30 µm, respectively, based on previous studies^17,19.
3. Express fluorescence intensity as a ratio of the increase in maximum evoked fluorescence to the basal level (ΔF/F0) or a ratio of the maximum evoked fluorescence to the basal level (Ft/F0), where F0 represents the maximum fluorescence intensity measured during the baseline period.
4. Measure the percentage of activated neurons and the intensity of evoked calcium transients to assess changes in neuronal responsiveness following somatic or CRD stimuli.
5. Define activation of a cell as an increase in fluorescence intensity (ΔF) ≥30% of the baseline (F0).
   NOTE: Before tracing visible cells, adjust the brightness of the images to the maximum level to aid in identifying cell borders. Do not trace cells if there is any uncertainty. Review the recorded movie forward and backward to ensure no cells are overlooked.