In Vivo Thoracic Dorsal Root Ganglia (DRG) Calcium Imaging and ECG Recording for Studying Peripheral Nerve Stimulation

All procedures followed the guidelines of the National Institutes of Health for the care and use of laboratory animals and were approved by the Animal Care and Use Ethics Committee of the Institute of Acupuncture & Moxibustion, China Academy of Chinese Medical Sciences. For in vivo calcium imaging of DRG neurons, adult Pirt-GCaMP6s mice (20-25 g, both sexes) were used. These mice were generated by crossing Pirt-Cre mice with Rosa26-loxP-STOP-loxP-GCaMP6s mice (see Table of Materials). They were housed in the animal facility of the Institute of Acupuncture and Moxibustion, China Academy of Chinese Medical Sciences, under controlled conditions (room temperature: 22-24 °C, humidity: 50%-60%, 12-h light-dark cycle), with ad libitum access to food and water. The animals underwent one week of acclimatization before experimentation. During experiments, animals were anesthetized, maintained at a warm temperature, and monitored for pain response to ensure adequate anesthesia depth. After the experiments, animals were euthanized under deep anesthesia (following institutionally approved protocols). The details of the reagents and the equipment used are listed in the Table of Materials.

1. Preoperative preparation

1. Verify the proper functioning of the anesthesia ventilation device, imaging system, and electrocardiogram monitor and activate them sequentially.
2. Administer an intraperitoneal injection of 1.25% tribromoethanol (0.2 mL per 10 g of body weight) to anesthetize the mouse). Assess the depth of anesthesia by pinching the mouse's paw pad.
   1. If signs of pain occur during the surgical procedure, administer an additional intraperitoneal injection of 0.2 mL of tribromoethanol solution, or use an anesthesia machine to deliver 0.5%-1% isoflurane to prevent muscle spasms and ensure adequate anesthesia depth.
3. Remove the hair from the anterior neck and posterior back of the mice.
4. Position the mouse in a supine position on a warming blanket and apply ointment to the mouse's eyes to prevent drying.

2. Tracheotomy

1. Disinfect the neck skin with iodine. Make a 1 cm vertical incision in the center of the skin anterior to the trachea.
2. Expand the skin incision and displace the glands in front of the trachea. Expose the digastric muscle, bluntly separate it, and expose the trachea. Gently separate the trachea from surrounding tissues.
3. Use spring scissors to create a transverse incision in the trachea. Insert an endotracheal tube into the trachea, directing it towards the lungs (Figure 1A). Secure the trachea with 3-0 surgical suture to prevent air leakage and accidental removal of the catheter.
4. Replace the paratracheal muscles and glands over the trachea. Close the neck skin using 6-0 sutures.

3. Exposure of thoracic vertebrae

1. Put the mouse in a prone position on a heated pad. Make a 2 cm longitudinal incision in the center of the nape of the neck, extending from the C6 to T3 vertebrae.
2. Carefully separate the fat and hibernating glands at the anterior aspect of the mouse's thoracic vertebrae, taking care to avoid the blood vessels beneath the glands. Use spring scissors to cut through the skin and muscle layers, including the trapezius muscle. Insert a retractor between the muscles to assist with further exposure.
3. Remove the muscles attaching to the head clamp and the straight portion of the long neck muscles, exposing the spinous processes of T2.
4. Displace the semispinalis and spinalis muscles to expose the vertebral arch from C6 to T3 (see Figure 1B).
   NOTE: During the process of exposing the thoracic vertebrae, ensure special attention is given to preserving the spinous process of T2, as it serves as the primary point of force application for subsequent exposure of T1-DRG.

4. Exposure of T1-DRG

1. Sever the connection between the vertebral arch plate and the articular process of T1. Use fine forceps to meticulously remove the left and right articular processes and mammillary processes of T1.
2. After clearing away the overlying connective tissue, carefully expose either the left or right T1 DRG, ensuring the epineurium's integrity on the chosen side.
3. Place a small cotton ball soaked in warm saline over the exposed DRG area to maintain moisture (see Figure 1C).
   NOTE: Surrounding tissues of the DRG can be covered with a hemostatic sponge to prevent blood leakage during surgery. Change moistened cotton balls as needed to maintain a clear view of the surgical area when soaked with hemorrhagic effusions.

5. Thoracic vertebrae fixation and ECG detection

1. Place the mouse on the stage of the custom spinal clamp with a heating pad (see Figure 1D).
2. Secure the mouse using two clips attached to the articular processes of C6 and T3 to minimize movement during the experiment (see Figure 1E).
3. Connect the ECG monitor with the negative pole on the right upper limb, the ground wire on the right lower limb, and the positive pole on the left lower limb (see Figure 1F).
4. Continuously replace the cotton balls soaked with saline until the surgical area is sufficiently clean.
   NOTE: Ensure thorough clearance of muscles and tissues surrounding the articular processes of C6 and T3, especially in the region where the custom spinal clamp is applied for spinal fixation, to prevent displacement during subsequent experiments.

6. Hardware and software setup for imaging

1. Position the spinal clamp with the secured mouse under the confocal microscope (see Figure 1G).
2. Connect the respiratory anesthesia machine and heating pad and adjust relevant parameters based on the mouse's physical condition to ensure animal welfare.
3. Place the 10x/0.32 long working distance air objective of the confocal microscope over the exposed T1-DRG for imaging (see Figure 1H).
4. Capture the entire T1-DRG by adjusting the stage z-axis up and down, with a step size of 25 µm and a resolution of 512 x 512 or 1024 x 1024 pixels.
5. Perform XYZT scanning of the DRG with 8 stacks comprising 1-2 baseline states, 3-6 brush or PENS stimuli, and 7-8 post-stimuli states.
   NOTE: To ensure a comprehensive and clear view of DRG morphology, position the DRG perpendicular to the lens. Adjust the tilt angle of each mouse using the customized stage according to practical requirements.

7. Somatic stimuli and imaging

1. Apply brush stimulation to the upper limb of the mouse to assess the responsiveness of the imaged DRG neurons (see Figure 1H).
2. Perform PENS-PC6 stimulation using a stimulator (see Figure 1F).
   NOTE: PC6 is located 1-2 mm proximal to the palmar crease of the wrist, between the ulnar flexor carpi tendon and the superficial flexor digitorum tendon. The stimulation parameters are set to 15 Hz and 1 mA for 3-6 stacks.

8. Data analysis and processing

1. In the confocal microscopy software, assess the calcium responses evoked by various stimuli by comparing the increase in green fluorescence intensity when neurons are stimulated by GCaMP upon calcium binding inside the cell (see Figure 2A-C).
2. Use Fiji software to manually trace the visible cells and measure their size and relative fluorescence intensity (see Figure 2D).
3. Express fluorescence intensity as the ratio of the maximum evoked fluorescence increase to the baseline level (Ft/F0), where the baseline (F0) represents the maximum fluorescence intensity measured during the baseline period.
4. Define cell activation as an increase in fluorescence intensity (Ft/F0) ≥130% of F0 (see Figure 2E,F).
5. Classify DRG neurons into small-sized neurons (<20 µm), medium-sized neurons (20-30 µm), and large-sized neurons (>30 µm) based on their diameter¹¹ (see Figure 2G) .
6. Observe changes in heart rate caused by somatic stimulation using the ECG recording software (see Figure 2H).
   NOTE: The heart rate of mice is easily influenced by external stimuli and respiratory anesthesia. When observing the heart rate response to a stimulus, wait for the heart rate to stabilize before administering the stimulus.