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

Summary

This study presents a surgical manipulation to expose the T-DRG in anesthetized mice for in vivo calcium imaging, along with synchronous ECG recordings. This method represents a cutting-edge tool for studying peripheral electric nerve stimulation and thoracic visceral organ inputs, as well as their interactions at the primary sensory level.

Abstract

The dorsal root ganglia (DRG), housing primary sensory neurons, transmit somatosensory and visceral afferent inputs to the dorsal horn of the spinal cord. They play a pivotal role in both physiological and pathological states, including neuropathic and visceral pain. In vivo calcium imaging of DRG enables real-time observation of calcium transients in single units or neuron ensembles. Accumulating evidence indicates that DRG neuronal activities induced by somatic stimulation significantly affect autonomic and visceral functions. While lumbar DRG calcium imaging has been extensively studied, thoracic segment DRG calcium imaging has been less explored due to surgical exposure and stereotaxic fixation challenges. Here, we utilized in vivo calcium imaging at the thoracic1 dorsal root ganglion (T1-DRG) to investigate changes in neuronal activity resulting from somatic stimulations of the forelimb. This approach is crucial for understanding the somato-cardiac reflex triggered by peripheral nerve stimulations (PENS), such as acupuncture. Notably, synchronization of cardiac function was observed and measured by electrocardiogram (ECG), with T-DRG neuronal activities, potentially establishing a novel paradigm for somato-visceral reflex in the thoracic segments.

Introduction

Dorsal root ganglia (DRG) neurons process afferent sensory information from both somatic and visceral receptors. Regulation of cardiac function involves not only primary sensory afferents from viscera but also somatosensory neurons within the same thoracic DRG segment (T-DRG). Recently published research in 'Circulation' has indicated that T-DRG plays a role in cardiac function regulation. Blocking Piezo1/IL-6 in T-DRG inhibited IL-6/STAT3 inflammatory signaling, thereby attenuating ventricular remodeling post-myocardial infarction (MI)¹. Additionally, Cui et al.² found in rats with MI that sympathetic sprouting and sympatho-sensory coupling occurred in T1-5 DRGs and upper limb skin, contributing to cardiac-related referred pain. Somatic stimulation in the referred pain area increased sympathetic discharge and regulated cardiac function. However, due to technical limitations, changes in T-DRG from cardiac and somatic inputs pre- and post-MI or somatic stimulation were observed only after animal sacrifice. Therefore, observing neural activities within T-DRG is crucial for understanding its intricate relationship with visceral function alterations.

In recent years, advancements in sensitive genetically encoded calcium indicators (GECIs)³, along with confocal microscopy and multi-photon imaging technology, have enabled scientists not only to describe neuronal activity and diameter, but also to combine genetic labeling techniques such as Pirt⁴ and neural tracing to observe specific neurons during imaging⁵. This integrated approach aids in uncovering deeper scientific principles. However, until recently, methods for studying in vivo calcium imaging of DRG have been predominantly limited to lumbar segments⁶.

The T-DRG in mice are relatively small and circular, located anterior and medial to the intervertebral foramen, with a total of 13 pairs. Due to the physiological curvature of the thoracic vertebrae, the space between adjacent thoracic vertebral segments, especially T1-5, is very narrow, increasing the difficulty of exposure. Fixation presents another challenge for stable calcium imaging of DRG in a single plane. Each side of the T-DRG is adjacent to two rib surfaces, and each rib surface of the vertebra is attached to the corresponding rib, further complicating the already cramped space. Building upon early calcium imaging methods for lumbar DRG⁷, this research team has developed an in vivo calcium imaging method for T-DRG using a custom-made spinal clamp. Additionally, numerous studies have demonstrated that peripheral electric nerve stimulation, such as acupuncture, can induce DRG neural activity and regulate visceral function^8,9,10. To better understand the relationship between DRG and acupuncture-mediated cardiac function regulation, synchronous recording of cardiac function during DRG calcium imaging has been implemented, offering novel research insights into how somatic stimulation-induced neuronal activity in T-DRG influences visceral function regulation.

This unique research details the exposure and fixation of mouse T-DRG, ensuring stable calcium imaging alongside cardiac function recording. This provides a cutting-edge scientific tool for studying the peripheral mechanisms of thoracic visceral functional changes and further exploring visceral-somatic afferent inputs.

Protocol

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 (5% isoflurane). 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 firmly pinching the mouse’s hind foot to ensure there is no response.
   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 ophthalmic 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 due to the nerve stimulation (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 (0.5%-1% isoflurane) 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.

Results

Following the described protocol, the T1-DRG of transgenic Pirt-GCaMP6s mice were exposed to various somatic stimuli. The aim of this experiment was to observe changes in the number and type of neurons and cardiac function induced by different stimuli.

As depicted in Figure 2A, under baseline conditions, most neurons in the T1-DRG did not exhibit GFP fluorescence. This baseline fluorescence could be influenced by two factors: the GCaMP expression level a...

Discussion

In this study, a method for calcium imaging of the thoracic segment T1 DRG is described, which has significant value for studying the afferent transmission of cardiopulmonary visceral sensory neurons and somato-visceral communication. Additionally, a general approach is presented for monitoring calcium activity in DRG neurons and changes in cardiac function simultaneously, enabling correlation analysis of neural activity and cardiac responses.

Calcium imaging techniques, using Ca²⁺ ...

Materials

Name	Company	Catalog Number	Comments
Acupuncture Needle	ZhongYanTianHe	0.25/13s
Anesthesia System	Kent Scientific	SomnoSuite
Animal Bio Amp	ADInstruments	NSW
Confocal Microscope	Leica	STELLARIS 8
DC Temperature Controller	FHC	40-90-8D
DC Temperature Controller Heating Pad	FHC	40-90-2-05
Fiji	National Institute of Health	N/A
Fine Forceps	RWD	F11028-13
Fine Ophthalmic Forceps	Jinzhong	JD1060
Gelatin Sponges	Coltene	274-007
Intubation Cannula	Harward Apparatus	73-2737
Isoflurane	RWD	R510
LabChart Professional Software	ADInstruments	Version 8.0
LAS X	Leica	N/A
Pirt-cre mice	Johns Hopkins University	N/A
Retractor	Fine Science Tools	16G212
Rosa-GCaMP6s  mice (AI96)	Jax Laboratory	28866
Spinal Clamp	N/A	N/A	Custom made
Spring Scissors	Jinzhong	YBC040
Stimulator	AMPI	Master-8
Tribromoethanol	Sigma	T48402
Wireless Biological Acquisition System	Kardiotek Biomedical Technologie	KLB-1