This study was funded by the National Natural Science Foundation of China (No. 82174518, 82074561, 82105029).

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 &#38; 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 &#176;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
<ol>
	<li>Verify the proper functioning of the anesthesia ventilation device, imaging system, and electrocardiogram monitor and activate them sequentially.</li>
	<li>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&#8217;s hind foot to ensure there is no response.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Remove the hair from the anterior neck and posterior back of the mice.</li>
	<li>Position the mouse in a supine position on a warming blanket and apply ophthalmic ointment to the mouse&#8217;s eyes to prevent drying.</li>
</ol>
2. Tracheotomy
<ol>
	<li>Disinfect the neck skin with iodine. Make a 1 cm vertical incision in the center of the skin anterior to the trachea.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Replace the paratracheal muscles and glands over the trachea. Close the neck skin using 6-0 sutures.</li>
</ol>
3. Exposure of thoracic vertebrae
<ol>
	<li>Put&#160;the mouse in a prone position on a heated pad. Make a 2&#160;cm longitudinal incision in the center of the nape of the neck, extending from the C6 to T3 vertebrae.</li>
	<li>Carefully separate the fat and hibernating glands at the anterior aspect of the mouse&#39;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.</li>
	<li>Remove the muscles attaching to the head clamp and the straight portion of the long neck muscles, exposing the spinous processes of T2.</li>
	<li>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.</li>
</ol>
4. Exposure of T1-DRG
<ol>
	<li>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.</li>
	<li>After clearing away the overlying connective tissue, carefully expose either the left or right T1 DRG, ensuring the epineurium&#39;s integrity on the chosen side.</li>
	<li>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.</li>
</ol>
5. Thoracic vertebrae fixation and ECG detection
<ol>
	<li>Place the mouse on the stage of the custom spinal clamp with a heating pad (see Figure 1D).</li>
	<li>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).</li>
	<li>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).</li>
	<li>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.</li>
</ol>
6. Hardware and software setup for imaging
<ol>
	<li>Position the spinal clamp with the secured mouse under the confocal microscope (see Figure 1G).</li>
	<li>Connect the respiratory anesthesia (0.5%-1% isoflurane) machine and heating pad and adjust relevant parameters based on the mouse&#39;s physical condition to ensure animal welfare.</li>
	<li>Place the 10x/0.32 long working distance air objective of the confocal microscope over the exposed T1-DRG for imaging (see Figure 1H).</li>
	<li>Capture the entire T1-DRG by adjusting the stage z-axis up and down, with a step size of 25 &#181;m and a resolution of 512 x 512 or 1024 x 1024 pixels.</li>
	<li>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.</li>
</ol>
7. Somatic stimuli and imaging
<ol>
	<li>Apply brush stimulation to the upper limb of the mouse to assess the responsiveness of the imaged DRG neurons (see Figure 1H).</li>
	<li>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.</li>
</ol>
8. Data analysis and processing
<ol>
	<li>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).</li>
	<li>Use Fiji software to manually trace the visible cells and measure their size and relative fluorescence intensity (see Figure 2D).</li>
	<li>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.</li>
	<li>Define cell activation as an increase in fluorescence intensity (Ft/F0) &#8805;130% of F0 (see Figure 2E,F).</li>
	<li>Classify DRG neurons into small-sized neurons (&#60;20 &#181;m), medium-sized neurons (20-30 &#181;m), and large-sized neurons (&#62;30 &#181;m) based on their diameter11 (see Figure 2G) .</li>
	<li>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.</li>
</ol>

Calcium Imaging of Thoracic Segment T1 DRG for Cardiopulmonary Sensory Neurons

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The authors declare no conflicts of interest.

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+ ...

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 &#39;Circulation&#39; 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 function8,9,10. 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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acupuncture Needle</td>
			<td>ZhongYanTianHe</td>
			<td>0.25/13s</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia System&#160;</td>
			<td>Kent Scientific</td>
			<td>SomnoSuite</td>
			<td></td>
		</tr>
		<tr>
			<td>Animal Bio Amp</td>
			<td>ADInstruments</td>
			<td>NSW</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Leica</td>
			<td>STELLARIS 8</td>
			<td></td>
		</tr>
		<tr>
			<td>DC Temperature Controller</td>
			<td>FHC</td>
			<td>40-90-8D</td>
			<td></td>
		</tr>
		<tr>
			<td>DC Temperature Controller Heating Pad</td>
			<td>FHC</td>
			<td>40-90-2-05</td>
			<td></td>
		</tr>
		<tr>
			<td>Fiji</td>
			<td>National Institute of Health</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps</td>
			<td>RWD</td>
			<td>F11028-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Ophthalmic Forceps&#160;</td>
			<td>Jinzhong</td>
			<td>JD1060</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin Sponges</td>
			<td>Coltene</td>
			<td>274-007</td>
			<td></td>
		</tr>
		<tr>
			<td>Intubation Cannula</td>
			<td>Harward Apparatus</td>
			<td>73-2737</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>RWD</td>
			<td>R510</td>
			<td></td>
		</tr>
		<tr>
			<td>LabChart Professional Software</td>
			<td>ADInstruments</td>
			<td>Version 8.0</td>
			<td></td>
		</tr>
		<tr>
			<td>LAS X</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Pirt-cre mice</td>
			<td>Johns Hopkins University</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractor</td>
			<td>Fine Science Tools</td>
			<td>16G212</td>
			<td></td>
		</tr>
		<tr>
			<td>Rosa-GCaMP6s&#160; mice (AI96)</td>
			<td>Jax Laboratory</td>
			<td>28866</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinal Clamp</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Custom made</td>
		</tr>
		<tr>
			<td>Spring Scissors</td>
			<td>Jinzhong</td>
			<td>YBC040</td>
			<td></td>
		</tr>
		<tr>
			<td>Stimulator</td>
			<td>AMPI&#160;</td>
			<td>Master-8&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Tribromoethanol</td>
			<td>Sigma</td>
			<td>T48402</td>
			<td></td>
		</tr>
		<tr>
			<td>Wireless Biological Acquisition System</td>
			<td>Kardiotek Biomedical Technologie</td>
			<td>KLB-1</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo thoracic dorsal root ganglia (drg) calcium imaging and ecg recording for studying peripheral nerve stimulation

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.

Following the described protocol, the T1-DRG of transgenic&#160;Pirt-GCaMP6s&#160;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...

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

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.

The dorsal root ganglia (DRG), housing primary sensory neurons, transmit somatosensory and visceral afferent inputs to the dorsal horn of the spinal cord. ...

in-vivo-thoracic-dorsal-root-ganglia-drg-calcium-imaging-ecg

Research

JoVE Journal

Neuroscience

592 Views.  China Academy of Chinese Medical Sciences. 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.

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

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after injury (Teng et al., 2006). It is recognized that transcriptional programs essential for neurite and axonal outgrowth are 
reactivated upon injury in the PNS (Makwana et al., 2005). However the tools available to analyze neuronal gene regulation in vivo are limited and 
often challenging.
The dorsal root ganglia (DRG) offer an excellent injury model system because both the CNS and PNS are innervated by a 
bifurcated axon originating from the same soma. The ganglia represent a discrete collection of cell bodies where all transcriptional events occur, 
and thus provide a clearly defined region of transcriptional activity that can be easily and reproducibly removed from the animal. Injury of nerve 
fibers in the PNS (e.g. sciatic nerve), where axonal regeneration does occur, should reveal a set of transcriptional programs that are distinct from 
those responding to a similar injury in the CNS, where regeneration does not take place (e.g. spinal cord). Sites for transcription factor binding, 
histone and DNA modification resulting from injury to either PNS or CNS can be characterized using chromatin immunoprecipitation (ChIP). 
Here, we describe a ChIP protocol using fixed mouse DRG tissue following axonal injury. This powerful combination provides a means for characterizing the pro-regeneration chromatin environment necessary for promoting axonal regeneration.

We present a method for chromatin immunoprecipitation from dorsal root ganglia tissue following axonal injury. The approach can be used to identify specific transcription factor binding sites and epigenetic modification of histone and DNA important for the regeneration of injured axons in both the peripheral and central nervous system.

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of primary afferent fibers mediated by GABAergic axo-axonal synapses (primary afferent depolarization). The strength of primary afferent depolarization can be measured by recording of volume-conducted potentials at the dorsal root (dorsal root potentials, DRP). Pathological changes of presynaptic inhibition are crucial in the abnormal central processing of certain pain conditions and in some disorders of motor hyperexcitability. Here, we describe a method of recording DRP in vivo in mice. The preparation of spinal cord dorsal roots in the anesthetized animal and the recording procedure using suction electrodes are explained. This method allows measuring GABAergic DRP and thereby estimating spinal presynaptic inhibition in the living mouse. In combination with transgenic mouse models, DRP recording may serve as a powerful tool to investigate disease-associated spinal pathophysiology. In vivo recording has several advantages compared to ex vivo isolated spinal cord preparations, e.g. the possibility of simultaneous recording or manipulation of supraspinal networks and induction of DRP by stimulation of peripheral nerves.

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

GABAergic presynaptic inhibition is a powerful inhibitory mechanism in the spinal cord important for motor and sensory signal integration in spinal cord networks. Underlying primary afferent depolarization can be measured by recording of dorsal root potentials (DRP). Here we demonstrate a method of in vivo recording of DRP in mice.

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of ...

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the study of nerve regeneration and myelination. The glial cells of the peripheral nervous system, Schwann cells (SC), are key facilitators of these processes; it is therefore crucial that the interactions of these cellular components are studied together. Direct contact between DRG neurons and glial cells provides additional stimuli sensed by specific membrane receptors, further improving the neuronal response. SC release growth factors and proteins in the culture medium, which enhance neuron survival and stimulate neurite sprouting and extension. However, SC require long proliferation time to be used for tissue engineering applications and the sacrifice of an healthy nerve for their sourcing. Adipose-derived stem cells (ASC) differentiated into SC phenotype are a valid alternative to SC for the set-up of a co-culture model with DRG neurons to study nerve regeneration. The present work presents a detailed and reproducible step-by-step protocol to harvest both DRG neurons and ASC from adult rats; to differentiate ASC towards a SC phenotype; and combines the two cell types in a direct co-culture system to investigate the interplay between neurons and SC in the peripheral nervous system. This tool has great potential in the optimization of tissue-engineered constructs for peripheral nerve repair.

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) are structures containing the sensory neurons of the peripheral nervous system. When dissociated, they can be co-cultured with SC-like adipose-derived stem cells (ASC), providing a valuable model to study in vitro nerve regeneration and myelination, mimicking the in vivo environment at the injury site.

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the ...

Hydraulic Extrusion of the Spinal Cord and Isolation of Dorsal Root Ganglia in Rodents

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may result in damage to the spinal cord caused by the dissection process. Here, we show how the spinal cord can be extruded using hydraulic pressure. Handling time is significantly reduced to only a few minutes, likely decreasing protein damage. The low risk of damage to the spinal cord tissue improves subsequent immunohistochemical analysis. By performing hydraulic spinal cord extrusion instead of traditional laminectomy, the rodents can further be used for DRG isolation, thereby lowering the number of animals and allowing analysis across tissues from the same rodent. We demonstrate a consistent method to identify and isolate the DRGs according to their localization relative to the costae. It is, however, important to adjust this method to the particular animal used, as the number of spinal cord segments, both thoracic and lumbar, may vary according to animal type and strain. In addition, we illustrate further processing examples of the isolated tissues.

Here, we present a protocol for hydraulic extrusion of the spinal cord as well as identification and isolation of specific dorsal root ganglia (DRGs) in the same rodent. Compared to standard spinal cord isolation methods, this method is significantly faster and reduces the risk of tissue damage.

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may ...

Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each other anatomically, it is important to identify an appropriate model to use for the study of axon regeneration. By using a suitable model, we can formulate a specific treatment based on the severity of injury, the neuronal cell type of interest, and the desired spinal tract for assessing regeneration. Within the sensory pathway, DRG neurons are responsible for relaying sensory information from the periphery to the CNS. We present here a protocol that uses a DRG injection with a viral vector and a concurrent dorsal root crush injury in the lower cervical spinal cord of an adult rat as a model to study sensory axon regeneration. As demonstrated using a control virus, AAV5-GFP, we show the effectiveness of a direct DRG injection in transducing DRG neurons and tracing sensory axons into the spinal cord. We also show the effectiveness of the dorsal root crush injury in denervating the forepaw as an injury model for evaluating axon regeneration. Despite the requirement for specialized training to perform this invasive surgical procedure, the protocol is flexible, and potential users can modify many parts to accommodate their experimental requirements. Importantly, it can serve as a foundation for those in search of a suitable animal model for their studies. We believe that this article will help new users to learn the procedure in a very efficient and effective manner.

This protocol presents the use of a dorsal root ganglion (DRG) injection with a viral vector and a concurrent dorsal root crush injury in an adult rat as a model to study sensory axon regeneration. This model is suitable for investigating the use of gene therapy to promote sensory axon regeneration.

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each ...

Dorsal Root Ganglia Isolation and Primary Culture to Study Neurotransmitter Release

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral tissues, such as skin, muscle and visceral organs, as well as the spinal dorsal horn of the central nervous system. Sensory neurons transmit somatic sensation, including touch, pain, thermal, and proprioceptive sensations. Therefore, DRG primary cultures are widely used to study the cellular mechanisms of nociception, physiological functions of sensory neurons, and neural development. The cultured neurons can be applied in studies involving electrophysiology, signal transduction, neurotransmitter release, or calcium imaging. With DRG primary cultures, scientists may culture dissociated DRG neurons to monitor biochemical changes in single or multiple cells, overcoming many of the limitations associated with in vivo experiments. Compared to commercially available DRG-hybridoma cell lines or immortalized DRG neuronal cell lines, the composition and properties of the primary cells are much more similar to sensory neurons in tissue. However, due to the limited number of cultured DRG primary cells that can be isolated from a single animal, it is difficult to perform high-throughput screens for drug targeting studies. In the current article, procedures for DRG collection and culture are described. In addition, we demonstrate the treatment of cultured DRG cells with an agonist of neuropeptide FF receptor type 2 (NPFFR2) to induce the release of peptide neurotransmitters (calcitonin gene-related peptide (CRGP) and substance P (SP)).

Dorsal root ganglia (DRG) primary cultures are frequently used to study physiological functions or pathology-related events in sensory neurons. Here, we demonstrate the use of lumbar DRG cultures to detect the release of neurotransmitters after neuropeptide FF receptor type 2 stimulation with a selective agonist.

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral ...

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for nerve fibers in the central and peripheral nervous systems. This method is particularly applicable to dorsal root ganglia (DRG), which are primary sensory neurons that exhibit a pseudo-unipolar structure of nervous processes. The patterns and features of the action potentials passed along axons are recordable in these neurons. The present study uses in vivo single-fiber recordings to observe the conduction failure of sciatic nerves in complete Freund&#8217;s adjuvant (CFA)-treated rats. As the underlying mechanism cannot be studied using in vivo single-fiber recordings, patch-clamp-recordings of DRG neurons are performed on preparations of intact DRG with the attached sciatic nerve. These recordings reveal a positive correlation between conduction failure and the rising slope of the after-hyperpolarization potential (AHP) of DRG neurons in CFA-treated animals. The protocol for in vivo single fiber-recordings allows the classification of nerve fibers via the measurement of conduction velocity and monitoring of abnormal conditions in nerve fibers in certain diseases. Intact DRG with attached peripheral nerve allows observation of the activity of DRG neurons in most physiological conditions. Conclusively, single-fiber recording combined with electrophysiological recording of intact DRGs is an effective method to examine the role of conduction failure during the analgesic process.

Single-fiber recording is an effective electrophysiological technique that is applicable to the central and peripheral nervous systems. Along with the preparation of intact DRG with the attached sciatic nerve, the mechanism of conduction failure is examined. Both protocols improve the understanding of the peripheral nervous system&#39;s relationship with pain.

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for ...

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.