The authors would like to acknowledge the Painless Research Foundation for support of the work. This work was also supported by the NIH grants R01 NS080921-01 and R21 NS079897-01A1.

Ethics Statement: All procedures for the maintenance and use of the experimental animals conformed to the regulations of UCSF Committees on Animal Research and were carried out in accordance with the guidelines of the NIH regulations on animal use and care (Publication 85 - 23, Revised 1996). The UCSF Institutional Animal Care and Use Committee approved the protocols used in this study.
1. Preparation of Instruments, Solutions and Dishes
<ol>
	<li>Prepare Artificial Cerebrospinal Fluid (aCSF).
	<ol>
		<li>Prepare 500 ml of 10x low cation solution, and 500 ml of 10x bicarbonate solution (see Table 1 and 2 for solution preparation). Store at 4 oC and use within one month.</li>
		<li>Prepare 600 ml of fresh aCSF by mixing 60 ml of 10x low cation solution with 60 ml of bicarbonate solution, and then add 480 ml of deionized water to reach a final volume of 600 ml. Bubble the aCSF with carbogen (95% O2 and 5% CO2) for at least 10 min before use.</li>
	</ol>
	</li>
	<li>Pull Patch Pipette from Capillary Glass.
	<ol>
		<li>Place thin walled capillary glass into a pipette puller to prepare recording pipettes. 
		Note: In our lab, the following setting parameters of the puller are used to achieve the ideal pipette shape: heat (510), velocity (32). The ideal pipette shape for intact DRG recording should have a gradual slender taper with a low cone angle, and this can best be achieved by trial and error.</li>
		<li>Ensure that the resistance of the pipette is 3-5 M&#937; when filled with internal solutions (see Table 3 for solution preparation). Please refer to Axon Guide for the detailed method of measuring pipette resistance 18.</li>
	</ol>
	</li>
	<li>Add 1 mg of collagenase to 400 &#181;l of aCSF, to give a final concentration of 13 units/ml. Then fill each glass pipette with about 20 &#181;l of collagenase solution. Store the filled pipettes in a -20 oC freezer and use them within three weeks.</li>
	<li>Prepare 200 ml of cold aCSF (about 4 oC). To cool the aCSF quickly place it into a beaker, seal with plastic wrap and place in a -20 oC freezer for approximately 20 min until it starts to freeze. Place the beaker in an ice bath and bubble with carbogen for 10 min. Bubble the remaining 400 ml of aCSF with carbogen at RT.</li>
	<li>While the aCSF is cooling, cut the end of a plastic disposable transfer pipette to enlarge the opening to a diameter of 5 mm (this will be used to transfer the DRG). Collect the following surgical instruments: scalpel (#15), Mayo straight and curved scissors, fine 2 mm tip rongeur, Adson (toothed) forceps, iris scissors, spring scissors and two fine forceps. Pour the oxygenated cold aCSF into 3 glass petri dishes (Outer diameter: 10 cm).</li>
</ol>
2. DRG Dissection
<ol>
	<li>Anesthetize a rat (200-220 g) with sodium pentobarbital (100 mg/kg, i.p.) and confirm that the level of anesthesia is adequate by testing pedal reflex and eye blink reflex.</li>
	<li>After verifying that the rat is anesthetized, shave the lumbar area and incise the skin and subcutaneous tissue along the midline. Detach the paraspinal muscles from the spinous processes at the L1-S2 level using a fine periosteal elevator.</li>
	<li>Use curved scissors to cut the spinous processes from L1 to S2. Use straight Mayo scissors to make transverse cuts in the exposed vertebral column at approximately L1 and S1 and remove this segment of the vertebral column en bloc and quickly submerge it into cold aCSF (prepared in step 1.5) for 2-3 min.&#160;Following the removal of lumbar DRGs, euthanasia is carried out by bilateral thoracotomy while the rat is still under deep pentobarbital&#160; anesthesia.&#160;</li>
	<li>Flush off the blood from the removed vertebral column and attached tissue and transfer to a petri dish with cold aCSF. Use a fine rongeur to remove the laminae. Cut the dura mater along the midline with microscissors to expose the spinal cord. Gently lift the spinal cord, and cut the nerve roots at their entry point in the spinal cord using iris scissors, then remove the spinal cord.</li>
	<li>Transfer the remaining vertebra with DRGs to the second petri dish filled with cold aCSF. Identify the L4 and L5 DRGs based on their relative position to transverse processes and sciatic nerve. 
	Note: Maintaining the sciatic nerve intact will help to identify L4 and L5 DRG (the sciatic nerve originates from the L4-6 spinal nerves).</li>
	<li>Use Dumont forceps and the Noyes scissors to free the DRGs from the surrounding connective tissues. Keep the nerve root and spinal nerve attached to the DRGs.</li>
	<li>Use the transfer pipette to move the DRGs into the third petri dish for further dissection.</li>
	<li>Under the dissection microscope, carefully remove as much as possible of the epineurium surrounding the DRG.
	<ol>
		<li>Use the Noyes spring scissors to cut an opening where the dorsal and ventral roots join the DRG and separate the ventral root from the DRG. Use Dumont #5 forceps with blunt tips to hold the epineurium, and use another fine forceps to roll the DRG from the epineurium.</li>
		<li>Continue to remove as much as possible of epineurium attached to DRGs using the fine forceps. 
		Note: A well-dissected DRG is important to obtain a good digestion in the next step.</li>
	</ol>
	</li>
</ol>
3. DRG Digestion
<ol>
	<li>Transfer the DRG to the recording chamber. Make sure the side of DRG where the nerve roots originate faces down. 
	Note: In this way, most neurons are accessible.</li>
	<li>Use an anchor to stabilize the DRG. Perfuse DRG with oxygenated aCSF at a rate of 0.5 ml/min through plastic tubes connected with a peristaltic pump, which is placed on a table next to the recording rig.</li>
	<li>Wait for 30 min to allow the cells to recover. Assess the quality of the DRG under infrared differential interface contrast (IR-DIC) optics at 40X objective magnification through a CCD camera. 
	Note: Good DRGs usually contains many round, well contrasted, neurons surrounded by satellite glial cells on its surface.</li>
	<li>Digest a small area of the surface of the DRG. 
	Note: The glass patch pipettes are placed in pipette holders, which are small couplings that allow tubing to be attached to the glass pipette with an airtight seal.
	<ol>
		<li>Individually connect two 1 ml syringes to the pipette holders through two pieces of small diameter rubber tubing. Put one glass pipette filled with collagenase into one of the pipette holders and an empty glass pipette into the other one.</li>
		<li>Use the micromanipulator to position the pipettes just above the DRG. Then, under microscope magnification, collide the tips of two pipettes against each other gently to enlarge the openings of pipette tips (Ideally a diameter of 5-10 &#181;m).</li>
		<li>Move the pipette filled with enzyme close to the surface of DRG and briefly apply positive pressure to the pipette containing collagenase through the tube connecting the holder and the syringe by displacing the plunger about 0.5 ml. 
		Note: Under pressure, the flow of the enzyme from the pipette will slightly enlarge the space between the DRG neurons, which can serve as sign for enzyme application.</li>
		<li>After 10 to 15 min, when debris of the remaining epineurium is observed, apply gentle negative pressure to the empty pipette to suck away the debris. 
		Note: In this way, the neurons and surrounding satellite glial cells will be clearly exposed and ready for patch recording.</li>
		<li>Discard the two pipettes into the sharps container.</li>
	</ol>
	</li>
</ol>
4. Patch Clamp Recordings
<ol>
	<li>Place the electrode solution (see Table 3) on ice to prevent degradation. Fill the glass patch pipette with filtered intracellular solution (syringe filter, pore size: 0.2 &#181;m) and place the pipette in the headstage pipette holder. Apply a gentle positive pressure to the pipette through a 5 ml syringe by displacing the plunger about 1 ml before lowering the pipette into bath solution. 
	Note: This prevents the pipette from becoming blocked.</li>
	<li>Choose &#34;V-clamp&#34; mode by turning the mode knob on the amplifier, then open &#34;Membrane test&#34; interface in the software. Move the pipette close to the target neuron under microscope inspection. 
	Note: In the intact DRG preparation, a thin layer of satellite glial cells encapsulates each neuron.</li>
	<li>Use positive pressure from the pipette to traverse the satellite glial cell layer until a sudden enlargement of space between the neuron and surrounding layer of satellite glial cells is observed. Keep moving the pipette towards the neuron until a &#34;dimple&#34; is observed on the neuron. 
	Note: Compared with recordings from disassociated neurons, the positive pressure needs to be a slightly larger, which will help to penetrate the satellite glial cell layer and increase the chances for successful patching.</li>
	<li>Reduce the positive pressure. Next, achieve a giga ohm seal with gentle suction. 
	Note: With this method, the success rate of recording is at least 70%.</li>
	<li>Obtain whole-cell recording configuration as described previously19.
	<ol>
		<li>Briefly, penetrate the neuron cell membrane via a short but strong suction. Alternatively, use the &#34;zap&#34; function on the amplifier while suction is applied.</li>
		<li>Once a whole cell mode is established, compensate whole-cell capacitance and series resistance (Rs) by turning the capacitance and resistance compensation knobs on the amplifier. 
		Note: Rs is normally 5-20 M&#937;.</li>
		<li>Abandon the cell if Rs is initially greater than 30 M&#937;; or Rs changes by more than 20% during the recording. Also measure the resting membrane potential; abandon a cell if the resting membrane potential is more than -50 mV.</li>
	</ol>
	</li>
	<li>Close the &#34;Membrane test&#34; window. Choose &#34;I-clamp normal&#34; mode by turning the mode knob on the amplifier.</li>
	<li>Click &#34;open protocol&#34; in the software, select and load the protocol for measuring rheobase. Click &#34;record&#34; to start recording. Measure input resistance and rheobase to examine the neuronal excitability by injecting a graded series of depolarizing currents in steps of 100 pA.
	<ol>
		<li>Click &#34;open protocol&#34; again and select the protocol for measuring membrane threshold. A 500 ms depolarizing ramp current (2,000 pA/s) will be injected to the neuron.</li>
		<li>Use data acquisition and analysis software (e.g. Clampfit) to analyze the recorded traces according to manufacturer&#39;s instructions. Use the software to calculate the input resistance (Rin) on the basis of the steady-state I-V relationship during the hyperpolarizing currents delivered. Move cursor to measure the value for rheobase and membrane threshold. 
		Note: The amplitude of current required to induce the AP is defined as the rheobase, and the lowest voltage for inducing AP is defined as membrane threshold.</li>
	</ol>
	</li>
	<li>Record the Ligand Induced Currents.
	<ol>
		<li>Fill a pipette with the specific agonists. 
		Note: In the current report, we used 100 &#181;M glutamate and 100 &#181;M AITC to induce the currents mediated by glutamate receptors and TRPA1 receptors respectively.</li>
		<li>Check the pipette to make sure there are no air bubbles inside. Place the pipette in the pipette holder. Connect the pipette holder with a tube connected to a drug dispensing system.</li>
		<li>Use the manipulator to move the pipette within 50 &#181;m of the neuron. Set the drug dispensing system pressure to 1 psi and the duration to 1 sec. Switch the recording mode to voltage clamp, and clamp at -70 mV. Briefly apply the pressure via the drug dispensing system to record a drug-induced current.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Basbaum, A. I., Bautista, D. M., Scherrer, G., Julius, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+and+molecular+mechanisms+of+pain.">Cellular and molecular mechanisms of pain.</a> Cell. 139, 267-284 (2009).</li><li>Caterina, M. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+capsaicin+receptor:+a+heat-activated+ion+channel+in+the+pain+pathway.">The capsaicin receptor: a heat-activated ion channel in the pain pathway.</a> Nature. 389, 816-824 (1997).</li><li>Gong, K., Bhargava, A., Jasmin, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GluN2B+N-methyl-D-aspartate+receptor+and+excitatory+amino+acid+transporter+3+are+upregulated+in+primary+sensory+neurons+after+7+days+of+morphine+administration+in+rats:+implication+for+opiate-induced+hyperalgesia.">GluN2B N-methyl-D-aspartate receptor and excitatory amino acid transporter 3 are upregulated in primary sensory neurons after 7 days of morphine administration in rats: implication for opiate-induced hyperalgesia.</a> Pain. 157, 147-158 (2016).</li><li>Gong, K., Kung, L. H., Magni, G., Bhargava, A., Jasmin, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+response+to+glutamate+in+small+diameter+dorsal+root+ganglion+neurons+after+sciatic+nerve+injury.">Increased response to glutamate in small diameter dorsal root ganglion neurons after sciatic nerve injury.</a> PloS one. 9, 95491 (2014).</li><li>Gong, K., Zou, X., Fuchs, P. N., Lin, Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minocycline+inhibits+neurogenic+inflammation+by+blocking+the+effects+of+tumor+necrosis+factor-alpha.">Minocycline inhibits neurogenic inflammation by blocking the effects of tumor necrosis factor-alpha.</a> Clin Exp Pharmacol Physiol. , (2015).</li><li>Ohtori, S., Takahashi, K., Moriya, H., Myers, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TNF-alpha+and+TNF-alpha+receptor+type+1+upregulation+in+glia+and+neurons+after+peripheral+nerve+injury:+studies+in+murine+DRG+and+spinal+cord.">TNF-alpha and TNF-alpha receptor type 1 upregulation in glia and neurons after peripheral nerve injury: studies in murine DRG and spinal cord.</a> Spine. 29, 1082-1088 (2004).</li><li>Waxman, S. G., Cummins, T. R., Dib-Hajj, S., Fjell, J., Black, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sodium+channels,+excitability+of+primary+sensory+neurons,+and+the+molecular+basis+of+pain.">Sodium channels, excitability of primary sensory neurons, and the molecular basis of pain.</a> Muscle nerve. 22, 1177-1187 (1999).</li><li>Zhang, J. M., Song, X. J., LaMotte, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+excitability+of+sensory+neurons+in+rats+with+cutaneous+hyperalgesia+produced+by+chronic+compression+of+the+dorsal+root+ganglion.">Enhanced excitability of sensory neurons in rats with cutaneous hyperalgesia produced by chronic compression of the dorsal root ganglion.</a> J Neurophysiol. 82, 3359-3366 (1999).</li><li>Dib-Hajj, S. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasticity+of+sodium+channel+expression+in+DRG+neurons+in+the+chronic+constriction+injury+model+of+neuropathic+pain.">Plasticity of sodium channel expression in DRG neurons in the chronic constriction injury model of neuropathic pain.</a> Pain. 83, 591-600 (1999).</li><li>Cummins, T. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+persistent+tetrodotoxin-resistant+sodium+current+in+SNS-null+and+wild-type+small+primary+sensory+neurons.">A novel persistent tetrodotoxin-resistant sodium current in SNS-null and wild-type small primary sensory neurons.</a> J Neurosci. 19, RC43 (1999).</li><li>Zheng, J. H., Walters, E. T., Song, X. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissociation+of+dorsal+root+ganglion+neurons+induces+hyperexcitability+that+is+maintained+by+increased+responsiveness+to+cAMP+and+cGMP.">Dissociation of dorsal root ganglion neurons induces hyperexcitability that is maintained by increased responsiveness to cAMP and cGMP.</a> J Neurophysiol. 97, 15-25 (2007).</li><li>Schoenen, J., Delree, P., Leprince, P., Moonen, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurotransmitter+phenotype+plasticity+in+cultured+dissociated+adult+rat+dorsal+root+ganglia:+an+immunocytochemical+study.">Neurotransmitter phenotype plasticity in cultured dissociated adult rat dorsal root ganglia: an immunocytochemical study.</a> J Neurosci Res. 22, 473-487 (1989).</li><li>Hanani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Satellite+glial+cells:+more+than+just+'rings+around+the+neuron'.">Satellite glial cells: more than just 'rings around the neuron'.</a> Neuron Glia Biol. 6, 1-2 (2010).</li><li>Takeda, M., Nasu, M., Kanazawa, T., Shimazu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+GABA(B)+receptors+potentiates+inward+rectifying+potassium+currents+in+satellite+glial+cells+from+rat+trigeminal+ganglia:+in+vivo+patch-clamp+analysis.">Activation of GABA(B) receptors potentiates inward rectifying potassium currents in satellite glial cells from rat trigeminal ganglia: in vivo patch-clamp analysis.</a> Neuroscience. 288, 51-58 (2015).</li><li>Zhang, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Altered+functional+properties+of+satellite+glial+cells+in+compressed+spinal+ganglia.">Altered functional properties of satellite glial cells in compressed spinal ganglia.</a> Glia. 57, 1588-1599 (2009).</li><li>Fan, N., Donnelly, D. F., LaMotte, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+compression+of+mouse+dorsal+root+ganglion+alters+voltage-gated+sodium+and+potassium+currents+in+medium-sized+dorsal+root+ganglion+neurons.">Chronic compression of mouse dorsal root ganglion alters voltage-gated sodium and potassium currents in medium-sized dorsal root ganglion neurons.</a> J Neurophysiol. 106, 3067-3072 (2011).</li><li>Fan, N., Sikand, P., Donnelly, D. F., Ma, C., Lamotte, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+Na++and+K++currents+in+small+mouse+dorsal+root+ganglion+neurons+after+ganglion+compression.">Increased Na+ and K+ currents in small mouse dorsal root ganglion neurons after ganglion compression.</a> J Neurophysiol. 106, 211-218 (2011).</li><li>Sherman-Gold, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Axon Guide. , (2008).</li><li>Cummins, T. R., Rush, A. M., Estacion, M., Dib-Hajj, S. D., Waxman, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-clamp+and+current-clamp+recordings+from+mammalian+DRG+neurons.">Voltage-clamp and current-clamp recordings from mammalian DRG neurons.</a> Nat Protoc. 4, 1103-1112 (2009).</li><li>Zhang, J. M., Donnelly, D. F., LaMotte, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch+clamp+recording+from+the+intact+dorsal+root+ganglion.">Patch clamp recording from the intact dorsal root ganglion.</a> J Neurosci Methods. 79, 97-103 (1998).</li><li>Benn, S. C., Costigan, M., Tate, S., Fitzgerald, M., Woolf, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+expression+of+the+TTX-resistant+voltage-gated+sodium+channels+Nav1.8+(SNS)+and+Nav1.9+(SNS2)+in+primary+sensory+neurons.">Developmental expression of the TTX-resistant voltage-gated sodium channels Nav1.8 (SNS) and Nav1.9 (SNS2) in primary sensory neurons.</a> J Neurosci. 21, 6077-6085 (2001).</li><li>Funakoshi, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+development+of+TRPV1-expressing+sensory+nerves+in+peripheral+organs.">Differential development of TRPV1-expressing sensory nerves in peripheral organs.</a> Cell Tissue Res. 323, 27-41 (2006).</li><li>Hayar, A., Gu, C., Al-Chaer, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+method+for+patch+clamp+recording+and+calcium+imaging+of+neurons+in+the+intact+dorsal+root+ganglion+in+rats.">An improved method for patch clamp recording and calcium imaging of neurons in the intact dorsal root ganglion in rats.</a> J Neurosci Methods. 173, 74-82 (2008).</li><li>Yagi, J., Sumino, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+a+hyperpolarization-activated+current+by+clonidine+in+rat+dorsal+root+ganglion+neurons.">Inhibition of a hyperpolarization-activated current by clonidine in rat dorsal root ganglion neurons.</a> J Neurophysiol. 80, 1094-1104 (1998).</li><li>Ma, C., Donnelly, D. F., LaMotte, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+visualization+and+functional+characterization+of+primary+somatic+neurons.">In vivo visualization and functional characterization of primary somatic neurons.</a> J Neurosci Methods. 191, 60-65 (2010).</li><li>Vit, J. P., Jasmin, L., Bhargava, A., Ohara, P. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Satellite+glial+cells+in+the+trigeminal+ganglion+as+a+determinant+of+orofacial+neuropathic+pain.">Satellite glial cells in the trigeminal ganglion as a determinant of orofacial neuropathic pain.</a> Neuron Glia Biol. 2, 247-257 (2006).</li></ol>

The authors declare that they have no competing financial interests.

We report a method to prepare whole DRGs for patch clamp studies. There are several key elements for preparing an ideal specimen. Firstly, it is important to dissect the DRGs with dorsal roots attached. After that, the epineurium need to be carefully removed while avoiding damage to the neurons. Finally, to expose the neurons and their surrounding satellite glial cells, it is necessary to digest the remaining connective tissue. Intact DRGs from adult rats prepared with the method described here will maintain good viabili...

Sensation is essential to an organism&#39;s survival and wellbeing. The transmission of stimuli is dependent on the sensory pathways starting at peripheral endings of axons from primary sensory neurons. Primary sensory neurons, with the exception of the mesencephalic nucleus of the trigeminal nerve, are located in the trigeminal ganglia and dorsal root ganglia (DRGs). They serve as gatekeepers of the sensory information 1. At the perikarial membrane, just as at the central and peripheral terminals, DRG neurons express receptors and ion channels, such as glutamate receptors, TNF alpha receptors, transient receptor potential cation channel subfamily V member 1 (TRPV1), sodium channels, etc. 2-7. Patch clamp recordings of the perikarial membrane allow understanding functional changes of many of these receptors and channels throughout the neuron.
The patch clamp recording technique is a powerful tool for studying the activities of channels or receptors and a great number of studies have been conducted by applying this technique on DRG neurons 8-10. In most studies the DRG is removed by cutting the dorsal rootlets and spinal nerve close to the ganglion. After mincing, the ganglion is then placed in digestive enzymes that result in dissociation of the DRG neurons, which may then be recorded immediately or cultured for several days prior to recording. Unfortunately, dissociation of DRG neurons involves a necessary axotomy close to the perikarya. Once dissociated and axotomized, DRG neurons undergo phenotypic changes as well as changes in membrane excitability 11,12. The loss of contact between the perikarya of individual neurons and the satellite glial cells that normally surround them is likely to contribute to these changes 13. The crosstalk between neurons and satellite glial cells is both essential in physiological conditions and in adaptation to pathological conditions such as those leading to intractable pain 14,15. It would be challenging to study the interaction between neurons and satellite glial cells using a dissociated DRG preparation.
Intact DRGs, on the other hand, provide closer to in vivo conditions. In the past several years, our laboratory, as well as some other groups, has been using intact DRGs from adult rats to investigate changes of primary sensory neurons in different conditions associated with chronic pain 3-5,11,15-17. Although the techniques used in these studies are somewhat established, a step-by-step description has not yet been published. In the present manuscript, we describe a convenient and fast way to prepare intact DRGs and their use for patch clamp recordings.

<table><tbody><tr><td>Pentobarbital sodium</td><td>vortech Pharmaceuticals</td><td></td><td></td></tr><tr><td>syringe</td><td>BD</td><td>309659</td><td>1 ml, 5 ml.</td></tr><tr><td>scalpel</td><td>BD</td><td></td><td>size: 15</td></tr><tr><td>Mayo straight scissor</td><td>Fine Science Tools</td><td>14010-15</td><td></td></tr><tr><td>Mayo curved scissor</td><td>Fine Science Tools</td><td>14011-15</td><td></td></tr><tr><td>Rongeur</td><td>Fine Science Tools</td><td>16021-14</td><td></td></tr><tr><td>Adson toothed forceps</td><td>Fine Science Tools</td><td>11027-12</td><td></td></tr><tr><td>Iris Scissor</td><td>Fine Science Tools</td><td>14084-08</td><td></td></tr><tr><td>Noyes spring scissor</td><td>Fine Science Tools</td><td>15124-12</td><td></td></tr><tr><td>Bone scissors</td><td>Fine Science Tools</td><td>16044-10</td><td>Special for cutting the bones.&amp;nbsp;</td></tr><tr><td>Forceps: Dumont, Dumoxel Biologie #5</td><td>Fine Science Tools</td><td>11252-30</td><td>These have the fine tips that do not need sharpening when first purchased.</td></tr><tr><td>periosteal elevator</td><td>Sklar</td><td>97-0530</td><td></td></tr><tr><td>Dissection microscope</td><td>WILD</td><td></td><td></td></tr><tr><td>Transfer pipette</td><td>Fisher brand</td><td>13-711-5AM</td><td></td></tr><tr><td>Petri dish (10 cm)</td><td>Pyrex</td><td></td><td>Glass petri dish can avoid damaging the tips of fine forceps</td></tr><tr><td>Collagenase (Liberase TM)</td><td>Roche</td><td>05-401-119-001</td><td>dissolve at the concentration of 13 U/ml, aliquot into glass pipette. Avoid repeated freeze and thaw.</td></tr><tr><td>filter</td><td>Thermo scientific</td><td>7232520</td><td>Filter the internal solutions for patch clamp recording to avoid clog.</td></tr><tr><td>Glass pipette</td><td>Sutter</td><td>BF150-110-7.5</td><td></td></tr><tr><td>Anchor</td><td>Havard apparatus</td><td>64-0250</td><td>stabilize the DRG to avoid drift.</td></tr><tr><td>Peristaltic pump</td><td>WPI</td><td></td><td></td></tr><tr><td>Pipette puller</td><td>Sutter</td><td>P97</td><td></td></tr><tr><td>Amplifier</td><td>Molecular devices</td><td>Axopatch 200B</td><td></td></tr><tr><td>Digitizer</td><td>Molecular devices</td><td>1440D</td><td></td></tr><tr><td>Microscope</td><td>NIKON</td><td>FN600</td><td></td></tr><tr><td>Micro-manipulator</td><td>Sutter</td><td>MPC200</td><td></td></tr><tr><td>microinjection dispense system</td><td>General Valve</td><td>Picrospitzer II</td><td>fast drug application system</td></tr><tr><td>Carbogen (95% O2, 5% CO2)</td><td>Local Medical Gas supplier</td><td></td><td></td></tr></tbody></table>

patch clamp recordings on intact dorsal root ganglia from adult rats

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of recordings are conducted on dissociated DRG neurons, which is a standard preparation for most laboratories. Neuronal properties, however, can be altered by axonal injury resulting from enzyme digestion used in acquiring dissociated neurons. Further, dissociated neuron preparations cannot fully represent the microenvironment of the DRG since loss of contact with satellite glial cells that surround the primary sensory neurons is an unavoidable consequence of this method. To overcome the limitations in using conventional dissociated DRG neurons for patch clamp recordings, in this report we describe a method to prepare intact DRGs and conduct patch clamp recordings on individual primary sensory neurons ex vivo. This approach permits the fast and straightforward preparation of intact DRGs, mimicking in vivo conditions by keeping DRG neurons associated with their surrounding satellite glial cells and basement membrane. Furthermore, the method avoids axonal injury from manipulation and enzyme digestion such as when dissociating DRGs. This ex vivo preparation can additionally be used to study the interaction between primary sensory neurons and satellite glial cells.

Figure 1 shows the process of preparing intact DRG for patch recording. Figure 1A shows the exposure and location of the ganglia after laminectomy. Figure1B shows L3, L4 and L5 DRGs with the nerve roots attached after removing the spinal cord. Then L4 and 5 DRGs are carefully dissected and freed from the vertebrae. Next, the epineurium, a transparent membrane surrounding the DRG, is removed (yellow arrow, Figure 1D). The ...

Watch this Scientific Journal Video about Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats at JoVE.com

Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats

This manuscript describes how to prepare intact dorsal root ganglia for patch clamp recordings. This preparation maintains the microenvironment for neurons and satellite glial cells, thus avoiding the phenotypic and functional changes seen using dissociated DRG neurons.

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of ...

patch-clamp-recordings-on-intact-dorsal-root-ganglia-from-adult-rats

Research

JoVE Journal

Biochemistry

14.2K Views.  University of California, San Francisco. This manuscript describes how to prepare intact dorsal root ganglia for patch clamp recordings. This preparation maintains the microenvironment for neurons and satellite glial cells, thus avoiding the phenotypic and functional changes seen using dissociated DRG neurons.

Video: Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

Neuroscience

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 ...

The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to network properties. We show how to prepare viable oblique slices of the spinal cord of young mice (P2 - P11). In this preparation, the motoneurons retain&#160;their axons coming out from the ventral roots of the&#160;spinal cord. Stimulation of these axons elicits back-propagating action potentials invading the motoneuron&#160;somas and exciting the motoneuron&#160;collaterals within the spinal cord. Recording of antidromic action potentials is an immediate, definitive and elegant way to characterize motoneuron identity, which surpasses&#160;other identification methods. Furthermore, stimulating the motoneuron collaterals is a simple and reliable way to excite the collateral targets of the motoneurons within the spinal cord, such as other motoneurons or Renshaw cells. In this protocol, we present antidromic recordings from the motoneuron somas as well as Renshaw cell excitation, resulting from ventral root stimulation.

We show how to prepare oblique slices of the spinal cord in young mice. This preparation allows for the stimulation of the ventral roots.

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...

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 ...

Isolating and Culturing Vestibular and Spiral Ganglion Somata from Neonatal Rodents for Patch-Clamp Recordings

The compact morphology of isolated and cultured inner ear ganglion neurons allows for detailed characterizations of the ion channels and neurotransmitter receptors that contribute to cell diversity across this population. This protocol outlines the steps necessary for successful dissecting, dissociating, and short-term culturing of the somata of inner ear bipolar neurons for the purpose of patch-clamp recordings. Detailed instructions for preparing vestibular ganglion neurons are provided with the necessary modifications needed for plating spiral ganglion neurons. The protocol includes instructions for performing whole-cell patch-clamp recordings in the perforated-patch configuration. Example results characterizing the voltage-clamp recordings of hyperpolarization-activated cyclic nucleotide-gated (HCN)-mediated currents highlight the stability of perforated-patch recording configuration in comparison to the more standard ruptured-patch configuration. The combination of these methods, isolated somata plus perforated-patch-clamp recordings, can be used to study cellular processes that require long, stable recordings and the preservation of intracellular milieu, such as signaling through G-protein coupled receptors.

Presented here are methods providing detailed instructions for dissecting, dissociating, culturing, and patch-clamp recording from vestibular ganglion and spiral ganglion neurons of the inner ear.

The compact morphology of isolated and cultured inner ear ganglion neurons allows for detailed characterizations of the ion channels and neurotransmitter ...

Understanding Motor Neuron Responses to Spinal Cord Stimulation

The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Author Spotlight: Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology 

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...

Calcium Imaging of Thoracic Segment T1 DRG for Cardiopulmonary Sensory Neurons

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.

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. ...

Patch-Clamp Recordings from the Dendrite of a Dopaminergic Neuron in a Brain Slice

Source: Engel, D. <a href="https://app.jove.com/v/54601">Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons.</a> J. Vis. Exp. (2016).
This video demonstrates a technique for patch-clamp recording from the dendritic membrane surface. This helps to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels in dopaminergic neurons.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

JoVE Encyclopedia of Experiments

Neurophysiology

Electrophysiology Techniques

Whole-Cell Patch Clamp Recording in a Substantia Gelatinosa Neuron of a Spinal Cord Slice

Source: Zhu, M., et al. <a href="https://app.jove.com/v/58479">Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons.</a> J. Vis. Exp. (2019)
This video demonstrates the procedure for performing electrophysiological recordings in substantia gelatinosa (SG) neurons of spinal cord slices, using whole-cell patch-clamp techniques to analyze neuron firing patterns. This method allows the study of neural responses to stimuli in SG neurons of the spinal cord.

Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats

Samenvatting

Meer video's verkennen