The study was funded (awarded to F-Q.Z.) by NIH (R01NS064288, R01NS085176, R01GM111514, R01EY027347), the Craig H. Neilsen Foundation and the BrightFocus Foundation.

All animal experiments were performed in accordance with the animal protocol approved by the Johns Hopkins Institutional Animal Care and Use Committee.
1. Materials and Reagents
<ol>
	<li>Animals
	<ol>
		<li>Use six-week-old female CF1 mice weighing 30&#8211;35 g for the experiments. 
		NOTE: The mice were group-housed (5 mice per cage) in individually ventilated sterilized cages with 1/4&#8221; corn cob bedding and 2&#8221; square nestlets for nesting. The cages were maintained in a controlled 12-h light-dark cycle and the room temperature was between 19.4 &#176;C and 25 &#176;C. The mice were fed with global rodent diet and automatic water supplying system.&#160;</li>
	</ol>
	</li>
	<li>Instruments
	<ol>
		<li>Use the following surgical instruments to efficiently conduct the surgery: disposable syringes (27 G, 1.0 mL) for intraperitoneal injection, fur clipper, stereo dissection microscope, micro-surgery forceps, micro-surgery scissors, and small bone rongeurs and sterilized non-woven sponges. 
		NOTE: All the instruments and materials are autoclaved before surgery or pre-sterilized by the producers. During surgery, the instruments are sprayed with 75% alcohol to maintain the sterilization.</li>
	</ol>
	</li>
	<li>Anesthetic Solutions
	<ol>
		<li>Use anesthetic solution #1 to induce the anesthesia and use anesthesia solution #2 before DRG injection to maintain the anesthesia.
		<ol>
			<li>To make anesthetic solution #1, dilute ketamine and xylazine in sterile saline at a final concentration of 10 mg/mL and 1.2 mg/mL.</li>
			<li>To make anesthetic solution #2, dilute avertin in sterile saline at a final concentration of 20 mg/mL.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>DNA Plasmid and RNA Oligos
	<ol>
		<li>Prepare the plasmids using the commercial kit following the manufacturer&#39;s protocols accordingly. Dilute the plasmid DNAs in sterile deionized water to the concentration of 2.0 &#181;g/&#181;L.</li>
		<li>Add Fast Green dye solution to reach a final concentration of 0.005&#8211;0.01% (v/v) for better visualization of the DRG outline during injection.</li>
		<li>Dissolve the siRNA or microRNA oligos in corresponding buffers provided by manufacturers to reach a final concentration at 50 &#181;M.</li>
	</ol>
	</li>
	<li>Micro-Injection Pipette
	<ol>
		<li>Pull the capillary glass using the micropipette puller and cut the tip of the pulled capillary glass pipette with microsurgery scissors to generate an opening with an approximate 50 &#181;m outer diameter. Then sterilize the glass pipette under UV light on a clean bench for 20&#8211;30 min approximately.</li>
		<li>Mount the sterilized glass pipette on the pipette holder connected to an intracellular microinjection dispense system. Set the parameters of the microinjection system at 30 psi of pressure and 8 ms of duration.</li>
	</ol>
	</li>
	<li>Electroporation System
	<ol>
		<li>Connect the electrodes to the square wave electroporation system and set the following parameters: 15 ms pulses at 35 V with 950 ms intervals for in vivo electroporation.</li>
	</ol>
	</li>
	<li>Perfusion and Fixing Solution
	<ol>
		<li>Dissolve the paraformaldehyde (PFA) powder in 1x PBS at a concentration of 4% (w/v). Store the PFA solution at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Experimental Procedures
<ol>
	<li>Surgical Exposure of the L4 and L5 DRGs
	<ol>
		<li>Anesthetize the mouse using an intraperitoneal injection of the anesthetic solution #1 prepared previously (with ketamine 100 mg/kg body weight and xylazine 12 mg/kg body weight).</li>
		<li>Shave the surgical area with the fur clipper. 
		NOTE: Since there will be another surgery for left sciatic nerve crush, the fur of the left posterior thigh can be shaved simultaneously.</li>
		<li>Place the mouse on the heated blanket (35.0 &#176;C) and monitor the rectal temperature with a temperature probe.</li>
		<li>To test the anesthesia, pinch the toes and tail of the mouse and observe the behavioral responses.</li>
		<li>Tape the four limbs of the mouse on the corkboard.</li>
		<li>Wipe the surgical area with iodophor solution and then with 75% alcohol to remove the iodophor before cutting the skin.</li>
		<li>Apply ophthalmic ointment on eyes to prevent dryness while under anesthesia.</li>
		<li>Before incision, mark both sides of the iliac crests with a fine marker pen. Draw a line connecting two points of iliac crests to facilitate identifying the positions of L5 DRGs.</li>
		<li>Make a 3 cm incision along the midline of low back with micro-scissors. Moreover, detach the paraspinous muscles, such as the musculus multi&#64257;dus and the musculus longissimus lumborum, from the L3 to S1 spinous processes, and expose the facet joints of L4-5 and L5-6.</li>
		<li>Use a micro-rongeur to remove the facet joints of L4-5 and L5-6. Moreover, remove the left neural arch of L4 and L5 to expose the dorsal side of the DRGs.</li>
	</ol>
	</li>
	<li>DRG Injection
	<ol>
		<li>Inject anesthetic solution #2&#160;(with avertin 200 mg/kg body weight)&#160; intraperitoneally to maintain the anesthesia before DRG injection.</li>
		<li>Load the DNA plasmids or RNA oligos (1 &#181;L per DRG) into the glass capillary pipette.</li>
		<li>Insert the tip of the capillary glass pipette carefully into the DRG.</li>
		<li>Gradually inject 1.0 &#181;L solution of DNA plasmids or RNA oligos into the DRG using the intracellular microinjection dispense systems (30 psi, 8 ms). 
		NOTE: The duration of the injection should last no less than 5 minutes.</li>
	</ol>
	</li>
	<li>Electroporation
	<ol>
		<li>Drop PBS on tips of the electrodes.</li>
		<li>Clean up the bleeding with sterilized square cotton gauze.</li>
		<li>Pinch the target DRG with the electrodes gently and apply 5 square electric pulses with the electroporation system.</li>
		<li>Close the muscle and skin layers respectively with 5-0 nylon sutures.</li>
		<li>Place the mouse on a heated blanket (35.0 &#176;C) under close attention until it has completely regained sufficient consciousness to maintain sternal recumbency. Return the mouse to the home cage after it has fully recovered from the anesthesia. 
		NOTE: It takes approximately 60 min for the mouse to completely recover from anesthesia on the 37 &#176;C blanket.&#160;Dissolve one pill (200 mg) ibuprofen into 1,000 ml water (0.2 mg/ml) for daily feeding to relieve the post-surgical pain.</li>
	</ol>
	</li>
	<li>Sciatic Nerve Crush
	<ol>
		<li>Two or three days (depending on the experimental design) after the DRG electroporation, anesthetize the mouse intraperitoneally with anesthetic solution #2&#160;(with avertin 400 mg/kg body weight).</li>
		<li>Tape the four limbs of the mouse on the corkboard.</li>
		<li>Make a 1 cm incision 0.5 cm to the left side along the midline. Cut the muscles, such as gluteus maximus muscle and piriformis muscle, longitudinally. Expose the segment of the sciatic nerve between the greater sciatic foramen and the sciatic notch.</li>
		<li>Crush the nerve with microsurgery forceps for 12 s and mark the crush site with a 10-0 nylon epineural suture. Make a knot on the dural membrane to mark the crush site.</li>
		<li>Close the muscle and skin layers with 5-0 nylon suture.</li>
		<li>Place the mouse on a heated blanket (35.0 &#176;C) with close attention until it has completely regained sufficient consciousness to maintain sternal recumbency. Return the mouse to the home cage after it has fully recovered from the anesthesia. 
		NOTE: It takes approximately 60 min for the mouse to completely recover from anesthesia on the 37 &#176;C blanket.&#160;Dissolve one pill (200 mg) ibuprofen into 1,000 ml water (0.2 mg/ml) for daily feeding to relieve the post-surgical pain.</li>
	</ol>
	</li>
	<li>Mouse Perfusion, DRG and Sciatic Nerve Harvest
	<ol>
		<li>Two or three days after the sciatic nerve crush (depending on the experimental design), anesthetize the mouse intraperitoneally with anesthetic solution #2.</li>
		<li>Perfuse the mouse transcardially with PBS (pH 7.4) followed by ice-cold 4% paraformaldehyde (PFA) (pH 7.5) in PBS.</li>
		<li>After perfusion, separate the DRGs together with the nerve roots and sciatic nerve before the knee carefully with micro-scissors and micro-forceps under the dissection microscope. Place the sciatic nerve directly in 4% PFA overnight at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Imaging and Measuring the Fluorescence-Labeled Sensory Axons
	<ol>
		<li>Strip off the attached tissue and membrane on the fixed sciatic nerve with micro-scissors and micro-forceps carefully under the dissection microscope. Then, exchange the 4% PFA with PBS and wash the nerve three times.</li>
		<li>Place the sciatic nerve on a slide and keep it straight. Add 80 &#181;L of antifade solution around the nerve, then lay a coverslip on it. Flatten the whole mounted tissue with pressure.</li>
		<li>Place the &#64258;attened tissues on the inverted epi&#64258;uorescent microscope equipped with an accessory for mosaic acquisition and image processing. 
		NOTE: The excitation and emission lengths are 488 nm and 509 nm. It is also possible to take multiple overlapping images manually and stitch the images together using ImageJ with the Mosaic plug-in.</li>
		<li>When measuring the length of regenerated axons, trace all identi&#64257;able &#64258;uorescently-labeled axons in the sciatic nerve from the crush site (marked with the 10-0 epineural suture) to the distal axon ends.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Saijilafu, , 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=PI3K-GSK3+signalling+regulates+mammalian+axon+regeneration+by+inducing+the+expression+of+Smad1.">PI3K-GSK3 signalling regulates mammalian axon regeneration by inducing the expression of Smad1.</a> Nature Communications. 4, 2690 (2013).</li><li>Zhang, B. Y., 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=Akt-independent+GSK3+inactivation+downstream+of+PI3K+signaling+regulates+mammalian+axon+regeneration.">Akt-independent GSK3 inactivation downstream of PI3K signaling regulates mammalian axon regeneration.</a> Biochemical and Biophysical Research Communications. 443 (2), 743-748 (2014).</li><li>Jiang, J. 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=MicroRNA-26a+supports+mammalian+axon+regeneration+in+vivo+by+suppressing+GSK3beta+expression.">MicroRNA-26a supports mammalian axon regeneration in vivo by suppressing GSK3beta expression.</a> Cell Death &amp; Disease. 6, 1865 (2015).</li><li>Krames, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+the+dorsal+root+ganglion+in+the+development+of+neuropathic+pain.">The role of the dorsal root ganglion in the development of neuropathic pain.</a> Pain Medicine. 15 (10), 1669-1685 (2014).</li><li>Salimzadeh, L., Jaberipour, M., Hosseini, A., Ghaderi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-viral+transfection+methods+optimized+for+gene+delivery+to+a+lung+cancer+cell+line.">Non-viral transfection methods optimized for gene delivery to a lung cancer cell line.</a> Avicenna Journal of Medical Biotechnology. 5 (2), 68-77 (2013).</li><li>Keeler, A. M., ElMallah, M. K., Flotte, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+Therapy+2017:+Progress+and+Future+Directions.">Gene Therapy 2017: Progress and Future Directions.</a> Clinical and Translational Science. 10 (4), 242-248 (2017).</li><li>Neumann, E., Schaeferridder, M., Wang, Y., Hofschneider, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene-Transfer+into+Mouse+Lyoma+Cells+by+Electroporation+in+High+Electric-Fields.">Gene-Transfer into Mouse Lyoma Cells by Electroporation in High Electric-Fields.</a> The EMBO Journal. 1 (7), 841-845 (1982).</li><li>Liu, C. M., 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=MicroRNA-138+and+SIRT1+form+a+mutual+negative+feedback+loop+to+regulate+mammalian+axon+regeneration.">MicroRNA-138 and SIRT1 form a mutual negative feedback loop to regulate mammalian axon regeneration.</a> Genes &amp; Development. 27 (13), 1473-1483 (2013).</li><li>Saito, T., Nakatsuji, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+gene+transfer+into+the+embryonic+mouse+brain+using+in+vivo+electroporation.">Efficient gene transfer into the embryonic mouse brain using in vivo electroporation.</a> Developmental Biology. 240 (1), 237-246 (2001).</li><li>Saijilafu, E. M., Hur, F. Q., Zhou, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+dissection+of+axon+regeneration+via+in+vivo+electroporation+of+adult+mouse+sensory+neurons.">Genetic dissection of axon regeneration via in vivo electroporation of adult mouse sensory neurons.</a> Nature Communications. 2, 543 (2011).</li><li>Pan, C., 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=Shrinkage-mediated+imaging+of+entire+organs+and+organisms+using+uDISCO.">Shrinkage-mediated imaging of entire organs and organisms using uDISCO.</a> Nature Methods. 13, 859 (2016).</li><li>Rao, R. D., Bagaria, V. B., Cooley, B. 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The authors have nothing to disclose.

Several surgical steps require particular attention. The L4 and L5 DRGs (location of somas), which dominate the sciatic nerve, need to be correctly identified and injected with gene constructs. Otherwise, the GFP-labeling will be absent in sciatic nerve axons. The iliac crests can be viewed as useful anatomical landmarks to pinpoint L4 and L5 DRGs. In most mice, the facet joint between L5 and L6 vertebrae is proximate to iliac crests12. Alternatively, L3 DRG can be chosen instead of L5, especially for assays such as immunohistochemistry or western blot13, as surgical exposure of L5 DRG usually meets great difficulties due to a deep location and rich blood supply. Also, the whole surgery should avoid damaging DRG surrounding structures, including the spinal cord and the nerve root. The second critical operational step worth noting is the injection. The Fast Green dye was mixed with the solution of gene constructs to visualize the solution ejected from the needle and diffuse in the DRG. A successful injection should show a clear round-shape DRG outlined by the fluorescent dye. If the solution with fluorescent dye leaks out from the DRG while injecting, a back-and-forth fine-tuning on the depth of the tip of micro-needle can ensure complete injection of all solution into the DRG capsule. Avoid impaling the DRG at more than three different sites. Further, mild bleeding is usually inevitable after DRG injection, as DRGs are tightly encapsulated with rete venosum13. It is important to stop bleeding so that there will not be blood insulating the surface of DRG from electrodes during electroporation. Successful electroporation depends on electric current transduction, and PBS solution can be applied on the electrode. Finally, the site on the sciatic nerve where is crushed with forceps should be marked with micro-suturing, because the injury site is invisible under the microscope and needs to be indicated as the starting point of axon regeneration for image analysis later. If the epineural suture knot at the crush site falls off after animal perfusion and dissection, it is critical to re-label the same site with a suture immediately on the PFA-fixed nerve when the crush-induced indentation is still identifiable.
Among transfection techniques, electroporation has a higher transfection rate. According to our study, the transfection rate of the DRG neuron for microRNAs or siRNAs after in vivo electroporation is close to 90%. More importantly, in vivo electroporation is far less time-consuming than virus-based methods, which require virus packaging of desired gene constructs. As far as we know, even though the lipofectamine-based in vivo transfection has less toxicity than electroporation, the lipofectamine does not work on DRG neurons particularly, neither in vivo nor in vitro. Noteworthy, the current methodology has several technical limitations. First, the duration of siRNA efficacy to knocking down gene-of-interest is shorter than the virus-based delivery of plasmids. Therefore, the whole procedure from electroporation to animal sacrifice has to be finished in 4&#8211;6 days14. Additionally, the surgery performed under the microscope requires practice and some manual dexterity. For a novice, the surgery duration is often longer than expected. The anesthetic dose administrated to the mouse has to be carefully controlled to prevent unwanted mortality. Finally, flattening the sciatic nerve causes overlapping of axons when conducting epifluorescent imaging. Imaging the unflattened nerves with a confocal microscope is an alternative option.
Anatomically, the DRG sensory neurons have two axonal branches - the peripheral descending branch and the ascending central branch projecting into the dorsal column of the spinal cord10. The current methodology also shows distinctive labeling of the dorsal column of the spinal cord. Thus, a similar methodology can be used as a model to investigate sensory axon regeneration after spinal cord injury. Combined with tissue-clearing techniques11, conventional confocal microscopy or light-sheet microscopy can be employed on the cleared spinal cord sample to build 3D reconstructed images of sensory axons within the dorsal column of the spinal cord.

Injuries in the nervous system caused by neural trauma or various neurodegenerative diseases usually result in defects in motor, sensory and cognitive functions. Recently, much effort has been devoted to regenerative potency re-establishment in adult neurons to restore the physiological functionsof injured neurons1,2,3. Sensory neurons in the DRG are a cluster of nerve cells that convey different sensory stimuli, such as pain, temperature, touch, or body posture, to the brain. Each of these neurons is pseudo-unipolar and contains a single axon that bifurcates with one branch extending toward the periphery and the other branch heading toward the spinal cord4. The adult sensory neurons in DRGs are among a few mature mammalian neurons known to regenerate their axons actively after injury. Hence, injuries of sensory axons have been extensively employed as a crucial model to study the mechanisms of axonal regeneration in vivo.
In vivo gene transfection techniques, which are usually less time-consuming to set up and more flexible than using transgenic animals, have been playing essential roles in studying the functions of genes and signaling pathways in the nervous system. The main techniques can be categorized into two approaches: instrument-based and virus-based5. Viral-based in vivo gene delivery in adult neurons can provide precise spatiotemporal manipulation of gene expression6. However, labor-intensive processes are involved in viral-based methods, such as the production and purification of viral particles containing the desired gene. In addition, many viral vectors could activate the immune system of the host, which may interfere with the data acquisition, data analysis and possibly mislead the interpretation of experimental results. Electroporation, a typical instrument-based transfection approach, uses an electrical pulse to increase the permeability of cell and nuclear membranes transiently, which favors the influx of gene vectors or small RNA oligos from space outside of the cells7. In vitro electroporation is widely recognized as a transient but highly efficient strategy for manipulating targeted gene expression in many cell types. Although in vivo electroporation only leads to transient gene expression with low transfecting efficiency compared to viral vectors, it has various advantages over viral approaches. For instance, it can be applied to almost all tissues and cells7,8,9. In addition, either plasmids encoding genes-of-interest or small RNA oligos (e.g., siRNAs, microRNAs) against certain transcripts can be injected into the target tissue directly and then electrically pulsed, which make the procedure less labor- and time- consuming. Moreover, transfecting multiple plasmids and RNA oligos simultaneously with single electroporation is possible.
We have established an in vivo electroporation approach to manipulate gene expression in adult mouse sensory neurons and successfully applied and validated such approach in numerous pioneer studies1,2,3,8,10. Here, we present a detailed protocol to facilitate the usage of this approach for future studies of mammalian axon regeneration.

<table border="0" cellpadding="0" cellspacing="0" style="width:1039px;" width="1037">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="26">
			<td height="26" style="height:27px;width:335px;">ECM 830 Square wave electroporation system</td>
			<td style="width:232px;">BTX Harvard Apparatus</td>
			<td style="width:121px;">45-0052</td>
			<td style="width:351px;">For in vivo electroporation</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:27px;width:335px;">Tweezertrodes electrodes</td>
			<td style="width:232px;">BTX Harvard Apparatus</td>
			<td style="width:121px;">45-0524</td>
			<td>For in vivo electroporation, 1 mm flat</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Picospritzer III</td>
			<td style="width:232px;">Parker Instrumentation</td>
			<td style="width:121px;">1096</td>
			<td>Intracellular Microinjection Dispense Systems</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Glass Capillary Puller</td>
			<td style="width:232px;">NARISHIGE</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Borosilicate Glass Capillaries</td>
			<td>World Precision Instruments, Inc.</td>
			<td style="width:121px;">1902328</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Stereo Dissection Microscope</td>
			<td style="width:232px;">Leica</td>
			<td style="width:121px;">M80</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Microsurgery Rongeur</td>
			<td style="width:232px;">F.S.T</td>
			<td style="width:121px;">16221-14</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Microsurgery Forceps</td>
			<td style="width:232px;">FST by DUMONT, Switzerland</td>
			<td style="width:121px;">11255-20</td>
			<td>Only for sciatic nerve crush</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Glass Capillary</td>
			<td style="width:232px;">World Precision Instruments, Inc.</td>
			<td style="width:121px;">TW100-4</td>
			<td>10 cm, standard wall</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Tape</td>
			<td style="width:232px;">Fisherbrand</td>
			<td style="width:121px;">15-901-30</td>
			<td>For fixing the mouse on the corkboard</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">2, 2, 2-Tribromoethanol (Avertin)</td>
			<td style="width:232px;">Sigma-Aldrich</td>
			<td style="width:121px;">T48402</td>
			<td>Avertin stock solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">2-methyl-2-butanol</td>
			<td style="width:232px;">Sigma-Aldrich</td>
			<td style="width:121px;">152463</td>
			<td>Avertin stock solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">siRNA Fluorescent Universal Negative Control #1</td>
			<td style="width:232px;">Sigma-Aldrich</td>
			<td style="width:121px;">SIC003</td>
			<td>Non-target siRNA with fluorescence</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;width:335px;">microRNA Mimic Transfection Control with Dy547</td>
			<td style="width:232px;">Dharmacon</td>
			<td style="width:121px;">CP-004500-01-05</td>
			<td>Non-target microRNA with fluorescence</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Plasmids preparation kit</td>
			<td style="width:232px;">Invitrogen Purelink</td>
			<td style="width:121px;">K210016</td>
			<td>GFP-coding plasmid preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Fast Green Dye</td>
			<td style="width:232px;">Millipore-Sigma</td>
			<td style="width:121px;">F7252</td>
			<td>For better visualization of the DRG outline during injection</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:335px;">Ketamine</td>
			<td style="width:232px;">Putney, Inc</td>
			<td style="width:121px;">NDC 26637-731-51</td>
			<td>Anesthesia induction</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:335px;">Xylazine</td>
			<td style="width:232px;">AnaSed</td>
			<td style="width:121px;">NDC 59399-110-20</td>
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investigating mammalian axon regeneration: in vivo electroporation of adult mouse dorsal root ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

To quantify the cytotoxicity of the current protocol and to validate that transfection rate of in vivo DRG electroporation is high enough, we injected and electroporated fluorescently-tagged microRNA or siRNA into L4 and L5 DRGs. The detached DRGs were processed through cryo-sectioning and immunohistochemistry (Figure 1A-B). When estimating the cell survival rate after injection and electroporation, the intact DRGs from L4 and L5 were harvested and processed through cryo-sectioning and immunohistochemistry. The neuron densities, reflected with Tuj1 staining, did not show a significant difference when compared to the neuron densities of intact DRGs, which indicates that the electroporation did not induce neuronal cell death (Figure 1C). The transfection rate was calculated as the ratio between the number of tagged RNA oligo neurons and the number of Tuj1-positive neurons. The mean transfection rate of miRNA is 80.7 &#177; 4.3% and the siRNA 94.2 &#177; 0.3% (Figure 1D). When applying the current in vivo electroporation method to study axon regeneration, the sciatic nerve was flattened and imaged. Every regenerated axon with distinctive trajectory and recognizable distal axon end can be traced from the crush site indicated by the suture knot (Figure 2). Additionally, we electroporated the DRGs with EGFP plasmids and harvested the spinal cord from T7 to L2 along with dorsal roots of L4 and L5 after 7 days. We used a well-established tissue clearing protocol &#8212; uDISCO to clear the spinal cord11. The EGFP-labeled dorsal column axons in the spinal cord were imaged with a confocal microscope and then identified in both the longitudinal and the sagittal projecting images (Figure 3). Alternatively, the confocal images can be processed with microscopy visualization software to reconstruct a 3D image (Movie 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58171/58171fig1.jpg" /> 
Figure 1: Immunohistochemistry of DRG cryo-section after the electroporation of fluorescently-tagged non-specific microRNA or siRNA in vivo. (A) Representative cryo-section of DRG tissue injected and electroporated with fluorescently-tagged (Dy547) non-target microRNAs. Left image: the image of the fluorescent signal (red color) of fluorescently-tagged (Dy547) non-target microRNAs. Middle image: immuno-staining (green color) of Tuj1 with an Alexa488 fluorophore on the secondary antibody. Right image: the merged image of the previous two channels. Scale bar = 100 &#181;m.(B) Representative cryo-section of DRG tissue injected and electroporated with fluorescently-tagged (Cy3) non-target siRNAs. Left image: the image of the fluorescent signal (red color) of fluorescently-tagged (Cy3) non-target siRNAs. Middle image: immuno-staining (green color) of Tuj1 with an Alexa488 fluorophore on the secondary antibody. Right image: the merged image of the previous two channels. The scale bar represents 100 &#181;m. (C) The neuron density of an intact DRG cryo-section is 809.6 &#177; 14.2 cells/mm2 (N = 6) and an injected DRG cryo-section is 801.6 &#177; 27.4 cells/mm2 (N = 6), Mean &#177; SEM, Student&#39;s t-test, NS: no significance. Three mice were performed in vivo DRG electroporation on the left L4 and L5 DRGs with tagged siRNA. Both left and right L4 and L5 DRGs are harvested after 48 h and processed through cryo-section and immunohistochemistry. Each DRG has been sectioned into approximate 60 slices and the thickness of the slice is 10 &#956;m. Three slices of each DRG are selected by 200 &#956;m and averaged. (D) The transfection rate of the tagged miRNA is 80.7 &#177; 4.3% (N = 4) and the tagged siRNA is 94.2 &#177; 0.3% (N = 6), Mean &#177; SEM. Three mice were performed in vivo DRG electroporation on the left L4 and L5 DRGs with tagged siRNA and two mice with tagged miRNA. The DRGs are harvested after 48 h and processed through cryo-section and immunohistochemistry. Each DRG has been sectioned into approximate 60 slices and the thickness of the slice is 10 &#956;m. Three slices of each DRG are selected by 200 &#956;m and averaged. <a href="https://www.jove.com/files/ftp_upload/58171/58171fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58171/58171fig2.jpg" /> 
Figure 2: Ectopically expressed EGFP displays regenerated neuronal axons in sciatic nerve after crushing injury. The axons in a flattened sciatic nerve suffered from crush injury are labeled with EGFP (green fluorescence) transported from the somas to axons. The red line indicates the position of the crushing injury site, which was originally surgically marked by a suturing knot. The white arrowhead, arrow, and star indicate three distinctive axon ends, all of which extend from the crush site. Scale bar = 500 &#181;m. <a href="https://www.jove.com/files/ftp_upload/58171/58171fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58171/58171fig3.jpg" /> 
Figure 3: Projection images of EGFP-expressing axons in the spinal cord after in vivo electroporation of DRGs. (A) Longitudinal projection of the EGFP-labeled axons within the dorsal column and the L4/L5 dorsal roots. Inset images A1 and A2 show magnified views of the two rectangular areas marked in panel (A). A1 exhibits a detailed view of axons in the dorsal column. A2 exhibits a detailed view of axons in the dorsal root entry zone (DREZ). Scale bar = 200 &#181;m. (B) Sagittal projection of the EGFP-labeled axons within the dorsal column. Scale bar = 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/58171/58171fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/58171/Movie_1.jpg" /> 
Movie 1: 3D reconstruction of EGFP-expressing axons in the spinal cord after in vivo electroporation of DRGs. <a href="https://www.jove.com/files/ftp_upload/58171/Movie_1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion at JoVE.com

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

investigating-mammalian-axon-regeneration-vivo-electroporation-adult

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.1K Views.  Johns Hopkins University School of Medicine. Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Video: Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

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 ...

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...

An Approach to Enhance Alignment and Myelination of Dorsal Root Ganglion Neurons

Axon regeneration is a chaotic process due largely to unorganized axon alignment. Therefore, in order for a sufficient number of regenerated axons to bridge the lesion site, properly organized axonal alignment is required. Since demyelination after nerve injury strongly impairs the conductive capacity of surviving axons, remyelination is critical for successful functioning of regenerated nerves. Previously, we demonstrated that mesenchymal stem cells (MSCs) aligned on a pre-stretch induced anisotropic surface because the cells can sense a larger effective stiffness in the stretched direction than in the perpendicular direction. We also showed that an anisotropic surface arising from a mechanical pre-stretched surface similarly affects alignment, as well as growth and myelination of axons. Here, we provide a detailed protocol for preparing a pre-stretched anisotropic surface, the isolation and culture of dorsal root ganglion (DRG) neurons on a pre-stretched surface, and show the myelination behavior of a co-culture of DRG neurons with Schwann cells (SCs) on a pre-stretched surface.

This protocol describes the isolation of dorsal root ganglion (DRG) neurons isolated from rats and the culture of DRG neurons on a static pre-stretched cell culture system to enhance axon alignment, with subsequent co-culture of Schwann Cells (SCs) to promote myelination.

Axon regeneration is a chaotic process due largely to unorganized axon alignment. Therefore, in order for a sufficient number of regenerated axons to ...

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 ...

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive targets for injection of novel therapeutics aimed at treating chronic pain. In small animal models, laminectomy has been used to facilitate DRG injection because it involves surgical removal of the vertebral bone surrounding each DRG. We demonstrate a technique for intraganglionic injection of lumbar DRG in a large animal species, namely, swine. Laminotomy is performed to allow direct access to DRG using standard neurosurgical techniques, instruments, and materials. Compared with more extensive bone removal via laminectomy, we implement laminotomy to conserve spinal anatomy while achieving sufficient DRG access. Intraoperative progress of DRG injection is monitored using a non-toxic dye. Following euthanasia on post-operative day 21, the success of injection is determined by histology for intraganglionic distribution of 4&#39;,6-diamidino-2-phenylindole (DAPI). We inject a biologically inactive solution to demonstrate the protocol. This method could be applied in future preclinical studies to target therapeutic solutions to DRG. Our methodology should facilitate testing the translatability of intraganglionic small animal paradigms in a large animal species. Additionally, this protocol may serve as a key resource for those planning preclinical studies of DRG injection in swine.

We describe a method for laminotomy in swine that provides access to lumbar dorsal root ganglia (DRG) for intraganglionic injection. Injection progress is monitored intraoperatively and histologically confirmed up to 21 days after surgery. This protocol could be used for future preclinical studies involving DRG injection.

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive ...

Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of the PNS are affected by injury and disease, affecting the nerve function and the capacity for regeneration. Neuronal immune cells are commonly analyzed by immunofluorescence (IF). While IF is essential for determining the location of the immune cells in the nerve, IF is only semi-quantitative and the method is limited to the number of markers that can be analyzed simultaneously and the degree of surface expression. In this study, flow cytometry was used for quantitative analysis of leukocyte infiltration into sciatic nerves or dorsal root ganglions (DRGs) of individual mice. Single cell analysis was performed using DAPI and several proteins were analyzed simultaneously for either surface or intracellular expression. Both sciatic nerves from one mouse that were treated according to this protocol generated &#8805; 30,000 single nucleated events. The proportion of leukocytes in the sciatic nerves, determined by expression of CD45, was approximately 5% of total cell content in the sciatic nerve and approximately 5-10% in the DRG. Although this protocol focuses primarily on the immune cell population within the PNS, the flexibility of flow cytometry to measure a number of markers simultaneously means that the other cells populations present within the nerve, such as Schwann cells, pericytes, fibroblasts, and endothelial cells, can also be analyzed using this method. This method therefore provides a new means for studying systemic effects on the PNS, such as neurotoxicology and genetic models of neuropathy or in chronic diseases, such as diabetes.

Quantitative analysis of cell content within the murine sciatic nerve is difficult due to the scarcity of the tissue. This protocol describes a method for tissue digestion and preparation that provides sufficient cells for flow cytometry analysis of immune cell populations from nerves of individual mice.

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of ...

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 ...