Name: investigating mammalian axon regeneration: in vivo electroporation of adult mouse dorsal root ganglion
Uploaded: 2018-09-01T14:30:00.000+00:00
Duration: 6 min 17 s
Description: Watch this Scientific Journal Video about Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion at JoVE.com

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>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Posterolateral+intertransverse+lumbar+fusion+in+a+mouse+model:+surgical+anatomy+and+operative+technique.">Posterolateral intertransverse lumbar fusion in a mouse model: surgical anatomy and operative technique.</a> Spine Journal. 7 (1), 61-67 (2007).</li><li>Dommisse, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+blood+supply+of+the+spinal+cord.+A+critical+vascular+zone+in+spinal+surgery.">The blood supply of the spinal cord. A critical vascular zone in spinal surgery.</a> The Journal of Bone and Joint Surgery. British Volume. 56 (2), 225-235 (1974).</li><li>Sorensen, D. 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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><tbody><tr><td>ECM 830 Square wave electroporation system</td><td>BTX Harvard Apparatus</td><td>45-0052</td><td>For &lt;em&gt;in vivo&lt;/em&gt; electroporation</td></tr><tr><td>Tweezertrodes electrodes</td><td>BTX Harvard Apparatus</td><td>45-0524</td><td>For &lt;em&gt;in vivo&lt;/em&gt; electroporation, 1 mm flat</td></tr><tr><td>Picospritzer III</td><td>Parker Instrumentation</td><td>1096</td><td>Intracellular Microinjection Dispense Systems</td></tr><tr><td>Glass Capillary Puller</td><td>NARISHIGE</td><td>PC-10</td><td></td></tr><tr><td>Borosilicate Glass Capillaries</td><td>World Precision Instruments, Inc.</td><td>1902328</td><td></td></tr><tr><td>Stereo Dissection Microscope</td><td>Leica</td><td>M80</td><td></td></tr><tr><td>Microsurgery Rongeur</td><td>F.S.T</td><td>16221-14</td><td></td></tr><tr><td>Microsurgery Forceps</td><td>FST by DUMONT, Switzerland</td><td>11255-20</td><td>Only for sciatic nerve crush</td></tr><tr><td>Glass Capillary</td><td>World Precision Instruments, Inc.</td><td>TW100-4</td><td>10 cm, standard wall</td></tr><tr><td>Tape</td><td>Fisherbrand</td><td>15-901-30</td><td>For fixing the mouse on the corkboard</td></tr><tr><td>2, 2, 2-Tribromoethanol (Avertin)</td><td>Sigma-Aldrich</td><td>T48402</td><td>Avertin stock solution</td></tr><tr><td>2-methyl-2-butanol</td><td>Sigma-Aldrich</td><td>152463</td><td>Avertin stock solution</td></tr><tr><td>siRNA Fluorescent Universal Negative Control #1</td><td>Sigma-Aldrich</td><td>SIC003</td><td>Non-target siRNA with fluorescence</td></tr><tr><td>microRNA Mimic Transfection Control with Dy547</td><td>Dharmacon</td><td>CP-004500-01-05</td><td>Non-target microRNA with fluorescence</td></tr><tr><td>Plasmids preparation kit</td><td>Invitrogen Purelink</td><td>K210016</td><td>GFP-coding plasmid preparation</td></tr><tr><td>Fast Green Dye</td><td>Millipore-Sigma</td><td>F7252</td><td>For better visualization of the DRG outline during injection</td></tr><tr><td>Ketamine</td><td>Putney, Inc</td><td>NDC 26637-731-51</td><td>Anesthesia induction</td></tr><tr><td>Xylazine</td><td>AnaSed</td><td>NDC 59399-110-20</td><td>Anesthesia induction</td></tr><tr><td>Acetaminophen</td><td>McNeil Consumer Healthcare</td><td>NDC 50580-449-36</td><td>Post-surgical pain relief</td></tr></tbody></table>

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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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

8.2K Views.  Johns Hopkins University School of Medicine. Electroporation is een effectieve aanpak te leveren van genen van belangen in cellen. Door deze aanpak in vivo toe te passen op de neuronen van volwassen muis achterwortelganglia ganglion (DRG), beschrijven we een model voor het bestuderen van axon regeneratie 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 ...

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

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

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

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

Live Imaging of Dorsal Root Axons after Rhizotomy

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

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

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

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 ...

Deciphering Axonal Pathways of Genetically Defined Groups of Neurons in the Chick Neural Tube Utilizing in ovo Electroporation

Employment of enhancer elements to drive expression of reporter genes in neurons is a widely used paradigm for tracking axonal projection. For tracking axonal projection of spinal interneurons in vertebrates, germ line-targeted reporter genes yield bilaterally symmetric labeling. Therefore, it is hard to distinguish between the ipsi- and contra-laterally projecting axons. Unilateral electroporation into the chick neural tube provides a useful means to restrict expression of a reporter gene to one side of the central nervous system, and to follow axonal projection on both sides 1 ,2-5. This video demonstrates first how to handle the eggs prior to injection. At HH stage 18-20, DNA is injected into the sacral level of the neural tube, then tungsten electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator. The egg is sealed with tape and placed back into an incubator for further development. Three days later (E6) the spinal cord is removed as an open book preparation from embryo, fixed, and processed for whole mount antibody staining. The stained spinal cord is mounted on slide and visualized using confocal microscopy.

Deciphering Axonal Pathways of Genetically Defined Groups of Neurons in the Chick Neural Tube Utilizing in ovo Electroporation

This video demonstrates how to visualize axonal pathways of genetically defined groups of neurons in the embryonic chick spinal cord utilizing in ovo electroporation of reporter genes under the control of specific enhancer elements.

Employment of enhancer elements to drive expression of reporter genes in neurons is a widely used paradigm for tracking axonal projection. For tracking ...

Biology

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

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 ...

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

Coculture of Axotomized Rat Retinal Ganglion Neurons with Olfactory Ensheathing Glia, as an In Vitro Model of Adult Axonal Regeneration

Olfactory ensheathing glia (OEG) cells are localized all the way from the olfactory mucosa to and into the olfactory nerve layer (ONL) of the olfactory bulb. Throughout adult life, they are key for axonal growing of newly generated olfactory neurons, from the lamina propria to the ONL. Due to their pro-regenerative properties, these cells have been used to foster axonal regeneration in spinal cord or optic nerve injury models.
We present an in vitro model to assay and measure OEG neuroregenerative capacity after neural injury. In this model, reversibly immortalized human OEG (ihOEG) is cultured as a monolayer, retinas are extracted from adult rats and retinal ganglion neurons (RGN) are cocultured onto the OEG monolayer. After 96 h, axonal and somatodendritic markers in RGNs are analyzed by immunofluorescence and the number of RGNs with axon and the mean axonal length/neuron are quantified.
This protocol has the advantage over other in vitro assays that rely on embryonic or postnatal neurons, that it evaluates OEG neuroregenerative properties in adult tissue. Also, it is not only useful for assessing the neuroregenerative potential of ihOEG but can be extended to different sources of OEG or other glial cells.

We present an in vitro model to assess olfactory ensheathing glia (OEG) neuroregenerative capacity, after neural injury. It is based on a coculture of axotomized adult retinal ganglion neurons (RGN) on OEG monolayers and subsequent study of axonal regeneration, by analyzing RGN axonal and somatodendritic markers.

Olfactory ensheathing glia (OEG) cells are localized all the way from the olfactory mucosa to and into the olfactory nerve layer (ONL) of the olfactory ...

Enrichment of Adult Mouse Dorsal Root Ganglia Neuron Cultures by Immunopanning

Dorsal root ganglia (DRGs) are peripheral structures adjacent to the dorsal horn of the spinal cord, which house the cell bodies of sensory neurons as well as various other cell types. Published culture protocols often refer to whole dissociated DRG cultures as being neuronal, despite the presence of fibroblasts, Schwann cells, macrophages, and lymphocytes. While these whole DRG cultures are sufficient for imaging applications where neurons can be discerned based on morphology or staining, protein or RNA homogenates collected from these cultures are not primarily neuronal in origin. Here, we describe an immunopanning sequence for cultured mouse DRGs. The goal of this method is to enrich DRG cultures for neurons by removing other cell types. Immunopanning refers to a method of removing cell types by adhering antibodies to cell culture dishes. Using these dishes, we can negatively select against and reduce the number of fibroblasts, immune cells, and Schwann cells in culture. This method allows us to increase the percentage of neurons in cultures.

This paper describes an immunopanning protocol for adult mouse dorsal root ganglia. By adhering antibodies to culture plates, we can negatively select and remove non-neuronal cells. We show that the cultures are enriched for neurons using this protocol, allowing for an in-depth study of neuronal responses to manipulation.

Dorsal root ganglia (DRGs) are peripheral structures adjacent to the dorsal horn of the spinal cord, which house the cell bodies of sensory neurons as ...

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

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