We thank Dr. Alan Tessler for comments and editorial help. This work was supported by NIH NS062320.

1. Microscope set up and imaging preparation
<ol>
 <li>Our imaging set up consists of a Leica MZ16 fluorescent stereomicroscope with a fast shutter and a cooled CCD camera controlled by Metamorph software.</li>
 <li>Prepare a thermostatically controlled heating pad and adjust output to 32.5&deg;C to maintain the animal's body temperature during and following surgery.</li>
 <li>Warm sterile Ringer's solution or artificial cerebrospinal fluid (ACSF) to 32.5&deg;C in advance for irrigation of spinal cord during surgery.</li>
 <li>Anesthetize the animal with an intraperitoneal injection of xylazine (8 mg/kg) and ketamine (120 mg/kg) cocktail.</li>
 <li>Shave the upper back with small animal clippers and spread one small drop of hair removal lotion over the shaved area with cotton-tipped swabs. Minutes later, remove the applied lotion using 70% ethanol soaked gauze sponges.</li>
</ol>
2. Laminectomy and surgical exposure of the L5 dorsal root
<ol>
 <li>Place the animal on a heated (32.5&deg; C) pad and disinfect the skin with 70% ethanol soaked swabs.</li>
 <li>Under bright field illumination on the stereomicroscope, perform a midline incision (2- to 3-cm) in the skin of the back. If necessary, use cotton-tipped swabs to stop bleeding.</li>
 <li>Reflect the spinal musculature to expose the underlying lumbar vertebrae.</li>
 <li>Expose the L3-S1 spinal segments by right-sided hemi-laminectomy using small rongeurs. A partial laminectomy exposing 4-6 segments of the lumbar and sacral spinal cord is created by removing the right dorsal portion of the L5 vertebrae at the level of the iliac crest of the hip (location of L5 DRG) rostrally to the L2 vertebrae (2 vertebrae caudal to the last rib). Perfuse the cavity with warm sterile Ringer's solution.</li>
 <li>Position the animal on a support cushion (rolled cotton gauze) to flatten the vertebral column. Use retraction hooks to widen the exposed area.</li>
 <li>At this point (approximately 30 minutes after the first i.p. injection of anesthetic, a supplement (0.5X) should be injected subcutaneously in order to keep the animal fully anesthetized. Alternatively, gas anesthesia (2-4% isoflurane in 0.5L/min oxygen) can be used for repeated anesthetization during imaging sessions longer than 1 hour. </li>
</ol>
3. Rhizotomy/Dorsal root crush 
<ol>
 <li>Switch to fluorescence excitation to visualize YFP labeled (+) axons.</li>
 <li>Using the tip of a Sub-Q (26ga.) needle, perform a small incision in the dura overlying the L5 dorsal root (DR). Perfuse repeatedly with Ringer's solution and clean gently with cotton-tipped swabs.</li>
 <li>Identify the site to be crushed and insert one side of a fine forceps (Dumont #5) subdurally.</li>
 <li>Close the forceps gently but firmly, holding the medial portion of the L5 root for 10 seconds, and then gently release the forceps.</li>
 <li>Wash repeatedly with physiological solution and clean gently with cotton-tipped swabs.</li>
</ol>
4. Image acquisition and post-op procedures
<ol>
 <li>Obtain multiple images of the whole exposed area including the crush site and the DREZ before and right after the crush at both low and high magnification.</li>
 <li>Images are acquired either as single snapshots or as multiple streams of 10 to 20 frames acquired within 30- to 40-ms exposure time. In-focus images are then selected, and an overview montage is created later using Photoshop. </li>
 <li>To minimize scar formation, tightly apply a piece of thin synthetic matrix membrane (Biobrane), followed by artificial dura, on top of the exposed spinal cord. Be sure to cut pieces to fit precisely in the exposed spinal cord window so that they are adherent to the spinal cord.</li>
 <li>Close the musculature with sterile 5-0 sutures and close the midline incision with wound clips. </li>
 <li>Inject Ringer's solution (0.3- to 0.5-mL subcutaneously), and administer Buprenorphine as postoperative analgesia (0.05 mg/kg) subcutaneously every 12 h for 2 days.</li>
 <li>Keep the animal on a heating pad (34&deg;-35&deg; C) until recovered.</li>
</ol>
5. Repeated imaging
<ol>
<li> Anesthetize the animal and remove the wound clips and sutures.</li>
<li> Remove artificial dura and thin synthetic matrix membrane patches and keep them in a sterile tube containing Ringer's solution for later reuse.</li>
<li> Gently remove any accumulated connective tissue scar with the bent tip of a Sub-Q needle and fine forceps. Perfuse frequently with warm Ringer's solution.</li>
<li> Re-expose the operative field, including crush site and DREZ, relocate YFP+ axons imaged in previous sessions, and repeat the procedures described in Section 4.</li>
</ol>
6. Representative results: 
We have observed that, whereas they did not regenerate after a transection injury, almost all YFP+ axons grew through the site of injury by 3 days after crush (Figure 1)11. Typically, the next day after crush, we observed dying-back degeneration of proximal stump axons and fragmentation/degeneration of the same axons distal to the crush, which confirmed that axons had been appropriately damaged (e.g., Figure 1; Day 3 and 5). Several additional criteria are applied to unambiguously distinguish regenerating axons from axons that were spared or had recovered from the injury. These included the following: (1) regenerating axons show an expansion of the non-fluorescent portion of the YFP+ axon at the crush site due to proximal and distal degeneration (in contrast to narrowing of the unlabeled gap due to fluorescent cytoplasm refilling the crush site if axons survived the injury); (2) regenerating axons are much thinner, less brightly fluorescent, and more undulating than axons that survived the injury; (3) regenerating neurites are thinner and more dimly fluorescent than the degenerating fluorescent fragments of axons through which they extended; (4) in contrast to surviving or spared axons, regenerating axons stop at the DREZ; and (5) in contrast to surviving or spared axons, regenerating axons do not exhibit nodes of Ranvier. Figure 1 shows four superficial YFP+ axons immediately after crush (A1; colored arrows). Three days after the crush, all four axons extend a single neurite that grows through the crush site (A2). Five days after crush, neurites remain stable and there is no additional growth from these or other proximal axons (A3).
The regenerating neurites that crossed the crush site elongated through much thicker and brighter fluorescent fragments of a degenerating axon (i.e., endoneurial tubes), and arrive at the DREZ as early as 4 days after the crush (about 3mm/2 days)11. Repeated imaging of these axons and their tips every two or three days for two more weeks (Figure 2) revealed that they did not grow forward or retract, but remained immobile. The only noticeable change was swelling of the tips and shafts of some axons. These observations therefore demonstrate surprisingly quick and chronic immobilization of the regenerating axons at the DREZ.
<img src="/files/ftp_upload/3126/3126fig1.jpg" alt="Figure 1"/> 
Figure 1: Repeated imaging of L5 DR YFP+ axons at the site of dorsal root crush over 5 days. The medial portion of the L5 root was crushed (red arrowhead) and imaged on Days 0, 3, and 5 after the crush. The area of the crush is magnified in the right panels (A1-A3).
<img src="/files/ftp_upload/3126/3126fig2.jpg" alt="Figure 2"/> Figure 2: Repeated imaging of axons arrived at the DREZ over 20 days after L5 root crush. Day 4, three axons (colored arrows) arrived at the DREZ. The tips of these axons remain in the same location and have a similar appearance in subsequent imaging sessions on Days 7, 9, 13, 15, and 20. Positions of an axon tip relative to other axon tips and landmarks (asterisks) were used to determine axon motility between imaging sessions.

<ol>
	<li>Feng, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+neuronal+subsets+in+transgenic+mice+expressing+multiple+spectral+variants+of+GFP.">Imaging neuronal subsets in transgenic mice expressing multiple spectral variants of GFP.</a> Neuron. 28, 41-51 (2000).</li><li>Lichtman, J. W., Sanes, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Watching+the+neuromuscular+junction.">Watching the neuromuscular junction.</a> J Neurocytol. 32, 767-775 (2003).</li><li>Bishop, D. L., Misgeld, T., Walsh, M. K., Gan, W. B., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axon+branch+removal+at+developing+synapses+by+axosome+shedding.">Axon branch removal at developing synapses by axosome shedding.</a> Neuron. 44, 651-661 (2004).</li><li>Balice-Gordon, R. J., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+vivo+visualization+of+the+growth+of+pre-+and+postsynaptic+elements+of+neuromuscular+junctions+in+the+mouse.">in vivo visualization of the growth of pre- and postsynaptic elements of neuromuscular junctions in the mouse.</a> J Neurosci. 10, 894-908 (1990).</li><li>Trachtenberg, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+in+vivo+imaging+of+experience-dependent+synaptic+plasticity+in+adult+cortex.">Long-term in vivo imaging of experience-dependent synaptic plasticity in adult cortex.</a> Nature. 420, 788-794 (2002).</li><li>Pan, F., Gan, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+imaging+of+dendritic+spine+development+in+the+mouse+cortex.">Two-photon imaging of dendritic spine development in the mouse cortex.</a> Dev Neurobiol. 68, 771-778 (2008).</li><li>Grutzendler, J., Gan, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+imaging+of+synaptic+plasticity+and+pathology+in+the+living+mouse+brain.">Two-photon imaging of synaptic plasticity and pathology in the living mouse brain.</a> NeuroRx. 3, 489-496 (2006).</li><li>Kerschensteiner, M., Schwab, M. E., Lichtman, J. W., Misgeld, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+vivo+imaging+of+axonal+degeneration+and+regeneration+in+the+injured+spinal+cord.">in vivo imaging of axonal degeneration and regeneration in the injured spinal cord.</a> Nat Med. 11, 572-577 (2005).</li><li>Misgeld, T., Nikic, I., Kerschensteiner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+vivo+imaging+of+single+axons+in+the+mouse+spinal+cord.">in vivo imaging of single axons in the mouse spinal cord.</a> Nat Protoc. 2, 263-268 (2007).</li><li>Davalos, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+in+vivo+imaging+of+densely+populated+glia,+axons+and+blood+vessels+in+the+mouse+spinal+cord+using+two-photon+microscopy.">Stable in vivo imaging of densely populated glia, axons and blood vessels in the mouse spinal cord using two-photon microscopy.</a> J Neurosci Methods. 169, 1-7 (2008).</li><li>Maio, D. D. i. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+vivo+imaging+of+dorsal+root+regeneration:+Rapid+immobilization+and+presynaptic+differentiation+at+the+CNS/PNS+border.">in vivo imaging of dorsal root regeneration: Rapid immobilization and presynaptic differentiation at the CNS/PNS border.</a> Journal of Neuroscience. 31, 4569-4582 (2011).</li></ol>

No conflicts of interest declared.

Imaging DR regeneration directly in living mice is particularly challenging because it requires a substantial dorsal laminectomy to monitor axon growth over a wide area followed by multiple invasive surgical and anesthetic procedures in subsequent imaging sessions. Several strategies helped to overcome these challenges. First, successful imaging required reducing mouse mortality (approximately 25%) by minimizing the duration of anesthesia and bleeding, and by meticulous post-op care. The mortality was also reduced by usi...

<table border="1">
	<tbody>
		<tr>
			<th>Name of the reagent</th>
			<th>Company</th>
			<th>Catalogue number</th>
			<th>Comments</th>
		</tr>
		<tr>
			<td>H line thy1-YFP (2-4 months old, either sex) </td>
			<td>Jackson Laboratory (Bar Harbor, ME)</td>
			<td>003782</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Xylazine (AnaSed injection, sterile solution) </td>
			<td>Lloyd Laboratories, (Shenandoah, LA)</td>
			<td>4811</td>
			<td>8 mg/kg</td>
		</tr>
		<tr>
			<td>Ketamine (Ketamine hydrochloride injection, USP) </td>
			<td>Hospira, Inc. (Lake Forest, IL)</td>
			<td>2051</td>
			<td>120 mg/kg</td>
		</tr>
		<tr>
			<td>Buprenorphine (Buprenex injectable) (0.05 mg/kg)</td>
			<td>Reckitt Benckiser Pharmaceuticals Inc.(Richmond, VA)</td>
			<td>7571</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Small animal hair clippers </td>
			<td>Oster Professional, (McMinnville, TN)</td>
			<td>76059-030</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hair removal lotion</td>
			<td>Church &amp; Dwight Co (Princeton, NJ)</td>
			<td>NAIR with Baby Oil</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gauze sponges</td>
			<td>Fisher Scientific, (Pittsburgh, PA)</td>
			<td>22-362-173</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cotton-tipped swabs</td>
			<td>Fisher Scientific, (Pittsburgh, PA)</td>
			<td>14-960-3Q</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>1 mL syringes</td>
			<td>Becton, Dickson and Company Franklin Lakes, NJ)</td>
			<td>309602</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Subcutaneous (Sub-Q) needles, 26ga.</td>
			<td>Becton, Dickson and Company (Franklin Lakes, NJ)</td>
			<td>305115</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Spring scissors and forceps</td>
			<td>Fine Science Tools, (Foster City, CA)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>2.5-mm curved rongeurs </td>
			<td>Fine Science Tools, (Foster City, CA)</td>
			<td>16221-14</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Lactated Ringer's Injection USP</td>
			<td>B. Braun Medical Inc., (Irvine, CA)</td>
			<td>BBR-L7502</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Sterile saline solution</td>
			<td>APP Pharmaceuticals, (Schaumburg, IL)</td>
			<td>918610</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Thin synthetic matrix membrane (Biobrane)</td>
			<td>Bertek Pharmaceuticals, (Morgantown, WV)</td>
			<td>62794-096-251</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Artificial dura</td>
			<td>Gore Preclude MVP Dura Substitute, W.L. Gore and Associates, (Flagstaff, AZ)</td>
			<td>1MVP40</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>5-0 silk sutures</td>
			<td>Ethicon, Inc. (Somerville, NJ)</td>
			<td>K-580</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Wound clips</td>
			<td>Perfect &#x2013; Ets Bruneau, (Burnea, France)</td>
			<td>A75</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fluorescent stereomicroscope</td>
			<td>Leica Microsystems, (Wetzlar, Germany)</td>
			<td>MZ16</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CCD camera</td>
			<td>Hamamatsu, (Bridgewater, NJ)</td>
			<td>ORCA-Rx2</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>World Precision Instruments (Sarasota, FL)</td>
			<td>ATC 1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Metamorph software</td>
			<td>Molecular Devices, (Sunnyvale, CA)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Photoshop</td>
			<td>Adobe Systems, San Jose, CA</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Live Imaging of Dorsal Root Axons after Rhizotomy at JoVE.com

Live Imaging of Dorsal Root Axons after Rhizotomy

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. ...

live-imaging-of-dorsal-root-axons-after-rhizotomy

Research

JoVE Journal

Neuroscience

13.8K Views.  Shriners Hospitals Pediatric Research Center and Department of Anatomy and Cell Biology. 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.

Video: Live Imaging of Dorsal Root Axons after Rhizotomy

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 ...

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 ...

Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors (nAChRs) is an important mechanism for neuromodulation by acetylcholine (ACh). The difficulty with access to probing the signaling mechanisms within intact axons and at nerve terminals both in vitro and in vivo has limited progress in the study of the pre-synaptic components of synaptic plasticity. Here we introduce a gene-chimeric preparation of ventral hippocampal (vHipp)&#8211;accumbens (nAcc) circuit in vitro that allows direct live imaging to analyze both the pre- and post-synaptic components of transmission while selectively varying the genetic profile of the pre- vs post-synaptic neurons. We demonstrate that projections from vHipp microslices, as pre-synaptic axonal input, form multiple, reliable glutamatergic synapses with post-synaptic targets, the dispersed neurons from nAcc. The pre-synaptic localization of various subtypes of nAChRs are detected and the pre-synaptic nicotinic signaling mediated synaptic transmission are monitored by concurrent electrophysiological recording and live cell imaging. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of glutamatergic synaptic plasticity in vitro.

We developed a gene-chimeric preparation of ventral hippocampal &#8211; accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors ...

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 ...

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 ...

Dorsal Root Ganglia Isolation and Primary Culture to Study Neurotransmitter Release

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

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

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

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

Rapid Isolation of Dorsal Root Ganglion Macrophages

There are growing interests to study the molecular and cellular interactions among immune cells and sensory neurons in the dorsal root ganglia after peripheral nerve injury. Peripheral monocytic cells, including macrophages, are known to respond to a tissue injury through phagocytosis, antigen presentation, and cytokine release. Emerging evidence has implicated the contribution of dorsal root ganglia macrophages to neuropathic pain development and axonal repair in the context of nerve injury. Rapidly phenotyping (or &#8220;rapid isolation of&#8221;) the response of dorsal root ganglia macrophages in the context of nerve injury is desired to identify the unknown neuroimmune factors. Here we demonstrate how our lab rapidly and effectively isolates macrophages from the dorsal root ganglia using an enzyme-free mechanical dissociation protocol. The samples are kept on ice throughout to limit cellular stress. This protocol is far less time consuming compared to the standard enzymatic protocol and has been routinely used for our Fluorescence-activated Cell Sorting analysis.

Here we present a mechanical dissociation protocol to rapidly isolate macrophages from the dorsal root ganglion for phenotyping and functional analysis.

There are growing interests to study the molecular and cellular interactions among immune cells and sensory neurons in the dorsal root ganglia after ...

In Vitro Myelination of Peripheral Axons in a Coculture of Rat Dorsal Root Ganglion Explants and Schwann Cells

The process of myelination is essential to enable rapid and sufficient signal transduction in the nervous system. In the peripheral nervous system, neurons and Schwann cells engage in a complex interaction to control the myelination of axons. Disturbances of this interaction and breakdown of the myelin sheath are hallmarks of inflammatory neuropathies and occur secondarily in neurodegenerative disorders. Here, we present a coculture model of dorsal root ganglion explants and Schwann cells, which develops a robust myelination of peripheral axons to investigate the process of myelination in the peripheral nervous system, study axon-Schwann cell interactions, and evaluate the potential effects of therapeutic agents on each cell type separately. Methodologically, dorsal root ganglions of embryonic rats (E13.5) were harvested, dissociated from their surrounding tissue, and cultured as whole explants for 3 days. Schwann cells were isolated from 3-week-old adult rats, and sciatic nerves were enzymatically digested. The resulting Schwann cells were purified by magnetic-activated cell sorting and cultured under neuregulin and forskolin-enriched conditions. After 3 days of dorsal root ganglion explant culture, 30,000 Schwann cells were added to one dorsal root ganglion explant in a medium containing ascorbic acid. The first signs of myelination were detected on day 10 of coculture, through scattered signals for myelin basic protein in immunocytochemical staining. From day 14 onward, myelin sheaths were formed and propagated along the axons. Myelination can be quantified by myelin basic protein staining as a ratio of the myelination area and axon area, to account for the differences in axonal density. This model provides experimental opportunities to study various aspects of peripheral myelination in vitro, which is crucial for understanding the pathology of and possible treatment opportunities for demyelination and neurodegeneration in inflammatory and neurodegenerative diseases of the peripheral nervous system.

In the coculture system of dorsal root ganglia and Schwann cells, myelination of the peripheral nervous system can be studied. This model provides experimental opportunities to observe and quantify peripheral myelination and to study the effects of compounds of interest on the myelin sheath.

The process of myelination is essential to enable rapid and sufficient signal transduction in the nervous system. In the peripheral nervous system, ...