Name: ethanol-induced cervical sympathetic ganglion block applications for promoting canine inferior alveolar nerve regeneration using an artificial nerve
Uploaded: 2018-11-30T13:00:00.000+00:00
Duration: 6 min 48 s
Description: Watch this Scientific Journal Video about Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve at JoVE.c

This work was supported by the Department of Bioartificial Organs in Kyoto University Institute for Frontier Medical Science. We would like to thank the veterinary staff of the Institute for Frontier Medical Science.

This study was conducted in accordance with the Guiding Principles for the Care and Use of Animals and approved by the Committee for Animal Research of Kyoto University (Kyoto, Japan; authorization number: R-16-16). All efforts were made to minimize animal suffering, and all sections of this report adhere to the ARRIVE (Animal Research: Reporting of in Vivo Experiments) guidelines.
1. Fabrication of the PGA-C tube
<ol>
	<li>To fabricate the artificial nerve conduit by means of an absorbable polyglycolic acid (PGA) tube, use a tubular braiding machine equipped with 48 spindles and five PGA fibers, comprised of bundles of 26 filaments (Figure 1)18.</li>
	<li>To render the PGA tube surface hydrophilic, expose it to plasma discharge.</li>
	<li>Use 1% v/w atelocollagen in hydrochloride solution7. 
	NOTE: Atelocollagen is extracted from porcine skin via enzyme treatment and subjected to a virus check. It mainly consists of type I (70&#8211;80%) and type III collagen, the ratio of which is described in detail elsewhere7. Prepare the collagen solution by dissolving 1 g collagen in 100 mL hydrochloride solution (pH = 3.0). Since the density of the hydrochloride solution is approximately 1.0, the w/w collagen concentration is almost 1%.</li>
	<li>Coat the tube with the collagen layers by repeatedly dipping it into the 1% collagen hydrochloride solution for 5 s each time.
	<ol>
		<li>After dipping, dry the tube on a clean bench at room temperature. Perform next dipping after ensuring the tube is completely dry (about 6 h for air-drying).</li>
		<li>Repeat the coating process 10 times.</li>
	</ol>
	</li>
	<li>Subject the PGA-C tube to 140 &#176;C for 24 h under vacuum (dehydrothermal treatment), in order to control bio-absorption and crosslinking of the collagen molecules. Perform the entire process under aseptic conditions. 
	NOTE: This procedure generates a tube of 14-mm final length, 3-mm inner diameter, and 50-&#956;m wall thickness.</li>
</ol>
2. Surgical Procedure Set-up
<ol>
	<li>Use adult male beagles weighing 9.0 to 13.0 kg.
	<ol>
		<li>House animals in separate cages, under controlled kennel conditions (12-h light and dark cycle).</li>
		<li>Provide solid food and water ad libitum.</li>
	</ol>
	</li>
	<li>Weigh the beagles.</li>
	<li>Autoclave all surgical instruments.</li>
	<li>Don sterilized gloves and disinfect all surfaces of the operating setting with an 80% ethanol solution. Discard the used gloves.</li>
	<li>Perform surgical handwashing.</li>
	<li>Put on a fresh mask, gown, and sterile gloves.</li>
</ol>
3. Anesthesia and Skin Preparation
<ol>
	<li>Anesthetize the dog with a mixture of 5 mg/kg ketamine hydrochloride and 1 mg/kg xylazine by an intramuscular injection.</li>
	<li>Intubate by a tracheal tube of 7.5 mm diameter and 25 cm length.</li>
	<li>Place the dog on the right lateral position. Maintain general anesthesia with 3.2% sevoflurane with oxygen (1.0 L/min).</li>
	<li>Use a heating pad to maintain body temperature at 37 &#176;C.</li>
	<li>Apply an ophthalmic gel over the anterior surface of the eyes to avoid corneal abrasion.</li>
	<li>Carefully shave the surgical field (left side chest area) using surgical clippers. Spray a substantial amount of alcoholic solution over the operative site. Wait at least 15 sec. Repeat the application 3 times.</li>
	<li>Record the heart rate and oxygen saturation during surgery.</li>
</ol>
4. Inferior Alveolar Nerve Reconstruction Using PGA-C tube: Development of the Reconstruction-only Model
<ol>
	<li>Inject 3 mL of 1% lidocaine using a 27 G needle to the left mandibular gingiva as a local anesthetic and analgesic.</li>
	<li>Perform a 5-cm transverse incision with a number 15 scalpel blade in the left mandibular gingiva, to expose the mandibles of the animal.</li>
	<li>Use piezoelectric ultrasonic vibrations to grind the proximal aspect of the mandible into a 3-cm &#215; 8-mm rectangle through the posterior mental foramen. 
	NOTE: The vibration frequency was 28 &#8210; 32 kHz.</li>
	<li>Remove the frontal part of the mandibular bone plate (dimensions, 3 cm &#215; 8 mm) to expose the left IAN (Figure 2A)18. 
	NOTE: The reconstruction site corresponds to the root apex of the first molar</li>
	<li>Transect the IAN with a scalpel to remove a 10-mm segment.</li>
	<li>Insert the proximal and distal stumps of the severed nerve into the nerve tube to a depth of 2 mm.</li>
	<li>Use 8-0 nylon sutures and a surgical microscope at 8X magnification to suture the tube to the proximal and distal nerve ends (Figure 2B)18.</li>
	<li>Return the bone plate to its original site in the mandible.</li>
	<li>Close the wound with 4-0 nylon sutures.</li>
	<li>One day after surgery, confirm that the mandibular bone plate is in its proper position.
	<ol>
		<li>4.10.1 Perform computed tomography (CT) imaging of the facial bone under anesthesia. Set CT parameters as follows: 120 kVp, 200 mAs, 0.5 mm/s, 0.5-mm slice thickness.
		<ol>
			<li>Administer anesthesia using a mixture of 5 mg/kg ketamine hydrochloride and 1 mg/kg xylazine (Figure 3).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Administer ampicillin (100 mg/day) as an antibiotic and acetaminophen (100 mg/day) as an analgesic for a week after surgery.</li>
</ol>
5. Ethanol-induced CSGB: Development of the Reconstruction + CSGB Model
<ol>
	<li>Perform IAN reconstruction as described in section 4 and allow a week for recovery.</li>
	<li>Anesthetize the animal with 1.5% sevoflurane in oxygen (4 L/min) and air (6 L/min). Shave and clean the intended surgical field, as described in section 3.</li>
	<li>Mark the incision line with a surgical skin marker by drawing a line on the left side chest area (Figure 4, the incision line is 20 cm in length).</li>
	<li>Inject 5 mL of 1% lidocaine using a 21 G needle to the left side chest area as a local anesthetic and analgesic.</li>
	<li>Incise the left side chest skin with a number 10 scalpel blade.</li>
	<li>Incise the fat layer with an electric scalpel to expose the muscle fascia.</li>
	<li>Expose the serratus ventralis muscle and scalenus muscle.</li>
	<li>Raise the serratus ventralis muscle and scalenus muscle from ventral to dorsal to expose the second and third ribs (Figure 5).</li>
	<li>Perform a left lateral thoracotomy at the second and third intercostal space to expose the left cervical sympathetic ganglion (Figure 6).</li>
	<li>Inject 0.2 mL of 99.5% ethanol into the cervical sympathetic ganglion using a 30 G needle under direct visualization (Figure 7).</li>
	<li>Close the intercostal space with interrupted 1-0 absorbable stitches.</li>
	<li>Close the skin with interrupted 3-0 nylon stitches.</li>
	<li>Administer ampicillin (100 mg/day) as an antibiotic and acetaminophen (100 mg/day) as an analgesic for a week after surgery.</li>
	<li>At 1 week after CSGB, measure facial skin temperature with infrared thermography to confirm the CSGB.</li>
</ol>
6. Electrophysiological Recordings
<ol>
	<li>To measure sensory nerve action potential (SNAP) and sensory nerve conduction velocity (SCV) of the IAN three months after reconstruction, anesthetize animals as described in section 3. 
	NOTE: SNAP and SCV should be measured on both the experimental and normal control sides for each dog in both treatment groups.</li>
	<li>Make an incision in the left mandibular gingiva with a number 10 scalpel blade.</li>
	<li>Carefully remove the mandibular bone plate to avoid physically damaging the regenerated nerve.</li>
	<li>Stimulate the IAN using a pair of needle electrodes, to record the SNAP and SCV.
	<ol>
		<li>Insert the electrodes proximally to the nerve conduit.</li>
		<li>Apply 10-kHz electrical stimulus 20 times.</li>
	</ol>
	</li>
	<li>Analyze the results.
	<ol>
		<li>Determine SNAP by calculating the average response amplitude to the electrical stimulation.</li>
		<li>Measure the peak latency and peak amplitude from the chart recordings.</li>
		<li>Calculate the recovery index with the following equation: peak amplitude of the left IAN of the reconstruction-only or reconstruction + CSGB group / peak amplitude of the normal control (central segment of the right IAN in the reconstruction-only group)19,20.</li>
	</ol>
	</li>
</ol>
7. Histological Analysis
<ol>
	<li>Section Preparation
	<ol>
		<li>Three months after reconstruction, harvest the left IAN, including 1 cm of the nerve on either side of the reconstructed site.</li>
		<li>Harvest the right IAN at the level corresponding to the harvest site on the left side.</li>
		<li>Prefix the harvested nerves by immersion in 2.5% glutaraldehyde in a 0.1 M cacodylate buffer solution (pH 7.4, 48 &#176;C, 24 h).</li>
		<li>Postfix with 2% osmium tetroxide solution (48 &#176;C, 4 h) and potassium ferrocyanide in 0.1 M phosphate buffer solution (pH 7.4, 2 h).</li>
		<li>Dehydrate the nerves with a series of graded ethanol solutions.</li>
		<li>Embed in epoxy resin (paraffin).</li>
		<li>Section the specimens at a thickness of 0.5 &#8210; 1.0 &#956;m.</li>
	</ol>
	</li>
	<li>Toluidine Blue Staining and Morphological Analysis
	<ol>
		<li>Stain sections with toluidine blue solution.</li>
		<li>Obtain microscopy images using an optical microscope, at 400X magnification at the following regions along the samples: left IAN, the center of the regenerated segment and 2 mm distally to the stump; right IAN, the center of the IAN segment corresponding to the harvest site on the left side.</li>
		<li>Select images of all regions with regenerated nerve fibers.
		<ol>
			<li>Randomly select 8 &#8210; 10 areas of 100 &#956;m &#215; 100 &#956;m containing regenerated nerve fibers.</li>
			<li>Perform morphological analysis using an appropriate software to measure the following parameters: myelinated nerve fiber diameter (&#956;m) and density (count/area), nerve tissue percentage, and G-ratio (myelinated axon diameter/myelinated nerve fiber diameter).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Immunostaining
	<ol>
		<li>Follow standard protocols for paraffin section staining.</li>
		<li>Incubate with primary antibodies for 30 min at 25 &#176;C.</li>
		<li>Wash with phosphate-buffered saline 3 times at 25 &#176;C.</li>
		<li>Incubate with secondary antibodies labeled with horseradish-peroxidase for 30 min at 25 &#176;C.</li>
		<li>Obtain images using a light microscope.</li>
	</ol>
	</li>
	<li>Transmission Electron Microscopy (TEM) 
	<ol>
		<li>Prepare nerves as described in step 7.1.</li>
		<li>Section nerves at a thickness of 70 &#8210; 90 &#956;m using an ultramicrotome.</li>
		<li>Stain sections with Reynold&#8217;s lead citrate and uranyl.</li>
		<li>Examine and image by transmission electron microscopy.</li>
	</ol>
	</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Radiofrequency+lesions+of+the+stellate+ganglion+in+chronic+pain+syndromes:+retrospective+analysis+of+clinical+efficacy+in+86+patients.">Radiofrequency lesions of the stellate ganglion in chronic pain syndromes: retrospective analysis of clinical efficacy in 86 patients.</a> Clinical Journal of Pain. 16 (2), 164-168 (2000).</li><li>Ohno, K., Oshita, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transdiscal+lumbar+sympathetic+block:+a+new+technique+for+a+chemical+sympathectomy.">Transdiscal lumbar sympathetic block: a new technique for a chemical sympathectomy.</a> Anesthesia and Analgesia. 85 (6), 1312-1316 (1997).</li><li>Slappendel, R., Thijssen, H. O., Crul, B. J., Merx, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+stellate+ganglion+in+magnetic+resonance+imaging:+a+quantification+of+the+anatomic+variability.">The stellate ganglion in magnetic resonance imaging: a quantification of the anatomic variability.</a> Anesthesiology. 83 (2), 424-426 (1995).</li><li>Wang, Y. C., Wei, S. H., Sun, M. H., Lin, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+mode+of+percutaneous+upper+thoracic+phenol+sympathicolysis:+report+of+50+cases.">A new mode of percutaneous upper thoracic phenol sympathicolysis: report of 50 cases.</a> Neurosurgery. 49 (3), 628-634 (2001).</li></ol>

The authors have nothing to disclose.

We present an efficient method for IAN regeneration by using a bioabsorbable nerve tube in combination with ethanol-induced CSGB. For this study we used dogs, since other animal models, like mice, rats, and rabbits, have a short life expectancy and small body size, and hence cannot be used to perform the precise surgical procedures. As the IAN is located within the mandibular canal surrounded by bone, a surgical technique is necessary to avoid nerve and blood vessel damage when performing nerve reconstruction. An important technical tip for the procedure is to carefully remove the mandibular bone plate in order to minimize the risk of nerve and vessel injury. Traditional burs and micro saws cannot distinguish between hard and soft tissue21. Additionally, these tools tend to slip causing damaging the adjacent tissue, especially the IAN, by accidental contact22. We therefore used piezoelectric surgical tools for the bone processing steps. This is a new and innovative bone surgery technique that uses ultrasonic microvibrations of specialized scalpels. Therefore, soft tissue are not damaged even upon accidental contact with the cutting tips23,24. Microvibrations of 60 - 200 &#181;m/s at 24 - 29 kHz are optimal for cutting elastic mineralized tissue while sparing elastic soft tissue; this is not possible at frequencies above 50 kHz25. Moreover, rotational burs or oscillating saws require a force to counteract the rotation or vibration of the instrument. Compared to these instruments, piezoelectric surgical tools do not need application of extra force and so safe and accurate bone processing is possible26. This is especially important in the hands of a novel user.
Another important aspect in our method is that the bone plate was not fixed using metallic plates but was placed in its original position in the mandible after the placement of the PGA-C tube. The reason for this was to avoid the risk of exposure of the plate to oral mucosal necrosis, which arises when using metallic plates for fixation. However, in some cases the bone plate deviated from the original site. Therefore, it is crucial to perform a CT scan of the facial bone to confirm that the mandibular bone plate is in the proper position after surgery. When using a metallic plate for fixation, tight suturing should be avoided as it may cause oral mucosal necrosis due to blood flow disturbances.
CSGB is an effective treatment for peripheral vascular diseases and pain syndromes of the face and neck13,14,15. However, the mechanisms underlying its therapeutic effects remain unclear. One reason for the lack of research on the therapeutic effects of CSGB is the difficulty in obtaining a consistent and uniform sympatholytic effect. For example, the spread of sympathetic blockade after percutaneous CSGB is not uniform27,28. Mullenheim et al.29 implanted dogs with a polyethylene catheter, after thoracotomy, under the fascia and alongside the upper sympathetic chain, and performed CSGB by injecting lidocaine via the catheter. Although this approach can spread sympathetic blockade at the targeted areas, it carries risks of catheter occlusion or dislocation, as well as infection, especially in long-term experiments. In our canine model of CSGB, the direct injection of 99.5% ethanol produced long-term increases in blood flow to the ipsilateral orofacial region. In our approach, block injection was administered under direct visualization, so CSGB could be performed with precision. Therefore, our approach reduces the risk of uneven spread of the sympatholytic effect and of the sympathetic blockade. This is considered to be advantageous especially in long-term experiments. Moreover, we used thermography to confirm the success of CSGB, since facial skin temperature increases on the block side upon successful CSGB. In this case, thermography is useful, because it is simple and noninvasive. Importantly, the nasal and not the facial skin temperature should be measured, since it is not affected by the dog&#39;s hair. Our model could contribute to further research on the therapeutic effects of CSGB. In our previous study, blood flow in the orofacial area increased for 6-11 weeks after cervical sympathetic ganglionectomy. Researchers can choose this alternative method30 if desired.
The limitation of this study is that ethanol-induced CSGB poses the risk of developing permanent Horner&#39;s syndrome (ptosis and miosis)31. Other methods, including radiofrequency ablation, phenol, and sympathetic ganglionectomy have been used for performing sympathectomies; however, specifically for sympathectomy of the cervical sympathetic ganglion, only radiofrequency ablation has been employed in clinical practice32,33,34,35. Therefore, radiofrequency ablation and local anesthetics can be considered as an alternative method to ethanol-induced CSGB. Future studies would be necessary to validate how nerve regeneration mediated by a bioabsorbable nerve tube can be enhanced by CSGB with a local anesthetic or radiofrequency ablation.

In many cases, traumatic injury of the inferior alveolar nerve (IAN) is iatrogenic, being frequently caused by the extraction of the third molar or the placement of dental implants1,2,3. Injury of the IAN can lead to deficits in thermal and touch sensations as well as paresthesia, dysesthesia, hypoesthesia, and allodynia. Nerve injury is treated not only by conservative therapy but also by other methods, including suturing and autograft placement. However, these methods have drawbacks, which often include the lack of symptom improvement and neurological defects at the donor site4,5,6.
The artificial nerve &#8212; polyglycolic acid-collagen (PGA-C) tube was originally developed in Japan. It is a bio-absorbable tube with its inner lumen filled with a spongiform collagen7. In animal experiments, this tube was used to enhance nerve regeneration in beagle dogs with peroneal nerve defect, and was shown to promote higher level of recovery than autologous nerve transplantation8. The clinical application of the PGA-C tube began in 2002 in patients with peripheral nerve injuries. Moreover, favorable clinical outcomes have been achieved in the treatment of trigeminal neuropathy (IAN and lingual nerve)9,10,11. A critical factor for successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue8. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region and increases blood flow to the respective innervated area12; thus, it has been used in the treatment of complex regional pain syndrome and circulatory insufficiency13,14,15. However, there have been only a few experimental investigations on the efficacy of CSGB in increasing blood flow16,17. To ensure adequate CSGB efficiency, the blockade must be applied together with local anesthetics once or twice daily for several weeks, thus posing a challenge when generating animal models to investigate this technique. To address this limitation, in a previous study, we developed a canine model of long-term increased blood flow in the orofacial region18. The model was generated by performing a CSGB by injecting 99.5% ethanol. We evaluated the oral mucosal blood flow and nasal skin temperature by laser Doppler flowmetry and infrared thermography once per week for 12 weeks. We found that the blood flow of the orofacial region was increased for 7 &#8211; 10 weeks in this model.
In the present study, we evaluated the effects of ethanol-induced CSGB on nerve regeneration. 
The PGA-C tube was implanted into beagle dogs across a 10-mm gap in the left IAN. A week later, CSGB was performed by injecting ethanol. Three months after surgery, we performed a variety of electrophysiological, histological, and morphological studies to evaluate the effects of CSGB on nerve regeneration. We provide a detailed protocol for IAN reconstruction using a PGA-C tube and ethanol-induced CSGB.

<table><tbody><tr><td>NMP Collagen PS</td><td>Nippon Meatpackers</td><td>301-84621</td><td>Atelocollagen extracted from young porcine skin by enzyme treatment</td></tr><tr><td>Surgical clippers</td><td>Roboz Surgical Instrument Company</td><td>RC-5903</td><td></td></tr><tr><td>Disposable scalpel (No.15)</td><td>Kai medical</td><td>219ABBZX00073000</td><td></td></tr><tr><td>VarioSurg3</td><td>Nakanishi</td><td>VS3-LED-HPSC, E1133</td><td>Piezoelectric surgery for bone processing</td></tr><tr><td>4-0 nylon sutures</td><td>Ethicon</td><td>8881H</td><td></td></tr><tr><td>8-0 nylon sutures</td><td>Ethicon</td><td>2775G</td><td></td></tr><tr><td>Isepamicin sulfate</td><td>Nichi-Iko</td><td>620005641</td><td></td></tr><tr><td>Disposable scalpel (No.10)</td><td>Kai medical</td><td>219ABBZX00073000</td><td></td></tr><tr><td>30-gauge needle</td><td>Nipro</td><td>1134</td><td></td></tr><tr><td>1-0 absorbable stitches</td><td>Ethicon</td><td>J347H</td><td></td></tr><tr><td>3-0 Nylon stitches</td><td>Ethicon</td><td>8872H</td><td></td></tr><tr><td>Neo Thermo</td><td>NEC Avio</td><td>TVS-700</td><td>Infrared thermography&amp;nbsp;</td></tr><tr><td>Neuropack &amp;Sigma;</td><td>NIHON KOHDEN</td><td>MEB-5504</td><td>Orthodromic recorder for electrophysiological recording</td></tr><tr><td>Toluidine Blue</td><td>Sigma-Aldrich</td><td>T3260-5G</td><td></td></tr><tr><td>Light microscope</td><td>Keyence</td><td>BZ-9000</td><td></td></tr><tr><td>Mouse anti-human neurofilament protein monoclonal antibody</td><td>DAKO</td><td>N1591</td><td></td></tr><tr><td>Polyclonal rabbit anti-S100 antibody</td><td>DAKO</td><td>Z0311</td><td></td></tr><tr><td>Transmission electron microscopy</td><td>Hitachi High Technologies</td><td>Hitachi H-7000</td><td></td></tr><tr><td>Dynamic cell count</td><td>Keyence</td><td>BZ-H1C</td><td>Software for morphological evaluation</td></tr></tbody></table>

ethanol-induced cervical sympathetic ganglion block applications for promoting canine inferior alveolar nerve regeneration using an artificial nerve

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen films. Favorable clinical outcomes have been achieved when using these tubes for the treatment of damaged inferior alveolar nerve (IAN). A critical factor for the successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region thus increasing blood flow in the area. To ensure an adequate effect, the blockade must be administered with local anesthetics one to two times a day for several consecutive weeks; this poses a challenge when creating animal models for investigating this technique. To address this limitation, we developed an ethanol-induced CSGB in a canine model of long-term increase in blood flow in the orofacial region. We examined whether IAN regeneration via PGA-C tube implantation can be enhanced by this model. Fourteen Beagles were each implanted with a PGA-C tube across a 10-mm gap in the left IAN. The IAN is located within the mandibular canal surrounded by bone, therefore we chose piezoelectric surgery, consisting of ultrasonic waves, for bone processing, in order to minimize the risk of nerve and vessel injury. A good surgical outcome was obtained with this approach. A week after surgery, seven of these dogs were subjected to left CSGB by injection of ethanol. Ethanol-induced CSGB resulted in improved nerve regeneration, suggesting that the increased blood flow effectively promotes nerve regeneration in IAN defects. This canine model can contribute to further research on the long-term effects of CSGB.

We observed an increase in the facial skin temperature of the blocked side 1 week after the left CSGB (Figure 8).
At 3 months post-reconstruction, the PGA-C tube at the reconstruction area was absorbed and regeneration of the inferior alveolar nerve was observed in the reconstruction-only and reconstruction + CSGB groups (Figure 9A, B)18.
SNAP was measurable in both reconstruction sides of the reconstruction + CSGB and reconstruction-only groups. The results of the electrophysiological evaluation are summarized in Table 118. The recovery index and SCV were significantly higher in the reconstruction + CSGB than in the reconstruction-only group.
We observed myelinated nerve fibers at the central and distal segments of the regenerated IAN in the reconstruction-only and reconstruction + CSGB groups (Figure 10A, B)18. The reconstruction-only and reconstruction + CSGB groups showed smaller regenerated myelinated nerve diameters as compared to the normal control group (central segment of the right IAN in the reconstruction group, Figure 10C). Immature myelinated nerve fibers were also observed.
Examination of the reconstruction-only, reconstruction + CSGB groups using TEM showed regenerated myelinated nerve fibers and Schwann cells (Figure 10D, E). Figure 10F shows these results of TEM for the normal control group (central segment of the right IAN in the reconstruction group).
The presence of regenerated axons and Schwann cells was confirmed at the central and distal segments of the reconstruction-only and reconstruction + CSGB groups, by staining with anti-neurofilament (NF) and anti-S100 antibodies, respectively (Figure 11)18.
Morphological evaluation results are summarized in Table 218. The myelinated nerve fiber diameter at the center of the regenerated left IAN segment was 4.27 &#177; 1.5 &#181;m in the reconstruction group and 5.11 &#177; 1.98 &#181;m in the CSGB group, while at the distal segment of the regenerated left IAN the diameter was 3.47 &#177; 1.21 &#181;m in the reconstruction group and 4.53 &#177; 1.36 &#181;m in the CSGB group. In both cases, the diameter was significantly larger in the CSGB group, which also demonstrated a significantly higher myelinated nerve fiber density and nerve tissue percentage in both the center and the distal segment of the regenerated left IAN. The G-ratio at the center of the regenerated left IAN was 0.75 &#177; 0.04 in the reconstruction group and 0.68 &#177; 0.05 in the CSGB group, while at the distal part it was 0.74 &#177; 0.04 in the reconstruction group and 0.69 &#177; 0.04 in the CSGB group. Thus, in both cases, the G-ratio was significantly smaller in the CSGB group.
The sample sizes for the reconstruction-only and reconstruction + CSGB groups were n = 7. The statistical analyses for the myelinated nerve fiber diameter and density, G-ratio, and SCV were performed using Dunnett&#39;s test. Analysis of the recovery index was performed using an unpaired Student&#39;s t-test. The level of statistical significance was set at 5% (p &#60; 0.05).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58039/58039fig1.jpg" /> 
Figure 1: Polyglycolic acid tube filled with collagen sponge. A) Gross image of the tube. The final dimensions of the nerve conduit were 14 mm length, 3 mm inner diameter, and 50 &#181;m wall thickness. B) Scanning electron micrograph of the tube. This figure was previously published by Shionoya et al.18 and is reprinted with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig1large.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/58039/58039fig2.jpg" /> 
Figure 2: Left inferior alveolar nerve (IAN) pre- and post-reconstruction. A) Pre-reconstruction image of the left IAN after being exposed by removal of bone. B) Post-reconstruction image of the left IAN reconstructed using a polyglycolic acid-collagen tube. This figure was previously published by Shionoya et al.18 and is reprinted with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig2large.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/58039/58039fig3.jpg" /> 
Figure 3: Computed tomography imaging of the facial bone after left inferior alveolar nerve reconstruction. The image shows that the mandibular bone plate is in the proper position. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58039/58039fig4.jpg" /> 
Figure 4: Pre-operative skin markings on the left side chest area prior to surgery. Photograph showing the skin markings before performing the cervical sympathetic ganglion block. The incision line is 20 cm in length. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58039/58039fig5.jpg" /> 
Figure 5: Operating view of the cervical sympathetic ganglion block: pre-thoracotomy. Image shows the second and third ribs after raising the serratus ventralis and scalenus muscles. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58039/58039fig6.jpg" /> 
Figure 6: Operating view of the cervical sympathetic ganglion block: post-thoracotomy. Image shows the left cervical sympathetic ganglion after lateral thoracotomy at the second and third intercostal space. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58039/58039fig7.jpg" /> 
Figure 7: Cervical sympathetic ganglion pre- and post-ethanol injection using a 30 G needle. A) Pre-ethanol injection image of the left cervical sympathetic ganglion. B) Post-ethanol injection image of the left cervical sympathetic ganglion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/58039/58039fig8.jpg" /> 
Figure 8: Thermogram after left cervical sympathetic ganglion block (CSGB). The thermogram was acquired one week after CSGB by ethanol injection. Note that the facial skin temperature on the left side is higher than on the contralateral side. The color bar indicates temperatures in &#176;C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/58039/58039fig9.jpg" /> 
Figure 9: Regenerated inferior alveolar nerve (IAN). A) Image of the IAN in the reconstruction-only group. B) Image of the IAN in the reconstruction + CSGB (cervical sympathetic ganglion block) group. Nerve regeneration (region between white arrowheads) is observed in both groups. This figure was previously published by Shionoya et al.18 and is reprinted with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/58039/58039fig10.jpg" /> 
Figure 10: Toluidine blue and transmission electron microscopy analysis of the regenerated inferior alveolar nerve (IAN). A - C) Semi-thin transverse sections of the IAN at 3 months post-reconstruction stained with toluidine blue. Images show the distal segments of the regenerated left IAN in each group, as indicated. D - F) Transmission electron microscopy images from semi-thin sections showing myelinated and non-myelinated nerve fibers (black and white arrowheads, respectively). Scale bars represent 50 &#181;m in (A) - (C) and 5 &#181;m (D) - (F). Normal control: central segment of the right IAN in the reconstruction-only group. This figure was previously published by Shionoya et al.18 and is reprinted with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/58039/58039fig11.jpg" /> 
Figure 11: Immunohistochemical analysis of distal segments of the regenerated left inferior alveolar nerve (IAN). A, B) Sections of the regenerated IAN at 3 months post-reconstruction stained with an anti-neurofilament (NF) antibody for the reconstruction-only group (A) and reconstruction + cervical sympathetic ganglion block (CSGB; B) groups. Black arrowheads indicate regenerated axons. C, D) Sections of the regenerated IAN at 3 months post-reconstruction stained with an anti-S-100 antibody for the reconstruction-only group (C) and reconstruction + CSGB (D) groups. White arrowheads indicate Schwann cells. Scale bars, 50 &#181;m. This figure was previously published by Shionoya et al.18 and is reprinted with permission. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58039/58039fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Sensory nerve conduction (m/s)</td>
			<td>Recovery index</td>
		</tr>
		<tr>
			<td>Normal control</td>
			<td>48.5 &#177; 2.8</td>
			<td>&#65293;</td>
		</tr>
		<tr>
			<td>Reconstruction-only group</td>
			<td>36.8 &#177; 2.9*</td>
			<td>0.22 &#177; 0.04</td>
		</tr>
		<tr>
			<td>Reconstruction+CSGB group</td>
			<td>42.0 &#177; 2.4*#</td>
			<td>0.35 &#177; 0.06#</td>
		</tr>
	</tbody>
</table>
Table 1: Electrophysiological findings in theinferior alveolar nerve (IAN) at 3 months after surgery. Data are presented as mean &#177; standard deviation (n = 7). Comparisons were made using an unpaired Student&#39;s t-test. IAN, inferior alveolar nerve; CSGB, cervical sympathetic ganglion block. *, p &#60; 0.05 in comparison with the normal control group; #, p &#60; 0.05 in comparison with the reconstruction-only group. Normal control: central segment of the right IAN in the reconstruction-only group; Recovery index: ratio of the peak amplitude of the left IAN of the reconstruction-only or reconstruction + CSGB group to the peak amplitude of the normal control. This table was previously published by Shionoya et al.18 and is reprinted with permission.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2"></td>
			<td>Myelinated nerve fiber diameter (&#956;m)</td>
			<td>Myelinated nerve fiber density (count/100 &#956;m2)</td>
			<td>Proportion of nerve tissue (%)</td>
			<td>G ratio</td>
		</tr>
		<tr>
			<td>Normal control</td>
			<td>Center</td>
			<td>8.83 &#177; 3.11</td>
			<td>103 &#177; 8</td>
			<td>41.3 &#177; 3.9</td>
			<td>0.62 &#177; 0.03</td>
		</tr>
		<tr>
			<td rowspan="2">Reconstruction-only group</td>
			<td>Center</td>
			<td>4.27 &#177; 1.5*</td>
			<td>126 &#177; 20*</td>
			<td>11.6 &#177; 2.1*</td>
			<td>0.75 &#177; 0.04*</td>
		</tr>
		<tr>
			<td>Distal</td>
			<td>3.47 &#177; 1.21*</td>
			<td>109 &#177; 17*</td>
			<td>7.3 &#177; 2.0*</td>
			<td>0.74 &#177; 0.04*</td>
		</tr>
		<tr>
			<td rowspan="2">Reconstruction+CSGB group</td>
			<td>Center</td>
			<td>5.11 &#177; 1.98*#</td>
			<td>140 &#177; 22*#</td>
			<td>15.9 &#177; 3.0*#</td>
			<td>0.68 &#177; 0.05*#</td>
		</tr>
		<tr>
			<td>Distal</td>
			<td>4.53 &#177; 1.36*$</td>
			<td>123 &#177; 15*$</td>
			<td>12.5 &#177; 2.1*$</td>
			<td>0.69 &#177; 0.04*$</td>
		</tr>
	</tbody>
</table>
Table 2: Morphological findings in the inferior alveolar nerve (IAN) at 3 months after surgery. Data are presented as mean &#177; standard deviation (n = 7). Comparisons were made using Dunnett&#39;s test. IAN, inferior alveolar nerve; CSGB, cervical sympathetic ganglion block. *, p &#60; 0.05 in comparison with the normal control group;*#, p &#60; 0.05 in comparison with the central segment of the left IAN in the reconstruction-only group;*$,p &#60; 0.05 in comparison with the distal end of the left IAN in the reconstruction-only group. Normal control: central segment of the right IAN in the reconstruction-only group; G ratio is the ratio of the myelinated axon diameter to the total myelinated fiber diameter. This table was previously published by Shionoya et al.18 and is reprinted with permission.

Watch this Scientific Journal Video about Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve at JoVE.c

Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

We evaluated the effect of cervical sympathetic ganglion block on nerve repair using artificial nerve conduits. Male beagle dogs were each implanted with an artificial nerve across a 10-mm gap in the left inferior alveolar nerve; left cervical sympathetic ganglion was blocked by injecting 99.5% ethanol via lateral thoracotomy.

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen ...

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Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.8K Views.  Nippon Dental University Hospital at Tokyo. Deze methode kan helpen bij het beantwoorden van belangrijke vragen over zenuwregeneratie in het kunstmatige orgaanveld met behulp van een kunstmatige zenuwleiding. Het belangrijkste voordeel van deze techniek is dat de piëzo-elektrische chirurgie kan worden gebruikt voor inferieure alveololische zenuwreconstructie om schade aan de zenuwen en bloedvaten te voorkomen. De implicaties van deze techniek breiden zich uit tot de behandeling van orofaciale neuropathie, omdat het het potentieel heef...

Video: Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of communication with the brain. One of the goals of our research is to promote axon regeneration to restore connectivity across the lesion site. To accomplish this we developed a peripheral nerve (PN) grafting technique where segments of sciatic nerve are either placed directly between the damaged ends of the spinal cord or are used to form a bridge across the lesion. There are several advantages to this approach compared to transplantation of other neural tissues; regenerating axons can be directed towards a specific target area, the number and source of regenerating axons is easily determined by tracing techniques, the graft can be used for electrophysiological experiments to measure functional recovery associated with axons in the graft, and it is possible to use an autologous nerve to reduce the possibility of graft rejection. In our lab we have performed both autologous (donor and recipient are the same animal) and heterologous (donor and recipient are different animals) grafts with comparable results. This approach has been used successfully in both acute and chronic injury situations. Regenerated axons that reach the distal end of the PN graft often fail to extend back into the spinal cord, so we use microinjections of chondroitinase to degrade inhibitory molecules associated with the scar tissue surrounding the area of SCI. At the same time we have found that providing exogenous growth and trophic molecules encourages longer distance axonal regrowth into the spinal cord. Several months after transplantation we perform a variety of anatomical, behavioral and electrophysiological tests to evaluate the recovery of function in our spinal cord injured animals. This experimental approach has been used successfully in several spinal cord injury models, at different levels of injury and in different species (mouse, rat and cat). Importantly, the peripheral nerve grafting approach is effective in promoting regeneration by acute and chronically injured neurons.

Traumatic injury to the spinal cord disrupts communication with the brain. To restore lost connectivity we utilize a peripheral nerve graft to provide a substratum for regenerating fibers in combination with neurotrophic factors and matrix-modulating enzymes to remove inhibitory molecules to promote long distance growth.

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of ...

Biology

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

An Ultrasonic Tool for Nerve Conduction Block in Diabetic Rat Models

Nerve conduction block with a high intensity-focused ultrasound (HIFU) transducer has been performed in normal and diabetic animal models recently. HIFU can reversibly block the conduction of peripheral nerves without damaging the nerves while using an appropriate ultrasonic parameter. Temporary and partial block of the action potentials of nerves shows that HIFU has the potential to be a useful clinical treatment for pain relief. This work demonstrates the procedures for suppressing the action potentials of neuropathic nerves in diabetic rats in vivo using an HIFU transducer. The first step is to generate adult male diabetic neuropathic rats by streptozotocin (STZ) injection. The second step is to evaluate the peripheral diabetic neuropathy in STZ-induced diabetic rats by an electronic von Frey probe and a hot plate. The final step is to record in vivo extracellular action potentials of the nerve exposed to HIFU sonication. The method showed here may benefit the study of ultrasound analgesic applications.

This work presents the methodology of applying high intensity-focused ultrasound to block the action potentials of diabetic neuropathic nerves.

Nerve conduction block with a high intensity-focused ultrasound (HIFU) transducer has been performed in normal and diabetic animal models recently. HIFU ...

Bioengineering

Minimally-invasive Technique for Injection into Rat Optic Nerve

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central nervous system in a minimally-invasive surgery without bone removal. This technique describes an approach to visualization and direct injection of the optic nerve following minor fascial dissection from the orbital ridge, using a conjunctival traction suture to gently pull the eye down and out. Representative examples of an injected optic nerve show successful injection of dyed beads.

Direct injection into the rat optic nerve is useful for regenerative research. We demonstrate a minimally-invasive technique for direct injection into a rat optic nerve that does not involve opening the skull. Using this method, surgical complications are minimized and recovery is more rapid.

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central ...

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

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

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

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

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

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 ...

Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial nerve can be manipulated along its entire length, from its intracranial segment to its extratemporal course. There are three primary types of nerve injury used for the experimental study of regenerative properties: nerve crush, transection, and nerve gap. The range of possible interventions is vast, including surgical manipulation of the nerve, delivery of neuroactive reagents or cells, and either central or end-organ manipulations. Advantages of this model for studying nerve regeneration include simplicity, reproducibility, interspecies consistency, reliable survival rates of the rat, and an increased anatomic size relative to murine models. Its limitations involve a more limited genetic manipulation versus the mouse model and the superlative regenerative capability of the rat, such that the facial nerve scientist must carefully assess time points for recovery and whether to translate results to higher animals and human studies. The rat model for facial nerve injury allows for functional, electrophysiological, and histomorphometric parameters for the interpretation and comparison of nerve regeneration. It thereby boasts tremendous potential toward furthering the understanding and treatment of the devastating consequences of facial nerve injury in human patients.

This protocol describes a reproducible approach to facial nerve surgery in the rat model, including descriptions of various inducible patterns of injury.

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial ...

Nerve Stimulator-guided Injection of Autologous Stem Cells Near the Equine Left Recurrent Laryngeal Nerve

Recurrent laryngeal neuropathy (RLN) commonly affects horses and is characterized by abnormal respiratory sounds and exercise intolerance. The recurrent laryngeal nerve shows lesions of demyelination. The benefit of applying stem cells to demyelinated nerves has been demonstrated in various animal models. The aim of the study was to test the feasibility and safety of a peri-neuronal injection of autologous muscle-derived mesenchymal stem cells to the left recurrent laryngeal nerve in healthy horses by using an electrical nerve stimulator.
Muscle-derived stems cell are obtained from five healthy Standardbred horses by sampling 20 mg of muscle tissue with a semi-automatic 14 G biopsy needle from the triceps muscle. Movements of the larynx are monitored via upper-airway video endoscopy. The left recurrent laryngeal nerve is approached with an insulated nerve block needle. Nerve stimulation is applied, starting at 2 mA, and the successful abduction of the left arytenoid is monitored. The stimulation intensity is reduced progressively. When a loss of the motor response is observed at 0.5 mA, 107 autologous muscle-derived stem cells are injected. Two examiners, who are blinded to the time point, score the laryngeal function of the horses prior to the treatment and at day 1, day 7, and day 28 after the injection of the cells. In a sixth horse, 1 mL of 2% lidocaine is injected to further confirm the correct positioning of the needle. This leads to a temporary paralysis of the left arytenoid cartilage.
This study proves that the recurrent laryngeal nerve can be approached with the help of an electrical nerve stimulator and that the electrical stimulation of the nerve is well tolerated by the horses. No modification of the laryngeal function was observed in any of the horses after the injection of the stem cells. Further studies should be conducted to describe the effects of a peri-neuronal injection of autologous muscle-derived mesenchymal stem cells to horses suffering from RLN.

Here, we present a protocol to inject autologous muscle-derived stem cells in the proximity to the recurrent laryngeal nerve with the help of an electrical nerve stimulator. This novel technique may become useful for the treatment of equine recurrent laryngeal neuropathy.

Recurrent laryngeal neuropathy (RLN) commonly affects horses and is characterized by abnormal respiratory sounds and exercise intolerance. The recurrent ...

Medicine

Partial Sciatic Nerve Ligation: A Mouse Model of Chronic Neuropathic Pain to Study the Antinociceptive Effect of Novel Therapies

Management of chronic pain remains challenging to this day, and current treatments are associated with adverse effects, including tolerance and addiction. Chronic neuropathic pain results from lesions or diseases in the somatosensory system. To investigate potential therapies with reduced side effects, animal pain models are the gold standard in preclinical studies. Therefore, well-characterized and well-described models are crucial for the development and validation of innovative therapies. 
Partial ligation of the sciatic nerve (pSNL) is a procedure that induces chronic neuropathic pain in mice, characterized by mechanical and thermal hypersensitivity, ongoing pain, and changes in limb temperature, making this model a great fit to study neuropathic pain preclinically. pSNL is an advantageous model to study neuropathic pain as it reproduces many symptoms observed in humans with neuropathic pain. Furthermore, the surgical procedure is relatively fast and straightforward to perform. Unilateral pSNL of one limb allows for comparison between the ipsilateral and contralateral paws, as well as evaluation of central sensitization. 
To induce chronic neuropathic hypersensitivity, a 9-0 non-absorbable nylon thread is used to ligate the dorsal third of the sciatic nerve. This article describes the surgical procedure and characterizes the development of chronic neuropathic pain through multiple commonly used behavioral tests. As a plethora of innovative therapies are now being investigated to treat chronic pain, this article provides crucial concepts for standardization and an accurate description of surgeries required to induce neuropathic pain.

Partial sciatic nerve ligation induces long-lasting chronic neuropathic pain, characterized by exaggerated responses to thermal and mechanical stimuli. This mouse model of neuropathic pain is commonly used to study innovative therapies for pain management. This article describes in detail the surgical procedure to improve standardization and reproducibility.

Management of chronic pain remains challenging to this day, and current treatments are associated with adverse effects, including tolerance and addiction. ...

Inducing a Facial Nerve Injury in a Rat Model

Source: Ali, S. A., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59224">Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration.</a> J. Vis. Exp. (2020)
This video demonstrates a surgical protocol on a rat model, detailing steps from anesthesia preparation to facial nerve injury induction. This protocol is useful for studying nerve regeneration and surgical techniques in medical research.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

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