The study has been funded by the Mont-le-Soie Equine Research Centre.

The commission for the ethical use of animals of the University of Liege approved the study protocol.
1. Muscle Microbiopsy
<ol>
	<li>Identify the sample site by visual inspection and palpation, in the long head of the triceps muscle, approximately midline between the point of the elbow and the point of the shoulder.</li>
	<li>Clip a zone of approximately 2 x 2 cm2 with an electrical clipper. Apply surgical scrub consisting of polyiodide liquid soap and alcohol compresses for 1 min over the clipped area. Inject subcutaneously 1 mL of 2% lidocaine solution with a 25 G needle in the center of the clipped and disinfected zone.</li>
	<li>Scrub hands with antibacterial hand soap. Put on sterile gloves. Use a disposable sterile drape as a sterile surface to place the biopsy needle and the trocar on.</li>
	<li>Arm the semi-automatic 14 G biopsy needle by pulling the spring mechanism as shown in Figures 1b and 1c. Release the biopsy needle to get familiar with its mechanism. Place the armed biopsy needle on the sterile surface. Get familiar with the trocar, which consists of a cannula and an obturator as shown in Figure 1a.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58023/58023fig1.jpg" /> 
Figure 1: The biopsy needle set. The biopsy needle set consists of (a) the cannula and its obturator and the biopsy needle (from top to bottom). The other panels show the biopsy needle in its (b) neutral and (c) armed position. <a href="https://www.jove.com/files/ftp_upload/58023/58023fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="5">
	<li>Prepare the trocar by assembling the cannula and the obturator and keep them in a locked position. Introduce the trocar perpendicularly to the skin through the formerly desensitized zone. Advance the trocar to a position where the tip of the trocar is approximately 1.5 cm deep in the muscle. Take the trocar out, dissociate the cannula from its obturator, and place the obturator on the sterile surface.</li>
	<li>Introduce the armed biopsy needle through the cannula. Introduce the needle and the cannula through the skin incision into the muscle. Advance the needle to a position where the tip of the needle is in the middle of the long head of the triceps muscle.</li>
	<li>Release the biopsy needle and then pull out the biopsy needle together with the cannula. Pull the biopsy needle out of the cannula. Leave the cannula on the sterile surface.</li>
	<li>Open the biopsy needle by pulling the spring with one hand and holding it with the other. 
	NOTE: The specimen becomes visible at the tip of the biopsy needle.</li>
	<li>With the help of a second person, open the sample tube that contains the sample medium. Use a 19 G hypodermic needle to remove the tiny muscle piece from the biopsy needle. Place the muscle sample in the sampling medium. Close the tube with the screw cap and gently tilt it 2x to ensure that the sample is floating in the medium.</li>
	<li>Arm the biopsy needle. Take the cannula from the sterile surface and reassemble the armed biopsy needle and cannula. Introduce the armed needle and the cannula through the skin incision as previously described. Repeat the sample procedure 2&#8211;3x until approximately 20 mg of muscle tissue is sampled. Close the sampling tube. Gently turn the tube 2x to mix the muscle pieces with the sampling medium.</li>
	<li>Ship the samples to the laboratory at a constant temperature between 4 to 8 &#176;C. 
	NOTE: The laboratory will then process the sample. The protocol can be paused here.</li>
	<li>The procedure is well tolerated and soreness at the sampling side is not observed. In the unlikely event of muscle soreness observed at the sampling site, administer appropriate analgesic treatment.</li>
</ol>
2. Treatment of the Cells
NOTE: The stem cells used in this study have been prepared according to the method described by Ceusters et al.7. Their article also describes the characterization of the cells.
<ol>
	<li>Start by preparing the specific culture medium, DF20, by adding 20% of heat-inactivated fetal bovine serum, 5 mL of penicillin (1,000 IU/mL)-streptomycin (10,000 &#181;g/mL), and 2.5 mL of amphotericin B (250 &#181;g/mL) to a bottle of 500 mL of a commercially available culture medium with glutamine and phenol red.</li>
	<li>Pour 150 &#181;L of the DF20 culture medium into each of the inner 16 wells of a 24-multi-well dish. Pour 1 mL of phosphate-buffered saline (PBS) [potassium chloride (200 mg/L), potassium phosphate monophasic (200 mg/L), sodium chloride (8,000 mg/L), and potassium phosphate diphasic (2,160 mg/L)] into the remaining outer wells.</li>
	<li>Rinse the tiny muscle pieces 2x in PBS. Separate them into very small pieces with the help of a scalpel and forceps and place a small piece (of the size of the tip of the scalpel blade) in each inner-16 well of the multi-well dish.</li>
	<li>Place the multi-well dish in an incubator maintained at the following conditions: 37 &#176;C, 21% O2, and 5% CO2. Monitor the wells every day under an inverted microscope and add 50 &#181;L of DF20 if necessary to prevent the cells from drying out. 
	NOTE: After approximately 10 d, there will be enough cells grown from the explant to allow the stem cell isolation.The protocol can be paused here.</li>
	<li>When a halo of cells is visible around the muscle explants, discard the explant of muscle tissue. Detach the cells by using 150 &#181;L of an EDTA-containing trypsin solution in each well: discard the culture medium, rinse the cells with 1 mL of PBS, discard the PBS, and add the trypsin solution for a maximum of 10 min at 37 &#176;C. Add culture medium to stop the action of the trypsin, harvest the cell suspension, and, then, centrifuge the cell suspension at 200 x g for 10 min at 37 &#176;C. Discard the supernatant and suspend the pellet in a balanced salt solution.</li>
	<li>Transfer the cell suspension on a discontinuous density gradient of 3 layers (15%, 25%, and 35%). Centrifuge the tube containing the cell suspension and the discontinuous density gradient solution at 1,250 x g for 20 min at 25 &#176;C. Do not use the brake of the centrifuge. Observe the cell fractions with different densities that will appear between the different layers of the discontinuous density gradient solution after the centrifugation.</li>
	<li>Continue the culture with the fraction between 15 and 25%. Transfer it to a tube and wash it with the balanced salt solution. Centrifuge the cell suspension at 200 x g for 10 min at 37 &#176;C. Discard the supernatant. Proceed with suspending the pellet in 1 mL of DF20. Prefill a cell culture flask, with a surface of 25 cm2, with 6 mL of DF20. Transfer the cells into the prefilled cell culture flask and culture them in an incubator with the following conditions: 37&#176;C, 21% O2, and 5% CO2. 
	NOTE: The protocol can be paused here.</li>
	<li>Monitor the cells every day under an inverted microscope. Change the culture medium if necessary (discard the old medium and add 6 mL of new culture medium). Determine the cellular confluence by observing the cell layers in the culture dishes, using an optical microscope at a predefined magnification. Estimate the surface of the dish that is occupied by the cells. Work at a fixed microscopic magnification for every observation to be able to evaluate the confluence. When the cells occupy 85% of the surface of the dish, proceed with a passage of the cells.</li>
	<li>When the cells are confluent, detach them using an EDTA-containing trypsin solution with the same protocol as described in step 2.4. Then, centrifuge the cell suspension at 200 x g for 10 min at 37 &#176;C. Discard the supernatant and proceed with the resuspension of the cells in 5 mL of DF20. Place them into a cell culture flask of 175 cm2 prefilled with 25 mL of DF20. Place the flask in an incubator with the following conditions: 37&#176;C, 21% O2, and 5% CO2. 
	NOTE: The protocol can be paused here.</li>
	<li>Monitor the cells every day under an inverted microscope. Change the medium if necessary (discard the old medium and add 30 mL of new medium). When the cells are confluent, detach them using trypsin-EDTA as described in step 2.4. Then, centrifuge the cell suspension at 200 x g for 10 min at 37 &#176;C. Discard the supernatant and suspend the cells in 6 mL of DF20. Place 1 mL of the cell suspension into six 175-cm2 flasks, each prefilled with 25 mL of DF20, and culture them at 37 &#176;C in a CO2 incubator (with 21% O2 and 5% CO2).</li>
	<li>Monitor the cells every day under an inverted microscope. Change the medium if necessary (discard the old medium and add 30 mL of new medium in each flask). When the cells are confluent, detach them using trypsin-EDTA as described in step 2.5. Centrifuge them 3x at 200 x g for 10 min at 37 &#176;C and discard the supernatant at each step.</li>
	<li>Count the cells with the help of a B&#252;rker counting chamber and a light microscope. Prepare a cellular suspension of 10 million cells/mL of a specific cryopreservation medium.</li>
	<li>Deep-freeze the therapeutic doses of 1 x 107 cells in 1 mL of cryopreservation medium and store them in the vapor phase of liquid nitrogen until their use. 
	NOTE: The protocol can be paused here.</li>
	<li>Ship the cells in the vial on dry ice for final use.</li>
</ol>
3. Injection of the Cells
NOTE: Two people are required to perform the injection of the stem cells.
<ol>
	<li>Warm the cell suspension progressively to room temperature. Aspirate the suspension into a 2 mL syringe.</li>
	<li>Sedate the horse by injecting detomidine 10 &#181;g/kg into the jugular vein with a 19 G hypodermic needle. Wait for 5 min to see the full effect of the sedation, which becomes visible by the characteristic head-down position of the horse.</li>
	<li>Clip a zone of 20 x 10 cm2 over the left laryngeal region of the horse with a clipper, as shown in Figure 2. Then, apply a surgical scrub of polyiodide soap and alcohol to the clipped region.</li>
	<li>Place a flexible standard video endoscope through the left nostril of the horse and advance it toward the nasopharynx. Adjust the position of the endoscope until a full view of both arytenoid cartilages and the epiglottis is obtained. Hold the endoscope in this position during the procedure and turn the screen of the video endoscope toward the operators.</li>
	<li>Connect the stimulation/injection needle to the nerve stimulator&#39;s negative electrode. Maintain the sterile conditions of the needle. Connect the positive electrode of the nerve stimulator to an electrode patch, which is stuck to the area of clipped skin.</li>
	<li>Connect the syringe containing the cell suspension to the injection line and prefill the tube to chase the air of the system. Remember the volume of the system and prepare a syringe with the same volume of sterile saline solution.</li>
	<li>If not familiar with this technique, use a disinfected 5&#160;to 7 MHz linear ultrasound transducer to visualize the anatomical structures in the region of interest. Use a sterile ultrasound coupling gel to increase the image quality. Visualize the larynx and the vessels that run in the dorsal region of the larynx.</li>
	<li>Once familiar with the anatomy of the region, introduce the stimulation/injection needle toward the dorsolateral aspect of the larynx. When approaching the dorsolateral aspect of the larynx, start the nerve stimulator in the 1 Hz stimulation mode with a current of 2 mA.</li>
	<li>Gently move the needle toward the position of the recurrent laryngeal nerve while observing the endoscopic view on the screen. As soon as the abduction movement of the left arytenoid cartilage is visible on the video screen, reduce the current until the movement disappears while keeping the needle in place.</li>
	<li>Identify the ideal position of the needle. Consider a loss of motor response at 0.5 mA as the ideal distance from the nerve. When arytenoid cartilage movements persist at currents lower than 0.4 mA, do not inject the cells, as that may result in an intraneuronal injection.</li>
	<li>When the loss of motor response at 0.5 mA is observed, inject the cell suspension containing 107 autologous stem cells and flush the injection line with the saline solution prepared in step 3.6. This will inject the rest of the cell suspension that remained in the injection tube.</li>
	<li>After the injection of the cells, take out the needle and the endoscope.</li>
	<li>Let the horse recover from the sedation. No further treatment is needed for the biopsy site.</li>
</ol>

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The authors have nothing to disclose.

This protocol describes the successful application of muscle-derived autologous stem cells to the recurrent laryngeal nerves in horses using a nerve stimulator-guided approach. The harvest of the muscle microbiopsy specimen, as well as the isolation, culture, and characterization of the stem cells, have been described in detail before, while the injection of these cells in proximity to a peripheral nerve is original. The electro-localization of the nerve with an electrical nerve stimulator has served for the present study and was used to inject mesenchymal stem cells in direct proximity to the recurrent laryngeal nerve. The motor response was monitored by video endoscopy through the nasopharynx. If, during the positioning process of the needle, other nerves were stimulated, the corresponding movement was visible on the endoscopy screen. A successful stimulation of the recurrent laryngeal nerve was characterized by the typical abduction of the arytenoid cartilage. In one horse, local anesthetic was injected to induce a transient paralysis of the arytenoid muscle. Although the successful implantation of the cells in proximity to the nerve has not been tested specifically in the present study, the electro-localization of the nerve and the injection at a low current proved that the stem cells were applied near the nerve. The proximity of the injectate to the nerve was further demonstrated by a successful motor block induced by an injection of a local anesthetic.
In the remaining five horses, the time from the first stimulation until the successful stem cell injection varied from 3 to 12 min. Horses were slightly sedated during the procedure, which increased their compliance with the electrical stimulation. None of the horses showed adverse reactions at any time, indicating that the duration of the stimulation can be prolonged if necessary to obtain good needle positioning. A steep learning curve is expected to be observed if the operator is using this technique regularly and in a larger number of horses with varying anatomy.
Horses commonly suffer from RLN, which is characterized by varying degrees of paralysis of the left arytenoid cartilage leading to abnormal respiratory sounds and exercise intolerance. A histology of affected nerves reveals typical lesions of peripheral neuropathy16. Peripheral neuropathy is a term used to describe various types of peripheral nerve lesions leading to impaired sensations, movements, or organ functions. The causes are highly variable and may include diabetes, nutrient deficiencies, a treatment with specific drugs, a traumatic injury, ischemia, or an infection, or they can be idiopathic. Independent of the cause, peripheral neuropathies share common histopathological signs, such as Wallerian degeneration, distal axonopathy, or segmental demyelination. It has been shown that the administration of mesenchymal stem cells to damaged peripheral nerves may be beneficial for their regeneration; however, the underlying mechanisms are not well understood. Traditionally, we believe that mesenchymal stem cells will only differentiate into progenitors of adipose, muscle, cartilage, and bone tissue, but under specific conditions, they have been shown to differentiate into myocytes17, astrocytes18, and myelinating cells of the peripheral19 and central nervous system19. During an in vitro culture, an exposure to neuropeptides will favor the differentiation of mesenchymal stem cells into cells expressing neuronal markers21,22. Several in vivo studies have demonstrated the beneficial effect of mesenchymal stem cells on nerve regeneration and a functional recovery in rat models of sciatic nerve injury10,23. This is probably caused by favorable environmental conditions that contribute to the differentiation of stem cells into tissue-specific cells24. Future studies should also investigate this technique for the administration of pre-differentiated cells in RLN-affected horses.
To the best of our knowledge, this is the first report that describes the use of a nerve stimulator to localize a peripheral nerve in order to inject a specific treatment into it. Other peripheral nerve pathologies in various species, such as neuropathic pain, can be treated by the administration of stem cells in proximity to the nerve25,26. Future studies should test the effect of this technique in clinical patients and investigate the risk of migration and the potential of differentiation of the injected cells.

RLN is a common pathology of the upper airway in horses characterized by varying degrees of arytenoid paralysis. The left side of the larynx is most commonly affected. The prevalence of the disease can reach up to 35% in certain horse populations. Although several hypotheses tried to explain the etiology and pathogenesis of this disease, the exact cause of RLN remains uncertain. The pathology is described as a distal axonopathy of the recurrent laryngeal nerve with lesions of demyelination but also some degree of remyelination. This axonopathy leads to the denervation of intrinsic laryngeal muscles and concomitant atrophy1,2. This pathology is often slowly progressive and may lead to a total loss of the arytenoid abduction of the affected side3.
Affected horses emit abnormal respiratory sounds during exercise and, sometimes, show exercise intolerance in more severe cases. The definitive diagnosis is made by the endoscopic examination on a non-sedated standing horse where a partial or total loss of laryngeal abduction is observed1,2,4. Currently, the most common treatment is laryngoplasty (also known as &#34;tie-back&#34;), which is sometimes associated with a ventriculo-cordectomy. Although the overall success rate of these surgeries is considered as good to excellent5, post-operative complications are very common. The most common complication is a gradual loss of abduction. Barnett et al.6 reported a loss of at least one grade of abduction within the first six weeks following the surgery in at least 76% of horses. Other complications, such as prosthesis failure, coughing, and airway contamination, are also reported5.
Mesenchymal stem cells (MSCs) have been a part of equine medicine in research and practice for more than a decade, although proven studies on their efficiency are still scarce. The two most commonly exploited sources of adult mesenchymal stem cells in horses are bone marrow and adipose tissue7. Both sampling techniques are relatively invasive and do not always lead to a sufficient number of cells. Recently, Ceusters et al.7 described the culture of stem cells derived from striated muscle tissue, which is obtained by a less invasive microbiopsy technique.
MSCs are capable of self-renewal, self-generation, multipotency, and differentiation7,8. Their ability to differentiate into all mesoderm lineages of fat, bone, muscle, and cartilage is now well established9. However, under specific environmental conditions, they can differentiate into non-mesenchymal lineages such as neurons, astrocytes, and myelinating cells of the peripheral nervous system and the spinal cord9,10. MSCs have already been used in several neuropathy models10,11. Their ability to migrate into areas of degenerated nervous tissue and to regenerate neural cells has been demonstrated after a systemic and local administration11. Moreover, Schwann-like cells derived from MSCs can recruit macrophages to remove cellular debris and secrete neurotrophic factors promoting axonal growth and remyelination9.
The aim of this study is to describe the technique of a nerve stimulator-guided injection of muscle-derived autologous stem cells near the left recurrent laryngeal nerve in healthy horses. Typically, a nerve stimulator connected to an injection needle is used for the electro-localization of peripheral nerves in order to apply local anesthetics to that area12. A weak direct current impulse is supplied in proximity to the nerve to induce a motor response. The ability to produce this motor response depends on several parameters, such as the conductive area of the needle, any impedance of the tissue, the current applied, the pulse duration, and the distance from the needle to the nerve. Nerve stimulator needles are designed to have a very restrictive conductive area at the tip of the needle, while the rest of the needle is insulated. This design helps to precisely localize the nerve. Modern nerve stimulators adapt to varying tissue impedances and deliver the constant amperage set on the machine. Furthermore, most machines use a pulse duration of 0.1 s, so that the determinative parameters are the current applied and the distance to the needle. The relationship between the needle-to-nerve distance and the current necessary to produce a motor response is described by Coulomb&#39;s law: E = kQ/r; where E is the required stimulation charge, k is Coulomb&#39;s constant, Q is the minimal required stimulation charge, and r is the distance between the two electrodes. In the electro-localization of nerves, the distance between the two electrodes is considered to be the needle-to-nerve distance12. Electrical charge dissipates following the rule of the inverse square of the needle-to-nerve distance13. Clinical practice has shown that the motor nerve stimulation at 0.5 mA is highly correlated to the successful nerve block and that a loss of motor signal at lower currents will prevent the user from administering accidental intraneuronal injections. The aim of this study is to test the feasibility and safety of this technique in a limited number of horses. If the feasibility and safety of this technique are confirmed, it can be easily transferred to affected horses. Further, equine RLN can serve as a model for peripheral degenerative neuropathy.

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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.

The commission for the ethical use of animals of the University of Liege approved the study protocol. Six horses from the herd of research horses at the Mont-le-Soie Equine Research Centre were included in this study. In the first horse, the recurrent laryngeal nerve was localized by electrostimulation. Figure 2 shows the setting used, with the stimulation needle inserted into the left dorsolateral aspect of the larynx of the horse in the nerve stimulator. When the loss of movement of the arytenoid cartilage is observed at 0.5 mA, 1 mL of 2% lidocaine is injected. This resulted in an absence of movement at higher currents and a transitory paralysis of the left arytenoid. The control endoscopy, 24 h later, revealed a complete recovery of the arytenoid function. The full protocol was successfully repeated in five horses. None of the horses showed a negative reaction to the muscle microbiopsy. In all horses, a sufficient number of cells were grown in the set time. The upper airway endoscopy was performed in all five horses and the laryngeal function scores of I or II.1 according to Robinson et al.14 were determined. All horses tolerated the nerve stimulation of the recurrent laryngeal nerve very well. No adverse reaction was observed during or after the stimulation or the injection of the stem cells.
All endoscopies were recorded and scored by two blinded clinicians. There was no difference between the pre-injection scores of the laryngeal functions and the scores obtained on day 1, 7, and 28 after the stem cell injection, as tested by Wilcoxon rank analysis15. Table 1 summarizes the laryngeal scores of all horses at different time points, as well as the time from the insertion of the needle to the injection of the stem cells and the current that provoked the arytenoid movement when the cells where injected.
The cells that have been used in the present study were prepared according to a protocol published previously7. The cells from all the fractions were able to trilineage differentiation, expressed CD90 and CD44, did not express CD45 and MHC II, and had clonogenic capacities. The cells of the 15&#8211;25% fraction have been chosen for further processing because they showed the greatest expression of CD90 and the biggest proliferative capacities.
The aim of the present study was not to test the efficiency of the stem cell application to regenerate a damaged peripheral nerve but only to test the feasibility of the procedure and to assess the safety of the stem cell injection to the recurrent laryngeal nerve in a small number of healthy horses. An absence of any functional changes on the endoscopic imaging in the five horses up to 28 days after the injection and an absence of any clinical signs in four of the five horses up to 1 year after the injection confirms the feasibility and the safety of the procedure. One horse had to be euthanized in the follow-up period for reasons unrelated to the study. Although the techniques prove to be safe in a small number of horses, the number of horses included in the present study is too low to demonstrate an absence of rare events.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td></td>
			<td>Score before injection</td>
			<td>Time to injection (min)</td>
			<td>Current at injection (mA)&#160;</td>
			<td>Score at day 1 post injection</td>
			<td>Score at day 7 post injection</td>
			<td>Score at day 28 post injection</td>
		</tr>
		<tr>
			<td>Horse 1 (Standardbred, mare, 16 years old)</td>
			<td>II.1</td>
			<td>12</td>
			<td>0.5</td>
			<td>I</td>
			<td>II.1</td>
			<td>II.1</td>
		</tr>
		<tr>
			<td>Horse 2 (Standardbred, mare, 22 years old)</td>
			<td>I</td>
			<td>3</td>
			<td>0.7</td>
			<td>I</td>
			<td>I</td>
			<td>I</td>
		</tr>
		<tr>
			<td>Horse 3 (Standardbred, mare, 11&#160; years old)</td>
			<td>II.1</td>
			<td>4</td>
			<td>0.5</td>
			<td>II.2</td>
			<td>II.1</td>
			<td>II.1</td>
		</tr>
		<tr>
			<td>Horse 4 (Standardbred, mare,&#160; 12 years old)</td>
			<td>I</td>
			<td>7</td>
			<td>0.5</td>
			<td>I</td>
			<td>II.1</td>
			<td>I</td>
		</tr>
		<tr>
			<td>Horse 5 (Standardbred, mare,&#160; 10 years old)</td>
			<td>I</td>
			<td>3</td>
			<td>0.7</td>
			<td>II.1</td>
			<td>I</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: Signalment and laryngeal function scores of the horses. This table shows the time of injection, the lowest current provoking any arytenoid movement before the injection, and the laryngeal scores of five healthy horses before and at day 1, 7, and 28 after the injection of autologous muscle-derived mesenchymal stem cells in proximity to the left laryngeal recurrent nerve. Horse #5 was not available for the control at day 28 after the injection for reasons unrelated to this study. The scores refer to the scores described by Robinson et al.14. Here, score I means that the movements of both arytenoid cartilages were synchronous and symmetrical, and a complete abduction of both arytenoid cartilages could be obtained and maintained. Score II.1 means that the movements of the arytenoid cartilages were asynchronous or may be asymmetric at certain times; however, a complete abduction of both arytenoid cartilages could be obtained and maintained.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58023/58023fig2.jpg" /> 
Figure 2: Setting during the injection of the stem cells. The nerve stimulation needle is introduced on the left side of the larynx and directed toward the left recurrent laryngeal nerve. The nerve stimulator is set at 0.8 mA. <a href="https://www.jove.com/files/ftp_upload/58023/58023fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Nerve Stimulator-guided Injection of Autologous Stem Cells Near the Equine Left Recurrent Laryngeal Nerve at JoVE.com

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

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 ...

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8.9K Views.  University of Liege. 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.

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

The Spared Nerve Injury (SNI) Model of Induced Mechanical Allodynia in Mice

Peripheral neuropathic pain is a severe chronic pain condition which may result from trauma to sensory nerves in the peripheral nervous system. The spared nerve injury (SNI) model induces symptoms of neuropathic pain such as mechanical allodynia i.e. pain due to tactile stimuli that do not normally provoke a painful response [1]. 
The SNI mouse model involves ligation of two of the three branches of the sciatic nerve (the tibial nerve and the common peroneal nerve), while the sural nerve is left intact [2]. The lesion results in marked hypersensitivity in the lateral area of the paw, which is innervated by the spared sural nerve. The non-operated side of the mouse can be used as a control. The advantages of the SNI model are the robustness of the response and that it doesn&#x2019;t require expert microsurgical skills.
The threshold for mechanical pain response is determined by testing with von Frey filaments of increasing bending force, which are repetitively pressed against the lateral area of the paw [3], [4]. A positive pain reaction is defined as sudden paw withdrawal, flinching and/or paw licking induced by the filament. A positive response in three out of five repetitive stimuli is defined as the pain threshold. 
As demonstrated in the video protocol, C57BL/6 mice experience profound allodynia as early as the day following surgery and maintain this for several weeks.

The Spared Nerve Injury SNI Model of Induced Mechanical Allodynia in Mice

The Spared Nerve Injury animal model is described here as a mouse model of peripheral neuropathic pain following partial denervation of the sciatic nerve by lesioning the tibial and common peroneal nerve branches, leaving the remaining sural nerve intact. Behavioral modification resulting from mechanical allodynia is quantified by von Frey filaments.

Peripheral neuropathic pain is a severe chronic pain condition which may result from trauma to sensory nerves in the peripheral nervous system. The spared ...

Chronic Constriction of the Sciatic Nerve and Pain Hypersensitivity Testing in Rats

Chronic neuropathic pain, resulting from damage to the central or peripheral nervous system, is a prevalent and debilitating condition, affecting 7-18% of the population1,2. Symptoms include spontaneous (tingling, burning, electric-shock like) pain, dysaesthesia, paraesthesia, allodynia (pain resulting from normally non-painful stimuli) and hyperalgesia (an increased response to painful stimuli). The sensory symptoms are co-morbid with behavioural disabilities, such as insomnia and depression. To study chronic neuropathic pain several animal models mimicking peripheral nerve injury have been developed, one of the most widely used is Bennett and Xie's (1988) unilateral sciatic nerve chronic constriction injury (CCI)3 (Figure 1). Here we present a method for performing CCI and testing pain hypersensitivity.
CCI is performed under anaesthesia, with the sciatic nerve on one side exposed by making a skin incision, and cutting through the connective tissue between the gluteus superficialis and biceps femoris muscles. Four chromic gut ligatures are tied loosely around the sciatic nerve at 1 mm intervals, to just occlude but not arrest epineural blood flow. The wound is closed with sutures in the muscle and staples in the skin. The animal is then allowed to recover from surgery for 24 hrs before pain hypersensitivity testing begins.
For behavioural testing, rats are placed into the testing apparatus and are allowed to habituate to the testing procedure. The area tested is the mid-plantar surface of the hindpaw (Figure 2), which falls within the sciatic nerve distribution. Mechanical withdrawal threshold is assessed by mechanically stimulating both injured and uninjured hindpaws using an electronic dynamic plantar von Frey aesthesiometer or manual von Frey hairs4. The mechanical withdrawal threshold is the maximum pressure exerted (in grams) that triggers paw withdrawal. For measurement of thermal withdrawal latency, first described by Hargreaves et al (1988), the hindpaw is exposed to a beam of radiant heat through a transparent glass surface using a plantar analgesia meter5,6. The withdrawal latency to the heat stimulus is recorded as the time for paw withdrawal in both injured and uninjured hindpaws. Following CCI, mechanical withdrawal threshold, as well as thermal withdrawal latency in the injured paw are both significantly reduced, compared to baseline measurements and the uninjured paw (Figure 3). The CCI model of peripheral nerve injury combined with pain hypersensitivity testing provides a model system to investigate the effectiveness of potential therapeutic agents to modify chronic neuropathic pain. In our laboratory, we utilise CCI alongside thermal and mechanical sensitivity of the hindpaws to investigate the role of neuro-immune interactions in the pathogenesis and treatment of neuropathic pain.

Due to the simplicity of surgery and the robust behavioural outcome, chronic constriction of the sciatic nerve is one of the pre-eminent animal models of neuropathic pain. Within 24 hrs following surgery, pain hypersensitivity is established and can be quantitatively measured using a von Frey aesthesiometer (mechanical test) and plantar analgesia meter (thermal test).

Chronic neuropathic pain, resulting from damage to the central or peripheral nervous system, is a prevalent and debilitating condition, affecting 7-18% of ...

Modeling Intracerebral Hemorrhage in Mice: Injection of Autologous Blood or Bacterial Collagenase

Spontaneous intracerebral hemorrhage (ICH) defines a potentially life-threatening neurological malady that accounts for 10-15% of all stroke-related hospitalizations and for which no effective treatments are available to date1,2. Because of the heterogeneity of ICH in humans, various preclinical models are needed to thoroughly explore prospective therapeutic strategies3. Experimental ICH is commonly induced in rodents by intraparenchymal injection of either autologous blood or bacterial collagenase4. The appropriate model is selected based on the pathophysiology of hemorrhage induction and injury progression. The blood injection model mimics a rapidly progressing hemorrhage. Alternatively, bacterial collagenase enzymatically disrupts the basal lamina of brain capillaries, causing an active bleed that generally evolves over several hours5. Resultant perihematomal edema and neurofunctional deficits can be quantified from both models. In this study, we described and evaluated a modified double injection model of autologous whole blood6 as well as an ICH injection model of bacterial collagenase7, both of which target the basal ganglia (corpus striatum) of male CD-1 mice. We assessed neurofunctional deficits and brain edema at 24 and 72 hr after ICH induction. Intrastriatal injection of autologous blood (30 &mu;l) or bacterial collagenase (0.075U) caused reproducible neurofunctional deficits in mice and significantly increased brain edema at 24 and 72 hr after surgery (p&lt;0.05). In conclusion, both models yield consistent hemorrhagic infarcts and represent basic methods for preclinical ICH research.

Clinically relevant animal models of intracerebral hemorrhage (ICH) are needed to extend our knowledge of hemorrhagic stroke and to examine novel therapeutic strategies. In this study, we describe and evaluate two ICH models that implement unilateral injections of either autologous whole blood or bacterial collagenase into the basal ganglia (corpus striatum) of mice.

Spontaneous intracerebral hemorrhage (ICH) defines a potentially life-threatening neurological malady that accounts for 10-15% of all stroke-related ...

The Sciatic Nerve Cuffing Model of Neuropathic Pain in Mice

Neuropathic pain arises as a consequence of a lesion or a disease affecting the somatosensory system. This syndrome results from maladaptive changes in injured sensory neurons and along the entire nociceptive pathway within the central nervous system. It is usually chronic and challenging to treat. In order to study neuropathic pain and its treatments, different models have been developed in rodents. These models derive from known etiologies, thus reproducing peripheral nerve injuries, central injuries, and metabolic-, infectious- or chemotherapy-related neuropathies. Murine models of peripheral nerve injury often target the sciatic nerve which is easy to access and allows nociceptive tests on the hind paw. These models rely on a compression and/or a section. Here, the detailed surgery procedure for the &#34;cuff model&#34; of neuropathic pain in mice is described. In this model, a cuff of PE-20 polyethylene tubing of standardized length (2 mm) is unilaterally implanted around the main branch of the sciatic nerve. It induces a long-lasting mechanical allodynia, i.e., a nociceptive response to a normally non-nociceptive stimulus that can be evaluated by using von Frey filaments. Besides the detailed surgery and testing procedures, the interest of this model for the study of neuropathic pain mechanism, for the study of neuropathic pain sensory and anxiodepressive aspects, and for the study of neuropathic pain treatments are also discussed.

Neuropathic pain is a consequence of a lesion or disease affecting the somatosensory system. The &#8220;cuff model&#8221; of neuropathic pain in mice consists of the implantation of a polyethylene cuff around the main branch of the sciatic nerve. Mechanical allodynia is tested using von Frey filaments.

Neuropathic pain arises as a consequence of a lesion or a disease affecting the somatosensory system. This syndrome results from maladaptive changes in ...

In Vivo Imaging and Tracking of Technetium-99m Labeled Bone Marrow Mesenchymal Stem Cells in Equine Tendinopathy

Recent advances in the application of bone marrow mesenchymal stem cells (BMMSC) for the treatment of tendon and ligament injuries in the horse suggest improved outcome measures in both experimental and clinical studies. Although the BMMSC are implanted into the tendon lesion in large numbers (usually 10 - 20 million cells), only a relatively small number survive (&#60;10%) although these can persist for up to 5 months after implantation. This appears to be a common observation in other species where BMMSC have been implanted into other tissues and it is important to understand when this loss occurs, how many survive the initial implantation process and whether the cells are cleared into other organs. Tracking the fate of the cells can be achieved by radiolabeling the BMMSC prior to implantation which allows non-invasive in vivo imaging of cell location and quantification of cell numbers.
This protocol describes a cell labeling procedure that uses Technetium-99m (Tc-99m), and tracking of these cells following implantation into injured flexor tendons in horses. Tc-99m is a short-lived (t1/2 of 6.01 hr) isotope that emits gamma rays and can be internalized by cells in the presence of the lipophilic compound hexamethylpropyleneamine oxime (HMPAO). These properties make it ideal for use in nuclear medicine clinics for the diagnosis of many different diseases. The fate of the labeled cells can be followed in the short term (up to 36 hr) by gamma scintigraphy to quantify both the number of cells retained in the lesion and distribution of the cells into lungs, thyroid and other organs. This technique is adapted from the labeling of blood leukocytes and could be utilized to image implanted BMMSC in other organs.

In Vivo Imaging and Tracking of Technetium-99m Labeled Bone Marrow Mesenchymal Stem Cells in Equine Tendinopathy

This protocol describes the radiolabeling of equine mesenchymal stem cells and their implantation into tendon injuries in the horse in order to determine cell survival and tissue distribution using gamma scintigraphy.

Recent advances in the application of bone marrow mesenchymal stem cells (BMMSC) for the treatment of tendon and ligament injuries in the horse suggest ...

Studying Pancreatic Cancer Stem Cell Characteristics for Developing New Treatment Strategies

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor initiation, metastasis and resistance to radio- and chemotherapy. Here we describe a specific methodology for culturing primary human pancreatic CSCs as tumor spheres in anchorage-independent conditions. Cells are grown in serum-free, non-adherent conditions in order to enrich for CSCs while their more differentiated progenies do not survive and proliferate during the initial phase following seeding of single cells. This assay can be used to estimate the percentage of CSCs present in a population of tumor cells. Both size (which can range from 35 to 250 micrometers) and number of tumor spheres formed represents CSC activity harbored in either bulk populations of cultured cancer cells or freshly harvested and digested tumors 1,2. Using this assay, we recently found that metformin selectively ablates pancreatic CSCs; a finding that was subsequently further corroborated by demonstrating diminished expression of pluripotency-associated genes/surface markers and reduced in vivo tumorigenicity of metformin-treated cells. As the final step for preclinical development we treated mice bearing established tumors with metformin and found significantly prolonged survival. Clinical studies testing the use of metformin in patients with PDAC are currently underway (e.g., NCT01210911, NCT01167738, and NCT01488552). Mechanistically, we found that metformin induces a fatal energy crisis in CSCs by enhancing reactive oxygen species (ROS) production and reducing mitochondrial transmembrane potential. In contrast, non-CSCs were not eliminated by metformin treatment, but rather underwent reversible cell cycle arrest. Therefore, our study serves as a successful example for the potential of in vitro sphere formation as a screening tool to identify compounds that potentially target CSCs, but this technique will require further in vitro and in vivo validation to eliminate false discoveries.

Pancreatic cancer stem cells (CSCs) can be expanded in vitro using the anchorage-independent sphere culture technique, which represents a powerful tool to study CSC biology and can serve as the first step to develop novel CSC-targeting therapies. Here the methodology for expanding, analyzing and targeting of pancreatic CSCs is provided.

Pancreatic ductal adenocarcinoma (PDAC) contains a subset of exclusively tumorigenic cancer stem cells (CSCs) which have been shown to drive tumor ...

Nerve-sparing Mid-urethral Obstruction (NeMO) in Female Small Rodents

Partial bladder outlet obstruction (pBOO) has a high prevalence, causes significant patient burden, and immense health care costs. The most common animal model to investigate bladder remodeling in pBOO are female rodents undergoing partial obstruction at the proximal urethra. Variability in the degree of obstruction and animal mortality are major concerns with proximal obstruction. Furthermore, dissecting around the proximal urethra and bladder neck jeopardizes bladder innervation.
We developed a nerve-sparing mid-urethral obstruction (NeMO) model for pBOO avoiding the disadvantages of the traditional model. We approached the urethra just inferior to the pubic symphysis, which obviated the need for laparotomy as well as for dissection in this area; also, the striated urethral sphincter remained untouched. We performed NeMO in female Sprague-Dawley rats (12 obstructions, 6 sham animals) as well as in female C57/bl6 mice (20 obstructions, 18 sham animals). After two weeks, we evaluated bladder function, bladder mass, and body mass.
We had no mortalities among obstructed- or sham-operated female rats; as described for the traditional proximal pBOO-method, we tied the suture around the proximal urethra and a temporarily placed 0.9 mm metal rod. NeMO induced an 85% increase in bladder mass after two weeks, average residual urine volume was 0.4 mL in partially obstructed rats while only 0.03 mL in sham animals. In mice, we tested 3 sizes of cannulas that we placed along the urethra when tying the suture. We found that using a 27-gauge cannula resulted in over 50% animal mortality; placing the 25-gauge cannula did not yield the desired response in increasing bladder mass; utilizing a 26-gauge cannula yielded favorable results with minimal animal mortality (1/8) yet a significant 2-fold increase in bladder mass.

Nerve-sparing Mid-urethral Obstruction NeMO in Female Small Rodents

Traditional modeling of partial bladder outlet obstruction in rodents is fraught with animal mortality. A denervation injury from dissection around the proximal urethra and bladder neck is also of major concern. We developed and evaluated a safe and reliable mid-urethral obstruction model, avoiding the shortcomings of the traditional model.

Partial bladder outlet obstruction (pBOO) has a high prevalence, causes significant patient burden, and immense health care costs. The most common animal ...

Cavernous Nerve Stimulation and Recording of Intracavernous Pressure in a Rat

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for erectile dysfunction. However, the methods used vary between laboratories, and pitfalls still exist. The goal of this study was to describe a surgical technique that would provide a reliable and reproducible model. By exposing the ischiocavernosus muscle at its point of insertion on the ischial tuberosity, the penile crus could be cannulated with minimal dissection and injury to the structures involved in erectile function. Repeated stimulation of the CN, without the need for lifting and drying, was achieved by using a 125 &#181;m bipolar silver electrode and biocompatible silicon glue to isolate the electrode-nerve complex. This method prevents neuropraxia by reducing stretching and drying the nerve and provides complete isolation of the nerve, negating electrical leakage and preventing stimulation of alternative pathways.

This study describes a simplified surgical procedure and technique for performing cavernous nerve stimulation with the isolation of the nerve-electrode complex using silicone glue and intracavernous pressure measurement.

The stimulation of the cavernous nerve (CN) and measurement of intracavernous pressure (ICP) have been used extensively to test and evaluate therapies for ...

Functional and Physiological Methods of Evaluating Median Nerve Regeneration in the Rat

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different methods of simulating postoperative physiotherapy are presented. Multiple standardized strategies are used to assess motor and sensory recovery using an MN model of peripheral nerve lesion and repair, thus permitting easy comparison of the results. Several options are included for providing a postoperative physiotherapy-like environment to rats that have undergone MN injuries. Finally, the paper provides a method to evaluate the recovery of the MN using several noninvasive tests (i.e., grasping test, pin prick test, ladder rung walking test, rope climbing test, and walking track analysis), and physiological measurements (infrared thermography, electroneuromyography, flexion strength evaluation, and flexor carpi radialis muscle weight determination). Hence, this model seems particularly appropriate to replicate a clinical scenario, facilitating extrapolation of results to the human species.
Although the sciatic nerve is the most studied nerve in peripheral nerve research, analysis of the rat MN presents various advantages. For example, there is a reduced incidence of joint contractures and automutilation of the affected limb in MN lesion studies. Furthermore, the MN is not covered by muscle masses, making its dissection easier than that of the sciatic nerve. In addition, MN recovery is observed sooner, because the MN is shorter than the sciatic nerve. Also, the MN has a parallel path to the ulnar nerve in the arm. Hence, the ulnar nerve can be easily used as the nerve graft for repairing MN injuries. Finally, the MN in rats is located in the forelimb, akin to the human upper limb; in humans, the upper limb is the site of most peripheral nerve lesions.

Presented is a protocol to produce different types of median nerve (MN) lesions and repair in the rat. Additionally, the protocol shows how to evaluate the functional recovery of the nerve using several noninvasive behavioral tests and physiological measurements.

The main goal of this investigation is to show how to create and repair different types of median nerve (MN) lesions in the rat. Moreover, different ...

Structured Motor Rehabilitation After Selective Nerve Transfers

After severe nerve injuries, selective nerve transfers provide an opportunity to restore motor and sensory function. Functional recovery depends both on the successful re-innervation of the targets in the periphery and on the motor re-learning process entailing cortical plasticity. While there is an increasing number of methods to improve rehabilitation, their routine implementation in a clinical setting remains a challenge due to their complexity and long duration. Therefore, recommendations for rehabilitation strategies are presented with the aim of guiding medical doctors and therapists through the long-lasting rehabilitation process and providing step-by-step instructions for supporting motor re-learning.
Directly after nerve transfer surgery, no motor function is present, and therapy should focus on promoting activity in the sensory-motor cortex areas of the paralyzed body part. After about two to six months (depending on the severity and modality of injury, the distance of nerve regeneration and many other factors), the first motor activity can be detected via electromyography (EMG). Within this phase of rehabilitation, multimodal feedback is used to re-learn the motor function. This is especially critical after nerve transfers, as muscle activation patterns change due to the altered neural connection. Finally, muscle strength should be sufficient to overcome gravity/resistance of antagonistic muscles and joint stiffness, and more functional tasks can be implemented in rehabilitation.

Here, we present a protocol for the motor rehabilitation of patients with severe nerve injuries and selective nerve transfer surgery. It aims at restoring the motor function proposing several stages in patient education, early-stage therapy after surgery and interventions for rehabilitation after successful re-innervation of the nerve&#8217;s target.

After severe nerve injuries, selective nerve transfers provide an opportunity to restore motor and sensory function. Functional recovery depends both on ...