S.A.A. is funded by the American Academy of Facial Plastic and Reconstructive Surgery Leslie Bernstein Grants Program.

All interventions were performed in strict accordance with the National Institutes of Health (NIH) guidelines. The experimental protocol was approved by the University of Michigan&#39;s Institutional Animal Care &#38; Use Committee (IACUC) prior to implementation. Ten-week-old adult female Sprague-Dawley rats were utilized.
1. Prior to the operative day
<ol>
	<li>Ensure an appropriate stock of sterilized surgical instruments, analgesic medications, anesthetic medication, and oxygen prior to the operating day. Please see Table of Materials for a complete list.</li>
</ol>
2. Preoperative setup
<ol>
	<li>Ensure an adequate working space, including room for at least two individuals (the surgeon and an assistant). 
	NOTE: There is need for a dedicated operating table, room for the anesthesia machine setup, and adequate storage space for sterilized and backup supplies.</li>
	<li>Calibrate an operating microscope for use during the procedures. Make sure the surgeon has the ability to adjust the handles of the microscope and the zoom/focus buttons by placing a sterilized cover over the handles/buttons 
	NOTE: We utilized sterilized aluminum foil over the handles/buttons.</li>
</ol>
3. Anesthesia and preparation
<ol>
	<li>Place the animal in the anesthesia chamber and induce general anesthesia via 1.8% isoflurane and 0.9 L/min oxygen.
	<ol>
		<li>Confirm an adequate plane of anesthesia via an assessment of spontaneous breathing and an evaluation of consciousness by assessing the animal&#39;s grimace response to a toe pinch.</li>
	</ol>
	</li>
	<li>Apply eye lubricant bilaterally to guard against corneal irritation or dryness.</li>
	<li>Shave the operative site(s) with a razor or automatic clipper.
	<ol>
		<li>Establish a method for rat identification at this time, either via an ear tag or tail label/marking.</li>
	</ol>
	</li>
	<li>Administer a subcutaneous injection of 0.05 mg/kg buprenorphine along the animal&#39;s back for prophylaxis against postoperative pain.</li>
</ol>
4. Surgical approach and injury patterns
<ol>
	<li>Transfer the animal to the operating table and continue the gas flow via a nosecone. Ensure that a warming pad is positioned underneath the animal and the sterile field to maintain its body temperature.</li>
	<li>Place sterilized gauze (rolled up and fastened with tape) to use as a neck roll for the rat; this will provide an enhanced exposure of the surgical field. Note that the appropriate positioning of the animal is paramount for efficient nerve identification and dissection.</li>
	<li>Prepare the animal&#39;s facial skin for the procedure. Use chlorhexidine or an iodine-based solution to scrub the surgical site 3x, alternating with 70% ethanol, to ensure disinfection.</li>
	<li>Plan and mark the surgical incision if desired. Manipulate the ipsilateral ear in an anterior-posterior direction to determine the natural folding of the postauricular skin.</li>
	<li>Fashion a 4-5 mm incision in the postauricular crease using sharp iris scissors or a number 15 blade. This can be expanded later in the procedure as necessary.</li>
	<li>Bluntly dissect through the immediate subcutaneous fascia and place a micro-Weitlaner retractor to enhance exposure. Note that there may be small caliber blood vessels in this area; these are best avoided by retracting superiorly or inferiorly via the Weitlaner retractor.</li>
	<li>Identify the anterior digastric muscle as it travels in an inferior-to-superior direction toward its insertion along the skull base.
	<ol>
		<li>Spread gently through the muscle belly along its insertion point to reveal the tendon of the anterior digastric belly. Note that the tendon appears as a filmy white process emanating from the muscle with a solid insertion onto the skull base.</li>
	</ol>
	</li>
	<li>After identification of the anterior digastric muscle and its tendon, adjust the Weitlaner retractor to further retract the muscle belly. Note that the subsequently exposed region is the three-dimensional space where the main trunk of the facial nerve lies. 
	NOTE: This region is bounded superiorly and medially by the skull base, laterally by the anterior digastric muscle, posteromedially by the ear canal, and inferiorly by the structures of the neck, including the superficial temporal artery.</li>
	<li>After adequate exposure, identify the main trunk of the facial nerve as it travels inferiorly from underneath the tendon of the digastric muscle, where it exits the stylomastoid foramen from the skull base. Note that the nerve appears as a pearly white cord, encased in the animal&#39;s parotid-masseteric fascia. Practice caution when further exposing the nerve, for the following reasons.
	<ol>
		<li>Avoid aggressive dissection, or perpendicular spreads, to guard against stretch-mediated neuropraxia injury.</li>
		<li>Avoid aggressive posteriorly and medially directed dissection to guard against violating the thin tissues overlying the ear canal as this could introduce middle ear flora into the surgical field.</li>
		<li>Avoid damaging the superficial temporal artery through broad medially and inferiorly directed dissection. Note that an injury will be identified by brisk, pulsatile bleeding.
		<ol>
			<li>If the artery is injured, apply prompt pressure with a cotton-tipped applicator or sterile gauze via forceps. Hemostatic agents or liquid fibrin sealant can be placed in near proximity. Keep in mind that the animal may require a subcutaneous injection of 0.9% sterile saline for fluid stabilization.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Trace the main trunk distally by dissecting along the nerve in an inferior direction, distally from the exit of the stylomastoid foramen.
	<ol>
		<li>Extend the original incision to allow for a full exposure of the nerve and its branches. Take care to avoid a disruption of the parotid gland as this could result in postoperative sialocele.</li>
	</ol>
	</li>
	<li>Induce the desired injury patterns as follows.
	<ol>
		<li>For a crush injury, use smooth-surfaced jeweler&#39;s forceps to firmly grasp the nerve and compress it9. Apply constant and reproducible pressure to the nerve for a period of 30 s to ensure an appropriate crush injury.</li>
		<li>For a simple transection, grasp the fascia overlying the nerve, or the immediate epineurium, with fine-toothed forceps, and use sharp microscissors to cleanly transect the nerve at the desired point with a single cut. Take care to avoid excess traction on the nerve with the forceps.</li>
		<li>For a nerve gap model, create the desired nerve gap using a similar method to the simple transection injury. Use the sterilized shaft of a cotton-tipped applicator-cut to the desired nerve gap length-intraoperatively to ensure similarity of injury pattern between animals.</li>
	</ol>
	</li>
</ol>
5. Wound closure
<ol>
	<li>Irrigate the wound with sterile saline and dry it with sterile gauze.</li>
	<li>Approximate the skin edges in a simple, subcuticular fashion with absorbable sutures, or use skin glue or wound clips, which are also acceptable for wound closure. Place a buried stitch by taking a deep-to-superficial bite of one skin edge and then a subsequent superficial-to-deep bite of the opposite skin edge.</li>
</ol>
6. Postoperative recovery
<ol>
	<li>Administer a subcutaneous injection of nonsteroidal anti-inflammatory analgesic (such as&#160;0.05 mg/kg of buprenorphine and 0.5 mg /kg Carprofen) for postoperative pain control. Place the injection along the animal&#39;s back.</li>
	<li>Cease the administration of the anesthetic agent and allow the animal to inhale oxygen for an additional 1 min.</li>
	<li>Place the animal in a warmed (via a heat lamp), aseptic cage devoid of bedding material to avoid accidental ingestion. Note that the animal will typically demonstrate signs of recovery within 1-2 min and can appear disoriented, with a delayed recovery of hind-leg function.</li>
	<li>Return the animals to their cages in the appropriate housing unit and administer postoperative analgesics on postoperative day #1 to ensure continued prophylaxis against pain.</li>
	<li>Monitor the animals 2x per day to evaluate for signs of malnourishment, corneal irritation, or surgical site infection, and maintain appropriate surgical logs.
	<ol>
		<li>Administer 0.9% sterile saline in a subcutaneous fashion if there is significant weight loss.</li>
		<li>Apply lubricating eye ointment daily until the animal&#39;s blink reflex is re-established.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The rat facial nerve injury model has emerged as the most versatile system for the evaluation of neurotrophic factors due to its surgical accessibility, branching pattern, and physiological significance27,29,33,34,35,36. The combination of video demonstration and application of transgenic animal data opens new possibilities for the scientific study of nerve regenerative phenomena. This model allows the systematic and detailed characterization of neural response to trauma, neurotrophic influences, immunomodulatory influences, and other aspects of the microenvironment. Although the primary objective of clinical facial nerve study relates to the recovery of motoneuron function, the model can also be used to better understand events at the neuromuscular junction, nuances of axonal transport, and the interplay of axonal-glial influences27,36,37,38,39,40,41,42,43,44. Mechanistic study using facial nerve has paradoxically led to important insights relating to central inflammation and neurodegenerative diseases, such as Alzheimer&#39;s disease and Parkinson&#39;s disease45,46,47. There, the medical implications of work conducted in this model have notable implications for better understanding and eventually improving clinical care for patients afflicted with both peripheral and central nervous system disorders affecting both supporting cells and neuronal tissues.
There is an initial learning curve with performing facial nerve surgery in the rat model. While it is not necessary for the scientist to be surgically trained, they should be comfortable working underneath an operating microscope and utilizing binocular vision. The scientist should further be comfortable working with their nondominant hand, as adjusting the retraction with the Weitlaner retractor in the contralateral hand can significantly improve visualization. This is particularly true for identifying the main trunk of the facial nerve, as it is possible for the novice surgeon to lose landmarks and become disoriented in the three-dimensional space where the nerve exits the skull base. However, once experience is gained and the surgeon is consistently able to identify the tendon of the digastric muscle, then the procedures are quite straight-forward. Where the procedure-a simple transection of the nerve-could take as long as 30 min when the scientist is initially learning, in the hands of an experienced surgeon, it can be as expedient as 5 min from incision to wound closure. With an experienced assistant managing the preparation and the anesthesia of the animals-as well as restocking supplies as needed-it is feasible to operate on several animals in a single sitting. If complex intraoperative maneuvers are performed, such as a stereotaxic intraneural injection, then the time required will increase.
This group has experience working with the adult Lewis and Sprague-Dawley rats and the adult Thy1-GFP rat. These models have demonstrated impressive resilience, both intraoperatively and postoperatively. It would be prudent to allow time for the acclimation of the animal (typically 1 week) if ordered from a separate facility as mandated by the NIH Guide for Care and Use of Laboratory Animals, as operating too soon after a transfer could result in excessive stress and poor postoperative health. With unilateral facial nerve injury, the rat does not demonstrate untoward signs of malnourishment or corneal irritation postoperatively. Furthermore, they tolerate repetitive periods of anesthesia quite well48, as previous protocols have demanded serial electrophysiological analyses of the return of whisker function until eventual sacrifice. Although the operators should attempt to maintain intraoperative sterility to the best of their abilities, we have not noted postoperative infections with any of the operated animals. On occasion, the animal will attempt to scratch at their incision site; however, it typically returns to baseline status with regrowth of hair within 1-2 weeks postoperatively. If the parotid gland is injured or inadvertently removed, then sialocele will result, which may necessitate repeat anesthesia for drainage.
Modifications can be made to the location of injury, type of injury, or intervention performed. Injury can be induced at any point along the course of the facial nerve, from intracranial facial nerve transection to injury to the main trunk or any of its peripheral branches49. The broad patterns of injury include crush injury, simple transection with or without repair, and gap-defect with or without repair or bridge11. The range of possible interventions is exhaustive. Briefly, intervention can be undertaken at the level of the nerve proper17, at the site of the injury23, or at the muscular end-organ50. The list of possible outcome parameters is equally lengthy. Workhorse histomorphometric parameters include axial-based quantification of nerve count, nerve density, and percentage of nerve, among others. Additional measures include histologic analyses of longitudinal sections to illustrate the extent of regeneration and neuromuscular junction quantification to demonstrate the reinnervation of target facial musculature51. Novel methods of assessing outcomes continue to be developed22. For example, Hadlock et al. have demonstrated a complex method for assessing the contracture of independent facial zones via scent- or puff-induced reactions; this has potential for the assessment and eventual treatment of troublesome synkinetic regeneration11.
As with any animal model, there are limitations in translating the results to human patients. The mouse and rat models both exhibit the superior regenerative potential inherent in the rodent nervous system; this property allows the rodent to demonstrate regenerative results that could not possibly be achieved in humans and higher animals24. Therefore, the facial nerve scientist must carefully select appropriate time-points for the evaluation of nerve regeneration and recovery; if a prolonged time-point is selected, the narrow window of opportunity where experimental groups may reliable demonstrate the effect from their unique intervention may be missed24.

Cranial nerve injury in the head and neck region can be secondary to congenital, infectious, idiopathic, iatrogenic, traumatic, neurologic, oncologic, or systemic etiologies1. Cranial nerve VII, or the facial nerve, is commonly affected. The incidence of facial nerve dysfunction can be significant, as it affects 20 to 30 per 100,000 people each year2. The main motor branches of the facial nerve are the temporal, zygomatic, buccal, marginal mandibular, and cervical branches; depending on the branch involved, the consequences can include oral incompetence or drooling, corneal dryness, visual field obstruction secondary to ptosis, dysarthria, or facial asymmetry2,3. Long-term morbidity includes the phenomenon of synkinesis, or involuntary movement of one facial muscle group, with attempted voluntary contraction of a distinct facial muscle group. Ocular-oral synkinesis is the most common of the aberrant regeneration as a sequela of facial nerve injury and causes functional impairment, embarrassment, diminished self-esteem, and poor quality of life3. Injury to individual branches dictates the functions that are selectively compromised.
The clinical treatment of facial nerve injury is not well standardized and is in need of further research to improve outcomes. Steroids can alleviate acute facial nerve swelling, whereas Botox is useful for temporizing synkinetic movements; but, the primary reconstructive options in the practitioner&#39;s armamentarium involve surgical intervention through nerve repair, substitution, or reanimation3,4,5,6. Depending on the type of facial nerve injury sustained, the facial nerve surgeon may utilize a number of options. For simple transection, nerve reanastomosis is useful whereas cable-graft repair is better suited for a nerve defect; for a restoration of function, the surgeon may choose either static or dynamic facial reanimation procedures. In many cases of facial nerve injury and subsequent repair, even in the hands of experienced facial nerve surgeons, the best outcome still results in persistent facial asymmetry and functional compromise7.
These suboptimal outcomes have spurred extensive research on facial nerve regeneration. Broad topics of interest include perfecting and innovating nerve repair techniques, determining the effect of various nerve regeneration factors, and assessing the potential of specific neural inhibitors to help combat the long-term outcome of synkinesis8,9,10,11. While in vitro models can be used to assess some characteristics of pro-growth or inhibitory factors, true translational research on this subject matter is best accomplished via translatable animal models.
The decision of which animal model to utilize can be challenging, as researchers have utilized both large animals, such as sheep and small animal models, such as mice12,13. While large animal models offer ideal anatomic visualization, their use requires specialized equipment and personnel not readily or easily available. Furthermore, powering a study to demonstrate effect could be highly cost-prohibitive and potentially not within the feasible scope of many scientific centers. Thus, the small animal model is most frequently utilized. The mouse model can be utilized for assessing a number of outcomes related to facial nerve surgery; however, the limited length of the nerve can restrict the scientist&#39;s ability to model certain patterns, such as large-gap injury14.
Thus, the rat murine prototype has emerged as the workhorse model through which the scientist can perform innovative surgical procedures or utilize inhibitory or pro-growth factors and assess effect across a broad range of outcome parameters. The rat facial nerve anatomy is predictably and easily approached in a reproducible fashion. Its larger scale, in comparison to the mouse model, allows for modeling of a wide range of surgical defects, ranging from simple transection to 5 mm gaps15,16. This further allows for the application of complex interventions at the defect site, including the topical placement of factor, intraneural injections of factor, and the placement of isografts or bridges17,18,19,20,21,22,23.
The docile nature of the rat, its reliable anatomy, and its propensity for effective nerve regeneration allows for the collection of many outcome measures in response to the aforementioned surgical patterns of injury24. Via the rat model, the facial nerve scientist is able to assess electrophysiologic responses to injury, nerve and muscle histologic outcomes via immunohistochemistry, functional outcomes via tracking movement of the vibrissal pad and assessing eye closure, and micro- and macroscopic changes via fluorescent or confocal microscopy, among others11,22,23,25,26,27,28,29. Thus, the following protocol will outline a surgical approach to the rat facial nerve and the injury patterns that can be induced.

<table><tbody><tr><td>1.8% isoflurane</td><td>VetOne</td><td>13985-030-40</td><td></td></tr><tr><td>11-0 nylon microsutures</td><td>AROSuture</td><td>TK-117038</td><td></td></tr><tr><td>4-0 monocryl suture</td><td>VWR</td><td>75982-084</td><td></td></tr><tr><td>Buprenorphine SR</td><td>ZooPharm</td><td>MIF 900-006</td><td></td></tr><tr><td>Carprofen</td><td>Sigma-Aldrich</td><td>MFCD00079028</td><td></td></tr><tr><td>Chlorhexidine</td><td>VWR</td><td>IC19135805</td><td></td></tr><tr><td>Jeweler forceps</td><td>VWR</td><td>21909-458</td><td></td></tr><tr><td>Micro Weitlaner retractor</td><td>VWR</td><td>82030-146</td><td></td></tr><tr><td>Micro-scissors</td><td>VWR</td><td>100492-348</td><td></td></tr><tr><td>Mini tenotomy scissors</td><td>VWR</td><td>89023-522</td><td></td></tr><tr><td>Number 15 scalpel blade</td><td>VWR</td><td>102097-834</td><td></td></tr><tr><td>Operating microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Petrolatum eye gel</td><td>Pharmaderm</td><td>B002LUWBEK</td><td></td></tr><tr><td>Sterile water</td><td>VWR</td><td>89125-834</td><td></td></tr><tr><td>Tissue adhesive</td><td>Vetbond, 3M</td><td>NC9259532</td><td></td></tr><tr><td>Water conductor pad</td><td>Aqua Relief System</td><td>ARS2000B</td><td></td></tr><tr><td>Bupivacaine</td><td></td><td></td><td>Use as a local analgesic</td></tr></tbody></table>

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.

Following the initial surgical procedure, there are two main types of outcome measures: serial measurements in the live animal and measurements that require sacrificing the animal. Examples of serial measurements include electrophysiological assays, such as a compound muscle action potential measurement30, assessments of facial muscle movement via laser-assisted or videography means9, or even repetitive live imaging of regrowth of the facial nerve in fluorescent transgenic animals31,32. Figure 1 illustrates live imaging of the main trunk of the facial nerve in an adult transgenic Thy1-GFP rat. A crush injury has been performed on the marginal mandibular branch, approximately 2-3 mm distal to the branch point of the first pes. Utilizing MetaMorph imaging software, we were able to quantify the fluorescent intensity at any point along the course of the facial nerve. In particular, it is possible to quantify fluorescence proximal and distal to an injury site, thereby serially assessing for the return of fluorescence as a marker for nerve regeneration. Figure 2 illustrates the gradual return of fluorescence (measured as a ratio of distal fluorescence to proximal fluorescence across a simple transection site) at the 1, 2, 3, and 4 week time-points in the Thy1-GFP rat.
Histomorphometric analysis of the nerve proper or muscle requires sacrificing the animal after a predetermined length of time to demonstrate the desired effect across groups. Figure 3 demonstrates cross-sectional images of the marginal mandibular division. This technique requires careful tissue handling, storage, preparation, sectioning, and staining to allow for a histomorphometric analysis across various groups. If appropriately performed, this technique allows for the quantification of the axonal diameter, amount of debris, nerve fiber, percentage of nerve, and density measurements.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59224/59224fig1.jpg" /> 
Figure 1:&#160;Facial nerve anatomy demonstrated in the Thy1-GFP rat 1 week after a crush injury at the marginal mandibular nerve. The site of the crush injury is demonstrated by the white arrow. Sections of the nerve are labeled as MT (main trunk), B (buccal), and MM (marginal mandibular) branches. The scale bar represents 1.5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59224/59224fig1large.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/59224/59224fig2.jpg" /> 
Figure 2: Quantification of return of fluorescence as a ratio of the fluorescent intensity of the site immediately distal to the transection injury over the intensity of the site proximal to the transection injury. Four animals were studied, and a crush injury was modeled. The graph is plotted as mean proportion &#177; standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59224/59224fig2large.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/59224/59224fig3.jpg" /> 
Figure 3: Axial cross-section of the marginal mandibular branch of the rat facial nerve following staining with toluidine blue. The scale bar represents 100 &#181;m.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59224/59224fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration at JoVE.com

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

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 ...

facial-nerve-surgery-rat-model-to-study-axonal-inhibition

Nanyang Technological University

Research

JoVE Journal

Neuroscience

7.4K Views.  Michigan Medicine. This protocol describes a reproducible approach to facial nerve surgery in the rat model, including descriptions of various inducible patterns of injury. 

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

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 ...

Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and either cut or crush it to induce peripheral nerve injury. Advantages of this surgery are its simplicity, high reproducibility, and the lack of effect on vital functions or mobility from the subsequent facial paralysis, thus resulting in a relatively mild surgical outcome compared to other nerve injury models. A major advantage of using a cranial nerve injury model is that the motoneurons reside in a relatively homogenous population in the facial motor nucleus in the pons, simplifying the study of the motoneuron cell bodies. Because of the symmetrical nature of facial nerve innervation and the lack of crosstalk between the facial motor nuclei, the operation can be performed unilaterally with the unaxotomized side serving as a paired internal control. A variety of analyses can be performed postoperatively to assess the physiologic response, details of which are beyond the scope of this article. For example, recovery of muscle function can serve as a behavioral marker for reinnervation, or the motoneurons can be quantified to measure cell survival. Additionally, the motoneurons can be accurately captured using laser microdissection for molecular analysis. Because the facial nerve axotomy is minimally invasive and well tolerated, it can be utilized on a wide variety of genetically modified mice. Also, this surgery model can be used to analyze the effectiveness of peripheral nerve injury treatments. Facial nerve injury provides a means for investigating not only motoneurons, but also the responses of the central and peripheral glial microenvironment, immune system, and target musculature. The facial nerve injury model is a widely accepted peripheral nerve injury model that serves as a powerful tool for studying nerve injury and regeneration.

We present a surgical protocol detailing how to perform a cut or crush axotomy on the facial nerve in the mouse. The facial nerve axotomy can be employed to study the physiological response to nerve injury and test therapeutic techniques.

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and ...

Medicine

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 ...

Sheath-Preserving Optic Nerve Transection in Rats to Assess Axon Regeneration and Interventions Targeting the Retinal Ganglion Cell Axon

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as glaucoma, trauma, and ischemic optic neuropathies injure RGC axons, disrupt transmission of visual stimuli, and cause vision loss. Animal models simulating RGC axon injury include optic nerve crush and transection paradigms. Each of these models has inherent advantages and disadvantages. An optic nerve crush is generally less severe than a transection and can be used to assay axon regeneration across the lesion site. However, differences in crush force and duration can affect tissue responses, resulting in variable reproducibility and lesion completeness. With optic nerve transection, there is a severe and reproducible injury that completely lesions all axons. However, transecting the optic nerve dramatically alters the blood brain barrier by violating the optic nerve sheath, exposing the optic nerve to the peripheral environment. Moreover, regeneration beyond a transection site cannot be assessed without reapposing the cut nerve ends. Furthermore, distinct degenerative changes and cellular pathways are activated by either a crush or transection injury.
The method described here incorporates the advantages of both optic nerve crush and transection models while mitigating the disadvantages. Hydrostatic pressure delivered into the optic nerve by microinjection completely transects the optic nerve while maintaining the integrity of the optic nerve sheath. The transected optic nerve ends are reapposed to allow for axon regeneration assays. A potential limitation of this method is the inability to visualize the complete transection, a potential source of variability. However, visual confirmation that the visible portion of the optic nerve has been transected is indicative of a complete optic nerve transection with 90-95% success. This method could be applied to assess axon regeneration promoting strategies in a transection model or investigate interventions that target the axonal compartments.

This protocol describes an optic nerve transection that preserves the optic nerve sheath in rats. Hydrostatic pressure from microinjections into the optic nerve generate a complete transection, allowing for sutureless reapposition of the transected optic nerve ends and direct targeting of the axonal compartment in a transection model.

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as ...

Electroporation and Nerve Injury in Mouse Levator Auris Longus Muscle

Combined In Vivo Electroporation and Short&#45;Term Reinnervation of the Cranial Levator Auris Longus Skeletal Muscle

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many organisms. At the NMJ, a presynaptic motor axon terminal contacts a muscle postsynaptic domain and is covered by terminal Schwann cells. The integrity and function of the NMJ is compromised under several conditions, including aging, neuromuscular diseases, and traumatic injuries. To analyze the potential contribution of muscle-derived proteins to NMJ maintenance and regeneration, an in vivo gene transfer strategy has been combined with the denervation of the cranial levator auris longus (LAL) muscle after mechanical nerve injury. Previous findings showed that the forced expression of control fluorescent proteins does not alter NMJ organization or neurotransmission. This procedure aims to describe a detailed method of in vivo electroporation of the LAL muscle followed by transection or crushing of the specific branch of the facial nerve innervating cranial muscles, leading to NMJ denervation and reinnervation, respectively. The combination of these experimental strategies in the convenient LAL muscle constitutes an efficient method to study the potential contribution of muscle protein overexpression or silencing in the context of short-term NMJ reinnervation.

Author Spotlight: Regenerative Roles of Muscle Proteins at Neuromuscular Junction Post-Nerve Injury

Here, we present a protocol that combines in vivo electroporation and denervation of the cranial levator auris longus (LAL) muscle. This procedure enables the study of the potential role of muscle-derived proteins in the regeneration of the neuromuscular synapse.

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many ...

Establishment of Facial Nerve Injury Rat Model for Idiopathic Facial Paralysis Research

Idiopathic facial paralysis is the most common type of facial nerve injury, accounting for approximately 70% of peripheral facial paralysis cases. This disease can not only lead to a change in facial expression but also greatly impact the psychology of patients. In severe cases, it can affect the normal work and life of patients. Therefore, the research on facial nerve injury repair has important clinical significance. In order to study the mechanism of this disease, it is necessary to carry out relevant animal experiments, among which the most important task is to establish an animal model with the same pathogenesis as human disease. The compression of the facial nerve within the petrous bone, especially the nerve trunk at the junction of the distal end of the internal auditory canal and the labyrinthine segment, is the pathogenesis of idiopathic facial paralysis. In order to simulate this common disease, a compression injury model of the main extracranial segment of the facial nerve was established in this study. The neurological damage was evaluated by behavioral, neuroelectrophysiological, and histological examination. Finally, 50 g constant force and 90 s clamp injury were selected as the injury parameters to construct a stable idiopathic facial paralysis model.

The present protocol establishes a facial nerve injury rat model using microscopy to investigate the diagnostic and therapeutic mechanisms of idiopathic facial paralysis.

Idiopathic facial paralysis is the most common type of facial nerve injury, accounting for approximately 70% of peripheral facial paralysis cases. This ...

Biology

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.

Wywoływanie uszkodzenia nerwu twarzowego u modelu szczura

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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JoVE Encyclopedia of Experiments

Neuropathology

Trauma

Induction of Neuropathic Pain in a Rat Model by Constriction of the Infraorbital Nerve

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/53167/chronic-constriction-injury-rat-s-infraorbital-nerve-ion-cci-to-study'>Source: Deseure, K., et al.&#160;Chronic Constriction Injury of the Rat's Infraorbital Nerve (IoN-CCI) to Study Trigeminal Neuropathic Pain. J. Vis. Exp. (2015).</a>This video demonstrates a method to induce neuropathic pain in a rat model by constricting the infraorbital nerve. After exposing the infraorbital nerve and tying ligatures to constrict it, the immune response at the injury site increases the excitability of neighboring neurons, which transmit pain signals to the brain.

Source: Deseure, K., et al.&#160;Chronic Constriction Injury of the Rat's Infraorbital Nerve (IoN-CCI) to Study Trigeminal Neuropathic Pain. J. Vis. Exp. ...

Neuromuscular Disorders

Modeling Sensory Axon Regeneration in a Rat Using Dorsal Root Ganglion Injection and Dorsal Root Crush Injury

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/55535/dorsal-root-ganglion-injection-dorsal-root-crush-injury-as-model-for'>Source: Cheah, M., et al. Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration. J. Vis. Exp. (2017).</a>This video demonstrates a step-by-step procedure for injecting a viral suspension into the dorsal root ganglia (DRGs) and inducing a controlled axonal injury via a dorsal root crush in an anesthetized rat. The protocol involves exposing the DRGs, delivering a growth-promoting viral vector to facilitate axonal regeneration, and severing axonal connections to the spinal cord to model sensory nerve injury and regeneration.

Source: Cheah, M., et al. Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration. J. Vis. Exp. (2017).This ...

Performing Crush Axotomy on the Facial Nerve in a Mouse Model

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/52382/facial-nerve-axotomy-mice-model-to-study-motoneuron-response-to'>Source: Olmstead, D. N., et al.,&#160;Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury. J. Vis. Exp.&#160;(2015).</a>This video demonstrates the facial nerve crush procedure in a mouse, where blunt dissection through key landmarks exposes the facial nerve trunk at the stylomastoid foramen. Forceps are then used to compress the nerve trunk, inducing axonal damage and disrupting neuronal signal transmission.

Source: Olmstead, D. N., et al.,&#160;Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury. J. Vis. Exp.&#160;(2015).This video ...