Name: facial nerve axotomy in mice: a model to study motoneuron response to injury
Uploaded: 2015-02-23T15:00:00.000+00:00
Duration: 10 min 11 s
Description: Watch this Scientific Journal Video about Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury at JoVE.com

This work is funded by NIH RO1 NS 40433 (K.J.J.).

All procedures executed are approved by the Indiana University School of Medicine Institutional Animal Care and Use Committee and follow National Institute of Health guidelines.
1. Surgical Technique
<ol>
	<li>Maintain aseptic technique during this procedure by using sterile gloves, instruments, and a sterile surgical field according to NIH guidelines25. Sterilize tools before beginning surgery by autoclaving them (see Table of Specific Reagents/Equipment for complete list). Use a glass bead sterilizer to sterilize tools during the operation.</li>
</ol>
2. Anesthesia and Preparation
<ol>
	<li>Anesthetize the mouse in an anesthesia box with a mixture of 0.9 L/min oxygen and 2.5% isoflurane using a veterinary isoflurane vaporizer system. Ensure that the mouse does not respond to changes in body position before removing it from the box.</li>
	<li>Apply ophthalmic ointment to the mouse&#8217;s eyes to protect them from drying out.</li>
	<li>Switch the gas flow from the box to the nose cone. Place the mouse squarely on its left side on a heated pad covered with a surgical pad and absorbent bench paper with its nose and mouth inside the cone. Continuously monitor the mouse&#8217;s breathing rhythm and rate and adjust isoflurane levels as needed (between 2.5 - 3% isoflurane) to maintain an adequate level of anesthesia, and use the toe pinch reflex to confirm total sedation.</li>
</ol>
3. Surgical Approach
<ol>
	<li>Align and focus the stereoscope with the surgical field. Adjust the nose cone and tape it down so that it is positioned along the edge of the visual field.</li>
	<li>With the mouse lying on its left side, tape the edge of the right ear to the nose cone, exposing the area behind the ear where the incision will be made. Ensure that the posterior auricular vein travels horizontally across the ear. Note that the correct placement of the animal and taping of the ear are crucial in order to quickly find the facial nerve.</li>
	<li>Wet the fur on and behind the ear with 70% ethanol and shave the surgical site using a razor or scalpel blade. Pre-wetting the fur&#160;makes shaving easier in this anatomical location.</li>
	<li>Clean the skin with an iodine solution, such as Betadine surgical scrub (7.5% povidone-iodine), followed by 70% ethanol. Repeat this cleaning two more times to thoroughly disinfect the area.</li>
	<li>To determine where to make the incision, trace the posterior auricular vein from the ear caudally to the area posterior to the ear protuberance. Using spring scissors, make a 4 mm incision 2 - 3 mm posterior to the protuberance.</li>
	<li>Dissect through the subcutaneous fat and fascia using blunt dissection. Avoid direct cutting with the scissors because blood vessels or muscle tissue could be easily damaged.</li>
	<li>If bleeding occurs, apply pressure to the surgical site with a sterile cotton swab for at least 30 sec. If significant fluid loss occurs, inject the mouse intraperitoneally with up to 0.5 ml of sterile 0.9% saline solution using a 25 or 27 G needle.</li>
	<li>Use several key landmarks, the spinal accessory nerve, ear canal, and anterior digastric muscle (described below), to locate the facial nerve. Dissect around these landmarks until the branches of the facial nerve are visualized. The nerve will appear as a significant solid white structure when it is revealed and a layer of fascia adheres it to the underlying structures.
	<ol>
		<li>Find the spinal accessory nerve, which travels from the caudal portion of the skull to innervate the trapezius muscle, once the subcutaneous fat and fascia have been dissected. The facial nerve is deep to the spinal accessory nerve.</li>
		<li>Find the cartilaginous ear canal that looks pearly white and can be seen rostral to the facial nerve.</li>
		<li>Find the muscle belly of the anterior digastric muscle that lies on top of and caudal to the facial nerve.</li>
	</ol>
	</li>
	<li>When the main branches of the facial nerve are visualized, trace them dorsally to find their origin from the stylomastoid foramen. Using fine tipped Dumont forceps #5/45 to hold the surgical site open, advance the spring scissor tips following the nerve&#8217;s path, then move the forceps dorsally to keep the newly advanced area open.</li>
	<li>Visualize the trunk of the facial nerve with the zygomatic, buccal, and marginal mandibular branches at this point. 
	NOTE: The temporal branch will be found closer to the foramen. The marginal mandibular nerve branches into its upper and lower parts closer to the jaw, thus those nerve branches&#160;will not be visible at this level.
	<ol>
		<li>If performing a nerve transection, stabilize the nerve gently with the fine tip forceps and cut the nerve with the spring scissors. Avoid applying too much traction to the nerve with the forceps to prevent avulsing the nerve from the brainstem. Push the stumps away from each other, or cut and remove a portion of the distal nerve to ensure that no reconnection can occur.</li>
		<li>If performing a crush injury, use Dumont #5/45 forceps to compress the nerve for 30 sec using constant pressure to sever all axons, then repeat this crush at a second angle perpendicular to the first crush site. Avoid applying variable amounts of pressure during the 30 sec crush, otherwise the injury will be inconsistent between animals.</li>
	</ol>
	</li>
</ol>
4. Closing and Recovery
<ol>
	<li>Reposition the fat and muscles over the underlying structures.</li>
	<li>Approximate the edges of the incision and close the wound using a 7.5 mm wound clip.&#160;Sutures or glue are also acceptable for wound closure.&#160; Postsurgical analgesics can be provided at this time.&#160; &#160;</li>
	<li>Remove the tape from the mouse&#8217;s ear. Turn off the isoflurane flow and allow the mouse to breathe pure oxygen for 30 sec to 1 min. Place the mouse in an empty cage with no bedding to recover from anesthesia.</li>
	<li>When the mouse is recovered, examine its behavior for confirmatory signs of facial paralysis. The whiskers will be paralyzed and angled back towards the cheek, the nose will be deviated, and the eye will not blink in response to a puff of air.</li>
	<li>House animals jointly after surgery if they are female. Avoid housing&#160;male mice jointly because they&#160;are more aggressive and tend to forcibly remove their cagemate&#8217;s wound clips, which leads to infection. Provide postsurgical analgesics at this time, if necessary.</li>
	<li>Monitor the mice once a day for several days after the operation to ensure that no infection or other complication occurs postoperatively. Remove wound clips 7 - 10 days after the surgery if they have not fallen out on their own.</li>
	<li>Apply lubricating eye ointment to the affected eye daily to prevent corneal complications, either until the eye blink reflex is recovered or until euthanasia.</li>
</ol>

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The authors have nothing to disclose.

The critical step for this protocol is positioning the mouse properly before surgery is begun. If the mouse is not lying flat on its side, the ear is not taped at the correct angle, or the incision is made in an incorrect location, then finding the facial nerve becomes much more difficult. When this technique is mastered, surgeries will take only minutes per mouse. 
Either sutures, glue, or wound clips can be used to close the wound. Wound clips are preferred because of the small size of incision and rapidity with which they can be applied. We have determined that if wound clips are used and the mice are housed together, female mice fare better than male mice because males housed together often remove their cagemate&#8217;s wound clips, leading to infection of the surgical site.
Modifications can be made to the injury, such as changing the location along the nerve to induce injury. For example, the injury could be administered along one of the branches of the facial nerve28, or the skull could be opened to deliver an intracranial injury29. In addition, a variety of treatments can be implemented, such as applying drugs to the nerve stump14, injecting medicines systemically30, or even performing cell transfers31. Motoneuron diseases such as amyotrophic lateral sclerosis can also be studied using the facial nerve axotomy model32. Ongoing applications of this technique involve studying nerve reconnection strategies after transection and the application of pharmaceutical agents to accelerate nerve regeneration after injury. 
One of the most useful aspects of this surgery is that it is easy to confirm successful axotomy by monitoring the mouse&#8217;s vibrissae orientation and movements as well as the eye blink reflex. 
The paralysis does not affect the animal&#8217;s vital functions and overall is well tolerated in the mouse. 
A major advantage of the facial nerve injury model is that the isolated study of motoneurons is possible. However, because this is a cranial nerve injury, results found using this method may not translate to spinal, autonomic, or sensory nerve injury models. To study these other nerve injury types, a sciatic33 or tibial34 nerve injury model is recommended. The drawback of these models is that there is crosstalk from the contralateral neurons in the spinal cord that can confound results. There is no crosstalk between the facial motor nuclei, so the contralateral facial motor nucleus can serve as a paired internal control. The facial nerve axotomy model presents many advantages for studying nerve injury and regeneration.

Many peripheral nerve injury models exist, but one that stands out for the study of motoneurons is the facial nerve axotomy model. The facial nerve, also known as cranial nerve VII, originates in the pons and innervates the muscles of facial expression1,2. In this surgical protocol, the facial nerve is exposed at its exit from the stylomastoid foramen and either cut or crushed. The severity of nerve injury can be classified following the Sunderland3 classifications, which differentiates the injury based on the intactness of the axons, endoneurium, perineurium, and epineurium, which are connective tissue layers that sequentially wrap around the axon bundles. In the crush injury (axonotmesis), the axons are severed, but the perineurium and epineurium are preserved. Complete functional recovery from facial nerve crush occurs in about 11 days because the intact nerve sheath serves as a conduit within which the axons regrow4,5. On the other hand, in the cut injury (neurotmesis), the axons and all 3 connective tissue layers are severed, and the entire distal nerve must regrow to restore musculature innervation. Surgical reconnection of the epineurium is often performed in human patients with nerve transection injuries, however the recovery outcomes are seldom optimal. Further study is required to understand why the nerve fails to regrow to its target and what therapies can be employed to improve and accelerate the regenerative process.
There are many advantages to studying nerve injury using the facial nerve axotomy model. First, the facial nerve axotomy procedure is quick, easy, and highly reproducible; and the resultant paralysis of the facial muscles does not impact vital functions and is well tolerated by the animal. Because this is a cranial nerve injury model, studying the motoneuron cell bodies is simplified because the motoneurons reside in a relatively homogenous population in the facial motor nucleus in the pons. The population does differ based on the subnuclear pattern within the facial motor nucleus, as there are seven subnuclei each specific to innervating a specific group of muscles, so subnuclear differences in response to axotomy may impact results2,6,7.
A major benefit of the facial nerve injury model is that the unaxotomized side can serve as a paired internal control because the nerve innervation is highly symmetrical and there is no crosstalk between the facial motor nuclei8. Another advantage of using this surgical method is the lack of direct trauma to the CNS or disruption of the blood brain barrier9. Complications such as excessive bleeding and infection are rare with this procedure.
A variety of analyses can be performed to assess the physiologic response to nerve injury. The recovery of the eye blink reflex and whisker activity can be used as a behavioral measure of functional recovery10,11. Video recording of vibrissae activity is currently the most powerful method for detecting recovery of facial nerve innervation12,13. After euthanasia, histological analysis of the brainstem can be performed on the motoneuron cell bodies within the facial motor nucleus. The facial motor nucleus is subdivided into seven subnuclei, each specific to certain facial muscles, allowing for differential examination of responses to injury2,6. Facial motoneurons can be counted to quantify cell survival, or immunohistochemistry can be used to identify biomarkers and specific cell populations14. The facial motor nucleus can be accurately microdissected using laser capture for molecular analysis of the cellular response to nerve injury15,16. Impacts of the facial nerve axotomy can be analyzed in the motor cortex17,18. Also, the nerve can be dissected to study Wallerian degeneration19 or axon regeneration20, and the muscles can be removed to study neuromuscular junctions21. The facial nerve axotomy can also be used to study the accompanying central and peripheral glial cells22, target musculature21, and the immune system response23. Although much has been accomplished in studying the facial nerve axotomy model24, further study of peripheral nerve injury is required because nerve damage is a significant problem for patients and current treatments fail to produce optimal results. This model is a powerful tool for examining the physiologic response to nerve injury and analyzing the effectiveness of nerve regeneration therapies.

<table><tbody><tr><td>Stereo microscope</td><td>Leica</td><td>M60</td><td></td></tr><tr><td>Labeling tape</td><td>Fisher Scientific</td><td>15-952</td><td></td></tr><tr><td>Vannas-T&amp;uuml;bingen spring scissors - straight/sharp/8.5 cm/5 mm cutting edge</td><td>Fine Science Tools</td><td>15003-08</td><td>Sterilize before use</td></tr><tr><td>Dumont #5/45 forceps - standard tips/angled 45&amp;deg;/Dumoxel/11 cm</td><td>Fine Science Tools</td><td>11251-35</td><td>Sterilize before use</td></tr><tr><td>Michel suture clips - 7.5 mm x 1.75 mm</td><td>Fine Science Tools</td><td>12040-01</td><td>Described as &quot;wound clip&quot; in protocol, sterilize before use</td></tr><tr><td>Hagenbarth cross action wound clip applier 5&quot;</td><td>George Tiemann &amp;amp; Co</td><td>160-910</td><td>Used to apply wound clip, sterilize before use</td></tr><tr><td>Michel suture clip applicator &amp;amp; remover - For 7.5 mm clips</td><td>Fine Science Tools</td><td>12029-12</td><td>Used to remove wound clip</td></tr><tr><td>0.9% Sodium chloride injection, USP</td><td>Hospira</td><td>0409-4888-10</td><td></td></tr><tr><td>Betadine, 16 oz, with dispenser</td><td>Fisher Scientific</td><td>19-027132</td><td></td></tr><tr><td>70% Ethanol</td><td></td><td></td><td></td></tr><tr><td>Glass bead sterilizer</td><td></td><td></td><td></td></tr></tbody></table>

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.

After the facial nerve axotomy is performed, motoneuron loss occurs as a result of the injury. Motoneuron survival after injury depends on many variables, such as gender, animal age at time of surgery, and the timepoint at which the motoneuron counts are done, and the Moran and Graeber review24 and Jinno and Yamada review22 both summarize motoneuron survival data. Typically, about 86% of motoneurons survive at 28 days post-axotomy14,15,26. Kinetics of motoneuron loss are described in Serpe et al. 2000. Figure 1 illustrates the variation in motoneuron survival in multiple genetically modified mice. No significant differences are observed in facial motoneuron counts of the control side, indicating that the genetic alterations do not impact baseline counts. Compared to the motoneuron survival in wild type mice (84% &#177; 2.0; Figure 1A,D), significant cell loss is observed in a mouse model of amyotrophic lateral sclerosis (SOD1G93A; 68% &#177; 1; Figure 1B,E) as well as the immunodeficient recombination-activating gene-2 knockout mouse (RAG-2-/-; 57% &#177; 2.5; Figure 1C,F)27.
Figure 2 demonstrates the laser capture microdissection technique applied to the facial motor nucleus. The entire facial motor nucleus can be captured (Figure 2A-C), or subnuclei can be collected separately (Figure 2D-F). For greater precision, motoneurons can be captured individually, and the remaining neuropil can be collected for analysis (Figure 2G-I). Figure 3 depicts qPCR results of the RNA material extracted from the subnuclear samples comparing the ventromedial and ventrolateral subnuclei. The four genes tested, &#946;II tubulin, growth associated protein-43 (Gap-43), hemopoietic- and neurologic-expressed sequence-1 (Hn1), and brain-derived neurotrophic factor (BDNF) are all associated with the nerve regeneration response and there are interesting differences between the two subnuclei and their gene expression profiles after axotomy16.
<img alt="Figure 1" src="/files/ftp_upload/52382/52382fig1highres.jpg" /> 
Figure 1. Representative coronal facial motor nucleus sections stained with thionin and quantified 28 days after facial nerve transection. Facial motor nuclei are shown from (A,D) WT, (B,E) SOD1G93A, and (C,F) RAG-2-/- mice (control side, axotomized side). Scale bars = 120 &#956;m. This figure has been modified from 27. <a href="https://www.jove.com/files/ftp_upload/52382/52382fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52382/52382fig2highres.jpg" /> 
Figure 2. Laser microdissection of the facial motor nucleus. (A) Thionin-stained section of axotomized facial nucleus, (B) with partial laser microdissection of the axotomized facial nucleus, and (C) collection of laser microdissected tissue. A template (D) of the subnuclei was superimposed on the computer screen to identify the ventromedial and ventrolateral facial subnuclei for laser microdissection (E,F). Facial motoneurons were laser microdissected based on their morphology with a visible nucleus and nucleolus (* indicates motoneuron, G,H), while FMN cell body fragments, indicated by the arrows (G), were laser microdissected separately and disposed of to eliminate FMN mRNA in the neuropil samples. After all FMN and cell body fragments were collected, the remaining facial nucleus tissue was laser microdissected as the neuropil sample (I). Scale bars = 100 &#956;m. This figure has been modified from 16. <a href="https://www.jove.com/files/ftp_upload/52382/52382fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52382/52382fig3highres.jpg" /> 
Figure 3. Pro-regeneration and pro-survival mRNA expression in the ventromedial (VM) and ventrolateral (VL) facial motor subnuclear regions following facial nerve axotomy. Average percent of mRNA expression &#177; SEM in the transected VM and VL facial subnuclei relative to the unoperated control subnuclei (A-D). Time-course of mRNA expression includes no injury (0), 3, 7, 14, and 28 dpo for &#946;II tubulin (A), GAP-43 (B), Hn1 (C), and BDNF (D). # represents significant differences of VL compared to VM, at p &#60; 0.05. This figure has been modified from 16.

Watch this Scientific Journal Video about Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury at JoVE.com

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

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 ...

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Universidad San Sebastian

Research

JoVE Journal

Medicine

12.8K Views.  Indiana University School of Medicine. 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. 

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

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 ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

Neuroscience

The Fibular Nerve Injury Method: A Reliable Assay to Identify and Test Factors That Repair Neuromuscular Junctions

The neuromuscular junction (NMJ) undergoes deleterious structural and functional changes as a result of aging, injury and disease. Thus, it is imperative to understand the cellular and molecular changes involved in maintaining and repairing NMJs. For this purpose, we have developed a method to reliably and consistently examine regenerating NMJs in mice. This nerve injury method involves crushing the common fibular nerve as it passes over the lateral head of the gastrocnemius muscle tendon near the knee. Using 70 day old female mice, we demonstrate that motor axons begin to reinnervate previous postsynaptic targets within 7 days post-crush. They completely reoccupy their previous synaptic areas by 12 days. To determine the reliability of this injury method, we compared reinnervation rates between individual 70 day old female mice. We found that the number of reinnervated postsynaptic sites was similar between mice at 7, 9, and 12 days post-crush. To determine if this injury assay can also be used to compare molecular changes in muscles, we examined levels of the gamma-subunit of the muscle nicotinic receptor (gamma-AChR) and the muscle-specific kinase (MuSK). The gamma-AChR subunit and MuSK to are highly upregulated following denervation and return to normal levels following reinnervation of NMJs. We found a close relationship between transcript levels for these genes and innervation status of muscles. We believe that this method will accelerate our understanding of the cellular and molecular changes involved in repairing the NMJ and other synapses.

We have developed a nerve injury method to reliably examine muscle reinnervation, and thus regeneration of neuromuscular junctions in mice. This technique involves injuring the common fibular nerve via a simple and highly reproducible surgery. Muscle reinnervation in then assessed by whole-mounting the extensor digitorum longus muscle.

The neuromuscular junction (NMJ) undergoes deleterious structural and functional changes as a result of aging, injury and disease. Thus, it is imperative ...

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

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

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

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

A Mouse Model of Direct Anastomosis via the Prespinal Route for Crossing Nerve Transfer Surgery

Crossing nerve transfer surgery has been a powerful approach for repairing injured upper extremities in patients with brachial plexus avulsion injuries. Recently, this surgery was creatively applied in the clinical treatment of brain injury and achieved substantial rehabilitation of the paralyzed arm. This functional recovery after the surgery suggests that peripheral sensorimotor intervention induces profound neuroplasticity to compensate for the loss of function after brain damage; however, the underlying neural mechanism is poorly understood. Therefore, an emergent clinical animal model is required. Here, we simulated clinical surgery to establish a protocol of direct anastomosis of bilateral brachial plexus nerves via the prespinal route in mice. Neuroanatomical, electrophysiological, and behavioral experiments helped identify that the transferred nerves of these mice successfully reinnervated the impaired forelimb and contributed to accelerating motor recovery after brain injury. Therefore, the mouse model revealed the neural mechanisms underlying rehabilitation upon crossing nerve transfer after central and peripheral nervous system injuries.

We simulated clinical surgery to establish a protocol of direct anastomosis of bilateral brachial plexus nerves via the prespinal route in mice, contributing to the study of the neural mechanisms underlying rehabilitation upon crossing nerve transfer after central and peripheral nervous system injuries.

Crossing nerve transfer surgery has been a powerful approach for repairing injured upper extremities in patients with brachial plexus avulsion injuries. ...

Traumatic Peripheral Nerve Injury in Mice

Traumatic peripheral nerve injury (TPNI) is a common cause of morbidity following orthopedic trauma. Reproducible and precise methods of injuring nerve and denervating muscle have long been a goal in musculoskeletal research. Many traumatically injured limbs have nerve trauma that defines the long-term patient outcome. Over several years, precise methods of producing microsurgical nerve injuries have been developed, including crush, lacerations, and nerve-gap grafting, allowing for reproducible outcome assessments. Moreover, newer methods are created for calibrated crush injuries that offer clinically relevant correlations with outcomes used to assess human patients. The principles of minimal manipulation to ensure low variability in nerve injury allow for adding still more associated tissue injuries into these models. This includes direct muscle crush and other components of limb injury. Finally, atrophy assessment and precise analysis of behavioral outcomes make these methods a complete package for studying musculoskeletal trauma that realistically incorporates all the elements of human traumatic limb injury.

The present protocol describes traumatic peripheral nerve injuries (TPNIs), including precisely calibrated crush, strictly aligned and misaligned laceration, as well as grafted and non-grafted gaps of the sciatic nerve in mice. Custom-designed sensors are developed to gauge nerve trauma, induced with commonly available tools to ensure reproducible post-TPNI outcomes.

Traumatic peripheral nerve injury (TPNI) is a common cause of morbidity following orthopedic trauma. Reproducible and precise methods of injuring nerve ...

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 ...

Establishing an Orofacial Muscle Fibrosis Model in Mice via a Freezing Injury

Freezing Injury in Mouse Masseter Muscle to Establish an Orofacial Muscle Fibrosis Model

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived limb muscles, the orofacial muscles originate from the branchial arches, with exclusive contributions from the cranial neural crest. A recent study has revealed that regeneration is also different in the orofacial muscle group. However, the underlying regulatory mechanism remains to be uncovered. Current skeletal muscle regeneration models mainly focus on the limb and trunk muscle. In this protocol, dry ice was used to induce freezing injury in the mouse masseter muscle and tibialis anterior muscle to create an orofacial muscle fibrosis model. The temporal dynamics of muscle satellite cells and fibro-adipogenic progenitors were different between the two muscles, leading to impaired myofiber regeneration and excessive extracellular matrix deposition. With the help of this model, a deeper investigation into muscle regeneration in the orofacial area could be carried out to develop therapeutic approaches for patients with orofacial diseases.

Author Spotlight: Exploring Orofacial Muscle Regeneration &#8211; Insights and Innovations

The goal of this protocol is to establish an orofacial muscle fibrosis model. Comparison of the histology between mice masseter and tibialis anterior muscle after freezing injury confirmed masseter muscle fibrosis. This model will facilitate further investigation into the mechanism underlying orofacial muscle fibrosis.

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived ...

Biology

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 ...

Inducing a Facial Nerve Injury in a Rat Model

Source: Ali, S. A., et al. <a href="https://app.jove.com/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.
1. ...