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 f...

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.

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	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">1.8% isoflurane</td>
			<td style="width:175px;">VetOne</td>
			<td style="width:182px;">13985-030-40</td>
			<td style="width:171px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">11-0 nylon microsutures</td>
			<td style="width:175px;">AROSuture</td>
			<td style="width:182px;">TK-117038</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">4-0 monocryl suture</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">75982-084</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Buprenorphine SR</td>
			<td style="width:175px;">ZooPharm</td>
			<td>MIF 900-006</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Carprofen</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:182px;">MFCD00079028</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Chlorhexidine</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">IC19135805</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Jeweler forceps</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">21909-458</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Micro Weitlaner retractor</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">82030-146</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Micro-scissors</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">100492-348</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Mini tenotomy scissors</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">89023-522</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Number 15 scalpel blade</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">102097-834</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Operating microscope</td>
			<td style="width:175px;">Leica</td>
			<td style="width:182px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Petrolatum eye gel</td>
			<td style="width:175px;">Pharmaderm</td>
			<td style="width:182px;">B002LUWBEK</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Sterile water</td>
			<td style="width:175px;">VWR</td>
			<td style="width:182px;">89125-834</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Tissue adhesive</td>
			<td style="width:175px;">Vetbond, 3M</td>
			<td style="width:182px;">NC9259532</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Water conductor pad</td>
			<td style="width:175px;">Aqua Relief System</td>
			<td style="width:182px;">ARS2000B</td>
			<td></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...

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 ...

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7.3K 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

An Optic Nerve Crush Injury Murine Model to Study Retinal Ganglion Cell Survival

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible vision loss. Examples of such diseases in human include traumatic optic neuropathy and optic nerve degeneration in glaucoma. It is characterized by typical changes in the optic nerve head, progressive optic nerve degeneration, and loss of retinal ganglion cells, if uncontrolled, leading to vision loss and blindness.
The optic nerve crush (ONC) injury mouse model is an important experimental disease model for traumatic optic neuropathy, glaucoma, etc. In this model, the crush injury to the optic nerve leads to gradual retinal ganglion cells apoptosis. This disease model can be used to study the general processes and mechanisms of neuronal death and survival, which is essential for the development of therapeutic measures. In addition, pharmacological and molecular approaches can be used in this model to identify and test potential therapeutic reagents to treat different types of optic neuropathy.
Here, we provide a step by step demonstration of (I) Baseline retrograde labeling of retinal ganglion cells (RGCs) at day 1, (II) Optic nerve crush injury at day 4, (III) Harvest the retinae and analyze RGC survival at day 11, and (IV) Representative result.

This protocol shows how to retrogradely label retinal ganglion cells, and how to subsequently make an optic nerve crush injury in order to analyze retinal ganglion cell survival and apoptosis. It is an experimental disease model for different types of optic neuropathy, including glaucoma.

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible ...

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

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

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

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

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

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

Visual Evoked Potential Recording in a Rat Model of Experimental Optic Nerve Demyelination

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to monitor the disease course in the follow-up period. Changes in the VEP parameters closely correlate with pathological damage in the optic nerve. This protocol provides a detailed description about the rodent model of optic nerve microinjection, in which a partial demyelination lesion is produced in the optic nerve. VEP recording techniques are also discussed. Using skull implanted electrodes, we are able to acquire reproducible intra-session and between-session VEP traces. VEPs can be recorded on individual animals over a period of time to assess the functional changes in the optic nerve longitudinally. The optic nerve demyelination model, in conjunction with the VEP recording protocol, provides a tool to investigate the disease processes associated with demyelination and remyelination, and can potentially be employed to evaluate the effects of new remyelinating drugs or neuroprotective therapies.

Focal demyelination is induced in the optic nerve using lysolecithin microinjection. Visual evoked potentials are recorded via skull electrodes implanted over the visual cortex to examine the signal conduction along the visual pathway in vivo. This protocol details the surgical procedures underlying electrode implantation and optic nerve microinjection.

The visual evoked potential (VEP) recording is widely used in clinical practice to assess the severity of optic neuritis in its acute phase, and to ...

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

Screening of Axonal Degeneration in Carpal Tunnel Syndrome Using Ultrasonography and Nerve Conduction Studies

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current diagnostic and severity gradation system cannot clearly indicate its coexistence, resulting in&#160;confusion of appropriate treatment prescription. There are also constraints in conventional methods for&#160;differentiation as well. This study aims at introducing an innovative, efficient, and quick screening protocol to differentiate axonal degeneration associated with CTS, using ultrasound and nerve conduction studies (NCS). It starts by using NCS to perform orthodromic stimulation at the wrist, to obtain the sensory conduction of the median and the ulnar nerves respectively. Meanwhile, the motor conduction&#160;of the median nerve is collected by stimulating the palm, wrist, and elbow, followed by the stimulation of the ulnar nerve at the wrist,&#160;below and above the elbow. Then, an ultrasound assessment is performed, using a linear array transducer, with cross-sectional area (CSA) and perimeter (P) at the wrist and at the one-third distal forearm calipered. Ratios (R-CSA, R-P) and changes from wrist to one-third distal forearm (&#916;CSA and &#916;P) are calculated according to a standard format. Potential axonal degeneration coexisting in CTS will be screened according to the criteria of NCS and cut-off values of ultrasound measurements established in a previous study. In terms of its noninvasiveness, low cost, convenience, and efficiency, it is easy to apply ultrasound complimentarily in clinical practice to prescreen patients with potential coexisting axonal degeneration. Nevertheless, the ultrasonographic imaging cannot directly reflect axonal degeneration. It still relies on conventional but invasive methods&#160;such as electromyography (EMG) and biopsy&#160;for confirmation if needed.

Here we present a protocol using nerve conduction studies and ultrasound to screen potential axonal degeneration associated with carpal tunnel syndrome. The criteria for differentiation are established. Compared to conventional approaches, this method is noninvasive, convenient, and efficient, with an overall satisfactory accuracy, sensitivity, and specificity.

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current ...

Thoracic Spinal Cord Hemisection Surgery and Open-Field Locomotor Assessment in the Rat

Spinal cord injury (SCI) causes disturbances in motor, sensory, and autonomic function below the level of the lesion. Experimental animal models are valuable tools to understand the neural mechanisms involved in locomotor recovery after SCI and to design therapies for clinical populations. There are several experimental SCI models including contusion, compression, and transection injuries that are used in a wide variety of species. A hemisection involves the unilateral transection of the spinal cord and disrupts all ascending and descending tracts on one side only. Spinal hemisection produces a highly selective and reproducible injury in comparison to contusion or compression techniques that is useful for investigating neural plasticity in spared and damaged pathways associated with functional recovery. We present a detailed step-by-step protocol for performing a thoracic hemisection at the T8 vertebral level in the rat that results in an initial paralysis of the hindlimb on the side of the lesion with graded spontaneous recovery of locomotor function over several weeks. We also provide a locomotor scoring protocol to assess functional recovery in the open-field. The locomotor assessment provides a linear recovery profile and can be performed both early and repeatedly after injury in order to accurately screen animals for appropriate time points in which to conduct more specialized behavioral testing. The hemisection technique presented can be readily adapted to other transection models and species, and the locomotor assessment can be used in a variety of SCI and other injury models to score locomotor function.

The rat thoracic spinal hemisection is a valuable and reproducible model of unilateral spinal cord injury to investigate the neural mechanisms of locomotor recovery and treatment efficacy. This article includes a detailed step-by-step guide to perform the hemisection procedure and to assess locomotor performance in an open-field arena.

Spinal cord injury (SCI) causes disturbances in motor, sensory, and autonomic function below the level of the lesion. Experimental animal models are ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Automated Gait Analysis to Assess Functional Recovery in Rodents with Peripheral Nerve or Spinal Cord Contusion Injury

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and a lot of comparative data has been published in the literature. This includes a multitude of assessment methods to study functional recovery following nerve injury and repair. Besides evaluation of nerve regeneration by means of histology, electrophysiology, and other in vivo and in vitro assessment techniques, functional recovery is the most important criterion to determine the degree of neural regeneration. Automated gait analysis allows recording of a vast quantity of gait-related parameters such as Paw Print Area and Paw Swing Speed as well as measures of inter-limb coordination. Additionally, the method provides digital data of the rats&#8217; paws after neuronal damage and during nerve regeneration, adding to our understanding of how peripheral and central nervous injuries affect their locomotor behavior. Besides the predominantly used sciatic nerve injury model, other models of peripheral nerve injury such as the femoral nerve can be studied by means of this method. In addition to injuries of the peripheral nervous systems, lesions of the central nervous system, e.g., spinal cord contusion can be evaluated. Valid and reproducible data assessment is strongly dependent on meticulous adjustment of the hard- and software settings prior to data acquisition. Additionally, proper training of the experimental animals is of crucial importance. This work aims to illustrate the use of computerized automated gait analysis to assess functional recovery in different animal models of peripheral nerve injury as well as spinal cord contusion injury. It also emphasizes the method&#8217;s limitations, e.g., evaluation of nerve regeneration in rats with sciatic nerve neurotmesis due to limited functional recovery. Therefore, this protocol is thought to help researchers interested in peripheral and central nervous injuries to assess functional recovery in rodent models.

Automated gait analysis is a feasible tool to evaluate functional recovery in rodent models of peripheral nerve injury and spinal cord contusion injury. While it requires only one setup to assess locomotor function in various experimental models, meticulous hard- and soft-ware adjustment and training of the animals is highly important.

Peripheral and central nerve injuries are mostly studied in rodents, especially rats, given the fact that these animal models are both cost-effective and ...