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

JoVE Journal
Neuroscience

Your institution must subscribe to JoVE's Neuroscience section to access this content.

Fill out the form below to receive a free trial or learn more about access:

 

Summary

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

Cite this Article

Copy Citation | Download Citations | Reprints and Permissions

Ali, S. A., Stebbins, A. W., Hanks, J. E., Kupfer, R. A., Hogikyan, N. D., Feldman, E. L., Brenner, M. J. Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration. J. Vis. Exp. (159), e59224, doi:10.3791/59224 (2020).

Abstract

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.

Introduction

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'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'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.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

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's Institutional Animal Care & Use Committee (IACUC) prior to implementation. Ten-week-old adult female Sprague-Dawley rats were utilized.

1. Prior to the operative day

  1. 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.

2. Preoperative setup

  1. 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.
  2. 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.

3. Anesthesia and preparation

  1. Place the animal in the anesthesia chamber and induce general anesthesia via 1.8% isoflurane and 0.9 L/min oxygen.
    1. Confirm an adequate plane of anesthesia via an assessment of spontaneous breathing and an evaluation of consciousness by assessing the animal's grimace response to a toe pinch.
  2. Apply eye lubricant bilaterally to guard against corneal irritation or dryness.
  3. Shave the operative site(s) with a razor or automatic clipper.
    1. Establish a method for rat identification at this time, either via an ear tag or tail label/marking.
  4. Administer a subcutaneous injection of 0.05 mg/kg buprenorphine along the animal's back for prophylaxis against postoperative pain.

4. Surgical approach and injury patterns

  1. 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.
  2. 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.
  3. Prepare the animal'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.
  4. 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.
  5. 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.
  6. 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.
  7. Identify the anterior digastric muscle as it travels in an inferior-to-superior direction toward its insertion along the skull base.
    1. 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.
  8. 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.
  9. 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's parotid-masseteric fascia. Practice caution when further exposing the nerve, for the following reasons.
    1. Avoid aggressive dissection, or perpendicular spreads, to guard against stretch-mediated neuropraxia injury.
    2. 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.
    3. Avoid damaging the superficial temporal artery through broad medially and inferiorly directed dissection. Note that an injury will be identified by brisk, pulsatile bleeding.
      1. 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.
  10. Trace the main trunk distally by dissecting along the nerve in an inferior direction, distally from the exit of the stylomastoid foramen.
    1. 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.
  11. Induce the desired injury patterns as follows.
    1. For a crush injury, use smooth-surfaced jeweler'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.
    2. 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.
    3. 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.

5. Wound closure

  1. Irrigate the wound with sterile saline and dry it with sterile gauze.
  2. 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.

6. Postoperative recovery

  1. Administer a subcutaneous injection of nonsteroidal anti-inflammatory analgesic (such as 0.05 mg/kg of buprenorphine and 0.5 mg /kg Carprofen) for postoperative pain control. Place the injection along the animal's back.
  2. Cease the administration of the anesthetic agent and allow the animal to inhale oxygen for an additional 1 min.
  3. 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.
  4. 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.
  5. Monitor the animals 2x per day to evaluate for signs of malnourishment, corneal irritation, or surgical site infection, and maintain appropriate surgical logs.
    1. Administer 0.9% sterile saline in a subcutaneous fashion if there is significant weight loss.
    2. Apply lubricating eye ointment daily until the animal's blink reflex is re-established.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

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.

Figure 1
Figure 1: 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. Please click here to view a larger version of this figure.

Figure 2
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 ± standard error of the mean. Please click here to view a larger version of this figure.

Figure 3
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 µm. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

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's disease and Parkinson'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.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have nothing to disclose.

Acknowledgments

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

Materials

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

DOWNLOAD MATERIALS LIST

References

  1. Chan, J. Y. K., Byrne, P. J. Management of facial paralysis in the 21st century. Facial Plastic Surgery. 27, (4), 346-357 (2011).
  2. Razfar, A., Lee, M. K., Massry, G. G., Azizzadeh, B. Facial Paralysis Reconstruction. Otolaryngologic Clinics of North America. 49, (2), 459-473 (2016).
  3. Couch, S. M., Chundury, R. V., Holds, J. B. Subjective and objective outcome measures in the treatment of facial nerve synkinesis with onabotulinumtoxinA (Botox). Ophthalmic Plastic and Reconstructive Surgery. 30, (3), 246-250 (2014).
  4. Wei, L. A., Diels, J., Lucarelli, M. J. Treating buccinator with botulinum toxin in patients with facial synkinesis: A previously overlooked target. Ophthalmic Plastic and Reconstructive Surgery. 32, (2), 138-141 (2016).
  5. Cooper, L., Lui, M., Nduka, C. Botulinum toxin treatment for facial palsy: A review. Journal of Plastic, Reconstructive and Aesthetic Surgery. 70, (6), 833-841 (2017).
  6. Choi, K. H., et al. Botulinum toxin injection of both sides of the face to treat post-paralytic facial synkinesis. Journal of Plastic, Reconstructive and Aesthetic Surgery. 66, (8), 1058-1063 (2013).
  7. Yang, X. N., et al. Peripheral nerve repair with epimysium conduit. Biomaterials. 34, (22), 5606-5616 (2013).
  8. Banks, C. A., et al. Long-term functional recovery after facial nerve transection and repair in the rat. Journal of Reconstructive Microsurgery. 31, (3), 210-216 (2015).
  9. Hadlock, T. A., Kowaleski, J., Lo, D., MacKinnon, S. E., Heaton, J. T. Rodent facial nerve recovery after selected lesions and repair techniques. Plastic and Reconstructive Surgery. 125, (1), 99-109 (2010).
  10. Hadlock, T., et al. The effect of electrical and mechanical stimulation on the regenerating rodent facial nerve. Laryngoscope. 120, (6), 1094-1102 (2009).
  11. Hadlock, T., et al. Functional assessments of the rodent facial nerve: A synkinesis model. Laryngoscope. 118, (10), 1744-1749 (2008).
  12. Diogo, C. C., et al. The use of sheep as a model for studying peripheral nerve regeneration following nerve injury: review of the literature. Neurological Research. 39, (10), 926-939 (2017).
  13. Wanner, R., et al. Functional and Molecular Characterization of a Novel Traumatic Peripheral Nerve–Muscle Injury Model. NeuroMolecular Medicine. 19, (2-3), 357-374 (2017).
  14. Olmstead, D. N., et al. Facial nerve axotomy in mice: A model to study motoneuron response to injury. Journal of Visualized Experiments. (96), e52382 (2015).
  15. Maeda, T., Hori, S., Sasaki, S., Maruo, S. Effects of tension at the site of coaptation on recovery of sciatic nerve function after neurorrhaphy: Evaluation by walking-track measurement, electrophysiology, histomorphometry, and electron probe X-ray microanalysis. Microsurgery. 19, (4), 200-207 (1999).
  16. Zhang, F., Inserra, M., Richards, L., Terris, D. J., Lineaweaver, W. C. Quantification of nerve tension after nerve repair: Correlations with nerve defects and nerve regeneration. Journal of Reconstructive Microsurgery. 17, (6), 445-451 (2001).
  17. Macfarlane, B. V., Wright, A., Benson, H. A. E. Reversible blockade of retrograde axonal transport in the rat sciatic nerve by vincristine. Journal of Pharmacy and Pharmacology. 49, (1), 97-101 (1997).
  18. Stromberg, B. V., Vlastou, C., Earle, A. S. Effect of nerve graft polarity on nerve regeneration and function. Journal of Hand Surgery. 4, (5), 444-445 (1979).
  19. Sotereanos, D. G., et al. Reversing nerve-graft polarity in a rat model: The effect on function. Journal of Reconstructive Microsurgery. 8, (4), 303-307 (1992).
  20. Whitlock, E. L., et al. Ropivacaine-induced peripheral nerve injection injury in the rodent model. Anesthesia and Analgesia. 111, (1), 214-220 (2010).
  21. Lloyd, B. M., et al. Use of motor nerve material in peripheral nerve repair with conduits. Microsurgery. 27, (2), 138-145 (2007).
  22. Kawamura, D. H., et al. Regeneration through nerve isografts is independent of nerve geometry. Journal of Reconstructive Microsurgery. 21, (4), 243-249 (2005).
  23. Brenner, M. J., et al. Repair of motor nerve gaps with sensory nerve inhibits regeneration in rats. Laryngoscope. 116, (9), 1685-1692 (2006).
  24. Brenner, M. J., et al. Role of timing in assessment of nerve regeneration. Microsurgery. 28, (4), 265-272 (2008).
  25. Heaton, J. T., et al. A system for studying facial nerve function in rats through simultaneous bilateral monitoring of eyelid and whisker movements. Journal of Neuroscience Methods. 171, (2), 197-206 (2008).
  26. Magill, C. K., Moore, A. M., Borschel, G. H., Mackinnon, S. E. A new model for facial nerve research: The novel transgenic Thy1-GFP rat. Archives of Facial Plastic Surgery. 12, (5), 315-320 (2010).
  27. Guntinas-Lichius, O., et al. Factors limiting motor recovery after facial nerve transection in the rat: Combined structural and functional analyses. European Journal of Neuroscience. 21, (2), 391-402 (2005).
  28. Skouras, E., et al. Manual stimulation, but not acute electrical stimulation prior to reconstructive surgery, improves functional recovery after facial nerve injury in rats. Restorative Neurology and Neuroscience. 27, (3), 237-251 (2009).
  29. Bischoff, A., et al. Manual stimulation of the orbicularis oculi muscle improves eyelid closure after facial nerve injury in adult rats. Muscle and Nerve. 39, (2), 197-205 (2009).
  30. Schulz, A., Walther, C., Morrison, H., Bauer, R. In vivo electrophysiological measurements on mouse sciatic nerves. Journal of Visualized Experiments. (86), e51181 (2014).
  31. Placheta, E., et al. Macroscopic in vivo imaging of facial nerve regeneration in Thy1-GFP rats. JAMA Facial Plastic Surgery. 17, (1), 8-15 (2015).
  32. Moore, A. M., et al. A transgenic rat expressing green fluorescent protein (GFP) in peripheral nerves provides a new hindlimb model for the study of nerve injury and regeneration. Journal of Neuroscience Methods. 204, (1), 19-27 (2012).
  33. Grosheva, M., et al. Early and continued manual stimulation is required for long-term recovery after facial nerve injury. Muscle and Nerve. 57, (1), 100-106 (2018).
  34. Grosheva, M., et al. Comparison of trophic factors' expression between paralyzed and recovering muscles after facial nerve injury. A quantitative analysis in time course. Experimental Neurology. 279, 137-148 (2016).
  35. Grosheva, M., et al. Local stabilization of microtubule assembly improves recovery of facial nerve function after repair. Experimental Neurology. 209, (1), 131-144 (2008).
  36. Angelov, D. N., et al. Mechanical stimulation of paralyzed vibrissal muscles following facial nerve injury in adult rat promotes full recovery of whisking. Neurobiology of Disease. 26, (1), 229-242 (2007).
  37. Tomov, T. L., et al. An example of neural plasticity evoked by putative behavioral demand and early use of vibrissal hairs after facial nerve transection. Experimental Neurology. 178, (2), 207-218 (2002).
  38. Streppel, M., et al. Focal application of neutralizing antibodies to soluble neurotrophic factors reduces collateral axonal branching after peripheral nerve lesion. European Journal of Neuroscience. 15, (8), 1327-1342 (2002).
  39. Peeva, G. P., et al. Improved outcome of facial nerve repair in rats is associated with enhanced regenerative response of motoneurons and augmented neocortical plasticity. European Journal of Neuroscience. 24, (8), 2152-2162 (2006).
  40. Pavlov, S. P., et al. Manually-stimulated recovery of motor function after facial nerve injury requires intact sensory input. Experimental Neurology. 211, (1), 292-300 (2008).
  41. Guntinas-Lichius, O., et al. Transplantation of olfactory ensheathing cells stimulates the collateral sprouting from axotomized adult rat facial motoneurons. Experimental Neurology. 172, (1), 70-80 (2001).
  42. Guntinas-Lichius, O., Angelov, D. N., Stennert, E., Neiss, W. F. Delayed hypoglossal-facial nerve suture after predegeneration of the peripheral facial nerve stump improves the innervation of mimetic musculature by hypoglossal motoneurons. Journal of Comparative Neurology. 387, (2), 234-242 (1997).
  43. Sinis, N., et al. Electrical stimulation of paralyzed vibrissal muscles reduces endplate reinnervation and does not promote motor recovery after facial nerve repair in rats. Annals of Anatomy. 191, (4), 356-370 (2009).
  44. Kiryakova, S., et al. Recovery of whisking function promoted by manual stimulation of the vibrissal muscles after facial nerve injury requires insulin-like growth factor 1 (IGF-1). Experimental Neurology. 222, (2), 226-234 (2010).
  45. Banati, R. B., et al. Early and rapid de novo synthesis of Alzheimer βA4-Amyloid precursor protein (APP) in activated microglia. Glia. 9, (3), 199-210 (1993).
  46. Blinzinger, K., Kreutzberg, G. Displacement of synaptic terminals from regenerating motoneurons by microglial cells. Zeitschrift für Zellforschung und Mikroskopische Anatomie. 85, (2), 145-157 (1968).
  47. Rieske, E., et al. Microglia and microglia-derived brain macrophages in culture: generation from axotomized rat facial nuclei, identification and characterization in vitro. Brain Research. 492, (1-2), 1-14 (1989).
  48. Matsumoto, K., et al. Peripheral nerve regeneration across an 80-mm gap bridged by a polyglycolic acid (PGA)-collagen tube filled with laminin-coated collagen fibers: A histological and electrophysiological evaluation of regenerated nerves. Brain Research. 868, (2), 315-328 (2000).
  49. Mattsson, P., Janson, A. M., Aldskogius, H., Svensson, M. Nimodipine promotes regeneration and functional recovery after intracranial facial nerve crush. Journal of Comparative Neurology. 437, (1), 106-117 (2001).
  50. Yian, C. H., Paniello, R. C., Gershon Spector, J. Inhibition of motor nerve regeneration in a rabbit facial nerve model. Laryngoscope. 111, (5), 786-791 (2001).
  51. Angelov, D. N., et al. Nimodipine accelerates axonal sprouting after surgical repair of rat facial nerve. Journal of Neuroscience. 16, (3), 1041-1048 (1996).

Comments

0 Comments


    Post a Question / Comment / Request

    You must be signed in to post a comment. Please sign in or create an account.

    Usage Statistics