The authors would like to thank Autumn Peterson for assistance with mouse photography, Daniel Thoresen for assistance with mice, and Drs. Nicole Ward and Abdelmadjid Belkadi for assistance with surgical denervation. These studies were supported by funding from the National Institute of Arthritis and Musculoskeletal and Skin Diseases (grants R00AR059796 and R01AR065409); the University of Michigan Department of Dermatology; the Biological Sciences Scholars Program; the Center for Organogenesis; the University of Michigan Comprehensive Cancer Center; and the John S. and Suzanne C. Munn Cancer Fund. S.C.P. was supported by funding from the National Institute of General Medical Sciences (grant T32 GM007315). This work was also supported by the NIH Intramural Research Program, Center for Cancer Research, National Cancer Institute.

All procedures described in this protocol were performed in accordance with regulations established by the University of Michigan Unit for Laboratory Animal Medicine.
1. Induce Genetic Recombination in Mice
Note: The Gli1tm3(cre/ERT2)Alj/J mouse strain (Gli1-CreERT2)13 enables targeting of tamoxifen-induced genetic recombination to TD epithelia. Cross this strain with B6.129S4-Gt(ROSA)26Sortm1Sor/J reporter mice (LacZ)22 to generate Gli1;LacZ animals to visualize TD cells by whole-mount staining below.
<ol>
	<li>Prepare tamoxifen solution to a concentration of 12.5 mg/ml in corn oil.
	<ol>
		<li>In a 1.5 ml tube, add up to 20 mg of crystalline tamoxifen, and then 1 ml of corn oil. Firmly tape the tube to a vortex mixer, and vortex continuously at the highest setting at RT until the tamoxifen has fully dissolved (2-4 hr), as confirmed by examining the tube under a dissecting microscope for the absence of tamoxifen particulates.</li>
		<li>Transfer the solution to a 15 ml tube and dilute the tamoxifen to a final concentration of 12.5 mg/ml with additional corn oil. Mix by vortexing the viscous solution for an additional 30 sec. Store this solution for up to 1 week at 4 &#176;C in the dark.</li>
	</ol>
	</li>
	<li>Inject the tamoxifen solution intraperitoneally into Gli1;LacZ mice, at a volume of 200 &#181;l per 20 g of mouse body weight, for an effective tamoxifen dose of 2.5 mg per 20 g mouse weight.</li>
</ol>
2. Harvest Skin Biopsies
Note: Depending on the experiment, harvest skin biopsies several days to weeks after tamoxifen induction.&#160;For all surgeries, follow standard protocols for rodent surgery, including using sterile gloves, wearing a surgical mask or hair net, and covering the animal with a sterile surgical drape during the procedure.
<ol>
	<li>Prepare 10x stock anesthetic solution by mixing 90 mg/ml ketamine and 6.5 mg/ml xylazine in water. Dilute this stock solution 1:10 into sterile PBS just prior to use, and store at RT in the dark for up to 8 months.
	<ol>
		<li>Alternatively, anesthetize mice by isoflurane inhalation, beginning with a gas concentration of 4% with oxygen to fully anesthetize the animal, and then subsequently lowering this to 1-2% for the duration of the procedure.</li>
	</ol>
	</li>
	<li>Inject the anesthetic solution intraperitoneally at a dose of 200 &#181;l per 20 g mouse body weight. Check that the animal has reached the proper plane of sedation by toe pinch assay, and confirm that heart and respiratory rates are normal (approximately 600 beats and 160 breaths per min, respectively).</li>
	<li>Use an electric clipper to remove the hair from the site of biopsy on the dorsal back skin, being careful not to nick or damage the underlying skin.</li>
	<li>Prepare the surgical site by wiping the shaved area in an anterior-to-posterior direction using Betadine and alcohol wipes. Ensure all hair clippings are removed from the site.</li>
	<li>Outline the biopsy site using a black marker,&#160;&#160;place the animal on a warming pad in an aseptic surgical area, and cover with a sterile surgical drape (for demonstration purposes, sterile drape was omitted to increase visibility).&#160; &#160; 
	Note: To obtain longitudinal sections of hair follicles, the longer edge of the biopsy (the edge to be sectioned for histology, ~1 cm) should run in an anterior-posterior direction (parallel to the direction of the hair follicles), parasagital to the dorsal midline (Figure 1A).</li>
	<li>Use a sterile #11 scalpel to make a full thickness excision along the marked area without damaging the underlying muscular fascia. 
	Note: The excised skin tissue includes the epidermis, dermis, subcutaneous fat and panniculus carnosus. Bleeding is typically minimal.</li>
	<li>Flatten the excised skin sample on a dry paper towel, dermis side down, trim away the excess paper towel, and store the sample in cold PBS for up to 1 hr if other samples need to be collected. When ready, proceed to Steps 3.1 or 3.2 to process samples for histology, or Step 4 for whole-mount &#946;-galactosidase (LacZ) staining.</li>
	<li>Suture close the biopsy site using 6-0 nylon sutures, in a simple interrupted pattern spaced roughly 3 mm apart.</li>
	<li>Do not return mice that have undergone surgery into the same cage as other animals until after full recovery.</li>
	<li>Monitor animals immediately after surgery until they regain consciousness, and also daily until the surgical area has healed, typically within 1 week. Use analgesics in accordance with designated institutional animal care and use guidelines if mice exhibit signs of pain or distress. Remove sutures within 7-10 days after surgery. 
	Note: If needed, prepare analgesic solution by diluting carprofen (50 mg/ml stock solution) 1:100 in sterile water.&#160;&#160;Inject the solution subcutaneously between the shoulder blades near the scruff of the neck, at a dose of 200 &#181;l per 20 g body weight (5 mg/kg mouse body weight).</li>
</ol>
3. Process Samples for Histology
Note: To fix and process the excised tissue, use either method below depending on application.
<ol>
	<li>To generate paraffin-embedded histological samples, fix the skin in 3.7% formalin in PBS O/N at RT and store in 70% ethanol for up to 2 weeks. Remove the paper towel before embedding into paraffin.</li>
	<li>For generating frozen histological samples, submerge the tissue in cold 4% paraformaldehyde in PBS and gently shake for 1 hr. Remove the solution and wash the sample with 3 changes of PBS, roughly 5 min each. Next, submerge the sample in 30% sucrose in PBS to cryoprotect the tissue (&#34;sucrose sinking&#34;).</li>
	<li>Incubate with gentle shaking O/N at 4 &#176;C. The next day, remove the paper towel and trim away excess adipose tissue from the dermal side of the skin. Embed the tissue directly into OCT and store the frozen block at -80 &#176;C. 
	Note: After sectioning, either paraffin or frozen samples can be stained by immunohistochemistry to identify TDs, Merkel cells and nerves using antibodies against Keratin 17, Keratin 8 and Neurofilament, respectively, as previously described5,19.</li>
</ol>
4. Visualize Samples by Whole-mount LacZ Staining
<ol>
	<li>Prepare X-gal staining solution.
	<ol>
		<li>Combine 0.94 g sodium phosphate monobasic, and 2.6 g sodium phosphate dibasic in 250 ml of sterile water. Adjust pH to 7.3. To this, add 0.5 ml of 1 M magnesium chloride, 0.528 g of potassium ferrocyanide, and 0.412 g of potassium ferricyanide. Add 250 &#181;l of octylphenyl-polyethylene glycol and 125 mg of deoxycholate. The base solution can be stored at 4 &#176;C for up to 6 months in the dark.</li>
		<li>Prepare 50x stock X-gal solution by adding dimethylformamide to the X-gal stock bottle to generate a 50 mg/ml solution. Store this solution at -20 &#176;C in the dark.</li>
		<li>Just prior to use, dilute stock X-gal solution 1:50 into X-gal base solution to generate staining solution. For smaller biopsies (&#60;1 cm2), aliquot 1-2 ml of staining solution per sample.</li>
	</ol>
	</li>
	<li>Fix the skin sample collected in Step 2.7 in a solution containing 2% paraformaldehyde/0.2% glutaraldehyde in cold PBS for 30 min, gently shaking on ice. For smaller biopsies (&#60;1 cm2), use 1-2 ml of fixative solution per sample. 
	Note: Alternatively, fix samples in 2-4% paraformaldehyde only, or in 0.5% glutaraldehyde only. Optimal fixation conditions depend on the tissue, degree of LacZ expression and application.</li>
	<li>Remove the fixative solution, and rinse samples with 3 changes of PBS, 5 min each, on a shaker at RT.</li>
	<li>Remove the paper towel underneath the sample and cut away excess adipose tissue from the dermal side of the skin by gripping the fat with blunt forceps and trimming with dissecting scissors.</li>
	<li>Submerge the sample in X-gal staining solution, and incubate at 37 &#176;C O/N. LacZ expression will be visible as a blue stain under a dissecting microscope (Figure 1B). 
	Note: If the signal intensity is weak, replace the staining solution the next day and repeat the O/N incubation. If the background staining is too intense, reduce the time of staining, or incubate the sample at RT instead of 37 &#176;C.</li>
	<li>Remove the staining solution and wash the samples in 3 changes of PBS containing 3% DMSO for approximately 5 min, gently shaking at RT.</li>
	<li>Wash samples in 2-3 changes of 70% ethanol for 5 min&#160;each. Store samples in 70% ethanol.</li>
</ol>
5. Surgical Denervation
<ol>
	<li>Anesthetize the animal as in Step 2.2 and shave the entire dorsal skin.</li>
	<li>Prepare the surgical area of the back skin using Betadine and alcohol wipes,&#160;and cover the animal with a sterile surgical drape (for demonstration purposes, sterile drape was omitted to increase visibility).&#160;&#160;Keep the animal warm using a heating pad while operating in an aseptic surgical area.</li>
	<li>Make an incision using a sterile #11 scalpel along the dorsal midline from the base of the neck to roughly 0.5 cm above the tail.</li>
	<li>Using blunt forceps, gently reflect the skin on the left side away from the flank to visualize the underlying tissue from the scapular fat pads near the neck to just above the hind limb. 
	Note: Dorsal cutaneous nerves appear as white strands that travel caudally through the translucent fascia of the trunk wall before making sharp bends and entering the loose connective tissue underneath the skin (Figure 2).</li>
	<li>Using ultra-fine forceps under a dissecting light microscope, remove the nerves exclusively from the left side of the animal located at anatomical sites T3-12 by plucking from where the segments bend at the trunk wall to their entry sites into the skin (Figure 2).
	<ol>
		<li>Orient forceps vertically and remove the nerves by grasping approximately 0.5 cm below their bend sites and pulling upwards, causing the nerve to stretch and separate from the surrounding tissue (Figure 2C-E). Be careful to avoid rupturing adjacent blood vessels.</li>
		<li>Continue until all nerves extending from the trunk wall to the skin are removed. Do not disrupt the nerves within the dense fascia of the trunk wall. Keep the tissue moist throughout the procedure by periodically applying drops of sterile 0.9% saline solution.</li>
		<li>Alternatively, remove nerves by grasping their proximal ends near the trunk wall with forceps and snipping with fine scissors. Afterwards, sever the distal ends near the skin (Figures 2F-H). Finally, remove the intervening nerve segments (Figure 2I).</li>
	</ol>
	</li>
	<li>Remove any nerves from the skin flap exposed in Step 5.4. These fibers comprise the distal branches of the dorsal cutaneous nerves and appear as white branching strands located sporadically within the connective tissue on the dermal side of the skin flap (Figures 2J-K).
	<ol>
		<li>To remove these fine branches, position the fine forceps roughly parallel to the dermal surface, grasp the nerves and pluck upwards to avoid disrupting blood vessels and puncturing the skin. Continue until all visible nerves have been removed.</li>
	</ol>
	</li>
	<li>Using blunt dissection, reflect the skin on the right side of the dorsal midline incision, but do not remove the nerves. This will serve as the contralateral sham-operated control.</li>
	<li>Suture along the dorsal midline in a simple interrupted pattern&#160;&#160;to close the incision. Monitor the animal during recovery and post-operatively as previously demonstrated (Steps 2.8-10). Remove sutures within 7-10 days after surgery.</li>
	<li>To functionally assess stable denervation up to several weeks after surgery, remove the hair from the dorsal skin using an electric clipper.</li>
	<li>Gently prick the denervated skin area using a hypodermic needle, and note whether the animal responds, typically by shuddering or turning its head. If the skin area has been stably denervated, the animal will exhibit little or no response.</li>
	<li>Using a black marker, outline the area of no response, as well as an area of similar size and location on the contralateral sham side.</li>
	<li>Collect biopsies from these sites as in Steps 2.1-2.9 for analysis. 
	Note: Alternatively, the entire dorsal back skin, including denervated and sham-operated regions, can be removed as a single sheet for whole-mount staining, similar to as described in Step 4.</li>
</ol>

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The authors have nothing to disclose.

Nerves serve crucial functions not only in sensation, but also in mammalian organ development, maintenance and regeneration13,24-27. As nerves have recently been implicated in diverse skin disorders, the techniques described here can be used to study the requirement for innervation in a variety of animal disease models. Indeed, the unilateral denervation technique allows for the direct comparison of skin with either intact or disrupted nerves from the same mouse. This provides an ideal internal control to comp...

Over the past few years, there has been a widening appreciation for the influence of nerves on diseases not typically regarded as classical neuropathies1-4. In the skin, recent experimental evidence has suggested that sensory nerves can modulate diverse pathologies ranging from psoriasis to cancer5-9. This has been demonstrated using techniques such as surgical denervation and pharmacological inhibition of neural function in rodents. In the case of psoriasis, these studies have provided a mechanistic framework for understanding why human psoriatic plaques regress following loss of neural function7,10-12.
Cutaneous nerves can also affect gene expression13,14 and are critical for mechanosensing in normal skin15. In particular, touch dome (TD) epithelia are comprised of a patch of columnar epidermal cells in juxtaposition with neuroendocrine Merkel cells innervated by slowly adapting type 1 (SA1) nerve fibers16-18. TDs mediate light touch sensation and have been shown to display Hedgehog pathway activity5,19. TD maintenance depends on innervation20,21, as nerves secrete Hedgehog ligands to sustain normal TDs and their associated Merkel cells19. In addition, innervation promotes Hedgehog-dependent tumor formation from TD epithelia5. Together, these studies reinforce the notion that intricate molecular interactions occurring between nerves and the surrounding cells in their niche are crucial for normal TD physiology as well as disease.
To interrogate the nature of these interactions, we describe here a series of in vivo techniques for manipulating gene expression in the TD, as well as for harvesting skin biopsies for TD visualization after lineage tracing. Finally, we describe procedures for performing unilateral surgical denervation, wherein nerves are removed from one side of the mouse dorsal skin, while leaving the contralateral side intact as a sham internal control. Several weeks after surgery, denervated and sham control skin are compared to assess changes that occur when nerves are ablated. Although these techniques are described in the context of normal TDs, the denervation procedure has been used to examine the requirement for nerves in mouse models of psoriasis6, wound healing13 and tumorigenesis5. Finally, since the skin is amenable to repeated biopsies, this provides an opportunity to monitor the long-term fates of labeled cells or to assess disease progression over multiple time points.

<table><tbody><tr><td>Alcohol prep pads</td><td>PDI</td><td>B339</td><td></td></tr><tr><td>AnaSed (Xylazine)</td><td>Lloyd</td><td>NADA 139-236</td><td></td></tr><tr><td>Antibody, anti-Keratin 8</td><td>Developmental Studies Hybridoma Bank</td><td>TROMA-I</td><td>rat antibody, use at 1:500 concentration</td></tr><tr><td>Antibody, anti-Keratin 17</td><td>Cell Signaling</td><td>#4543</td><td>rabbit antibody, use at 1:1,000 concentration</td></tr><tr><td>Antibody, anti-Neurofilament</td><td>Cell Signaling</td><td>C28E10</td><td>rabbit antibody, use at 1:500 concentration</td></tr><tr><td>Betadine prep pads</td><td>Medline</td><td>MDS093917</td><td></td></tr><tr><td>Carprofen (Rimadyl)</td><td>Zoetis</td><td></td><td></td></tr><tr><td>Cordless rechargable clipper</td><td>Wahl</td><td>trimmer model 8900</td><td></td></tr><tr><td>Corn Oil</td><td>Sigma-Aldrich</td><td>C8267</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM1860</td><td></td></tr><tr><td>DAPI</td><td>ThermoFisher Scientific</td><td>D1306</td><td>use at 1:1,000 concentration</td></tr><tr><td>Deoxycholate</td><td>Sigma-Aldrich</td><td>D6750</td><td></td></tr><tr><td>Depilatory Cream</td><td>Nair</td><td>N/A</td><td></td></tr><tr><td>Dimethylforamide</td><td>Sigma-Aldrich</td><td>319937</td><td></td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D8418</td><td></td></tr><tr><td>Glutaraldehyde</td><td>Sigma-Aldrich</td><td>G5882</td><td></td></tr><tr><td>ImmEdge Pen</td><td>Vector Laboratories</td><td>H-4000</td><td></td></tr><tr><td>Ketamine HCl</td><td>Hospira</td><td>NDC 0409-2051-05</td><td></td></tr><tr><td>Magnesium chloride</td><td>Sigma</td><td>M8266</td><td></td></tr><tr><td>Micro cover glass</td><td>VWR</td><td>48404-454</td><td></td></tr><tr><td>Micro Slides</td><td>VWR</td><td>48311-703</td><td></td></tr><tr><td>10% Neutral Buffered Formalin</td><td>VWR</td><td>BDH0502-4LP</td><td></td></tr><tr><td>6-0 nylon sutures</td><td>DemeTECH</td><td>NL166012F4P</td><td></td></tr><tr><td>Octylphenyl-polyethylene glycol</td><td>Sigma-Aldrich</td><td>I8896</td><td></td></tr><tr><td>O.C.T. Compound</td><td>Sakura Tissue-Tek</td><td>4583</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td></td></tr><tr><td>Pottasium ferrocyanide</td><td>Sigma-Aldrich</td><td>P9387</td><td></td></tr><tr><td>Pottasium ferricyanide</td><td>Sigma-Aldrich</td><td>702587</td><td></td></tr><tr><td>Sodium phosphate monobasic</td><td>Sigma-Aldrich</td><td>P9791</td><td></td></tr><tr><td>Sodium phosphate dibasic</td><td>Sigma-Aldrich</td><td>S5136</td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>84097</td><td></td></tr><tr><td>Tamoxifen</td><td>Sigma-Aldrich</td><td>T5648-1G</td><td></td></tr><tr><td>Ultra fine forceps</td><td>Dumont</td><td>0103-5-PO</td><td></td></tr><tr><td>Vectashield</td><td>Vector Laboratories</td><td>H1000</td><td></td></tr><tr><td>X-gal</td><td>Roche</td><td>10 651 745 001</td><td>Disolve in dimethylforamide to create 50x stock prior to use</td></tr></tbody></table>

cutaneous surgical denervation:  a method for testing the requirement for nerves in mouse models of skin disease

Cutaneous somatosensory nerves function to detect diverse stimuli that act upon the skin. In addition to their established sensory roles, recent studies have suggested that nerves may also modulate skin disorders including atopic dermatitis, psoriasis and cancer. Here, we describe protocols for testing the requirement for nerves in maintaining a cutaneous mechanosensory organ, the touch dome (TD). Specifically, we discuss methods for genetically labeling, harvesting and visualizing TDs by whole-mount staining, and for performing unilateral surgical denervation on mouse dorsal back skin. Together, these approaches can be used to directly compare TD morphology and gene expression in denervated as well as sham-operated skin from the same animal. These methods can also be readily adapted to examine the requirement for nerves in mouse models of skin pathology. Finally, the ability to repeatedly sample the skin provides an opportunity to monitor disease progression at different stages and times after initiation.

By generating mice expressing tamoxifen-inducible Gli1-CreERT2 and a LacZ reporter allele, it is possible to visualize TD epithelia and track the fates of these cells over time. The entire denervation procedure typically can be completed within 1 hr per mouse and should cause minimal distress to the animal.
Our previous studies have indicated that nerves are crucial for maintaining both normal TDs as well as th...

Watch this Scientific Journal Video about Cutaneous Surgical Denervation:  A Method for Testing the Requirement for Nerves in Mouse Models of Skin Disease at JoVE.com

Cutaneous Surgical Denervation:  A Method for Testing the Requirement for Nerves in Mouse Models of Skin Disease

This article includes detailed protocols for genetic labeling of mouse skin, surgical denervation, skin biopsy and visualizing labeled epithelia by whole-mount &#946;-galactosidase staining. These methods can be used to test the requirement for nerves in mouse models of normal and pathological skin.

Cutaneous somatosensory nerves function to detect diverse stimuli that act upon the skin. In addition to their established sensory roles, recent studies ...

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Research

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Medicine

9.7K Views.  University of Michigan. This article includes detailed protocols for genetic labeling of mouse skin, surgical denervation, skin biopsy and visualizing labeled epithelia by whole-mount β-galactosidase staining. These methods can be used to test the requirement for nerves in mouse models of normal and pathological skin.

Video: Cutaneous Surgical Denervation:  A Method for Testing the Requirement for Nerves in Mouse Models of Skin Disease

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 ...

Tibial Nerve Transection - A Standardized Model for Denervation-induced Skeletal Muscle Atrophy in Mice

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of denervation-induced skeletal muscle atrophy in rodents. Although originally developed and used extensively in the rat due to its larger size, the tibial nerve in mice is big enough that it can be easily manipulated with either crush or transection, leaving the peroneal and sural nerve branches of the sciatic nerve intact and thereby preserving their target muscles. Thus, this model offers the advantages of inducing less morbidity and impediment of ambulation than the sciatic nerve transection model and also allows investigators to study the physiologic, cellular and molecular biologic mechanisms regulating the process of muscle atrophy in genetically engineered mice. The tibial nerve supplies the gastrocnemius, soleus and plantaris muscles, so its transection permits the study of denervated skeletal muscle composed of fast twitch type II fibers and/or slow twitch type I fibers. Here we demonstrate the tibial nerve transection model in the C57Black6 mouse. We assess the atrophy of the gastrocnemius muscle, as a representative muscle, at 1, 2, and 4 weeks post-denervation by measuring muscle weights and fiber type specific cross-sectional area on paraffin-embedded histologic sections immunostained for fast twitch myosin.

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of skeletal muscle atrophy. The model surgical protocol is described and demonstrated in C57Black6 mice. &#160;

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of denervation-induced skeletal muscle atrophy in rodents. ...

The Sciatic Nerve Cuffing Model of Neuropathic Pain in Mice

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

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

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

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Neuroscience

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

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

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

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

Establishing a Mouse Model of a Pure Small Fiber Neuropathy with the Ultrapotent Agonist of Transient Receptor Potential Vanilloid Type 1

Patients with diabetes mellitus (DM) or those experiencing the neurotoxic effects of chemotherapeutic agents may develop sensation disorders due to degeneration and injury of small-diameter sensory neurons, referred to as small fiber neuropathy. Present animal models of small fiber neuropathy affect both large- and small-diameter sensory fibers and thus create a neuropathology too complex to properly assess the effects of injured small-diameter sensory fibers. Therefore, it is necessary to develop an experimental model of pure small fiber neuropathy to adequately examine these issues. This protocol describes an experimental model of small fiber neuropathy specifically affecting small-diameter sensory nerves with resiniferatoxin (RTX), an ultrapotent agonist of transient receptor potential vanilloid type 1 (TRPV1), through a single dose of intraperitoneal injection, referred to as RTX neuropathy. This RTX neuropathy showed pathological manifestations and behavioral abnormalities that mimic the clinical characteristics of patients with small fiber neuropathy, including intraepidermal nerve fiber (IENF) degeneration, specifically injury in small-diameter neurons, and induction of thermal hypoalgesia and mechanical allodynia. This protocol tested three doses of RTX (200, 50, and 10 &#181;g/kg, respectively) and concluded that a critical dose of RTX (50 &#181;g/kg) is required for the development of typical small fiber neuropathy manifestations, and prepared a modified immunostaining procedure to investigate IENF degeneration and neuronal soma injury. The modified procedure is fast, systematic, and economic. Behavioral evaluation of neuropathic pain is critical to reveal the function of small-diameter sensory nerves. The evaluation of mechanical thresholds in experimental rodents is particularly challenging and this protocol describes a customized metal mesh that is suitable for this type of assessment in rodents. In summary, RTX neuropathy is a new and easily established experimental model to evaluate the molecular significance and intervention underlying neuropathic pain for the development of therapeutic agents.

This study establishes an experimental model of pure small fiber neuropathy with resiniferatoxin (RTX). A unique dose of RTX (50 &#181;g/kg) is optimal for developing a small fiber neuropathy model that mimics patient characteristics and could help investigate the nociceptive molecular significance underlying neuropathic pain.

Patients with diabetes mellitus (DM) or those experiencing the neurotoxic effects of chemotherapeutic agents may develop sensation disorders due to ...

Modified Spared Nerve Injury Surgery Model of Neuropathic Pain in Mice

Spared nerve injury (SNI) is an animal model that mimics the cardinal symptoms of peripheral nerve injury for studying the molecular and cellular mechanism of neuropathic pain in mice and rats. Currently, there are two types of SNI model, one to cut and ligate the common peroneal and the tibial nerves with intact sural nerve, which is defined as SNIs in this study, and another to cut and ligate the common peroneal and the sural nerves with intact tibial nerve, which is defined as SNIt in this study. Because the sural nerve is purely sensory whereas the tibial nerve contains both motor and sensory fibers, the SNIt model has much less motor deficit than the SNIs model. In the traditional SNIt mouse model, the common peroneal and the sural nerves are cut and ligated separately. Here a modified SNIt surgery method is described to damage both common peroneal and sural nerves with only one ligation and one cut with a shorter procedure time, which is easier to perform and reduces the potential risk of stretching the sciatic or tibial nerves, and produces similar mechanical hypersensitivity as the traditional SNIt model.

The modified surgery is a simplified method for mouse or rat spared nerve injury model that requires only one ligation and one cut to injure both common peroneal and sural nerves.

Spared nerve injury (SNI) is an animal model that mimics the cardinal symptoms of peripheral nerve injury for studying the molecular and cellular ...

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 ...

Partial Sciatic Nerve Ligation: A Mouse Model of Chronic Neuropathic Pain to Study the Antinociceptive Effect of Novel Therapies

Management of chronic pain remains challenging to this day, and current treatments are associated with adverse effects, including tolerance and addiction. Chronic neuropathic pain results from lesions or diseases in the somatosensory system. To investigate potential therapies with reduced side effects, animal pain models are the gold standard in preclinical studies. Therefore, well-characterized and well-described models are crucial for the development and validation of innovative therapies. 
Partial ligation of the sciatic nerve (pSNL) is a procedure that induces chronic neuropathic pain in mice, characterized by mechanical and thermal hypersensitivity, ongoing pain, and changes in limb temperature, making this model a great fit to study neuropathic pain preclinically. pSNL is an advantageous model to study neuropathic pain as it reproduces many symptoms observed in humans with neuropathic pain. Furthermore, the surgical procedure is relatively fast and straightforward to perform. Unilateral pSNL of one limb allows for comparison between the ipsilateral and contralateral paws, as well as evaluation of central sensitization. 
To induce chronic neuropathic hypersensitivity, a 9-0 non-absorbable nylon thread is used to ligate the dorsal third of the sciatic nerve. This article describes the surgical procedure and characterizes the development of chronic neuropathic pain through multiple commonly used behavioral tests. As a plethora of innovative therapies are now being investigated to treat chronic pain, this article provides crucial concepts for standardization and an accurate description of surgeries required to induce neuropathic pain.

Partial sciatic nerve ligation induces long-lasting chronic neuropathic pain, characterized by exaggerated responses to thermal and mechanical stimuli. This mouse model of neuropathic pain is commonly used to study innovative therapies for pain management. This article describes in detail the surgical procedure to improve standardization and reproducibility.

Management of chronic pain remains challenging to this day, and current treatments are associated with adverse effects, including tolerance and addiction. ...

Targeting Mouse Sympathetic Neurons: A Lumbar Sympathectomy Approach

Surgical Lumbar Sympathectomy in Mice

Peripheral nerve injuries are common, and full functional recovery after injury is achieved in only 10% of patients. The sympathetic nervous system plays many critical roles in maintaining bodily homeostasis, but it has rarely been studied in the context of peripheral nerve injury. The extent of postganglionic sympathetic neuronal functions in distal targets in the periphery is currently unclear. To better explore the role of sympathetic innervation of peripheral targets, a surgical &#34;knock-out&#34; model provides an alternative approach. Although this can be achieved chemically, chemical destruction of postganglionic sympathetic neurons can be nonspecific and dose-dependent. The use of a surgical lumbar sympathectomy in mice, once thought to be &#34;virtually not practicable&#34; in small animals, allows for specific targeting of postganglionic sympathetic neurons that innervate the hind limbs. This manuscript describes how to surgically remove the L2-L5 lumbar sympathetic ganglia from a mouse as a survival surgery, which reliably decreases the hind paw sweat response and the number of sympathetic axons in the sciatic nerve.

Author Spotlight: Exploring Plasticity of Sympathetic Neurons

This manuscript presents a protocol for surgically removing the postganglionic lumbar sympathetic neurons from a mouse. This procedure will facilitate a multitude of studies aimed at investigating the role of sympathetic innervation in distal tissue targets.

Peripheral nerve injuries are common, and full functional recovery after injury is achieved in only 10% of patients. The sympathetic nervous system plays ...

Cutaneous Surgical Denervation: A Method for Testing the Requirement for Nerves in Mouse Models of Skin Disease

Summary

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Chapters in this video

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

Tibial Nerve Transection - A Standardized Model for Denervation-induced Skeletal Muscle Atrophy in Mice

The Sciatic Nerve Cuffing Model of Neuropathic Pain in Mice

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

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

Establishing a Mouse Model of a Pure Small Fiber Neuropathy with the Ultrapotent Agonist of Transient Receptor Potential Vanilloid Type 1

Modified Spared Nerve Injury Surgery Model of Neuropathic Pain in Mice

Traumatic Peripheral Nerve Injury in Mice

Partial Sciatic Nerve Ligation: A Mouse Model of Chronic Neuropathic Pain to Study the Antinociceptive Effect of Novel Therapies

Surgical Lumbar Sympathectomy in Mice