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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Surgical Denervation
<ol>
	<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>Prepare the surgical area of the back skin using Betadine and alcohol wipes, and cover the animal with a sterile surgical drape (for demonstration purposes, sterile drape was omitted to increase visibility).&#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 1).</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 1).
	<ol>
		<li>Orient forceps vertically and removes 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 1C-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 1F-H). Finally, remove the intervening nerve segments (Figure 1I).</li>
	</ol>
	</li>
	<li>Remove any nerves from the skin flap exposed in Step 1.4. 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 1J-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; to close the incision 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; 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>
	<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>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>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>6-0 nylon sutures</td>
			<td>DemeTECH</td>
			<td>NL166012F4P</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra fine forceps</td>
			<td>Dumont</td>
			<td>0103-5-PO</td>
			<td></td>
		</tr>
	</tbody>
</table>

testing sensory nerve function in a surgically denervated mouse model

Source: Peterson, S. C. et al., <a href="https://app.jove.com/v/54050">Cutaneous Surgical Denervation: A Method for Testing the Requirement for Nerves in Mouse Models of Skin Disease.</a> J. Vis. Exp. (2016)
This video demonstrates testing sensory nerve function in a mouse model by surgically denervating one side and leaving the other as a control. The response to mechanical stimulation confirms the role of sensory nerves.

<img alt="Figure 1" src="/files/ftp_upload/22852/22852_Figure 1.jpg" /> 
Figure 1. Two approaches for denervating dorsal skin. (A) Cartoon diagram of innervated mouse skin. Red dotted lines indicate a single excision made along the dorsal midline to expose the underlying musculature on the trunk wall (purple) as well as the dermis (grey, asterisk) beneath the reflected skin. Dorsal cutaneous nerves traveling caudally appear to &#34;bend&#34; as they leave the trunk w...

Testing Sensory Nerve Function in a Surgically Denervated Mouse Model

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Begin with an anesthetized mouse with exposed dorsal skin. Disinfect the area and cover it. 
 Make a midline incision from the neck to just above the tail. 
 Pull the left-side skin to create a skin flap and expose the underlying tissue. 
 Place the mouse under a microscope and locate the sensory nerves. Carefully pull and remove these nerves without disrupting the blood vessels. 
 Moisten the tissue and remove the remaining nerves from the skin flap, creating a denervated area. 
 On the right side, leave the nerves intact as a control. Suture the skin incision. 
 Allow the mouse to recover. 
To assess stable denervation several weeks after surgery, use the same denervated mouse with shaved dorsal skin. 
 Using a needle, prick at the control area, generating a mechanical stimulation. 
 This excites the underlying nerves, sending a signal to the brain and causing the mouse to respond. 
 However, pricking the denervated area fails to generate a response, confirming the role of sensory nerves.

testing-sensory-nerve-function-in-a-surgically-denervated-mouse-model

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

61 Views. Źródło: Peterson, S. C. et al., Chirurgiczne odnerwienie skóry: metoda testowania zapotrzebowania na nerwy w mysich modelach chorób skóry. J. Vis. Exp. (2016) Ten film demonstruje testowanie funkcji nerwu czuciowego w modelu mysim poprzez chirurgiczne odnerwienie jednej strony i pozostawienie drugiej jako kontroli. Reakcja na stymulację mechaniczną potwierdza rolę nerwów czuciowych. 

Video: Testowanie funkcji nerwów czuciowych w chirurgicznie odnerwionym modelu myszy - Concept

Subcutaneous Trigeminal Nerve Field Stimulation for Refractory Facial Pain

Chronic or neuropathic trigeminal facial pain can be challenging to treat. Neurosurgical procedures should be applied when conservative treatment fails. Neuromodulation techniques for chronic facial pain include deep brain stimulation and motor cortex stimulation, which are complex to perform. Subcutaneous nerve field stimulation is certified for chronic back pain and is the least invasive form of neuromodulation. We applied this technique to treat chronic and neuropathic trigeminal pain as an individual therapy concept. First, trial stimulation is performed. Subcutaneous leads are placed in the painful trigeminal dermatome under local anesthesia. The leads are connected to an external neurostimulator that applies constant stimulation. Patients undergo a 12 day outpatient trial to assess the effect of the stimulation. Electrodes are removed after the trial. If the patient reports pain reduction of at least 50% in intensity and/or attack frequency, a reduction in medication or increase in quality of life, permanent implantation is scheduled. New electrodes are implanted under general anesthesia and are subcutaneously tunneled to an infraclavicular internal pulse generator. Patients are able to turn stimulation on and off and to increase or decrease the stimulation amplitude as needed.
This technique represents a minimal invasive alternative to other more invasive means of neuromodulation for trigeminal pain such as motor cortex stimulation or deep brain stimulation.

Treating chronic or neuropathic facial pain can be challenging when medical or standard treatment fails. Subcutaneous nerve field stimulation is the least invasive form of neuromodulation and is used for chronic back pain. We applied this technology to treat chronic and neuropathic trigeminal facial pain.

Chronic or neuropathic trigeminal facial pain can be challenging to treat. Neurosurgical procedures should be applied when conservative treatment fails. ...

JoVE Journal

Medicine

The Muscle Cuff Regenerative Peripheral Nerve Interface for the Amplification of Intact Peripheral Nerve Signals

Robotic exoskeletons have gained recent acclaim within the field of rehabilitative medicine as a promising modality for functional restoration for those individuals with extremity weakness. However, their use remains largely confined to research institutions, frequently operating as a means of static extremity support as motor detection methods remain unreliable. Peripheral nerve interfaces have arisen as a potential solution to this shortcoming; however, due to their inherently small amplitudes, these signals can be difficult to differentiate from background noise, lowering their overall motor detection accuracy. As current interfaces rely on abiotic materials, inherent material breakdown can occur alongside foreign body tissue reaction over time, further impacting their accuracy. The Muscle Cuff Regenerative Peripheral Nerve Interface (MC-RPNI) was designed to overcome these noted complications. Consisting of a segment of free muscle graft secured circumferentially to an intact peripheral nerve, the construct regenerates and becomes reinnervated by the contained nerve over time. In rats, this construct has demonstrated the ability to amplify a peripheral nerve's motor efferent action potentials up to 100 times the normal value through the generation of compound muscle action potentials (CMAPs). This signal amplification facilitates high accuracy detection of motor intent, potentially enabling reliable utilization of exoskeleton devices.

This manuscript provides an innovative method for developing a biologic peripheral nerve interface termed the Muscle Cuff Regenerative Peripheral Nerve Interface (MC-RPNI). This surgical construct can amplify its associated peripheral nerve's motor efferent signals to facilitate accurate detection of motor intent and the potential control of exoskeleton devices.

Robotic exoskeletons have gained recent acclaim within the field of rehabilitative medicine as a promising modality for functional restoration for those ...

Bioengineering

Improved Renal Denervation Mitigated Hypertension Induced by Angiotensin II Infusion

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the regulatory mechanism of RDN on hypertension remains elusive. Thus, it&#39;s essential to establish a simpler RDN model in mice. In this study, osmotic mini pumps filled with Angiotensin II were implanted in 14-week-old C57BL/6 mice. One week after the implantation of the mini-osmotic pump, a modified RDN procedure was performed on bilateral renal arteries of the mice using phenol. Age-sex-matched mice were given saline and served as sham group. Blood pressure was measured at baseline and every week subsequently for 21 days. Then, renal artery, abdominal aorta and heart were collected for histological examination using H&#38;E and Masson staining. In this study, we present a simple, practical, repeatable, and standardized RDN model, which can control hypertension and alleviate cardiac hypertrophy. The technique can denervate peripheral renal sympathetic nerves without renal artery damage. Compared to previous models, the modified RDN facilitates the study of the pathobiology and pathophysiology of hypertension.

Here, we present a protocol for renal sympathetic denervation (RDN) in mice with hypertension induced by Angiotensin II infusion. The procedure is repeatable, convenient and allows to study the regulatory mechanisms of RDN on hypertension and cardiac hypertrophy.

The benefits of renal sympathetic denervation (RDN) on blood pressure have been proved in a large number of clinical trials in recent years. However, the ...

Testing Sensory Nerve Function in a Surgically Denervated Mouse Model

Transkrypcja

Testing Sensory Nerve Function in a Surgically Denervated Mouse Model