This work was supported by the NIH National Institute of Neurological Disorders and Stroke under award number K01NS124912 and in part by a developmental grant from the NIH-funded Emory Specialized Center of Research Excellence in Sex Differences U54AG062334 and the Medical Scientist Training Program of Emory University School of Medicine. Thank you to David Kim, postbaccalaureate, for sectioning sciatic nerves and to HaoMin SiMa, research specialist, for 3D printing a phone mount for our stereo microscope that allowed for the filming of&#160;the video.

All experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Emory University (under the IACUC protocol number PROTO202100177). Four adult female wild-type C57BL/6J mice, aged 14 weeks and weighing between 16-21 g, were used in this study. The details of the reagents and equipment used here are listed in the Table of Materials.
1. Presurgical preparation
<ol>
	<li>Autoclave the surgical tools: 1 pair of sharp scissors, 2 fine-tipped tweezers, 1 needle driver.</li>
	<li>Warm a heating pad to 37 &#176;C and place it under the surgical tabletop.</li>
	<li>Spray the surgical area, including the anesthesia induction chamber, with disinfectant and wipe it down thoroughly with paper towels.</li>
	<li>Wash hands and don proper personal protective equipment for mouse handling.</li>
	<li>Administer meloxicam orally at 5 mg/kg body weight, one drop at a time as the mouse swallows. Allow plenty of time for the animal to completely swallow the medicine to avoid aspiration during induction.</li>
	<li>Induce the mouse under anesthesia with 3% isoflurane in 1 L/min of oxygen (following institutionally approved protocols).</li>
	<li>Once the mouse is anesthetized (no reaction to toe pinch), place the mouse on the surgical tabletop, supine, with their nose in a fitted nose cone.</li>
	<li>Once the mouse is properly fitted in the nose cone, adjust the isoflurane to 2% for maintenance of anesthesia.</li>
	<li>Place ophthalmic lubricant on the mouse&#39;s eyes to prevent keratoconjunctivitis sicca (dry eye).</li>
	<li>Ensure the mouse is properly secured with surgical tape to the surgical tabletop.</li>
	<li>Monitor continuously for ease of breathing and adequate breathing rate.</li>
	<li>Shave the fur from the mouse&#39;s abdomen from the level of the genitals to below the ribs.</li>
	<li>Once the fur is removed, first wipe the surgical area in a circular motion, from central to peripheral, with 70% ethanol. Then, wipe the surgical area with betadine in the same circular motion. Repeat both the ethanol and betadine steps&#160;a total of 3 times.</li>
	<li>Put on a surgical mask, a hair cover, a sterile gown, and sterile gloves.</li>
	<li>Drape the mouse with a sterile surgical drape with an appropriately sized hole cut out of the middle to visualize the surgical field. The rhomboid shape can be made by folding the drape in half and cutting an isosceles triangle with a height of ~10 mm and a base of ~15 mm.</li>
</ol>
2. Incisions
<ol>
	<li>Using a pair of sharp scissors and fine-tipped tweezers, create a midline incision from ~1 mm above the level of the pubic symphysis to ~2 mm below the ribs.</li>
	<li>Identify the midline fascia (linea alba) between the bilateral rectus abdominis muscles. Using the tweezers, lift the abdominal muscles away from the underlying organs and cut along the linea alba to enter the abdominal cavity. 
	NOTE: The linea alba can be identified by gently pulling the bilateral rectus abdominus muscles laterally to reveal a thinner fascia line that lies longitudinally along the midline32. Making the incision through the linea alba allows the muscle to be closed more easily.</li>
	<li>Retract the abdominal muscles and skin with 5-0 sutures laterally to visualize the next steps properly.</li>
</ol>
3. Identification of the L2-L5 lumbar sympathetic ganglia
<ol>
	<li>The lumbar sympathetic ganglia lie posterior to the abdominal aorta and inferior vena cava. To visualize the aorta, partially remove the intestines from the abdominal cavity. 
	NOTE: The L2-L5 level sympathetic ganglia traverse from the level of the aortic bifurcation into the iliac vessels to the level of the left renal artery21,33.
	<ol>
		<li>Lay a sterile cotton square saturated in sterile saline on the drape, lying superiorly and to the left of the incision. Carefully, with&#160;2 sterile cotton-tipped applicators, push the colon and small intestine partially&#160;out onto the saline-soaked cotton square. 
		NOTE: Ensure that the cecum and appendix are out of the abdominal cavity and that the descending colon can be visualized.</li>
		<li>Cover the exposed intestines with another saline-saturated cotton square.</li>
		<li>Periodically monitor peristalsis throughout the surgery, which can be identified by rhythmic movement of the intestines.</li>
	</ol>
	</li>
	<li>Identify the descending colon, which is the portion of the intestine that is traveling towards the rectum and anus and may contain visible fecal matter, and deflect this structure to the left with closed tweezers to reveal the abdominal aorta and inferior vena cava. These two vessels are bound by connective tissue&#160;and should move as one unit.
	<ol>
		<li>With a pair of tweezers, lift the abdominal vessels by their surrounding connective tissue. While this is lifted, carefully place a 5-0 nylon suture through the connective tissue. 
		CAUTION: Do NOT puncture the abdominal vessels with the needle: This will lead to rapid decompensation of the mouse and likely death.</li>
		<li>Once the suture has been inserted through the connective tissue, deflect the abdominal vessels to the left. This will give a triangular window, allowing direct visualization of the bilateral psoas muscles. 
		NOTE: Due to the intimate connections that the ganglia have with the abdominal vessels, the sympathetic ganglia, though typically located midline, will be deflected slightly to the left (mouse&#39;s right) along with the vessels. Once the abdominal vessels are deflected, one should visualize a triangle, with the two sides on the left consisting of the deflected abdominal vessels and the right side defined by the mouse&#39;s midline (between the bilateral psoas muscles). The sympathetic ganglia lie in the middle of this triangle (Figure 1A). They travel vertically side-by-side and appear translucent. They may overlap a small vessel that dives into the mouse&#39;s midline and lies flat on its right psoas muscle. Use tweezers to bluntly dissect any overlying fascia to reveal the ganglia (Figure 1B).</li>
	</ol>
	</li>
	<li>Identify the left renal artery and the left testicular or ovarian artery. 
	NOTE: These are large vessels that will form a quadrangle, the sides of which consist of the left renal artery superiorly, the animal&#39;s midline and abdominal aorta/inferior vena cava laterally, and the left testicular or ovarian artery inferiorly. The L2 level ganglia are large and lie within this quadrangle (Figure 1A). Carefully blunt dissect to enter this quadrangle with tweezers and identify the bilateral L2 ganglia (Figure 1B).</li>
	<li>Once the lower and upper parts of the ganglia are identified, grab the inferior aspect of the visible bilateral ganglia with a pair of tweezers and pull upwards. Visualization of more inferiorly-located ganglia, such as L4 and L5, can be increased by gently pushing any organs obstructing the view inferiorly with the second pair of tweezers. 
	NOTE: Visualizing the midline perforating vessel mentioned in step 3.2.2&#160;will ensure a more complete lumbar sympathectomy. Holding a second pair of tweezers in the other hand, pull the fascia and neuronal connections holding the ganglia in the abdomen away while simultaneously pulling up on the ganglia themselves with the first pair of tweezers. Only one pair of tweezers should hold onto the ganglia, while the other clears the fascia and neuronal connections. Once at the level of the left testicular or ovarian artery, it may be helpful for L2 identification if the lower chain ganglia are still intact, as the L2 ganglia will move when pulled by the lower chain. The L3-5 ganglia can be broken at the testicular or ovarian artery before extracting the L2 ganglia from the quadrangle with tweezers. The only ganglia present within the quadrangle are the L2 ganglia.</li>
	<li>This procedure should have minimal blood loss. If bleeding occurs, ensure adequate hemostasis prior to closure.</li>
</ol>
4. Skin closure
<ol>
	<li>After adequate hemostasis is achieved, remove the suture holding the abdominal vessels in place.</li>
	<li>Using 2 sterile cotton-tipped applicators, carefully replace the exteriorized intestines into the abdominal cavity.</li>
	<li>With a 5-0 absorbable suture, perform a running stitch to approximate the abdominal muscles.</li>
	<li>With 5-0 nylon suture, use simple interrupted stitches to close the skin.</li>
	<li>Apply a generous layer of antibiotic ointment, such as Neosporin, to the incision site.</li>
	<li>Place the mouse in a clean cage on a heating pad.</li>
	<li>Observe the mouse every 15 min until it is awake and ambulatory. This usually takes less than 30 min.</li>
</ol>
5. Pilocarpine sweat assay
NOTE: To assess for depletion of sympathetic functional activity following a lumbar sympathectomy, a pilocarpine sweat assay was utilized 7 days post-lumbar sympathectomy.
<ol>
	<li>Prepare a 1% pilocarpine hydrochloride solution in 0.9% NaCl alongside a mixture of 10% potato starch in castor oil.</li>
	<li>Using a paintbrush, cover the plantar surface of the foot with betadine. Allow this layer to dry completely.</li>
	<li>Once the betadine layer is dry, use a separate paintbrush to apply a layer of 10% starch in castor oil.</li>
	<li>Administer 0.25 &#181;L/g body weight of 1% pilocarpine subcutaneously using a syringe into the loose skin over the neck. Start a timer immediately after pilocarpine administration.</li>
	<li>At 8 min post-injection, take photos of the plantar surface of the foot.</li>
	<li>Using Fiji, count the number of dark spots located on all six (6) footpads (Figure 2A).</li>
</ol>
6. Immunohistochemistry
NOTE: To assess the degeneration of sympathetic axons in the peripheral nerves post-lumbar sympathectomy, the bilateral sciatic nerves were harvested on postoperative day 21.
<ol>
	<li>Anesthetize the mice (following step 1 of the protocol), harvest the bilateral sciatic nerves, and then promptly euthanize the animal (following institutionally approved protocols).</li>
	<li>Place the sciatic nerves directly into 4% paraformaldehyde in 0.01 M phosphate-buffered saline (PBS) for 20 min, then transfer them to 20% sucrose in 0.1 M PBS for overnight cryoprotection at 4 &#176;C.</li>
	<li>Section the sciatic nerves longitudinally using a cryostat at 20 &#181;m, and place the sections on charged slides.</li>
	<li>Block the nerve sections with 10% normal goat serum (NGS) in tris buffered saline with 1% Tween 20 (TBST) for 1 h at room temperature.</li>
	<li>Remove the blocking buffer and replace it with the primary antibodies rabbit anti-tyrosine hydroxylase and chicken anti-neurofilament-heavy diluted in the blocking buffer (10% NGS in TBST) at 1:750 and 1:1000, respectively. Allow the primary antibodies to incubate overnight in a humidity chamber at room temperature.</li>
	<li>Wash slide with TBST 3 times, 10 min each wash, before applying the secondary antibodies goat anti-rabbit 647 and goat anti-chicken 488, both diluted at 1:200 in the blocking buffer. Allow the secondary antibodies to incubate in a humidity chamber for 2 h at room temperature.</li>
	<li>Wash the slide with TBST 4 times, 10 min each wash, and allow the slide to dry in a light-protected area prior to mounting.</li>
	<li>Image nerve sections at least 40 &#181;m apart on a fluorescent microscope on the 10x objective.</li>
	<li>On Fiji, straighten the nerve sections and draw three randomly placed vertical lines spanning the width of the section. Count the number of axons crossing each vertical line and divide that by the width of the section at the vertical line. Repeat for each of the three vertical lines per section. Average the three values obtained per section, and further average the values of the three sections per nerve.</li>
</ol>

Targeting Mouse Sympathetic Neurons: A Lumbar Sympathectomy Approach

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wallerian+degeneration:+Gaining+perspective+on+inflammatory+events+after+peripheral+nerve+injury.">Wallerian degeneration: Gaining perspective on inflammatory events after peripheral nerve injury.</a> J Neuroinflammation. 8 (1), 1-13 (2011).</li><li>Babetto, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeting+nmnat1+to+axons+and+synapses+transforms+its+neuroprotective+potency+in+vivo.">Targeting nmnat1 to axons and synapses transforms its neuroprotective potency in vivo.</a> J Neuroscience. 30 (40), 13291-13304 (2010).</li><li>Brumovsky, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dorsal+root+ganglion+neurons+and+tyrosine+hydroxylase-an+intriguing+association+with+implications+for+sensation+and+pain.">Dorsal root ganglion neurons and tyrosine hydroxylase-an intriguing association with implications for sensation and pain.</a> Pain. 157 (2), 314 (2016).</li><li>Tian, T., Harris, A., Owyoung, J., Sima, H., Ward, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conditioning+electrical+stimulation+fails+to+enhance+sympathetic+axon+regeneration.">Conditioning electrical stimulation fails to enhance sympathetic axon regeneration.</a> bioRxiv. , (2023).</li><li>Tian, T., Ward, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+ThCre:+Mtmg+mouse+has+sparse+expression+in+the+sympathetic+nervous+system.">The ThCre: Mtmg mouse has sparse expression in the sympathetic nervous system.</a> bioRxiv. , 2023 (2023).</li><li>Ohman-Gault, L., Huang, T., Krimm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transcription+factor+Phox2b+distinguishes+between+oral+and+non-oral+sensory+neurons+in+the+geniculate+ganglion.">The transcription factor Phox2b distinguishes between oral and non-oral sensory neurons in the geniculate ganglion.</a> J Comparative Neurol. 525 (18), 3935-3950 (2017).</li><li>Pattyn, A., Morin, X., Cremer, H., Goridis, C., Brunet, J. -. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+homeobox+gene+phox2b+is+essential+for+the+development+of+autonomic+neural+crest+derivatives.">The homeobox gene phox2b is essential for the development of autonomic neural crest derivatives.</a> Nature. 399 (6734), 366-370 (1999).</li><li>Fran&#231;ois, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sympathetic+innervation+of+the+interscapular+brown+adipose+tissue+in+mouse.">Sympathetic innervation of the interscapular brown adipose tissue in mouse.</a> Ann N Y Acad Sci. 1454 (1), 3-13 (2019).</li></ol>

The authors have no conflicts of interest to disclose.

The lumbar sympathetic ganglia are very small structures located behind many critical abdominal organs and large vessels. Therefore, this procedure requires significant precision and accuracy. Much of the difficulty lies in identifying the sympathetic ganglia intraoperatively. It is suggested that the learner first be able to identify the ganglia in a mouse cadaver prior to attempting this procedure in a live mouse. Troubleshooting will often need to occur when identifying the sympathetic ganglia after the diversion of t...

Peripheral nerve injuries (PNIs) can lead to motor, sensory, and sympathetic deficits in distal tissue targets that rarely fully functionally recover1. PNI research has often focused on the&#160;motor and sensory regeneration; however, nearly one-quarter of the rat sciatic nerve consists of unmyelinated sympathetic axons2.&#160;The role of sympathetic innervation in the peripheral tissues, nevertheless, is not fully understood3. The sympathetic nervous system plays a major role in maintaining bodily homeostasis, participating in immune regulation, thermoregulation, vascular tone, mitochondrial biogenesis, and more4,5,6,7,8,9,10,11. When sympathetic innervation at the neuromuscular junction is lost, persistent muscle weakness and synaptic instability are observed despite the maintenance of motoneuron innervation12. This sympathetic regulation of synaptic transmission at the neuromuscular junction has been shown to decline with aging13,14, which contributes to sarcopenia, defined as the age-dependent reduction in muscle mass, force, and power15. A better understanding of the role of sympathetic innervation of peripheral tissues is necessary for the development of therapies that will optimize functional outcomes for patients with PNIs and other forms of sympathetic dysfunction.
A sympathectomy is a powerful experimental tool that will allow for investigations of the role of sympathetic innervation in distal target tissues. Specifically, removal of the L2-L5 level sympathetic ganglia removes a majority of the sympathetic innervation to the lower limbs, which is especially useful for investigators interested in the sciatic nerve.
This protocol details the removal of L2-L5 level postganglionic sympathetic neurons from a mouse as a survival surgery. This procedure requires rodent microsurgical skills and familiarity with mouse anatomy, and when performed effectively, does not cause any visible phenotypic differences. A surgical lumbar sympathectomy has been used in rodent research, more so in rats than in mice16,17,18,19,20,21; however, a detailed protocol describing the protocol does not currently exist. Previous studies utilizing the lumbar sympathectomy have primarily focused on the role of sympathetic innervation in the pain response, which is generally attenuated by sympathectomy in various nerve injury models. Fewer studies have used this technique in mice22, likely due to the smaller size of anatomic landmarks, as the use of surgical sympathectomy was believed to be &#34;virtually not practicable&#34; in small animals23,24. Localized sympathectomies in the form of microsympathectomies have also been utilized in rodent models, also mostly in the context of pain behaviors25,26,27. The microsympathectomy, in contrast to the total lumbar sympathectomy, utilizes a dorsal approach through which a segment of the gray ramus to a specific spinal nerve is disconnected and removed, allowing for a very targeted sympathectomy that will avoid wider spread side effects.
Because mouse models are critical for many studies requiring genetic manipulation, this procedure will have versatile applications beyond the breadth of peripheral nerve injuries as well. Using a transabdominal approach, the lumbar sympathetic ganglia can be reliably visualized and resected from the mouse with no apparent adverse effects. Although protocols for the chemical destruction of postganglionic sympathetic neurons are available, such as the use of 6-hydroxydopamine (6-OHDA)23,24, this surgical procedure allows for anatomically specific targeting of the postganglionic lumbar sympathetic ganglia. The use of a surgical sympathectomy also avoids the nonspecific and dose-dependent concerns related to pharmacological methods28,29.
The use of chemical sympathectomies via administration of 6-OHDA was described in 1967 as a simple way to achieve selective destruction of adrenergic nerve endings since surgical sympathectomies in small animals were not favored23,24. 6-OHDA is a catecholaminergic neurotoxin that is endogenously formed in patients with Parkinson&#39;s disease, and its toxicity is derived from its ability to form free radicals and inhibit the electron transport chain in mitochondria30,31. Through norepinephrine uptake-1 transport mechanisms, 6-OHDA is able to accumulate within noradrenergic neurons, such as postganglionic sympathetic neurons28. Eventually, the neuron is destroyed by 6-OHDA; however, terminals in the peripheral nervous system do regenerate, with the restoration of functional activity even when the amine levels are still reduced. Different dosage thresholds are also present for different organs in response to 6-OHDA, and higher doses of 6-OHDA have been shown to exhibit more nonspecific effects, extending its neurotoxic consequences to non-catecholamine-containing neurons and even non-neuronal cells. Aside from noradrenergic neurons, dopaminergic neurons are affected by 6-OHDA as well29, making the chemical sympathectomy ultimately less specific to postganglionic sympathetic neurons than the surgical sympathectomy.
Therefore, a surgical lumbar sympathectomy enables the targeted ablation of the sympathetic innervation to the lower limbs, which can be combined with a variety of experimental techniques and genetic manipulations in the mouse to study how the sympathetic nervous system contributes to various injury and disease states.

<table><tbody><tr><td>5-0 absorable suture</td><td>CP Medical</td><td>421A</td><td></td></tr><tr><td>5-0 nylon suture</td><td>Med-Vet International</td><td>MV-661</td><td></td></tr><tr><td>70% ethanol</td><td>Sigma-Aldrich</td><td>E7023-4L</td><td></td></tr><tr><td>Anesthesia Induction Chamber</td><td>Kent Scientific VetFlo</td><td>VetFlo-0530XS</td><td></td></tr><tr><td>Anesthesia Vaporizer</td><td>Kent Scientific VetFlo</td><td>13-005-202</td><td></td></tr><tr><td>Betadine</td><td>HealthyPets</td><td>BET16OZ</td><td></td></tr><tr><td>C57BL/6J mice</td><td>Jackson Laboratory</td><td>#000664</td><td></td></tr><tr><td>Chicken anti-neurofilament-heavy</td><td>Abcam</td><td>ab72996</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM1850</td><td></td></tr><tr><td>Data Analysis Software</td><td>Prism</td><td></td><td></td></tr><tr><td>Fine-tipped tweezers</td><td>World Precision Instruments</td><td>500233</td><td></td></tr><tr><td>Fluorescent microscope</td><td>Nikon</td><td>Ti-E</td><td></td></tr><tr><td>Goat anti-chicken 488</td><td>Invitrogen</td><td>A32931</td><td></td></tr><tr><td>Goat anti-rabbit 647</td><td>Invitrogen</td><td>A21245</td><td></td></tr><tr><td>Heating pad</td><td>Braintree Scientific</td><td>39DP</td><td></td></tr><tr><td>Image Analysis Software</td><td>Fiji</td><td></td><td></td></tr><tr><td>Imaging Software</td><td>Nikon</td><td>NIS-Elements</td><td></td></tr><tr><td>Isoflurane</td><td>Med-Vet International</td><td>RXISO-250</td><td></td></tr><tr><td>Meloxicam</td><td>Med-Vet International</td><td>RXMELOXIDYL32</td><td></td></tr><tr><td>Needle driver</td><td>Roboz Surgical Store</td><td>RS-7894</td><td></td></tr><tr><td>Normal Goat Serum</td><td>Abcam</td><td>ab7481</td><td></td></tr><tr><td>Ophthalmic ointment</td><td>Refresh</td><td>Refresh P.M.</td><td></td></tr><tr><td>&lt;em&gt;Phox2bCre:tdTomato&lt;/em&gt; mutant mice</td><td>Jackson Laboratory&amp;nbsp;</td><td>#016223, #007914</td><td></td></tr><tr><td>Pilocarpine hydrochloride</td><td>Sigma-Aldrich</td><td>P6503</td><td></td></tr><tr><td>Rabbit anti-tyrosine hydroxylase</td><td>Abcam</td><td>ab112</td><td></td></tr><tr><td>Small straight scissors&amp;nbsp;</td><td>Fine Science Tools</td><td>14084-09</td><td></td></tr><tr><td>Sterile cotton swabs 2x2</td><td>Dynarex</td><td>3252</td><td></td></tr><tr><td>Sterile cotton tipped applicators</td><td>Dynarex</td><td>4301</td><td></td></tr><tr><td>Sterile drape</td><td>Med-Vet International</td><td>DR4042</td><td></td></tr><tr><td>Sterile saline solution</td><td>Med-Vet International</td><td>1070988-BX</td><td></td></tr><tr><td>&lt;em&gt;ThCre:mTmG&lt;/em&gt; mutant mice</td><td>Mutant Mouse Resource and Research Centers</td><td>strain #017262-UCD</td><td>Jackson Laboratory, strain #007576</td></tr><tr><td>&lt;em&gt;ThCre:tdTomato &lt;/em&gt;mutant mice</td><td>European Mouse Mutant Archive</td><td>strain #00254</td><td>Jackson Laboratory, strain #007914</td></tr></tbody></table>

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.

This protocol describes the surgical removal of postganglionic lumbar sympathetic neurons from a mouse. Two mice received lumbar sympathectomies, and two mice served as controls. To achieve a successful surgical lumbar sympathectomy, adequate visualization of at least the L2 and L3 bilateral lumbar sympathetic ganglia must be achieved, as seen in Figure 1. Removal of the L4 and L5 ganglia would achieve complete sympathetic denervation of the lower body; however, visualization of the lower ga...

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.

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 ...

Research

JoVE Journal

Neuroscience

2.0K Views.  Emory University School of Medicine. Ons onderzoek richt zich op de plasticiteit van sympathische neuronen. Enkele van de vragen waarin we geïnteresseerd zijn, zijn onder meer hoe postganglionische sympathische axonen regenereren in vergelijking met andere neurontypen, en hoe sympathische innervatie de functie van distale weefsels die ze innerveren reguleert. Het is bekend dat het sympathische zenuwstelsel verantwoordelijk is voor een breed scala aan homeostatische mechanismen.Meer recen...

Video: Auteur in de schijnwerpers: onderzoek naar plasticiteit van sympathische neuronen

Modified Technique for Coronary Artery Ligation in Mice

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI we need better knowledge about pathophysiology of myocardial ischemia. This knowledge may be valuable to define new therapeutic targets for innovative cardiovascular therapies4. Experimental MI model in mice is an increasingly popular small-animal model in preclinical research in which MI is induced by means of permanent or temporary ligation of left coronary artery (LCA)5. In this video, we describe the step-by-step method of how to induce experimental MI in mice.
The animal is first anesthetized with 2% isoflurane. The unconscious mouse is then intubated and connected to a ventilator for artificial ventilation. The left chest is shaved and 1.5 cm incision along mid-axillary line is made in the skin. The left pectoralis major muscle is bluntly dissociated until the ribs are exposed. The muscle layers are pulled aside and fixed with an eyelid-retractor. After these preparations, left thoracotomy is performed between the third and fourth ribs in order to visualize the anterior surface of the heart and left lung. The proximal segment of LCA artery is then ligated with a 7-0 ethilon suture which typically induces an infarct size ~40% of left ventricle. At the end, the chest is closed and the animals receive postoperative analgesia (Temgesic, 0.3 mg/50 ml, ip). The animals are kept in a warm cage until spontaneous recovery.

The surgical procedure used to induce experimental myocardial infarction in mice begins with left thoracotomy between the third and the fourth ribs in order to visualize the anterior surface of the heart and left lung. The left coronary artery is ligated, the chest is closed and the mouse is allowed to recover spontaneously.

Myocardial infarction (MI) is one of the most important causes of mortality in humans1-3. In order to improve morbidity and mortality in patients with MI ...

Medicine

Surgical Transplantation of Mouse Neural Stem Cells into the Spinal Cords of Mice Infected with Neurotropic Mouse Hepatitis Virus

Mice infected with the neurotropic JHM strain of mouse hepatitis virus (MHV) develop pathological and clinical outcomes similar to patients with the demyelinating disease Multiple Sclerosis (MS). We have shown that transplantation of NSCs into the spinal cords of sick mice results in a significant improvement in both remyelination and in clinical outcome. Cell replacement therapies for the treatment of chronic neurologic diseases are now a reality and in vivo models are vital in understanding the interactions between the engrafted cells and host tissue microenvironment. This presentation provides an adapted method for transplanting cells into the spinal cord of JHMV-infected mice. In brief, we provide a procedure for i) preparation of NSCs prior to transplant, ii) pre-operative care of mice, iii) exposure of the spinal cord via laminectomy, iv) stereotactic injection of NSCs, and iv) post-operative care.

The transplantation of mouse neural stem cells (NSCs) into the spinal cords of mice with established demyelination is detailed. The preparation of NSCs, the laminectomy of thoracic vertebra 9 (T9), and transplantation of NSCs is outlined along with the pre- and post-operative care of the mice.

Mice infected with the neurotropic JHM strain of mouse hepatitis virus (MHV) develop pathological and clinical outcomes similar to patients with the ...

Surgical Technique for Superior Cervical Ganglionectomy in a Murine Model

Growing evidence suggests that the sympathetic nervous system plays an important role in cancer progression. Adrenergic innervation regulates salivary gland secretion, circadian rhythm, macular degeneration, immune function, and cardiac physiology. Murine surgical sympathectomy is a method for studying the effects of adrenergic innervation by allowing for complete, unilateral adrenergic ablation while avoiding the need for repeated pharmacologic intervention and the associated side effects. However, surgical sympathectomy in mice is technically challenging because of the small size of the superior cervical ganglion. This study describes a surgical technique for reliably identifying and resecting the superior cervical ganglion to ablate the sympathetic nervous system. The successful identification and removal of the ganglion are validated by imaging the fluorescent sympathetic ganglia using a transgenic mouse, identifying post-resection Horner's syndrome, staining for adrenergic markers in the resected ganglia, and observing diminished adrenergic immunofluorescence in the target organs following sympathectomy. This model enables future studies of cancer progression as well as other physiological processes regulated by the sympathetic nervous system.

The present protocol describes a mouse model of the ablation of adrenergic innervation by identifying and resecting the superior cervical ganglion.

Growing evidence suggests that the sympathetic nervous system plays an important role in cancer progression. Adrenergic innervation regulates salivary ...

Cancer Research

Permanent Ligation of the Left Anterior Descending Coronary Artery in Mice: A Model of Post-myocardial Infarction Remodelling and Heart Failure

Heart failure is a syndrome in which the heart fails to pump blood at a rate commensurate with cellular oxygen requirements at rest or during stress. It is characterized by fluid retention, shortness of breath, and fatigue, in particular on exertion. Heart failure is a growing public health problem, the leading cause of hospitalization, and a major cause of mortality. Ischemic heart disease is the main cause of heart failure.
Ventricular remodelling refers to changes in structure, size, and shape of the left ventricle. This architectural remodelling of the left ventricle is induced by injury (e.g., myocardial infarction), by pressure overload (e.g., systemic arterial hypertension or aortic stenosis), or by volume overload. Since ventricular remodelling affects wall stress, it has a profound impact on cardiac function and on the development of heart failure. A model of permanent ligation of the left anterior descending coronary artery in mice is used to investigate ventricular remodelling and cardiac function post-myocardial infarction. This model is fundamentally different in terms of objectives and pathophysiological relevance compared to the model of transient ligation of the left anterior descending coronary artery. In this latter model of ischemia/reperfusion injury, the initial extent of the infarct may be modulated by factors that affect myocardial salvage following reperfusion. In contrast, the infarct area at 24 hr after permanent ligation of the left anterior descending coronary artery is fixed. Cardiac function in this model will be affected by 1) the process of infarct expansion, infarct healing, and scar formation; and 2) the concomitant development of left ventricular dilatation, cardiac hypertrophy, and ventricular remodelling.
Besides the model of permanent ligation of the left anterior descending coronary artery, the technique of invasive hemodynamic measurements in mice is presented in detail.

Heart failure is the leading cause of hospitalization and a major cause of mortality. A model of permanent ligation of the left anterior descending coronary artery in mice is applied to investigate ventricular remodelling and cardiac dysfunction post-myocardial infarction. The technique of invasive hemodynamic measurements in mice is presented.

Heart failure is a syndrome in which the heart fails to pump blood at a rate commensurate with cellular oxygen requirements at rest or during stress. It ...

Developing Efficient and Precise Mouse Model for Spinal Cord Glioma

Surgical Transplantation of Tumor Cells into the Spinal Cord of Mice

Spinal cord gliomas are commonly malignant tumors of the spinal cord, leading to a high rate of disability. However, uniform treatment guidelines and comprehensive data on spinal cord gliomas remain limited due to the lack of suitable preclinical animal models. Developing a simple and reproducible animal model has become essential for advancing basic and translational research. A murine model is ideal, as the murine spinal cord shares structural similarities with the human spinal cord. This protocol describes the generation of a reproducible murine model of spinal cord glioma by directly injecting tumor cells into the intervertebral space using the spinous process of the seventh cervical vertebra as a guide. Compared to other methods, this approach is more effective and convenient, involving a smaller incision, reduced invasiveness and blood loss, faster recovery, and more stable tumor formation. This model is expected to advance the understanding of disease mechanisms, optimize surgical strategies, and support the development of therapeutic drugs for spinal cord gliomas.

The present protocol describes the development of a reproducible murine model of spinal cord glioma by injecting tumor cells into the intervertebral space, offering a more effective and less invasive approach for research and therapeutic development.

Spinal cord gliomas are commonly malignant tumors of the spinal cord, leading to a high rate of disability. However, uniform treatment guidelines and ...

Blocking Lymph Flow by Suturing Afferent Lymphatic Vessels in Mice

Lymphatic vessels are critical in maintaining tissue fluid balance and optimizing immune protection by transporting antigens, cytokines, and cells to draining lymph nodes (LNs). Interruption of lymph flow is an important method when studying the function of lymphatic vessels. The afferent lymphatic vessels from the murine footpad to the popliteal lymph nodes (pLNs) are well-defined as the only routes for lymph drainage into the pLNs. Suturing these afferent lymphatic vessels can selectively prevent lymph flow to the pLNs. This method allows for interference in lymph flow with minimal damage to the lymphatic endothelial cells in the draining pLN, the afferent lymphatic vessels, as well as other lymphatic vessels around the area. This method has been used to study how lymph impacts high endothelial venules (HEV) and chemokine expression in the LN, and how lymph flows through the adipose tissue surrounding the LN in the absence of functional lymphatic vessels. With the growing recognition of the importance of lymphatic function, this method will have broader applications to further unravel the function of lymphatic vessels in regulating the LN microenvironment and immune responses.

A protocol to block lymph flow by surgical suturing of afferent lymphatic vessels is presented.

Lymphatic vessels are critical in maintaining tissue fluid balance and optimizing immune protection by transporting antigens, cytokines, and cells to ...

Immunology and Infection

Targeting Gray Rami Communicantes in Selective Chemical Lumbar Sympathectomy

Chemical lumbar sympathectomy (CLS) is a commonly used, minimally invasive procedure for the treatment of conditions including ischemic diseases of the lower extremities, hyperhidrosis, etc. It is commonly practiced to position the puncture needle tip in front of the anterior fascia of the psoas major muscle and inject the inactivating agent around the sympathetic trunk, which is defined as conventional CLS. Although relatively rare, ureteropelvic damage is the most frequently reported complication of conventional CLS and can cause serious harm to patients. We found that injecting the inactivating agent behind the anterior fascia, which only targets gray rami communicantes, helped achieve therapeutic efficacy in&#160;vasodilation, sweat reduction, and pain relief comparable to conventional CLS, and serious complications were largely reduced. We define this procedure as selective CLS. Here, we present a protocol of selective CLS. The precise needle tract and accurate evaluation of the spreading of the contrast agent are critical to ensure that the drug is injected behind the anterior fascia of the psoas major muscle. The needle tip is at approximately one-third the dividing line of the vertebral body in the lateral view of a lumbar X-ray. The contrast is mainly confined around the needle tip and spreads outward and downward along the psoas muscle fibers. In this way, the anterior fascia provides a natural barrier for the ureteropelvic area, and the psoas major muscle provides a natural barrier for the lumbar nerve root. There are several highlights of this article, including 1) a detailed description of the selective CLS procedures, 2) an explanation of the anatomical basis for the implementation of selective CLS, and 3) an explanation of the differences between selective and conventional CLS.

We present a protocol for selective chemical lumbar sympathectomy (CLS), which inactivates only gray rami communicantes and not the sympathetic trunk. Selective CLS can help achieve therapeutic efficacy in vasodilation, sweat reduction, and pain relief that are comparable to conventional CLS, and serious complications, particularly ureteropelvic damages, can be reduced.

Chemical lumbar sympathectomy (CLS) is a commonly used, minimally invasive procedure for the treatment of conditions including ischemic diseases of the ...

A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore potential treatment options. This mouse model allows researchers to induce significant lymphedema lasting at least 8 weeks. Lymphedema is induced using a combination of fractioned radiotherapy and surgical ablation of lymphatics. This model requires that the mice get a dose of 10 Gray (Gy) radiation before and after surgery. The surgical part of the model involves ligation of three lymph vessels and extraction of two lymph nodes from the mouse hindlimb. Having access to microsurgical tools and a microscope is essential, due to the small anatomical structures of mice. The advantage of this model is that it results in statistically significant lymphedema, which provides a good basis for evaluating different treatment options. It is also a great and easily available option for microsurgical training. The limitation of this model is that the procedure can be time consuming, especially if not practiced in advance. The model results in objectively quantifiable lymphedema in mice, without causing severe morbidity and has been tested in three separate projects.

This animal model enables researchers to induce statistically significant secondary lymphedema in the hindlimb of mice, lasting at least 8 weeks. The model can be used to study the pathophysiology of lymphedema and to investigate novel treatment options.

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore ...

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. ...

A Mouse Model of Lumbar Spine Instability

Intervertebral disc degeneration (IDD) is a common pathological change leading to low back pain. Appropriate animal models are desired for understanding the pathological processes and evaluating new drugs. Here, we introduced a surgically induced lumbar spine instability (LSI) mouse model that develops IDD starting from 1 week post operation. In detail, the mouse under anesthesia was operated by low back skin incision, L3&#8211;L5 spinous processes exposure, detachment of paraspinous muscles, resection of processes and ligaments, and skin closure. L4&#8211;L5 IVDs were chosen for the observation. The LSI model develops lumbar IDD by porosity and hypertrophy in endplates at an early stage, decrease in intervertebral disc volume, shrinkage in nucleus pulposus at an intermediate stage, and bone loss in lumbar vertebrae (L5) at a later stage. The LSI mouse model has the advantages of strong operability, no requirement of special equipment, reproducibility, inexpensive, and relatively short period of IDD development. However, LSI operation is still a trauma that causes inflammation within the first week post operation. Thus, this animal model is suitable for study of lumbar IDD.

We developed a lumbar intervertebral disc degeneration mouse model by resection of L3&#8211;L5 spinous processes along with supra- and inter-spinous ligaments and detachment of paraspinous muscles.

Intervertebral disc degeneration (IDD) is a common pathological change leading to low back pain. Appropriate animal models are desired for understanding ...

Surgical Lumbar Sympathectomy in Mice

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