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 the intestines. To ensure adequate visualization, the descending colon must be visible. The intestinal diversion outside of the abdominal cavity must include the cecum. Additionally, the bladder may be full and enlarged, which can obstruct the view of the descending colon and, subsequently, the descending abdominal aorta and inferior vena cava. Ideally, the bladder is manually expressed once the mouse is under anesthesia prior to making the initial skin incision. However, should the bladder not be expressed and the intraoperative view be blocked, gentle expression with 2 sterile cotton-tipped applicators is possible. Take care not to damage the ureters, which are attached to the posterior aspect of the bladder. Once the abdominal aorta and inferior vena cava are identified posterior to the descending colon, adequate deflection and stabilization of the vessels is critical for visualization of the sympathetic ganglia. The fascia is relatively tough, and if the suture is placed solely through the layer of fascia and neuronal connections and not through the vessel walls, the vessels should be pulled at least 1 mm from the midline. The fascia overlying the sympathetic ganglia can be gently dissected away with tweezers. Should any small vessel break during the procedure, apply pressure with a sterile cotton-tipped applicator for at least 10 s to ensure adequate hemostasis before closure. If the large abdominal vessels are punctured during the surgery, the mouse should be quickly euthanized.
Although this method allows for direct visualization and removal of the sympathetic ganglia, thus allowing for more specific targeting of these postganglionic sympathetic neurons compared to chemical sympathectomies23, there are some limitations. This surgery results in the removal of the L2-L5 lumbar sympathetic ganglia; however, due to the visual obstruction caused by vital urogenital organs, the lowest ganglia, such as the L4-L5 level ganglia, are more difficult to visualize and remove during this procedure. The majority of the neurons innervating the lower limbs are located in the large L2 ganglia. The L5 ganglia are extremely small and contain fewer neurons, as seen in previous fluorescent retrograde tracing experiments38; however, failure to remove these neurons may distort the analysis of results from the most distal structures, such as the feet. To ensure better results when studying the distal targets in the feet, it would be advisable to check the extracted tissue to count the number of ganglia that have been removed. This can be done via a fluorescent reporter mouse in which the postganglionic sympathetic neurons can easily be visualized or via a fluorescent retrograde tracer injected into the distal target prior to the surgery. Some fluorescent reporter mice for this purpose include the ThCre:mTmG (see Table of Materials), which has extremely sparse labeling of tyrosine hydroxylase-positive neurons39, the Phox2bCre:tdTomato (see Table of Materials), which has moderate labeling of postganglionic sympathetic neurons40,41, and the ThCre:tdTomato (see Table of Materials), which has extensive labeling of the postganglionic sympathetic neurons42.&#160;This is not an exhaustive list of available lines.
Additionally, due to the large incision required for this surgical procedure, mice may need a longer recovery period before further experiments can be performed. Mice should be monitored for at least 3 days for proper defecation, urination, and feeding. Furthermore, improper hemostasis can lead to unaddressed intraabdominal bleeding, which can result in mouse fatality at unexpected time points. Therefore, ensuring that hemostasis is achieved prior to surgical closure is critical for the success of future experiments. The sympathetic neurons can also be targeted chemically, as aforementioned24.
The use of surgical sympathectomies will allow for a multitude of studies that aim to investigate the role of postganglionic sympathetic innervation in distal targets, such as the role of sympathetic innervation at the neuromuscular junction12. Additionally, fluorescent retrograde tracing techniques can be employed to quantify sympathetic reinnervation of distal tissues as&#160;this protocol can be adapted to extract&#160;the ganglia&#160;en bloc from paraformaldehyde-fixed mice38. In the context of peripheral nerve injuries, this surgical &#34;knock-out&#34; model will allow for better characterization of expected sympathetic functional recovery in previously innervated tissues.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>Phox2bCre:tdTomato mutant mice</td>
			<td>Jackson Laboratory&#160;</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&#160;</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>ThCre:mTmG 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>ThCre:tdTomato 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 ganglia may be obstructed by the urogenital organs. Previous retrograde tracing studies have shown that the majority of neurons in the L2-L5 ganglia are located in the L2 and L3 ganglia. Although only female, age-matched mice were used to achieve the representative results, this surgical procedure has been reproduced many times in both male and female mice with no significant anatomical differences observed in terms of the locations of the testicular or ovarian artery and the sympathetic ganglia.
Upon removal of the ganglia, the sweating response in response to pilocarpine rapidly decreases, with significant differences seen by postoperative day 7 (Figure 2). Postoperative day 7 was chosen due to the exponential decrease in sympathetically-mediated sweating activity within 7 days after various sciatic nerve injuries in rats34. Because axon degeneration can take up to 14 days to complete and due to potential difficulty visualizing the L4 and L5 ganglia intraoperatively (bladder and genital organs may be obstructing the view), the sweating response may not be completely ablated35,36.
The tyrosine hydroxylase-positive (TH+) axon density was significantly decreased by postoperative day 21 with no change in the neurofilament-heavy chain (NF-H) density (Figure 3). TH+ axons are unlikely to be completely depleted due to the presence of TH+ sensory axons innervating low threshold mechanoreceptors and potential difficulties visualizing the full lumbar sympathetic chain37.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66821/66821fig01v2.jpg" /> 
Figure 1: Intraoperative view of the lumbar sympathetic ganglia. (A) Schematic with expected landmarks. (B) Representative intraoperative image with labeled landmarks. The lumbar sympathetic ganglia are located behind the descending abdominal aorta and inferior vena cava (deflected). The ganglia are pulled to the mouse&#39;s right with the deflection of the abdominal vessels and are located on the right psoas, traversing inferiorly and superiorly. The bilateral L2 ganglia can be visualized between the left renal artery and the left testicular or ovarian artery, and the L3 ganglia are seen inferiorly to the left testicular or ovarian artery. Scale bar: 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66821/66821fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66821/66821fig02.jpg" /> 
Figure 2: Surgical lumbar sympathectomy reduces the number of sweating spots on the hind paw. (A) Schematic of footpad locations on the plantar surface of the mouse&#39;s hind paw. (B) A representative image of the sweating response 8 min after the injection of pilocarpine in an intact animal. (B&#39;) Zoomed-in image with white arrows indicating a select number of sweating spots. (C) A representative image of the sweating response 8 min after the injection of pilocarpine in a sympathectomized animal&#160;on postoperative day 7. (C&#39;) Zoomed-in image with white arrows indicating a select number of sweating spots. (D) The total number of sweating spots on all six footpads of the hind paw in intact versus sympathectomized mice. Data is shown with the line representing the median. Mann-Whitney test. *p &#60; 0.05. Scale bars: 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66821/66821fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66821/66821fig03.jpg" /> 
Figure 3: Surgical lumbar sympathectomy (Sympx) reduces the number of sympathetic axons in the sciatic nerve. (A) Tyrosine hydroxylase (TH)-positive axons per &#181;m of the sciatic nerve (width of the nerve) on postoperative day 21. Representative nerve sections of TH staining in an intact (B) and sympathectomized (B&#39;) mouse. (C) Neurofilament-heavy chain (NF-H)-positive axons per &#181;m of the sciatic nerve. Representative nerve section of NF-H staining in an intact (D) and sympathectomized (D&#39;) mouse. Magenta dashed lines represent possible randomly placed vertical lines that span the width of the nerve section used to calculate the axon densities in the nerve. Magenta arrows indicate a select number of axons that cross the randomly placed vertical lines. Data is shown as mean &#177; SEM. Unpaired t-test. **p &#60; 0.01. Scale bars: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66821/66821fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

Nanyang Technological University

Research

JoVE Journal

Neuroscience

1.7K Views.  Emory University School of Medicine. 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.

Video: Author Spotlight: Exploring Plasticity of Sympathetic Neurons

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 ...

Video-oculography in Mice

Eye movements are very important in order to track an object or to stabilize an image on the retina during movement. Animals without a fovea, such as the mouse, have a limited capacity to lock their eyes onto a target. In contrast to these target directed eye movements, compensatory ocular eye movements are easily elicited in afoveate animals1,2,3,4. Compensatory ocular movements are generated by processing vestibular and optokinetic information into a command signal that will drive the eye muscles. The processing of the vestibular and optokinetic information can be investigated separately and together, allowing the specification of a deficit in the oculomotor system. The oculomotor system can be tested by evoking an optokinetic reflex (OKR), vestibulo-ocular reflex (VOR) or a visually-enhanced vestibulo-ocular reflex (VVOR). The OKR is a reflex movement that compensates for "full-field" image movements on the retina, whereas the VOR is a reflex eye movement that compensates head movements. The VVOR is a reflex eye movement that uses both vestibular as well as optokinetic information to make the appropriate compensation. The cerebellum monitors and is able to adjust these compensatory eye movements. Therefore, oculography is a very powerful tool to investigate brain-behavior relationship under normal as well as under pathological conditions (f.e. of vestibular, ocular and/or cerebellar origin).
Testing the oculomotor system, as a behavioral paradigm, is interesting for several reasons. First, the oculomotor system is a well understood neural system5. Second, the oculomotor system is relative simple6; the amount of possible eye movement is limited by its ball-in-socket architecture ("single joint") and the three pairs of extra-ocular muscles7. Third, the behavioral output and sensory input can easily be measured, which makes this a highly accessible system for quantitative analysis8. Many behavioral tests lack this high level of quantitative power. And finally, both performance as well as plasticity of the oculomotor system can be tested, allowing research on learning and memory processes9.
Genetically modified mice are nowadays widely available and they form an important source for the exploration of brain functions at various levels10. In addition, they can be used as models to mimic human diseases. Applying oculography on normal, pharmacologically-treated or genetically modified mice is a powerful research tool to explore the underlying physiology of motor behaviors under normal and pathological conditions. Here, we describe how to measure video-oculography in mice8.

Video-oculography is a very quantitative method to investigate ocular motor performance as well as motor learning. Here, we describe how to measure video-oculography in mice. Applying this technique on normal, pharmacologically-treated or genetically modified mice is a powerful research tool to explore the underlying physiology of motor behaviors.

Eye movements are very important in order to track an object or to stabilize an image on the retina during movement. Animals without a fovea, such as the ...

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

The spinal motoneuron has long been a good model system for studying neural function because it is a neuron of the central nervous system with the unique properties of (1) having readily identifiable targets (the muscle fibers) and therefore having a very well-known function (to control muscle contraction); (2) being the convergent target of many spinal and descending networks, hence the name of "final common pathway"; and (3) having a large soma which makes it possible to penetrate them with sharp intracellular electrodes. Furthermore, when studied in vivo, it is possible to record simultaneously the electrical activity of the motoneurons and the force developed by their muscle targets. Performing intracellular recordings of motoneurons in vivo therefore put the experimentalist in the unique position of being able to study, at the same time, all the compartments of the "motor unit" (the name given to the motoneuron, its axon, and the muscle fibers it innervates1): the inputs impinging on the motoneuron, the electrophysiological properties of the motoneuron, and the impact of these properties on the physiological function of the motoneurons, i.e. the force produced by its motor unit. However, this approach is very challenging because the preparation cannot be paralyzed and thus the mechanical stability for the intracellular recording is reduced. Thus, this kind of experiments has only been achieved in cats and in rats. However, the study of spinal motor systems could make a formidable leap if it was possible to perform similar experiments in normal and genetically modified mice.
For technical reasons, the study of the spinal networks in mice has mostly been limited to neonatal in vitro preparations, where the motoneurons and the spinal networks are immature, the motoneurons are separated from their targets, and when studied in slices, the motoneurons are separated from most of their inputs. Until recently, only a few groups had managed to perform intracellular recordings of motoneurons in vivo2-4 , including our team who published a new preparation which allowed us to obtain very stable recordings of motoneurons in vivo in adult mice5,6. However, these recordings were obtained in paralyzed animals, i.e. without the possibility to record the force output of these motoneurons. Here we present an extension of this original preparation in which we were able to obtain simultaneous recordings of the electrophysiological properties of the motoneurons and of the force developed by their motor unit. This is an important achievement, as it allows us to identify the different types of motoneurons based on their force profile, and thereby revealing their function. Coupled with genetic models disturbing spinal segmental circuitry7-9, or reproducting human disease10,11, we expect this technique to be an essential tool for the study of spinal motor system.

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

This new method permits the simultaneous intracellular recording of a single adult mouse motoneuron and the measurement of the force produced by its muscle fibers. The combined investigation of the electrical and mechanical properties of motor units in normal and genetically modified animals is a breakthrough for the study of the neuromuscular system.

The  spinal motoneuron has long been a good model system for studying neural  function because it is a neuron of the central nervous system with the ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated thinning of the skull. Cranial window surgery is typically performed with a high speed rotary drill, and many investigators find it challenging to prevent the drill from damaging the delicate dura and blood vessels. Extensive training and practice is required to remove the bone without damage to underlying tissue and thus cranial window surgery can be difficult, time consuming, and produce tissue damage. Piezoelectric surgery, which is extensively used for maxillofacial and dental surgery, utilizes ultrasonic vibrations to remove bone without damaging soft tissues. We have developed a method applying piezoelectric surgery to improve cranial window surgery in mice in preparation for multiphoton imaging. Comparisons within our lab find that the method requires less surgery time and has a lower average rate of complications due to dural bleeding than cranial window surgery with a rotary drill.

Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice

Piezoelectric surgery has led to improvements in human maxillofacial and dental surgery. We have developed a protocol to optimize piezoelectric surgery for cranial window surgery in mice.

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated ...

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive targets for injection of novel therapeutics aimed at treating chronic pain. In small animal models, laminectomy has been used to facilitate DRG injection because it involves surgical removal of the vertebral bone surrounding each DRG. We demonstrate a technique for intraganglionic injection of lumbar DRG in a large animal species, namely, swine. Laminotomy is performed to allow direct access to DRG using standard neurosurgical techniques, instruments, and materials. Compared with more extensive bone removal via laminectomy, we implement laminotomy to conserve spinal anatomy while achieving sufficient DRG access. Intraoperative progress of DRG injection is monitored using a non-toxic dye. Following euthanasia on post-operative day 21, the success of injection is determined by histology for intraganglionic distribution of 4&#39;,6-diamidino-2-phenylindole (DAPI). We inject a biologically inactive solution to demonstrate the protocol. This method could be applied in future preclinical studies to target therapeutic solutions to DRG. Our methodology should facilitate testing the translatability of intraganglionic small animal paradigms in a large animal species. Additionally, this protocol may serve as a key resource for those planning preclinical studies of DRG injection in swine.

We describe a method for laminotomy in swine that provides access to lumbar dorsal root ganglia (DRG) for intraganglionic injection. Injection progress is monitored intraoperatively and histologically confirmed up to 21 days after surgery. This protocol could be used for future preclinical studies involving DRG injection.

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive ...

Assessing Pupil-linked Changes in Locus Coeruleus-mediated Arousal Elicited by Trigeminal Stimulation

Current scientific literature provides evidence that trigeminal sensorimotor activity associated with chewing may affect arousal, attention, and cognitive performance. These effects may be due to widespread connections of the trigeminal system to the ascending reticular activating system (ARAS), to which noradrenergic neurons of the locus coeruleus (LC) belongs. LC neurons contain projections to the whole brain, and it is known that their discharge co-varies with pupil size. LC activation is necessary for eliciting task-related mydriasis. If chewing effects on cognitive performance are mediated by the LC, it is reasonable to expect that changes in cognitive performance are correlated to changes in task-related mydriasis. Two novel protocols are presented here to verify this hypothesis and document that chewing effects are not attributable to aspecific motor activation. In both protocols, performance and pupil size changes observed during specific tasks are recorded before, soon after, and half an hour following a 2 min period of either: a) no activity, b) rhythmic, bilateral handgrip, c) bilateral chewing of soft pellet, and d) bilateral chewing of hard pellet. The first protocol measures level of performance in spotting target numbers displayed within numeric matrices. Since pupil size recordings are recorded by an appropriate pupillometer that impedes vision to ensure constant illumination levels, task-related mydriasis is evaluated during a haptic task. Results from this protocol reveal that 1) chewing-induced changes in performance and task-related mydriasis are correlated and 2) neither performance nor mydriasis are enhanced by handgrip. In the second protocol, use of a wearable pupillometer allows measurement of pupil size changes and performance during the same task, thus allowing even stronger evidence to be obtained regarding LC involvement in the trigeminal effects on cognitive activity. Both protocols have been run in the historical office of Prof. Giuseppe Moruzzi, the discoverer of ARAS, at the University of Pisa.

To verify whether trigeminal effects on cognitive performance involve locus coeruleus activity, two protocols are presented that aim to evaluate possible correlations between the performance and task-related pupil size changes induced by chewing. These protocols may be applied to conditions in which locus coeruleus contribution is suspected.

Current scientific literature provides evidence that trigeminal sensorimotor activity associated with chewing may affect arousal, attention, and cognitive ...

Quantitative Measurement of Intrathecally Synthesized Proteins in Mice

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the CSF protein composition delivers crucial information in basic neuroscience research as well as neurological diseases. One caveat is that proteins measured in CSF may derive from both intrathecal synthesis and transudation from serum, and protein analysis of CSF can only determine the sum of these two components. To discriminate between protein transudation from the blood and intrathecally produced proteins in animal models as well as in humans, CSF protein profiling measurements using conventional protein analysis tools must include the calculation of the albumin CSF/serum quotient (Qalbumin), a marker of the integrity of the blood-brain interface (BBI), and the protein index (Qprotein/Qalbumin), an estimate of intrathecal protein synthesis. This protocol illustrates the entire procedure, from CSF and blood collection to quotients and indices calculations, for the quantitative measurement of intrathecal protein synthesis and BBI impairment in mouse models of neurological disorders.

Elevated spinal fluid protein levels can either be the result of diffusion of plasma protein across an altered blood-brain barrier or intrathecal synthesis. An optimized testing protocol is presented in this article that helps to discriminate both cases and provides quantitative measurements of intrathecally synthesized proteins.

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Surgical Lumbar Sympathectomy in Mice

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

Video-oculography in Mice

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Assessing Pupil-linked Changes in Locus Coeruleus-mediated Arousal Elicited by Trigeminal Stimulation

Quantitative Measurement of Intrathecally Synthesized Proteins in Mice

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice