Q. W. was supported by NIH T32CA009685. R. J. W. was supported by NIH R01CA219534. The Memorial Sloan Kettering Cancer Center Core Facilities were supported by NIH P30CA008748.

The animal procedures described here were approved by the Institutional Animal Care and Use Committee at the Memorial Sloan Kettering Cancer Center. Eight-week-old male and female NSG mice were used here. The animals were obtained from a commercial source (see Table of Materials). The instruments are sterilized, the surgical working surface is disinfected, the animal&#39;s skin surface is disinfected, and the surgeon wears sterile gloves throughout the procedure.
1. Preparation of the mice and preoperative setup
<ol>
	<li>On the day before surgery, anesthetize the mouse with 2% isoflurane in an induction chamber (3.75 in width x 9 in depth x 3.75 in height, see Table of Materials). 
	&#8203;NOTE: A surgical plane of anesthesia is usually achieved in 3-5 min, depending on the individual animal. Assess the adequacy of anesthesia by toe pinch, and increase the isoflurane percentage as appropriate.
	<ol>
		<li>Shave the ventral aspect of the neck or use a chemical hair removal agent according to the manufacturer&#39;s instructions (see Table of Materials).</li>
	</ol>
	</li>
	<li>On the day of the surgery, anesthetize the mouse with 2% isoflurane in an induction chamber. Assess the adequacy of anesthesia by toe pinch, and increase the isoflurane percentage as appropriate.</li>
	<li>Administer 2 mg/kg of meloxicam subcutaneously for preemptive systemic analgesia. Apply topical ophthalmic ointment (see Table of Materials) to prevent ocular injuries and dryness under anesthesia.</li>
	<li>Place the mouse under a dissecting microscope on its dorsal side and provide thermal support. Maintain inhalational anesthesia with 2%-2.5% isoflurane using a precision vaporizer and nose cone. Gently secure both forelimbs with hypoallergenic tape (see Table of Materials).</li>
	<li>Clean the shaved, ventral aspect of the neck with povidone-iodine, and then wipe with 70% alcohol. Repeat this process two more times. Ensure that the surgical site is free from any loose hair. 
	NOTE:&#160;A pair of short curved forceps may also be used. Ensure to use a pair of fine or ophthalmic forceps to adequately work in this confined space. Additional pre-operative setup can be included as per the institutional guidelines.&#160;</li>
</ol>
2. Dissection
<ol>
	<li>Make a 1.5 cm midline skin incision on the ventral aspect of the neck using small scissors from approximately 2 mm below the chin to 2 mm above the sternal notch.</li>
	<li>Retract the edges of the skin laterally with forceps to expose the underlying fascia and submandibular salivary glands. Separate the skin from the underlying fascia by inserting pointed scissors under the skin on each side and spreading. Pull down the submandibular glands caudally with forceps to reveal the underlying muscles.</li>
	<li>Locate the junction of the posterior belly of the digastric muscle and omohyoid muscle (Figure 1A, black circle). The anterior jugular vein is seen running longitudinally and lateral to the omohyoid muscle. 
	NOTE: The omohyoid muscle covers the trachea longitudinally, while the digastric muscle lies transversely at the cranial aspect of the trachea (Figure 1C).
	<ol>
		<li>Insert the tip of 45&#176; angled forceps at this junction, lateral to the anterior jugular vein, to pierce and spread an opening in the overlying deep cervical fascia.</li>
	</ol>
	</li>
	<li>Keep this window created in step 2.3.1 open with the 45&#176; angled forceps. Expand this opening wider by performing spreading maneuvers with a pair of curved forceps in the other hand.</li>
</ol>
3. Identification and resection of the ganglion
<ol>
	<li>Locate the superior cervical ganglion (SCG) on the lateral wall of the revealed space. It appears as a round, pearly tissue. 
	NOTE: If the SCG is not identified, the tissues in this space need to be examined more laterally and superiorly. The SCG may be easily confused with fat, which is often present in this region. Fat has a slightly yellow tinge, while in contrast, the SCG appears pearl white.</li>
	<li>While maintaining the opening with forceps with the other hand, gently grasp the SCG with forceps, and pull it out of the opening to bring it into better view.</li>
	<li>Once the SCG is in view, grasp the lateral base of the SCG, where it is still attached to the surrounding tissues. Using the other hand, slowly and gently retract the SCG in a ventral and caudal direction.
	<ol>
		<li>Retract the SCG multiple times to gradually avulse the ganglion little by little. Keep the ganglion intact during this maneuver to ensure no residual ganglion remnants are left behind. 
		NOTE: Pull the ganglion gently, as bleeding may occur during this step. If minor bleeding occurs, use oxidized regenerated cellulose or a small strip of sterile gauze to hold pressure over the opening for 30 s to 1 min. Then, slowly lift the gauze and reassess. Repeat the process of holding pressure over the opening as necessary until the bleeding has stopped.</li>
	</ol>
	</li>
	<li>Slowly release the other forceps holding the base of the ganglion. Check for bleeding by looking for blood pooling. 
	NOTE: Slight oozing at this time is normal. Monitor and ensure there is no persistent or significant bleeding before closing and finishing the procedure. If this occurs, hold pressure over the opening, as described in step 3.3.1.</li>
	<li>Move the salivary glands back into their normal anatomic positions. Approximate and close the skin using simple interrupted 5-0 nylon sutures (see Table of Materials).</li>
	<li>Place the mouse in a clean cage by itself to allow for full recovery from the anesthesia. 
	NOTE: It may take 5-15 min for the mouse to awaken fully from the anesthesia. Do not leave the mouse unattended until it has regained sufficient consciousness to maintain sternal recumbency. Do not place the mouse in a cage with other mice until it has fully recovered. Assess the mouse for postsurgical recovery at least once every 24 h for 72 h.</li>
</ol>

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The authors have nothing to disclose.

This protocol describes a mouse model for the surgical unilateral ablation of SCG input. This technique allows for studying the effects of adrenergic innervation in various settings. In addition, the resected sympathetic ganglion can also be grown in 3D matrigel culture for in vitro experiments30.
Studies involving SCGx have mostly been performed in rats, as their larger anatomy allows for easier anatomic visualization and dissection. While SCGx in mice has been described before by Ziegler et al.26 and briefly reported in other studies21,22,27,28, the technique was based on that used in rats, in which the CCA is exposed and dissected prior to the resection of the SCG. In contrast to the rat model, the CCA in mice is smaller and thinner, rendering the dissection more difficult and, therefore, more prone to the serious complication of major bleeding from the CCA. Additionally, the exposure of the CCA requires more extensive manipulation, including the displacement of the sternocleidomastoid muscle, as well as the dissection and lateral rotation of the salivary gland26. In contrast, the present method uses the anterior jugular vein instead of the CCA as the anatomical landmark. In comparison to the CCA, the anterior jugular vein is located more superficially and extends further cranially (Figure 2A). This offers a few advantages. First, this landmark is more readily seen without the dissection and displacement of the salivary gland and sternocleidomastoid muscle, making the surgery less invasive; this protocol, therefore, only requires the salivary to be pulled down slightly (step 2.2). The minimal dissection also shortens the surgery time and the duration of anesthesia for the animal. Furthermore, by avoiding extensive dissection of the CCA, this minimizes the chances of injuring the CCA, which can lead to major and fatal bleeding in serious cases. Manipulation of the CCA is inevitable, as the SCG is located at the bifurcation of the CCA, but by approaching this region posteromedially via a piercing next to the anterior jugular vein rather than opening the fascia directly overlying the CCA, this protocol minimizes the contact with and, therefore, the risk of injury to this major artery.
Two major challenges are faced when performing this surgery. The first is the successful identification of the SCG, especially given the very small size of the anatomical landmarks and the ganglion itself in mouse models. The careful dissection and identification of the landmarks are, therefore, essential. In step 2.3, angled forceps must be inserted to pierce the deep cervical fascia at the angle of the posterior belly of the digastric and omohyoid muscles. During this step, the anterior jugular vein is usually seen running alongside the lateral edge of the omohyoid muscle and should be kept medial to the insertion point (Figure 1); this is an important landmark and will help entry into the correct space to find the SCG. If the SCG is not seen in the lateral region of this space, the tissues need to be explored more laterally and superiorly. During this dissection, the carotid sheath is visualized lateral to the field of view to avoid bleeding of the surrounding tissues and to help identify the SCG medial to this structure.
The second major challenge for this procedure is managing the risk of bleeding. There are multiple critical vascular structures adjacent to the SCG, including the CCA, the external carotid artery, and the internal jugular vein. In our experience, if bleeding occurs, it is encountered intraoperatively rather than postoperatively. Bleeding might be encountered during the step of unclamping the forceps in step 3.4. Injury to the vessels is most likely to occur when trying to peel and gently avulse the ganglion from the surrounding vessels and tissues. Active bleeding might not be seen immediately because a pair of forceps is clamped near the vessels in that region. Therefore, bleeding may be identified once the forceps are released, and it is important to inspect the area carefully after the ganglion is removed. In the rare instance of exsanguination due to a tear in a major vessel, holding pressure over the area is futile because of the rapid rate of bleeding. In this situation, the surgery must be terminated, and the mouse must be euthanized.
Given the challenges of SCG identification and possible bleeding complications, it is recommended to first practice SCG dissection and removal on cadaveric mice to become familiar with the anatomy prior to performing the experimental survival surgery.
This method may also be affected by the surgeon&#39;s handedness. The procedure is easier to perform on the same side as the surgeon&#39;s dominant hand. For example, when performing SCGx on the right side of the mouse, the surgeon&#39;s left hand would be used to grasp the base of the ganglion, and the right hand would be used to peel out the ganglion, thus meaning the surgery would require more finesse with the right hand. If bilateral SCGx is to be performed, it may be more time-consuming and require more training to perform on the surgeon&#39;s non-dominant side.
This surgical technique of SCGx in a murine model enables future experimental studies examining the effects of the sympathetic nervous system in both oncologic and physiologic settings. The mouse model has multiple advantages over other in vivo models, including the low cost, the ease of handling, and the amenability to genetic manipulation, thus enabling more powerful experimental models to be created.

Multiple studies have reported that the nerves in the tumor microenvironment play an active role in supporting tumor progression. The ablation of adrenergic sympathetic nerves has been shown to impair tumor development and dissemination in prostate and gastric cancer in vivo1,2,3, while the pharmacological blockade of adrenergic receptors inhibits tumor growth in head and neck cancer4. Sympathetic neural involvement has also been described in pancreatic, cervical, and basal cell carcinoma progression5,6,7.
Within the sympathetic nervous system, the superior cervical ganglion (SCG) is the only ganglion of the sympathetic trunk that innervates the head. The SCG regulates various physiologic functions, such as salivary secretion and circadian rhythm, and directly innervates the cervical lymph nodes8,9,10. The SCG has also been implicated in pathologic processes such as macular degeneration11 and the progression of aortic dissection12. Additionally, resection of the SCG has been reported to aggravate ischemia reperfusion-induced acute kidney injury13 and also alter the gut microbiota in rats14.
The complete ablation of the SCG in a mouse model would represent a valuable experimental technique to enable cancer and autonomic nervous system research. While many studies have utilized pharmacological adrenergic receptor blockade as an adrenergic ablation15,16,17,18,19,20, surgical resection allows for complete, unilateral adrenergic ablation while avoiding the need for repeated pharmacologic intervention and the associated side effects21,22,23.
Surgical resection of the SCG has been described in rats24, and most reports studying the effect of superior cervical ganglionectomy (SCGx) have employed the rat model. Compared to the rat model, SCGx is technically more challenging in mice due to the small size of the SCG. However, mice are comparatively easier to handle, more cost-effective, and more amenable to genetic manipulation. Garcia et al. were one of the first to report SCGx in mice, and it was found to affect insulin release25. More recently, Ziegler et al. described SCGx in mice based on the published technique described for rats24,26. This and other articles describe a method in which the common carotid artery (CCA) is first identified and dissected, and the SCG is subsequently removed from the bifurcation of the CCA21,22,27,28. In this article, a less invasive and safer technique is described in mice that avoids the dissection of the CCA, thereby minimizing the most serious complication of this procedure &#8211; bleeding from an injury to the CCA.

<table><tbody><tr><td>Anti-Tyrosine Hydroxylase Antibody</td><td>EMD Millipore</td><td>AB152</td><td></td></tr><tr><td>Artificial Tears Lubricant Ophthalmic Ointment</td><td>Akorn</td><td>59399-162-35</td><td></td></tr><tr><td>Curity 2 x 2 Inch Gauze Sponge 8 Ply, Sterile</td><td>Covidien</td><td>1806</td><td></td></tr><tr><td>Derf Needle Holder</td><td>Thomas Scientific</td><td>1177K00</td><td></td></tr><tr><td>Dissecting Microscope</td><td></td><td></td><td></td></tr><tr><td>Dumont #5/45 Forceps</td><td>Fine Science Tools</td><td>11251-35</td><td></td></tr><tr><td>Dumont #7b Forceps</td><td>Fine Science Tools</td><td>11270-20</td><td></td></tr><tr><td>ETHILON Nylon Suture</td><td>Ethicon</td><td>698H</td><td></td></tr><tr><td>Fine Scissors - ToughCut</td><td>Fine Science Tools</td><td>14058-09</td><td></td></tr><tr><td>Hypoallergenic Surgical Tape</td><td>3M Blenderm</td><td>70200419342</td><td></td></tr><tr><td>Induction Chamber, 2 Liter</td><td>VetEquip</td><td>941444</td><td></td></tr><tr><td>Isoflurane</td><td>Baxter</td><td>1001936060</td><td></td></tr><tr><td>Nair</td><td>Church &amp;amp; Dwight Co., Inc</td><td>40002957</td><td>chemical hair removing agent</td></tr><tr><td>NORADRENALINE RESEARCH ELISA</td><td>Labor Diagnostika Nord (Rocky Mountain Diagnostics)</td><td>BA E-5200</td><td></td></tr><tr><td>NSG Mouse</td><td>Jackson Laboratory</td><td>JAX:005557</td><td></td></tr><tr><td>Povidone-Iodine Swabstick</td><td>PDI</td><td>S41350</td><td></td></tr><tr><td>Webcol Alcohol Preps</td><td>Covidien</td><td>5110</td><td></td></tr></tbody></table>

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.

This protocol describes the surgical removal of the SCG in a mouse model. Figure 2 illustrates the anatomical landmarks, including the CCA, the anterior jugular vein, and the SCG. With dissection (Figure 2A), the right anterior jugular vein can be seen coursing alongside the lateral border of the trachea. As it is located deeper than the anterior jugular vein, the left CCA and its bifurcation into the internal carotid artery (ICA) and external carotid artery (ECA) are only faintly seen lateral to the vein. When examining this in NSG.B6-P0TdTomato transgenic mouse (a P0-Cre TdTomato mouse where the Schwann cells are fluorescent red, unpublished work) with red fluorescent neurons under a fluorescent microscope, the fluorescent vagus nerve can be seen coursing laterally to the CCA, and the fluorescent SCG can be seen at the bifurcation of the CCA, lateral to the anterior jugular vein (Figure 2B).
After the resection of the SCG in a normal mouse and a transgenic mouse, the resected tissue was confirmed by its red fluorescence compared to the non-fluorescent SCG control (Figure 3A) and immunofluorescent staining for tyrosine hydroxylase (TH), a marker for adrenergic nerves13,29 (Figure 3B).
If the procedure is performed correctly, the mouse develops ipsilateral Horner&#39;s syndrome immediately after surgery upon regaining full consciousness24. Ptosis, the drooping of the eyelid, was observed, which is a sign of Horner&#39;s syndrome (Figure 4B).
The submandibular salivary gland is one of the tissues innervated by the SCG. To validate successful SCGx, immunofluorescence staining for TH was performed on the right submandibular salivary gland following right SCGx and confirmed the successful ablation of the adrenergic signaling with absent TH nerve staining (right side of the dotted line, Figure 5A). In contrast, the left control submandibular gland (no SCGx) maintained its adrenergic input and intact TH nerve staining (left side of the dotted line, Figure 5A). These findings were confirmed by quantification (Figure 5B). ELISA quantification of norepinephrine13,30,31 in these tissues further confirmed a significant reduction in norepinephrine expression in the submandibular gland on the side of SCGx in contrast to the control sham surgery side (Figure 6). The quantification for both was analyzed by an unpaired, two-tailed Student&#39;s t-test.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64527/64527fig01.jpg" /> 
Figure 1: The left anterior jugular vein serves as an anatomical landmark. (A) The left anterior jugular vein (blue arrow) can be seen coursing longitudinally and alongside the lateral edge of the omohyoid muscle. When piercing the deep cervical fascia between the angle of the posterior belly of the digastric and omohyoid muscles, the piercing should also be lateral to the anterior jugular vein (black circle). (B) When the deep cervical fascia is slightly stretched, the vagus nerve (white arrow) and the common carotid artery with its bifurcation (red arrow) can also be seen. (C) Stylized illustration of (B). Scale bar = 100 &#181;m. Abbreviation: M = muscle. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig01large.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/64527/64527fig2v4.jpg" /> 
Figure 2: The SCG and its relation to anatomical landmarks in a transgenic mouse with fluorescent neurons. (A)&#160;Dissection in a transgenic mouse with red fluorescent neurons, illustrating the right common carotid artery (red arrow pointing to its bifurcation) and the anterior jugular vein. The common carotid artery bifurcates into the external carotid artery (ECA) and internal carotid artery (ICA). The yellow arrow points to the vagus nerve running laterally to the common carotid artery. The distance between the common carotid artery and the vagus nerve appears wider here as the mouse&#39;s head is turned to capture all the structures in this image. (B)&#160;The same dissection examined with fluorescent imaging. The vagus nerve (yellow arrow) has red fluorescence and is again seen running laterally to the common carotid artery (red arrow pointing to its bifurcation). The fluorescent SCG is located at the bifurcation of the carotid artery (yellow arrowhead). The anterior jugular vein (blue arrow) runs medial to the common carotid artery. The deep cervical fascia overlying these structures can be seen with its glistening reflection. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig2v4large.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/64527/64527fig03.jpg" /> 
Figure 3: Microscopic images of the resected ganglion. (A) Resected superior cervical ganglia under fluorescent microscopy. The left shows the SCG resected from a normal mouse, serving as a non-fluorescent control. The right shows a fluorescent SCG resected from a transgenic mouse with red fluorescent neurons. Scale bar = 500 &#181;m. (B) Immunofluorescent staining for tyrosine hydroxylase (TH), a marker for adrenergic nerves, in the resected red fluorescent ganglion (P0). Scale bar = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64527/64527fig04.jpg" /> 
Figure 4: Development of Horner&#39;s syndrome following SCGx. (A) A normal mouse before SCGx. (B) The development of ptosis (black arrow), the drooping of the eyelid, following ipsilateral SCGx, which is a sign of Horner&#39;s syndrome. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64527/64527fig05v2.jpg" /> 
Figure 5: Immunofluorescence and corresponding H&#38;E staining for the adrenergic marker in the target tissue after SCGx versus sham surgery. (A)&#160;Left, immunofluorescence staining for tyrosine hydroxylase (TH) in the submandibular salivary gland following SCGx or sham surgery. Right, the corresponding H&#38;E staining of the same tissue. Scale bar = 200 &#181;m. (B)&#160;Quantification of the TH staining. Data represent mean &#177; SEM. Statistical analysis by unpaired, two-tailed Student&#39;s t-test, p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig05largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64527/64527fig06.jpg" /> 
Figure 6: ELISA quantification of norepinephrine in the salivary gland after SCGx versus sham surgery. Data represent mean &#177; SEM. Statistical analysis by unpaired, two-tailed Student&#39;s t-test, p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/64527/64527fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Surgical Technique for Superior Cervical Ganglionectomy in a Murine Model at JoVE.com

Surgical Technique for Superior Cervical Ganglionectomy in a Murine Model

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 ...

surgical-technique-for-superior-cervical-ganglionectomy-murine

Nanyang Technological University

Research

JoVE Journal

Cancer Research

4.3K Views.  Memorial Sloan Kettering Cancer Center. Het sympathische zenuwstelsel speelt een belangrijke rol bij de progressie van kanker en andere ziekten. Chirurgische sympathectomie is een waardevolle methode om deze processen te bestuderen door volledige unilaterale adrenerge ablatie te induceren. Deze techniek vermijdt de bijwerkingen geassocieerd met farmacologische remming van het sympathische zenuwstelsel.Bovendien is het minder invasief dan eerder beschreven methoden omdat het dissectie van de halsslagader voorkomt. Bij het uit...

Video: Surgical Technique for Superior Cervical Ganglionectomy in a Murine Model

Murine Superficial Lymph Node Surgery

In the field of immunology, to understand the progression of an immune response against a vaccine, an infection or a tumour, the response is often followed over time. Similarly, the study of lymphocyte homeostasis requires time course experiments. Performing these studies within the same mouse is ideal to reduce the experimental variability as well as the number of mice used. Blood withdrawal allows performance of time course experiments, but it only gives information about circulating lymphocytes and provides a limited number of cells1-4. Since lymphocytes circulating through the body and residing in the lymph nodes have different properties, it is important to examine both locations. The sequential removal of lymph nodes by surgery provides a unique opportunity to follow an immune response or immune cell expansion in the same mouse over time. Furthermore, this technique yields between 1-2x106 cells per lymph node which is sufficient to perform phenotypic characterization and/or functional assays. Sequential lymph node surgery or lymphadenectomy has been successfully used by us and others5-11. Here, we describe how the brachial and inguinal lymph nodes can be removed by making a small incision in the skin of an anesthetised mouse. Since the surgery is superficial and done rapidly, the mouse recovers very quickly, heals well and does not experience excessive pain. Every second day, it is possible to harvest one or two lymph nodes allowing for time course experiments. This technique is thus suitable to study the characteristics of lymph node-residing lymphocytes over time. This approach is suitable to various experimental designs and we believe that many laboratories would benefit from performing sequential lymph node surgeries.

To follow the progression of an immune response over time within the same mouse, lymph nodes can be sequentially removed by surgery. Here, we describe how this technique can be performed.

In the field of immunology, to understand the progression of an immune response against a vaccine, an infection or a tumour, the response is often ...

Immunology and Infection

Protocol for Culturing Sympathetic Neurons from Rat Superior Cervical Ganglia (SCG)

The superior cervical ganglia (SCG) in rats are small, glossy, almond-shaped structures that contain sympathetic neurons.  These neurons provide sympathetic innervations for the head and neck regions and they constitute a well-characterized and relatively homogeneous population (4).  Sympathetic neurons are dependent on nerve growth factor (NGF) for survival, differentiation and axonal growth and the wide-spread availability of NGF facilitates their culture and experimental manipulation (2, 3, 6).  For these reasons, cultured sympathetic neurons have been used in a wide variety of studies including neuronal development and differentiation, mechanisms of programmed and pathological cell death, and signal transduction (1, 2, 5, and 6).  Dissecting out the SCG from newborn rats and culturing sympathetic neurons is not very complicated and can be mastered fairly quickly.  In this article, we will describe in detail how to dissect out the SCG from newborn rat pups and to use them to establish cultures of sympathetic neurons.  The article will also describe the preparatory steps and the various reagents and equipment that are needed to achieve this.

Protocol for Culturing Sympathetic Neurons from Rat Superior Cervical Ganglia SCG

This is a protocol describing how to isolate and culture primary sympathetic neurons from superior cervical ganglia (SCG) of newborn rat pups.

The superior cervical ganglia (SCG) in rats are small, glossy, almond-shaped structures that contain sympathetic neurons.  These neurons provide ...

Biology

A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers that cause LMD are breast and lung cancers and melanoma. Patients diagnosed with LMD have a very poor prognosis and generally survive for only a few weeks or months. One possible reason for the lack of efficacy of systemic therapy against LMD is the failure to achieve therapeutically effective concentrations of drug in the CSF because of an intact and relatively impermeable blood-brain barrier (BBB) or blood-CSF barrier across the choroid plexus. Therefore, directly administering drugs intrathecally or intraventricularly may overcome these barriers. This group has developed a model that allows for the effective delivery of therapeutics (i.e., drugs, antibodies, and cellular therapies) chronically and the repeated sampling of CSF to determine drug concentrations and target modulation in the CSF (when the tumor microenvironment is targeted in mice). The model is the murine equivalent of a magnetic resonance imaging-compatible Ommaya reservoir, which is used clinically. This model, which is affixed to the skull, has been designated as the &#34;Murine Ommaya.&#34; As a therapeutic proof of concept, human epidermal growth factor receptor 2 antibodies (clone 7.16.4) were delivered into the CSF via the Murine Ommaya to treat mice with LMD from human epidermal growth factor receptor 2-positive breast cancer. The Murine Ommaya increases the efficiency of drug delivery using a miniature access port and prevents the wastage of excess drug; it does not interfere with CSF sampling for molecular and immunological studies. The Murine Ommaya is useful for testing novel therapeutics in experimental models of LMD.

Here, we describe a murine xenograft model that functionally resembles an Ommaya reservoir in patients. We developed the Murine Ommaya to study novel therapeutics for the universally fatal leptomeningeal disease.

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers ...

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

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

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

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

Medicine

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These neurotropic cancer cells have a unique ability to migrate unidirectionally along nerves towards the central nervous system (CNS). The dorsal root ganglia (DRG)/cancer cell model is a three dimensional (3D) in vitro model frequently used for studying the interaction between neural stroma and cancer cells. In this model, mouse or human cancer cell lines are grown in ECM adjacent to preparations of freshly dissociated cultured DRG. In this article, the DRG isolation protocol from mice, and implantation in petri dishes for co-culturing with pancreatic cancer cells are demonstrated. Five days after implantation, the cancer cells made contact with the DRG neurites. Later, these cells formed bridgeheads to facilitate more extensive polarized, neurotropic migration of cancer cells.

In Vitro Modeling of Cancerous Neural Invasion: The Dorsal Root Ganglion Model

This video article shows the use of the dorsal root ganglia (DRG)/cancer cell model in pancreatic ductal adenocarcinoma.

One way that solid tumors disseminate is through neural invasion. This route is well-known in cancers of the head and neck, prostate, and pancreas. These ...

Neuroscience

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Stereotaxic surgery to target brain sites in mice is commonly guided by skull landmarks. Access is then obtained via burr holes drilled through the skull. This standard approach can be challenging for targets in the caudal brainstem and upper cervical cord due to specific anatomical challenges as these sites are remote from skull landmarks, leading to imprecision. Here we outline an alternative stereotaxic approach via the cisterna magna that has been used to target discrete regions of interest in the caudal brainstem and upper cervical cord. The cisterna magna extends from the occipital bone to the atlas (i.e., the second vertebral bone), is filled with cerebrospinal fluid, and is covered by dura mater. This approach provides a reproducible route of access to select central nervous system (CNS) structures that are otherwise hard to reach due to anatomical barriers. Furthermore, it allows for direct visualization of brainstem landmarks in close proximity to the target sites, increasing accuracy when delivering small injection volumes to restricted regions of interest in the caudal brainstem and upper cervical cord. Finally, this approach provides an opportunity to avoid the cerebellum, which can be important for motor and sensorimotor studies.

Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Stereotaxic surgery to target brain sites in mice commonly involves access through the skull bones and is guided by skull landmarks. Here we outline an alternative stereotaxic approach to target the caudal brainstem and upper cervical spinal cord via the cisterna magna that relies on direct visualization of brainstem landmarks.

Stereotaxic surgery to target brain sites in mice is commonly guided by skull landmarks. Access is then obtained via burr holes drilled through the skull. ...

Location, Dissection, and Analysis of the Murine Stellate Ganglion

The autonomic nervous system is a substantial driver of cardiac electrophysiology. Especially&#160;the role of its sympathetic branch is an ongoing matter of investigation in the pathophysiology of ventricular arrhythmias (VA). Neurons in the stellate ganglia (SG)&#160;&#8211;&#160;bilateral star-shaped structures of the sympathetic chain &#8211;&#160;are an important component of the sympathetic infrastructure. The SG are a recognized target for&#160;treatment via cardiac sympathetic denervation in patients with therapy-refractory VA. While neuronal remodeling and glial activation in the SG have been described in patients with VA, the underlying cellular and molecular processes that potentially precede the onset of arrhythmia are only insufficiently understood and should be elucidated to improve autonomic modulation. Mouse models allow us to study sympathetic neuronal remodeling, but identification of the murine SG is challenging for the inexperienced investigator. Thus, in-depth cellular and molecular biological studies of the murine SG are lacking for many common cardiac diseases. Here, we describe a basic repertoire for dissecting and studying the SG in adult mice for analyses at RNA level (RNA isolation for gene expression analyses, in situ hybridization), protein level (immunofluorescent whole mount staining), and cellular level (basic morphology, cell size measurement). We present potential solutions to overcome challenges in the preparation technique, and how to improve staining via quenching of autofluorescence. This allows for the visualization of neurons as well as glial cells via established markers in order to determine cell composition and remodeling processes. The methods presented here allow characterizing the SG to gain further information on autonomic dysfunction in mice prone to VA and can be complemented by additional techniques investigating neuronal and glial components of the autonomic nervous system in the heart.

Pathophysiological changes in the cardiac autonomic nervous system, especially in its sympathetic branch, contribute to the onset and maintenance of ventricular arrhythmias. In the present protocol, we show how to characterize murine stellate ganglia to improve the understanding of the underlying molecular and cellular processes.

The autonomic nervous system is a substantial driver of cardiac electrophysiology. Especially&#160;the role of its sympathetic branch is an ongoing matter ...

Low-input Nucleus Isolation and Multiplexing with Barcoded Antibodies of Mouse Sympathetic Ganglia for Single-nucleus RNA Sequencing

The cardiac autonomic nervous system is crucial in controlling cardiac function, such as heart rate and cardiac contractility, and is divided into sympathetic and parasympathetic branches. Normally, there is a balance between these two branches to maintain homeostasis. However, cardiac disease states such as myocardial infarction, heart failure, and hypertension can induce the remodeling of cells involved in cardiac innervation, which is associated with an adverse clinical outcome.
Although there are vast amounts of data for the histological structure and function of the cardiac autonomic nervous system, its molecular biological architecture in health and disease is still enigmatic in many aspects. Novel technologies such as single-cell RNA sequencing (scRNA-seq) hold promise for the genetic characterization of tissues at single-cell resolution. However, the relatively large size of neurons may impede the standardized use of these techniques. Here, this protocol exploits droplet-based single-nucleus RNA sequencing (snRNA-seq), a method to characterize the biological architecture of cardiac sympathetic neurons in health and disease. A stepwise approach is demonstrated to perform snRNA-seq of the bilateral superior cervical (SCG) and stellate ganglia (StG) dissected from adult mice.
This method enables long-term sample preservation, maintaining an adequate RNA quality when samples from multiple individuals/experiments cannot be collected all at once within a short period of time. Barcoding the nuclei with hashtag oligos (HTOs) enables demultiplexing and the trace-back of distinct ganglionic samples post sequencing. Subsequent analyses revealed successful nuclei capture of neuronal, satellite glial, and endothelial cells of the sympathetic ganglia, as validated by snRNA-seq. In summary, this protocol provides a stepwise approach for snRNA-seq of sympathetic extrinsic cardiac ganglia, a method that has the potential for broader application in studies of the innervation of other organs and tissues.

This protocol describes the detailed, low-input sample preparation for single-nucleus sequencing, including the dissection of mouse superior cervical and stellate ganglia, cell dissociation, cryopreservation, nucleus isolation, and hashtag barcoding.

The cardiac autonomic nervous system is crucial in controlling cardiac function, such as heart rate and cardiac contractility, and is divided into ...

Targeting Mouse Sympathetic Neurons: A Lumbar Sympathectomy Approach

Surgical Lumbar Sympathectomy in Mice

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

Author Spotlight: Exploring Plasticity of Sympathetic Neurons

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

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

Surgical Technique for Superior Cervical Ganglionectomy in a Murine Model

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