Surgical Lumbar Sympathectomy in Mice

All experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Emory University (under the IACUC protocol number PROTO201700371). 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

1. Autoclave the surgical tools: 1 pair of sharp scissors, 2 fine-tipped tweezers, 1 needle driver.
2. Warm a heating pad to 37 °C and place it under the surgical tabletop.
3. Spray the surgical area, including the anesthesia induction chamber, with disinfectant and wipe it down thoroughly with paper towels.
4. Ensure proper gowning and hand washing prior to handling the mouse.
5. Feed the mouse with meloxicam at 5 mg/kg body weight, and allow plenty of time for the mouse to completely swallow the medicine to avoid aspiration during induction.
6. Induce the mouse under anesthesia with 3% isoflurane in 1 L/min of oxygen (following institutionally approved protocols).
7. 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.
8. Once the mouse is properly fitted in the nose cone, adjust the isoflurane to 2% for maintenance of anesthesia.
9. Place eye gel on the mouse's eyes to prevent keratoconjunctivitis sicca (dry eye).
10. Ensure the mouse is properly secured with surgical tape to the surgical tabletop.
11. Monitor continuously for ease of breathing and adequate breathing rate.
12. Shave the fur from the mouse's abdomen from the level of the genitals to below the ribs.
13. 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 a total of 3 times.
14. 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.

2. Incisions

1. 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.
2. 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.
3. Retract the abdominal muscles and skin with 5-0 sutures laterally to visualize the next steps properly.

3. Identification of the L2-L5 lumbar sympathetic ganglia

1. 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 artery^21,33.
   1. Lay a sterile cotton square saturated in sterile saline on the drape, lying superiorly and to the left of the incision. Carefully, with 2 sterile cotton-tipped applicators, push the colon and small intestine partially 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.
   2. Cover the exposed intestines with another saline-saturated cotton square.
   3. Periodically monitor peristalsis throughout the surgery, which can be identified by rhythmic movement of the intestines.
2. 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 and should move as one unit.
   1. 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.
   2. 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'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'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's midline and lies flat on its right psoas muscle. Use tweezers to bluntly dissect any overlying fascia to reveal the ganglia (Figure 1B).
3. 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'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).
4. 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 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.
5. This procedure should have minimal blood loss. If bleeding occurs, ensure adequate hemostasis prior to closure.

4. Skin closure

1. After adequate hemostasis is achieved, remove the suture holding the abdominal vessels in place.
2. Using 2 sterile cotton-tipped applicators, carefully replace diverted intestines into the abdominal cavity.
3. With a 5-0 absorbable suture, perform a running stitch to approximate the abdominal muscles.
4. With 5-0 nylon suture, use simple interrupted stitches to close the skin.
5. Apply a generous layer of antibiotic ointment, such as Neosporin, to the incision site.
6. Place the mouse in a clean cage on a heating pad.
7. Observe the mouse every 15 min until it is awake and ambulatory. This usually takes less than 30 min.

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.

1. Prepare a 1% pilocarpine hydrochloride solution in 0.9% NaCl alongside a mixture of 10% potato starch in castor oil.
2. Using a paintbrush, cover the plantar surface of the foot with betadine. Allow this layer to dry completely.
3. Once the betadine layer is dry, use a separate paintbrush to apply a layer of 10% starch in castor oil.
4. Administer 0.25 µL/g body weight of 1% pilocarpine subcutaneously using a syringe into the loose skin over the neck. Start a timer immediately after pilocarpine administration.
5. At 8 min post-injection, take photos of the plantar surface of the foot.
6. Using Fiji, count the number of dark spots located on all six (6) footpads (Figure 2A).

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.

1. Anesthetize the mice (following step 1 of the protocol), harvest the bilateral sciatic nerves, and then promptly euthanize the animal (following institutionally approved protocols).
2. 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 °C.
3. Section the sciatic nerves longitudinally using a cryostat at 20 µm, and place the sections on charged slides.
4. 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.
5. 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.
6. 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.
7. 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.
8. Image nerve sections at least 40 µm apart on a fluorescent microscope on the 10x objective.
9. 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.