Murine Model for Selective Transection of Intestinal Autonomic Nerves

Summary

This protocol aims to enable precise, selective transection of intestine-innervating sympathetic and parasympathetic nerves in mice using the superior mesenteric artery as a consistent anatomical landmark, providing a focused and reproducible model to study autonomic regulation of intestinal physiology and disease.

Abstract

Autonomic nerves comprise the sympathetic and parasympathetic systems, which act in a largely antagonistic manner to regulate diverse physiological functions. With growing interest in the roles of autonomic innervation in intestinal physiology and pathology, there is an increasing need for experimental models that enable precise manipulation of gut-directed sympathetic and parasympathetic inputs. Here, we present a surgical protocol to selectively transect the sympathetic and parasympathetic nerves innervating the intestine. Using the superior mesenteric artery (SMA) as a reproducible anatomical landmark, this approach allows reliable identification of intestinal autonomic nerve bundles and subsequent targeted transection. In contrast to broader denervation approaches such as ganglionectomy or subdiaphragmatic vagotomy, this approach is intended to achieve a more anatomically localized transection of intestine-directed autonomic bundles, which may help reduce extra-intestinal interference. The injury is controllable and is typically associated with minimal bleeding, supporting procedural consistency and postoperative recovery. Although the surgery requires appropriate microsurgical training, the protocol is adaptable, enabling users to tailor key parameters to specific experimental aims. Importantly, this model provides a practical platform to investigate the roles of sympathetic and parasympathetic nerves in regulating intestinal function under both physiological and disease conditions. This article aims to describe the procedure in detail to facilitate its adoption by new users in their research.

Introduction

The gastrointestinal tract is under continuous control of the autonomic nervous system (ANS), with sympathetic and parasympathetic pathways coordinating fundamental aspects of intestinal physiology, including motility, secretion, epithelial barrier function, and immune homeostasis^1,2. In the colon, extrinsic sympathetic inputs can exert region-dependent effects across multiple cellular compartments, underscoring that broad ANS perturbations may obscure site-specific intestinal phenotypes¹. Conversely, parasympathetic circuits, classically mediated by vagal signaling and downstream cholinergic mechanisms, are increasingly recognized as integral to maintaining intestinal immune balance and shaping mucosal responses, further motivating experimental strategies that can disentangle sympathetic versus parasympathetic contributions with anatomic precision².

Beyond physiological regulation, autonomic activity has emerged as a relevant modifier of intestinal disease biology, including colorectal cancer (CRC). Recent work in CRC demonstrates a bidirectional sympathetic nerve–mesenchymal program, in which β2-adrenergic signaling and NGF-expressing cancer-associated fibroblasts reinforce neural–stromal interactions and promote tumor progression, providing direct evidence that sympathetic circuits can be functionally coupled to CRC pathogenesis³. In parallel, cholinergic signaling has been linked to CRC growth and tumor-associated immune features, as shown by pharmacologic inhibition of muscarinic receptor 3 in an orthotopic mouse model and by connecting cholinergic signaling with immune checkpoint expression and other malignant hallmarks^4,5. Importantly, autonomic influences on intestinal disease may also converge with other modulators, such as psychological stress and the gut microbiota. The enteric nervous system can relay stress signals to intestinal inflammation⁶, and microbial signals can engage gut–brain circuits that tune gut-extrinsic sympathetic neurons, collectively supporting a multi-factorial framework in which neural regulation intersects with immune and microbial ecology^6,7.

These converging lines of evidence highlight the need for experimental approaches that can isolate gut-directed autonomic effects with sufficient anatomical precision to support causal inference across both physiological and pathological settings. Despite rapid progress in circuit mapping and neuromodulation, many commonly used interventions remain limited in organ specificity. Systemic pharmacologic manipulation, chemical sympathectomy, and subdiaphragmatic vagotomy can influence broad autonomic outputs and thereby alter immune and metabolic set points beyond the intestine, complicating causal attribution to colon-specific neural inputs⁸. For example, vagal circuits can suppress inflammatory cytokine production⁹, and vagal stimulation can modulate pancreatic endocrine function and β cell dynamics, illustrating that upstream or non-selective interventions may introduce extra-intestinal effects that confound interpretation of intestinal phenotypes¹⁰. These limitations are particularly consequential for studies aiming to isolate colon-restricted autonomic mechanisms in physiology and pathology. The overall goal of the method is therefore to enable anatomically guided, selective transection of gut-directed sympathetic and parasympathetic fibers in mice, providing a practical model for interrogating intestinal autonomic regulation with reduced off-target impact.

In summary, this protocol reports a more precise murine intestinal autonomic denervation approach with minimal disruption to the autonomic innervation of other abdominal organs, thereby enhancing interpretability by reducing potential bias from extra-intestinal organ effects.

Protocol

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Beijing Friendship Hospital, Capital Medical University (NO. 24-2026). Adult wild-type male C57BL/6J mice were used in this study (8–10 weeks old, 20–30 g). Detailed information on reagents and equipment is provided in the Table of Materials.

Inclusion criteria: Mice of the same strain, sex, and age range were used within each experiment to minimize biological variability. Only healthy animals demonstrating normal activity, grooming behavior, food and water intake, and stool output during the acclimatization period were included. In addition, mice were acclimated to the animal facility for at least 7 days before surgery.

Exclusion criteria: Mice were excluded in cases of uncontrolled bleeding or major vascular injury during surgery, intestinal perforation, or significant tissue damage incompatible with survival. Animals that failed to recover from anesthesia, defined as not becoming ambulatory within 2 h after surgery, were also excluded. Additional exclusion criteria included postoperative complications such as persistent wound dehiscence, severe infection, or weight loss exceeding 15% within 48 h or 20% over any 72 h period. Mice with surgical misidentification of the nerve branch, confirmed by post-hoc histological evaluation, were also excluded from the study.

1. Preoperative preparation

1. Sterilize all surgical instruments.
2. Administer preventive analgesia to all mice 30 min prior to surgery to avoid pain sensitization. Administer meloxicam (5 mg/kg body weight) via subcutaneous injection.
3. Prewarm the heating pad to 37 °C and place it underneath the surgical platform.
4. Spray the surgical area (including the anesthesia induction chamber) with 75% ethanol, and wipe thoroughly with paper towels. Place sufficient paper towels at the bottom of the induction chamber.
5. Inspect the anesthesia induction chamber and tubing to confirm there are no leaks. Refill and ensure an adequate supply of isoflurane. Induce anesthesia with 3%–3.5% isoflurane delivered in 1 L/min oxygen. Monitor the mouse continuously; a surgical plane is typically reached within 1–2 min.
6. Place the mouse in the supine position with all four limbs fully extended, and secure with hypoallergenic tape. Fit and secure the nose cone. Reduce isoflurane to 1.5%–2.5% for maintenance. Assess anesthetic depth by the absence of a pedal withdrawal response (no reaction to toe pinch) and regular respiration (55–65 breaths/min).
   NOTE: During prolonged microsurgical procedures, anesthetic depth is continuously monitored beyond the initial surgical plane. In addition to assessing the pedal withdrawal reflex and respiratory rate every 5–10 min, we monitor respiratory pattern (e.g., regularity and depth) and heart rate via visual observation or contact with the thoracic wall. Mucous membrane color (assessed from the paws or lips) is also checked periodically to ensure adequate oxygenation. If the mouse shows any signs of lightening anesthesia, such as increased respiratory rate (>80 breaths/min), spontaneous movements, or a return of the pedal reflex, the isoflurane concentration is increased in increments of 0.25%–0.5% until an adequate surgical plane is restored. Conversely, if respiration becomes shallow or irregular, or if the heart rate drops markedly, isoflurane is reduced by 0.25%–0.5% until stable cardiorespiratory function returns. This dynamic adjustment ensures that the mouse remains at an appropriate anesthetic depth throughout the procedure while minimizing cardiorespiratory depression.
7. Apply ophthalmic ointment to both eyes to prevent corneal drying and injury under anesthesia.
8. Shave the abdominal hair from the level of the pubic symphysis to the area below the costal margin using an electric hair shaver. Then, move the surgical platform under a stereomicroscope.
9. Disinfect the surgical site using povidone-iodine swabs in an outward, concentric pattern (from the center to the periphery), covering the area from below the xiphoid to the pubic symphysis and laterally to both flanks. Repeat 3 times. Then wipe with 70% ethanol in the same manner. Ensure the field is free of any loose hair.
10. Don a surgical mask and disposable cap, wear a sterile surgical gown, and put on sterile gloves.
11. Cover the mouse with a sterile fenestrated drape. Adjust the fenestration to approximately 2 × 3 cm and align it with the abdominal midline to fully expose the operative field. Secure the four corners of the drape.

2. Laparotomy and exposure

1. Using sterile forceps, gently lift a small fold of skin just cranial to the pubic symphysis. With sterile sharp-tipped scissors, make a small initial incision at the apex (~0.3 cm), then extend the skin incision cranially along the midline for 1.5–2.0 cm.
   NOTE: After inserting the sharp scissor tips into the initial opening, angle the tips upward and perform a controlled “push-cut” along the undersurface of the skin to avoid cutting too deeply and injuring the abdominal musculature or viscera. Use forceps and scissors to bluntly dissect the loose subcutaneous connective tissue from the underlying abdominal wall musculature. Dissect slightly wider than the skin incision to facilitate subsequent exposure.
2. Approximately 0.5 cm caudal to the xiphoid process, carefully insert the tips of sterile fine forceps into the linea alba and lift a small tent of tissue.
3. Using sterile, sharp-tipped scissors, create a small opening (2–3 mm) at the lifted site. At this point, the smooth peritoneum and/or underlying organs (e.g., a liver lobe) should be visible.
4. Insert one blade of sterile blunt-tipped scissors into the peritoneal cavity, keeping it closely opposed to the abdominal wall. Under direct visualization, extend the incision along the linea alba toward the cranial direction until the total length matches the skin incision.
   NOTE: Keep the scissor tip angled upward throughout and proceed only under direct vision to ensure that no bowel loops or blood vessels are beneath the scissors.

3. Identification of the superior mesenteric artery and nerve transection

1. Gently retract the abdominal wall laterally using sterile tape strips to expose the peritoneal cavity.
2. Apply moderate pressure with a cotton swab to the abdominal wall to allow the intestines to exteriorize. Gently move the small intestine, colon, and cecum to the left side of the mouse.
   NOTE: Cover the exposed bowel with a sterile cotton pad or gauze moistened with warm (≈37 °C) sterile saline to prevent tissue drying and heat loss.
3. Using the right hand, gently displace the stomach, duodenum, and colon en bloc toward the left upper quadrant with a sterile, moistened cotton swab (or blunt-tipped micro forceps). The abdominal aorta, anterior to the spine, should be clearly visible at this stage, sometimes partially covered by a small amount of retroperitoneal fat.
4. Identify the left renal vein and the celiac trunk. The superior mesenteric artery (SMA) arises as a single arterial branch from the anterior wall of the abdominal aorta, typically ~2–4 mm caudal to the celiac trunk and at or slightly above the level of the left renal vein.
   NOTE: The left renal vein is a dark, thick vein coursing horizontally across the anterior aspect of the abdominal aorta before draining into the inferior vena cava. The celiac trunk is usually the first major arterial branch arising from the anterior wall of the abdominal aorta, located just cranial (superior) to the left renal vein, and commonly trifurcates into the hepatic, splenic, and left gastric arteries. In contrast, the SMA is a single trunk without hepatic/splenic branches. It enters the root of the mesentery and supplies the jejunum, ileum, and proximal colon; tracing its course into the mesentery can help confirm correct identification.
5. Identify the milky-white, cord-like bundles running parallel to the SMA as the target nerve fibers. The nerve bundle located superior to the SMA corresponds to the parasympathetic fibers, whereas the bundle located inferior to the SMA corresponds to the sympathetic fibers (Figure 1A).
6. Using micro forceps and micro scissors, very gently separate the fat and lymphatic tissue surrounding the SMA to fully expose the nerve fibers. Gently lift the sympathetic or parasympathetic nerve bundle with micro forceps to separate it from the SMA, then transect it quickly and carefully (Figure 1).
   NOTE: During dissection, keep the tips of the forceps and scissors oriented toward the lateral wall of the artery and avoid directing the instruments vertically into deeper planes. If major bleeding occurs (e.g., hemorrhage from the SMA or abdominal aorta that cannot be controlled within 30 s by applying pressure with a sterile cotton swab), the procedure is immediately terminated. The animal is humanely euthanized while still under anesthesia and is excluded from all subsequent analyses. This endpoint is predefined in accordance with IACUC-approved humane endpoints to prevent unnecessary suffering. Additional troubleshooting guidance for common intraoperative complications is provided in Supplementary File 1.

4. Closure of the abdominal cavity

1. Remove all intra-abdominal gauze/cotton pads. Carefully inspect the abdominal cavity for any active bleeding, with particular attention to the SMA dissection field, the nerve transection site, and the abdominal wall cut edges.
2. Gently irrigate the peritoneal cavity with a small volume of warm sterile saline (0.5–1.0 mL), then carefully wick away excess fluid using a sterile dry cotton swab.
3. Perform an instrument and gauze/cotton count to ensure that no foreign material is retained within the abdominal cavity.
4. Using 6-0 absorbable suture, close the peritoneum and muscle layer with a continuous running stitch starting from one end of the incision. Each bite must incorporate the peritoneum and the abdominal wall musculature on both sides.
   NOTE: Lift the wound edges with forceps and place each stitch under direct visualization to avoid catching underlying bowel or omentum. If needed, gently displace the intestines away from the incision using a moistened cotton swab.
5. Close the skin with interrupted sutures using 5-0 silk braided non-absorbable suture. Place sutures at ~2 mm intervals and tie knots with moderate tension, approximating (but not strangulating) the skin edges to avoid ischemia.
   NOTE: Poor wound apposition may result in an incisional hernia. Ensure the muscle layer is tightly sealed, and the skin edges are well aligned.
6. Gently clean the closed skin incision once more with a povidone-iodine swab.
7. Mark the mouse (ear or tail) with the date of surgery.
8. Remove the nose cone and transfer the mouse to a clean, soft recovery cage placed on a 37 °C warming pad (single housing). Maintain the animal in lateral recumbency to reduce the risk of aspiration.
   NOTE: Confirm that the warming pad remains stable at 37 °C to prevent hypothermia. If necessary, administer 1 mL of warm sterile saline subcutaneously for volume support.
9. Continuously monitor the mouse until it has fully regained consciousness and can maintain a prone position independently.
   NOTE: Postoperative analgesia was continued with meloxicam (5 mg/kg, subcutaneous, once every 24 h for up to 48 h after surgery), with additional veterinary assessment if signs of persistent pain or distress were observed.

5. Postoperative monitoring parameters

NOTE: Postoperative monitoring was performed twice daily for the first 72 h and then daily for the duration of the study. Parameters assessed included:

1. Monitor general appearance, including grooming, coat condition, posture, and activity level.
2. Measure body weight daily; define a loss of more than 15% of preoperative body weight within 48 h or 20% over any 72 h period as a humane endpoint.
3. Inspect the surgical site for signs of infection, dehiscence, or discharge.
4. Monitor behavior, including spontaneous movement, response to handling, and signs of distress (e.g., vocalization and restlessness).
5. Evaluate any animal showing persistent signs of pain or distress despite analgesia by veterinary staff and, if necessary, euthanize the animal.

Results

Using the SMA as a consistent anatomical landmark, intestinal autonomic nerve bundles can be identified as thin, pearly-white structures running parallel to the vessel. Based on their anatomical position relative to the SMA, the superior bundle was designated as the presumed parasympathetic bundle and the inferior bundle as the presumed sympathetic bundle (Figure 1A,D). A positive intraoperative result is achieved when the target bundle is clearly visualized and completely i...

Discussion

The murine SMA arises from the anterior wall of the abdominal aorta, inferior to the celiac trunk, and provides arterial inflow to major abdominal viscera, including the intestine¹⁴. Anterior to the SMA lie the colon and its mesentery, whereas posteriorly it is closely related to the left renal vein and the abdominal aorta; the superior mesenteric vein typically courses along its right side. In mice, the SMA is extremely delicate and is surrounded by critical organs and major vessels. Importantly,...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
5-0 silk braided non-absorbable suture	Ethicon	W500
6-0 PGA Suture	Jinhuan	CR631
Anesthesia Induction Chambers	RWD Life Science	V100
Animal Anesthesia Machine	RWD Life Science	R500IP
Anti-Choline Acetyltransferase antibody	Abcam	ab181023
Artificial Tears Lubricant Ophthalmic Ointment	Akorn	59399-162-35
C57BL/6J Mice	Shanghai Model Organisms CenterTechnology	SM-001
Cholera Toxin Subunit B (CTB), Alexa FluorTM 555 Conjugate	Thermo Scientific	C34776
Compound Microscope	Yuyan Instruments	512
DAPI and Hoechst Nucleic Acid Stains	Invitrogen	D3571
Fast Green	Thermo Scientific	F7252
Fine Hemostatic Forceps	Jinzhong surgical instrument	JCG020
Hypoallergenic Surgical Tape	3M Blenderm	70200419342
Isoflurane	RWD Life Science	R510-22-10
Micro Forceps(Curved)	Jinzhong surgical instrument	WCC200
Micro Forceps(Straight)	Jinzhong surgical instrument	WCC190
Micro Needle holders	Jinzhong surgical instrument	WBA050
Micro scissors	Jinzhong surgical instrument	Y3F010
Micro syringe	Hamilton	7632-01
Ophthalmic Scissors(pointed tip)	Jinzhong surgical instrument	JC2303
Ophthalmic Scissors(rounded tip)	Jinzhong surgical instrument	JC2303
Povidone-Iodine Swabstick	Winner Medical	206
Tissue and Cell Fixative (4% Paraformaldehyde, PFA)	MedChemExpress	HY-DY3003
Tissue OCT-Freeze Medium	Sakura	C2077
Tyrosine Hydroxylase (E2L6M) Rabbit Monoclonal Antibody	Cell Signaling Technology	58844S