Standardized Ileal Bladder Augmentation For Enterocystoplasty In Rats via Midline Laparotomy

Summary

This protocol describes a standardized rat enterocystoplasty model using a pedicled, vascularized, and detubularized ileal patch with temporarily externalized urinary diversion via a cannula tube to achieve reproducible bladder augmentation.

Abstract

Bladder augmentation using intestinal segments remains a cornerstone reconstructive procedure for patients with low-capacity or poorly compliant bladders when conservative therapies fail. Despite its clinical relevance, experimental progress and in-depth study are limited by the lack of standardized, reproducible protocols that reliably model surgical principles in a controlled preclinical setting. Here, we present a step-by-step rat enterocystoplasty protocol that uses a vascularized, detubularized ileal patch as a pedicled flap to augment the native bladder. The procedure includes midline laparotomy, controlled exposure, and stabilization of the bladder with holding sutures, urinary diversion via an exteriorized cannula tube Foley, isolation of a short ileal segment with an intact mesenteric pedicle, detubularization, and tension-free anastomosis with the bladder wall. Emphasis is placed on maintaining mesenteric perfusion, preventing pedicle torsion, ensuring watertight suturing, and securing catheter patency, which are critical for procedural success and reproducibility. Representative intraoperative validation using hyperspectral imaging (HSI) in eight non-survival rats demonstrated preserved tissue oxygenation and perfusion of the bowel segment after key operative steps, supporting tissue viability at the end of this procedure. Thus, this protocol provides a standardized experimental approach for reproducible surgical implementation of rat enterocystoplasty. Potential downstream applications of the model include the investigation of metabolic and electrolyte alterations, infection susceptibility, mucus production, and induction of dysplasia or malignancy, and remain to be addressed in future survival investigations.

Introduction

A physiological urinary bladder function is fundamental to maintaining a high quality of life, as effective urine storage and voiding are critical for continence, social well-being, and overall health. Disorders affecting bladder function can result in substantial morbidity, including urinary incontinence, recurrent infections, renal impairment, and a profound negative impact on daily life.

Both congenital and acquired bladder pathologies can cause significant functional impairment, often necessitating surgical intervention when conservative and pharmacological treatments prove insufficient. Congenital anomalies such as posterior urethral valves¹, bladder and cloacal exstrophy, and epispadias^2,3 are frequently associated with low-capacity, poorly compliant bladders, thereby increasing the risk of upper urinary tract deterioration. Similarly, neurological conditions, including myelodysplasia⁴, multiple sclerosis⁵, and spinal cord injury⁶, often lead to neurogenic bladder dysfunction, impairing both storage and emptying mechanisms. Additionally, infectious etiologies such as tuberculosis⁷ and schistosomiasis⁸ may induce bladder wall fibrosis, reducing compliance and predisposing to secondary complications. In refractory cases, bladder augmentation serves as an effective strategy to improve storage capacity, reduce intravesical pressure, and preserve renal function, ultimately enhancing quality of life and mitigating long-term complications.

According to the American Urological Association (AUA) guidelines on overactive bladder, bladder augmentation (enterocystoplasty) is indicated for patients with diminished bladder capacity, poor compliance, or detrusor overactivity refractory to conservative management⁹. Originally described by Tizzoni and Foggi in 1888 using a canine model¹⁰ and later adapted for human application by von Mikulicz in 1899¹¹, enterocystoplasty remains the standard surgical approach for reducing urinary storage pressures and mitigating the risks of renal damage and incontinence associated with various bladder dysfunctions, including neurogenic bladder, bladder exstrophy, and posterior urethral valves^12,13.

The conventional enterocystoplasty technique involves bivalving the bladder in either the coronal or sagittal plane down to the trigone, followed by the incorporation of a detubularized bowel segment¹². The ileum is the most frequently utilized intestinal segment, typically harvested 25-40 cm proximal to the ileocecal valve¹². When the ileum is unsuitable, the sigmoid colon serves as the preferred alternative¹².

While this procedure offers long-term durability and high patient satisfaction rates¹⁴, the introduction of gastrointestinal tissue into the urinary tract presents notable drawbacks, including metabolic disturbances, bladder stone formation, and an elevated risk of malignancy^15,16,17.

Given these potential complications, there is a critical need for controlled studies to explore the underlying mechanisms, refine surgical techniques, and evaluate alternative augmentation materials. A rodent model provides a structured and reproducible environment to investigate these aspects, enabling researchers to analyze metabolic adaptations following augmentation, including acid-base balance and electrolyte disturbances. Furthermore, it allows the examination of bladder tissue remodeling over time, providing insight into fibrosis, urothelial regeneration, and the integration of augmented segments. Additionally, given the increased risk of malignancy associated with enterocystoplasty, a rodent model offers the opportunity to study histopathological changes over time and identify early markers of carcinogenesis. Importantly, such a model also facilitates research into biomaterial and tissue engineering applications, potentially paving the way for alternative augmentation materials with lower carcinogenic and immunogenic risks¹².

Although experimental bladder augmentation models in rodents and other species have been reported, a directly comparable, standardized step-by-step protocol for ileal enterocystoplasty in rats via midline laparotomy is lacking. Earlier rat studies established classical augmentation cystoplasty as an experimental model and described variants such as seromuscular and laser-assisted enterocystoplasty^18,19,20. Subsequent work addressed long-term histopathological and bacteriological changes, tumor development, transport physiology, and functional outcomes, while additional reconstructive modifications such as partial bladder transplantation and photodynamic replacement of intestinal mucosa with urothelium were also explored^{21,22,23,24,25,26,27}. Taken together, these studies show that rat augmentation models are existent, but heterogeneous in scope and mainly focused on feasibility, biological sequelae, physiology, or specific reconstructive modifications^{18,19,20,21,22,23,24,25,26,27}. Therefore, the objective of this work is to present a detailed procedural guide for performing standardized enterocystoplasty via midline laparotomy in rats. The protocol has been established in a non-survival setting. Applications that require long-term observation are plausible, but might require adaptation of this protocol to a survival-study setting with postoperative recovery and longitudinal follow-up. These could include evaluation of functional outcomes, postoperative failure mechanisms, tissue regeneration over time, or tissue-engineering integration.

The present model was designed to reproduce key technical principles of clinical enterocystoplasty in a controlled rat setting while remaining feasible for standardized experimental implementation. A detubularized ileal patch was selected because detubularization reduces intrinsic bowel contractility and better reflects the reconstructive principle commonly used in clinical augmentation. The use of a pedicled, vascularized segment allows preservation of mesenteric perfusion and enables direct assessment of tissue viability during and after graft preparation. Externalized urinary diversion via a cannula tube Foley was incorporated to provide controlled drainage of the bladder during the procedure and to facilitate standardized intraoperative assessment of the reconstructed segment.

Protocol

All animal procedures outlined in this document were carried out within accredited facilities and have been granted approval by the institutional animal care and use committee (IACUC) of the Baden-Württemberg Regional Council in Karlsruhe, Germany (35-9185.81/G-62/23). Experimental animals were handled in accordance with institutional protocols and in compliance with German legislation governing animal welfare, as well as adhering to the guidelines set forth by the European Community Council (2010/63/EU) and the ARRIVE guidelines. Male Sprague-Dawley rats (n = 8), aged 8–12 weeks and weighing 400 g, obtained from Janvier Labs, were used following a one-week acclimatization period.

1. Anesthesia and analgesia

1. Induce anesthesia by placing the animal in an induction chamber and using 4 vol% isoflurane diluted in 100% oxygen at a flow rate of 5 L/min.
2. For dissociative anesthesia, analgesia, and sedation, administer a subcutaneous injection of 100 mg/kg ketamine (10% solution; equaling 0.35 mL for 400 g of body weight) and 4 mg/kg xylazine (1:10 dilution of 2% solution; equaling 0.75 mL for 400 g)^28,29,30,31upon loss of righting reflex. Administer all anesthetic and analgesic agents on a mg/kg body-weight basis.
   NOTE: Any stated injection volumes are approximate examples for rats of the body weight of 400 g and must be calculated individually for each animal.
3. Confirm analgesic depth by the absence of a firm toe-pinch response using forceps and apply corneal ointment to prevent corneal desiccation.
4. Achieve additional analgesia through a subcutaneous injection of 5 mg/kg carprofen (1:10 dilution of 5% solution; equaling 0.35 mL for 400 g).
5. Maintain narcosis with supplemental isoflurane via a neonatal face mask as required throughout the surgery.
6. Perform intraoperative temperature management by placing the animal on a heating pad set to 40°C. Additionally, place a rectal temperature probe, but it is not obligatory.
7. Provide perioperative and postoperative fluid support by subcutaneous injection of 37 °C isotonic saline solution of 25 mL/kg per hour (equaling 10 mL for 400 g) split up into injections every 20 min. This should include a final injection after abdominal closure, calculated for the expected remaining narcosis of usually 30 minutes.

2. Operating site and instrument preparation

1. Prepare the scrub table with instruments and materials, including:
   Atraumatic preparation forceps
   Blunt overholt clamps
   Preparation scissors
   Double-armed non-resorbable monofilament 6-0 suture
   Single-armed fast resorbable polyfilament 4-0 suture
   A 20 Gauge catheter as an improvised Foley catheter
   Humidified cotton wool swabs
   Gauze swabs
   Isotonic sodium chloride solution
2. Mount the animal on the rodent surgical exposure apparatus as described in previous publications^30,31. Shave the abdominal access area and sterilize it by performing alternating triple scrubs with 70% ethanol and povidone-iodine or chlorhexidine-based antiseptic swabs, working in concentric circles from the incision site outward. Repeat three times.
3. Cover the animal with sterile drapes, and leave only the surgical field exposed.

3. Surgical access via midline laparotomy

1. Perform a median abdominal laparotomy of approximately 5 cm using fine scissors and incise the linea alba longitudinally in the midline to access the peritoneal cavity.
2. Use preparation hooks and moist compresses to retract the abdominal wall and expose the peritoneal cavity.

4. Identification and preparation of the small bowel and the urinary bladder (Figure 1)

1. Identify the small bowel and gently mobilize a loop using atraumatic preparation forceps and humidified cotton swabs (Figure 1A.1).
2. Approximate the small bowel loop to the bladder to identify an appropriately located
3. small bowel segment of approximately 12 mm in length for the augmentation (Figure 1A.2). This should be the first mechanically suitable bowel loop of at least 2 cm proximal to the ileocecal junction.
4. Mobilize the urinary bladder by dissecting the surrounding connective tissue using preparation scissors (Figure 1B).
5. Place stay sutures at the 3, 6, 9, and 12 o'clock positions on the bladder wall using single-armed non-resorbable monofilament 4-0 sutures (Figure 1C.1–1C.5).
6. Grasp the bladder dome with atraumatic forceps and create an approximately 4 mm transverse defect in between the stay sutures using dissecting scissors (Figure 1D.1–1D.2).
7. Replace the 4 stay sutures to involve the cut edge: sequentially pull each suture through. One suture at a time, guide the needle around the incision margin, take a bite from the luminal side directly at the edge, then exit through the adjacent seromuscular layer to loop, retract, and expose the margin (Figure 1E.1–1E.6).

5. Placement and fixation of the cannula tube Foley for external urinary diversion (Figure 1)

1. Insert the intravenous cannula (G20) through the bladder lumen and advance it from the intraluminal side through the abdominal wall to the exterior (Figure 1F.1–1F.3).
2. Remove the metal cannula from the tube while stabilizing the cannula tube with a forceps as an improvised Foley catheter (Figure 1F.4–1F.5).
3. To secure the cannula tube Foley to the skin, pass a non-resorbable monofilament 4-0 suture through the tube lumen and anchor it with an adequate safety margin from the external opening to minimize the risk of subsequent dislodgement (Figure 1G.1–1G.6).
4. Flush the cannula tube Foley with saline to confirm patency, using the tube of a second intravenous cannula one size larger (e.g., G18) than the initial cannula (Figure 1G.1–1G.9).
5. Suture the cannula tube Foley intravesically to the bladder wall using a fast resorbable 4-0 polyfilament suture, maintaining a safety margin from the cannula tube hub to reduce the risk of accidental dislodgement (Figure 1H.1–1H.4).
6. Grasp the cannula tube hub with forceps and detach it from the tubing using scissors (Figure 1H.5–1H.7).
7. Place the cannula tube Foley into the bladder (Figure 1H.8–1H.9).

Figure 1: Small bowel and bladder preparation with placement of an externalized urine diversion cannula tube Foley. (A) Identification and approximation of the small bowel and bladder(A.1–A.2). (B) Mobilization of the bladder by dissection of the peritoneum (B.1–B.4). (C) Placement of stay sutures at the bladder dome (C.1–C.5). (D) Dissection of the bladder wall (cystotomy) (D.1–D.2). (E) Replacement of stay sutures (E.1–E.6). (F) Placement of the cannula for external urinary diversion through the cystotomy (F.1–F.5). (G) Fixation of the external cannula tube segment at the abdominal wall (G.1–G.9). (H) Fixation and final intravesical placement of the cannula tube tip (H.1–H.9). (I) Schematic illustration summarizing the described preparation and cannula tube placement as a Foley. The thick dotted line depicts the small intestine. The thin dotted line depicts the bladder. Black squares depict the image section magnified in the subsequent photo. Please click here to view a larger version of this figure.

6. Isolation of the small bowel segment (Figure 2)

1. Place the small-bowel loop on a sterile gauze swab, assess its mobility toward the urinary bladder, and select an appropriate segment of approximately 12 mm in length (Figure 2A.1–2A.4).
2. Transect the selected bowel segment from the remaining intestine while preserving its mesenteric blood supply and maintain correct orientation to prevent subsequent twisting of the vascular pedicle (Figure 2B.1–1B.4).
3. Gently express residual luminal contents from both transected bowel ends using a cotton swab (Figure 2C.1–2C.6).
4. Reflect both adjacent limbs of the small-bowel loop upward so that only the selected segment remains on the gauze swab (Figure 2D–2E).

Figure 2: Identification and isolation of a small bowel segment for the bladder augmentation. (A) Mobilization and identification of a suitable small bowel segment for bladder augmentation (A.1–A.4). (B) Transection of the selected small bowel segment from the remaining intestinal loop (B.1–B.4). (C) Evacuation of residual luminal contents (C.1–C.4). (D) Separation and positioning of the oral and aboral ends of the remaining small bowel loop. (E) Isolated small bowel segment prepared for subsequent augmentation. (F) Schematic illustration summarizing segment selection and isolation. The thick dotted line depicts the isolated small bowel segment. Please click here to view a larger version of this figure.

7. Preparation of the bowel segment and bladder augmentation (Figure 3)

1. Open the isolated bowel segment along the antimesenteric border using scissors, thereby creating a detubularized patch of approximately 12 mm × 8 mm (Figure 3A.1–3A.3).
2. Excise the newly formed corner edges of the graft (Figure 3B.1–3B.3).
3. Proceed with anastomosis of the small-bowel segment using a double-armed 6-0 monofilament suture. At the 12 o'clock position, pass the needle through the bladder wall from the extraluminal to the intraluminal side (Figure 3C.1–3C.2).
4. Pass the needle through the lower right corner of the small-bowel segment from the intraluminal to the extraluminal side. Ensure the double-armed suture is pulled through to approximately its midpoint, as this defines the superior anchoring point (Figure 3C.3–3C.4).
5. Tighten the suture to approximate the bowel graft to the bladder, resulting in an approximately 45° anti-clockwise rotation of the segment. Tie the knot on the extraluminal side (Figure 3D.1–3D.2) and pass the second needle around the mesentery of the bowel segment to enable circumferential suturing of the graft from both sides (Figure 3E.1).
6. Begin circumferential anastomosis of the bladder augmentation patch: sequentially suture the bowel segment into the bladder defect using an outside-to-inside bite at the bladder edge, followed by an inside-to-outside bite through the bowel segment, then proceed with the next stitch.
7. Sequentially release the stay sutures (Figure 3E.1–3E.15). Use slightly larger bites for the small bowel patch than for the bladder; however, it is intended that there be substantially more length to cover on the patch, so that unsutured excess small bowel length will remain.
8. After completing circumferential closure of the bladder defect, tie off one suture end. Using the remaining suture, close the residual free portion of the bowel segment in a conical ("pointed-cap-like") fashion and tie off the suture (Figure 3F.1–3F.6).
9. Return the bladder to its anatomical position and reposition the intestine, ensuring that the bowel is not twisted (Figure 3G.1–3G.2).

Figure 3: Construction of the small bowel patch and circumferential bladder augmentation anastomosis. (A) Dissection and preparation of the small bowel segment (graft preparation) (A.1–A.3). (B) Excision of the corner edges of the graft (B.1–B.3). (C) Initiation of the anastomosis by placing the anchoring stitch between the graft and the bladder opening (C.1–C.4). (D) Fixation of the anchoring point to align graft and cystotomy margins (D.1–D.2). (E) Circumferential anastomosis of the bladder augmentation patch to the bladder wall (E.1–E.15). (F) Closure of the remaining small bowel segment tissue in a conical, pointed-cap configuration (F.1–F.6). (G) Repositioning of the organs to their anatomical position (G.1–G.2). (H) Schematic illustration summarizing graft preparation and bladder augmentation anastomosis. The thick dotted line depicts the small bowel segment. The thin dotted line depicts the opening of the bladder wall. The white arrow depicts the needle. Please click here to view a larger version of this figure.

8. End-to-end small-bowel anastomosis (Figure 4)

1. Use the double-armed non-resorbable monofilament 6-0 suture cut in half for the anastomosis. Pass the needle through the antimesenteric border of both bowel ends. Do not tighten the knot yet, but use a parachute technique by placing a clamp on the suture segment between the bowel ends to preserve visualization (Figure 4A.3–4A.6).
2. Place a suture through the mesenteric border of both bowel ends and clamp in a similar fashion (Figure 4A.7–4A.10).
3. Place two sutures through the anterior wall of both small-bowel ends to facilitate subsequent closure, and secure the suture ends with clamps (Figure 4B.1–4B.4).
4. Reflect both bowel ends upward in order to expose the posterior small bowel wall. Analogous to the anterior wall, place two sutures through the posterior wall and secure the suture ends with clamps (Figure 4C.1–4C.4).
5. Once all sutures are placed, sequentially tie off all six sutures (Figure 4D.1–4D.6).
6. Close the mesenteric defect with 2–3 interrupted fast resorbable polyfilament 4-0 sutures to prevent subsequent herniation (Figure 4F.1–4F.8).

Figure 4: End-to-end small bowel anastomosis and mesenteric defect closure. (A) Placement of alignment sutures at the antimesenteric and mesenteric borders (A.1–A.10). (B) Placement of two anterior-wall approximation sutures (B.1–B.4). (C) Exposure of the posterior wall and placement of two posterior-wall approximation sutures (C.1–C.4). (D) Sequential tying of all six sutures to complete the anastomosis (D.1–D.6). (E) Enlarged end-to-end anastomosis (E.1–E.2). (F) Closure of the mesenteric defect with interrupted sutures to prevent internal herniation (F.1–F.8). (G) Schematic illustration of the end-to-end small bowel anastomosis. Please click here to view a larger version of this figure.

9. Terminal steps and considerations

1. Close the abdominal wall in a multilayer fashion under sterile conditions, in accordance with previously published protocols^29,30,31.
2. For non-survival procedures or scheduled termination after follow-up, euthanize animals under deep anesthesia by sharp cardiectomy, cervical dislocation, or guillotine decapitation.
3. If required, excise the bladder for subsequent anatomical evaluation (e.g., assessment of the bladder augmentation site and bowel-bladder anastomosis), biomolecular analyses, or histopathological imaging.
4. For this purpose, remove the augmented bladder en bloc by careful stepwise dissection under direct visualization, with gentle release of surrounding adhesions, identification of the native bladder, augmented bowel patch, and anastomotic borders, followed by transection of the distal bladder outlet and adjacent mesenteric or bowel attachments as required, while avoiding compression or distortion of the specimen to preserve tissue architecture for subsequent analysis.

Results

To confirm tissue viability after this protocol, tissue oxygenation and perfusion were assessed during the procedure in eight rats (Table 1). Before enterocystoplasty, hyperspectral imaging (HSI) of the small bowel was performed to visualize tissue oxygen saturation (StO₂) and perfusion (NIR) (Figure 5A.1). As a negative control, maximal hypoperfusion was induced by clamping of the aorta (Figure 5A.2). HSI of re...

Discussion

This work provides a reproducible surgical protocol for enterocystoplasty in rats using a midline laparotomy, extracorporeal cannulation, and incorporation of a detubularized ileal segment. The technique follows the principles described by Tizzoni and Foggi for bladder augmentation and adapts them to a rodent model by combining precise anesthesia, aseptic preparation, exteriorized cannulation for urine diversion, and the harvesting of a vascularized small‑bowel patch.

This protocol was c...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Atraumatic preparation forceps	Aesculap	FB395R	DE BAKEY ATRAUMATA atraumatic forceps, straight
Blunt overholt clamps	Aesculap	BJ012R	BABY-MIXTER preparation and ligature clamp, bent, 180 mm
Cannula tube foley for flusing and diversion	Mdk Mart	23-CA-CD-UL-CD	Cannula G18 (only the tube is used to flush the diversion cannula)
Carprofen	cp pharma	115	Carprofen 0,5% (5% with 50 mg/mL diluted 1:10) (Carprosol)
Cotton whool swabs	Amazon	ASIN: B0FPDB8ZGS	Non-sterile cotton whool swabs (autoclave for survival experiments)
Fast resorbable polyfilament 4-0 sutures	COVIDIEN	SV-494	Single-armed polyfilament surgical suture from resorbable polyglactin with one P-13 needle
Fixation rods	legefirm	‎500343896	Tuning forks used as y-shaped metal fixation rods
Foley catheter	B. Braun	4242010-02	20 Gauge 50 mm Introcan Safety 2 Catheter as diversion cannula
Gauze swabs	Medrull	ASIN: B09164D1JG	Sterile gauze swabs
Heating pad	Royal Gardineer	IP67	Royal Gardineer Heating Pad Size S, 20 Watt
Hyperspectral imaging camera system	Diaspective Vision	TIVITA Tissue Halogen	HSI system for validation
Isoflurane	Piramal Critical Care	PZN / EAN 09714675 / 4150097146757	100% isoflurane 250 mL
Isoflurane vaporizer	UNO ROESTVASTSTAAL BV	180000002	Isoflurane vaporizer
Isotonic sodium chloride solution	B. Braun	ASIN: B007PZJOQ4	0,9% sodium chloride solution
Ketamine	cp pharma	1202	Ketamine 10% (100 mg/mL)
Metal cannula	BD (Beckton, Dickinson)	301300	BD Microlance 3 cannula 20 Gauge as preparation hooks
Neontal face mask	asia connection	ME03016-0	Neontal face mask size #0 for neonate
Non-resorbable monofilament 4-0 sutures	COVIDIEN	SP-670	Single-armed monofilament surgical suture from non-resorbable poly-propylene with one C-16 needle
Non-resorbable monofilament 6-0 sutures	COVIDIEN	VP-733-X	Double-armed monofilament surgical suture from non-resorbable poly-propylene with two CV-22 needles
Ophthalmic ointment	Bayer Vital GmbH	1578681	5% dexpanthenol
Plastic perfusor tube	M. Schilling GmbH	S702NC150	Connecting tube COEX 150 cm
Preparation scissors	Aesculap	BC177R	JAMESON preparation scissors, bent, fine model, blunt/blunt, 150 mm (6")
Rat induction chamber	wpiinc	EZ-1785	Induction chamber for narcosis
Steel plate	Maschinenbau Feld GmbH	C010206	Galvanized sheet plate, 40 x 50 cm, thickness 4.0 mm
Xylazine	cp pharma	1206	Xylazine 0,2% (2% with 20 mg/mL diluted 1:10) (Xylavet)