Vesicostomy in the Rat Model for Low-Pressure Urinary Drainage in Neurogenic Bladder Research

Summary

This protocol provides a standardized method for performing cutaneous vesicostomy and postoperative wound management in rats, enabling continuous low-pressure urinary drainage for preclinical spinal cord injury research.

Abstract

The rat is the primary animal model for preclinical research on spinal cord injury (SCI)-induced neurogenic lower urinary tract dysfunction (nLUTD). Owing to its anatomical and functional similarity to humans, rats closely recapitulate key pathological consequences of SCI. However, a crucial translational gap remains: whereas standard bladder management in humans typically relies on continuous or intermittent catheterization, comparable approaches are not feasible in rodents. The common practice of manual bladder expression generates exceptionally high intravesical pressures, which may introduce methodological bias by independently promoting pathological bladder remodeling after SCI. This protocol describes cutaneous vesicostomy in rats, together with postoperative wound management, to establish continuous low-pressure urinary drainage for preclinical SCI research. Cutaneous vesicostomy—the surgical creation of a stoma between the bladder dome and the lower abdominal wall—enables continuous urinary drainage, thereby maintaining a low-resistance drainage route and bypassing the urethra. In this way, excessive urine retention and high intravesical pressures can be avoided, approximating the concept of continuous low-pressure decompression used clinically during the early post-injury phase with suprapubic or transurethral urinary diversion, while recognizing that normal storage and continence mechanisms are abolished. Therapeutic effects that might otherwise be obscured by repetitive overfilling and high-pressure manual bladder expression may thus be more readily detected. Moreover, vesicostomy offers a conceptual strategy to mitigate sex-related constraints in preclinical SCI research. The considerably greater urethral length in male rodents results in higher outflow resistance, making manual bladder expression substantially more difficult and effectively confining most preclinical studies to female animals, despite the predominance of male patients among clinical SCI cases. As an alternative drainage strategy, vesicostomy has the potential to broaden experimental bladder management paradigms and enhance the translational relevance of future studies, while its feasibility and outcomes in male rats will need to be established in dedicated follow-up work.

Introduction

Healthy bladder function relies on the coordinated function of the bladder muscle and urethral sphincter, achieved through a complex neuronal network involving the brain, central nervous system (CNS), and peripheral nervous system (PNS), which enables conscious control of the lower urinary tract (LUT)^1,2. The complexity of these mechanisms and their switch-like circuitry³ renders bladder function susceptible to a wide range of injuries and diseases. Accordingly, following spinal cord injury (SCI), almost all patients experience impaired bladder sensation and control^4,5. The term neurogenic bladder encompasses a broad spectrum of dysfunctions affecting urine storage and/or voiding, depending on the characteristics of the injury. In particular, suprasacral injuries lead to detrusor overactivity in combination with sphincter overactivity, often accompanied by loss of coordination between the two—termed detrusor-sphincter dyssynergia (DSD)—resulting in excessive urine retention and high intravesical pressures^6,7. This can severely damage the LUT, threaten the integrity of the upper urinary tract (UUT), and, if left untreated, result in life-threatening complications⁸. Intermittent or continuous urinary drainage via catheterization represents the current standard of bladder management in humans following SCI, preventing bladder overfilling through a safe, pressure-free procedure^9,10,11,12.

With no effective treatment currently available for SCI and predominantly symptomatic management of associated neurogenic LUT dysfunction (nLUTD), the development of novel therapeutic strategies remains a major research priority. Animal models enable investigation of disease mechanisms at a level of complexity comparable to that in humans while providing exceptional intragroup homogeneity under standardized experimental conditions¹³. Among these, the rat is the primary preclinical model, with the majority of SCI studies conducted in this species^14,15. This preference reflects its anatomical and functional similarity to humans, reproducible pathological response, and widespread availability. Nevertheless, an important translational gap remains that may limit the interpretation and clinical relevance of bladder-related findings derived from rodent models.

Although rats develop nLUTD following SCI that resembles key features observed in humans, bladder management differs substantially between species. Because intermittent catheterization and other permanent urinary drainage approaches are generally not feasible in rats, urine evacuation is routinely achieved by manual bladder expression¹⁶. This technique, analogous to the Credé maneuver¹⁷, which is generally not recommended in clinical practice, generates markedly elevated intravesical pressures. Consequently, it may introduce methodological bias by independently promoting pathological bladder remodeling after SCI, thereby reducing the translational relevance of experimental findings or obscuring genuine therapeutic effects. An additional limitation arises from sex-related anatomical differences. The considerably greater urethral length in male rodents results in higher outflow resistance, making manual bladder expression substantially more difficult and effectively restricting most preclinical SCI studies to female animals¹⁴. Given that the majority of clinical SCI cases occur in males^18,19, this discrepancy further emphasizes the need for alternative bladder management strategies in preclinical rat SCI models.

Cutaneous vesicostomy involves the surgical creation of a stoma between the bladder and the lower abdominal wall by advancing the bladder dome to the skin and securing it with sutures. This enables continuous urinary drainage while maintaining a low-pressure drainage route and bypassing the urethra. Primarily used in children with complex medical conditions or when conventional drainage options are unsuitable, cutaneous vesicostomy provides reliable urinary diversion with minimal maintenance requirements and a low complication rate^20,21,22. Beyond its clinical application, vesicostomy has also been successfully performed in rodent models^23,24,25,26, although not in the context of SCI.

In a recent study¹⁶, we addressed the translational limitations associated with manual bladder expression in preclinical SCI models and introduced vesicostomy in rats together with a postoperative wound management regimen that enabled extended follow-up. Direct comparison of the two bladder management approaches demonstrated greater structural bladder alterations following manual bladder expression, whereas vesicostomy animals exhibited values closer to those of healthy controls across several parameters. Most notably, bladder wall thickness increased from a mean of 768.26 µm (SD 28.05) in healthy rats to 1181.94 µm (SD 289.56) two months after SCI with manual bladder expression, with some animals reaching values of up to 1485 µm. In contrast, rats managed by vesicostomy showed a mean bladder wall thickness of 963.08 µm (SD 68.82), and only one animal exceeded 1000 µm.

This study provides a standardized protocol for performing vesicostomy in rats, including perioperative care and postoperative wound management, to enable reproducible continuous low-pressure urinary drainage in preclinical SCI research. The protocol is intended to facilitate reproduction of the technique and assist investigators in determining its suitability for studies in which high-pressure manual bladder expression may introduce methodological bias or in which the inclusion of male animals is desirable. The aim of this project was to establish a standardized vesicostomy protocol for rats with SCI and to evaluate its feasibility and safety over a two-month follow-up period. In addition, a standardized postoperative wound management regimen was applied to minimize skin-related complications and maintain stomal patency through regular calibration.

Protocol

Three-month-old female Lewis rats were used for this study. Although this protocol was primarily developed to avoid manual bladder expression following SCI, details of the contusion SCI procedure are not provided here but have been described previously. All procedures were approved by the Austrian Governmental Legal Entity on Animal Research (BMWFW-2022-0.062.928 and BMWFW-2023-0.848.340) and were conducted in accordance with local guidelines for animal use and the ARRIVE guidelines. Overall, the vesicostomy procedure requires approximately 15 min.

1. Preoperative Preparation

1. Prepare a sterile working area, including a microscope or magnification glasses (2.5–3.5× magnification), all required surgical instruments, and a heating pad.
2. Prewarm the heating pad to 30°C–35°C (hand-warm) and maintain this temperature throughout the surgery. Replace the heating pad if its temperature drops below 30°C during prolonged surgical procedures.

2. Animal Preparation

1. Anesthesia
   1. Use a general anesthetic regimen appropriate for the surgical procedure.
   2. Prepare a fresh anesthetic mixture containing medetomidine (0.15 mg/kg), midazolam (2 mg/kg), and fentanyl (0.005 mg/kg) according to the provided calculation table (Table 1). Select the anesthetic dose and corresponding final injection volume based on the body weight closest to the animal’s exact weight, and administer the mixture intramuscularly using a 1 mL syringe fitted with a 27G needle.
2. Surgical Preparation
   1. Once the rat is anesthetized, shave the lower abdomen to the vaginal region.
   2. Wash the surgical field twice with warm soapy water.
   3. Wash the surgical field twice with warm clean water.
   4. Disinfect the surgical field twice with a mixture of 7.5% povidone-iodine and 70% alcohol, including the vaginal region.
   5. Administer 1.5 mg enrofloxacin and 0.2 mg long-acting meloxicam, dissolved in 0.2 mL pre-warmed 0.9% sodium chloride (NaCl), subcutaneously.
   6. Administer 3 mL pre-warmed 0.9% NaCl subcutaneously to compensate for fluid loss.
   7. Apply 2% dexpanthenol ophthalmic ointment to both eyes to prevent corneal desiccation.
   8. Position the animal supine on the pre-warmed heating pad and adjust the microscope or magnification glasses.
   9. Confirm an adequate surgical plane of anesthesia by performing a nociceptive withdrawal test on both hindlimbs before beginning surgery.
   10. If the withdrawal reflex is still present, wait 2 min and repeat the nociceptive withdrawal test. Continue this procedure at 2-min intervals until the reflex is absent.

Body weight (g)	Medetomidine (1 mg/mL), mL	Midazolam (5 mg/mL), mL	Fentanyl (0.05 mg/mL), mL	Total volume (mL)
100	0.015	0.04	0.010	0.065
125	0.019	0.05	0.013	0.082
150	0.023	0.06	0.015	0.098
175	0.026	0.07	0.018	0.114
200	0.030	0.08	0.020	0.130
225	0.034	0.09	0.023	0.147
250	0.037	0.10	0.025	0.163
275	0.041	0.11	0.028	0.179
300	0.045	0.12	0.030	0.195
325	0.049	0.13	0.033	0.212
350	0.052	0.14	0.035	0.227
375	0.056	0.15	0.038	0.244
400	0.060	0.16	0.040	0.260
425	0.063	0.17	0.043	0.276
450	0.067	0.18	0.045	0.292
475	0.071	0.19	0.048	0.309
500	0.074	0.20	0.051	0.325
525	0.078	0.21	0.053	0.341
550	0.082	0.22	0.055	0.357
575	0.086	0.23	0.058	0.374
600	0.089	0.24	0.060	0.389

Table 1: Intramuscular anesthesia dosing for rats. Volumes of medetomidine (1 mg/mL), midazolam (5 mg/mL), and fentanyl (0.05 mg/mL) required to prepare the intramuscular anesthetic mixture for rats weighing 100–600 g. The total injection volume represents the sum of the individual drug volumes.

3. Bladder Preparation

1. To allow a tension-free vesicostomy, make a 3 mm longitudinal midline skin incision using a No. 11 scalpel blade while stabilizing the skin with hooked (Adson) forceps, approximately two-thirds of the distance between the second and third teat pairs (counted from caudal to cranial). This incision corresponds to the level of the bladder dome.
2. Identify the subcutaneous abdominal muscles and the linea alba. Incise the outer fascia with the scalpel, bluntly dissect the abdominal muscle layers using micro scissors, then carefully incise the peritoneum and open the abdominal cavity using hooked micro forceps.
3. To identify the bladder within the abdominal cavity, gently elevate both sides of the abdominal muscle layers at the incision to improve visualization and locate the bladder beneath the fatty tissue. If the bladder cannot be identified, perform transurethral catheterization and repeat the localization.

4. Transurethral Catheter Placement

1. Fill the bladder through a transurethral catheter to facilitate bladder identification.
2. Apply lubricating gel to a 24G peripheral venous catheter (without stylet) and to the urethral meatus. Lift the periurethral skin with small serrated forceps and insert the catheter at approximately 110° to facilitate atraumatic placement.
3. Advance the catheter atraumatically through the urethra to the level of the pubic bone. Lower the catheter until it is parallel to the animal’s body and continue advancing it gently into the bladder. If resistance is encountered approximately 1 cm into the urethra, withdraw the catheter completely and repeat the insertion. If the catheter advances smoothly for at least 2 cm, proceed with bladder filling.
4. Confirm successful catheter placement by slowly infusing sodium chloride into the bladder without resistance. If catheter placement is unsuccessful, the syringe becomes blocked, and slight withdrawal of the catheter results in leakage of sodium chloride from the urethra.

5. Bladder Positioning

1. Fill the bladder with a maximum of 400 µL of prewarmed 0.9% NaCl. Ensure that filling proceeds smoothly without increased infusion pressure. Visually monitor bladder filling and confirm that the bladder remains centered within the abdominal cavity. Note that the physiological bladder volume depends on the animal’s age, body weight, and strain and may vary. For 12-week-old Lewis rats (LEW/Crl) weighing 200–230 g, the maximum physiological bladder volume is approximately 500 µL. Leave the transurethral catheter in place and refill the bladder if necessary.
2. Inspect the bladder and identify the planned vesicostomy site below the bladder dome on the ventral bladder wall.
3. Place two superficial stay sutures using 6-0 coated Vicryl at the 3 and 9 o’clock positions on either side of the planned incision site. Secure the suture ends with clamps to maintain bladder position.

6. Bladder Vesicostomy

1. Carefully grasp the bladder dome with hooked micro forceps and make a 3 mm longitudinal incision using a No. 11 scalpel blade. Ensure that the incision transects all layers of the bladder wall to allow immediate urine drainage.
2. Inspect the incision and, if necessary, enlarge it with micro scissors to approximately 3 mm.
3. Use the stay sutures to position the bladder for suturing. Remove the transurethral catheter.
4. Begin suturing with 6-0 coated Vicryl at the 12 o’clock position, passing the suture from the luminal side of the bladder wall through the skin while excluding the muscle layer. Place subsequent sutures at the 6, 8, 10, 2, and 4 o’clock positions until the bladder opening is completely approximated to the skin. Remove the stay sutures once the bladder is securely fixed.
5. Consider the vesicostomy complete once the bladder mucosa is fully apposed to the abdominal skin without visible gaps and the 3–5 mm vesicostomy opening can be readily intubated with a calibration stick.
6. Place four stainless-steel wire sutures approximately 4–5 mm from the stomal edge to discourage the animal from licking or chewing the surgical site. Pass each wire through the skin using a 20G needle, secure it with two simple overhand knots, and trim the wire ends to approximately 2 mm.

7. Calibration and Wound Dressing

1. Clean the surrounding skin with clean water, followed by incubation for 10 min with a mucosa-compatible antiseptic agent, such as 0.2% sodium hypochlorite solution.
2. Insert a sterile calibration stick of approximately 6–11 Charrière (1 Charrière = approximately 0.33 mm diameter) into the vesicostomy to assess stomal diameter.
3. Use a calibration stick that passes smoothly while filling the stomal lumen without causing inward invagination of the wound edges.
4. Apply topical skin adhesive generously to the shaved abdominal skin, the tissue surrounding the vesicostomy, and the stomal opening.
5. Continuously rotate the calibration stick within the vesicostomy until the adhesive has dried (approximately 10 s), preferably with assistance from a second person, to prevent closure of the vesicostomy. Representative images of the vesicostomy immediately after surgery and after healing are shown in Figure 1.

Figure 1. Representative appearance of the vesicostomy immediately after surgery and following healing. Representative images of the vesicostomy site immediately after surgery and wound dressing at day post-injury (DPI) 0 (left) and after healing at DPI 53, shortly before completion of the 56-day (2-month) follow-up period (right). The inset shows a higher-magnification view of the vesicostomy immediately after surgery. Scale bars = 1 cm (overview images) and 2 mm (inset). Please click here to view a larger version of this figure.

8. Antagonism

1. Antagonize general anesthesia (no earlier than 1 h after anesthetic administration) by administering a mixture of atipamezole (0.76 mg/kg) and flumazenil (0.2 mg/kg) subcutaneously using a 1 mL syringe fitted with a 27G needle according to Table 2.
2. To allow stress-free recovery, place the animal in its home cage before administering the antagonistic drug mixture. Monitor the animal continuously until full consciousness is regained. Keep the home cage under a heating lamp in a separate recovery room for 3 h.

Body weight (g)	Atipamezole (5 mg/mL), mL	Flumazenil (0.1 mg/mL), mL	Total volume (mL)
100	0.015	0.20	0.215
125	0.019	0.25	0.269
150	0.023	0.30	0.323
175	0.026	0.35	0.376
200	0.030	0.40	0.430
225	0.034	0.45	0.484
250	0.037	0.50	0.537
275	0.041	0.55	0.591
300	0.045	0.60	0.645
325	0.049	0.65	0.699
350	0.052	0.70	0.752
375	0.056	0.75	0.806
400	0.060	0.80	0.860
425	0.063	0.85	0.913
450	0.067	0.90	0.967
475	0.071	0.95	1.021
500	0.074	1.00	1.074
525	0.078	1.05	1.128
550	0.082	1.10	1.182
575	0.086	1.15	1.236
600	0.089	1.20	1.289

Table 2: Subcutaneous anesthetic antagonism dosing for rats. Volumes of atipamezole (5 mg/mL) and flumazenil (0.1 mg/mL) required to prepare the subcutaneous anesthetic antagonist mixture for rats weighing 100–600 g. The total injection volume represents the sum of the individual drug volumes.

9. Waste Disposal

1. Dispose of sharps (e.g., syringe needles and stainless-steel wire remnants) in an approved sharps container. Dispose of all biological and surgical waste according to institutional biosafety regulations.

10. Postoperative Management

1. Postoperative Care
   1. Administer enrofloxacin (1.5 mg, once daily, subcutaneously) and long-acting meloxicam (0.2 mg, twice daily, subcutaneously) for the first 5 days after surgery. Supplement pre-warmed 0.9% NaCl subcutaneously as required to compensate for fluid loss.
   2. Continue low-dose enrofloxacin (0.5 mg, once daily, subcutaneously) throughout the follow-up period.
   3. Resume or intensify antibiotic and/or analgesic treatment if signs of pain or infection are observed during daily examinations. In such cases, administer the same antibiotic and analgesic doses used during the initial 5-day postoperative period.
   4. Initiate a 5-day course of antibiotics at the standard dose if urine characteristics change (e.g., hematuria or cloudy urine persisting for 3 days) and/or if the condition of the skin surrounding the vesicostomy changes (e.g., reddened skin persisting for 2 days without improvement). Resume analgesic treatment during periods of hematuria and/or upon the onset of skin plaque formation, and continue treatment until the symptoms resolve.
   5. Perform vesicostomy calibration daily. Begin with a calibration stick at least one size smaller than the previously documented stomal diameter and apply lubricating gel before insertion. If the calibration stick passes smoothly, repeat the procedure with the next larger size and record the largest size that passes smoothly.
   6. Reduce the calibration frequency to every second day no earlier than 3 weeks after surgery and only if the calibration size has remained stable for at least 3 consecutive days.
   7. Once the skin adhesive begins to detach (typically 3–5 days after surgery), clean the surrounding skin daily with a mucosa-compatible antiseptic agent, such as 0.04% sodium hypochlorite solution, followed by application of an oil-based moisturizing protective film. Assess wound healing according to the wound scoring system and representative wound appearances shown in Figure 2A,B.
   8. Remove the stainless-steel wires 7 days after surgery.
2. Postoperative Monitoring
   1. Monitor all animals closely throughout the postoperative period.
   2. Examine each animal at least twice daily for signs of pain or discomfort according to the Rat Grimace Scale²⁷, abnormal swelling, bleeding, other changes at the surgical site, and excessive porphyrin discharge. Assess the vesicostomy using the wound scoring system shown in Figure 2A,B and record the calibration size as shown in Figure 2C.
   3. Maintain detailed records for each animal and regularly document the following parameters:
      1. Body weight (relative to the preoperative value)
      2. General condition (fur, eyes, whiskers, ears, and respiration)
      3. Walking behavior (e.g., stilted or unsteady gait)
      4. Urine characteristics (e.g., discoloration or strong odor)
      5. Vesicostomy calibration size
      6. Condition of the skin surrounding the vesicostomy

Figure 2. Postoperative wound healing and vesicostomy calibration during follow-up. (A) Wound score of the vesicostomy site during the 7-week follow-up period. (B) Representative images of wound scores 1–4, illustrating the progression from normal wound appearance to ulcer formation. Animals with open skin lesions were excluded according to the predefined study criteria. Scale bars = 5 mm. (C) Vesicostomy calibration size measured in Charrière during the follow-up period. Data in panels A and C are presented as mean ± standard deviation (SD). Please click here to view a larger version of this figure.

Results

A total of 18 rats underwent vesicostomy. To establish the technique, the procedure was first performed in six animals without SCI to verify its basic safety and feasibility and to evaluate postoperative management in healthy animals. Subsequently, vesicostomy was performed in combination with a thoracic contusion SCI at the T8/T9 vertebral level in 12 animals. Of these, six animals were used to further refine the postoperative wound management protocol for SCI animals. The resulting regimen was then applied to a second ...

Discussion

This protocol provides the first detailed description of vesicostomy in rodents as a method for continuous, low-pressure urinary drainage following SCI. Beyond describing the operative technique, we provide a comprehensive postoperative wound management protocol, which proved critical for achieving prolonged and successful follow-up. Because the rat is a small animal model and the objective is to create a minimal bladder opening to limit confounding influences and promote wound healing, magnifying glasses (2.5–3.5&...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% sodium chloride	Fresenius Kabi, Graz, Austria	—	Sterile saline
20G hypodermic needle	Servoprax, Wesel, Germany	REF L1 0201	Used for stainless-steel wire placement
24G peripheral venous catheter	B. Braun, Sempach, Switzerland	REF 4254074B	Used as a transurethral catheter (without stylet)
27G hypodermic needle	B. Braun, Melsungen, Germany	REF 4657705	Used for injections
6-0 coated Vicryl suture	Ethicon, Puerto Rico, USA	REF V134H	Used for bladder fixation
Actimaris Wound Irrigation Solution forte	ActiMaris, Appenzell, Switzerland	REF 30350	0.2% sodium hypochlorite solution
Actimaris Wound Irrigation Solution sensitive	ActiMaris, Appenzell, Switzerland	REF 30300	0.04% sodium hypochlorite solution
Atipamezole (Antisedan 5 mg/mL)	Orion Pharma, Espoo, Finland	No catalog number available	Anesthetic antagonist
Bladder calibration sticks (6–11 Charrière)	No manufacturer available	No catalog number available	Used for vesicostomy calibration
Carbon steel scalpel blade No. 11	Aesculap, Tuttlingen, Germany	REF BB511	Scalpel blade
Dexpanthenol ophthalmic ointment (2%)	Fresenius Kabi, Graz, Austria	REF RA4302436	Eye ointment
Enrofloxacin (Baytril 25 mg/mL)	Elanco, Monheim, Germany	No catalog number available	Antibiotic
Female Lewis rats (12 weeks old)	Charles River Laboratories, Sulzfeld, Germany	—	Experimental animals
Fentanyl (50 µg/mL)	hameln pharma, Hameln, Germany	No catalog number available	Anesthetic
Flumazenil (0.1 mg/mL)	Fresenius Kabi, Graz, Austria	No catalog number available	Anesthetic antagonist
Heating pad	Lenric, Littlehampton, UK	No catalog number available	Used to maintain body temperature
Hooked (Adson) forceps	Fine Science Tools, Heidelberg, Germany	11019-12	Tissue handling
Hooked micro forceps	Medicon, Tuttlingen, Germany	13.71.62	Microsurgical forceps
Long-acting meloxicam (Metacam 5 mg/mL)	Boehringer Ingelheim, Ingelheim am Rhein, Germany	No catalog number available	Analgesic
Lubricating gel (Cathejell C)	Montavit, Absam, Austria	No catalog number available	Used for transurethral catheterization and vesicostomy calibration
Medetomidine (Sedator 1 mg/mL)	Dechra, Aulendorf, Germany	No catalog number available	Anesthetic
Microscope or surgical magnification glasses (2.5–3.5×)	Evident Europe GmbH, Vienna, Austria	Olympus SZ61	Used during microsurgery
Micro scissors (Bonn scissors)	Fine Science Tools, Heidelberg, Germany	14084-08	Microsurgical scissors
Mosquito hemostat	Fine Science Tools, Heidelberg, Germany	13009-12	Used to secure stay sutures
Oil-based moisturizing protective film	Panin, Rovigo, Italy	No catalog number available	Postoperative skin protection
Povidone-iodine solution (7.5%)	B. Braun, Melsungen, Germany	No catalog number available	Skin antiseptic
Scalpel handle No. 3	Fine Science Tools, Heidelberg, Germany	10003-12	Compatible with No. 11 scalpel blade
Small serrated forceps	Fine Science Tools, Heidelberg, Germany	11050-10	Used for periurethral skin manipulation
Stainless-steel wire	Vömel, Kronberg, Germany	No catalog number available	Used for protective wire sutures
Syringe, 1 mL	Becton Dickinson, Vienna, Austria	REF 303172	Used for anesthetic and antagonist administration
Topical skin adhesive	Ethicon, Puerto Rico, USA	REF AHVM12	Skin adhesive