1Children's Hospital Boston, Harvard Medical School, 2Tufts University
Tu, D. D., Seth, A., Gil, E. S., Kaplan, D. L., Mauney, J. R., Estrada Jr., C. R. Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models. J. Vis. Exp. (66), e3981, doi:10.3791/3981 (2012).
Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and expels it with a precisely orchestrated contraction. A number of congenital and acquired urological anomalies including posterior urethral valves, benign prostatic hyperplasia, and neurogenic bladder secondary to spina bifida/spinal cord injury can result in pathologic tissue remodeling leading to impaired compliance and reduced capacity1. Functional or anatomical obstruction of the urinary tract is frequently associated with these conditions, and can lead to urinary incontinence and kidney damage from increased storage and voiding pressures2. Surgical implantation of gastrointestinal segments to expand organ capacity and reduce intravesical pressures represents the primary surgical treatment option for these disorders when medical management fails3. However, this approach is hampered by the limitation of available donor tissue, and is associated with significant complications including chronic urinary tract infection, metabolic perturbation, urinary stone formation, and secondary malignancy4,5.
Current research in bladder tissue engineering is heavily focused on identifying biomaterial configurations which can support regeneration of tissues at defect sites. Conventional 3-D scaffolds derived from natural and synthetic polymers such as small intestinal submucosa and poly-glycolic acid have shown some short-term success in supporting urothelial and smooth muscle regeneration as well as facilitating increased organ storage capacity in both animal models and in the clinic6,7. However, deficiencies in scaffold mechanical integrity and biocompatibility often result in deleterious fibrosis8, graft contracture9, and calcification10, thus increasing the risk of implant failure and need for secondary surgical procedures. In addition, restoration of normal voiding characteristics utilizing standard biomaterial constructs for augmentation cystoplasty has yet to be achieved, and therefore research and development of novel matrices which can fulfill this role is needed.
In order to successfully develop and evaluate optimal biomaterials for clinical bladder augmentation, efficacy research must first be performed in standardized animal models using detailed surgical methods and functional outcome assessments. We have previously reported the use of a bladder augmentation model in mice to determine the potential of silk fibroin-based scaffolds to mediate tissue regeneration and functional voiding characteristics.11,12 Cystometric analyses of this model have shown that variations in structural and mechanical implant properties can influence the resulting urodynamic features of the tissue engineered bladders11,12. Positive correlations between the degree of matrix-mediated tissue regeneration determined histologically and functional compliance and capacity evaluated by cystometry were demonstrated in this model11,12. These results therefore suggest that functional evaluations of biomaterial configurations in rodent bladder augmentation systems may be a useful format for assessing scaffold properties and establishing in vivo feasibility prior to large animal studies and clinical deployment. In the current study, we will present various surgical stages of bladder augmentation in both mice and rats using silk scaffolds and demonstrate techniques for awake and anesthetized cystometry.
1. Surgical Preparation and Anesthesia
2. Incision and Exposure of the Bladder
3. Anastomosis of the Scaffold
4. Incisional Closure
The steps for cystostomy catheter placement for cystometric analysis are as follows:
5. Tunneling the Cystostomy Catheter
6. Placing the Cystostomy Tube
7. Testing the Catheter and Closing the Abdominal Incision
8. Closing the Dorsal Incision and Securing the Catheter (For rats)
8.* Closing the Dorsal Incision and Securing the Catheter (For mice or rats)
9. Representative Results - Surgical Methods
The reconstructed bladder should be as water-tight as possible to avoid complications related to a significant urinary leak (Figure 3). Pain or discomfort usually manifest as shivering or scratching and gnawing at the abdominal incision. This can be managed with daily subcutaneous injections of a non-steroidal anti-inflammatory such as meloxicam (0.5-1.0 mg/kg subcutaneous). Typically, the animals only require the injections for the first 3 days post-operatively. This can be supplemented with an opioid, such as buprenorphine (0.05-0.1 mg/kg subcutaneous every 8-12 hr) as needed. The animals should be monitored 3 times daily for the first 3 post-operatively days, twice daily for post-operative days 3-5 and then daily thereafter to evaluate for pain, signs of infection, adequate wound healing, activity, grooming, and skin turgor. Antibiotics (Baytril, 5mg/kg subcutaneous every 24 hr in a volume not exceeding 0.1 mL) are given for the first 72 hr following surgery, as surgical prophylaxis against infection. Signs of normal recovery are normal ambulation and activity levels, appropriate feeding and drinking, absence of pain or distress (no vocalization) and normal socialization with cagemates. A recovery time of at least 5-7 days should be given before cystometric analysis, to allow for bladder healing and decreased inflammation which could potentially affect the results.
10. Awake Cystometric Analysis
11. Unconscious Cystometric Analysis (No suprapubic catheter)
12. Representative Results - Cystometric Analyses
Urodynamic tracings can then be analyzed to derive parameters such as voided volumes, compliance, peak voiding pressures, inter contraction interval, micturition cycle time and post void residual volumes.
Cystometrogram can be divided into a filling and a voiding phase. A normal filling phase is the portion of the micturition cycle in which bladder fills with very little change in intravesical pressure. A normal voiding phase of the tracing consists of a steady rise in the intravesical pressure corresponding to the detrusor contraction. The highest pressure reached during the voiding phase of the tracing is termed the peak voiding pressure. A high peak voiding pressure could suggest an obstructive voiding pattern, a hypercontractile bladder or a kink in the SP catheter. Compliance can be calculated by acquiring the ratio of the volume instilled during the filling phase and the change in pressure (compliance = ΔV/ΔP). A hypocompliant bladder is one that is unable to accommodate adequate urinary volumes at low pressures. The intercontraction interval can be calculated by analyzing the time between two contractions as seen on the cystometrogram. A short intercontraction interval is suggestive of an irritable bladder. The micturition cycle time refers to the time it takes for an entire filling and voiding phase to complete and can be easily ascertained by analyzing the tracing. At the conclusion of the cystometry, post-void residual (PVR) can be obtained. This is done by aspirating the suprapubic catheter upon the completion of a detrusor contraction. These parameters help the investigator objectively study the bladder dynamics as the bladder fills and empties.
Figure 1. Photograph of the abdominal incision and extrusion of the bladder.
Figure 2. Bladder incision with exposure of the bladder lumen.
Figure 3. Integration of the implant onto the bladder wall.
Figure 4. Photograph of the closed incision.
Figure 5. Flared end of the PE-50 tubing.
Figure 6. PE-50 tubing (catheter) through the dorsal incision.
Figure 7. Pursestring suture.
Figure 8. Securing the catheter to the bladder.
Figure 9. Secured catheter hub.
Figure 10. Coiled tubing in subcutaneous pouch.
Figure 11. Dorsal incisional closure.
Figure 12. Example cystometric set-up.
Figure 13. Representative cystometry tracing.
Cystometric evaluations of biomaterial configurations following implantation and bladder augmentation in small animal models represents an important validation step in identifying optimal structural and mechanical characteristics of matrix designs for use in clinical situations. In this study, we describe surgical methods for performing bladder augmentation in mice and rats as well as cystometric techniques to determine urodynamic properties of engineered organs for functional assessments. We have utilized these techniques in multiple experiments involving both mice and rats, with each experiment consisting of 30+ rodents without significant issues. Our research laboratory is a diverse conglomerate of basic scientists and physician surgeons, and surgeons with at least 5-6 years of post-graduate surgical training performed the procedural aspects of these experiments.
Regardless of the type of biomaterial used, the major difference between augmenting the bladder in rats versus mice is the size of the bladder. Due to smaller bladder size, dissection and incorporation of the biomaterial is more technically difficult in the mouse. To aid in visualization, a surgical microscope can be used. Since the size of the bladder in rats is larger, it is more amenable to situations where more than one procedure has to be performed on the bladder (e.g. augmentation and placement of cystostomy catheter). Additionally, the protocol above describes use of PE-50 tubing for the rat 13, however, even larger size catheters, up to PE-100 have been used, especially for long-term studies 14. In mice, a smaller caliber such as PE-10 tubing can be utilized 15,16, but it should be kept in mind that smaller, more pliable tubes may not transmit pressure changes to the transducer accurately. Also, the alternative method of securing the catheter on the dorsum (step 8* above) is done in mice due to their smaller body size and the blunt tip needle and IV cap are too cumbersome. The disadvantage of this is the need for anesthesia to extract the end of the catheter in the subcutaneous pouch prior to cystometry.
Studies have shown that in the initial first days (0-4 days) after placement of the catheters, cystometry revealed high bladder pressures and overactivity with low voiding volumes. These findings appeared to stabilize around the sixth to seventh day14,17 and therefore, is probably the ideal timing for cystometric evaluation. However, most reports in the literature perform cystometry within the first 3 days of catheterization 18, and this accounts for the wide variation in the above parameters relative to time. Leaving the suprapubic catheter for a duration longer than 3 days carries with it morbidities such as risk of stones, dislodgement, infection, hematuria and occlusion of catheter with debris.
Different infusion rates during cystometry have been described from 1-3mL/hr for mice15,16 and 10-11mL/hr for rats 13,19,20. Supraphysiologic infusion rates can cause falsely elevated pressures 14. We use an infusion rate of 12.5 μL/min (0.75 mL/hr) for mice and 100 μL/min (6 mL/hr) for rats in our setup, but lower rates can also be utilized. The temperature of the physiologic saline should be at least room temperature, although warm (37°) saline is more optimal in order to avoid bladder overactivity provoked with instilling cold solution. In awake cystometry, it is crucial to allow for stabilization of the voiding pattern as the animal becomes adjusted to the cage, which in our experience requires a period of ~10-20 minutes. Following this, regular micturition cycles can be recorded for 45-120 minutes or at minimum 3-4 voiding cycles. The animal should be observed in real-time since the animal is freely moving and complications such as twisting or kinking of the catheter can alter cystometric analysis. Limiting environmental noise during cystometry is desired to decrease animal movement and subsequent artifacts. Unconscious cystometry does not have the attendant problems as awake cystometry, but multiple anesthetics have been shown to inhibit spontaneous bladder contractions. This inhibition corresponds directly to the expected duration of action of the anesthetic drugs, i.e. when the anesthetic effect subsides, spontaneous contractions resume 14. Moreover, pressures measured when the bladder overflowed, were statistically greater in anesthetized rats, both alive and post-mortem, indicating an effect on the passive compliance properties of the bladder wall. This effect is seen with pentobarbital21, ketamine, and chloralose IM/IP, in addition to inhaled halothane and intrathecal nesacaine 14. A more extensive study of various anesthetics confirm this finding with suppression of the micturition reflex for both inhalational (isoflurane and methoxyflurane) and barbiturate (pentobarbital and thiobutabarbital) anesthetics under moderate anesthesia levels17. This effect was observed with even light or sedative levels of anesthesia with medications such as fentanyl-droperidol and ketamine-diazepam, and as in the previous study, as the anesthesia effect subsided, so did the inhibition17. For this procedure, urethane intraperitoneal injections can be used since it has been demonstrated that reflex micturition is preserved while also allowing for adequate anesthesia17,22. Moreover, no effect is observed with respect to micturition pressures23. Suprapubic catheter placement for cystometry is described here, since intraurethral catheterization has been shown to have higher bladder pressure curves and lower flow rates consistent with relative bladder outlet obstruction 24. Moreover, intraurethral catheterization is only feasible in anesthetized animals, and even then, catheterization can be difficult, especially in male rodents and mice.
In conclusion, the choice of which model to use for bladder augmentation and/or cystometric analysis is dependent on the goals of the specific study. From a technical standpoint the rat model clearly holds the advantage for the reasons discussed above. However, the mouse model can be used in studies evaluating the roles of specific gene-encoded end products in diseases of the urinary tract, due to their susceptibility for genetic manipulation. This is not generally feasible in the rat.
Awake cystometry most accurately mimics the normal physiologic state in which these animals undergo their micturition cycles, and so, is likely to give a more reliable physiologic determination of bladder function. Moreover, the confounding variable of direct effects of anesthetics on bladder function is avoided.
No conflicts of interest declared.
These studies were funded, in part, by the Children's Hospital Boston Urology Endowment Revenue Fund and the National Institutes of Health grants NIBIB P41-EB002520 (Kaplan); NIDDK T32-DK60442 (Freeman); NIDDK 1K99-DK083616 (Mauney). We acknowledge Dr. Peter Zvara from the University of Vermont for assistance in establishing the technique for cystostomy tube placement and cystometry.
|Shaving shears||Preparation of rat/mouse for surgery|
|Sterile drapes, betadine, 70% ethanol, sterile gauze||Preparation of sterile surgical field|
|Scalpel blade||Skin incision|
|forceps with teeth||Manipulating skin|
|Fine forceps||Atraumatic (no teeth), no serrations or with fine serrations to manipulate|
|Small needle driver||Sharp tissue dissection|
|Metzenbaum scissors||Bldder incision|
|Tenotomy scissors||For retraction sutures and to develop subcutaneous tunnel (cystostomy catheter)|
|Small curved clamps||Subcutaneous tunnel (cystostomy catheter)|
|6-0 polypropylene sutures||Bladder stay sutures and pursestring suture|
|7-0 polyglactin suture||Anastomosis of scaffold to bladder|
|4-0 polyglactin suture||Closure of muscle/skin|
|3-0 or 4-0 Silk suture||Securing catheter tip to skin|
|Needles and syringes:|
|18 Gauge needle||Piercing the bladder for cystostomy catheter|
|25 and 30 Gauge needles||Testing bladder for leakage|
|1 mL saline filled syringe|
|22 Gauge blunt tip needle|
|Lighter||Flaring PE-50 tubing|
|Small curved clamp||Developing subcutaneous tunnel|
|MLT844 ADInstruments data capture and LabChart software||Pressure data acquisition|
|Harvard 22 syringe pump (Harvard Apparatus, Holliston, MA)||Fluid infusion pump|
|Anesthetics (Unconscious cystometry):|
|Isoflurane||Induction/maintenance of general anesthesia|
|Bupivicaine or equivalent||Local anesthesia|