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.

Surgical Methods
1. Surgical Preparation and Anesthesia
<ol>
	<li>Set up the sterile surgical field with the necessary surgical instruments: shaving shears, forceps with teeth, fine atraumatic forceps, fine needle driver, gauze, Metzenbaum scissors, tenotomy scissors, scalpel blade, 30 gauge hypodermic needle, saline filled 1 mL syringe, four 6-0 polypropylene sutures, 7-0 polyglactin suture, 4-0 polyglactin suture.</li>
	<li>Anesthetize the animal with isoflurane inhalation in the induction chamber. Confirm complete induction of the animal before transferring to the surgical field. Ensure that the inhalational anesthesia tube is in appropriate position to have continuous anesthesia.</li>
	<li>Place the animal supine on the sterile drape. 
	[For cystometric analysis, see section below in tunneling the cystostomy catheter.]</li>
	<li>Use the shaving shears to remove the fur from the lower abdomen.</li>
	<li>Prep the abdomen with betadine and 70% ethanol.</li>
	<li>Prior to incision, an analgesic such as buprenorphine (0.05-0.1 mg/kg) can be injected subcutaneously for perioperative pain control.</li>
</ol>
2. Incision and Exposure of the Bladder
<ol>
	<li>Make a 1-2 cm (dependant on size of animal and whether rat or mouse) lower midline incision with the scalpel through the skin. Deepen the incision at the lower portion of the incision through the rectus muscle taking care not to injure the underlying bowel or bladder.</li>
	<li>Using toothed forceps, elevate the rectus muscle and dissect free the posterior surface of the muscle with fine Metzenbaum scissors.</li>
	<li>Incise the remainder of the muscle in the midline for the entire length of your skin incision.</li>
	<li>Deliver the bladder through the incisional wound (Figure 1). The bladder is usually the most dependent organ in the pelvis. (In the male, the prostate is actually more dependent and is larger than the decompressed bladder.)</li>
	<li>Place one stay suture through the posterior wall of the bladder, and then another through the anterior wall of the bladder using 6-0 polypropylene suture. Place additional sutures laterally. Do not tie these sutures. When the sutures are held taut, the bladder will have a square configuration measuring approximately 1 cm2 (Figure 2). Be careful not to have too much tension on these sutures as they can easily be pulled through the bladder tissue.</li>
	<li>Incise the bladder longitudinally through the anterior bladder wall (just inferior to the dome of the bladder) in the midline for approximately 1 cm (1.5-2 cm in the rat bladder).</li>
</ol>
3. Anastomosis of the Scaffold
<ol>
	<li>Using fine scissors, trim the silk scaffold to the approximate area of the bladder defect.</li>
	<li>Using 7-0 polyglactin suture, start at one corner of the scaffold and suture it to the bladder in a continuous, running fashion to create a watertight seal all the way around the defect (Figure 3).</li>
	<li>Test the integrity of the anastomosis by filling the bladder with sterile saline by instilling it through the wall of the bladder with a 30 gauge hypodermic needle. If a leak is found, this can be closed with an additional interrupted 7-0 polyglactin suture to close the gap.</li>
	<li>Reduce the reconstructed bladder back into the abdomen.</li>
</ol>
4. Incisional Closure
<ol>
	<li>Prior to closure of the abdominal wall, inject the rectus muscle and subcutaneous tissue with bupivicaine for local anesthesia (&lt; 3 mg/kg of 0.25%).</li>
	<li>Reapproximate the rectus muscle with a continuous, running 4-0 polyglactin suture.</li>
	<li>Close the skin with a continuous, running 4-0 polyglactin suture.</li>
	<li>Clean and dry the incision (Figure 4).</li>
	<li>Transfer the animal into a warm, clean cage for awakening from the anesthesia.</li>
</ol>
The steps for cystostomy catheter placement for cystometric analysis are as follows:
5. Tunneling the Cystostomy Catheter
<ol>
	<li>Set up the sterile surgical field with the necessary surgical instruments: shaving shears, forceps with teeth, fine atraumatic forceps, fine needle driver, gauze, Metzenbaum scissors, tenotomy scissors, small curved clamp, scalpel blade, 6-0 polypropylene, 4-0 polyglactin suture, 3-0 silk suture (4-0 silk suture for mice), 18G needle, 22G blunt tip needle, 25G needle, 1 mL saline filled syringe, polyethylene tubing 50 (PE-50) cut for a length of ~10 cm.</li>
	<li>Flare the end of the PE-50 tubing by gently exposing it to a flame. Be careful not to melt the end off or occlude the lumen (this can be verified by injecting saline through a 25G needle connected to the "non-flared" end and ensuring flow). This serves as an anchor to maintain the tube within the bladder (Figure 5).</li>
	<li>Anesthetize the animal as above in step 1.2.</li>
	<li>Use the shaving shears to remove the fur from both the dorsum of the animal between the scapula and the lower abdomen on the ventrum.</li>
	<li>Prep the areas with betadine and 70% ethanol. Place the animal prone on the drape.</li>
	<li>Make a 1 cm incision on the dorsum between the scapula. Using the Metzenbaum scissors, develop a plane between the skin and the underlying muscle by placing the tips of the scissors in the plane and spreading them to create a tunnel around to the ventral abdomen.</li>
	<li>Reposition the animal supine. Make your abdominal incision and expose the bladder as above in steps 2.1-2.4. Reduce the bladder back into the abdomen.</li>
	<li>Place a small clamp into your subcutaneous tunnel created in step 5.6 starting from your dorsal skin incision. Using your fingers to protect the intra-abdominal contents, pierce through the abdominal wall with the tips of the clamp into the abdomen.</li>
	<li>Grasp the smooth end of the PE-50 tubing with the clamp and pull it back through the dorsal incision. Ensure that the bulbed end is not pulled past the abdominal wall (Figure 6).</li>
</ol>
6. Placing the Cystostomy Tube
<ol>
	<li>Deliver the bladder through the incision. From this point on, the bladder should be handled with fine forceps to prevent trauma to the bladder that could cause damage or inflammation that can skew your cystometric results or result in additional discomfort for the animal post-operatively.</li>
	<li>Make note of the proposed site for the cystostomy tube. It should be placed at the dome of the bladder (superior to the augment segment). This will prevent kinking or occlusion of the tube.</li>
	<li>Using the 6-0 polypropylene suture, place a pursestring stitch at the dome of the bladder in the following manner: place the first throw through the wall of the bladder longitudinally, lateral to the proposed site for the cystostomy tube. Leave a small clamp on the loose end so that the suture is not inadvertently pulled all the way through. Place the next throw in a transverse direction, starting first going into the bladder wall slightly lateral to the exit of your first throw.</li>
	<li>Do not pull the suture taut. Place your next suture parallel to your first (longitudinally) going into the bladder just cephalad to the exit site of your last, transverse suture. The fourth throw will start lateral to the exit of the last stitch and end next to the entrance for your very first throw. Done correctly, this forms a circumferential square around the proposed catheter site (Figure 7).</li>
	<li>Using an 18G needle, pierce the bladder wall in the center of the pursestring suture. Be careful not to pierce too deeply (just enough to be intraluminal). Place the tips of your fine forceps in the opening and gently spread to widen the hole.</li>
	<li>Insert the bulbed end of the catheter into the defect in the bladder until it is intraluminal. Pull the pursestring suture tight around the catheter and tie it down. This should cinch the bladder wall around the catheter keeping it in place (Figure 8).</li>
	<li>Take one end of the suture and wrap it around the catheter once and tie this down to further secure the catheter.</li>
</ol>
7. Testing the Catheter and Closing the Abdominal Incision
<ol>
	<li>With a 1 mL syringe and 25G needle, insert the needle into the tubing and slowly inject saline to distend the bladder. Observe for leaking around the catheter. Once you see leaking from the urethra, aspirate the saline to decompress the bladder again.</li>
	<li>Close the abdominal incision as above in steps 4.1-4.4.</li>
</ol>
8. Closing the Dorsal Incision and Securing the Catheter (For rats)
<ol>
	<li>Reposition the animal prone.</li>
	<li>Cut the catheter tubing at the level of the skin with the scissors. Insert a 22G blunt tip needle into the tubing.</li>
	<li>Close the skin over the tubing with 4-0 polyglactin in a running fashion. Leave the needle hub extruding from the skin.</li>
	<li>Place an intravenous line cap on the blunt needle. Using 3-0 silk suture, secure the catheter tip to the skin (Figure 9).</li>
	<li>Clean the incision. Transfer the rat into a warm, clean cage for awakening from the anesthesia.</li>
</ol>
8.* Closing the Dorsal Incision and Securing the Catheter (For mice or rats)
<ol>
	<li>*Occlude the distal end of the catheter by kinking it or using a flame to melt the end.</li>
	<li>*Coil the end of the tubing and leave it in the subcutaneous pouch on the dorsum of the animal (do NOT cut tubing to shorten it) (Figure 10).</li>
	<li>*Close the skin over the tubing with 4-0 polyglactin in a running fashion (Figure 11).</li>
	<li>*On the day of cystometry, prepare the dorsal incision with betadine and 70% ethanol. Open the dorsal incision under anesthesia and remove the coiled tubing from the subcutaneous pouch. Close the incision. Awaken the animal from anesthesia and perform cystometry when it is fully awake.</li>
</ol>
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.
Cystometric Analyses
10. Awake Cystometric Analysis
<ol>
	<li>Setup described with MLT844 ADInstruments with data capture and analysis with LabChart v6 (ADInstruments) and infusion with a Harvard 22 syringe pump (Harvard Apparatus, Holliston, MA), although other comparable systems are available (Figure 12).</li>
	<li>Calibrate both volume and pressure based on the specifications of the cystometric system used.</li>
	<li>Place the animals in metabolic cages (cages with a wire mesh floor) which are suspended over a scale. The scale is connected to a transducer.</li>
	<li>Purge the system of any air bubbles and ensure continuous flow from the infusion pump.</li>
	<li>Connect the data capture system to a computer and observe for data tracings. Adjust the scale accordingly. Bladder pressure and urine volume will be continuously recorded.</li>
	<li>Access the suprapubic catheters with a 27G needle connected via a T-tube to the pressure transducer and the infusion pump. Begin the infusion of physiologic saline at 12.5 &mu;L/min for the mouse and 100 &mu;L/min for the rat.</li>
	<li>Allow the voiding pattern tracing to stabilize (bladder pressure rise, followed by a void). This usually takes approximately 10-20 minutes. Record the micturition cycles for 45-120 minutes or at least 3-4 voiding cycles.</li>
	<li>Observe entire procedure in real-time to troubleshoot for complications that will lead to artifact (i.e. catheter kinking, obstruction, etc; see below discussion).</li>
	<li>Stop the infusion, disconnect the catheter from the system, and return the animal to its cage.</li>
</ol>
11. Unconscious Cystometric Analysis (No suprapubic catheter)
<ol>
	<li>Anesthetize the animal with urethane (1-2 g/kg) intraperitoneal injection (IP).</li>
	<li>Expose the bladder as above in steps 1.3-2.4.</li>
	<li>Calibrate the system as in step 9.2. Prepare the system as in step 9.4-9.5.</li>
	<li>Insert a 27G needle connected via a T-tube to the pressure transducer and infusion pump into the lateral aspect of the bladder.</li>
	<li>Record the micturition cycles for 45-90 minutes.</li>
	<li>Stop the infusion, remove the needle from the bladder and euthanize the animal.</li>
</ol>
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 = &Delta;V/&Delta;P). 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.
<img alt="Figure 1" src="/files/ftp_upload/3981/3981fig1.jpg" /> 
Figure 1. Photograph of the abdominal incision and extrusion of the bladder.
<img alt="Figure 2" src="/files/ftp_upload/3981/3981fig2.jpg" /> 
Figure 2. Bladder incision with exposure of the bladder lumen.
<img alt="Figure 3" src="/files/ftp_upload/3981/3981fig3.jpg" /> 
Figure 3. Integration of the implant onto the bladder wall.
<img alt="Figure 4" src="/files/ftp_upload/3981/3981fig4.jpg" /> 
Figure 4. Photograph of the closed incision.
<img alt="Figure 5" src="/files/ftp_upload/3981/3981fig5.jpg" /> 
Figure 5. Flared end of the PE-50 tubing.
<img alt="Figure 6" src="/files/ftp_upload/3981/3981fig6.jpg" /> 
Figure 6. PE-50 tubing (catheter) through the dorsal incision.
<img alt="Figure 7" src="/files/ftp_upload/3981/3981fig7.jpg" /> 
Figure 7. Pursestring suture.
<img alt="Figure 8" src="/files/ftp_upload/3981/3981fig8.jpg" /> 
Figure 8. Securing the catheter to the bladder.
<img alt="Figure 9" src="/files/ftp_upload/3981/3981fig9.jpg" /> 
Figure 9. Secured catheter hub.
<img alt="Figure 10" src="/files/ftp_upload/3981/3981fig10.jpg" /> 
Figure 10. Coiled tubing in subcutaneous pouch.
<img alt="Figure 11" src="/files/ftp_upload/3981/3981fig11.jpg" /> 
Figure 11. Dorsal incisional closure.
<img alt="Figure 12" src="/files/ftp_upload/3981/3981fig12.jpg" /> 
Figure 12. Example cystometric set-up.
<img alt="Figure 13" src="/files/ftp_upload/3981/3981fig13.jpg" /> 
Figure 13. Representative cystometry tracing.

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No conflicts of interest declared.

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 &mu;L/min (0.75 mL/hr) for mice and 100 &mu;L/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&deg;) 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.

<table border="1">
 <tbody>
 <tr>
 <td>Materials:</td>
 <td>Description/Use:</td>
 </tr>
 <tr>
 <td>Shaving shears</td>
 <td>Preparation of rat/mouse for surgery</td>
 </tr>
 <tr>
 <td>Sterile drapes, betadine, 70% ethanol, sterile gauze</td>
 <td>Preparation of sterile surgical field</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Instruments:</td>
 </tr>
 <tr>
 <td>Scalpel blade</td>
 <td>Skin incision</td>
 </tr>
 <tr>
 <td>forceps with teeth</td>
 <td>Manipulating skin</td>
 </tr>
 <tr>
 <td>Fine forceps</td>
 <td>Atraumatic (no teeth), no serrations or with fine serrations to manipulate</td>
 </tr>
 <tr>
 <td>Small needle driver</td>
 <td>Sharp tissue dissection</td>
 </tr>
 <tr>
 <td>Metzenbaum scissors</td>
 <td>Bldder incision</td>
 </tr>
 <tr>
 <td>Tenotomy scissors</td>
 <td>For retraction sutures and to develop subcutaneous tunnel (cystostomy catheter)</td>
 </tr>
 <tr>
 <td>Small curved clamps</td>
 <td>Subcutaneous tunnel (cystostomy catheter)</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Sutures:</td>
 </tr>
 <tr>
 <td>6-0 polypropylene sutures</td>
 <td>Bladder stay sutures and pursestring suture</td>
 </tr>
 <tr>
 <td>7-0 polyglactin suture</td>
 <td>Anastomosis of scaffold to bladder</td>
 </tr>
 <tr>
 <td>4-0 polyglactin suture</td>
 <td>Closure of muscle/skin</td>
 </tr>
 <tr>
 <td>3-0 or 4-0 Silk suture</td>
 <td>Securing catheter tip to skin</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Needles and syringes:</td>
 </tr>
 <tr>
 <td>18 Gauge needle</td>
 <td>Piercing the bladder for cystostomy catheter</td>
 </tr>
 <tr>
 <td>25 and 30 Gauge needles</td>
 <td>Testing bladder for leakage</td>
 </tr>
 <tr>
 <td>1 mL saline filled syringe</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>22 Gauge blunt tip needle</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Cystostomy catheter:</td>
 </tr>
 <tr>
 <td>PE-50 tubing</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Lighter</td>
 <td>Flaring PE-50 tubing</td>
 </tr>
 <tr>
 <td>Small curved clamp</td>
 <td>Developing subcutaneous tunnel</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Cystometry:</td>
 </tr>
 <tr>
 <td>MLT844 ADInstruments data capture and LabChart software</td>
 <td>Pressure data acquisition</td>
 </tr>
 <tr>
 <td>Harvard 22 syringe pump (Harvard Apparatus, Holliston, MA)</td>
 <td>Fluid infusion pump</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td colspan="2">Anesthetics (Unconscious cystometry):</td>
 </tr>
 <tr>
 <td>Isoflurane</td>
 <td>Induction/maintenance of general anesthesia</td>
 </tr>
 <tr>
 <td>Urethane</td>
 <td>Unconconscious cystometry</td>
 </tr>
 <tr>
 <td>Bupivicaine or equivalent</td>
 <td>Local anesthesia</td>
 </tr>
 <tr>
 <td>Meloxicam</td>
 <td>Post-operative analgesia</td>
 </tr>
 <tr>
 <td>Buprenorphine</td>
 <td>Post-operative analgesia</td>
 </tr>
 </tbody>
</table>

evaluation of biomaterials for bladder augmentation using cystometric analyses in various rodent models

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.

Watch this Scientific Journal Video about Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models at JoVE.com

Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models

Surgical stages of bladder augmentation are described using 3-D scaffolds in murine and rat models. To test the efficacy of biomaterial configurations for use in bladder augmentation, techniques for both awake and anesthetized cystometry are presented.

Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and ...

evaluation-biomaterials-for-bladder-augmentation-using-cystometric

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Bioengineering

19.1K Views.  Harvard Medical School. Surgical stages of bladder augmentation are described using 3-D scaffolds in murine and rat models. To test the efficacy of biomaterial configurations for use in bladder augmentation, techniques for both awake and anesthetized cystometry are presented.

Video: Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models

Fabrication of Micro-tissues using Modules of Collagen Gel Containing Cells

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized tissues.  The modules can be made of collagen as well as other gelable or crosslinkable materials.  They are approximately 2 mm in length and 0.7 mm in diameter upon fabrication but shrink in size with embedded cells or when the modules are coated with endothelial cells.  The modules individually are small enough that the embedded cells are within the diffusion limit of oxygen and other nutrients but modules can be packed together to form larger tissues that are perfusable.  These tissues are modular in construction because different cell types can be embedded in or coated on the modules before they are packed together to form complex tissues.  There are three main steps to making the modules: (1) neutralizing the collagen and embedding cells in it, (2) gelling the collagen in the tube and cutting the modules and (3) coating the modules with endothelial cells.

Creation of micro-tissues using cylindrical collagen gels, called modules, that contain embedded cells and which surface is coated with endothelial cells.

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized ...

Polymer Microarrays for High Throughput Discovery of Biomaterials

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a combinatorial library of materials to be screened in a parallel, high throughput format1. Herein is described the formation and characterization of a polymer microarray using an on-chip photopolymerization technique 2. This involves mixing monomers at varied ratios to produce a library of monomer solutions, transferring the solution to a glass slide format using a robotic printing device and curing with UV irradiation. This format is readily amenable to many biological assays, including stem cell attachment and proliferation, cell sorting and low bacterial adhesion, allowing the ready identification of 'hit' materials that fulfill a specific biological criterion3-5. Furthermore, the use of high throughput surface characterization (HTSC) allows the biological performance to be correlated with physio-chemical properties, hence elucidating the biological-material interaction6. HTSC makes use of water contact angle (WCA) measurements, atomic force microscopy (AFM), X-ray photoelectron spectroscopy (XPS) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). In particular, ToF-SIMS provides a chemically rich analysis of the sample that can be used to correlate the cell response with a molecular moiety. In some cases, the biological performance can be predicted from the ToF-SIMS spectra, demonstrating the chemical dependence of a biological-material interaction, and informing the development of hit materials5,3.

A description of the formation of a polymer microarray using an on-chip photopolymerization technique. The high throughput surface characterization using atomic force microscopy, water contact angle measurements, X-ray photoelectron spectroscopy and time of flight secondary ion mass spectrometry and a cell attachment assay is also described.

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a ...

A Full Skin Defect Model to Evaluate Vascularization of Biomaterials In Vivo

Insufficient vascularization is considered to be one of the main factors limiting the clinical success of tissue-engineered constructs. In order to evaluate new strategies that aim at improving vascularization, reliable methods are required to make the in-growth of new blood vessels into bio-artificial scaffolds visible and quantify the results. Over the past couple of years, our group has introduced a full skin defect model that enables the direct visualization of blood vessels by transillumination and provides the possibility of quantification through digital segmentation. In this model, one surgically creates full skin defects in the back of mice and replaces them with the material tested. Molecules or cells of interest can also be incorporated in such materials to study their potential effect. After an observation time of one&#8217;s own choice, materials are explanted for evaluation. Bilateral wounds provide the possibility of making internal comparisons that minimize artifacts among individuals as well as of decreasing the number of animals needed for the study. In comparison to other approaches, our method offers a simple, reliable and cost effective analysis. We have implemented this model as a routine tool to perform high-resolution screening when testing vascularization of different biomaterials and bio-activation approaches.

A Full Skin Defect Model to Evaluate Vascularization of Biomaterials In Vivo

Vascularization is key to approaches in successful tissue engineering. Therefore, reliable technologies are required to evaluate the development of vascular networks in tissue-constructs. Here we present a simple and cost-effective method to visualize and quantify vascularization in vivo.

Insufficient vascularization is considered to be one of the main factors limiting the clinical success of tissue-engineered constructs. In order to ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Synthesis of Multi-walled Carbon Nanotubes Modified with Silver Nanoparticles and Evaluation of Their Antibacterial Activities and Cytotoxic Properties

In this study, multi-walled carbon nanotubes (MWCNTs) were treated with an aqueous sulfuric acid solution to form an oxygen-based functional group. Silver MWCNTs were prepared by the reductive deposition of silver from an aqueous solution of AgNO3 on the oxidized MWCNTs. Given the unique color of the CNTs, it was not possible to apply them to the minimum inhibitory concentration or mitochondrial toxicity assays to evaluate the toxicity and antibacterial properties, since they would interfere with the assays. The inhibition zone and minimum bactericidal concentration for the Ag-MWCNTs were measured and Live/Dead and Trypan Blue assays were used to measure the toxicity and antibacterial properties without interfering with the color of the CNTs.

In this study, antimicrobial nanomaterials were synthesized by acidic oxidation of multiwalled carbon nanotubes and subsequent reductive deposition of silver nanoparticles. Antimicrobial activity and cytotoxicity tests were performed with the as-prepared nanomaterials.

In this study, multi-walled carbon nanotubes (MWCNTs) were treated with an aqueous sulfuric acid solution to form an oxygen-based functional group. Silver ...

Biological Compatibility Profile on Biomaterials for Bone Regeneration

Large non-union bone fractures are a significant challenge in orthopedic surgery. Although auto- and allogeneic bone grafts are excellent for healing such lesions, there are potential complications with their use. Thus, material scientists are developing synthetic, biocompatible biomaterials to overcome these problems. In this study, we present a multidisciplinary platform for evaluating biomaterials for bone repair. We combined expertise from bone biology and immunology to develop a platform including in vitro osteoclast (OC) and osteoblast (OB) assays and in vivo mouse models of bone repair, immunogenicity, and allergenicity. We demonstrate how to perform the experiments, summarize the results, and report on biomaterial biocompatibility. In particular, we tested OB viability, differentiation, and mineralization and OC viability and differentiation in the context of &#946;-tricalcium phosphate (&#946;-TCP) disks. We also tested a &#946;-TCP/Collagen (&#946;-TCP/C) foam which is a commercially available material used clinically for bone repair in a critical-sized calvarial bone defect mouse model to determine the effects on the early phase of bone healing. In parallel experiments, we evaluated immune and allergic responses in mice. Our approach generates a biological compatibility profile of a bone biomaterial with a range of parameters necessary for predicting the biocompatibility of biomaterials used for bone healing and repair in patients.

The number of novel biomaterials engineered for repairing large bone lesions is continuously expanding with the aim to enhance bone healing and overcome the complications associated with bone transplantation. Here, we present a multidisciplinary strategy for pre-clinical biocompatibility testing of biomaterials for bone repair.

Large non-union bone fractures are a significant challenge in orthopedic surgery. Although auto- and allogeneic bone grafts are excellent for healing such ...

Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many parameters that efficiently measure photosynthesis. Leaf reflectance analysis provides several vegetation indices in ecology and agriculture, including the photochemical reflectance index (PRI), which can be used as an indicator of thermal energy dissipation during photosynthesis because it correlates with non-photochemical quenching (NPQ). However, since NPQ is a composite parameter, its validation is required to understand the nature of the PRI parameter. To obtain physiological evidence for evaluation of the PRI parameter, we simultaneously measured chlorophyll fluorescence and leaf reflectance in xanthophyll cycle defective mutant (npq1) and wild-type Arabidopsis plants. Additionally, the qZ parameter, which likely reflects the xanthophyll cycle, was extracted from the results of chlorophyll fluorescence analysis by monitoring relaxation kinetics of NPQ after switching the light off. These simultaneous measurements were carried out using a pulse-amplitude modulation (PAM) chlorophyll fluorometer and a spectral radiometer. The fiber probes from both instruments were positioned close to each other to detect signals from the same leaf position. An external light source was used to activate photosynthesis, and the measuring lights and saturated light were provided from the PAM instrument. This experimental system enabled us to monitor light-dependent PRI in the intact plant and revealed that light-dependent changes in PRI differ significantly between the wild type and npq1 mutant. Furthermore, PRI was strongly correlated with qZ, meaning that qZ reflects the xanthophyll cycle. Together, these measurements demonstrated that simultaneous measurement of leaf reflectance and chlorophyll fluorescence is a valid approach for parameter evaluation.

We describe a new technical approach to study photosynthetic responses in higher plants involving simultaneous measurements of chlorophyll a fluorescence and leaf reflectance using a PAM and a spectral radiometer for the detection of signals from the same leaf area in Arabidopsis.

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many ...

Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows assessment of bone substitution materials for oral and craniofacial bone regeneration in terms of bone growth and neovascularization support. Once animals are anesthetized and ventilated (endotracheal intubation), four cylinders made of polyether ether ketone (PEEK) are screwed onto the skull, on both sides of the median and coronal sutures. Five intramedullary holes are drilled within the bone area delimited by each cylinder, allowing influx of bone marrow cells. The material samples are placed into the cylinders which are then closed. Finally, the surgical site is sutured, and animals are awaken. Bone growth may be assessed on live animals by using microtomography. Once animals are euthanized, bone growth and neovascularization may be evaluated by using microtomography, immune-histology and immunofluorescence. As the evaluation of a material requires maximum standardization and calibration, the calvarial model appears ideal. Access is very easy, calibration and standardization are facilitated by the use of defined cylinders and four samples may be assessed simultaneously. Furthermore, live tomography may be used and ultimately a large decrease in animals to be euthanized may be anticipated.

Here we present a surgical protocol in rabbits with the aim to assess bone substitution materials in terms of bone regeneration capacities. By using PEEK cylinders fixed onto rabbit skulls, osteoconduction, osteoinduction, osteogenesis and vasculogenesis induced by the materials may be evaluated either on live or euthanized animals.

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows ...

Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested cell culture fluid. While traditional methods like centrifugation or filtration are widely implemented for cell removal, the equipment for these processes have large footprints and operation can involve contamination risks and filter fouling. Additionally, traditional methods may not be ideal for continuous bioprocessing schemes for primary clarification. Thus, an alternate application using acoustic (sound) waves was investigated to continuously separate cells from the cell culture fluid. Presented in this study is a detailed protocol for using a bench-scale acoustic wave separator (AWS) for the primary separation of culture fluid containing a monoclonal IgG1 antibody from a CHO cell bioreactor harvest. Representative data are presented from the AWS and demonstrate how to achieve effective cell clarification and product recovery. Finally, potential applications for AWS in continuous bioprocessing are discussed. Overall, this study provides a practical and general protocol for the implementation of AWS in primary clarification for CHO cell cultures and further describes its application potential in continuous bioprocessing.

Presented here is a protocol for the primary clarification of CHO cell culture using an acoustic separator. This protocol can be used for the primary clarification of shake flask cultures or bioreactor harvests and has the potential application for continuous clarification of the cell bleed material during perfusion bioreactor operations.

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested ...

In vivo Structural Assessments of Ocular Disease in Rodent Models using Optical Coherence Tomography

Spectral-domain optical coherence tomography (SD-OCT) is useful for visualizing retinal and ocular structures in vivo. In research, SD-OCT is a valuable tool to evaluate and characterize changes in a variety of retinal and ocular disease and injury models. In light induced retinal degeneration models, SD-OCT can be used to track thinning of the photoreceptor layer over time. In glaucoma models, SD-OCT can be used to monitor decreased retinal nerve fiber layer and total retinal thickness and to observe optic nerve cupping after inducing ocular hypertension. In diabetic rodents, SD-OCT has helped researchers observe decreased total retinal thickness as well as decreased thickness of specific retinal layers, particularly the retinal nerve fiber layer with disease progression. In mouse models of myopia, SD-OCT can be used to evaluate axial parameters, such as axial length changes. Advantages of SD-OCT include in vivo imaging of ocular structures, the ability to quantitatively track changes in ocular dimensions over time, and its rapid scanning speed and high resolution. Here, we detail the methods of SD-OCT and show examples of its use in our laboratory in models of retinal degeneration, glaucoma, diabetic retinopathy, and myopia. Methods include anesthesia, SD-OCT imaging, and processing of the images for thickness measurements.

Here, we describe the use of spectral-domain optical coherence tomography (SD-OCT) to visualize retinal and ocular structures in vivo in models of retinal degeneration, glaucoma, diabetic retinopathy, and myopia.

Spectral-domain optical coherence tomography (SD-OCT) is useful for visualizing retinal and ocular structures in vivo. In research, SD-OCT is a valuable ...