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A comparison of engineered urinary bladder and intestinal smooth muscle for urinary bladder wall replacement in a rabbit model
Journal of Pediatric Surgery (2006) 41, 2090 – 2094
www.elsevier.com/locate/jpedsurg
A comparison of engineered urinary bladder and intestinal smooth muscle for urinary bladder wall replacement in a rabbit modelB Jin-Yao Lai MD*, Pei-Yeh Chang MD, Jer-Nan Lin MD Department of Pediatric Surgery, Chang-Gung Children’s Hospital, School of Medicine, Chang-Gung University, Kweishan, Taoyuan 333, Taiwan Index words: Urinary bladder; Small intestine; Polyglycolic acid; Cell culture; Bladder augmentation
Abstract Background/Purpose: The small intestine is the most common resource for bladder augmentation. Little is known whether intestinal smooth muscle cells (SMCs) may be engineered into bladder tissue. We investigated the phenotypic and functional characteristics of engineered bladder and intestinal SMCs as bladder wall replacement in a rabbit model. Methods: One month after an initial 70% partial cystectomy, 3 autoaugmentation surgeries were performed, including traditional autoaugmentation (TA, n = 6), TA using engineered bladder SMCs (TA + B, n = 6), and TA using intestinal SMCs (TA + I, n = 6). All were followed up by bladder volume measurement and retrieved on the first, third, and sixth month. The grafts and the native bladder wall were evaluated with immunocytochemistry and electrical field stimulation (EFS). Statistical analysis was performed using analysis of variance. Results: Both the TA + I and TA + B groups showed significant and similar bladder capacity increment in all time points. The engineered muscle cells demonstrated the typical bcontraction-relaxationQ response to supramaximal EFS. There were no statistical differences in both the TA + I and TA + B groups in contractility force. Conclusion: Engineered SMCs derived from urinary bladder and small intestine could retain their phenotype after implantation in vivo. Both exhibited a similar degree of contractility to EFS. These results suggest that there are no phenotypic or functional differences between muscle cells obtained from the 2 different organs. Both have the potential to be engineered into normal bladder tissues. D 2006 Elsevier Inc. All rights reserved.
Presented at the 39th Annual Meeting of the Pacific Association of Pediatric Surgeons, May 14 –18, 2006, Taipei, Taiwan. B Supported by grants from the National Science Council of the Republic of China (No. NSC90-2314-B-182A-024) and from Chang Gung Memorial Hospital (CMRP 1296). * Corresponding author. Tel.: +886 3 328 1200x8227; fax: +886 3 328 7261. E-mail address: [email protected] (J.-Y. Lai). 0022-3468/$ – see front matter D 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.jpedsurg.2006.08.013
Conventional enterocystoplasty is associated with a variety of complications [1-3], which in turn stimulated the development of alternative methods for bladder augmentation. Autoaugmentation, which preserves intact urothelial layers, prevents the complications related to the intestinal mucosa. However, the bladder volume increment after the autoaugmentation is often limited without a satisfactory bladder wall support [4-7]. Autoaugmentation with
Urinary bladder wall replacement in a rabbit model
2091
demucosalized seromuscular colonic or gastric flaps offers good bladder wall support. These flaps prevent adhesions, shrinkage, perforation, and even leakage of the protruded mucosa. Disadvantages of these flaps include operative time, technical difficulties, and postoperative complications [8,9]. We have demonstrated that engineered bladder smooth muscle cells (B-SMCs) could be an effective bladder wall support during bladder autoaugmentation in a rabbit model [10]. Clinically, the small intestine is the most common resource for bladder augmentation. It is not known whether isolated intestinal SMCs (I-SMCs) may be engineered into a functional muscular tissue. We attempted to determine the phenotypic and functional characteristics of the engineered B-SMCs and I-SMCs as bladder wall replacement in a rabbit model.
1. Materials and methods 1.1. Polyglycolic acid scaffolds Unwoven sheets of polyglycolic acid polymers (Smith and Nephew, Heslington, York, UK) were trimmed to 4 4-cm patches. The scaffolds were designed to degrade via hydrolysis during a 4-week period. The scaffolds were coated with a liquefied copolymer PLGA (Sigma Chemical Co, St Louis, Mo), 80 mg/mL, to achieve adequate mechanical characteristics. The polymers were sterilized in ethylene oxide and stored under sterile condition until used.
1.2. Animals Experiments were performed on male New Zealand white rabbits with body weight of about 3 kg. This project was approved by the Animal Research and Care Committee at Chang-Gung Children’s Hospital. All animals were in good health.
1.4. Small I-SMC seeding and implantation A piece of the small intestinal wall, about 2 2 cm in size, was obtained at the same time. The muscle layer was stripped from the mucosa. The I-SMCs were cultured by the same technique used with B-SMCs. The muscle cells were expanded until the desired cell numbers were obtained. The seeding density was the same as with the B-SMCs.
1.5. Bladder autoaugmentation One month after PC, all rabbits were divided into 3 groups. A 4 4-cm muscular defect was created at the anterior bladder surface without perforation into the mucosal layer. In the group using traditional autoaugmentation (TA, n = 6), the bulging mucosa was left uncovered. In TA + B (B-SMCs, n = 6) and TA + I (I-SMCs, n = 6), the cell-seeded scaffolds were patched to the prolapsed mucosa, respectively. Perivesical fat was used to cover the implanted grafts as a source of blood supply. After the operation, the rabbits were allowed to resume normal diet. Bladder volume measurements were performed in all animals on 1, 3, and 6 months time point. At the time of harvest, gross appearance of the grafts was identified by the silk stitches that were put during surgery.
1.6. Bladder volume measurements Bladder volume measurements were performed by using a 7-F double-lumen transurethral catheter. The bladder was emptied and the intravesical pressure was recorded as resting pressure. The pressure was then recorded during instillation of prewarmed saline solution at constant rates until the pressure of 40 cm H2O was reached. The bladder capacity at 40 cm H2O was used as bbladder volume.Q
1.7. Histologic and immunocytochemical analyses 1.3. Bladder SMC seeding and implantation All rabbits received 70% partial cystectomy (PC). Bladder specimens were obtained during PC. The B-SMCs were cultured by using a previously described technique [10,11]. The muscle cells were expanded until the desired cell numbers were obtained.
Table 1
The scaffolds were placed in 10% neutral buffered formalin for histologic (hematoxylin and eosin) and immunohistochemical evaluation (smooth muscle a-actin and pancytokeratin AE1/AE3). We used 3,3V-diaminobenzidine tetrahydrochloride as the enzyme substrate, which yielded granular brown deposits for positive interpretation.
The changes in bladder volume in percentage
Preoperative Partial cystectomy First month Third month Sixth month
Control
TA
TA + B
TA + I
100 27.94 35.50 37.52 36.55
100 34.71 37.74 42.62 41.14
100 35.51 54.03 73.25 76.48
100 35.12 51.30 71.42 76.91
F F F F
14.05 5.41 6.48 0.42
* Indicates P b .05 as compared with the control group.
F F F F
4.51 4.75 8.35 1.25
F F F F
5.16 1.12* 3.71* 3.10*
F F F F
5.01 2.79* 4.81* 3.65*
2092
J.-Y. Lai et al.
2.1. Gross appearance of the bladder Both the TA + B and TA + I groups could retain grossly normal appearance. The ureters and kidneys were grossly normal without evidence of hydronephrosis. There was no bladder calculus in all groups.
2.2. Bladder volume profile
Fig. 1 Microscopic findings of the TA + I augmentation grafts at the sixth month. A, The histology at the anastomosis shows unorganized muscle bundles (arrowhead) and the marking stitch (arrow) (immunocytochemistry smooth muscle a-actin, original magnification 100). B, The center of the augmentation part shows similar unorganized muscle bundles all over the graft (immunocytochemistry smooth muscle a-actin, original magnification 200).
1.8. Tissue bath test The tissue bath test was described previously [18]. In brief, modified Krebs solution was used for the tissue bath. A 50-mL double-chambered quiet bath was used as a working chamber. The gas flow induced circulation of the Krebs solution, which was warmed to 378C by an external heating circuit. A 1 0.5-cm strip, either from native or augmented bladder, was mounted in the tissue bath. An isometric force displacement transducer was connected to the other side by means of two 5-0 braided silk. An electric stimulator served as the source of electrical field stimulation (EFS). The transducer signal was fed into a chart recorder. For the electrical field study, we used 100 V at 10, 30, and 50 Hz for stimulation. The weight of the strip was measured after the contractility test. The contractility strength under supramaximal stimulation (50Hz) was determined for statistical analysis. The contraction strength was expressed as gram force per gram of tissue (g/g).
Preoperatively, the bladder capacity in the control, TA, TA + B, and TA + I groups was 80.5 F 11.2, 69.2 F 9.5, 68.5 F 5.2, and 72.2 F 5.2 mL, respectively. After PC, the capacity decreased to 25.3 F 2.8 mL (27.9% F 14% of original capacity), 24.0 F 4.4 mL (34.7% F 4.5%), 24.2 F 2.7 mL (35.5% F 5.2%), and 25.2 F 2.4 mL (35.1% F 5.0%), respectively. Since then, the bladder capacity increased slowly with time until the last time point in all groups. At the 6th month, the volume became 32 F 4.2 mL (36.5% F 0.4%, control group), 30.0 F 1.4 mL (41.1% F 1.3%, TA group), 55.5 F 4.9 mL (76.5% F 3.1%, TA + B group), and 58.52 F 4.9 mL (76.9% F 3.7%, TA + I group), respectively. Both the TA + B and TA + I groups, but not the TA group, demonstrated persistent increment in bladder volume with time as compared with the control group ( P b .05 at the first, third, and sixth month time point). However, there was no statistical difference between the TA + B and TA + I group ( P = .95) (Table 1).
2.3. Histology Microscopically, all augmented parts were covered with normal epithelial layer through all time points. Unorganized regenerated smooth muscle bundles were detected in both the TA + B and TA + I group 3 and 6 months after the surgery (Fig. 1). The TA group had only disorientated fibers. Only scanty small nerve fibrils were visible over the augmentation parts in all groups.
2.4. Supramaximal muscle contraction All augmentation grafts in the TA + B and TA + I groups demonstrated the typical bcontraction-relaxationQ response to supramaximal EFS. The contraction waves were amplitude
1.9. Statistical analysis Statistical evaluations were performed using analysis of variance, and P value less than .05 was considered as significant.
2. Results All animals survived the surgical procedures and were able to void spontaneously after the operation. There was no urine leakage in all groups.
Fig. 2 The changes in supramaximal muscle contractility at 50 Hz in different time points.
Urinary bladder wall replacement in a rabbit model dependent. Most of the strips reached maximal contractility at 30 to 50 Hz. The contraction strength of the native bladder varied from 4.5 F 0.42 to 4.87 F 0.67 g/g during all time points. The contractility of augmentation grafts was low initially. During the first month, the strength was only 0.78 F 0.10 g/g (TA + B group) and 0.96 F 0.35 g/g (TA + I group), respectively. Since then, the strength increased with time until the last time point in all groups. During the sixth month, the contractility became 2.21 F 0.47 g/g (TA + B group) and 2.95 F 0.41 g/g (TA + I group), respectively. There was no statistical difference in the contraction strength between the TA + B and TA + I groups ( P = .23). The cell-seeded scaffolds showed contractile responses at approximately 45% to 60% amplitude as compared with native bladder strips (Fig. 2).
3. Discussion Tissue-engineering techniques using autologous cells have been previously used in the creation of neo-bladders [12]. The animal models underwent an initial biopsy of tissue from normal bladders with cell expansion in vitro and cell seeding to the scaffolds. The engineered bladders showed satisfactory bladder volume with time. However, biopsy on the bladder may increase the incidence of bladder wall fibrosis and perivesical adhesion. It might be detrimental to future augmentation procedure. Questions remain regarding the replacement of B-SMCs by other types of SMCs, such as I-SMCs. The present study was performed to elucidate some of the phenotypic and functional characteristics of bladder and intestinal muscle cells for the engineering of urinary bladder wall. Bladder SMCs, when cultured in vitro and seeded in a collagen lattice, demonstrate loss of contractile response to pharmacologic agonists, such as carbachol [13]. Bladder SMCs, like vascular SMCs, assume a dedifferentiated or proliferative phenotype when placed in culture. It is well known that cultures of dispersed vascular SMCs invariably cause rapid modulation from the contractile to the proliferative phenotype. This change involves loss of contractile ability, decreased contractile protein content, and increased expression of rough endoplasmic reticulum [14-17]. Previous studies have shown that tissue-engineered B-SMCs seeded on either PGA polymer or collagen matrix could regain the contractile phenotype both in vitro and in vivo in a mice model [18]. Our further studies demonstrated that engineered B-SMCs seeded on PGA polymers could offer satisfactory architectural support by smooth muscle regeneration and significantly increase the bladder volume in a rabbit model [10]. In this study, the bladder wall support from I-SMCs seeded on PGA polymer was as effective as that from B-SMCs in terms of bladder volume increment. The tissue bath study provided a reliable quantitative measurement of the augmentation parts to characterize the function of the engineered grafts. Every muscle strip, either
2093 from the native bladder or augmentation parts, could be evaluated for its contractility by EFS. Although the same number of SMCs was seeded to every scaffold initially after incubation and implantation, one would expect a certain variation of SMC number in each graft owing to different cell proliferative and regenerative rates. The individual graft variation was reduced by dividing the contraction force by the weight of each strip (gram of contraction force/gram per strip). The EFS results in the present study are similar to those described previously [18]. The engineered grafts could retain their contractile phenotype in vivo, no matter what the source of the SMCs is. The contraction force could increase slowly with time. The regenerated SMCs, when exposed to the cyclic bstretch-and-relaxationQ patterns associated with bladder filling and emptying, may be evoked to an increased contractile ability [19]. Both the TA + B and TA + I groups demonstrated similar but lower contractile force (45%-60%) as compared with native bladder. The lower contractility could be attributed to the unorganized muscle bundle in the augmentation grafts. In conclusion, engineered bladder muscle derived from the urinary bladder and small intestine could retain their phenotype after implantation in vivo. Both groups exhibited a similar degree of contractility to electrical stimulation. These results suggest that there are no phenotypic or functional differences between muscle cells obtained from the 2 different organs. Both of them may have the potential to be engineered into normal bladder tissues.
Acknowledgments The authors thank Mr James C Shaw for his technical assistance in this experiment.
References [1] Atala A, Bauer SB, Hendren WH, et al. The effect of gastric augmentation on bladder function. J Urol 1993;149:1099 - 102. [2] Kaefer M, Tobin MS, Hendren WH, et al. Continent urinary diversion: the Children’s Hospital experience. J Urol 1997;157:1394 - 9. [3] McDougal WS. Long-term metabolic effects of urinary diversion: a comparison of myelomeningocele patients managed by clean intermittent catheterization and urinary diversion. J Urol 1992;147:1343 - 7. [4] Snow BW, Cartwright PC. Bladder autoaugmentation. Urol Clin North Am 1996;23:323 - 31. [5] MacNeily AE, Afshar K, Coleman GU, et al. Autoaugmentation by detrusor myotomy: its lack of effectiveness in the management of congenital neuropathic bladder. J Urol 2003;170:1643 - 6. [6] Perovic SV, Djordjevic ML, Kekic ZK, et al. Bladder autoaugmentation with rectus muscle backing. J Urol 2002;168:1877 - 80. [7] Marte A, Di Meglio D, Cotrufo AM, et al. A long-term follow-up of autoaugmentation in myelodysplastic children. BJU Int 2002;89: 928 - 31. [8] Dewan PA. Autoaugmentation demucosalized enterocystoplasty. World J Urol 1998;16:255 - 61.
2094 [9] Nguyen DH, Mitchell ME, Horowitz M, et al. Demucosalized augmentation gastrocystoplasty with bladder autoaugmentation in pediatric patients. J Urol 1996;156:206 - 9. [10] Lai JY, Chang PY, Lin JN. Bladder autoaugmentation using various biodegradable scaffolds seeded with autologous smooth muscle cells in a rabbit model. J Pediatr Surg 2005;40:1869 - 73. [11] Freeman MR, Yoo JJ, Raab G, et al. Heparin-binding EGF-like growth factor is an autocrine factor for human urothelial cells and is synthesized by epithelial and smooth muscle cells in the human bladder. J Clin Invest 1997;99:1028 - 36. [12] Oberpenning F, Meng J, Yoo JJ, et al. De novo reconstitution of a functional mammalian urinary bladder by tissue engineering. Nat Biotechnol 1999;17:149 - 55. [13] Kropp BP, Zhang Y, Tomasek JJ, et al. Characterization of cultured bladder smooth muscle cells: assessment of in vitro contractility. J Urol 1999;162:1779 - 84.
J.-Y. Lai et al. [14] Chamley-Campbell JH, Campbell GR, Ross R. Phenotype-dependent response of cultured aortic smooth muscle to serum mitogens. J Cell Biol 1981;89:379 - 83. [15] Owens GK. Regulation of differentiation of vascular smooth muscle cells. Physiol Rev 1995;75:487 - 517. [16] Thyberg J. Differentiated properties and proliferation of arterial smooth muscle cells in culture. Int Rev Cytol 1996;169:183 - 265. [17] Bowers CW, Dahm LM. Maintenance of contractility in dissociated smooth muscle: low density cultures in a defined medium. Am J Physiol 1993;264(1 Pt 1):C229 - 36. [18] Lai JY, Yoon CY, Yoo JJ, et al. Phenotypic and functional characterization of in vivo tissue engineered smooth muscle from normal and pathological bladders. J Urol 2002;168:1853 - 7. [19] De Filippo RE, Atala A. Stretch and growth: the molecular and physiologic influences of tissue expansion. Plast Reconstr Surg 2002;109:2450 - 62.
www.elsevier.com/locate/jpedsurg
A comparison of engineered urinary bladder and intestinal smooth muscle for urinary bladder wall replacement in a rabbit modelB Jin-Yao Lai MD*, Pei-Yeh Chang MD, Jer-Nan Lin MD Department of Pediatric Surgery, Chang-Gung Children’s Hospital, School of Medicine, Chang-Gung University, Kweishan, Taoyuan 333, Taiwan Index words: Urinary bladder; Small intestine; Polyglycolic acid; Cell culture; Bladder augmentation
Abstract Background/Purpose: The small intestine is the most common resource for bladder augmentation. Little is known whether intestinal smooth muscle cells (SMCs) may be engineered into bladder tissue. We investigated the phenotypic and functional characteristics of engineered bladder and intestinal SMCs as bladder wall replacement in a rabbit model. Methods: One month after an initial 70% partial cystectomy, 3 autoaugmentation surgeries were performed, including traditional autoaugmentation (TA, n = 6), TA using engineered bladder SMCs (TA + B, n = 6), and TA using intestinal SMCs (TA + I, n = 6). All were followed up by bladder volume measurement and retrieved on the first, third, and sixth month. The grafts and the native bladder wall were evaluated with immunocytochemistry and electrical field stimulation (EFS). Statistical analysis was performed using analysis of variance. Results: Both the TA + I and TA + B groups showed significant and similar bladder capacity increment in all time points. The engineered muscle cells demonstrated the typical bcontraction-relaxationQ response to supramaximal EFS. There were no statistical differences in both the TA + I and TA + B groups in contractility force. Conclusion: Engineered SMCs derived from urinary bladder and small intestine could retain their phenotype after implantation in vivo. Both exhibited a similar degree of contractility to EFS. These results suggest that there are no phenotypic or functional differences between muscle cells obtained from the 2 different organs. Both have the potential to be engineered into normal bladder tissues. D 2006 Elsevier Inc. All rights reserved.
Presented at the 39th Annual Meeting of the Pacific Association of Pediatric Surgeons, May 14 –18, 2006, Taipei, Taiwan. B Supported by grants from the National Science Council of the Republic of China (No. NSC90-2314-B-182A-024) and from Chang Gung Memorial Hospital (CMRP 1296). * Corresponding author. Tel.: +886 3 328 1200x8227; fax: +886 3 328 7261. E-mail address: [email protected] (J.-Y. Lai). 0022-3468/$ – see front matter D 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.jpedsurg.2006.08.013
Conventional enterocystoplasty is associated with a variety of complications [1-3], which in turn stimulated the development of alternative methods for bladder augmentation. Autoaugmentation, which preserves intact urothelial layers, prevents the complications related to the intestinal mucosa. However, the bladder volume increment after the autoaugmentation is often limited without a satisfactory bladder wall support [4-7]. Autoaugmentation with
Urinary bladder wall replacement in a rabbit model
2091
demucosalized seromuscular colonic or gastric flaps offers good bladder wall support. These flaps prevent adhesions, shrinkage, perforation, and even leakage of the protruded mucosa. Disadvantages of these flaps include operative time, technical difficulties, and postoperative complications [8,9]. We have demonstrated that engineered bladder smooth muscle cells (B-SMCs) could be an effective bladder wall support during bladder autoaugmentation in a rabbit model [10]. Clinically, the small intestine is the most common resource for bladder augmentation. It is not known whether isolated intestinal SMCs (I-SMCs) may be engineered into a functional muscular tissue. We attempted to determine the phenotypic and functional characteristics of the engineered B-SMCs and I-SMCs as bladder wall replacement in a rabbit model.
1. Materials and methods 1.1. Polyglycolic acid scaffolds Unwoven sheets of polyglycolic acid polymers (Smith and Nephew, Heslington, York, UK) were trimmed to 4 4-cm patches. The scaffolds were designed to degrade via hydrolysis during a 4-week period. The scaffolds were coated with a liquefied copolymer PLGA (Sigma Chemical Co, St Louis, Mo), 80 mg/mL, to achieve adequate mechanical characteristics. The polymers were sterilized in ethylene oxide and stored under sterile condition until used.
1.2. Animals Experiments were performed on male New Zealand white rabbits with body weight of about 3 kg. This project was approved by the Animal Research and Care Committee at Chang-Gung Children’s Hospital. All animals were in good health.
1.4. Small I-SMC seeding and implantation A piece of the small intestinal wall, about 2 2 cm in size, was obtained at the same time. The muscle layer was stripped from the mucosa. The I-SMCs were cultured by the same technique used with B-SMCs. The muscle cells were expanded until the desired cell numbers were obtained. The seeding density was the same as with the B-SMCs.
1.5. Bladder autoaugmentation One month after PC, all rabbits were divided into 3 groups. A 4 4-cm muscular defect was created at the anterior bladder surface without perforation into the mucosal layer. In the group using traditional autoaugmentation (TA, n = 6), the bulging mucosa was left uncovered. In TA + B (B-SMCs, n = 6) and TA + I (I-SMCs, n = 6), the cell-seeded scaffolds were patched to the prolapsed mucosa, respectively. Perivesical fat was used to cover the implanted grafts as a source of blood supply. After the operation, the rabbits were allowed to resume normal diet. Bladder volume measurements were performed in all animals on 1, 3, and 6 months time point. At the time of harvest, gross appearance of the grafts was identified by the silk stitches that were put during surgery.
1.6. Bladder volume measurements Bladder volume measurements were performed by using a 7-F double-lumen transurethral catheter. The bladder was emptied and the intravesical pressure was recorded as resting pressure. The pressure was then recorded during instillation of prewarmed saline solution at constant rates until the pressure of 40 cm H2O was reached. The bladder capacity at 40 cm H2O was used as bbladder volume.Q
1.7. Histologic and immunocytochemical analyses 1.3. Bladder SMC seeding and implantation All rabbits received 70% partial cystectomy (PC). Bladder specimens were obtained during PC. The B-SMCs were cultured by using a previously described technique [10,11]. The muscle cells were expanded until the desired cell numbers were obtained.
Table 1
The scaffolds were placed in 10% neutral buffered formalin for histologic (hematoxylin and eosin) and immunohistochemical evaluation (smooth muscle a-actin and pancytokeratin AE1/AE3). We used 3,3V-diaminobenzidine tetrahydrochloride as the enzyme substrate, which yielded granular brown deposits for positive interpretation.
The changes in bladder volume in percentage
Preoperative Partial cystectomy First month Third month Sixth month
Control
TA
TA + B
TA + I
100 27.94 35.50 37.52 36.55
100 34.71 37.74 42.62 41.14
100 35.51 54.03 73.25 76.48
100 35.12 51.30 71.42 76.91
F F F F
14.05 5.41 6.48 0.42
* Indicates P b .05 as compared with the control group.
F F F F
4.51 4.75 8.35 1.25
F F F F
5.16 1.12* 3.71* 3.10*
F F F F
5.01 2.79* 4.81* 3.65*
2092
J.-Y. Lai et al.
2.1. Gross appearance of the bladder Both the TA + B and TA + I groups could retain grossly normal appearance. The ureters and kidneys were grossly normal without evidence of hydronephrosis. There was no bladder calculus in all groups.
2.2. Bladder volume profile
Fig. 1 Microscopic findings of the TA + I augmentation grafts at the sixth month. A, The histology at the anastomosis shows unorganized muscle bundles (arrowhead) and the marking stitch (arrow) (immunocytochemistry smooth muscle a-actin, original magnification 100). B, The center of the augmentation part shows similar unorganized muscle bundles all over the graft (immunocytochemistry smooth muscle a-actin, original magnification 200).
1.8. Tissue bath test The tissue bath test was described previously [18]. In brief, modified Krebs solution was used for the tissue bath. A 50-mL double-chambered quiet bath was used as a working chamber. The gas flow induced circulation of the Krebs solution, which was warmed to 378C by an external heating circuit. A 1 0.5-cm strip, either from native or augmented bladder, was mounted in the tissue bath. An isometric force displacement transducer was connected to the other side by means of two 5-0 braided silk. An electric stimulator served as the source of electrical field stimulation (EFS). The transducer signal was fed into a chart recorder. For the electrical field study, we used 100 V at 10, 30, and 50 Hz for stimulation. The weight of the strip was measured after the contractility test. The contractility strength under supramaximal stimulation (50Hz) was determined for statistical analysis. The contraction strength was expressed as gram force per gram of tissue (g/g).
Preoperatively, the bladder capacity in the control, TA, TA + B, and TA + I groups was 80.5 F 11.2, 69.2 F 9.5, 68.5 F 5.2, and 72.2 F 5.2 mL, respectively. After PC, the capacity decreased to 25.3 F 2.8 mL (27.9% F 14% of original capacity), 24.0 F 4.4 mL (34.7% F 4.5%), 24.2 F 2.7 mL (35.5% F 5.2%), and 25.2 F 2.4 mL (35.1% F 5.0%), respectively. Since then, the bladder capacity increased slowly with time until the last time point in all groups. At the 6th month, the volume became 32 F 4.2 mL (36.5% F 0.4%, control group), 30.0 F 1.4 mL (41.1% F 1.3%, TA group), 55.5 F 4.9 mL (76.5% F 3.1%, TA + B group), and 58.52 F 4.9 mL (76.9% F 3.7%, TA + I group), respectively. Both the TA + B and TA + I groups, but not the TA group, demonstrated persistent increment in bladder volume with time as compared with the control group ( P b .05 at the first, third, and sixth month time point). However, there was no statistical difference between the TA + B and TA + I group ( P = .95) (Table 1).
2.3. Histology Microscopically, all augmented parts were covered with normal epithelial layer through all time points. Unorganized regenerated smooth muscle bundles were detected in both the TA + B and TA + I group 3 and 6 months after the surgery (Fig. 1). The TA group had only disorientated fibers. Only scanty small nerve fibrils were visible over the augmentation parts in all groups.
2.4. Supramaximal muscle contraction All augmentation grafts in the TA + B and TA + I groups demonstrated the typical bcontraction-relaxationQ response to supramaximal EFS. The contraction waves were amplitude
1.9. Statistical analysis Statistical evaluations were performed using analysis of variance, and P value less than .05 was considered as significant.
2. Results All animals survived the surgical procedures and were able to void spontaneously after the operation. There was no urine leakage in all groups.
Fig. 2 The changes in supramaximal muscle contractility at 50 Hz in different time points.
Urinary bladder wall replacement in a rabbit model dependent. Most of the strips reached maximal contractility at 30 to 50 Hz. The contraction strength of the native bladder varied from 4.5 F 0.42 to 4.87 F 0.67 g/g during all time points. The contractility of augmentation grafts was low initially. During the first month, the strength was only 0.78 F 0.10 g/g (TA + B group) and 0.96 F 0.35 g/g (TA + I group), respectively. Since then, the strength increased with time until the last time point in all groups. During the sixth month, the contractility became 2.21 F 0.47 g/g (TA + B group) and 2.95 F 0.41 g/g (TA + I group), respectively. There was no statistical difference in the contraction strength between the TA + B and TA + I groups ( P = .23). The cell-seeded scaffolds showed contractile responses at approximately 45% to 60% amplitude as compared with native bladder strips (Fig. 2).
3. Discussion Tissue-engineering techniques using autologous cells have been previously used in the creation of neo-bladders [12]. The animal models underwent an initial biopsy of tissue from normal bladders with cell expansion in vitro and cell seeding to the scaffolds. The engineered bladders showed satisfactory bladder volume with time. However, biopsy on the bladder may increase the incidence of bladder wall fibrosis and perivesical adhesion. It might be detrimental to future augmentation procedure. Questions remain regarding the replacement of B-SMCs by other types of SMCs, such as I-SMCs. The present study was performed to elucidate some of the phenotypic and functional characteristics of bladder and intestinal muscle cells for the engineering of urinary bladder wall. Bladder SMCs, when cultured in vitro and seeded in a collagen lattice, demonstrate loss of contractile response to pharmacologic agonists, such as carbachol [13]. Bladder SMCs, like vascular SMCs, assume a dedifferentiated or proliferative phenotype when placed in culture. It is well known that cultures of dispersed vascular SMCs invariably cause rapid modulation from the contractile to the proliferative phenotype. This change involves loss of contractile ability, decreased contractile protein content, and increased expression of rough endoplasmic reticulum [14-17]. Previous studies have shown that tissue-engineered B-SMCs seeded on either PGA polymer or collagen matrix could regain the contractile phenotype both in vitro and in vivo in a mice model [18]. Our further studies demonstrated that engineered B-SMCs seeded on PGA polymers could offer satisfactory architectural support by smooth muscle regeneration and significantly increase the bladder volume in a rabbit model [10]. In this study, the bladder wall support from I-SMCs seeded on PGA polymer was as effective as that from B-SMCs in terms of bladder volume increment. The tissue bath study provided a reliable quantitative measurement of the augmentation parts to characterize the function of the engineered grafts. Every muscle strip, either
2093 from the native bladder or augmentation parts, could be evaluated for its contractility by EFS. Although the same number of SMCs was seeded to every scaffold initially after incubation and implantation, one would expect a certain variation of SMC number in each graft owing to different cell proliferative and regenerative rates. The individual graft variation was reduced by dividing the contraction force by the weight of each strip (gram of contraction force/gram per strip). The EFS results in the present study are similar to those described previously [18]. The engineered grafts could retain their contractile phenotype in vivo, no matter what the source of the SMCs is. The contraction force could increase slowly with time. The regenerated SMCs, when exposed to the cyclic bstretch-and-relaxationQ patterns associated with bladder filling and emptying, may be evoked to an increased contractile ability [19]. Both the TA + B and TA + I groups demonstrated similar but lower contractile force (45%-60%) as compared with native bladder. The lower contractility could be attributed to the unorganized muscle bundle in the augmentation grafts. In conclusion, engineered bladder muscle derived from the urinary bladder and small intestine could retain their phenotype after implantation in vivo. Both groups exhibited a similar degree of contractility to electrical stimulation. These results suggest that there are no phenotypic or functional differences between muscle cells obtained from the 2 different organs. Both of them may have the potential to be engineered into normal bladder tissues.
Acknowledgments The authors thank Mr James C Shaw for his technical assistance in this experiment.
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