The authors would like to acknowledge the staff at the Department of Experimental Medicine (AEM), University of Copenhagen, for assistance with planning and carrying out animal surgeries and husbandry, and ELLA-CS, s.r.o, Hradec Kralove, Czech Republic, for providing customized biodegradable stents used in the study. Financial support was provided by the Swedish Society of Medical Research, Promobilia Foundation, Rydbeck Foundation, Samariten Foundation, The Foundation for Pediatric Health Care, Foundation Frimurare Barnhuset in Stockholm, and the Novo Nordisk Foundation (NNFSA170030576).

This experiment was carried out in an AAALAC accredited experimental facility in accordance with the European legislation on laboratory use of animal subjects and after ethical permission granted by the Danish Ministry of Food and Agriculture (Ref. no. 2022-15-0201-01206).
1. Surgical procedure
<ol>
	<li>Animal preparation
	<ol>
		<li>Fast a female full-grown G&#246;ttingen minipig for at least 12 h preoperatively.</li>
		<li>Prepare the surgical table with all sterile utensils as described below.</li>
		<li>For full-grown standard-size minipigs, sedate the animal by intramuscular injection with 1.0-1.4 mL/10 kg with a solution of 125 mg of zolazepam and 125 mg of tiletamine suspended in 1.25 mL of ketamine (100 mg/mL), 6.25 mL of xylazine (20 mg/mL), 1.25 mL of methadone (10 mg/mL) and 2 mL of butorphanol (10 mg/mL) (later on referred as sedation mixture).</li>
		<li>Perform visual-guided endotracheal intubation and add capnography as an anesthesia monitoring parameter and confirm proper tube placement. Confirm anesthesia by vital signs and eye and interdigital reflex testing. Apply ophthalmic ointment bilaterally.</li>
		<li>Install bilateral ear vein catheters and sustain anesthesia with propofol (10-15 mg/kg/h) and fentanyl (5-15 mg/kg/h).</li>
		<li>Insert an 8 Fr urinary catheter and fill the bladder with 250 mL of physiologically temperate isotonic saline using an appropriately sized luer lock syringe.</li>
		<li>Place the pig in the supine position, then raze and scrub the abdomen. After two further rounds of skin cleaning with 70% ethanol and chlorhexidine scrub in alternating rounds.&#160;Frame the surgical field with sterile draping.</li>
	</ol>
	</li>
	<li>Tissue harvesting and surgical scaffold implantation&#160;
	<ol>
		<li>Perform a standard lower midline laparotomy with scalpel and cautery, dividing the skin, muscle, and peritoneum, and pull the intraperitoneal urinary bladder to the incision.</li>
		<li>Perform prophylactic hemostasis with electrocautery on the anterior bladder wall and excise a 2 cm2 full-wall segment, leaving a proximal opening of 1 cm2 while closing the remaining bladder wall with a fast-resorbable monofilament&#160;running suture.</li>
		<li>Carefully dissect the mucosal layer of the resected specimen and mince a 2 cm2 mucosal specimen into 1 mm2 micrografts for scaffold embedding (described below in section 2).</li>
		<li>After completing the scaffold, anastomose the tubular construct to the remaining opening on the anterior bladder wall with a slow-resorbable monofilament running suture.</li>
		<li>Use a peritoneal flap from the pubovesical ligament to patch the tubular scaffold and place an intraluminal 14 Fr antegrade colonic enema (ACE) stopper into the tubular scaffold.</li>
		<li>Ligate the distal end of the conduit with a slow-resorbable 4-0 monofilament suture to prevent urine from leaking out, and inject a total of 250 mL of sterile saline with syringes via the bladder catheter to confirm anastomotic patency.</li>
		<li>Bluntly dissect a trans-fascial channel laterally to the midline, 2-3 cm caudally to the caudal mammary gland on the right side, and place the conduit in a subcutaneous pocket. Fixate the distal conduit with two transcutaneous non-resorbable monofilament sutures to mark the location at skin level.</li>
		<li>Close the anterior muscle fascia of the abdominal muscle with a slow resorbable monofilament running suture, adapt the subcutis with a fast-resorbable braided running suture, and close the skin with a non-resorbable monofilament running suture.</li>
		<li>After discontinuing anesthesia, extubate the animal and observe it in the stables until it is fully ambulant and safely able to drink and eat.</li>
	</ol>
	</li>
</ol>
2. Scaffold construction
<ol>
	<li>Preparation of the composite scaffold
	<ol>
		<li>Prior to surgery (maximum 2 h), prepare a liquid solution of rat-tail collagen type I as previously described17. In brief, add 4:1 of 10x minimum essential medium (MEM) to the collagen solution and approximate the pH to 7.4 with 1 M NaOH, and finally add 1x MEM, aiming for a final collagen concentration of 1.64 mg/mL. Store the solution in a sterile vial on ice until further use.</li>
		<li>After surgical tissue resection and mincing, manually place the mucosal particles (i.e., micrografts) onto a 2 cm x 6 cm fitted biodegradable mesh with a 1:6 expansion rate (e.g., a 2 cm2 mucosal tissue is expanded to a 12 cm2 mesh) using forceps.</li>
		<li>Prepare a sterile rectangular steel mold measuring 1 cm x 3 cm x 6 cm (height x width x length) on top of a sterile steel plate and place the mesh into the steel mold with the micrografts facing upwards. Gently pour 20 mL of the collagen solution into the mold, making sure not to flush the micrografts off the mesh. Transfer the entire construct to a 38 &#176;C sterile heating chamber and leave to solidify for five minutes.</li>
		<li>After sufficient solidification, slide the hydrogel onto a nylon mesh resting on a perforated steel plate and gently remove the mold.</li>
		<li>Expel water from the hydrogel by placing a nylon mesh and then a steel plate on top of the gel, and then passively compress with a 120 g weight (in this case equivalent to the steel mold used for embedding) placed on top of the steel plate for 5 min.</li>
		<li>After compression, roll the flattened scaffold around a biodegradable stent, micrografts facing the stent, measuring 5 cm x 0.6 cm (length x inner diameter), and suture the scaffold in place longitudinally with a slow-resorbable monofilament running suture. The completed conduit is now ready for surgical implantation.</li>
	</ol>
	</li>
</ol>
3. Postoperative management
<ol>
	<li>Analgesia and antibiotic prophylaxis
	<ol>
		<li>Administer buprenorphine (0.05-0.1 mg/kg/8 h intravenously) for the first 3 days, meloxicam (0.4 mg/kg/day intramuscular or orally) for the first 4 days, and trimethoprim (2.7 mg/kg/day intramuscular or 4.2 mg/kg/day orally) and sulfadoxin (13.3 mg/kg/day intramuscular or 20.8 mg/kg/day orally) for the first 5 days. Administer the intramuscular injections postoperatively while the animal is still anesthetized.</li>
		<li>Single-house the animals to avoid nibbling of external vein catheters and suture material. Provide visual contact with neighboring minipigs through plexiglass windows and the possibility of snout contact between pens. Provide daily fresh straw and hay, as well as toys and water supply ad libitum and feed twice daily.</li>
		<li>Monitor the animals daily for natural behavior, eating habits, urine- and stool production, and assess body weight weekly.</li>
		<li>At the end of the observational period (6 weeks), sedate the animals with 1-1.4 mL/10 kg intramuscular injection of sedation mixture and terminate the animal with a lethal pentobarbital injection (100 mg/kg intravenously).</li>
	</ol>
	</li>
</ol>
4. Postmortem assessments
<ol>
	<li>Gross anatomy
	<ol>
		<li>After termination, dissect the distal conduit at skin level and remove the ACE stopper. Close the urethra with a plastic clamp and inject 250 mL of a 1:20 contrast solution of iohexol in isotonic saline via the distal conduit opening using a catheter.</li>
		<li>Assess the animal with a 64-slice computed tomography scanner. Visualize images using multiplanar reconstruction and analyze all images using medical image processing software.</li>
		<li>Perform an endoscopic examination of the bladder and conduit lumina with a 16.2 Fr flexible cystoscope via the native urethra.</li>
		<li>Resect the conduit en bloc while carefully evaluating any gross anatomical findings. Additionally, resect full-wall bladder biopsies with a 2 cm margin to the conduit anastomosis and process in a similar fashion for reference values.</li>
	</ol>
	</li>
	<li>Histological processing
	<ol>
		<li>Fix the excised specimen in 10% formalin for 24 h.</li>
		<li>Divide the conduit orthogonally with a scalpel into equal-sized separate sections of proximal, medial, and distal conduit segments. Dehydrate the specimens with increasing ethanol concentrations and embed them in paraffin before microtome sectioning.</li>
		<li>Stain 5 &#181;m sections with hematoxylin and eosin (H&#38;E) and pancytokeratin CK-AE and scan with a digital histology slide scanner.</li>
	</ol>
	</li>
</ol>

Accessing Collagen Scaffold for Single-Stage Urinary Diversion Surgery in Pigs

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compressed+collagen+constructs+with+optimized+mechanical+properties+and+cell+interactions+for+tissue+engineering+applications.">Compressed collagen constructs with optimized mechanical properties and cell interactions for tissue engineering applications.</a> Int J Biol Macromol. 108, 158-166 (2018).</li><li>Chamorro, C. I., Zeiai, S., Engberg, G. R., Fossum, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minced+tissue+in+compressed+collagen:+A+cell-containing+biotransplant+for+single-staged+reconstructive+repair.">Minced tissue in compressed collagen: A cell-containing biotransplant for single-staged reconstructive repair.</a> J Vis Exp. 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I., Fossum, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advancing+autologous+urothelial+micrografting+and+composite+tubular+grafts+for+future+single-staged+urogenital+reconstructions.">Advancing autologous urothelial micrografting and composite tubular grafts for future single-staged urogenital reconstructions.</a> Sci Rep. 13 (1), 15584 (2023).</li><li>Willacy, O., Juul, N., Taouzlak, L., Chamorro, C. I., Ajallouiean, F., Fossum, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+perioperative+layered+autologous+tissue+expansion+graft+for+hollow+organ+repair.">A perioperative layered autologous tissue expansion graft for hollow organ repair.</a> Heliyon. 10 (3), e25275 (2024).</li><li>Chamorro, C. 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The authors have no conflicts of interest to declare.

This protocol presents a simple and approachable technique for future reconstructive surgeries. A common drawback in tissue engineering, including autologous cell expansion, is the expensive and substantial prefatory steps required before surgical implantation. Autologous micrografting may simplify many of these steps and potentially allow for single-staged procedures. By auto-transplanting complex histological entities, pro-regenerative paracrine signaling is induced18. In previous studies, we experienced that micrografts alone are vulnerable to the physical environments unless suitably attached to a scaffold15,19. Collagen has been studied as a viable environment for tissue expansion in vitro and was chosen for our purpose due to its favorable biocompatibility and commercial availability. The composite scaffold presented here has previously been optimized during in vitro experiments evaluating variations in micrograft embedding and collagen concentrations20,21,22. Before in vivo testing, the scaffold properties regarding permeability, biomechanics, and degradation have been evaluated in vitro20. Furthermore, the in vivo scaffold-based tissue expansion was previously validated in rodent and rabbit models21,22.
The surgical model was chosen to evaluate a tubular version of the scaffold, mimicking the clinical setting of a urinary diversion for neurogenic bladder dysfunction in pediatric or adolescent patients. The critical steps include the exact dissection of the mucosal micrografts and maintaining a moist environment from the time of resection to the scaffold embedding. Another critical step includes proper hydrogel solidification; careful pipetting of the collagen ensures that air bubbles are not formed within the gel, and correct temperature settings and component solutions ensure that the gel properly solidifies. Failure to obtain a solidified gel will increase the risk of collagen delamination and micrograft detachment. For the surgical part, careful handling during implantation is crucial to avoid damaging the micrografts due to mechanical trauma or dissociation. Before closing the abdomen, fluid patency should be carefully addressed by insufflating the bladder with fluids.
Limitations to the technique include the thickness of the scaffold, which intuitively has upper limits regarding the diffusion of nutrients from the external environment to the micrografts. On the other hand, a reduction in scaffold thickness may lead to inappropriately high permeability and urine leakage. Our current composition is based on previous in vitro assessments, where cell regeneration in varying collagen concentrations was compared20. Micrografting of autologous tissues also relies on healthy graft tissue, making the current procedure unsuited for malignant diseases where the risk of cancerous re-transplantation cannot be properly ruled out23; nevertheless, the current technique was designed for cases with functional voiding disabilities where this is not considered a risk. Although the model mimics several steps from the clinical setting (i.e., the appendicovesicostomy procedure), this experiment does not utilize a fully functional stoma for urinary diversion since the conduit is ligated distally. Also, as clinical complications can occur life-long, a 6 week observational period may provide limited knowledge on specific outcomes on strictures and continency. Therefore, an additional 6-month followup could be added to the study after anastomosing the healed conduit to the skin level.
The perspective of this technique relates to the simple design, enabling universal applications in case the micrograft tissue-origin and supporting biomaterial is replaced with other relevant alternatives. These components can be modified to suit organ-specific purposes related to scaffold strength, elasticity, and biodegradation. Finally, the accessible and low-cost expenses allow for reproducibility and a broadened translation of the technique.

In urogenital malformations, reconstructive surgery can be required to restore functional anatomy, often on a vital indication1,2. Conventional surgical approaches have utilized native tissues from other organ systems (such as the gastrointestinal tract) to reconstruct the malformed or missing organs; however, often with the risk of severe postoperative complications3,4. In the case of urinary diversion for patients with neurogenic bladder dysfunction in need of long-term catheterization, the appendix or re-tailored small bowel segments are often used to construct a urinary conduit5,6. Tissue engineering offers an alternative grafting tissue that can be tailored to meet organ-specific characteristics, thereby minimizing postoperative morbidity for the patients7,8. Whereas scaffolds of various kinds can be implanted on their own, additional scaffold cellularization, preferably with autologous cells, has been shown to improve the regenerative outcomes after implantation9,10,11,12,13,14. Nevertheless, tissue-engineered scaffolds are often comprised of complex and costly components, and secondly, the requirements for ex vivo cell culturing and scaffold seeding are laborious and resource-intensive. These factors have hindered the clinical translation of tissue-engineered scaffolds despite several decades of research within the area. By reducing the complexity as well as monetary and materialistic requirements, tissue-engineered scaffolds could be implemented in modern surgery on a broad scale, addressing both rare and more common procedures.
Collagen has previously been established as a viable platform for cell expansion and, furthermore, acts as a favorable bio-adhesive when attaching cells or tissue onto a scaffold for surgical implantation15,16,17. Perioperative autologous micrografting circumvents the need for ex vivo cell culturing by harvesting the tissue of interest during the primary procedure and re-implanting it directly. By mincing the resected tissue into smaller particles, the surface area and the growth potential is increased, allowing for a larger expansion ratio onto the scaffold18. The collagen-based scaffold does not adhere specifically to urogenital reconstructions but can theoretically apply to multiple areas of hollow-organ reconstruction.
In this manuscript, we present both a protocol for the construction of a tubular scaffold, combining collagen with embedded autologous urothelial micrografts, and a minipig model evaluating the technical feasibility and safety, as well as the regenerative performance, of the scaffold in vivo. The model was evaluated in 10 full-grown female minipigs using the protocol and method presented here. The main advantage of the scaffold is the simplicity of the construct and the single-staged implantation, sparing the patient of several subsequent surgical procedures. The procedure can be performed in conventional surgical settings by regular surgical personnel and requires standard equipment and materials. The animal model allows for a controlled environment for studying the implantation while the animal readily returns to normal behavior, with the added possibility of implementing variations to the scaffold and the procedure.

<table><tbody><tr><td>10x MEM</td><td>Gibco, Thermo Fisher Scientific, Waltham, US</td><td>2517592</td><td>Collagen preparation</td></tr><tr><td>1x MEM</td><td>Gibco, Thermo Fisher Scientific, Waltham, US</td><td>2508924</td><td>Collagen preparation</td></tr><tr><td>Ambu aScope 4 Cysto</td><td>Ambu A/S, Ballerup, DK</td><td>1000682507</td><td>Cystoscope</td></tr><tr><td>Aquaflush ACE stopper</td><td>Abena, Taastrup, DK</td><td>ACE12/220501</td><td>ACE stopper</td></tr><tr><td>Borgal vet inj opl 200 + 40 mg/mL</td><td>Ceva Animal Health A/S</td><td>510460</td><td>Sulfonamide/Trimethoprim</td></tr><tr><td>Bupaq multidose vet 0.3 mg/mL</td><td>Salfarm Danmark A/S, DK</td><td>502763</td><td>Buprenorphin</td></tr><tr><td>Butomidor vet inj 10 mg/mL</td><td>Salfarm Danmark A/S, DK</td><td>531943</td><td>Buthorphanol</td></tr><tr><td>Comfortan vet inj 10 mg/mL</td><td>Dechra Veterinary Products A/S, DK</td><td>492312</td><td>Metadone</td></tr><tr><td>Ethilon suture 3-0</td><td>Ethicon, Johnson &amp;amp; Johnson, New Brunswick, US</td><td>SGBCXV</td><td>Monofilament non-resorbable</td></tr><tr><td>Fentanyl inj 50 &amp;micro;g/mL(hamel)</td><td>Hameln Pharma ApS, DK</td><td>432520</td><td>Fentanyl</td></tr><tr><td>Ketador vet inj 100 mg/mL</td><td>Salfarm Danmark A/S, DK</td><td>115727</td><td>Ketamine</td></tr><tr><td>Metacam inj 20 mg/mL t.cattle/pig/horse</td><td>Boehringer Ingelheim Animal, DE</td><td>6443</td><td>Meloxcicam</td></tr><tr><td>Metacam oral suspension 15 mg/mL pigs</td><td>Boehringer Ingelheim Animal, DE</td><td>482780</td><td>Meloxcicam</td></tr><tr><td>Omnipaque</td><td>GF Healthcare, Oslo, NO</td><td>16173849</td><td>Contrast for CT</td></tr><tr><td>Pancytokeratin CK-AE</td><td>DAKO Agilent, US</td><td>GA053</td><td>Clone AE1/AE3</td></tr><tr><td>PDS suture 3-0</td><td>Ethicon, Johnson &amp;amp; Johnson, New Brunswick, US</td><td>SEMMTQ</td><td>Monofilament slow-resorbable</td></tr><tr><td>Prolene suture 4-0</td><td>Ethicon, Johnson &amp;amp; Johnson, New Brunswick, US</td><td>PGH187</td><td>Monofilament non-resorbable</td></tr><tr><td>Propolipid t.inj/inf 10 mg/mL</td><td>Fresenius Kabi, DK</td><td>21636</td><td>Propofol</td></tr><tr><td>Rat-tail collagen type I</td><td>First Link Ltd, Wolverhampton, UK</td><td>60-30-810</td><td>2.06 mg/mL protein in 0.6% acetic acid</td></tr><tr><td>Suprim vet&amp;nbsp; 20 + 100 mg (Solution for use in drinking water)</td><td>Dechra Veterinary Products A/S, DK</td><td>33661</td><td>Sulfonamide/Trimethoprim</td></tr><tr><td>SX-ELLA Degradable Biliary DV stent</td><td>ELLA-CS, Trebes, CZ</td><td>S23000056-01</td><td>&amp;oslash; 6 mm x 60 mm</td></tr><tr><td>Vicryl mesh</td><td>Ethicon, Johnson &amp;amp; Johnson, New Brunswick, US</td><td>VM1208</td><td>Mesh</td></tr><tr><td>Vicryl suture 4-0</td><td>Ethicon, Johnson &amp;amp; Johnson, New Brunswick, US</td><td>SMBDGDR0</td><td>Braided fast-resorbable</td></tr><tr><td>Xysol vet inj 20 mg/mL</td><td>ScanVet Animal Health A/S, DK</td><td>54899</td><td>Xylazine</td></tr><tr><td>Zoletil 50 vet plv/sol t.inj 25 + 25 mg/mL</td><td>Virbac Danmark A/S, DK</td><td>568527</td><td>Tiletamine and Zolazepam</td></tr></tbody></table>

surgical model for single-staged tissue-engineered urothelial tubes in minipigs

Reconstructive surgeries are often challenged by a lack of grafting tissue. In the treatment of urogenital malformations, the conventional solution has been harvesting gastrointestinal tissue for non-orthotopic reconstruction due to its abundance to reestablish normal function in the patient. The clinical outcomes after rearranging native tissues within the body are often associated with significant morbidity; thus, tissue engineering holds specific potential within this field of surgery. Despite substantial advances, tissue-engineered scaffolds have not yet been established as a valid surgical treatment alternative, mainly due to the costly and complex requirements of materials, production, and implantation. In this protocol, we present a simple and accessible collagen-based tubular scaffold embedded with autologous organ-specific tissue particles, designed as a conduit for urinary diversion. The scaffold is constructed during the primary surgical procedure, comprises commonly available surgical materials, and requires conventional surgical skills. Secondly, the protocol describes an animal model designed to evaluate the short-term in vivo outcomes post-implantation, with the possibility of additional variations to the procedure. This publication aims to demonstrate the procedure step-by-step, with special attention to the use of autologous tissue and a tubular form.

In this study, in vivo urothelial tissue expansion is achieved in a collagen-based tubular scaffold. By embedding the scaffold with autologous tissue particles, harvested and processed perioperatively, the procedure allows for single-staged scaffold implantation without the need for concomitant immunosuppressive treatment postoperatively. Surgical handling is enabled by reinforcing the scaffold with a biodegradable mesh and stent (Figure 1). After 6 weeks of observation, the macroscopical tissue evaluation revealed no signs of host rejection or infection, and the tubular scaffold presents patent and unobstructed (Figure 2). From histological evaluations, a stratified luminal epithelium of urothelial origin is seen covering the entirety of the scaffold, and remnants of the reinforcing biomaterials are still visible after 6 weeks (Figure 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66936/66936fig01.jpg" /> 
Figure 1: Scaffold construction and implantation. The bladder tissue is dissected perioperatively (top left). The minced mucosal micrografts are expanded onto a surgical mesh (top middle) and embedded in solidified collagen (top right). The collagen has been compressed to expel water, and a stent is prepared (bottom left). The scaffold is tubularized around the stent, and an ACE stopper is placed inside the stent (bottom middle). The bladder is partially closed, and the construct is finally incorporated into the bladder at the original site of tissue excision (bottom right). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66936/66936fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66936/66936fig02.jpg" /> 
Figure 2: Scaffold macroscopical evaluation. After 6 weeks, the animal is euthanized, and the scaffold (arrow) is dissected at the skin level (top left). The bladder is filled with contrast (yellow) and CT scanning is performed to evaluate the conduit (arrow) for patency and signs of stricture formation (top right). A cystoscopy is performed via the urethra to evaluate the bladder and the anastomosis (arrow) after 6 weeks (bottom left). The conduit is once more tested for patency by inserting a catheter (arrow) via the external opening and into the bladder (bottom right). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66936/66936fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66936/66936fig03.jpg" /> 
Figure 3: Scaffold microscopical evaluation. The resected conduit is fixated, and orthogonal transverse sections are performed to evaluate the conduit in the proximal-distal direction. After 6 weeks, the conduit lumen (1) is evaluated to confirm epithelialization (magnified top). Remnants of the biodegradable stent (2) and mesh materials (magnified bottom) are still visible at this point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66936/66936fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Surgical Model for Single-Staged Tissue-Engineered Urothelial Tubes in Minipigs

Tissue-engineered implants for reconstructive surgery rarely progress beyond preclinical trials due to laborious ex vivo culturing, which includes complex and expensive scaffold components. Here, we present a single-staged procedure designed for urinary diversion with an accessible collagen-based tubular scaffold containing autologous micrografts.

Reconstructive surgeries are often challenged by a lack of grafting tissue. In the treatment of urogenital malformations, the conventional solution has ...

surgical-model-for-single-staged-tissue-engineered-urothelial-tubes

Nanyang Technological University

Research

JoVE Journal

Bioengineering

261 Views.  Copenhagen University Hospital - Rigshospitalet. Om te beginnen, snel een volgroeid Göttingen minivarken 12 uur voor de operatie. Na het uitvoeren van een standaard laparotomie op de onderste middellijn bij een verdoofd varken, trekt u de interperitoneale urineblaas naar de wond. Voer nu profylactische hemostase uit op de voorste blaaswand en snijd de detrusorspier in met diathermie.Knip vervolgens de binnenste slijmvlieslaag met een schaar af om een volledig wandblaassegment weg te snijden. Sluit d...

Video: Auteur in de schijnwerpers: innovatieve benaderingen voor het verbeteren van de resultaten bij reconstructieve chirurgie voor aangeboren misvormingen en verworven aandoeningen

An Orthotopic Model of Murine Bladder Cancer

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and subsequent cell adhesion. After their bladders are transurethrally catheterized and drained, animals are again catheterized to permit insertion of a platinum wire into bladders without damaging the urethra or bladder. The catheters are made of Teflon to serve as an insulator for the wire, which will conduct electrical current into the bladder to create a burn injury. An electrocautery unit is used to deliver 2.5W to the exposed end of the wire, burning away extracellular layers and providing attachment sites for carcinoma cells that are delivered in suspension to the bladder through a subsequent catheterization. Cells remain in the bladder for 90 minutes, after which the catheters are removed and the bladders allowed to drain naturally. The development of tumor is monitored via ultrasound. Specific attention is paid to the catheterization technique in the accompanying video.

Bladder tumors are established in female mice in a minimally invasive fashion through catheterization, local cauterization, and subsequent adhesion of carcinoma cells to the burn sites.

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and ...

Medicine

Basic Surgical Techniques in the G&ouml;ttingen Minipig: Intubation, Bladder Catheterization, Femoral Vessel Catheterization, and Transcardial Perfusion

The emergence of the G&ouml;ttingen minipig in research of topics such as neuroscience, toxicology, diabetes, obesity, and experimental surgery reflects the close resemblance of these animals to human anatomy and physiology 1-6.The size of the G&ouml;ttingen minipig permits the use of surgical equipment and advanced imaging modalities similar to those used in humans 6-8. The aim of this instructional video is to increase the awareness on the value of minipigs in biomedical research, by demonstrating how to perform tracheal intubation, transurethral bladder catheterization, femoral artery and vein catheterization, as well as transcardial perfusion. 
Endotracheal Intubation should be performed whenever a minipig undergoes general anesthesia, because it maintains a patent airway, permits assisted ventilation and protects the airways from aspirates. Transurethral bladder catheterization can provide useful information about about hydration state as well as renal and cardiovascular function during long surgical procedures. Furthermore, urinary catheterization can prevent contamination of delicate medico-technical equipment and painful bladder extension which may harm the animal and unnecessarily influence the experiment due to increased vagal tone and altered physiological parameters. Arterial and venous catheterization is useful for obtaining repeated blood samples and monitoring various physiological parameters. Catheterization of femoral vessels is preferable to catheterization of the neck vessels for ease of access, when performing experiments involving frame-based stereotaxic neurosurgery and brain imaging. When performing vessel catheterization in survival studies, strict aseptic technique must be employed to avoid infections6. Transcardial perfusion is the most effective fixation method, and yields preeminent results when preparing minipig organs for histology and histochemistry2,9. For more information about anesthesia, surgery and experimental techniques in swine in general we refer to Swindle 2007. Supplementary information about premedication and induction of anesthesia, assisted ventilation, analgesia, pre- and postoperative care of G&ouml;ttingen minipigs are available via the internet at <a href="http://www.minipigs.com">http://www.minipigs.com</a>10. For extensive information about porcine anatomy we refer to Nickel et al. Vol. 1-511.

Basic Surgical Techniques in the G&#246;ttingen Minipig: Intubation, Bladder Catheterization, Femoral Vessel Catheterization, and Transcardial Perfusion

The use of domestic and miniature pigs in science has increased significantly in recent years. By demonstrating how to perform intubation, transurethral bladder catheterization, femoral artery and vein catheterization, as well as transcardial perfusion, we aim to further increase the value of G&ouml;ttingen minipigs in biomedical research.

The emergence of the G&ouml;ttingen minipig in research of topics such as neuroscience, toxicology, diabetes, obesity, and experimental surgery reflects ...

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.

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 ...

Tissue Engineering of a Human 3D in vitro Tumor Test System

Cancer is one of the leading causes of death worldwide. Current therapeutic strategies are predominantly developed in 2D culture systems, which inadequately reflect physiological conditions in vivo. Biological 3D matrices provide cells an environment in which cells can self-organize, allowing the study of tissue organization and cell differentiation. Such scaffolds can be seeded with a mixture of different cell types to study direct 3D cell-cell-interactions. To mimic the 3D complexity of cancer tumors, our group has developed a 3D in vitro tumor test system.	
	Our 3D tissue test system models the in vivo situation of malignant peripheral nerve sheath tumors (MPNSTs), which we established with our decellularized porcine jejunal segment derived biological vascularized scaffold (BioVaSc). In our model, we reseeded a modified BioVaSc matrix with primary fibroblasts, microvascular endothelial cells (mvECs) and the S462 tumor cell line. For static culture, the vascular structure of the BioVaSc is removed and the remaining scaffold is cut open on one side (Small Intestinal Submucosa SIS-Muc). The resulting matrix is then fixed between two metal rings (cell crowns).	
	Another option is to culture the cell-seeded SIS-Muc in a flow bioreactor system that exposes the cells to shear stress. Here, the bioreactor is connected to a peristaltic pump in a self-constructed incubator. A computer regulates the arterial oxygen and nutrient supply via parameters such as blood pressure, temperature, and flow rate. This setup allows for a dynamic culture with either pressure-regulated pulsatile or constant flow.	
	In this study, we could successfully establish both a static and dynamic 3D culture system for MPNSTs. The ability to model cancer tumors in a more natural 3D environment will enable the discovery, testing, and validation of future pharmaceuticals in a human-like model.

Tissue Engineering of a Human 3D in vitro Tumor Test System

Methods to create human 3D tumor tissues as test systems are described. These technologies are based on a decellularized Biological Vascularized Scaffold (BioVaSc), primary human cells and a tumor cell line, which can be cultured under static as well as under dynamic conditions in a flow bioreactor.

Cancer is one of the leading causes of death worldwide.  Current therapeutic strategies are predominantly developed in 2D culture  systems, which ...

Minimally Invasive Establishment of Murine Orthotopic Bladder Xenografts

Orthotopic bladder cancer xenografts are the gold standard to study molecular cellular manipulations and new therapeutic agents&#160;in vivo. Suitable cell lines are inoculated either by intravesical instillation (model of nonmuscle invasive growth) or intramural injection into the bladder wall (model of invasive growth). Both procedures are complex and highly time-consuming. Additionally, the superficial model has its shortcomings due to the lack of cell lines that are tumorigenic following instillation. Intramural injection, on the other hand, is marred by the invasiveness of the procedure and the associated morbidity for the host mouse.
With these shortcomings in mind, we modified previous methods to develop a minimally invasive approach for creating orthotopic bladder cancer xenografts. Using ultrasound guidance we have successfully performed percutaneous inoculation of the bladder cancer cell lines UM-UC1, UM-UC3 and UM-UC13 into 50 athymic nude. We have been able to demonstrate that this approach is time efficient, precise and safe. With this technique, initially a space is created under the bladder mucosa with PBS, and tumor cells are then injected into this space in a second step. Tumor growth is monitored at regular intervals with bioluminescence imaging and ultrasound. The average tumor volumes increased steadily in in all but one of our 50 mice over the study period.
In our institution, this novel approach, which allows bladder cancer xenograft inoculation in a minimally-invasive, rapid and highly precise way, has replaced the traditional model.

The established technique to inoculate primary invasive orthotopic bladder cancer xenografts requires laparotomy and mobilization of the bladder. This procedure inflicts significant morbidity on the mice, is technically challenging and time-consuming. We therefore developed a high-precision, percutaneous approach utilizing ultrasound guidance.

Orthotopic bladder cancer xenografts are the gold standard to study molecular cellular manipulations and new therapeutic agents&#160;in vivo. Suitable ...

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among laboratories. The goal of this study was to describe the microsurgical procedure used to implant an intravesical tube and the experimental technique for recording urinary bladder pressure in an awake, freely moving mouse. In addition, experimental data is presented to show how surgery, as well as tubing type and size, affect lower urinary tract function and recording sensitivity. The effect of tube diameter on pressure recording was assessed in both polyethylene and polyurethane tubing with different internal diameters. Subsequently, the best performing tube from both materials was surgically implanted into the dome of the urinary bladder of male C57BL/6 mice. Twelve-hour, overnight micturition frequency was recorded in healthy, intact animals and animals 2, 3, 5, and 7 days post-surgery. At harvest, bladders were assessed for signs of swelling using gross observation and were subsequently processed for pathological analysis. The greatest extent of bladder swelling was observed on day 2 and 3, which correlated with behavioral voiding data showing significantly impaired bladder function. By day 5, bladder histology and voiding frequency had normalized. Based on the literature and evidence provided by our studies, we propose the following steps for in vivo recording of intravesical pressure and voided volume in an awake mouse: 1) Perform the surgery using an operating microscope and microsurgical tools, 2) Use polyethylene-10 tubing to minimize movement artifacts, and 3) Perform cystometry on post-operative day 5, when bladder swelling resolves.

This study describes the surgical procedures and experimental techniques for performing awake cystometry in a freely moving mouse. In addition, it provides experimental evidence to support its optimization and standardization.

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among ...

Urethral Stricture Induction Followed by Buccal Mucosa Graft Urethroplasty in a Rat Model

Urethral reconstruction is an important area of expertise for urologists. The buccal mucosa is considered the best option when urethral grafting is necessary, although in some cases, it is inappropriate or needs to be optimized to repair a given stricture. Therefore, developing innovative procedures and evaluating their putative success in experimental models is crucial to fit the clinical need. With this goal, this study describes a protocol in which urethral stricture was induced by electrocautery in Wistar rats. Urethral reconstruction was performed 1 week later with a buccal mucosa graft, harvested from the lower lip, and placed in a ventral onlay fashion. A retrograde urethrogram showed a significant improvement in urethral diameter after urethroplasty compared to the respective value after stricture induction. Additionally, the graft placement was assessed by blood perfusion analysis using laser Doppler. As expected, a dark blue area corresponded to the non-vascularized buccal mucosa graft. This procedure can successfully simulate the normal pathophysiological process of urethral injury and tissue modulation, as well as urethral reconstruction using a buccal mucosa graft in a reproducible manner, and serve as the basis for future research based on tissue engineering or urethral grafts.

In the present protocol, a urethral stricture induction was developed in Wistar rats, followed by urethral reconstruction with a buccal mucosa graft. A retrograde urethrogram and laser Doppler assessment were performed, validating urethral reconstruction (after stricture formation) and graft placement.

Urethral reconstruction is an important area of expertise for urologists. The buccal mucosa is considered the best option when urethral grafting is ...

Efficient Rat Model for Urethral Stricture: Electric Knife Method

Iatrogenic Injury Recapitulated: Electroexcision Technique for Urethral Stricture Modeling in Rats

Urethral stricture (US) is a common clinical condition in urology, characterized by high prevalence and morbidity across all ages. Current treatments for US, such as urethral dilatation and internal urethrotomy, fail to fully resolve the condition and are associated with high rates of recurrence and complications. 
Additionally, the pathogenesis of US is not well understood. To explore the pathogenesis of US and develop new therapeutic strategies, it is crucial to establish a standardized rat model that accurately reflects the clinical manifestations. This study outlines a straightforward and repeatable method for inducing US in rats using a high-frequency electric knife. The method involves making a longitudinal incision with the electric knife set to a unipolar mixed cutting mode at 4 W, which inflicts significant urethral damage. Histopathological analysis shows thickening of the urothelium, inflammatory infiltration, and disorganized collagen fibers. This model effectively replicates iatrogenic injury through electroexcision in the rat urethra. In summary, this study successfully establishes a new, efficient, and stable rat model of US that closely mimics the clinical scenario, providing a valuable tool for further research into the mechanisms and novel treatments for US.

In this study, we present a novel, efficient, and stable rat model of urethral stricture created through electroexcision of the rat urethra, which effectively simulates iatrogenic injury observed in clinical settings.

Urethral stricture (US) is a common clinical condition in urology, characterized by high prevalence and morbidity across all ages. Current treatments for ...

An Immature Murine Model of Reversible Unilateral Ureteral Obstruction

Unilateral ureteral obstruction (UUO) is a common cause of chronic kidney disease (CKD), leading to the progression of renal interstitial fibrosis and ultimately resulting in irreversible kidney damage. The alleviation of UUO is crucial. Several animal models of reversible unilateral ureteral obstruction (RUUO) have been established in the literature, enabling the observation of structural changes and functional damage while also simulating physiological and pathophysiological changes following the relief of ureteral obstruction. In this study, a reversible obstruction model was established in the unilateral murine ureter using a silicone tube. Significant renal damage was observed prior to obstruction relief, with partial recovery noted afterward. Unlike UUO, this model prevents progressive hydronephrosis, leading to distinct pathological outcomes. This simple surgical procedure demonstrates a high success rate and holds promise as a classical model for investigating reversible obstructive nephropathy and potential treatments for renal interstitial fibrosis. Furthermore, it provides a practical platform for studying the mechanisms of recovery from obstructive nephropathy, renal cell regeneration, and tissue remodeling.

The present protocol describes a step-by-step, reproducible model of unilateral ureteral obstruction.

Unilateral ureteral obstruction (UUO) is a common cause of chronic kidney disease (CKD), leading to the progression of renal interstitial fibrosis and ...

Patient-derived Tumor Xenografting: A Technique to Generate Experimental Mouse Model for Evaluating Urothelial Cell Carcinoma

Source: Moret, R. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/59223" target="_blank">Patient-derived Orthotopic Xenograft Models for Human Urothelial Cell Carcinoma and Colorectal Cancer Tumor Growth and Spontaneous Metastasis.</a> J. Vis. Exp. (2019)
This video describes the procedure to generate an experimental xenograft murine model, which can be used for studying urothelial cell carcinoma progression.

Surgical Model for Single-Staged Tissue-Engineered Urothelial Tubes in Minipigs

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