Alimentary Tract

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64 Alimentary Tract Richard M. Day University College London, London, United Kingdom

INTRODUCTION The alimentary tract is a hollow organ that starts at the mouth and terminates at the anus. It conducts a number of highly complex and diverse functions that are regulated by distinct cellular and functional differences along the tract length, which allow it to provide the body with nutrients, water, and electrolytes. To achieve this, it is essential that the lumenal contents be propelled along at a rate that will allow efficient digestion and absorption to take place while also enabling waste products to be stored and excreted in a controlled manner. In addition to this, an important symbiotic relationship exists between bacterial species that colonize the alimentary tract of the host [1]. Therefore, the surface of the alimentary tract must also provide a barrier against unwanted entry of organisms and toxins. If the barrier function is breached, specialized cells and tissues within the gut wall provide an important component of the immune system to protect the host. Dysfunction of the alimentary tract may result from a variety of congenital and acquired conditions that can affect any of its physiological functions. This chapter will discuss knowledge regarding tissue engineering different regions of the alimentary tract, highlighting successful strategies as well as failures and some obstacles that have yet to be overcome in this rapidly evolving field.

ESOPHAGUS The esophagus is a muscular tube measuring approximately 25 cm in length in adult humans. It functions primarily as a conduit to connect the pharynx with the stomach, providing coordinated peristaltic contractions in response to swallowing to propel food into the stomach. The esophageal mucosa is lined by stratified, squamous, nonkeratinized epithelium. The submucosa contains muscle, nerve, blood vessels, lymphatics, and mucosal glands. The muscularis has two layers consisting of an outer longitudinal layer and an inner circular layer. Both layers consist of striated muscle in the upper portion and smooth muscle in the lower third, continuous with the muscle layers of the stomach. The myenteric plexus exists between the muscle layers. The esophagus has no serosa and its vascular supply is less extensive compared with the intraabdominal portions of the gut. Sphincters at the upper and lower ends of the esophagus ensure food is transferred appropriately between it and the pharynx or stomach. The upper esophageal sphincter, found in the upper 3e4 cm of the esophagus, and the lower esophageal sphincter, located 2e5 cm above the gastroesophageal junction, remain tonically and strongly constricted to prevent air from entering the esophagus during respiration between swallowing and reflux of stomach contents into the esophagus between peristaltic waves, respectively. Regenerative medicine techniques are being explored for a number of conditions affecting the esophagus. Gastroesophageal reflux disease is one of the most common disorders affecting the gastrointestinal tract, resulting from lower esophageal sphincter incompetence. Medical therapy is generally safe and effective in most cases, but for patients for whom this option fails, antireflux surgery or endoscopic procedures that involve injecting bulking materials may be used in an attempt to narrow the lumen of the lower esophagus. Attempts have been made to restore physiological function of the esophagus using regenerative medicine. The feasibility of using a suspension of muscle precursor cells to restore gastroesophageal function in a model of gastroesophageal reflux disease has been explored Principles of Regenerative Medicine, Third Edition https://doi.org/10.1016/B978-0-12-809880-6.00064-3

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[2]. Muscle precursor cells isolated from expanded satellite cells derived from skeletal muscle fibres were injected into the gastroesophageal junction after cryoinjury. Histology showed an increase in myofibers at the site of injection that had fused into newly formed or preexisting myofibers. The need remains to demonstrate that cells injected in this manner can contribute to functional improvement of damaged esophageal sphincter, but the feasibility of using this approach offers a promising therapy for this common condition. Esophageal reconstruction is a requirement for congenital esophageal atresia, burns, malignancy, or severe benign disease. Surgical techniques include stretching, circular myotomy, and interposition of stomach or colon, but these approaches are frequently associated with complications including stricture, leakage, elongation, and gastroesophageal reflux. An artificial esophageal construct has been sought for many years. To be effective, the construct must be implantable without rejection and biocompatible to support appropriate tissue growth, and retain biomechanical characteristics of native esophageal tissue, i.e., be soft and elastomeric while maintaining a tubular structure when implanted in vivo. Attempts to tissue engineer replacement esophageal tissue have included both patch and circumferential implantation of constructs composed of synthetic as well as natural scaffold materials. A full-length of tissue engineered esophagus has not been produced, but a number of incremental advances toward this goal have been achieved. Early attempts exploring the use of a nondegradable prosthetic tubes in canine models were of limited success [3]. Surgical reconstruction techniques of the esophagus after resection for structures and malignancies have involved the transfer of small segments or patches of skin and other tissues on a vascular pedicle, with moderate results [4e6]. Tissue engineered sheets of autologous oral mucosal epithelial cells were successfully transplanted by endoscopy in a canine model [7]. The transplanted sheets adhered to the underlying esophageal muscle layers created by endoscopic submucosal dissection and enhanced wound healing without postoperative stenosis. Because the interaction between the epithelium and mesenchymal cells is thought to reduce fibrosis and scarring that cause stenosis, the investigators suggested that this approach may offer a novel therapy to reduce scarring and prevent painful constriction that can be associated with endoscopic submucosal dissection for the removal of large esophageal cancers. A variety of scaffold materials to support cell and tissue esophageal constructs have been investigated [8e12]. Acellular scaffolds composed of extracellular matrix components have been explored primarily because they are assumed to be advantageous over synthetic scaffold materials owing to their ability to promote cell attachment, growth, and cellecell signaling among different tissue components. Decellularized esophageal tissue can be produced via repeated detergent-enzymatic treatment that results in a scaffold with biocompatibility suitable for the growth of esophageal epithelial cells [13,14]. Based on these preclinical findings, it has been envisaged that human donor esophageal tissue might one day be used in a manner similar to that described for the tissue engineering of human airway tissue [15]. Scaffolds derived from small intestinal submucosa (SIS) have been widely investigated for tissue engineering replacement esophageal constructs. SIS consists of extracellular matrix material harvested from porcine small intestine and has been used extensively in tissue engineering experiments since it was originally described by Matsumoto and colleagues in 1966 for use in large vein replacement in dogs [16e20]. It has been successfully applied to regenerative medicine applications in humans, including repair of hernias, diaphragms, and tympanic membranes, and for large wound coverage [21e24]. The success with using SIS as a scaffold to promote tissue regeneration appears to relate to the retention of collagen (types I, II, and V), growth factors (transforming growth factor, fibroblast growth factor 2, and vascular endothelial growth factor), glycosaminoglycans (hyaluronic acid, chondroitin sulfate, and heparin sulfate), proteoglycans, and glycoproteins (fibronectin) during the fabrication process [25,26]. It is thought that the resulting scaffold has a composition closely resembling native extracellular matrix, which makes it ideally suited for the attachment and growth of new tissue. The extent of circumferential replacement of esophageal tissue appears to have an impact on the outcome of attempts to tissue engineer esophagus, with patches producing better results compared with tubular segments. Lopes and colleagues successfully used SIS patches to repair defects to the anterior wall of cervical or abdominal esophagus in rats without signs of stenosis over 150 days [27]. Likewise, Badylak and colleagues used SIS patches (or urinary bladder submucosa) to repair esophageal defects created in dogs without clinical signs of esophageal dysfunction [28]. However, the latter study reported signs of stenosis in dogs receiving complete circumferential segmental grafts of SIS [10]. Doede and colleagues reported similar findings, with severe stenosis occurring when relatively short tubular lengths (4 cm) of SIS were used in alloplastic esophageal replacement in piglets [28]. Thus, although scaffolds consisting of only extracellular matrix have shown the capacity to promote cell growth in vitro and tissue regeneration of patch defects in vivo, replacement of circumferential defects without stricture

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formation remains difficult to achieve. The outcome of repairing full circumferential defects using SIS may be improved by optimizing the poor mechanical properties of the scaffold material. To address this, hybrid scaffolds composed of SIS combined with synthetic polyesters (poly[3-hydroxybutyrate-co-3-hydroxyhexanoate] and poly [lactide-co-glycolide]) were assessed for their feasibility as a tissue engineering scaffold for esophageal constructs [29]. Improved biocompatibility was observed with the hybrid scaffold compared with scaffolds produced from the synthetic material alone. Epithelial-mesenchymal cell signaling is likely to have an important role in facilitating reconstruction of the esophageal construct after implantation. A similar effect has been shown in bladder reconstruction, in which the presence of urothelium led to infiltration of fibroblasts into acellular matrices and apparent transdifferentiation into a smooth muscle phenotype [30]. Signaling from the mesenchymal cell population appears to be equally important in promoting growth of overlying epithelium [31]. Complete reepithelialization with little inflammatory response and evidence of skeletal muscle regeneration was observed when bone marrow mesenchymal stem cells were seeded onto the SIS scaffold implanted in a canine model [32]. Moreover, the presence of epithelialmesenchymal signaling may prevent stricture formation in an esophageal construct, a problem frequently encountered with many of the scaffolds tested [10,33]. Similar signaling properties have been demonstrated in bladder reconstruction in which acellular collagen scaffolds seeded with urothelium and smooth muscle cells prevented tissue contraction [34]. Likewise, the interaction of muscle with the ablumenal surface of esophageal scaffolds at the time of implantation of partially circumferential grafts appears to have accounted for the reduced stricture formation observed in a canine model of esophageal reconstruction described by Badylak and colleagues [11]. It can be concluded from these observations that careful consideration of the order in which cells are added to the tissue engineered construct will improve the likelihood of achieving a successful outcome. In addition to SIS, gastric acellular matrix was used as scaffold by Urita and colleagues to regenerate esophagus in a rat model [35]. Grafts of gastric acellular matrix were used to patch defects in the abdominal esophagus and animals were killed at points between 1 week and 18 months. Although regeneration of the muscle layer or lamina muscularis did not occur, there was no evidence of stenosis or dilatation at the graft site. The matrix obtained in this study was from whole stomachs, but the investigators suggested that gastric acellular matrix may provide an autologous source of naturally derived extracellular matrix scaffold in a clinical setting, because the portion of stomach destroyed to obtain the matrix is minimal. It remains to be seen whether this approach is feasible in a larger animal model, but the use of autologous acellular matrix scaffolds avoids concerns related to the use of xenogenic scaffold materials such as porcine-derived SIS. In addition to the risk of transmitting viral pathogens and prions, cultural and religious beliefs may need to be considered when using acellular matrix scaffolds derived from certain species. In addition to SIS, other biological materials have been investigated for esophageal tissue engineering. Extracellular matrix scaffold has been generated from ovine forestomach tissue [36]. Moving away from mammalian sources of scaffold material, Franck and colleagues reported that a bilayer silk fibroin matrix composed of porous silk fibroin foam annealed to a homogeneous silk fibroin film exhibited improved cell attachment and spontaneous differentiation of esophageal epithelial cells toward a suprabasal cell lineage compared with SIS scaffolds [37]. Although the esophagus can be considered one of the less complex regions in the alimentary tract, several significant hurdles still need to be overcome before tissue engineering and clinical replacement of full-length esophageal segments become a clinical reality in humans. Unlike patch grafts, replacement of longer lengths of tissue will be unable to rely on adjacent esophagus to cover the surface area of larger scaffolds via guided tissue regeneration. Improved methods from isolating and expanding the different esophageal cell populations will therefore be a prerequisite for successful tissue engineering of larger constructs. Kofler and colleagues identified subsets of ovine esophageal epithelial cells that may help achieve this [38]. PCK-26epositive esophagus epithelial cells demonstrated high proliferative capacity and uniform coverage on collagen scaffolds, which the investigators suggested could have an important role for successfully tissue engineering esophagus. Further refinement of the scaffold material may also improve epithelialization. For example, the inclusion of copper into acellular porcine SIS scaffolds was reported to enhance epithelialization of the scaffold in a canine model of esophageal repair [39]. Failure to regenerate a functioning muscle layer may not be problematic for short or noncircumferential grafts, but for longer lengths of esophagus the presence of an innervated functional muscle layer will be essential. A retrospective study investigating the temporal appearance and spatial distribution of nervous tissue in a canine model of esophageal reconstruction using porcine urinary bladder submucosa showed the presence of nerve tissue within sites of the remodeling scaffold [40]. Although the study was unable to demonstrate whether the nervous tissue was functional or to distinguish among the various subsets of neurons, it opens the possibility of using similar models to identify mechanisms that promote innervation that will facilitate the tissue engineering of functional tissue. Peristalsis of food also depends on the correct orientation of muscle fibers in the wall of the alimentary tract.

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To address this, promising results have been obtained with orienting smooth muscle tissue on unidirectional scaffolds for tissue engineered esophagus in rats [41]. Oriented stands of smooth muscle mimicking the configurations found in the native organ were engineered when cells were seeded onto unidirectional scaffolds. These were assembled with esophageal epithelium to create a hybrid approach.

STOMACH Gastric disease affects approximately 10% of the world’s population and includes gastritis, peptic ulcers, and gastric cancer. Insufficient stomach mass, which may arise from gastrectomy or congenital microgastria, is associated with increased patient morbidity. The stomach functions as a digestive organ and reservoir and is anatomically divided into four regions (cardiac, fundus, corpus, and pylorus). The gastric glands are tubular structures whose cellular composition and function are specialized according to each region of the stomach. Such specialization, combined with the harsh environment created by the lumenal contents, makes tissue engineering of the stomach as whole challenging. The size and shape of the stomach vary, depending on its contents. The stomach wall contains outer longitudinal and inner circular layers of smooth muscle, with an innermost layer of oblique muscle fibers. These layers facilitate important functions including storage of ingested food in the stomach until it can be accommodated in the lower portion of the alimentary tract, mixing of the food to form chyme, and regulation of food transit into the small intestine at an optimal rate for digestion and absorption. Stomach emptying is controlled by the gastric food volume and the release of the hormone gastrin, as well as feedback signals from the duodenum. Tissue engineered neostomach constructs to patch partial gastrectomy have been explored in a canine model using a two-part sheet composed of an outer layer of collagen sponge and a temporary inner silicone sheet to protect the collagen from degradation by the acidic stomach juices and provide mechanical support [42]. After removal of the silicone sheet at 4 weeks, evidence of stomach regeneration was observed and complete coverage of the scaffold had occurred by 16 weeks, confirmed by the presence of mucosa and a thin muscular layer. Acid production capacity was present in the regenerated stomach wall but the contractile response to acetylcholine was poor [43]. Technical difficulties associated with suturing and endoscopic removal of the silicone sheet in this model were addressed by creating a tissue engineered sheet without silicone that had sufficient strength to allow suturing and resist anastomotic dehiscence [44]. The silicone sheet was replaced by a biodegradable copolymer of poly(D,L-lactide) and ε-caprolactone (PDLCL) on the mucosal side of the collagen scaffold, both of which were completely absorbed at 16 weeks’ implantation. Although regeneration of the stomach mucosa was observed, the replacement of the silicone sheet with PDLCL did not provide sufficient mechanical strength to prevent significant shrinkage of the scaffold. The feasibility of creating new stomach tissue using stomach-derived organoid units harvested from neonatal and adult rats has been investigated [45]. The organoid units were seeded onto polymer scaffold tubes to form constructs that were implanted into the omentum of adult syngeneic rats. At 4 weeks, the construct was anastomosed to the small intestine. Histology of the tissue engineered stomach tissue was similar to native stomach, with gastric pits, squamous epithelium, and positive staining for a-actin smooth muscle in the muscularis and gastrin indicating the presence of a well-developed gastric epithelium. The same approach has been used to tissue engineer stomach neoconstructs in an autologous large animal model [46]. A limitation of approaches using autologous or allogenic organoid units is the tissue source. To address this, three-dimensional gastric organoid tissue from human pluripotent stem cells (PSCs) have been generated by temporal manipulation of the fibroblast growth, WNT, bone morphogenetic protein, retinoic acid and epidermal growth factor (EGF) signaling pathways [47]. The primitive gastric organoids exhibited molecular and morphogenetic developmental stages similar to those observed in the developing antrum of murine stomach. The proliferative zones contained LGR5þ cells, mucus-secreting antral and pylorus cells, and gastric endocrine cells; however, the acid secreting (corpus) region was not developed in this model. A critical factor for the development of all the stomach regions from stem cells appears to be the Barx-1 gene [48]. Noguchi and colleagues demonstrated Barx-1einducing culture conditions generated spheroids of fully functional stomach-like tissue in vitro from mouse embryonic stem cells. The spheroids were able to develop into functional corpus and antrum tissue that secreted pepsinogen and acid. The relevance of mesenchymal stem cells to optimize and condition the cellular milieu within the tissue engineered construct has also been shown with stomach tissue engineering. Tissue regeneration after the creation of a full-thickness stomach defect in rats was enhanced when SIS scaffolds were used in conjunction with mesenchymal

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stem cells [49]. Whereas contractility in response to a muscarinic receptor agonist, a nitric oxide precursor, or electrical field stimulation was observed in all groups, smooth muscle layers were both longer and better structured compared with SIS grafts not seeded with mesenchymal stem cells.

SMALL INTESTINE The small intestine in adults measures approximately 6 m in length from the duodenojejunal flexure to the ileocecal valve. Its primary function is absorption of nutrients from the lumen, a process facilitated by specialized mucosal surface features (folds of Kerckring, villi, and microvilli) that increase the absorptive surface area about 600-fold to approximately 250 m2. The mucosa is lined with epithelium overlying lamina propria containing vascular and reticular stroma, large aggregates of lymphoid tissue called Peyer patches, and a strip of smooth muscle called the muscularis mucosa. Intestinal stem cells reside at the base of epithelial invaginations into the mucosa called crypts and develop into all four lineages of epithelial cells that line the intestine [50]. Epithelial cells migrate along the cryptevillus axis, differentiating and maturing toward the lumen of the bowel where they become senescent over the course of a few days and are shed into the lumen of the bowel. A lack of intestinal epithelial stem cell markers has hampered identification and isolation of pure populations of cells for regenerative medicine purposes. Studies have shown that Musashi-1 may be a marker of intestinal stem cells [51,52], and a Sox9-(enhanced green fluorescent protein [EGFP]) mouse model has been used to enrich multipotent intestinal epithelial stem cells [53]. Using a culture system that mimics the native intestinal epithelial stem cell niche, these cells are capable of generating “organoids” that contain all four epithelial cell types of the small intestinal epithelium. Furthermore, Sox9-EGFP multipotent intestinal epithelial stem cells express CD24, which may facilitate their enrichment by fluorescence activated cell sorting using widely available antibodies. The submucosa consists of fibrous connective tissue that supplies blood and lymphatic vessels to the mucosa. The muscularis propria consists of an inner layer of circular muscle and an outer longitudinal muscle layer. The muscularis propria is covered by the adventitia, a layer of loose connective tissue, and the serosa, a mesothelial lining of peritoneum. The function of the small intestine cannot be replaced by transposing another part of the gut. Intestinal ischemia and bowel resection for tumors and inflammatory bowel disease can result in short bowel syndrome when more than 75% of the small intestine is lost. Short bowel syndrome is often associated with intestinal failure and the requirement of lifelong nutritional support (total parenteral nutrition), which is frequently accompanied by severe complications such as liver failure, line sepsis, and poor long-term survival rates. The length of residual intestine is critical for these patients; thus, techniques for increasing absorptive surface area have been sought for many years. Surgical options for increasing the absorptive surface or slowing the transit time to enhance absorption have been reported, but these approaches require longer residual intestinal segments and most have only limited long-term clinical success [54e57]. Small bowel transplantation is a viable option for some patients, but this procedure has limitations including the availability of donor tissue, the need for long-term immunosuppression, graft versus host disease, and potential posttransplant lymphoproliferative disorder [58]. The amount of small bowel required for successful nutritional rehabilitation depends on factors including the patient’s age, the amount of small bowel present, the presence or absence of the ileocecal valve, and the amount of large bowel present. Therefore, small bowel elongation of just a few centimeters could allow many patients to become independent of total parenteral nutrition. Distraction enterogenesis has been devised as a novel method to increase intestinal length by applying linearly directed force, resulting in increased surface area and epithelial cell proliferation [59,60]. Devices used for distraction enterogenesis include extralumenal, radially selfexpanding shape memory polymer cylinders [61]; biodegradable springs composed of polycaprolactone created to lengthen intestinal segments mechanically while avoiding the need for subsequent retrieval [62]; and doubleballoon catheter devices [63]. Biological mechanisms that account for distraction enterogenesis are unknown [64], but the effect can be enhanced by adding exogenous glucagon-like peptide 2 [65] or by codelivering microspheres that provide sustained release of basic fibroblast growth factor (bFGF), which results in improved vascularity [66]. The use of growth factoreembedded scaffold materials is an effective method for improving the short half-lives of growth factors. An example used to achieve growth factoreembedded scaffolds for distraction enterogenesis include subcritical CO2 to embed heparin-binding EGF-like growth factor into polyglycolic acid/ poly-L-lactic acid scaffolds [67]. Local delivery of the trophic growth factor improved structure of the tissue engineered intestine.

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Several different approaches, using either guided tissue regeneration or tissue-engineering neotissue constructs, were taken to regenerate small intestine that used combinations of various synthetic and natural scaffold materials, different cell types, and surgical procedures. Early attempts to patch bowel defects using the serosal surface of another piece of intestine resulted in its being covered with regenerated mucosa [68,69]. This paved the way for other researchers to investigate a variety of scaffold materials, such as polytetrafluoroethylene tubing, for the ingrowth of neointestine via guided tissue regeneration [70]. The use of nonresorbable materials for studying intestinal morphogenesis and regeneration continues to be interesting [71], but the use of resorbable scaffold biomaterials for intestinal tissue engineering has become the predominant approach. Chen and Badylak used SIS to patch partial defects in the small bowel wall of a canine model [72]. Histological evaluation showed the presence of mucosa, varying amounts of smooth muscle, sheets of collagen, and an outer serosal layer. However, in the same study, attempts to use a tubular configuration of SIS were unsuccessful. The tubes either leaked or became obstructed; this occurred primarily because the SIS was unable to maintain lumenal patency when exposed to the moist lumenal contents. Similar limitations with the mechanical integrity of the scaffold material were reported by Pahari and colleagues, who used guided tissue regeneration to create a segment of new intestine in rats using acellular dermal matrix (AlloDerm) rolled into tubes [73]. Building on the work, elongation of the intestine was achieved using an acellular biologic scaffold to create autologous bioartificial intestinal segments (BIS) [74]. The BIS were demonstrated to have functional absorptive characteristics [75]. Other approaches using biologically derived scaffolds included the use of allogenic aortic graft segments interposed in an excluded small bowel segment wrapped in omentum, which resulted in intestinal-like wall transformation of the aortic graft [76]. In an attempt to maintain an open lumen in the tissue engineered intestine, Hori and colleagues reported that scaffolds composed of sheets of acellular collagen sponge wrapped on a temporary silicone stent and covered with omentum guided tissue regeneration of almost all layers of the gastrointestinal tract in a canine model, but only a thin muscularis mucosa was present and the muscularis propria was absent [77]. The same group explored the addition of mesenchymal stem cells seeded onto a collagen scaffold, which it was hypothesized might differentiate “site-specifically” into muscle cells and regenerate the muscle layer [78]. Intestinal regeneration occurred but muscle regeneration in an organized manner was not observed. Wang and colleagues used a rat model to evaluate the feasibility of regenerating tubular intestine using sheets of rat-derived SIS wrapped around a silicone stent [79]. The tubular graft was interposed in the middle of a Thiry-Vella loop (a defunctionalized segment of ileum that is brought out as a double ileostomy) in Lewis rats. The silicone stent was left in place for 3 weeks to maintain lumenal patency during tissue regeneration. At 4 weeks, an epithelial layer had begun to form and completely covered the lumenal surface by 12 weeks. The neomucosa had a typical morphology containing goblet cells, Paneth cells, enterocytes, and enteroendocrine cells. Although the regenerated bowel contained bundles of smooth muscle-like cells, especially near the sites of anastomosis, the quantity and organization of the muscle layer differed from those found in native small intestine; they were predominantly circular muscle with no longitudinal muscle. The use of a ThiryVella loop in the model created by Wang may have facilitated mucosal development in the neointestine by protecting it from alimentary transit and creating an isolated environment that avoided the food stream and digestive enzymes. Lee and colleagues observed only minimal intestinal regeneration in a rat model used to evaluate SIS scaffolds. From this, they concluded that SIS scaffolds alone were insufficient to regenerate small intestine and suggested that the use of appropriate progenitor cells is probably necessary to facilitate the regeneration of small intestine [80]. Many of the studies reporting intestinal tissue engineering strategies are based on methodologies used in the pioneering work conducted by Vacanti and colleagues in the 1980s and 1990s that combined intestinal tissue with scaffolds [81]. Important to these studies were previous investigations by Tait and colleagues that showed intestinal tissue could be separated by enzymatic digestion to produce organoid units [82]. These clusters of cells contained all of the elements of the intestinal mucosa including stem cells and mesenchyme, which could be used to regenerate intestinal neomucosa expressing digestive enzyme activities and glucose transport capacity similar to those of agematched native intestinal mucosa. When organoid units were subcutaneously grafted, they displayed different epithelial populations consistent with epithelial transit amplifying and stem cell populations [83]. Subsequent studies demonstrated that transplanting organoid units onto biodegradable polymer scaffolds followed by implantation into the omentum of syngeneic adult animals resulted in the formation of neointestinal cysts attached to a vascular pedicle with mucosa facing a lumen that contained mucoid material [84]. The mucosa of the neointestine created with this technique showed morphological similarities to native intestine, including the formation of a primitive cryptevillus axis lined with columnar epithelial cells and goblet cells and a polarized epithelium with the brush border enzyme sucrase expressed at the apical surface and laminin at the basolateral surface, and

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transepithelial resistance similar to that of native intestine [85]. For all of these promising findings, it is essential that the tissue engineered construct facilitates nutrient absorption. Anastomosis of the tissue engineered cyst-like structures to native jejunum in adult rats provided continuity with the native intestinal tract, which resulted in a more developed neomucosa containing significant increases in villus number, villus height, and surface length of the cyst compared with nonanastomosed cysts [86]. The investigators postulated that anastomosis may have facilitated neomucosal growth in the cysts by draining lumenal contents or via stimulatory factors present in the lumenal contents of the native intestine in continuity with the neomucosa. The anastomosed neointestine was also shown to express of Naþ-dependent glucose transporter SGLT1 [87] and a mucosal immune system with intraepithelial and lamina propria immune cells similar to that of native jejunum [88]. The native small intestine has a great adaptive and compensatory capacity in response to massive small bowel resection, which is considered to be controlled by humoral factors. The mucosa of the neointestine was also shown to possess this adaptive capacity after massive small bowel resection, resulting in a significant regenerative stimulus for the morphogenesis and differentiation of the tissue engineered intestine [89]. An improvement in intestinal function, capable of facilitating patient recovery after massive small bowel resection, was putatively demonstrated when cysts containing neointestine were anastomosed to native small bowel during an 85% enterectomy in rats [90]. The study showed that animals with tissue engineered intestine returned to their preoperative weight more rapidly compared with animals undergoing small bowel resection alone. These findings significant because they are the first to suggest that tissue engineered intestine may provide a therapeutic intervention for managing patients with short bowel syndrome. Although it is tempting to speculate that the observed effects resulted from neointestine restoring absorptive function after small bowel resection, the mechanism underlying the beneficial effects remain uncertain [91]. It has been postulated that the amount of intestine replaced by the anastomosed neointestine (approximately 4 cm) was far shorter than the amount resected, probably approximately 10% of the original length and is unlikely to have added sufficient mucosal surface area to account for the increase in postoperative weight observed. Furthermore, the improved nutrition may have resulted from the tissue engineered intestine slowing intestinal transit, leading to increased absorption and weight gain, a principle that could be achieved with simpler remedial surgical procedures [91]. To gain a better understanding of the mechanisms underlying the formation of neointestine, the model has been transitioned from a rat to a mouse model, which has enabled the use of transgenic tools for lineage tracing. This demonstrated that epithelium, muscularis, nerves, and blood vessels are derived from multicellular organoid units derived from donor small intestines of transgenic mice [92]. Studies also investigated the effects of donor age and region where intestine crypts are harvested [93]. In mice, higher efficiency of enterosphere formation was observed with crypts harvested from tissue collected from the proximal small intestine, and also in young mice. A significant drawback with this approach is the need for large amounts of donor tissue to harvest a sufficient number of organoid units to seed scaffolds that will generate a relatively short length of neointestine, which is likely to offer only limited therapeutic value [91]. A solution might exist with the use of yet unexplored alternative sources of intestinal epithelial stem cells, such as bone marrowederived cells and PSCs circulating in the peripheral blood [94,95]. New methods for generating organoid units that address the limitations of harvesting donor intestinal tissue have also been explored. McCracken and colleagues reported a protocol for differentiating human PSCs into threedimensional (3D) human intestinal tissue, developing into intestinal tissue containing all major types of intestinal epithelial cells and mesenchymal components [96]. To address challenges associated with the limited availability of autologous donor intestine in patients with short bowel syndrome, protocols exist for generating enteroids from minimal quantities of starting material [97]. Also, refinement of the harvested donor stem cells or manipulation of growth factors in the local environment may provide a method for enhancing the quality of the neointestine. The morphology of tissue engineered intestine was improved by seeding scaffolds with intestinal stem celleenriched crypts [98]. Greater circumferential mucosal engraftment and an average villous height closer to native intestine were achieved with the purified crypts collected using a filtration-based system compared with scaffolds seeded with a villous fraction containing differentiated epithelial cells. Likewise, manipulation of the expression of growth factors that control the growth and differentiation of the intestine during development might provide a valuable approach for improving the formation of tissue engineered small intestine. Tissue engineered organoid units overexpressing fibroblast growth factor 10 resulted in larger tissue engineered constructs, with longer villi and a greater proportion of proliferating epithelial cells [99]. Organoid units derived from human postnatal, small bowel resection specimens, were seeded onto biodegradable scaffolds and implanted into nonobese diabetic/severe combined immunodeficiency g chainedeficient mice [100]. After 4 weeks, the tissue engineered small intestine contained the four major types of differentiated intestinal epithelial cells, muscularis, and intestinal subepithelial myofibroblasts.

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Bone marrowederived epithelial cell adhesion moleculeepositive, and CD133-positive cells were used to recellularize human cadaveric small bowel specimens that had been chemically decellularized [101]. After recellularization, the tissue engineered small intestine contained mucin-positive goblet cells, cytokeratin 18epositive epithelial cells in villi, and smooth muscle cells in muscularis mucosa. Cryopreservation of the organoid units could be used to delay production of tissue engineered small intestine, a procedure that could be particularly helpful in patients who are critically ill and require delayed autologous implantation of a tissue engineered construct. Cryopreservation using vitrification led to higher viability of organoid units compared with standard snap-freezing; the thawed organoid units were capable of producing tissue engineered small intestine [102]. Another important aspect of intestinal tissue engineering is the ability for the neointestine to repair, regenerate, and remodel. The latter is particularly important when considering the use of engineered intestinal tissue for children, in whom the length of the intestine increases significantly during development. The trophic effects of glucagon-like peptide-2 (GLP-2) have been evaluated on neointestinal growth [103]. GLP-2 is an endogenous regulatory peptide with potent trophic effects on intestinal mucosal growth and an ability to modulate the expression of Naþ-glucose cotransporter 1 (SGLT1). Adult rats with neointestinal implants that received subcutaneous injections of a GLP-2 analog twice daily for 10 days had enhanced mucosal growth and increased expression of SGLT1 compared with control rats. These findings indicate that the neointestine is capable of responding to external regulator signals that could be used to further expand the surface of the neointestine. There is a lack of preclinical models for observing intestinal tissue regeneration and improved intestinal function on a scale that can be feasibly translated into humans. Intestinal tissue engineering has been investigated using a large-animal model designed to emulate conditions required for human therapy [46]. Tissue scaffolds were seeded with organoid units isolated from the jejunum of 6-week-old piglets and implanted into the omentum of that animal. However, the study provided only limited information on issues related to the scaling-up of a technique for use in humans because the neointestine was not anastomosed to the native intestine and the scaffolds used were similar in size to those used in previous small-animal models. A functional mucosal barrier is an essential element of intestinal tissue engineering for which scalability also needs to be considered. Although transepithelial resistance of the mucosa created in neointestinal cysts is similar to that of native intestine [85], the creation of larger intestinal constructs will require rapid coverage of the scaffold surface to ensure the barrier function is established. This process might be accelerated by including materials in the scaffold that promote epithelial cell spreading. Yoshida and colleagues investigated the effect of transplanting organoid units onto denuded colonic mucosa of syngeneic recipient rats [104]. The addition of bFGF facilitated neomucosal growth and improved restoration of intestinal epithelial cell coverage over the denuded mucosa compared with the control group. Other approaches might include including inorganic materials into hybrid scaffolds, such as bioactive glass, shown to increase epithelial cell migration via bFGF in an indirect manner [105]. As well as stimulating regeneration of the mucosa, delivery of growth factors may provide a strategy for regenerating the muscularis propria. Local delivery of bFGF from scaffolds, via either incorporation into the collagen coating of scaffolds or encapsulation into microspheres, was also shown to increase the engraftment and density of seeded smooth muscle cells and blood vessel formation after 28 days’ implantation in the omentum of rats [106]. Rapid vascular in growth into the tissue engineered intestine will be essential to maintain the viability and engraftment of cells seeded on the scaffold. Gardner-Thorpe and colleagues observed that tissue engineered intestine exhibited lower levels of bFGF and vascular endothelial growth factor (VEGF) and a fixed capillary density compared with native juvenile bowel [107]. This led that group to evaluate a polymeric microsphere system to deliver encapsulated VEGF and stimulate angiogenesis in the maturing neointestine [108]. Capillary density in the muscular and connective tissue layers was significantly increased in the presence of microspheres containing VEGF, as were the size and weight of the constructs. Interestingly, the rate of epithelial cell proliferation also increased in constructs implanted with VEGF-releasing microspheres, possibly related to the improved vascularization of the construct providing greater nutritional support to the rapidly proliferating epithelium. The need for neovascularization is not restricted to tissue engineering tissues of the alimentary tract. A number of different approaches are being used to tackle this problem [109]. It remains to be seen whether any of these approaches will provide a sufficient stimulus to promote arteriogenesis required for sufficient vascularization of larger tissue constructs. Furthermore, a functional lymphatic system in the neointestine is essential to establish normal nutrient absorption, fluid homeostasis, and immunological functions. Lymphangiogenesis is reported to occur in the neointestine created by the organoid unit-cyst model in rats [110]. Although angiogenesis has been demonstrated in intestinal tissue engineering using small-animal models, it is not certain whether the provision of thin-walled endothelium lined structures will be sufficient to support the functionality of a larger tissue construct. Therefore,

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techniques to promote the formation of medium-sized blood vessels via arteriogenesis are likely to be required to facilitate complete integration of large-scale intestinal constructs with a functional capacity. The small bowel has an extensive vascular system fed by arcades of arteries in the mesentery derived from the superior mesenteric artery. Translation of the existing tissue engineering models to a scale suitable for implantation into humans will require the formation of a similar vascular system consisting of medium-sized blood vessels to maintain viability of a larger tissue construct as well as enable absorption of fluid and dissolved nutrient material from the intestine into the portal blood, which will require a vascular system similar to that found in native intestine. One approach to enabling immediate perfusion of the tissue engineered construct might involve using the existing vascular system in decellularized tissue. Preservation of the vascular structure in decellularized porcine small bowel has been used to engineer tissue via innate vascularization [111]. The decellularized scaffold was repopulated by endothelial cells and exhibited patent vessels after arterial and venous microanastomosis. Improved methods for seeding and maturing larger tissue engineered intestinal constructs will be needed to ensure that the limited cells available are delivered to the tissue construct efficiently and uniformly. These obstacles may be overcome with the development of bioreactor systems that will assist with the long-term culture and bioengineering of tissues by providing an in vitro environment that is similar to normal physiological conditions. Kim and colleagues designed a perfusion bioreactor specifically for intestinal tissue engineering [112]. The use of bioreactors that provide magnetic force may also be used to deliver cells labeled with superparamagnetic iron oxide nanoparticles into hollow tubular scaffolds with more uniform distribution [113]. Techniques used to tissue engineer vascular grafts might also provide solutions that can be translated to intestinal tissue engineering. For example, centrifugal casting onto decellularized laser-porated natural scaffolds has been reported to enable the rapid fabrication of tubular tissue in a bioreactor-free manner [114]. The type of scaffold material chosen for tissue engineering is an important consideration. An optimal scaffold material must be capable of withstanding the intestinal microenvironment, which poses significant challenges in terms of biocompatibility, mechanical properties, and longevity. It must allow transplanted cells to engraft and proliferate rapidly while enabling tissue perfusion of nutrients and remodeling to ensure complete integration with the host. The composition, geometry, and topography of scaffolds used for intestinal tissue engineering may influence the properties of cells grown on their surface. Compared with natural extracellular matrixederived scaffolds, biodegradable synthetic polymer scaffolds provide more control over scaffold properties, such as scaffold architecture, degradation rates, and mechanical properties. Boomer and colleagues evaluated a selection of synthetic tubular scaffolds composed of poly(glycolic acid), poly(ε-caprolactone), poly(L-lactic acid), and polyurethane with either nanofiber or macrofiber structures [115]. Implantation of the scaffolds into the peritoneal cavity of rats revealed different rates of tissue infiltration and scaffold degradation. The inclusion of extracellular matrix components such as hyaluronic acid (a nonsulfated glycosaminoglycan found ubiquitously in connective, epithelial, and neural tissue) was found to enhance the physicochemical properties of gelatinecollagen scaffolds, including attachment, growth, and viability of Caco-2 cells [116]. Histological organization of cells resembling intestinal circular and longitudinal smooth muscle has also been achieved using scaffolds that consist of two layers of orthogonally oriented fibers [117]. The function of the geometry of the cryptevillus microenvironment in regulating intestinal cell proliferation and differentiation was explored by Wang and colleagues [118]. Caco-2 cells migrating over microwell structures showed increased metabolic activity and lower levels of differentiation compared with cells cultured on flat surfaces, which suggested that the structure of crypts may have a role in retaining a proliferative phenotype. Likewise, scaffold architecture is a parameter that can be used to enhance cell infiltration and mass transfer of nutrients to ensure the viability of tissue is maintained. The porosity of the scaffold has a critical role in cell survival and ultimately the viability of the tissue construct. This was illustrated in a study looking at the inclusion of 250-mm pores in multilayered electrospun scaffolds [119]. When implanted in vivo, scaffolds with greater macroporosity were associated with increased blood vessel development and improved survival of intestinal smooth muscle cells, which suggested that macropore connectivity can be optimized to enhance angiogenesis and improve cell viability. Compression molding combined with particulate leaching has been used to create multilayered scaffolds with differential porosities and pore sizes [120]. These structural features were found to influence the behavior and interaction (bridging versus penetration) of different cell types found within the small intestine (epithelial and smooth muscle cells). The impact of the spatial geometry and mechanics of the microenvironment are likely to have an important role in the physiological functionality of tissue engineered constructs. DiMarco and colleagues used a combination of experimental and finite element analysis to investigate critical variables that control intestinal organoid contraction [121]. Adjustment of ambient oxygen concentration, tailoring the density of the collagen type I matrix, addition of R-spondin1, and culture geometry were found to influence contractile behavior of the organoids. Contractile

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behavior occurred only within a narrow range of collagen densities, which the investigators suggested acted as a permissive switch to enable contraction. The inclusion of biomimetic topographical features might provide further control of cell behavior. To replicate the irregular multiscale features of native intestine, Koppes and colleagues created replicas of decellularized porcine small intestine by chemically depositing Parylene C to create molds for polydimethylsiloxane (PDMS) substrates [122]. The PDMS substrates exhibited multiscale folds, crypt and villus structures, and submicron features of the basement membrane. Finer control over the impact of microenvironmental cues is likely to improve scaffold performance, which affects tissue formation, development, differentiation, and functional behavior. Culture of primary intestinal organoids in recombinantly engineered extracellular matrix allowed improved physical manipulation of the scaffolds biomechanical cues while retaining the efficiency of organoid formation to match that obtained with natural collagen I matrices [123]. This provided a means to optimize the properties of the scaffolds to achieve improved matrix performance and identification of microenvironmental cues crucial for bioengineering tissue constructs. Porous protein scaffold systems composed of silk fibroin have also been used to replicate the tissue architecture and microenvironments of the intestine [124]. This type of insight into how properties of the scaffold materials influence the functional behavior of cells is crucial if physiological features such as the stem cell niche, cryptevillus axis, and peristalsis are to be achieved in the tissue engineered construct. Combined with the revolution in additive manufacturing, an icreased understanding of the biochemical and mechanical cues that control tissue regeneration will lead to a step change in state-of-the-art technology available to create scaffolds tailored to replicate the native scaffold of tissues in terms of topographical, mechanical, and biochemical features. Lee and colleagues fabricated scaffolds with a high surface areaeto-volume ratio using 3D printing technology [125]. The growth of smooth muscle cells in vitro was found to be influenced by the geometry of the scaffold. Scaffolds with small villi (0.5 mm) had increased cell density compared with scaffolds containing large villi (1 mm) after 14 days of culture. A 3D printable Matrigeleagarose system was described for the printing of intestinal epithelial cells [126]. The agarose component provided mechanical stability for the 3D printed structure and the Matrigel provides essential protein components for cell growth and spreading. The development of bioinks with improved mechanical properties and biocompatibility will enable additive manufacturing such as 3D printing to make tremendous opportunities for regenerative medicine [127]. Instilling peristaltic activity to the tissue engineered intestine to establish gut motility will require correctly oriented smooth muscle cell regeneration and reinnervation. Advances have been made in technologies to achieve peristalsis through the combination of autologous smooth muscle cells and biomaterials to produce patch or tubular constructs [128]. Maintaining native smooth muscle organization appears to be critical to achieving functionally contracting smooth muscle [129]. Intact smooth muscle strips retained neural and glial markers and exhibited periodic contraction whereas smooth muscle cells cultured after enzymatic digestion did not. Innervation is not only an unmet need for bioengineering gastrointestinal tissue constructs. Enteric neuropathies such as achalasia, gastroparesis, intestinal pseudoobstruction, and chronic constipation are functional gastrointestinal disorders that result from primary and secondary forms of degenerative disease that affect the nerves and muscles in the gastrointestinal tract [130]. Mouse enteric neural crest cells transplanted into aganglionic gut spread along the endogenous myenteric plexus to form functionally integrated branching networks closely associated with endogenous neural glial networks, providing evidence for the use of enteric neural stem cell therapies [131]. The appendix, a vestigial organ, might provide a potential source of autologous neural stem cells for enteric neural cell therapy [132]. Geisbauer and colleagues investigated whether a mixture of enteric cells isolated from longitudinal and circular muscle of the gut could be used as a potential source of neural crest stem cells for cell therapy. When the isolated cells were mixed in collagen containing bFGF and injected into an aganglionic segment of jejunum, ganglion-like structures were generated with elongated synapses [133]. Innervation of tissue engineered constructs is fundamental to achieving or restoring gastrointestinal transit. Colon smooth muscle cells cultured in composite chitosan scaffolds were innervated by differentiated functional neurons derived from cocultured neural progenitor cells [134]. Likewise, human smooth muscle cells and neural progenitor cells were engineered into innervated sheets of smooth muscle and wrapped around tubular chitosan scaffolds [135]. After subcutaneous implantation, the construct became vascularized and the lumenal patency was maintained. In addition to regulating peristalsis, the enteric nervous system in the intestine controls villi activity and the modulation of secretions from gut epithelial cells. Gut endocrine cells has an important role in regulating gastrointestinal activity by releasing serotonin, secretin, cholecystokinin, gastrin, and enteroglucagon and will be an essential component of the tissue engineered intestine. Nakase and colleagues investigated the regeneration of endocrine cells and the nerve system in a canine patch model of tissue engineered small intestine using a collagen sponge scaffold loaded with autologous gastric smooth muscle cells [136]. At 24 weeks after implantation of the scaffolds into the

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middle of an isolated ileal loop, the location and number of endocrine cells staining positive for chromogranin A were almost identical to those of native mucosa. Nerve fibers were present in the regenerated smooth muscle layer and villi, but the myenteric plexus of Auerbach and the submucosal plexus of Meissner were not visible. The density of smooth muscle cells implanted into the scaffolds did not affect the thickness of the regenerated smooth muscle layer, which remained approximately half that of the native smooth muscle layer, indicating that other cues will be necessary to increase its thickness. The investigators suggested that the thickness of the muscle layer might be limited by the blood supply available to the regenerating tissue, which might be increased by the delivery of angiogenic factors from the scaffold. Grikscheit and colleagues also reported that ganglion cells were distributed in the locality of the Auerbach and Meissner’s plexuses in tissue engineered small intestine [90]. Regeneration of the small intestine remains at an early stage of clinical development and has yet to provide a clear demonstration of improvement in nutrient absorption that will be valuable in a clinical therapeutic setting in humans. Although no models have unequivocally demonstrated functional neointestine with peristaltic activity, they indicate that it is feasible to engineer tubular segmental replacement of small bowel that incorporates innervated smooth muscle layers. Based on these findings, it may be possible to achieve incremental steps toward tissue engineering the intestine. For example, combining existing and established surgical procedures with the principles of tissue engineering and regenerative medicine may improve existing clinical outcome measures. An example of this type of approach was demonstrated by Nakao and colleagues, who showed that the longitudinal staples used during Bianchi’s procedure could be replaced with SIS graft [137]. Refinement of other existing techniques could yield further advances toward viable options for tissue engineered intestinal constructs.

COLON The colon is an important organ for water and sodium resorption and a storage pouch for waste products. Patients who undergo total colectomy are at risk for significant morbidities [138]. The surgical creation of an ileal pouch to create a reservoir provides only a limited solution and patients may still experience inflammation of the pouch (pouchitis), malabsorption, diarrhea, cramping abdominal pain, and fever [139]. Tissue engineering approaches similar to those used for the small intestine have been applied to the colon [140]. Consequently, many of the same challenges exist. Tissue engineered colon was achieved by seeding organoid units harvested from the sigmoid colon of neonatal Lewis rats, adult rats, and tissue-engineered colon itself onto a polymer scaffold that was implanted into the omentum of syngeneic adult Lewis rats. Tissue-engineered colon was generated from each of the donor tissue sources and the resulting architecture of the neocolon was similar to that of native tissue. When anastomosed to the native bowel, there was gross evidence of fluid absorption by the tissue engineered colon. The choice of scaffold material used for colon tissue engineering will have an impact on the viability of the neocolon. Two of the main biological scaffold materials explored in gastrointestinal tissue engineering are SIS and chitosan. Relatively few studies have made direct comparisons of these materials in terms of their biocompatibility. Denost and colleagues looked at the in vitro and in vivo properties of two bioscaffolds composed of these materials [141]. No substantial difference was observed in vitro in terms of cell attachment and proliferation, but the chitosan hydrogel facilitated improved healing of a preclinical in vivo model of colonic wall defect, including regeneration of the smooth muscle layer. A significant drawback reported with biological scaffolds such as chitosan is their weak mechanical properties. To enhance mechanical strength, Zakhem and Bitar reported using chitosan fibers circumferentially aligned around tubular chitosan scaffolds [142]. Tensile strength and strain, burst pressure, and Young’s modulus were all increased in scaffolds that contained the fibers.

ANAL CANAL Controlled storage and timely disposal of feces relies largely on the appropriate function of sphincter muscles that constrict the anal canal and maintain fecal continence. Fecal incontinence is a common disease, particularly in aging societies in which it has a huge impact on quality of life and incurs colossal health costs. Conservative estimates indicate that approximately 2% of community-dwelling adults experience regular fecal incontinence [143]. This figure increases to 50% in the institutionalized and geriatric population [144]. Conservative treatments for fecal

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incontinence are ineffective in patients with more than mild symptoms and surgical interventions produce poor long-term benefit, with frequent complications [145]. Despite holding considerable promise, cell therapy for incontinence affecting the alimentary tract remains relatively unexplored in humans [146,147]. Although the technical feasibility of injecting autologous myoblasts for treating fecal incontinence in humans has been demonstrated, these studies have been unable to demonstrate integration of cells into the damaged sphincter or a direct improvement in the functional integrity of the sphincter muscle. A fibrin-based bioengineered in vitro model of the internal anal sphincter (IAS) was described that demonstrated physiological functionality. This type of approach is likely to offer value in studying complex physiological mechanisms underlying sphincter malfunction [148,149]. The bioengineered sphincter was surgically implanted into the subcutaneous tissues of syngeneic mice and responded to the local delivery of bFGF, resulting in improved muscle viability, vascularization, and survival of the graft [150]. This field of research has continued to advance; the first study demonstrated the feasibility of transplanting bioengineered intrinsically innervated human IAS into mice. Isolated human IAS smooth muscle cells were cocultured with immortomouse fetal enteric neurons. The construct implanted into RAG1/ mice became neovascularized and the physiological function of the myogenic and neuronal components was retained. The IAS construct exhibited characteristics of IAS physiology [151]. The feasibility of implanting a bioengineered sphincter construct consisting of human IAS smooth muscle cells was explored in the perianal region of athymic rats [152]. The constructs were well-tolerated and the recipients were able to produce stool normally. For this study, vascularization was increased by delivering platelet-derived growth factor. In addition to the anal sphincter, proof-of-concept studies have demonstrated that it is feasible to bioengineer autologous bioengineered innervated pylorus constructs that consist of circumferentially aligned smooth muscle cells that exhibit tonic contractile phenotype and basal tone [153]. However, the feasibility of scaling-up this type of approach from a rodent model to humans remains uncertain. Although it is technically possible to bioengineer rings of muscle in vitro on a scale comparable to that of human sphincter muscle [154], innervation, vascularization, and cell viability in larger constructs have yet to be tested in larger-sized preclinical models. Regenerative medicine may also offer solutions to conditions in which existing medical and surgical procedures have failed. A condition in which this affects the alimentary tract is perianal fistulas that result from a connection between the anal canal and the perianal skin surface, creating an abnormal passageway for the discharge of pus, blood, and in some cases feces, resulting in significant morbidity. The goals of fistula treatment are eradication of perineal sepsis and fistula closure while posing a minimal risk for causing sphincter muscle damage. A difficulty in treating perianal fistulas is avoiding abscess formation caused by healing of the skin before closure of the tract. To address this, collagen anal fistula plugs have been devised to treat fistulas. Although early studies reported good healing rates with little or no risk to continence, long-term follow-up has revealed variable and disappointing success rates (24e78%) [155]. Reports of the plugs failing owing to dislodgment from the tracts indicate that this approach may not provide an ideal scaffold material to promote guided tissue regeneration and closure of the tract [155]. A possible solution to this problem is the use of scaffold materials that provide both optimal conditions for rapid cell infiltration when implanted into tissue cavities and mechanical strength to maintain an open scaffold structure [156].

IN VITRO MODELS Improved understanding of gastrointestinal developmental biology opens up new opportunities for creating 3D tissue constructs that can be used to modeling disease, understand embryonic development, and provide construct sources for therapeutic applications [157]. Principles of regenerative medicine are increasingly being used to fabricate biomimetic models of the gastrointestinal tract. Challenges that exist with developing in vitro tissue models of gastrointestinal tissue include mimicking the 3D microenvironment, interactions among different cell types, and the microbiome. New technologies are being applied to address these, including using microfluidics to create channels lined by living cells in microengineered biomimetic systems that might offer new opportunities to replace conventional animal models in preclinical toxicology testing. This approach has been applied to a variety of organs including the intestine to provide organs-on-chips that exhibit physiological properties including peristalsis-like movement [158]. Dynamic culture in a defined perfusion bioreactor has also been reported to result in tissue models that are physiologically closer to native small intestine [159]. Microfluidic cell culture devices have been designed that contain villi- and crypt-like structures that resulted in epithelial cells tightly connecting to each other and displaying absorption and paracellular transport function [160]. The incorporation of physiological parameters also

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appears to be important for establishing 3D in vitro tissue models of small intestine. Bioreactors used to culture decellularized segments of porcine jejunum have been used to coculture human Caco-2 cells with human microvascular endothelial cells. Compared with routine static Caco-2 assays, culture under dynamic conditions resulted in cell morphology that more closely resembled normal primary enterocytes [161]. Recapturing essential features of the cellular microenvironment is essential if in vitro tissue engineered models are to be used for functional studies such as cell growth, differentiation, absorption, or hostemicrobial interactions. The inclusion of native features such as accurately sized intestinal villi has been shown to facilitate cell differentiation along the villous axis [162]. Methods used to realize such structures include 3D natural and synthetic hydrogels created using a combination of laser ablation and sacrificial molding to achieve microscale structures that mimic the density and size of human intestinal villi [163]. The microbiome of the gastrointestinal tract is increasingly being recognized as a critical component to maintaining physiological homeostasis. Biomimetic in vitro intestinal models for investigating the adhesion and invasion profile of commensal and pathogenic organisms therefore have significant value in understanding microbe-induced intestinal disorders. To explore this interaction, synthetic 3D tissue scaffolds that support coculture of epithelial cell types have been used to provide microbial niches along the cryptevillus axis for modeling the interaction of a variety of commensal and pathogenic organisms [164].

CONCLUSION The alimentary tract is a complex organ that is essential for maintaining physiological homeostasis. Tissue engineering and regenerative medicine for hollow visceral organs have been proposed as an approach for replacing damaged or diseased tissue, as demonstrated in humans with bladder [165] and airway tissue [15]. Although these “first-in-human” studies have rightly attracted much attention, there is a long way to go until a similar approach becomes routine in the relatively complex structures of the alimentary tract. The past few decades have delivered a series of important studies that have used the innate ability of the alimentary tract to regenerate. Further studies are needed to demonstrate whether these approaches are transferable and of clinical value to humans. Fundamental challenges such as scalability will need to be addressed to enable the results obtained in small-animal models to progress into preclinical models applicable to humans. This will require refinement of the scaffolds used and the ability to seed limited quantities of cells available in an efficient manner onto the scaffolds. These are challenges that can be overcome and will allow regenerative medicine to progress in the alimentary tract in humans.

References [1] Sonnenburg JL, Ba¨ckhed F. Diet-microbiota interactions as moderators of human metabolism. Nature July 7, 2016;535(7610):56e64. [2] Fascetti-Leon F, Malerba A, Boldrin L, Leone E, Betalli P, Pasut A, et al. Murine muscle precursor cells survived and integrated in a cryoinjured gastroesophageal junction. J Surg Res December 2007;143(2):253e9. [3] Fukushima M, Kako N, Chiba K, Kawaguchi T, Kimura Y, Sato M, et al. Seven-year follow-up study after the replacement of the esophagus with an artificial esophagus in the dog. Surgery January 1983;93(1 Pt 1):70e7. [4] Jurkiewicz MJ. Reconstructive surgery of the cervical esophagus. J Thorac Cardiovasc Surg November 1984;88(5 Pt 2):893e7. [5] Harii K, Ebihara S, Ono I, Saito H, Terui S, Takato T. Pharyngoesophageal reconstruction using a fabricated forearm free flap. Plast Reconstr Surg April 1985;75(4):463e76. [6] Kakegawa T, Machi J, Yamana H, Fujita H, Tai Y. A new technique for esophageal reconstruction by combined skin and muscle flaps after failure in primary colonic interposition. Surg Gynecol Obstet June 1987;164(6):576e8. [7] Ohki T, Yamato M, Murakami D, Takagi R, Yang J, Namiki H, et al. Treatment of oesophageal ulcerations using endoscopic transplantation of tissue-engineered autologous oral mucosal epithelial cell sheets in a canine model. Gut December 2006;55(12):1704e10. [8] Shinhar D, Finaly R, Niska A, Mares AJ. The use of collagen-coated vicryl mesh for reconstruction of the canine cervical esophagus. Pediatr Surg Int March 1998;13(2e3):84e7. [9] Yamamoto Y, Nakamura T, Shimizu Y, Takimoto Y, Matsumoto K, Kiyotani T, et al. Experimental replacement of the thoracic esophagus with a bioabsorbable collagen sponge scaffold supported by a silicone stent in dogs. ASAIO J August 1999;45(4):311e6. [10] Badylak S, Meurling S, Chen M, Spievack A, Simmons-Byrd A. Resorbable bioscaffold for esophageal repair in a dog model. J Pediatr Surg July 2000;35(7):1097e103. [11] Badylak SF, Vorp DA, Spievack AR, Simmons-Byrd A, Hanke J, Freytes DO, et al. Esophageal reconstruction with ECM and muscle tissue in a dog model. J Surg Res September 2005;128(1):87e97. [12] Lynen Jansen P, Klinge U, Anurov M, Titkova S, Mertens PR, Jansen M. Surgical mesh as a scaffold for tissue regeneration in the esophagus. Eur Surg Res April 2004;36(2):104e11. [13] Ozeki M, Narita Y, Kagami H, Ohmiya N, Itoh A, Hirooka Y, et al. Evaluation of decellularized esophagus as a scaffold for cultured esophageal epithelial cells. J Biomed Mater Res December 15, 2006;79(4):771e8.

1144

64. ALIMENTARY TRACT

[14] Marzaro M, Vigolo S, Oselladore B, Conconi MT, Ribatti D, Giuliani S, et al. In vitro and in vivo proposal of an artificial esophagus. J Biomed Mater Res June 15, 2006;77(4):795e801. [15] Macchiarini P, Jungebluth P, Go T, Asnaghi MA, Rees LE, Cogan TA, et al. Clinical transplantation of a tissue-engineered airway. Lancet December 13, 2008;372(9655):2023e30. [16] Matsumoto T, Holmes RH, Burdick CO, Heisterkamp CA, O’Connell TJ. The fate of the inverted segment of small bowel used for the replacement of major veins. Surgery September 1966;60(3):739e43. [17] Badylak SF, Lantz GC, Coffey A, Geddes LA. Small intestinal submucosa as a large diameter vascular graft in the dog. J Surg Res July 1989; 47(1):74e80. [18] Kropp BP, Badylak S, Thor KB. Regenerative bladder augmentation: a review of the initial preclinical studies with porcine small intestinal submucosa. Adv Exp Med Biol 1995;385:229e35. [19] Dalla Vecchia L, Engum S, Kogon B, Jensen E, Davis M, Grosfeld J. Evaluation of small intestine submucosa and acellular dermis as diaphragmatic prostheses. J Pediatr Surg January 1999;34(1):167e71. [20] De Ugarte DA, Puapong D, Roostaeian J, Gillis N, Fonkalsrud EW, Atkinson JB, et al. Surgisis patch tracheoplasty in a rodent model for tracheal stenosis. J Surg Res June 1, 2003;112(1):65e9. [21] Puccio F, Solazzo M, Marciano P. Comparison of three different mesh materials in tension-free inguinal hernia repair: prolene versus Vypro versus surgisis. Int Surg August 2005;90(3 Suppl.):S21e3. [22] Spiegel JH, Kessler JL. Tympanic membrane perforation repair with acellular porcine submucosa. Otol Neurotol July 2005;26(4):563e6. [23] Grethel EJ, Cortes RA, Wagner AJ, Clifton MS, Lee H, Farmer DL, et al. Prosthetic patches for congenital diaphragmatic hernia repair: surgisis vs Gore-Tex. J Pediatr Surg January 2006;41(1):29e33. discussion 29. [24] Smith MD, Campbell RM. Use of a biodegradable patch for reconstruction of large thoracic cage defects in growing children. J Pediatr Surg January 2006;41(1):46e9. discussion 46. [25] Hodde JP, Badylak SF, Brightman AO, Voytik-Harbin SL. Glycosaminoglycan content of small intestinal submucosa: a bioscaffold for tissue replacement. Tissue Eng 1996;2(3):209e17. [26] Voytik-Harbin SL, Brightman AO, Kraine MR, Waisner B, Badylak SF. Identification of extractable growth factors from small intestinal submucosa. J Cell Biochem December 15, 1997;67(4):478e91. [27] Lopes MF, Cabrita A, Ilharco J, Pessa P, Patrı´cio J. Grafts of porcine intestinal submucosa for repair of cervical and abdominal esophageal defects in the rat. J Invest Surg April 2006;19(2):105e11. [28] Doede T, Bondartschuk M, Joerck C, Schulze E, Goernig M. Unsuccessful alloplastic esophageal replacement with porcine small intestinal submucosa. Artif Organs April 2009;33(4):328e33. [29] Fan M-R, Gong M, Da L-C, Bai L, Li X-Q, Chen K-F, et al. Tissue engineered esophagus scaffold constructed with porcine small intestinal submucosa and synthetic polymers. Biomed Mater February 2014;9(1):015012. [30] Master VA, Wei G, Liu W, Baskin LS. Urothlelium facilitates the recruitment and trans-differentiation of fibroblasts into smooth muscle in acellular matrix. J Urol October 2003;170(4 Pt 2):1628e32. [31] Rheinwald JG, Green H. Serial cultivation of strains of human epidermal keratinocytes: the formation of keratinizing colonies from single cells. Cell November 1975;6(3):331e43. [32] Tan B, Wei R-Q, Tan M-Y, Luo J-C, Deng L, Chen X-H, et al. Tissue engineered esophagus by mesenchymal stem cell seeding for esophageal repair in a canine model. J Surg Res June 1, 2013;182(1):40e8. [33] Nakase Y, Nakamura T, Kin S, Nakashima S, Yoshikawa T, Kuriu Y, et al. Intrathoracic esophageal replacement by in situ tissue-engineered esophagus. J Thorac Cardiovasc Surg October 2008;136(4):850e9. [34] Yoo JJ, Meng J, Oberpenning F, Atala A. Bladder augmentation using allogenic bladder submucosa seeded with cells. Urology February 1998;51(2):221e5. [35] Urita Y, Komuro H, Chen G, Shinya M, Kaneko S, Kaneko M, et al. Regeneration of the esophagus using gastric acellular matrix: an experimental study in a rat model. Pediatr Surg Int January 2007;23(1):21e6. [36] Lun S, Irvine SM, Johnson KD, Fisher NJ, Floden EW, Negron L, et al. A functional extracellular matrix biomaterial derived from ovine forestomach. Biomaterials June 2010;31(16):4517e29. [37] Franck D, Chung YG, Coburn J, Kaplan DL, Estrada CR, Mauney JR. In vitro evaluation of bi-layer silk fibroin scaffolds for gastrointestinal tissue engineering. J Tissue Eng November 5, 2014;5. 2041731414556849. [38] Kofler K, Ainoedhofer H, Ho¨llwarth ME, Saxena AK. Fluorescence-activated cell sorting of PCK-26 antigen-positive cells enables selection of ovine esophageal epithelial cells with improved viability on scaffolds for esophagus tissue engineering. Pediatr Surg Int January 2010;26(1): 97e104. [39] Tan B, Wang M, Chen X, Hou J, Chen X, Wang Y, et al. Tissue engineered esophagus by copperesmall intestinal submucosa graft for esophageal repair in a canine model. Sci China Life Sci February 2014;57(2):248e55. [40] Agrawal V, Brown BN, Beattie AJ, Gilbert TW, Badylak SF. Evidence of innervation following extracellular matrix scaffold-mediated remodelling of muscular tissues. J Tissue Eng Regen Med December 2009;3(8):590e600. [41] Saxena AK, Kofler K, Aino¨dhofer H, Ho¨llwarth ME. Esophagus tissue engineering: hybrid approach with esophageal epithelium and unidirectional smooth muscle tissue component generation in vitro. J Gastrointest Surg June 2009;13(6):1037e43. [42] Hori Y, Nakamura T, Matsumoto K, Kurokawa Y, Satomi S, Shimizu Y. Experimental study on in situ tissue engineering of the stomach by an acellular collagen sponge scaffold graft. ASAIO J June 2001;47(3):206e10. [43] Hori Y, Nakamura T, Kimura D, Kaino K, Kurokawa Y, Satomi S, et al. Functional analysis of the tissue-engineered stomach wall. Artif Organs October 2002;26(10):868e72. [44] Araki M, Tao H, Sato T, Nakajima N, Hyon S-H, Nagayasu T, et al. Development of a new tissue-engineered sheet for reconstruction of the stomach. Artif Organs October 16, 2009;33(10):818e26. [45] Grikscheit T, Srinivasan A, Vacanti JP. Tissue-engineered stomach: a preliminary report of a versatile in vivo model with therapeutic potential. J Pediatr Surg September 2003;38(9):1305e9. [46] Sala FG, Kunisaki SM, Ochoa ER, Vacanti J, Grikscheit TC. Tissue-engineered small intestine and stomach form from autologous tissue in a preclinical large animal model. J Surg Res October 2009;156(2):205e12.

REFERENCES

1145

[47] McCracken KW, Cata´ EM, Crawford CM, Sinagoga KL, Schumacher M, Rockich BE, et al. Modelling human development and disease in pluripotent stem-cell-derived gastric organoids. Nature December 18, 2014;516(7531):400e4. [48] Noguchi TK, Ninomiya N, Sekine M, Komazaki S, Wang P-C, Asashima M, et al. Generation of stomach tissue from mouse embryonic stem cells. Nat Cell Biol August 2015;17(8):984e93. [49] Nakatsu H, Ueno T, Oga A, Nakao M, Nishimura T, Kobayashi S, et al. Influence of mesenchymal stem cells on stomach tissue engineering using small intestinal submucosa. J Tissue Eng Regen Med March 2015;9(3):296e304. [50] Booth C, Potten CS. Gut instincts: thoughts on intestinal epithelial stem cells. J Clin Invest June 2000;105(11):1493e9. [51] Kayahara T, Sawada M, Takaishi S, Fukui H, Seno H, Fukuzawa H, et al. Candidate markers for stem and early progenitor cells, Musashi-1 and Hes1, are expressed in crypt base columnar cells of mouse small intestine. FEBS Lett January 30, 2003;535(1e3):131e5. [52] Potten CS, Booth C, Tudor GL, Booth D, Brady G, Hurley P, et al. Identification of a putative intestinal stem cell and early lineage marker; musashi-1. Differentiation January 2003;71(1):28e41. [53] Gracz AD, Ramalingam S, Magness ST. Sox9 expression marks a subset of CD24-expressing small intestine epithelial stem cells that form organoids in vitro. Am J Physiol Gastrointest Liver Physiol May 2010;298(5):G590e600. [54] Bianchi A. Experience with longitudinal intestinal lengthening and tailoring. Eur J Pediatr Surg August 1999;9(4):256e9. [55] Thompson JS. Surgical approach to the short-bowel syndrome: procedures to slow intestinal transit. Eur J Pediatr Surg August 1999;9(4): 263e6. [56] Weber TR. Isoperistaltic bowel lengthening for short bowel syndrome in children. Am J Surg December 1999;178(6):600e4. [57] Javid PJ, Kim HB, Duggan CP, Jaksic T. Serial transverse enteroplasty is associated with successful short-term outcomes in infants with short bowel syndrome. J Pediatr Surg June 2005;40(6):1019e23. discussion 1023. [58] Sudan D. The current state of intestine transplantation: indications, techniques, outcomes and challenges. Am J Transplant September 2014; 14(9):1976e84. [59] Miyasaka EA, Okawada M, Utter B, Mustafa-Maria H, Luntz J, Brei D, et al. Application of distractive forces to the small intestine: defining safe limits. J Surg Res October 2010;163(2):169e75. [60] Koga H, Sun X, Yang H, Nose K, Somara S, Bitar KN, et al. Distraction-induced intestinal enterogenesis: preservation of intestinal function and lengthening after reimplantation into normal jejunum. Ann Surg February 1, 2012;255(2):302e10. [61] Fisher JG, Sparks EA, Khan FA, Dionigi B, Wu H, Brazzo J, et al. Extraluminal distraction enterogenesis using shape-memory polymer. J Pediatr Surg June 2015;50(6):938e42. [62] Sullins VF, Wagner JP, Suwarnasarn AT, Lee SL, Wu BM, Dunn JCY. A novel biodegradable device for intestinal lengthening. J Pediatr Surg January 2014;49(1):109e13. discussion 113. [63] Demehri FR, Wong PM, Freeman JJ, Fukatsu Y, Teitelbaum DH. A novel double-balloon catheter device for fully endoluminal intestinal lengthening. Pediatr Surg Int December 2014;30(12):1223e9. [64] Okawada M, Maria HM, Teitelbaum DH. Distraction induced enterogenesis: a unique mouse model using polyethylene glycol. J Surg Res September 2011;170(1):41e7. [65] Sueyoshi R, Ralls MW, Teitelbaum DH. Glucagon-like peptide 2 increases efficacy of distraction enterogenesis. J Surg Res September 2013; 184(1):365e73. [66] Rouch JD, Scott A, Jabaji ZB, Chiang E, Wu BM, Lee SL, et al. Basic fibroblast growth factor eluting microspheres enhance distraction enterogenesis. J Pediatr Surg June 2016;51(6):960e5. [67] Liu Y, Nelson T, Cromeens B, Rager T, Lannutti J, Johnson J, et al. HB-EGF embedded in PGA/PLLA scaffolds via subcritical CO2 augments the production of tissue engineered intestine. Biomaterials October 2016;103:150e9. [68] Kobold EE, Thal AP. A simple method for the management of experimental wounds of the duodenum. Surg Gynecol Obstet March 1963;116: 340e4. [69] Binnington HB, Siegel BA, Kissane JM, Ternberg JL. A technique to increase jejunal mucosa surface area. J Pediatr Surg October 1973;8(5): 765e9. [70] Watson LC, Friedman HI, Griffin DG, Norton LW, Mellick PW. Small bowel neomucosa. J Surg Res March 1980;28(3):280e91. [71] Jwo S-C, Chiu J-H, Ng K-K, Chen H-Y. Intestinal regeneration by a novel surgical procedure. Br J Surg May 2008;95(5):657e63. [72] Chen MK, Badylak SF. Small bowel tissue engineering using small intestinal submucosa as a scaffold. J Surg Res August 2001;99(2):352e8. [73] Pahari MP, Raman A, Bloomenthal A, Costa MA, Bradley SP, Banner B, et al. A novel approach for intestinal elongation using acellular dermal matrix: an experimental study in rats. Transplant Proc August 2006;38(6):1849e50. [74] Pahari MP, Brown ML, Elias G, Nseir H, Banner B, Rastellini C, et al. Development of a bioartificial new intestinal segment using an acellular matrix scaffold. Gut June 2007;56(6):885e6. [75] Cicalese L, Corsello T, Stevenson HL, Damiano G, Tuveri M, Zorzi D, et al. Evidence of absorptive function in vivo in a neo-formed bioartificial intestinal segment using a rodent model. J Gastrointest Surg January 2016;20(1):34e42. discussion 42. [76] Chouillard E, Chahine E, Allaire E, Filaire-Legendre A, Van Nhieu JT, Martinod E. Small bowel in vivo bioengineering using an aortic matrix in a porcine model. Surg Endosc February 22, 2016;30(11):4742e9. [77] Hori Y, Nakamura T, Matsumoto K, Kurokawa Y, Satomi S, Shimizu Y. Tissue engineering of the small intestine by acellular collagen sponge scaffold grafting. Int J Artif Organs January 2001;24(1):50e4. [78] Hori Y, Nakamura T, Kimura D, Kaino K, Kurokawa Y, Satomi S, et al. Experimental study on tissue engineering of the small intestine by mesenchymal stem cell seeding. J Surg Res February 2002;102(2):156e60. [79] Wang ZQ, Watanabe Y, Noda T, Yoshida A, Oyama T, Toki A. Morphologic evaluation of regenerated small bowel by small intestinal submucosa. J Pediatr Surg December 2005;40(12):1898e902. [80] Lee M, Chang PCY, Dunn JCY. Evaluation of small intestinal submucosa as scaffolds for intestinal tissue engineering. J Surg Res June 15, 2008;147(2):168e71. [81] Vacanti JP, Morse MA, Saltzman WM, Domb AJ, Perez-Atayde A, Langer R. Selective cell transplantation using bioabsorbable artificial polymers as matrices. J Pediatr Surg January 1988;23(1 Pt 2):3e9. [82] Tait IS, Flint N, Campbell FC, Evans GS. Generation of neomucosa in vivo by transplantation of dissociated rat postnatal small intestinal epithelium. Differentiation April 1994;56(1e2):91e100.

1146

64. ALIMENTARY TRACT

[83] Slorach EM, Campbell FC, Dorin JR. A mouse model of intestinal stem cell function and regeneration. J Cell Sci September 1999;112 Pt 18: 3029e38. [84] Choi RS, Vacanti JP. Preliminary studies of tissue-engineered intestine using isolated epithelial organoid units on tubular synthetic biodegradable scaffolds. Transplant Proc March 1997;29(1e2):848e51. [85] Choi RS, Riegler M, Pothoulakis C, Kim BS, Mooney D, Vacanti M, et al. Studies of brush border enzymes, basement membrane components, and electrophysiology of tissue-engineered neointestine. J Pediatr Surg July 1998;33(7):991e6. discussion 996. [86] Kaihara S, Kim SS, Benvenuto M, Choi R, Kim BS, Mooney D, et al. Successful anastomosis between tissue-engineered intestine and native small bowel. Transplantation January 27, 1999;67(2):241e5. [87] Tavakkolizadeh A, Berger UV, Stephen AE, Kim BS, Mooney D, Hediger MA, et al. Tissue-engineered neomucosa: morphology, enterocyte dynamics, and SGLT1 expression topography. Transplantation January 27, 2003;75(2):181e5. [88] Perez A, Grikscheit TC, Blumberg RS, Ashley SW, Vacanti JP, Whang EE. Tissue-engineered small intestine: ontogeny of the immune system. Transplantation September 15, 2002;74(5):619e23. [89] Kim SS, Kaihara S, Benvenuto MS, Choi RS, Kim BS, Mooney DJ, et al. Regenerative signals for intestinal epithelial organoid units transplanted on biodegradable polymer scaffolds for tissue engineering of small intestine. Transplantation January 27, 1999;67(2):227e33. [90] Grikscheit TC, Siddique A, Ochoa ER, Srinivasan A, Alsberg E, Hodin RA, et al. Tissue-engineered small intestine improves recovery after massive small bowel resection. Ann Surg November 2004;240(5):748e54. [91] Warner BW. Tissue engineered small intestine: a viable clinical option? Ann Surg November 2004;240(5):755e6. [92] Sala FG, Matthews JA, Speer AL, Torashima Y, Barthel ER, Grikscheit TC. A multicellular approach forms a significant amount of tissueengineered small intestine in the mouse. Tissue Eng July 2011;17(13e14):1841e50. [93] Fuller MK, Faulk DM, Sundaram N, Shroyer NF, Henning SJ, Helmrath MA. Intestinal crypts reproducibly expand in culture. J Surg Res November 1, 2012;178(1):48e54. [94] Rizvi AZ, Swain JR, Davies PS, Bailey AS, Decker AD, Willenbring H, et al. Bone marrow-derived cells fuse with normal and transformed intestinal stem cells. Proc Natl Acad Sci U S A April 18, 2006;103(16):6321e5. [95] Zhao Y, Glesne D, Huberman E. A human peripheral blood monocyte-derived subset acts as pluripotent stem cells. Proc Natl Acad Sci U S A March 4, 2003;100(5):2426e31. [96] McCracken KW, Howell JC, Wells JM, Spence JR. Generating human intestinal tissue from pluripotent stem cells in vitro. Nat Protoc December 2011;6(12):1920e8. [97] Cromeens BP, Liu Y, Stathopoulos J, Wang Y, Johnson J, Besner GE. Production of tissue-engineered intestine from expanded enteroids. J Surg Res July 2016;204(1):164e75. [98] Liu Y, Rager T, Johnson J, Enmark J, Besner GE. Enriched intestinal stem cell seeding improves the architecture of tissue-engineered intestine. Tissue Eng August 2015;21(8):816e24. [99] Torashima Y, Levin DE, Barthel ER, Speer AL, Sala FG, Hou X, et al. Fgf10 overexpression enhances the formation of tissue-engineered small intestine. J Tissue Eng Regen Med February 2016;10(2):132e9. [100] Levin DE, Barthel ER, Speer AL, Sala FG, Hou X, Torashima Y, et al. Human tissue-engineered small intestine forms from postnatal progenitor cells. J Pediatr Surg January 2013;48(1):129e37. [101] Patil PB, Chougule PB, Kumar VK, Almstro¨m S, Ba¨ckdahl H, Banerjee D, et al. Recellularization of acellular human small intestine using bone marrow stem cells. Stem Cells Transl Med April 2013;2(4):307e15. [102] Spurrier RG, Speer AL, Grant CN, Levin DE, Grikscheit TC. Vitrification preserves murine and human donor cells for generation of tissueengineered intestine. J Surg Res August 2014;190(2):399e406. [103] Ramsanahie A, Duxbury MS, Grikscheit TC, Perez A, Rhoads DB, Gardner-Thorpe J, et al. Effect of GLP-2 on mucosal morphology and SGLT1 expression in tissue-engineered neointestine. Am J Physiol Gastrointest Liver Physiol December 2003;285(6):G1345e52. [104] Yoshida A, Noda T, Tani M, Oyama T, Watanabe Y, Kiyomoto H, et al. The role of basic fibroblast growth factor to enhance fetal intestinal mucosal cell regeneration in vivo. Pediatr Surg Int August 2009;25(8):691e5. [105] Moosvi SR, Day RM. Bioactive glass modulation of intestinal epithelial cell restitution. Acta Biomater January 2009;5(1):76e83. [106] Lee M, Wu BM, Stelzner M, Reichardt HM, Dunn JCY. Intestinal smooth muscle cell maintenance by basic fibroblast growth factor. Tissue Eng August 2008;14(8):1395e402. [107] Gardner-Thorpe J, Grikscheit TC, Ito H, Perez A, Ashley SW, Vacanti JP, et al. Angiogenesis in tissue-engineered small intestine. Tissue Eng December 2003;9(6):1255e61. [108] Rocha FG, Sundback CA, Krebs NJ, Leach JK, Mooney DJ, Ashley SW, et al. The effect of sustained delivery of vascular endothelial growth factor on angiogenesis in tissue-engineered intestine. Biomaterials July 2008;29(19):2884e90. [109] Day RM, Boccaccini AR, Shurey S, Roether JA, Forbes A, Hench LL, et al. Assessment of polyglycolic acid mesh and bioactive glass for softtissue engineering scaffolds. Biomaterials December 2004;25(27):5857e66. [110] Duxbury MS, Grikscheit TC, Gardner-Thorpe J, Rocha FG, Ito H, Perez A, et al. Lymphangiogenesis in tissue-engineered small intestine. Transplantation April 27, 2004;77(8):1162e6. [111] Mertsching H, Schanz J, Steger V, Schandar M, Schenk M, Hansmann J, et al. Generation and transplantation of an autologous vascularized bioartificial human tissue. Transplantation July 27, 2009;88(2):203e10. [112] Kim SS, Penkala R, Abrahimi P. A perfusion bioreactor for intestinal tissue engineering. J Surg Res October 2007;142(2):327e31. [113] Gonzalez-Molina J, Riegler J, Southern P, Ortega D, Frangos CC, Angelopoulos Y, et al. Rapid magnetic cell delivery for large tubular bioengineered constructs. J R Soc Interface November 7, 2012;9(76):3008e16. [114] Kasyanov VA, Hodde J, Hiles MC, Eisenberg C, Eisenberg L, De Castro LEF, et al. Rapid biofabrication of tubular tissue constructs by centrifugal casting in a decellularized natural scaffold with laser-machined micropores. J Mater Sci Mater Med January 2009;20(1):329e37. [115] Boomer L, Liu Y, Mahler N, Johnson J, Zak K, Nelson T, et al. Scaffolding for challenging environments: materials selection for tissue engineered intestine. J Biomed Mater Res November 2014;102(11):3795e802. [116] Shabafrooz V, Mozafari M, Ko¨hler GA, Assefa S, Vashaee D, Tayebi L. The effect of hyaluronic acid on biofunctionality of gelatin-collagen intestine tissue engineering scaffolds. J Biomed Mater Res September 2014;102(9):3130e9.

REFERENCES

1147

[117] Kobayashi M, Lei NY, Wang Q, Wu BM, Dunn JCY. Orthogonally oriented scaffolds with aligned fibers for engineering intestinal smooth muscle. Biomaterials August 2015;61:75e84. [118] Wang L, Murthy SK, Fowle WH, Barabino GA, Carrier RL. Influence of micro-well biomimetic topography on intestinal epithelial Caco-2 cell phenotype. Biomaterials December 2009;30(36):6825e34. [119] Walthers CM, Nazemi AK, Patel SL, Wu BM, Dunn JCY. The effect of scaffold macroporosity on angiogenesis and cell survival in tissueengineered smooth muscle. Biomaterials June 2014;35(19):5129e37. [120] Knight T, Basu J, Rivera EA, Spencer T, Jain D, Payne R. Fabrication of a multi-layer three-dimensional scaffold with controlled porous micro-architecture for application in small intestine tissue engineering. Cell Adh Migr June 2013;7(3):267e74. [121] DiMarco RL, Su J, Yan KS, Dewi R, Kuo CJ, Heilshorn SC. Engineering of three-dimensional microenvironments to promote contractile behavior in primary intestinal organoids. Integr Biol (Camb) February 2014;6(2):127e42. [122] Koppes AN, Kamath M, Pfluger CA, Burkey DD, Dokmeci M, Wang L, et al. Complex, multi-scale small intestinal topography replicated in cellular growth substrates fabricated via chemical vapor deposition of Parylene C. Biofabrication August 22, 2016;8(3):035011. [123] DiMarco RL, Dewi RE, Bernal G, Kuo C, Heilshorn SC. Protein-engineered scaffolds for in vitro 3D culture of primary adult intestinal organoids. Biomater Sci October 15, 2015;3(10):1376e85. [124] Chen Y, Lin Y, Davis KM, Wang Q, Rnjak-Kovacina J, Li C, et al. Robust bioengineered 3D functional human intestinal epithelium. Sci Rep September 16, 2015;5:13708. [125] Lee M, Wu BM, Dunn JCY. Effect of scaffold architecture and pore size on smooth muscle cell growth. J Biomed Mater Res December 15, 2008;87(4):1010e6. [126] Fan R, Piou M, Darling E, Cormier D, Sun J, Wan J. Bio-printing cell-laden Matrigel-agarose constructs. J Biomater Appl September 16, 2016; 31(5):684e92. [127] Wengerter BC, Emre G, Park JY, Geibel J. Three-dimensional printing in the intestine. Clin Gastroenterol Hepatol August 2016;14(8):1081e5. [128] Basu J, Mihalko KL, Payne R, Rivera E, Knight T, Genheimer CW, et al. Regeneration of rodent small intestine tissue following implantation of scaffolds seeded with a novel source of smooth muscle cells. Regen Med November 2011;6(6):721e31. [129] Walthers CM, Lee M, Wu BM, Dunn JCY. Smooth muscle strips for intestinal tissue engineering. PLoS One December 8, 2014;9(12):e114850. [130] Knowles CH, Lindberg G, Panza E, De Giorgio R. New perspectives in the diagnosis and management of enteric neuropathies. Nat Rev Gastroenterol Hepatol April 2013;10(4):206e18. [131] Cooper JE, McCann CJ, Natarajan D, Choudhury S, Boesmans W, Delalande J-M, et al. In vivo transplantation of enteric neural crest cells into mouse gut; engraftment, functional integration and long-term safety. PLoS One January 29, 2016;11(1):e0147989. [132] Zakhem E, Rego SL, Raghavan S, Bitar KN. The appendix as a viable source of neural progenitor cells to functionally innervate bioengineered gastrointestinal smooth muscle tissues. Stem Cells Transl Med June 2015;4(6):548e54. [133] Geisbauer CL, Wu BM, Dunn JCY. Transplantation of enteric cells into the aganglionic rodent small intestines. J Surg Res July 2012;176(1): 20e8. [134] Zakhem E, Raghavan S, Bitar KN. Neo-innervation of a bioengineered intestinal smooth muscle construct around chitosan scaffold. Biomaterials February 2014;35(6):1882e9. [135] Zakhem E, Elbahrawy M, Orlando G, Bitar KN. Successful implantation of an engineered tubular neuromuscular tissue composed of human cells and chitosan scaffold. Surgery December 2015;158(6):1598e608. [136] Nakase Y, Nakamura T, Kin S, Nakashima S, Yoshikawa T, Kuriu Y, et al. Endocrine cell and nerve regeneration in autologous in situ tissueengineered small intestine. J Surg Res January 2007;137(1):61e8. [137] Nakao M, Ueno T, Oga A, Kuramitsu Y, Nakatsu H, Oka M. Proposal of intestinal tissue engineering combined with Bianchi’s procedure. J Pediatr Surg April 2015;50(4):573e80. [138] Papa MZ, Karni T, Koller M, Klein E, Scott D, Bersuk D, et al. Avoiding diarrhea after subtotal colectomy with primary anastomosis in the treatment of colon cancer. J Am Coll Surg March 1997;184(3):269e72. [139] Meagher AP, Farouk R, Dozois RR, Kelly KA, Pemberton JH. J ileal pouch-anal anastomosis for chronic ulcerative colitis: complications and long-term outcome in 1310 patients. Br J Surg June 1998;85(6):800e3. [140] Grikscheit TC, Ochoa ER, Ramsanahie A, Alsberg E, Mooney D, Whang EE, et al. Tissue-engineered large intestine resembles native colon with appropriate in vitro physiology and architecture. Ann Surg July 2003;238(1):35e41. [141] Denost Q, Adam J-P, Pontallier A, Montembault A, Bareille R, Siadous R, et al. Colorectal tissue engineering: a comparative study between porcine small intestinal submucosa (SIS) and chitosan hydrogel patches. Surgery December 2015;158(6):1714e23. [142] Zakhem E, Bitar KN. Development of chitosan scaffolds with enhanced mechanical properties for intestinal tissue engineering applications. J Funct Biomater October 13, 2015;6(4):999e1011. [143] Perry S, Shaw C, McGrother C, Matthews RJ, Assassa RP, Dallosso H, et al. Prevalence of faecal incontinence in adults aged 40 years or more living in the community. Gut April 2002;50(4):480e4. [144] Nelson RL. Epidemiology of fecal incontinence. Gastroenterology January 2004;126(1 Suppl. 1):S3e7. [145] Tan JJY, Chan M, Tjandra JJ. Evolving therapy for fecal incontinence. Dis Colon Rectum November 2007;50(11):1950e67. [146] Frudinger A, Ko¨lle D, Schwaiger W, Pfeifer J, Paede J, Halligan S. Muscle-derived cell injection to treat anal incontinence due to obstetric trauma: pilot study with 1 year follow-up. Gut January 2010;59(1):55e61. [147] Frudinger A, Pfeifer J, Paede J, Kolovetsiou-Kreiner V, Marksteiner R, Halligan S. Autologous skeletal-muscle-derived cell injection for anal incontinence due to obstetric trauma: a 5-year follow-up of an initial study of 10 patients. Colorectal Dis September 2015;17(9):794e801. [148] Hecker L, Baar K, Dennis RG, Bitar KN. Development of a three-dimensional physiological model of the internal anal sphincter bioengineered in vitro from isolated smooth muscle cells. Am J Physiol Gastrointest Liver Physiol August 2005;289(2):G188e96. [149] Somara S, Gilmont RR, Dennis RG, Bitar KN. Bioengineered internal anal sphincter derived from isolated human internal anal sphincter smooth muscle cells. Gastroenterology July 2009;137(1):53e61. [150] Hashish M, Raghavan S, Somara S, Gilmont RR, Miyasaka E, Bitar KN, et al. Surgical implantation of a bioengineered internal anal sphincter. J Pediatr Surg January 2010;45(1):52e8. [151] Raghavan S, Gilmont RR, Miyasaka EA, Somara S, Srinivasan S, Teitelbaum DH, et al. Successful implantation of bioengineered, intrinsically innervated, human internal anal sphincter. Gastroenterology July 2011;141(1):310e9.

1148

64. ALIMENTARY TRACT

[152] Raghavan S, Miyasaka EA, Gilmont RR, Somara S, Teitelbaum DH, Bitar KN. Perianal implantation of bioengineered human internal anal sphincter constructs intrinsically innervated with human neural progenitor cells. Surgery April 2014;155(4):668e74. [153] Rego SL, Zakhem E, Orlando G, Bitar KN. Bioengineered human pyloric sphincters using autologous smooth muscle and neural progenitor cells. Tissue Eng January 2016;22(1e2):151e60. [154] Parmar N, Day RM. Appropriately sized bioengineered human external anal sphincter constructs. Surgery January 2015;157(1):177e8. [155] Adamina M, Hoch JS, Burnstein MJ. To plug or not to plug: a cost-effectiveness analysis for complex anal fistula. Surgery January 2010; 147(1):72e8. [156] Blaker JJ, Pratten J, Ready D, Knowles JC, Forbes A, Day RM. Assessment of antimicrobial microspheres as a prospective novel treatment targeted towards the repair of perianal fistulae. Aliment Pharmacol Ther September 1, 2008;28(5):614e22. [157] Howell JC, Wells JM. Generating intestinal tissue from stem cells: potential for research and therapy. Regen Med November 2011;6(6): 743e55. [158] Huh D, Kim HJ, Fraser JP, Shea DE, Khan M, Bahinski A, et al. Microfabrication of human organs-on-chips. Nat Protoc November 2013;8(11): 2135e57. [159] Schweinlin M, Wilhelm S, Schwedhelm I, Hansmann J, Rietscher R, Jurowich C, et al. Development of an advanced primary human in vitro model of the small intestine. Tissue Eng September 2016;22(9):873e83. [160] Chi M, Yi B, Oh S, Park D-J, Sung JH, Park S. A microfluidic cell culture device (mFCCD) to culture epithelial cells with physiological and morphological properties that mimic those of the human intestine. Biomed Microdevices 2015;17(3):9966. [161] Pusch J, Votteler M, Go¨hler S, Engl J, Hampel M, Walles H, et al. The physiological performance of a three-dimensional model that mimics the microenvironment of the small intestine. Biomaterials October 2011;32(30):7469e78. [162] Costello CM, Hongpeng J, Shaffiey S, Yu J, Jain NK, Hackam D, et al. Synthetic small intestinal scaffolds for improved studies of intestinal differentiation. Biotechnol Bioeng June 2014;111(6):1222e32. [163] Sung JH, Yu J, Luo D, Shuler ML, March JC. Microscale 3-D hydrogel scaffold for biomimetic gastrointestinal (GI) tract model. Lab Chip February 7, 2011;11(3):389e92. [164] Costello CM, Sorna RM, Goh Y-L, Cengic I, Jain NK, March JC. 3-D intestinal scaffolds for evaluating the therapeutic potential of probiotics. Mol Pharm July 7, 2014;11(7):2030e9. [165] Atala A, Bauer SB, Soker S, Yoo JJ, Retik AB. Tissue-engineered autologous bladders for patients needing cystoplasty. Lancet April 15, 2006; 367(9518):1241e6.