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a McGowan Institute for Regenerative Medicine, Department of Surgery, University of Pittsburgh, Pittsburgh, Pennsylvania
b Heart, Lung, and Esophageal Institute, University of Pittsburgh Medical Center, Pittsburgh, Pennsylvania
c Veterans' Affairs Pittsburgh Health Care System, Pittsburgh, Pennsylvania
d Department of Pediatrics, National Jewish Medical Research Center, Denver, Colorado
Accepted for publication April 21, 2008.
* Address correspondence to Dr Badylak, McGowan Institute for Regenerative Medicine, University of Pittsburgh, 100 Technology Drive, Ste 200, Pittsburgh, PA 15219 (Email: badylaks{at}upmc.edu).
Presented at the Forty-fourth Annual Meeting of The Society of Thoracic Surgeons, Fort Lauderdale, FL, Jan 28–30, 2008.
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Abstract |
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Methods: A 1-cm wide x 2-cm-long defect was created in the ventral trachea of 15 dogs and repaired with one of three acellular biologic scaffolds: urinary bladder matrix (UBM), UBM crosslinked with carbodiimide (UBMC), and decellularized tracheal matrix (DTM). The grafts were evaluated periodically using bronchoscopy and by macroscopic and microscopic morphologic examination at either 2 months or 6 months.
Results: The UBM, UBMC, and DTM groups showed no evidence of stenosis or tracheomalacia. The UBM, UBMC, and DTM groups all showed deposition of organized collagenous tissue at the site of scaffold placement and an intact epithelial layer. Scattered areas of mucociliary differentiation were present at the edges of the graft site. There was no evidence cartilage observed within the remodeled tissue at 6 months.
Conclusions: ECM scaffolds promote healing of significantly sized tracheal defects without stricture and with some, but not all, of the necessary structures required for tracheal reconstruction, including complete coverage with ciliated epithelium and dense organized collagenous tissue.
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Introduction |
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Despite advances in surgical techniques, resection of airway stricture with primary repair can result in excessive anastomotic tension, tissue ischemia, and failure to heal. Although stents can be used effectively as a palliative therapy, they may be associated with a high complication rate in the treatment of nonmalignant airway strictures [4]. Stated differently, the surgical management of benign and malignant tracheal pathology represents a significant and clinically relevant problem, and alternative approaches are needed for tracheal reconstruction.
A variety of regenerative medicine approaches have been proposed for long (>5 cm), full circumferential tracheal replacement, ranging from collagen scaffolds supported by silicone stents to cartilaginous tubes created by in vitro culture methods [5–8]. However, these tracheal replacement strategies have been inadequate as a result of incomplete epithelialization with associated stricture, or a lack of mechanical integrity resulting in tracheomalacia [9].
Biologic scaffolds composed of naturally occurring extracellular matrix (ECM) promote site-specific tissue remodeling in both preclinical studies and for numerous clinical applications [10]. ECM scaffolds are degraded rapidly with concomitant site-specific deposition of host tissues. Degradation appears to be critical to the constructive remodeling response as matricryptic peptides formed as a result of parent molecule breakdown exhibit inherent bioactivity, including bacteriostasis [11, 12] and chemotaxis for differentiated and progenitor cells [13]. For repair of tracheal defects, one form of ECM scaffold, small intestinal submucosa (SIS-ECM), has shown moderate success for small defects (less than half the circumference) and defects with adequate structural support, but did not fully restore functional tracheal tissue [14–16].
The purpose of the present study was to determine whether an ECM scaffold with specific features, such as a basement membrane ECM or cartilaginous ECM, might enhance the remodeling response for a complex tissue like the trachea. Three different forms of ECM scaffold were used to repair a partial (ie, noncircumferential) defect in the ventral trachea in a canine model. In one group, the defect was repaired with an 8-layer multilaminate ECM scaffold referred to as urinary bladder matrix (UBM-ECM), which retains a basement membrane structure [17]. The second group was repaired with UBM-ECM patch, an 8-layer UBM-ECM patch that was crosslinked with 10mM carbodiimide (UBMC) to provide additional mechanical support to the device. The third group was repaired with a decellularized full thickness patch of porcine tracheal matrix (DTM) that contained both a basement membrane and the remnants of tracheal cartilaginous rings.
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Material and Methods |
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Harvest of Porcine Tissues
Urinary bladders and tracheas were harvested from market-weight pigs (approximately 110 to 130 kg) immediately after slaughter at an abattoir and transported to the laboratory on ice. Residual external connective tissues were then removed. The urinary bladders were further cleaned by repeated rinses to remove all residual urine. The urothelial layer was removed by soaking of the material in 1N saline for 15 minutes. All tissues were frozen at –80°C until time for continued processing, usually no more than 2 months.
UBM-ECM Device Preparation
Urinary bladders were thawed and the tunica serosa, tunica muscularis externa, tunica submucosa, and most of the muscularis mucosa were mechanically delaminated from the bladder tissue. The remaining basement membrane of the tunica epithelialis mucosa and the subjacent tunica propria, collectively termed urinary bladder matrix (UBM), were then decellularized and disinfected by immersion in 0.1% (v/v) peracetic acid, 4% (v/v) ethanol, and 96% (v/v) deionized water for 2 hours. The UBM-ECM material was then washed twice for 15 minutes with phosphate-buffered saline (pH = 7.4) and twice for 15 minutes with deionized water.
Multilayer tubes were created by wrapping hydrated sheets of UBM around a 22-mm perforated tube/mandrel that was covered with umbilical tape for 8 complete revolutions (ie, a layer tube) [18]. The constructs were subjected to a vacuum of 710 to 740 mm Hg (Leybold, Export, PA, Model D4B) for 10 to 12 hours to remove the water and form a tightly coupled multilaminate device. Each tubular device was cut to approximately 2-cm-wide x 3-cm-long pieces that were terminally sterilized with ethylene oxide.
For the carbodiimide crosslinked group, the 2- x 3-cm UBM devices were immersed in 10mM 1-ethyl-3-(dimethylaminopropyl)-carbodiimide hydrochloride (Sigma-Aldrich, St. Louis, MO) for 24 hours at room temperature. Each device was rinsed three times with sterile deionized water for 15 minutes to remove any residual crosslinking agent. The devices were vacuum-pressed around the same mandrel as previously described to dry specimens and were terminally sterilized with ethylene oxide.
Tracheal Tissue Matrix Scaffold Preparation
The tracheas underwent the following chemical treatment to decellularize the tissue: 1 hour in 0.25% Trypsin/0.05% ethylenediaminetetraacetic acid at 37°C, 48 hours in deionized water at 4°C, and 48 hours in 3% Triton X-100 at 4°C. The tissue was rinsed heavily to remove residual detergent and then disinfected with peracetic acid as described for UBM-ECM. The DTM material was frozen at –20°C and lyophilized. Devices were then cut (2 x 3 cm) from the ventral portion of the trachea (ie, from the portion that contained cartilaginous tissue) and were terminally sterilized with ethylene oxide.
Surgical Procedure
All animals were sedated with acepromazine (0.1 mg/kg intramuscular), followed by intravenous administration of thiopental (12 to 25 mg/kg), intubation and isoflurane (1.5–3%) maintenance of surgical place anesthesia. Cefazolin (15 mg/kg intravenous) was administered before surgical preparation and skin incision.
Aseptic technique was used to expose the proximal cervical trachea through a midline neck incision. A 1-cm-wide x 2-cm-long portion of the ventral tracheal wall was surgically removed. The defect was then repaired with one of the ECM scaffold devices with at least 5 mm of overlap around the edges of the defect (Fig 1A). All grafts were secured using absorbable 4–0 polydioxanone suture (PDS; Ethicon, Somerville, NJ). Non-resorbable Prolene suture (Ethicon) were used to mark the corners of the repair site. The scaffold placement site was tested for air leaks by submerging in saline while applying a Valsalva maneuver. The repair was also examined by bronchoscopy to verify graft placement and airway patency.
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Clinical and Bronchoscopic Assessment
Each animal had a predetermined follow-up period of either 2 or 6 months, at which time the dog was euthanized. Bronchoscopic examinations were conducted at 1, 2 and 6 months after the procedure to evaluate scaffold remodeling. Airway stenoses were evaluated using a flexible bronchoscope, and visually quantified as a percentage decrease in the ventrodorsal diameter of the trachea. Strictures were classified as mild (< 25%), moderate (25% to 50%) or severe (> 50%). Documented bronchoscopic data included appearance of the graft surface and its relationship to the native trachea, signs of inflammation (eg, exudate, granulation tissue), presence or absence of tracheomalacia, and stricture formation.
Morphologic Assessment
Animals were euthanized by sedation with acepromazine (0.01 mg/kg subcutaneous) and butorphanol (0.05 mg/kg), induced anesthesia with isoflurane (5%) for 5 minutes, and intravenous administration of pentobarbital sodium (390 mg/4.5 kg). The scaffold placement site, including native tracheal tissue surrounding the graft site, was harvested immediately after euthanasia. The excised sample was exposed by a longitudinal incision in the distal-to-proximal direction along the membranous portion of the trachea. The exposed mucosal surface was examined and photographed.
The tissue was immersed in 10% neutral buffered formalin for histologic preparation. Tissue was trimmed longitudinally, sectioned, and stained by conventional histologic staining with Masson's trichrome and periodic acid Schiff (PAS), and with immunohistochemical staining for p63. The p63 staining was performed after antigen retrieval with citric acid buffer (pH = 6.0). The primary antibody was a mouse antihuman p63 (Dako Inc, Carpinteria, CA) at a 1:25 dilution for 1 hour at room temperature. The secondary antibody used was antimouse immunoglobulin (Ig) G at 1:500, followed by detection with HRP-streptavidin also at 1:500. Diaminobenzidine was the substrate, and the slides were counterstained with Harris' hematoxylin. The areas examined included the native tissue, the proximal and distal interfaces between the remodeled and native tissue, and the middle region of the remodeled area.
Cell counts were performed for secretory cells identified by positive PAS staining, and basal cells were identified by p63-positive staining within longitudinal sections of the remodeled tissue in each specimen using 200x images in the MetaVue Software package (Molecular Devices, Sunnyvale, CA). The length of the scaffold was also measured so that the number of cells could be related to the distance from the edge of the remodeled graft as defined by the presence of host cartilage. For qualitative comparison, the cell counts were interpolated for cell counts over every 1000 µm and the average number of cells in that region was averaged for each type of scaffold at each time point and presented from the proximal edge of the defect to the point of interest.
Semiquantitative assessment of the presence of ciliated cells was conducted. A score from 0 to 3 was assigned to each 200x field (Table 1). An interpolated score was then calculated for each 1000-µm region across the remodeled area.
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Results |
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Bronchoscopic Examination
Bronchoscopic evaluation of the remodeled tracheal defect sites repaired with UBM, UBMC, and DTM groups all showed a smooth, shiny surface consistent with the presence of an epithelial layer and neovascularization at 1 month. In the UBMC group, there was evidence a whitish exudate throughout the first 2 months. One dog in the UBMC group and 1 dog in the DTM group showed mild asymptomatic tracheal stenosis at 1 month, but in both cases the stenosis was completely resolved at 2 months. A small asymptomatic nodule at the distal aspect of the graft site developed in 1 dog in the UBM group, which may have been related to the sutures. At 6 months, the remodeled patch appeared similar to that observed at 2 months, with the exception that the vascular response was less pronounced and the newly formed tissue had more of a whitish appearance (Fig 1B).
Macroscopic Observations
For all three groups, gross observations showed remodeling of the scaffold with dense, fibrous connective tissue with a shiny epithelial layer covering the entire surface (Fig 1C). New blood vessels were evident in the remodeled tissue, and these vessels were slightly more visible at 2 months compared with 6 months. There was evidence of inflammation localized around the sutures at 2 months, but this also diminished by 6 months because the sutures had resorbed. The remodeled scaffold material in all groups was pliable without visible evidence of mechanical support from cartilage formation.
Microscopic Observations
Histologic examination of the UBM, UBMC, and DTM grafts showed complete degradation and replacement of the device with host tissue by 2 months after the procedure (Figs 2A,
3A, and 4A, respectively). The site of remodeling for all three devices showed dense, organized collagenous tissue with complete epithelialization. In all three groups, there was histologic evidence of neovascularization at 2 months that diminished slightly by 6 months. The UBMC group showed slightly increased mononuclear cell numbers in the dense collagenous tissue subjacent to the epithelium compared with either the UBM or DTM groups. There was no evidence of cartilage formation within any of the remodeled scaffold materials.
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UBMC
For the UBMC group, ciliated epithelial cells were observed at the edges of the remodeled scaffold, but no ciliated cells were present in the center of the patch (Fig 3A). The number of ciliated cells present increased from 2 months to 6 months (Fig 3B). At both 2 and 6 months, the number and distribution of secretory cells was less than observed in native trachea but was similar to that observed for the UBM group at 2 months (Fig 3C). There was no change in the number or distribution of secretory cells between the 2- and 6-month time points. The remodeled UBMC showed no basal cells over most of the remodeled area at 2 months, but by 6 months, there was a uniform distribution of basal cells similar to that observed for the UBM group (Fig 3D).
DTM
The DTM group showed large regions with no ciliated epithelial cells in the central portion of the remodeled graft at 2 months, and this deficiency was more pronounced after 6 months (Figs 4A and B). The DTM group showed the fewest number of secretory cells of the three groups at both time points, with a complete lack of secretory cells in the middle portion of the remodeled graft at 6 months (Fig 4C). At both time points, the middle portion of the remodeled grafts showed little or no presence of basal cells (Fig 4D).
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Comment |
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The results of the present study are similar to those observed when SIS-ECM was used for patch tracheoplasty in both a rat and rabbit model [14, 15, 19]. The remodeled SIS-ECM formed a fibrous connective tissue that showed ciliated epithelialization. No cartilage formation was found even when thyroid or auricular cartilage was colocalized at the site of remodeling [15, 19]. In fact, the presence of the cartilaginous tissue contributed to an increased rate of complications. The only study in which neochondrogenesis has been reported associated with SIS-ECM repair of the trachea was in a porcine model in which SIS-ECM was used to coat a self-expanding metallic stent [16]. However, the presence of the metallic stent led to significant luminal narrowing.
The use of carbodiimide to chemically crosslink UBM in the present study was intended to slow enzymatic degradation and increase the strength of the scaffold [20]. It was hypothesized that the crosslinked scaffold would remain at the site of implantation longer and provide mechanical support. Although a temporal and quantitative evaluation of scaffold degradation was not conducted in the present study, there was no histologic evidence to suggest delayed degradation of the UBMC scaffolds.
We hypothesized that the DTM graft would provide mature epithelial development and chondrogenesis due to the organ-specific composition and organization of the ECM. In fact, epithelialization of the DTM scaffold was qualitatively worse than either UBM or UBMC, with less coverage with ciliated epithelium, fewer secretory cells, and fewer basal cells. The decreased population of basal cells may have contributed to the lack of ciliated and secretory cells, because basal cells serve as a progenitor cell for the tracheal epithelium [21, 22].
A recent study showed that lyophilization of UBM caused irreversible changes to the ultrastructure of UBM and slowed cell growth on and penetration into the scaffold in vitro [23]. A similar phenomenon may explain the suboptimal results observed for the lyophilized DTM in vivo. Decellularized allogenic canine tracheas in a hydrated form have been evaluated for repair of full circumferential resections of the intrathoracic trachea with considerable success [24, 25]. The detergent treatment was similar to that used in the present study. A key aspect of this approach was the viability of the cartilage within the allograft.
The presence of chondrocyte material within the cartilage was highly predictive for successful patency of the tracheal allograft [24]. It is not clear whether the chondrocytes were viable and therefore participated in the remodeling response, or if the preserved mechanical integrity of the cartilage simply allowed the graft to maintain enough strength to resist the negative pressures of respiration. If the integrity of the cartilage was the critical factor determining success, then the lack of viable cartilage due to lyophilization may partially explain the lack of cartilage formation for the DTM in the current study. Lyophilization of the DTM likely disrupted glycosaminoglycans in the cartilage and compromised the ability of the scaffold to provide mechanical support during the remodeling process. It is unknown whether a dog would reject cellular material remaining in viable porcine cartilage, but in the canine allograft experiments, the chondrocytes did not express major histocompatibility complex [25].
The current study showed that the presence of a vacuum-pressed UBM graft or a lyophilized form of decellularized tracheal matrix is insufficient to promote the complete regeneration of functional trachea tissue. However, the results shed light on modifications that may be incorporated to an ECM-based airway replacement graft to increase its regenerative potential. For instance, the use of a hydrated form of ECM with preserved cartilage integrity may be necessary in order to realize functional tracheal replacement.
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Discussion |
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DR GILBERT: Indeed we are working towards a full circumferential tracheal replacement. At this point in time, we are focusing our efforts on the graft decellularization process. In the experiments I discussed today, the decellularization of the porcine tracheal graft was extensive, resulting in near complete destruction of cellular components found in the cartilaginous rings and the surrounding tracheal tissue. In addition, the graft was lyophilized or freeze dried. This process may also have had a negative impact on the structural properties of the porcine tracheal graft, giving it a softer consistency once rehydrated in vivo.
Future experiments may involve the use of hydrated porcine tracheal grafts submitted to a less aggressive decellularization process. We would hope to preserve the integrity of the cartilaginous components while completely eliminating the epithelium. Thank you for your question.
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Acknowledgments |
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References |
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This article has been cited by other articles:
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  J. Schanz, M. Hampel, H. Mertsching, and T. Walles Experimental Tracheal Patching Using Extracellular Matrix Scaffolds Ann. Thorac. Surg., April 1, 2009; 87(4): 1321 - 1322. [Full Text] [PDF]
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T. W. Gilbert Reply Ann. Thorac. Surg., April 1, 2009; 87(4): 1322 - 1323. [Full Text] [PDF]
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