Acellular dermal matrices (ADMs) are commonly used in wound healing and tissue repair to facilitate wound closure and regenerative remodeling.1–3 The extracellular matrix (ECM), a major component of ADMs, provides structure, cell-signaling cues, and mechanical support to facilitate the healing process.4–8 Key dermal ECM components include collagens, elastin, glycosaminoglycans (GAGs), and hyaluronic acid (HA).4,9–11 The ECM can sequester and control the bioavailability of growth factors that modulate cellular responses by serving as a growth factor reservoir.4,5,12 Apart from providing biological cues, the ECM imparts mechanical properties in the form of structural, tensile, and compressive support.13,14 Its architecture influences material stiffness, which regulates cell behavior by affecting cytoskeletal reorganization and cell signaling,15,16 whereas an open microstructure can facilitate host cell infiltration.17 These native dermal properties can guide cell behavior and tissue remodeling in a wound care setting.
Exogenous scaffolds replace or replicate native ECM by restoring structural and functional requirements.1,12,18 They also provide a barrier to protect wounds from infection and desiccation. Scaffold origins can be cellular or acellular and originate from biological, synthetic, or composite materials.1,19 Although synthetic scaffolds are reproducible and uniform, they lack the biological advantages of native dermal matrices.5,19 ADMs can be processed to preserve the dermal structure and leverage the dermal biology to reduce scarring and improve tissue regeneration.2,20–22
The structure of human dermis can be divided into 2 layers: papillary or superficial and reticular.1 The fibrils present in the papillary dermis are smaller compared with the reticular dermis. When the papillary dermis is injured (superficial cut or burn), it can often regenerate without a scar. The reticular dermis is the deeper and thicker region composed of dense collagen fibers, elastin, and woven reticular fibers. These characteristics provide this region with strength, extensibility, and elasticity.23 In a deep wound, this framework is missing, which can lead to scarring. By using an organized structure, this can coordinate new tissue repair and potentially address scarring.
ADM processing aims to remove cellular material to reduce immunogenicity and decontaminates or sterilizes the graft to limit disease transmission.2,24 If not designed appropriately, however, the processing can negatively impact the endogenous matrix proteins and natural architecture that can hamper host cell integration and result in encapsulation and foreign body response.25,26 Aseptic tissue processing utilizes gentle decontamination steps to ensure tissue safety, while preserving the matrix configuration.
In this study, we investigate the hypothesis that aseptically processed reticular dermal grafts provide a scaffold possessing biological and mechanical properties that can support wound healing. This unique deeper cut reticular dermis retains architectural elements (open structure), mechanical properties (elasticity, organized collagen and elastin), and key matrix proteins to support physiological cellular responses during regenerative remodeling.
MATERIALS AND METHODS
Human dermal tissue was screened and recovered following industry standard guidelines. Human reticular ADMs (HR-ADMs; AlloPatch Pliable, Musculoskeletal Transplant Foundation, Edison, N.J.) was processed aseptically (Fig. 1) without terminal sterilization at Musculoskeletal Transplant Foundation (Edison, N.J.). The tissue was decellularized and disinfected with peracetic acid–based solution and every lot was assessed as per 〈USP-71〉 Sterility Tests. The papillary dermis was prepared in a similar method as a comparison. Both reticular and papillary layers were cut to final tissue specifications of 0.4–1.0 mm thick.
Prehydrated papillary and reticular dermis samples (n = 3 donors) were fixed in 10% formalin, embedded in paraffin, sectioned into 5 μm thick cross-sections, and stained for hematoxylin and eosin (H&E) by Premier Laboratory, LLC (Boulder, Colo.).
Scanning electron microscopy (SEM) imaging was performed to visualize the microstructure of HR-ADM in comparison to papillary dermis (n = 3 donors). Samples were fixed in 10% formalin for 24 hours, rinsed with water twice for 15 minutes, and dehydrated in 50%, 70%, 80%, 95%, and 100% ethanol successively for 15 minutes each. These dry samples were coated under vacuum using a Balzer MED 010 evaporator (Technotrade International, Manchester, NH) with platinum alloy to a thickness of 25 nm and immediately flash carbon coated under vacuum. Samples were examined in a JSM-5910 SEM (JEOL USA, Inc., Peabody, Mass.) at an accelerating voltage of 25 kV. Imaging was conducted at 250×.
The material porosity (n = 3 donors) was determined through gravimetric method assuming the material is close to density of collagen (1.34 g/cm3), as collagen is the largest component of dermal tissue. The density of the tissue (using a ratio of dry and wet tissue densities) was calculated as per Loh and Choong.27 The pore size range was evaluated by mercury intrusion porosimeter (Quantachrome, Fla.) using standard techniques. Mercury is forced into the dermal sample under high pressure through the porosimeter. The pressure needed to force mercury into the sample is inversely proportional to the pore size.
Biomechanical properties of dermal tissue were evaluated using a MTS 858 Mini-Bionix tensile testing system (MTS, Eden Prairie, Minn.) with a calibrated 1 kN load cell. Dermal grafts (n = 3 donors) were cut into multiple specimens (4–12) using a custom punch shaped and sized to match the type V (microtensile testing) specimen specified by the American Society for Testing and Materials (ASTM) D638 guidelines for evaluating material properties (3.18 mm width; 0.4–1.0 mm thick). Sample thickness was measured and then loaded into tensile grips. Specimens were pulled under tensile load at a rate of 50.8 mm/min until failure. Ultimate tensile stress (UTS), Young’s modulus, and the percentage elongation at break were examined and normalized to cross-sectional area.
Matrix Protein Characterization
Immunohistochemistry staining was performed at HistoTox Labs, Inc. (Boulder, Colo.). Operators were blinded to HR-ADMs (sample 1) and unprocessed reticular dermis (sample 2) for collagens I, III, IV, and VI and elastin. The levels of GAG and HA in HR-ADMs were compared with unprocessed reticular dermis (n = 3 donors). GAGs were quantified using the Blyscan GAG assay (Biocolor Life Science Assays; Carrickfergus, UK/Fisher Science, Houston, Tex.). Samples were extracted in papain (125 μg/mL in 0.1 M phosphate buffer) for 2 hours at 65°C and centrifuged (10,000 rpm; 10 minutes). The dye-binding assay was performed and absorbances were read at 656 nm. HA was quantified using an enzyme-linked immunosorbent assay (Corgenix, Broomfield, Colo.). Samples were extracted (24 hours, 4°C in 1 M sodium chloride and sodium bicarbonate solution), homogenized for 5 minutes in a bullet blender (Next Advance, N.Y.), and centrifuged (10,000 rpm; 10 minutes). Absorbances were read at 450 nm.
HR-ADM samples were air dried overnight, weighed (21–25 mg), and rinsed in 0.9% saline solution. Samples (n = 3 donors) were then enzymatically digested (6 hours, 37°C water bath) in a collagenase type 1A (6.65 U/mL final enzyme solution; Sigma, St. Louis, Mo.) and thermolysin (15 U/mL final enzyme solution; Sigma, St. Louis, Mo.) solution in tricine buffer (pH 7.5). The filtered extract was mixed with ninhydrin (0.016 g/1 mL solution)–hydrindantin (0.0024 g/1 mL solution) (Sigma, St. Louis, Mo.) in ethylene glycol monoethyl solution and 4 N sodium acetate buffer (pH 5.5) that reacts with the released amino acids, producing a deep purple color proportional to the amount of peptides released. The standard curve was established with L-leucine (stock solution 2.0 mg/mL; Sigma, St. Louis, Mo.) and sample absorbances were read at 570 nm. The controls were crosslinked28 and denatured dermis samples. Unprocessed dermal tissue was crosslinked (16 hours, room temperature) with 0.025% glutaraldehyde (Sigma, St. Louis, Mo.) solution, followed by a 2-hour rinse step to remove residual glutaraldehyde. The denatured condition (representing harsh chemical processing) was prepared by crosslinking as above and then boiling (at 100°C) the rinsed samples for 5 minutes. These samples were digested as stated above, reacted, and read at 570 nm.
Cell Behavior Characterization
Normal human dermal fibroblasts (NHDFs; Lonza, Walkersville, Md.) were cultured (0.2 million cells/7 mm disk) on HR-ADMs in fibroblast growth medium (FGM-2) (Lonza) at 37°C and 5% CO2 in a humidified atmosphere. Cell attachment and matrix production were assessed over time (0, 7, 14 days). H&E, collagen IV, and fibronectin staining were performed by IHC World, LLC (Ellicott City, Md.), with standard histology techniques. Human umbilical vascular endothelial cells (HUVECs) were cultured (0.2 million cells/7 mm disk) on HR-ADMs to examine angiogenic capacity through tubular formation (CD31, AbCam, Cambridge, Mass.) and secretion of functional angiogenic factor, von Willebrand Factor (vWF; AbCam, Cambridge, Mass) on adhered cells through 4', 6-diamidino-2-phenylinadole (Life Technologies, Carlsbad, Calif) staining. Confocal imaging (Rutgers University, Piscataway, N.J.) was performed to visualize tubular network formation and vWF secretion.
All values are reported as average and SD. A student t-test (unpaired) was used to compare mechanical evaluation of dermal tissue (HR-ADM versus papillary), and enzymatic degradation analysis (unprocessed dermis to HR-ADM, crosslinked and denatured), with P < 0.05 being considered significant.
Unique Feature of HR-ADM
H&E staining of HR-ADM revealed an open, uniform architecture (no orientation or polarity; Fig. 2). In contrast, the papillary graft was asymmetrical (epidermal-facing side was dense versus the open dermal-facing side), resulting in directionality (distinct orientation or polarity). Higher magnification (20×) images clearly demonstrated the consistent open, interconnected network of HR-ADM compared with the asymmetrical papillary dermis (See PDF, Supplemental Digital Content 1, which reveals distinct structural differences between papillary and reticular dermal structures, http://links.lww.com/PRSGO/A274). Additionally, SEM imaging (Fig. 3) confirmed the open architecture present in HR-ADMs compared with papillary dermis. The porosity of HR-ADM was 88% ± 4% and of papillary dermis was 82% ± 6%. The pore size range as determined by mercury intrusion for HR-ADMs was 2.7–500 μm, whereas that determined for papillary dermis was 0.8–500 μm (Table 1). This open, interconnected network in HR-ADMs was seen in Figure 2, and this pore size range supports cell infiltration as evidenced in Figure 7.
The HR-ADM thickness was 0.8 ± 0.2 mm (n = 4 donors; 4–8 samples/donor), whereas the papillary dermis thickness was 0.7 ± 0.1 mm (n = 3 donors; 8–12 samples/donor). HR-ADMs exhibited lower UTS values (7 ± 2 MPa) and Young’s modulus (6 ± 1 MPa) compared with the papillary dermis UTS (14 ± 3 MPa) and Young’s modulus (15 ± 3 MPa). HR-ADMs had significantly lower UTS (P = 0.03) and Young’s modulus (P = 0.019) values compared with papillary dermis (Table 1). These lower biomechanical properties of HR-ADMs were similar to those reported for fetal porcine dermis as an elastic biomaterial comparison. The percentage elongation at break was significantly greater (P = 0.03) for HR-ADM (131% ± 15%) compared with papillary dermis (104% ± 2%), and is expected in an elastic scaffold. Generally, open porous scaffolds under tension align first, stretch, and break, whereas dense scaffolds, which have some orientation, load first and then break resulting in lower percentage elongation at break. Therefore, the biomechanical testing confirmed that the HR-ADMs are flexible structures, exhibiting low stiffness and increased elasticity.
Native ECM Components Preserved
Immunohistochemistry staining qualitatively revealed the retention of organized collagen types I and VI and elastin (Fig. 4) in unprocessed reticular dermis and HR-ADMs after aseptic processing (See PDF, Supplemental Digital Content 2, which demonstrates collagen III and IV retention, http://links.lww.com/PRSGO/A275). Although there was some reduction in staining intensity for collagen III, the majority of the ECM components in HR-ADMs are similar to unprocessed tissue. Additionally, GAGs and HA were present and quantified in unprocessed reticular dermis (2.5 ± 0.1 mg/g; 1000 ± 88 μg/g) and retained in HR-ADMs (2.7 ± 0.6 mg/g; 272 ± 51 μg/g), respectively (Fig. 5). Although lower HA levels were present in the processed HR-ADM as compared with the unprocessed sample, a considerable amount of HA is retained. These critical ECM components provide an organized architecture to support cellular activities.
To verify that aseptic processing preserves the native dermal components, enzymatic degradation studies examined the release of peptides in unprocessed and processed tissue samples along with controls (crosslinked, denatured dermis) representing process-altered tissue. Peptide release for aseptically processed HR-ADMs (4.5 ± 0.4 mg/g of tissue) was similar (no significant difference) to that for unprocessed reticular tissue (4.0 ± 0.5 mg/g of tissue; Fig. 6). Crosslinking dermis resulted in significantly (P = 0.004) lower peptide (0.6 ± 0.1 mg/g of tissue) release (resistance to degradation) due to the crosslinked collagen structure, whereas denatured dermis yielded significantly (P = 0.013) higher peptide (6.6 ± 0.6 mg/g of tissue) release (degraded collagen); both reflecting altered tissue components. Therefore, aseptic processing preserves the native dermal components, whereas other processing methods may alter it.
In Vitro Cell Studies
Histological and confocal imaging demonstrated that both NHDFs and HUVECs readily attached to HR-ADMs. Histology analysis of NHDFs revealed cell attachment and infiltration into the graft (Fig. 7). Immunohistochemistry analysis confirmed fibroblasts secreted an abundance of collagen IV in a multilayered network on top and within the open HR-ADM network as early as day 7 (See PDF, Supplemental Digital Content 3, which displays fibronectin secretion and day 14 images, http://links.lww.com/PRSGO/A276). HUVECs also readily attached (4', 6-diamidino-2-phenylinadole staining) with endothelial marker CD31, highlighting distinct, sustained tubular, network formation, and vWF, which is secreted by functional endothelial cells and confirming angiogenic capacity (Fig. 8). Similar observations were found on both sides of the HR-ADM. Both fibroblasts and endothelial cells are functional on HR-ADMs by attaching and secreting matrix proteins, which support granulation and angiogenic activities.
ADMs are used to protect the wound surface, maintain hydration, and provide a conducive microenvironment for dermal repair and regeneration.2,6,18,21,29 Traditionally, these matrices are obtained from the papillary dermal layer and are processed by methods that can alter the native dermal architecture and tissue quality, thereby impacting host engraftment and tissue remodeling. The novel dermal graft (HR-ADM) obtained from the deep reticular dermis layer used in this study was aseptically processed, and this preserved the native architecture and key ECM components that facilitate graft integration.
Histological analysis of the papillary dermis revealed an asymmetrical network (dense on one side, open on the other); this architectural polarity within the papillary region has been reported previously.1 The heterogeneous nature can impact cell infiltration and native tissue remodeling. The novel HR-ADM provided a uniform, open network, ensuring a homogeneous framework. The absence of any graft asymmetry or orientation can be beneficial in the clinical setting, facilitating the ease of use.
It is well known that open scaffold architectures modulate cell–matrix interactions and augment cellular activities and newly formed tissue.30,31 Increasing porosity can significantly improve cellular infiltration and tissue integration,30 whereas the intrinsic mechanical properties (stiffness, elongation) can regulate cellular behavior (proliferation, cell–matrix integration).32–34 This study demonstrated that HR-ADMs had an open, interconnected network with elastic biomechanical properties that are similar to fetal skin and significantly lower than papillary dermis. From literature, papillary dermis exhibited biomechanical properties which are in alignment with our study; similar Young’s modulus (18.4 MPa) for an ADM,35 obtained from papillary dermis, and UTS values (22 ± 8 MPa36 and 13–30 MPa13) were also observed. In contrast, HR-ADMs behaved similarly to fetal porcine tissue having low biomechanical properties: UTS (2.1 ± 0.3 MPa) and Young’s modulus (5.9 ± 1.5 MPa).37 Exhibiting similar biomechanical properties to fetal porcine tissue may culminate in reduced scar formation.37–39 As human wounds heal, the stiffness has shown to increase from 18 to 40 kPa,40 indicating wound bed fibrosis and scarring.41,42 Now, depending on clinical applications, different biomechanical properties are necessary. In the wound setting, graft strength is not critical, whereas the elasticity, flexibility, and conformability to the wound topography and irregular wound sizes are advantageous. Hence, HR-ADMs provide a promising elastic scaffold for wound repair.
This study also demonstrated that aseptic processing preserved ECM components important for wound healing, including collagens and elastin. They provide a stable, organized structure along with signaling cues that facilitate wound healing.4,12 Collagens are instrumental in supporting the wound healing phases8,22; homing inflammatory cells3,7; supporting fibroblast attachment/granulation10,43; and facilitating keratinocyte migration.44 Fetal fibroblasts have been shown to express more collagen III than collagen I and promote a more reticular deposition of fibers in a basket-weave orientation, which can assist in minimizing scar formation.45,46 This type of reticular collagen network can help promote regeneration versus repair and minimize scarring; that is distinctly different from disorganized, parallel bundles of collagen that cause scarring.38 Elastin provides scaffold elasticity and also mediates cellular activities by regulating the activity of TGFβs and its presence/organization minimizes scar formation.47–49 Therefore, the preserved organized, basket-weave collagen and elastin structure present in HR-ADMs may promote regenerative healing.
GAGs and HA, which are important biological components for both adult and fetal wound healing,20,50 are also retained in HR-ADMs. Exogenous addition of HA has reduced scar formation in adults by harnessing local growth factors and modulating cell behavior.51,52 GAGs also have an effect on chronic inflammatory response and fibrotic encapsulation, which otherwise may progress to implant failure or scarring.53 GAGs and HA can influence the ECM structure, assembly, and hydration38; impact inflammation50,51; and foster granulation and protect cells from free-radical damage.25 The retention of both GAGs and HA in HR-ADMs is predictive of clinical utility in facilitating wound healing.
These key biological ADM properties are beneficial to be retained through the processing steps to remove bioburden and minimize immunogenicity. Terminal sterilization and harsh chemical treatments can modify the scaffold structure and degradation characteristics by resident enzymes found in wounds.54–56 Furthermore, these treatments can damage the collagen structure and other bioactive components, hampering cell adhesion and cell–ECM interactions.54,55 Crosslinking agents (such as glutaraldehyde28) strengthen scaffold biomechanical properties; however, they reduce the ability of cell–matrix interactions by impairing the collagen structure and cell-binding sites, yielding poor clinical properties.26 Consequently, tissue processing strategies must balance bioburden reduction and cell removal with maintenance of scaffold integrity. This study verified aseptic processing retained the native dermal components. Furthermore, enzymatic degradation of HR-ADM yielded similar peptide release compared with unprocessed tissue, whereas crosslinking or denaturing dermis significantly altered peptide release.
Further evidence that aseptic tissue processing preserved the native architecture and biological components comes from in vitro fibroblast and endothelial cell studies in HR-ADMs. The open architecture of HR-ADMs (2.7–500 μm pore size range) and retained ECM components supported cell attachment and infiltration. Both cell types readily attached and were functional on the HR-ADMs by secreting an abundance of new matrix proteins (collagen IV, fibronectin, vWF) on top and within the graft. The secreted matrix proteins are critical in supporting granulation and angiogenesis4,5,57,58 and stimulate and guide other cellular responses.10,59
In summary, aseptically processed HR-ADMs provide a unique, biologically and mechanically advantageous scaffold for wound repair. The in vitro findings are supported by the clinical findings where HR-ADMs combined with standard of care performed significantly better than standard of care alone in the treatment of chronic diabetic foot ulcers.60 Further in vitro studies are needed to characterize the cell behavior and functionality of these biologically and mechanically stable novel reticular dermal grafts in a chronic setting.
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Supplemental Digital Content
Copyright © 2016 The Authors. Published by Wolters Kluwer Health, Inc. on behalf of the American Society of Plastic Surgeons. All rights reserved.