Approximately 30 million people in the United States have some form of liver disease.1 Because the only approved cure for end-stage liver disease or acute liver failure is whole or partial organ transplantation, there remains a significant demand on transplantable donor organs.2 This lack of available organs has led to approximately 27 000 deaths annually in the United States alone.1,3 A number of strategies, ranging from cell4-8 to engineered liver tissue transplantation, are currently under investigation to help alleviate the demand for donor organs.8-12
Regardless of the approach, the success of any cell therapy hinges on the long-term survival and function of the transplanted cells. Ectopic transplantation of single and multilayer sheets of hepatocytes has shown some success in maintaining liver tissue function.13 However, given the space constraints to reasonably scale up this technique, sheets would have to be stacked. This will inevitably result in the diffusion barrier being breeched, thus necessitating the incorporation of vasculature. Another approach has been the use of biomaterials as scaffolds to engineer 3-dimensional liver tissues.14 However, for these to function, the implants must be sufficiently thin. Furthermore, studies have shown that such implants function better when placed in a heavily vascularized area,14,15 which usually requires an invasive surgery.
In an effort to improve the efficacy and function of the engineered tissues, decellularized liver organs have been used as scaffolds.16,17 Decellularized livers can provide the structural and biochemical cues necessary to maintain viability and function of primary hepatocytes.18,19 Furthermore, if used as intact structures, they provide an existing architecture that facilitates vascularization and provides liver-specific geometrical cues. In this approach, because of the existing shortage of human organs, porcine or other xenogeneic livers have to be used. Thus, it is necessary to ensure proper decellularization to preserve the extracellular matrix (ECM) while ensuring the destruction of xenogeneic DNA. Although low DNA content (under 50 ng double stranded DNA per mg ECM) may not have triggered an immune response in tested animals, it can still carry the risk of immune rejection in humans.20,21
Bioengineered devices that can facilitate vascularization of the implant while supporting the viability and function of transplanted cells could be a potential solution to improve the outcome of cell transplantation.22 Enabling vascularization of the implant will allow the implant size to be scaled and a large number of cells to be housed within the device, which is necessary to improve the therapeutic outcome. In this study, we describe the use of a dual-compartment device for minimally invasive hepatocyte transplantation. Recently, we used such an approach successfully to support bone marrow transplantation.23 Building upon this dual-compartment concept, we optimized the biomaterial composition and dimensions to develop constructs with higher cell-carrying capacity, capable of maintaining long-term hepatocyte function and promoting vascularization. The dual-compartment system consists of an outer interconnected, macroporous solid structure to promote vascularization and/or to house supporting cells and a hollow inner compartment to load the donor cells. When implanted subcutaneously in mice, the cell-loaded dual-compartment device supported the viability and sustained function of transplanted primary human hepatocytes (from 2 different donors) for at least 1 month (the longest experimental time investigated).
MATERIALS AND METHODS
Polyethylene Glycol Diacrylate Synthesis
Polyethylene glycol diacrylate (PEGDA) (Mn = 10 kDA) oligomer was prepared according to a previously reported method.24 Briefly, 18.0 g of PEG was dissolved in 300 mL of toluene in a 500-mL round bottomed flask in an oil bath heated at 125°C. The solution was refluxed for 4 hours with vigorous stirring. Traces of water in the reaction mixture were removed by azeotropic distillation. Upon cooling, the solution to room temperature (RT), 3.262 g (32.2 mmol, 4.493 mL) of triethylamine was added to it with vigorous stirring. Then the flask was moved to an ice bath and stirred for 30 minutes. 2.918 g (32.2 mmol, 2.452 mL) of acryloyl chloride in 15 mL of anhydrous dichloromethane was then added to the reaction mixture dropwise over 30 minutes. After keeping the reaction mixture in the ice bath for another 30 minutes, the flask was heated to 45°C overnight. The reaction mixture was then cooled to RT, and the quaternary ammonium salt was removed from the mixture by filtration. The filtrate was condensed using a rotary evaporator and precipitated in excess diethyl ether. The white precipitate was collected by filtration and vacuum dried at 40°C for 24 hours. The resultant PEGDA oligomer was purified by precipitation followed by column chromatography and dialysis before its usage. The purified PEGDA was lyophilized and stored at −20°C.
Hyaluronic Acid Methacrylate Synthesis
Sodium Hyaluronate (Lifecore Biomedical), Research Grade, 41 to 65 kDa Mw (500 mg) was dissolved in DI water (25 mL). Methacrylate (MA) anhydride (8 mL) was added into the hyaluronic acid (HA) solution (drop-by-drop manner), pH was adjusted to 8, and the reaction was carried out at 4°C for 24 hours. pH was checked frequently and adjusted to 8 as needed. After 24 hours, the resulting mixture was purified using membrane dialysis (3.5-5 kDa) against Milli-Q water for 3 days, lyophilized, and stored at −20°C.
Porous Scaffold Formation
The porous scaffolds (either PEGDA or PEGDA/hyaluronic acid MA [HAMA] copolymer) were made by the leaching of polymethyl MA (PMMA) beads to create the macroporous structure.25 Briefly, 160 μm PMMA beads were packed in a 10-mm diameter by 3 mm height mold. Eighty microliters of 20% acetone solution (in ethanol) was added to the PMMA filled mold before it was placed in a 37°C oven for 10 minutes. To this PMMA filled mold, a PEGDA/HAMA solution (10%/5% w/v mixture in phosphate-buffered saline [PBS]) or a 10% PEGDA (w/v mixture in PBS) containing 0.005% (w/v) Irgacure (ie, a photoinitiator) was added and UV polymerized for 10 minutes. Acetone was used to dissolve the PMMA beads to create the macroporous hydrogel structures (Figure 1A). The structure was sterilized using multiple ethanol washes, followed by multiple PBS washes.
A 7-mm punch was used to cut out the center of the porous gels to leave a hollow ring. The 7-mm inner portion that was removed was sliced to make caps for the hollow ring. Both the hollow rings and the caps were sterilized with ethanol then washed multiple times with PBS. Under sterile conditions, the ring and the caps were dried to remove the solvent from the pores. Then, supporting cells (some combination of human umbilical vein endothelial cells [HUVECs], human bone marrow stromal cells [hMSCs], or mouse embryonic fibroblasts [MEFs]), were loaded into the outer compartment and the caps. The primary human hepatocytes were thawed in thawing media (MCHT50; Lonza) and centrifuged for 10 minutes at 100 g. Next, fibrinogen (8 mg/mL) and thrombin (2 U/mL) were added to the cell pellet. Fibrin was allowed to form at 37°C and the system was maintained at this temperature for up to 30 minutes to complete the reaction. Once the gel was formed, it was placed into the inner cored out compartment of the porous gel (now consisting of the hollow ring and the bottom cap). The top cap is then placed to seal the dual-compartment system. This resulted in a cell-laden fibrin gel inner compartment, encased by a macroporous gel outer compartment (Figures 1 B and C). Fibrin was then used around the caps as an added precaution to seal them in place. Each device was loaded with approximately 5 million hepatocytes (or 5-20 million when testing loading capacity) and approximately 5 × 105 supporting cells. Human umbilical vein endothelial cells, hMSCs, and MEFs were used either alone (5 × 105 HUVECs) or in combination (2.5 × 105 HUVECs plus 2.5 × 105 hMSCS or MEFs) as supporting cells. To determine the effect of the scaffold to facilitate in vivo vascularization, acellular PEGDA and PEGDA/HAMA constructs were used.
Dual-compartment systems made from PEGDA macroporous hydrogels and PEGDA/HAMA macroporous hydrogels were assembled as described above, however, without the inner fibrin compartment. Gels were flash frozen and lyophilized for 2 days. Then 5 gels for each group were weighed to get the dry weight. Gels were then placed in PBS and weighed periodically over a 48-hour period to determine the wet weight. The wet weight was divided by the dry weight to get the swelling ratio and swelling kinetics.26
Scanning Electron Microscopy
The microstructure of the PEGDA/HAMA hydrogels was examined using a scanning electron microscope (SEM). Briefly, samples were thinly sectioned, flash frozen, and lyophilized for 2 days. Then using a sputter coater (Emitech, K575X), Iridium was coated onto samples for 7 seconds. The iridium-coated samples were imaged using an SEM machine (Phillips XL30 ESEM). The diameter of the interconnected pores was measured using ImageJ by randomly selecting 10 pores from each of the 3 different SEM and bright-field images, respectively, and presented as mean ± standard deviation (n = 30).
Primary human hepatocytes from 2 different donors (Donor 1, HUM4100 and Donor 2 HUM4113) were acquired from LONZA (formerly TRL). The cells were thawed in thawing media (MCHT50, LONZA) and immediately encapsulated into fibrin gel and loaded into scaffolds. The cells encapsulated in fibrin gel were loaded as described above. The cell-laden dual-compartment scaffolds were cultured in 4 parts maintenance media (MM250, LONZA) and 1 part HUVEC media (components described below).
Human Umbilical Vein Endothelial Cells were obtained from American Type Culture Collection and cultured in HUVEC medium containing 79% M199 medium (Gibco), 10% fetal bovine serum (FBS) (Gibco), 10% endothelial cell growth medium (GM) (Cell Application, Inc.), and 1% penicillin/streptomycin (Gibco). Human umbilical vein endothelial cells used in this study were limited to cells between passages 3 and 5.
Mouse embryonic fibroblasts were cultured in GM, composed of Dulbecco Modified Eagle high glucose medium (Hyclone) supplemented with 10% FBS (Gibco) and 1% penicillin/streptomycin (Gibco). The cells were grown on 0.1% (w/v) Gelatin coated dishes to 70% confluency.
Human bone marrow stromal cells were acquired from the Institute for Regenerative Medicine, Texas A&M University (Donor 8013 L). Cells were cultured in GM composed of Dulbecco Modified Eagle high glucose medium (Hyclone) supplemented with 16.5% FBS (Gibco) and 1% penicillin/streptomycin (Gibco). The cells were passaged at 70% confluence and used for experiments at passages 4 to 5.
Subcutaneous Implantation of Devices
All animal procedures were approved by the Institutional Animal Care and Use Committee of the University of California, San Diego and performed in accordance with the National Institutes of Health and national and international guidelines for laboratory animal care. Subcutaneous implantation of the cell-laden dual-compartment devices was performed 3 days after assembly and culture in vitro. Recipient mice were administered with ketamine (100 mg/kg) and xylazine (10 mg/kg), and the fur on the back was shaved. Mice were then placed on a heating pad and a 1 cm-long incision was made in the back of the mice, and 1 subcutaneous pouch was inserted by blunt dissection using a 1-cm-wide spatula on the left side of the mouse. The process was repeated on the right side. A total of 2 devices were implanted per mouse. The skin was sutured once the devices were implanted. After the surgery, mice were housed in separated cages.
Sandwich enzyme-linked immunosorbent assay (ELISA) was performed to assess the albumin production of the implanted cells as a function of time (over 28 days). In short, once a week, blood was collected from the mice via the tail vein using heparinized capillary tubes. The blood was then centrifuged at 14 900g at 4°C for 15 minutes to separate and extract the serum. The serum was assessed for human albumin by using the Human Albumin ELISA Quantitation Set (Bethyl Labs, catalogue no. E80-129) according to the manufacturer's protocol.
Acellular implants were placed in mice for 3, 7, 14, and 28 days to assess cell infiltration and vascularization over time. At each time point, the implants were retrieved and washed with PBS. Then, gross images of the implant were taken before fixing them for immunostaining. The number of visible vessels that contained blood were counted and divided by the total surface area.
Postimplantation constructs were retrieved, washed with PBS, and fixed with 4% paraformaldehyde (PFA, Sigma Aldrich) at 4°C overnight. Samples were then incubated in optimum cutting temperature (Tissue-Tek O.C.T. Compound; Sakura, Torrance, CA) at 4°C overnight on a rocker. Samples were transferred to a mold and frozen with 2-methylbutane and liquid nitrogen, and stored at −80°C until sectioning. For cryosectioning, frozen tissue blocks were sectioned with a cryotome cryostat (at −20°C) to 20-μm thicknesses. For immunofluorescent staining, sections were treated with 20 μg/mL proteinase K, permeabilized with 0.5% Triton X-100 (4 minutes, RT), treated with NABH4 (30 minutes, RT), and blocked with 3% (w/v) bovine serum albumin (BSA; 60 minutes, RT) in PBS. Sections were stained for either CD31 (platelet endothelial cell adhesion molecule; 1:100; Santa Cruz) or fluorescein isothiocyanate conjugated human albumin (1:100; Bethyl Labs) or cytokeratin-18 (R&D Systems) overnight at 4°C. An appropriate secondary antibody Alexa Fluor 488 (1:200; Thermo Fisher) along with Hoechst 33342 (2 μg/mL; Thermo Fisher) was used to bind primary antibodies for 1 hour at RT. Then samples were imaged with a fluorescent microscope.
All experiments were independently repeated at least twice with replicate samples as indicated in figure captions. Statistical analysis was performed using one-way analysis of variance and Tukey post hoc test for group comparisons to determine statistical significance (P < 0.05). Errors bars represent SEM. GraphPad Prism 5 software was used to determine all statistical analysis.
Development and Characterization of the Dual-compartment System
The dual-compartment device was developed as a cell-carrying device to successfully transplant primary human hepatocytes (Figure 1A and B). The dual-compartment structure consisted of a 3-mm height by 5-mm radius, interconnected macroporous hydrogel (outer compartment), with a 1.5-mm height by 3.5-mm radius hollow interior (inner compartment) for cell loading (Figure 1C). Macroporous hydrogels with interconnected pores were fabricated by PMMA templating of PEGDA or PEGDA-co-HAMA crosslinked networks. The SEM images were used to verify the presence of an interconnected macroporous network. The SEM images suggest that the gels had both larger pores, which were about 140 μm in diameter, and smaller pores, which were about 85 μm in diameter (Figure 2A) with an average pore size of 120 μm. We also determined the swelling ratio of the macroporous hydrogels (PEGDA/HAMA and PEGDA) (Figure 2B). The PEGDA/HAMA macroporous hydrogels had a higher swelling ratio compared to PEGDA-alone structures. Similarly, PEGDA/HAMA structures swelled and equilibrated faster compared with PEGDA-alone structures.
Porous PEGDA/HAMA Facilitates Vascular Formation
To examine the ability of the macroporous structures to promote vascularization, both PEGDA and PEGDA/HAMA structures were implanted in vivo without the presence of any exogenous cells. In addition to the appearance, we used immunofluorescent staining for 4′,6-diamidino-2-phenylindole (DAPI) (stains nucleus) and CD31 (a vascular marker) to assess host cell infiltration and vascular formation as a function of postimplantation time (3-28 days). Figure 3 shows the representative whole-mount images of the excised implants and the staining results. Analyses of the implants after 3 days of subcutaneous implantation showed minimal to no vascularization. Although the sections were positive for DAPI staining, indicating that host cells had infiltrated the implant, no positive CD31 was observed. At the next experimental time point, day 7, the presence of vascular structures could be clearly seen in the PEGDA/HAMA constructs. The constructs appeared pink with clearly visible vascular structures filled with blood. The presence of vascular structures was further confirmed by the positive CD31 staining. As the days increased, increased vessel formation (Figure 3 A) as well as more CD31 positive cells was observed (Figure 3 B). By day 14, larger blood vessels filled with blood were observed, whereas multiple smaller vessels filled with blood were seen at day 28. Interestingly, no such vascularization was observed with PEGDA constructs (Figure S1,https://links.lww.com/TP/B593). The PEGDA constructs appeared as transparent as before implantation.
Effect of Supporting Cells on Function In Vivo
Because the supporting cells could play a key role in maintaining hepatocyte function,14,27 we have compared the effect of (i) MEFs plus HUVECs and (ii) hMSC plus HUVECs on the functionality of hepatocytes, based on albumin secretions. At 1 week posttransplantation, both MEFs and hMSCs-supported implants showed similar levels of albumin secretion, which was only slightly higher compared with the HUVECs only group (Figures 4 A and B). All groups showed significantly higher albumin productions compared with week 1. Among the different groups, implants loaded with MEFs or hMSCs produced more albumin than the HUVECs only group. Between the hMSCs and MEFs, the group containing hMSCs produced more albumin than the group containing MEFs. Even though there was no statistically significant difference between the albumin secretions among the week 3 hMSCs plus HUVECs group and the MEFs plus HUVECs, all subsequent experiments were carried out using a combination of HUVECs and hMSCs as the supporting cells. In addition to presence of human albumin in the host peripheral blood, we also stained the excised implants and they were positive for human-specific albumin (Figure 4 A).
Donor-independent Function of Dual-compartment System In Vivo
After characterizing and developing the dual-compartment system, we wanted to ensure that the device could support cells from multiple donors. To this end, we used the dual-compartment system to transplant cells from 2 different donors (Table S1,https://links.lww.com/TP/B593). The implant function was assessed for functionality via albumin secretions over a 1-month period postimplantation (Figure 5 and 6). Albumin analysis of host serum indicated the presence of human-specific albumin in the circulation of the host at day 7 postsubcutaneous implantation. The amount of albumin increased from day 7 to day 15 and remained more or less stable for the rest of the month (Figure 5B). On day 28, the implants were retrieved and analyzed. The gross picture of both the implants indicated vascular formation (Figure 6B column 1). The DAPI and albumin staining of the implant (Figure 5A and 6B column 2) showed that in all implants, the albumin staining was concentrated within the inner compartment. The cells in the inner compartment of the implant were also positive for cytokeratin-18, another hepatocyte-specific marker (Figure 6B column 3). The implants were also positive for CD31 staining, which was used to identify vascular cells (Figure 6B column 4). The staining results corroborated the presence of vascular networks observed earlier by us in the whole-mount images.
This study describes the application of a dual-compartment device, containing an outer vascularizable layer and an inner cell-loading compartment, for hepatocyte transplantation. The hollow core structure of the inner compartment enables loading of a large number of cells. At the current dimensions, the inner compartment can be easily loaded with up to 20 million hepatocytes (largest number tested for loading, Figure S2,https://links.lww.com/TP/B593). Furthermore, our results demonstrated that the vascularization of the implant in the subcutaneous space supported donor cell function for 1 month.
The design of the dual-compartment system, especially the hydrogel composition and architecture played an integral role in host cell infiltration, and device vascularization. Previously, we have shown that mineralized macroporous hydrogels promote host cell infiltration.23,28 The findings that the PEGDA macroporous hydrogels were not able to promote vascularization in vivo suggest that pore architecture alone is not sufficient to facilitate vascular formation. The addition of HA to the PEGDA allowed cell infiltration and implant vascularization. This could be due to various reasons, such as the biological functions of HA and its ability to interact with cell surface receptor CD44.29,30 Furthermore, the addition of HA could have promoted vascular formation, as studies have shown that HA fragments exhibit proangiogenic effects.29,31,32 While vascularization of the implant enables long-term maintenance of donor cells, the faster swelling kinetics of the macroporous dual-compartment device could be playing an important role in maintaining the viability of the donor cells in the initial days of the implantation (ie, before the implant was vascularized) through enhanced nutrient transport to the cells. Because these implants are large and thick, to avoid necrotic cores, fast swelling is important to ensure absorption and transport of nutrients throughout the structure.
Although the liver has many functions, here we chose to use human serum albumin secretions to characterize the function of the implants. This allowed us to monitor function of the same implant, with minimal interference, over multiple time points. For our proof-of-concept study, ELISA analysis of albumin secretions for both donors followed the trend of rising after day 7 and then remaining stable for the rest of the experiment duration. Based on albumin synthesis and secretions, there were no statistically significant differences between the cells from different donors.
Although this proof-of-concept study used primary hepatocytes, the dual-compartment system could be extended toward other cells. The modular assembly of the device allows parallel optimization of the outer and inner compartments to increase overall function of the device. Tuning pore size or material composition to accelerate host cell infiltration and vascularization can optimize the outer compartment. Similarly, the dimensions of the inner compartment could be increased to improve the number of donor cells that can be housed, while maintaining the overall outer dimensions. Increasing the dimensions of the inner compartment without changing the overall dimensions of the device involves decreasing the thickness of the outer compartment, which could improve diffusion of nutrients to the cells in the inner compartments. Further studies are needed to determine the upper cell-loading limit of these devices without compromising their viability or function. It is estimated that a delivery of 1 to 10 billion functioning cells is needed to achieve therapeutic effects in lieu of solid organ transplantation.33 Even with the vascularization, housing such a large number of cells within a single device could be challenging. However, because the dimensions of the device can be tuned, it is possible to increase the dimension of the inner compartment to accommodate more cells. If a diffusion limitation is reached leading to death and compromised function of the transplanted cells, then multiple devices could be transplanted. Another key parameter that determines successful cell transplantation is the longevity of the donor cells. The diminished function of the transplanted cells with time is often thought to be associated with lack of vascularization. Studies have shown that cell transplantation approaches that incorporated vascularization resulted in improvements to the viability and function of the transplanted cells. The results described in this study show the viability and function of the transplanted cells for a month. Though the vascularization of the implant suggests the possibility of survival and function of the transplanted cells beyond a month, additional studies are needed to assess the potential of the device to support long-term viability and function of the transplanted cells. Nonetheless, the dual-compartment system described in this study offers a promising tool for cell transplantation. Furthermore, its function can be easily extended for applications in drug screening, personalized medicine, and as a platform to screen for key components (eg, ECM composition, stiffness) of the microenvironment that are necessary to maintain long-term function of donor cells. It can also be used as an in vivo experimental tool to study how donor phenotype can affect transplantation success.
In conclusion, developing successful biomaterial devices for transplantable liver cell therapies requires an approach that can transplant large numbers of cells and be integrated with the host to facilitate formation of the functional vasculature needed to maintain the viability and function of transplanted cells. To meet these criteria, we have utilized a dual-compartment biomaterial device for the transplantation of human primary hepatocytes. This device enabled minimally invasive (subcutaneous implant) cell transplantation and maintained the function of transplanted hepatocytes for at least 1 month. The dual-compartment device described here is robust and scalable. Furthermore, the modular assembly of the device can be used as a tool to create and optimize scalable vascularized 3-dimensional liver tissues for extended applications in creating humanized tissue models for investigating disease pathology, drug testing, and personalized medicine.
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