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To achieve the task of fabricating functional tissues, scaffold materials that can be sufficiently vascularized to mimic functionality and complexity of native tissues are yet to be developed. Here, we report development of synthetic, biomimetic hydrogels that allow the rapid formation of a stable and mature vascular network both in vitro and in vivo. Hydrogels were fabricated with integrin binding sites and protease-sensitive substrates to mimic the natural provisional extracellular matrices, and endothelial cells cultured in these hydrogels organized into stable, intricate network of capillary-like structures. The resulting structures were further stabilized by recruitment of mesenchymal progenitor cells that differentiated into smooth muscle cell lineage and deposited collagen IV and laminin in vitro. In addition, hydrogels transplanted into mouse cornea were infiltrated with host vasculature, resulting in extensive vascularization with functional blood vessels. These results indicate that these hydrogels may be useful for applications in basic biological research, tissue engineering, and regenerative medicine.
Most successes in tissue engineering have been limited to thin or avascular tissues such as skin, bladder, and cartilage as these constructs are relatively simple in design without requirement for intricate blood vessels [1-4]. Although there have been recent advances in vascularization of tissue constructs in vivo , development of complex tissues or organs such as heart, kidney, liver and lung has been elusive due to lack of proper formation of vasculature in the engineered constructs. Thus, the most impending challenge in creating more complex and clinically relevant tissues is vascularization of engineered tissues.
The process of angiogenesis is achieved by complex interactions among endothelial cells (ECs), the interstitial extracellular matrix (ECM), and the neighboring mural cell types via various growth factors [6, 7]. In the initial phase of angiogenesis, vascular endothelial growth factor (VEGF) activates ECs from their normal quiescent states. When activated, the ECs proliferate and secrete various proteases, including matrix metalloproteinases (MMPs), to degrade the basement membrane and ECM. ECs migrate through and extend sprouts to build tubular structures. These nascent vessels are stabilized by recruitment of mural cells such mesenchymal stem cells (MSCs), which differentiate into pericytes and deposit new ECM proteins form basal lamina [8, 9].
As these multi-components are all essential part of neovascularization, the design of pro-angiogenic tissue constructs need to address each component in order to truly mimic the physiological microenvironment in which ECs can form functional blood vessels. The objectives of the present study were to integrate cellular, biochemical, and biophysical cues in synthetic biomaterials to achieve extensive vascularization both in vitro and in vivo. To achieve this, hydrogels mimicking natural provisional ECM were synthesized and assessed as a scaffold for angiogenesis.
To fabricate synthetic ECM-mimicking biomaterials, we used protease-sensitive PEG hydrogels, first reported by West and Hubbell . Protease-sensitive peptides were introduced into the backbone of PEG to render PEG-hydrogels biodegradable in response to cellular proteases. These types of materials can also be modified with cell adhesive sequences for use as scaffolds in regenerative medicine [11-16]. In the present study, we examined proteolytically-degradable and cell-adhesive PEG hydrogels as provisional matrices for angiogenesis both in vitro and in vivo. To form and stabilize new blood vessels in hydrogels, we exploited cellular interactions between ECs and MSCs; co-culture of ECs and MSCs in hydrogels led to formation of extensive tubule-like structures that were stabilized, not subject to regression, for long term culture in vitro. In addition, we demonstrate vascularization of these hydrogels with functional blood vessels in vivo.
Human umbilical vein endothelial cells (HUVECs) were obtained from Cambrex (East Rutherford, NJ). The cells were grown on endothelial cell medium EGM-2 (Cambrex) supplemented with 2 mM L-glutamine, 1000 U/mL penicillin, and 100 mg/L streptomycin (Sigma, St. Louis, MO), and they were used through passage 8. 10T1/2 cells (American Type Culture Collection, Rockville, MD) were grown and maintained in DMEM supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 1000 U/mL penicillin, and 100 mg/L streptomycin, and they were used through passage 18. All cells were incubated at 37 °C in a 5% CO2 environment.
A MMP-sensitive peptide sequence, GGGPQG↓IWGQGK was synthesized by solid phase peptide synthesis based on standard Fmoc chemistry using an Apex396 peptide synthesizer (Aapptec, Louisville, KY, USA). Hydrogels with this peptide in the polymer backbone have been shown to be completely degraded by matrix metalloproteinases (MMP) . Following purification, synthesis of the peptide was confirmed with matrix-assisted laser desorption ionization time of flight mass spectrometry (MALDI-ToF; Bruker Daltonics, Billerica, MA, USA)
Synthesis of the ABA block copolymers with the peptide linker was completed as described previously . Briefly, the MMP-sensitive peptide sequence was reacted with acrylate-PEG-N-hydroxysuccinimide (acrylate-PEG-NHS, 3400 Da; Nektar) in a 1:2 (peptide: PEG) molar ratio in 50 mM sodium bicarbonate buffer (pH 8.5) for 2 h. This step conjugates a PEG-monoacrylate chain to the N-terminus and to the amine group on the lysine at the C-terminus of the peptide. The resulting product was then dialyzed (MWCO 5,000; Spectrum Laboratories, Inc., Rancho Dominguez, CA, USA) to remove unreacted peptide and PEG moieties, and the product was lyophilized, frozen, and stored under argon at - 80 °C. The cell adhesive ligand, Arg-Gly-Asp-Ser, (RGDS, American Peptide, Sunnyvale, CA) was conjugated to acrylate-PEG-NHS in 1:1 molar ratio under the similar conditions. Fluorophore-tagged PEG-RGDS was synthesized by reacting PEG-RGDS with Alexa Fluor-488 carboxylic acid (Invitrogen) in 10-fold dye molar excess in DMF. VEGF165 (Sigma) was conjugated to acrylate-PEG-SMC in 200:1 molar ratio at pH 8.5 at 4 °C for 4 days, followed by final 4 hours at 24 °C. The resulting PEG-VEGF solution was then lyophilized and reconstituted in HEPES buffered saline (HBS) with 0.1% BSA at 4°C until use. A gel permeation chromatography system equipped with UV-vis and evaporative light scattering detectors (Polymer Laboratories, Amherst, MA) was used to analyze the products.
The hydrogel precursor solution was prepared in 10 mM HBS (pH 7.4) with PEG-RGDS (3.5 μmol/mL), a photoinitiator, Irgacure-2969 (0.3 mg/ml; Ciba Corporations, Basel, Switzerland), and varying amount of MMP-sensitive PEG-GGGPQGIWGQGK-PEG. In this work, MMP-sensitive PEG precursor was added to the final concentrations of 7.5 mg/ml, 10 mg/ml, 12.5 mg/ml, and 15 mg/ml to fabricate hydrogels with polymer weight percentages of 7.5%, 10%, 12.5%, and 15%, respectively. For HUVECs only culture, 6×106 cells were added to the hydrogel precursor solution. For HUVEC and 10T1/2 co-culture, 4.8×106 HUVECs and 1.2×106 10T1/2 cells were added. On coverglasses that had been cleaned with 70% ethanol and sterilized under UV lamp over night, the PEG precursor-cell suspension was dispensed as 5 μl droplets and photopolymerized by exposure to long-wavelength UV light (365 nm, 10 mW/cm2) for 7 min. The hydrogels were then immersed with EGM-2 media and incubated at 37 °C in a humidified atmosphere containing 5% CO2. For PEG hydrogels used in corneal angiogenesis assay in vivo, VEGF was mixed into the polymer solution to create concentrations of 512 ng VEGF per gel and 1.9 ng PEG-VEGF per gel.
The swelling ratio and water content of the hydrogels were then determined as described previously . The compressive moduli of the different hydrogels with and without cells were determined using a 10 N load cell at a crosshead speed of 0.5 mm/min in an Instron-3342 (Canton, MA) mechanical tester as previously described . Degradation rates of MMP-sensitive PEG hydrogels formulated with varying polymer weight percentages were measured by monitoring the release of tryptophan (W) in the MMP-sensitive peptide sequence with a UV-Vis spectrophotometer (Carey 5000 Varian, Walnut Creek, CA) at 280 nm.
Endothelial tubule formation was evaluated by measuring total tubule length formed and the number of branching points over 6 days. On day 1, 3, and 6, cells in the hydrogels were stained with 2 μM calcein AM and visualized with a confocal microscopy (Zeiss LIVE 5, Carl Zeiss, Thornwood, NY) with z stack depth ranging 100-200 m. Scion Image (Scion Corporation, Frederick, MD) was used to trace and measure the total tubule length and the number of branching points in each area. The data were normalized to the z stack depth in each area to account for the different optical slice volumes of the images.
To record time-lapse images during endothelial tubule formation, HUVECs and 10T1/2 were pre-labeled with 50 μg CellTracker Green CMFDA (5-chloromethylfluorescein diacetate; Invitrogen) and 50 μg CellTracker Red CMTPX (Invitrogen), respectively, per manufacturer's instructions prior to encapsulation in the hydrogels. After photoencapsulation of the cells, the hydrogels were incubated in EGM-2 media for 5 hrs before time-lapse imaging. A confocal microscope (Zeiss LIVE 5) equipped with a motorized XYZ stage and a temperature hood was used to record time-lapse images. T he temperature hood was perfused with 5% CO2 during the experiments. Confocal images were obtained at 1 hr intervals for 66 hrs, and the motorized stage allowed automatic collection of images from pre-recorded, multiple locations.
The hydrogels were prepared as 5 μl droplets each with either HUVEC monoculture (24×106 cells/ml), 10T1/2 mono-culture (6×106 cells/ml), or HUVECs and 10T1/2 co-culture (24×106 and 6×106 cells/ml, respectively). After 6 days, the cells were isolated from the bulk of hydrogels by degrading away the hydrogels with 2 mg/ml collagenase-1 solution (Sigma) for 10 min at 37 °C. Cell suspensions from four 5 μl hydrogel droplets were pooled for each cell culture condition to obtain enough cell lysate for Western blotting. In parallel experiments, cells were plated on 6 well tissue culture plates with either HUVEC mono-culture (24×104 cells/well), 10T1/2 mono-culture (6×104 cells/well), or HUVECs and 10T1/2 co-culture (24×104 and 6×104 cells/well, respectively). SDS-PAGE and Western blotting were performed following a stand protocol. The primary antibodies used for Western blotting include antibodies against smooth muscle α-actin (SM-αactin, 1:3000; ABCam, Cambridge, MA), calponin (1:500; ABCam), and caldesmon (1:500; Sigma). The secondary antibody was used at 1:2000, 1:1000, and 1:1000 dilutions, respectively.
Immunofluorescence staining was performed to confirm differentiation of 10T1/2 cells into SMC lineages and to visualize deposition of ECM proteins adjacent to the tubular structures. On day 1 and 6, the hydrogels containing HUVEC and 10T1/2 co-culture were fixed, permeabilized, and stained with primary and secondary antibodies. The primary antibodies used were mouse anti-SMα-actin (Sigma), rabbit anti-CD31 (Bethyl), rabbit anti-collagen-type IV (ABCam), and rabbit anti-laminin (Sigma) IgGs. The secondary antibodies were anti-rabbit and anti-mouse antibodies conjugated with either Alexa flour 488, Alexa fluor 594, or rhodamine (Invitrogen). For some samples, actin cytoskeletons and nuclei were stained with TRITC-conjugated phalloidin (5 U/ml, Sigma) for 1 h and DAPI (300 nM, Invitrogen) for 5 min.
Using a modified version of the previously described corneal micropocket angiogenesis assay , hydrogels were implanted into mouse cornea in Flk1-myrmCherry transgenic mouse, which exhibits EC-specific expression of a myristoylated mCherry fluorescence protein in EC membrane . Briefly, mice were anesthetized, and a partial thickness incision was made into the mouse cornea. The micropocket was created using a von Graef knife, and hydrogels were implanted into the micropocket immediately after UV photopolymerization. Seven days post-implantation, some mice were injected intravenously with dextran-Texas red (MW 70KDa), and the mice were sacrificed with CO2 asphyxiation. Eyes from the mice were enucleated and fixed in 4% paraformaldehyde for one hour at 4°C. Corneal flat-mount preparations were made, and imaging was performed using a Zeiss LSM 510 META inverted microscope system (Carl Zeiss Inc) equipped with a Zeiss Plan-Apochromat 20×/0.75 NA objective. 543-nm and 488-nm lasers were used to excite the Flk1-myrmCherry and fluorophore-tagged PEG-RGDS, respectively.
Statistical analysis was performed with Jmp 5.1 (SAS Institute Inc, Cary, NC). Data sets were analyzed using two-way analysis of variance (ANOVA), followed by Tukey's HSD test for multiple comparisons. P-values less than 0.05 were considered statistically significant. All values are reported as mean ± standard deviation.
To fabricate synthetic ECM-mimicking hydrogels used in the current studies, we incorporated an MMP-sensitive sequence (GGGPQG↓IWGQGK) into the backbone of PEG block polymers with acrylate terminal groups, which allow crosslinking of precursors into networks [10, 11, 21] (Figure 1A). GGGPQG↓IWGQGK is a mutated version of α1(I) collagen chain for increased degradation kinetics with various MMPs [11, 22]. In addition, a ligand for integrin, RGDS, was grafted onto PEG chains to support integrin-mediated cellular adhesion and migration while VEGF was incorporated into hydrogels to promote robust angiogenesis [12, 23].
We first characterized protease-mediated degradation of hydrogels. Degradation of the hydrogels was monitored by measuring release of tryptophan from the MMP-sensitive sequences in hydrogels incubated with collagenase, a subset of MMP family of proteases (Figure 1B). Hydrogels with 10% polymer weight percentage incubated with collagenase solution underwent complete degradation within 1.5 hrs, but not with either plasmin or the buffer solution even after 21 hrs, demonstrating specificity of degradation mediated by MMP family of proteases. Susceptibility of hydrogels to proteolysis was correlated with their polymer weight percentage as denser networks took progressively longer time to degrade (Figure 1C). As the polymer weight percentage was increased from 7.5% to 15%, the compressive modulus of the hydrogels was increased from 30.2 ± 6.05 kPa to 110 ± 23.8 kPa (Figure 1G). Encapsulation of cells did not alter the compressive moduli of hydrogels. The swelling ratios for the 7.5, 10, 12.5, and 15% hydrogels were 27.9 ± 8.37, 17.0 ± 2.59, 14.7 ± 3.11, and 13.2 ± 2.33, and their water contents were 96.2 ± 1.11, 94.0 ± 0.903, 92.9 ± 1.47, and 92.3 ± 1.28 %, respectively.
To form and stabilize new blood vessels in hydrogels, we exploited heterotypic cell interactions between ECs and mesenchymal progenitor cells. ECs are able to form primitive tubule structures when cultured alone; however, for the rudimentary blood vessels to become matured and stabilized, they require interaction with mural cell types. 10T1/2 cells, a subset of multipotent mesenchymal progenitor cells, are known to support endothelial tubule formation in naturally occurring ECM-derived materials [24-26].
MMP-sensitive PEG hydrogel precursors were formulated with human umbilical vein endothelial cells (HUVECs) and 10T1/2 cells (30 million cell/ml with HUVEC:10T1/2 ratio of 4:1) and exposed to long wavelength UV to photopolymerize the precursors into hydrogels. The cells encapsulated in hydrogels were cultured, and examined for their angiogenic responses at various time points (Figure 1D, E, F). In order to find the optimal mechanical composition of the hydrogels for the angiogenic responses, tube formation by HUVECs and 10T1/2 cells were quantified in hydrogels with 7.5% to 15% polymer weight percentage. In hydrogels with an intermediate polymer weight percentage of 10%, HUVECs and 10T1/2 cells formed primitive tubule-like structures by day 3 and continued to mature into extensive interconnections throughout the 3D network of hydrogels over 6 days. In soft, compliant hydrogels with 7.5% polymer weight, the cells underwent self-assembly to form thick interconnections, but the resulting structures rapidly regressed as the gels were completely degraded within 6 days. In stiffer hydrogels with 12.5 or 15% polymer weight, the degree of tubule formation was reduced as cells failed to migrate as much in softer matrices, and the cell clusters remained short and rudimentary. In contrast, HUVECs and 10T1/2 cells co-cultured on tissue culture plates for 14 days did not form any tubule-like structures but remained as monolayers. The varying degrees of angiogenic responses in these hydrogels did not seem to result from different cellular viability as more than 80% of cells remained viable after 6 days in all hydrogel formulations tested, which is within a typical range reported in other literatures using hydrogel systems (Supplement Figure 1) [27, 28]. These results show that angiogenesis organized and promoted by HUVECs and 10T1/2 cells are well recapitulated in proteolytically-degradable hydrogels and that there is a particular formulation of hydrogels with an intermediate rigidity that maximizes these angiogenic responses.
These results do not demonstrate mechanical properties of hydrogels as a sole determinant of angiogenic potential since other parameters such as density of cell adhesive ligand or degradable sites were not held constant. However, since 10% polymer weight PEG hydrogels produced robust angiogenic responses, all the subsequent studies were performed with this particular formulation of hydrogels unless noted otherwise.
To better understand the dynamics of angiogenesis in the hydrogels, the process of tubule formation was captured by time-lapse confocal microscopy. In hydrogels with HUVECs in mono-culture condition (30 million cells/ml), the cells started migrating toward each other to form large clusters (t = 5 hr) and organized into primitive tubule-like structures (t = 21 hr) (Figure 2A, C, and Supporting Video 1). However, at later time points (t = 53 and 69 hr), the structures lost their tubular morphology and rapidly regressed into individual cell clusters. The total tubule length decreased over 6 day culture period, while the number of branching points stayed at minimal levels (Supplementary Figure 2).
In contrast, HUVECs and 10T1/2 cells co-cultured in the hydrogels (30 million cell/ml with HUVEC:10T1/2 ratio of 4:1) formed large clusters that rapidly organized into tubule-like structures throughout the hydrogels (t = 21 and 37 hr) (Figure 2B, and Supporting Video 2). The tubule-like structures were maintained stably even at later time points (t = 53 and 69 hr) with extensive branches and typical length reaching 700 μm (Figure 2C, and Supplementary Figure 2).
The endothelial tubular structures formed were reminiscent of capillary structures in their cellular organizations and morphology. 10T1/2 cells were frequently observed to align adjacent to and migrate rapidly along the tubular structures that were assembled by HUVECs. Figure 2D shows HUVECs undergoing lumen formation and 10T1/2 cells adjacent to the ECs supporting the process. Subsets of the HUVECs had multiple small vesicles, while others had their vesicles fused together intercellularly to form lumens enclosed by multiple ECs (Figure 2D, E), and these processes of lumen formation and perivascular association of mural cell types parallel a series of physiological events that accompany neovascularization as described recently in vivo . In addition, HUVECs and 10T1/2 cells kept in culture for 28 days maintained their extensive network of tubular structures throughout the hydrogels (Figure 2F), demonstrating their durability. The hydrogels remained intact and stable when examined up to 1 month. Taken together, these results suggest that 10T1/2 cells enhance endothelial tubule formation and maintenance in hydrogels.
We next addressed the question whether 10T1/2 MSCs differentiate toward SMC lineages in co-culture conditions with HUVECs. 10T1/2 cells are known to up-regulate expressions of smooth muscle marker proteins, such as SM-myosin, SM22α, and calponin when co-cultured together with ECs . We indeed observed dramatic up-regulation of SMC protein markers such as SM α-actin, calponin, and caldesmon in the co-culture conditions in hydrogels, but not in mono-culture conditions of either cell type as examined by Western blotting (Figure 3A). Similar results were obtained from cells cultured in tissue culture plates. Over-proliferation of either cell type contributing to this result can be ruled out since time-lapse confocal images revealed a minimal number of cell division during 72 hrs of culture as reported in the literature .
Differentiation of 10T1/2 cells into SMC lineages was further confirmed by immunofluorescence staining. There was minimal level of SM α-actin staining on day 1 in the hydrogels with co-culture conditions, while anti-CD31 staining showed HUVECs dispersed in the matrix (Figure 3B). After 6 days, anti-CD31 staining showed HUVECs in tubule-like structures, whereas the cells stained with anti-SM α-actin antibody aligned adjacent to the HUVECs (Figure 3C). Taken together, these results suggest that 10T1/2 precursor cells differentiate toward SMC lineages in co-culture conditions and associate closely with the tubule-like structures composed of HUVECs.
Collagen-type IV and laminin, both the major components of basal lamina of blood vessels [31, 32], were found to be secreted and deposited along the tubule-like structures (Figure 3D-G). This indicates that these cells actively remodel the matrices by secreting their own set of ECM proteins, which are known to serve as reservoir for various growth factors and provides structural integrity to newly formed capillaries .
Proteolytically-degradable PEG hydrogels were implanted into mouse cornea using a micropocket angiogenesis assay  and examined 7 days post-implantation (Figure 5A,B). We utilized Flk1-myrmCherry transgenic mice, which exhibit EC-specific expression of a reporter mCherry in EC membrane  and allow monitoring of host vasculature invading into the hydrogels by confocal microscopy. In order to recruit the host blood vessels into hydrogels, pro-angiogenic growth factor, VEGF, was incorporated into the hydrogels both in a soluble and an immobilized form via PEG linkage as described previously . In the absence of VEGF, there was not any significant angiogenic response (Supplement Figure 3). VEGF released from non-degradable hydrogels lacking MMP-sensitive peptide sequence promoted neovascularization underneath, but not in the hydrogels as shown by the vessels in red in the depth profile graph (Figure 4A-D). In contrast, MMP-sensitive PEG hydrogels with 10% polymer weight had significant portions of the scaffold degraded after 7 days, and robust neovascularization was observed (Figure 4E-H). The newly formed vessels infiltrated into the core of hydrogels as shown in green in the depth profile graph (Figure 4H). Dextran-Texas red (MW 70KDa) was injected intravenously in the host mice to delineate functionality of the blood vessels formed in the hydrogels. Figure 5C shows that indeed the blood vessels in the hydrogels were functional and perfused with the host's circulatory system. More rigid hydrogels with 15% polymer weight showed the initial stages of vessel penetration into the hydrogels (Figure 4L arrow); however, cells failed to invade into the core of hydrogels (Figure 4I-L). These data in corroboration with the in vitro results indicate that degradation of hydrogels and the ensuing formation of functional blood vessels can be controlled by provision of appropriate biophysical and biochemical cues in the hydrogels.
This work aimed to combine cellular, biochemical, and biomechanical cues to promote neovascularization in synthetic biomaterials. By providing appropriate cellular and molecular components in microenvironment that mimics physiological landscape of angiogenesis, we have been able to recapitulate and promote blood vessel formation both in vitro and in vivo in completely synthetic biomaterials. These results suggest applications of these systems for angiogenesis and anti-angiogenesis studies including its development as a diagnostic platform for screening angiogenic and anti-angiogenic compounds as well as clinical applications in tissue engineering and regenerative medicine.
HUVECs were stained with CellTracker green prior to encapsulation into the hydrogels as indicated in the Materials and Methods section and imaged over 70 hrs. Scale bars = 50 μm.
HUVECs and 10T1/2 cells were stained with CellTracker green and red, respectively, prior to encapsulation into the hydrogels as indicated in the Materials and Methods section and imaged over 70 hrs. Scale bars = 50 μm.
Hydrogel formulations with varying polymer weight percentage did not have any significant effect on cellular viability after 6 days in culture. Data represent mean ± SD (n = 11).
Hydrogels seeded with either A) HUVECs alone or B) HUVECs and 10T1/2 cells were cultured, and the resulting tubule-like structures were visualized by calcein AM staining over 6 days. Tubule-like structures formed by HUVECs in mono culture condition regressed by day 6, whereas there was more robust tubule formation by HUVECs co-cultured with 10T1/2 cells in hydrogels. C, D) 10T1/2 cells enhanced the total tubule length formed and the number of branching points during the 6 day culture period. Data represent mean ± SD (n = 4). * P < 0.05, analyzed by two-way ANOVA followed by Tukey's HSD test. Scale bars = 100 μm.
Hydrogel formulated with non-degradable PEGDA chains without exogenous VEGF did not promote neovascularization during 7 days after implantation of hydrogels.
This work was supported by grants from NIH and NSF. The authors would like to thank Iris Kim, Melissa Scott, and Tegy Vadakkan for technical support.
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