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. Author manuscript; available in PMC: 2016 Dec 1.
Published in final edited form as: Arterioscler Thromb Vasc Biol. 2015 Oct 8;35(12):2677–2685. doi: 10.1161/ATVBAHA.115.306362

Three-dimensional vascular network assembly from diabetic patient-derived induced pluripotent stem cells

Xin Yi Chan 1, Rebecca Black 1, Kayla Dickerman 1, Joseph Federico 1, Mathieu Levesque 3,4, Jeff Mumm 3,4, Sharon Gerecht 1,2,*
PMCID: PMC4603427  NIHMSID: NIHMS726214  PMID: 26449749

Abstract

Objective

In diabetics, hyperglycemia results in deficient endothelial progenitors and cells, leading to cardiovascular complications. We aim to engineer three-dimensional (3D) vascular networks in synthetic hydrogels from type-1 diabetes (T1D) patient-derived human induced pluripotent stem cells (hiPSCs), to serve as a transformative autologous vascular therapy for diabetic patients.

Approach and Results

We validated and optimized an adherent, feeder free differentiation procedure to derive early vascular cells (EVCs) with high portions of VEcad+ cells from hiPSCs. We demonstrate similar differentiation efficiency from hiPSCs derived from healthy donor and T1D patients. T1D-hiPSC-derived VEcad+ cells can mature to functional endothelial cells (ECs) expressing mature markers: von Willebrand factor and eNOS, are capable of lectin binding and acetylated low density lipoprotein uptake, form cords in Matrigel and respond to tumor necrosis factor alpha. When embedded in engineered hyaluronic acid (HA) hydrogels, T1D-EVCs undergo morphogenesis and assemble into 3D networks. When encapsulated in a novel hypoxia-inducible (HI) hydrogel, T1D-EVCs respond to low oxygen and form 3D networks. As xenografts, T1D-EVCs incorporate into developing zebrafish vasculature.

Conclusion

Using our robust protocol, we can direct efficient differentiation of T1D-hiPSC to EVCs. Early ECs derived from T1D-hiPSC are functional when mature. T1D-EVCs self-assembled into 3D networks when embedded in HA and HI hydrogels. The capability of T1D-EVCs to assemble into 3D networks in engineered matrices and to respond to a hypoxic microenvironment is a significant advancement for autologous vascular therapy in diabetic patients and has broad importance for tissue engineering.

Keywords: induced pluripotent stem cells, Endothelial cell differentiation, Type I Diabetes, vascular therapy, 3D vascular network

Introduction

Cardiovascular diseases (CVD) are currently a leading cause of death in the United States.1 Studies have demonstrated that due to hyperglycemia, diabetes patients have a higher risk in developing CVD.2, 3 Under prolong hyperglycemia conditions, physiological changes in diabetic patients cause aberrant angiogenesis in both micro and macro vasculatures, resulting in CVD such as coronary heart disease and peripheral arterial disease.4, 5 High blood glucose also impairs endothelial progenitor cell (EPC) functionality and causes poor neovascularization in response to hypoxia, directly affecting wound healing processes in diabetic patients.6, 7 Thus, generation of vasculatures holds great promise in treating diabetic patients with CVD and wound healing complications. 8-10

Stem cells, defined by their ability to self-renew and differentiate into multiple mature cell types, are emerging as a promising source for non-invasive approaches to tissue regeneration including vascular regeneration.11 Autologous vascular stem cell therapy could contribute as a potential source for endothelial cells (ECs) to treat CVD. One such example would be endogenous EPCs. Both animal and clinical studies have demonstrated that EPCs can be isolated, expanded in vitro and transplanted to improve revascularization in ischemic CVD.12-15 Theoretically, EPCs represent as a great candidate for vascular therapy. Unfortunately, the functions of EPCs are compromised under high glucose conditions in diabetes.16-18 An alternate source of stem cells for vascular therapy is human induced pluripotent stem cells (hiPSCs). The discovery of the hiPSCs19, 20 makes stem cell therapy particularly attractive for individuals with diabetes offering not only an autologous therapy approach but also the potential of reversing the stress effects of high glucose, caused by diabetes, in reprogramed hiPSCs.21 Thus, hiPSCs may provide a renewable stem cell source for diabetic patients that could improve wound-healing outcomes.

In previous work, we established a step-wise differentiation scheme to generate a bicellular population of early vascular cells (EVCs) consisting of vascular endothelial cadherin-positive (VEcad+) cells (early ECs) and platelet-derived growth factor β-positive (PDGFRβ+) cells (early pericytes) from healthy hiPSC lines. EVCs can be further differentiated to mature ECs or pericytes. When encapsulated in a synthetic hyaluronic acid (HA) hydrogel, EVCs self-assemble to functional three-dimensional (3D) vascular networks.8 In the present study, using the same differentiation scheme, we investigated if EVCs can be derived from Type I diabetes (T1D) patient-derived hiPSCs, matured into functional ECs and assembled into vascular networks in synthetic hydrogel in response to hypoxic conditions. We examined the differentiation of T1D hiPSCs to EVCs, further increased the portion of VEcad+ cells in the EVCs and whether the T1D EVCs can be matured to functional ECs. We next determined the ability of T1D EVCs encapsulated in a synthetic HA hydrogel to undergo morphogenesis and self-organize to form 3D vascular networks. We also examined the ability of T1D EVCs to undergo morphogenesis to form 3D vascular networks in response to hypoxic conditions using a novel HI hydrogel. Finally, we demonstrated the in vivo functionality of T1D-EVCs using a zebrafish xenograft model. The current work shows that T1D-hiPSCs can differentiate into EVCs, mature into functional ECs, generate vascular networks in deliverable hydrogel as a response to hypoxia and integrate into host vasculature networks in vivo. Overall, this approach provides us a unique opportunity to advance autologous therapy for diabetic-vascular complications, especially diabetic wound healing.

Materials and Methods

Materials and Methods are available in the online-only Data Supplement.

Results

Validation and Optimization of Early Vascular Cell (EVC) differentiation from hiPSCs

In previous works, we established a two-step, adherent culture protocol for the controlled differentiation of hESCs and healthy hiPSCs into a bi-cellular EVC population.8, 22, 23 During the first six days of differentiation, seeded single-cell hiPSCs were induced to become mesodermal. Upon reseeding on day 6, they proceed to differentiate into EVCs for another six days (Figure SIA). This EVC population consists of VEcad+ early ECs and PDGFRβ+ early pericytes. With our goal to develop a therapeutic solution for diabetic-related vascular complications, we first sought to demonstrate EVC differentiation from T1D-hiPSCs and to increase the EC yield in our EVC cultures. We reasoned that higher EC yield will more closely mimic the in vivo EC to pericyte ratio in the average microvessel.24 In the optimization process, we used healthy individual-derived BC1-hiPSC, a completely sequenced hiPSC cell line derived via non-viral reprogramming techniques.8, 25, 26 It was previously shown that the addition of the Rho associated coiled coil containing kinases (ROCKs) inhibitor Y27632 improves the survival of the single-cell hiPSCs.27 Another study demonstrated that the initial cell seeding density affects the differentiation efficiency of pancreatic endocrine cells.28 Thus, as a comparison to our original culture conditions (5×104 cells/cm2; control) we supplemented the differentiation media with ROCK inhibitor Y27632 (5×104 cells/cm2 +RI) and also tested higher density culture conditions by increasing cell seeding density to 1×105 cells/cm2 (Figure SIA). On day 1, cultures seeded at a higher density, 1×105 cells/cm2, resulted in an overall increase in total cell attachment (Figure SIBi-ii). When ROCK inhibitor was added to the original culture conditions, cells attached to the plate at around 50-60% confluency (Figure SIBiii). Interestingly, ROCK inhibitor supplemented cells appeared more compact and less spindle-like, adapting a cobble stone-like morphology (Figure SIBiii). Also, addition of ROCK inhibitor increased cell survival and cell attachment of BC1-hiPSC, resulting in a ≥3-fold difference in the total number of cells attached between control and ROCK suppressed cultures (Figure SIC). By increasing cell-seeding density from 5×104 cells/cm2 to 1×105 cells/cm2, there was a 2-fold increase in total cell attachment (Figure SIC). On day 12 of differentiation, cells in culture using the original low density protocol (control) were not confluent and appeared mostly elongated and spindle-like (Figure SIDi). On the contrary, cells seeded either at a higher density or with ROCK inhibitor appeared as patches of cobblestone-like cells surrounded by cells that are spindle-like and elongated (Figure SIDii- and 1Diii, respectively). Importantly, we found an increased total yield of VEcad+ cells in the modified differentiation protocols. The total yield of VEcad+ cells for ROCK inhibitor supplemented differentiation is 41.76±17.05% (n=3) and for high initial cell seeding density is 62.0.0±14.9% (n=6) compared to our original protocol, which yielded 18.9±14.9% VEcad+ cells (n=4; Figure SIE).

EVC differentiation from T1D patient-derived hiPSC

We examined whether hiPSCs derived from patients with vascular complications have a similar differentiation potential to BC1-hiPSC derived from a healthy donor. Here, we utilized two T1D patient-derived hiPSC lines, H2.1 and 1018S, to generate a bi-cellular population of EVCs.29-31 We compared the original differentiation protocol and the improved differentiation protocols focusing on supplementation with ROCK inhibitor. Similar to the BC1-hiPSC, in comparison to cultures without ROCK inhibitor addition (Figure 1Ai), the ROCK inhibitor supplemented cells attached better and had a morphology that is more relaxed and less spindle-like on day 1 (Figure 1Aii and SIIAi). ROCK inhibition resulted in a ≥2-fold increase of the total number of attached T1D-hiPSC cells (Figure 1B). On day 12 of differentiation, compared to cells cultured using the original protocol, where most of the cells appeared to be spindle-like and elongated (Figure 1Ci), the cells cultured with the ROCK inhibitor appeared to be grouped in cobblestone-like clusters, suggestive of an endothelial-like morphology (Figure 1Cii and SIIAii). Supplementing ROCK inhibitor in T1D-hiPSC cultures resulted in a similar enrichment in endothelial differentiation, generating a higher yield of VEcad+ cells (Figure 1D and SIIB). At the end of the EVC differentiation step (day 12), we successfully generated 48.5±19.7% VEcad+ cells for H2.1 (60.9±22.4% for 1018S) and 73.21±11.25% PDGFRβ+ cells for H2.1 (64.11±18.4% for 1018S; Figure SIIIA and SIIC). In the optimized protocol we observed VEcad+ cells that are also positive for the endothelial cell marker CD31 (43.0±15.0%, Figure SIIIB). Similar to the BC1 hiPSC-derived EVCs, a small subset of T1D-derived VEcad+ cells also express PDGFRβ (Figure SIIIC). Immunofluorescence staining showed that the T1D hiPSC-derived EVCs (here after, shortened to T1D-EVCs) contain early ECs with the typical cobblestone morphology and both VEcad and CD31 localized to intercellular junctions, as well as elongated SM22α+ cells, indicating pericytes (Figure SIIID-E; SIID-E). Altogether, we conclude that the effect of ROCK inhibition in T1D-hiPC and BC1-hiPSC cultures is similar: an improvement in the total number of surviving cells and EC differentiation.

Figure 1. EVC Differentiation of T1D-hiPSCs.

Figure 1

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(A) LM images of the differentiating EVCs on day 1 in two different conditions. (i) Control: 5×104 cells/cm2 and (ii) 5×104 cells/cm2 and ROCK inhibitor supplement. Black arrowheads indicate spindle-like morphology. (B) Quantification of cells attachment after 24 hours in the two conditions examined. (C) LM images of the differentiating EVCs on day 1 in two different conditions. White arrowheads indicate cobble stone-like morphology. (i) Control: 5×104 cells/cm2 and (ii) 5×104 cells/cm2 and ROCK inhibitor supplement. (D) Representative flow cytometry histogram of VEcad and PDGFRβ expression in T1D-EVCs derived in the two conditions. Scale bars are 100 μm. Significance levels were set at *p<0.05, **p<0.01, and ***p<0.001.

EC maturation of T1D-EVCs

We next examined the endothelial maturation and functionality potential of T1D-EVCs. We speculated that VEcad+ cells derived from T1D-hiPSCs can mature into normally functional ECs. VEcad+ cells were sorted using magnetic sorting and purification of more than 90% VEcad expression of the sorted cells was confirmed using flow cytometry (Figure 2A). Sorted VEcad+ cells were subcultured and expanded for another 6 days, in which they maintained the cobblestone morphology of typical ECs (Figure 2B), compared to VEcad- cells, where most cells appeared fibroblast-like (Figure 2B). Sorted ECs exhibited typical membrane expression of VEcad and CD31 as well as cytoplasmic punctate expression of von Willebrand factor (vWF) and cytoplasmic endothelial nitric oxide synthase (eNOS) – both mature EC markers. Sorted cells demonstrated the ability to bind to lectin Ulex europaeus, uptake acetylated low density lipoprotein (acLDL; Figure 2C) and form cord when seeded on top of Matrigel (Figure 2D). In addition, T1D-EVCs responded to the inflammatory cytokine, tumor necrosis factor alpha (TNFα) by upregulating expression of intercellular adhesion molecule 1 (ICAM-1; Figure 2E). Overall, T1D-EVCs exhibit mature endothelial characteristics consistent with ECs derived from healthy BC1-hiPSC control cultures.7

Figure 2. Maturation of functional T1D-ECs from T1D-EVCs.

Figure 2

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(A) Representative flow cytometry histogram for VEcad expression of magnetic sorted VEcad+ cells. (B) LM images of sorted VEcad+ and VEcad- cells that were subcultured for an additional 6 days. (C) IF images of sorted and subcultured VEcad+ cells for: VEcad, CD31, vWF, eNOS, binding of Ulex europaeus fluorescein-conjugated lectin, and uptake of acLDL. Marker in red or green as indicated on each figure panel; nuclei in blue. (D) LM image of T1D-ECs forming cord on Matrigel. (E) T1D-ECs responded to TNFα via upregulation of ICAM-1 expression. Significance levels were set at *p<0.05, **p<0.01, and ***p<0.001.

Vascular assembly of T1D-EVCs in HA hydrogels

In order to assess the potential of T1D-EVCs to self-assemble into a microvascular bed we examined network formation in a synthetic HA-based hydrogel. These engineered HA hydrogels present adhesion and degradation motifs, allowing sequential progression in vascular morphogenesis.8, 32 We tracked the process of T1D-EVCs network formation every day for a total of 3 days. During the first two days after encapsulation of T1D-EVCs in HA-based hydrogels, we observed vacuole formation in many of the cells, presumably early ECs (Figure SIVA). In some cases, larger lumen formed when multiple vacuoles merged and coalesced into a larger structure (Figure SIVAiii-iv). Within the same time frame, we also observed tubulogenesis, in which many of the cells have sprouted. On day 3, an extensive, multicellular network was observed within the hydrogel, which appears to be a complex vascular network (Figure SIVB-C). Lumens were detected within the 3D vascular networks (Figure SIVD). By tracking the progression of 3D network formation of encapsulated T1D-EVCs, we observed cellular behavior and network progression that resembles typical vascular morphogenesis.33 Overall, we demonstrated that T1D-EVCs are able to self-assemble into vascular networks in a synthetic hydrogel.

Three-dimensional T1D-EVC network formation in HI hydrogel

Since EPC-mediated blood vessel formation in diabetic patients is deficient in response to hypoxia,34 we sought to determine if T1D-EVCs retain similar deficits. We employed a novel HI hydrogel system developed in our laboratory35 to encapsulate T1D-EVCs and examined their responsiveness to a hypoxic microenvironment, similar to diabetic wounds. Our positive control was healthy BC1-EVCs.8, 36 EVC derivatives were encapsulated in either non-hypoxic or hypoxic hydrogel (see Materials and Methods), and the process of morphogenesis and network formation was tracked every 24 hours for a total of 3 days. During the encapsulation period, we recorded dissolved oxygen (DO) levels of EVCs encapsulated in the hypoxic and non-hypoxic hydrogel at a range of 0.1 to 5.0% and 10 to 15%, respectively for the first 24 hours (Figure SVA-B). We found that in the hypoxic hydrogel, EVCs progressed through all stages of vascular morphogenesis, starting from the formation of individual vacuoles that coalesce to form lumens in the first day (Figure SVIAi-iii and SVC) and sprouting within the second day of encapsulation (Figure SVIAiv and SVD). On day 3, the encapsulated EVCs eventually formed extensive 3D networks that are lumenized, indicating complete tube formation (Figure SVIB-D). In comparison, within the first two days of non-hypoxic hydrogel encapsulation, EVCs formed vacuoles and lumens but did not sprout (Figure SVE-F). On day 3, T1D-EVCs encapsulated in non-hypoxic hydrogel had limited tube formation (Figure SVIE and F). Overall, we observed more complex 3D T1D-EVC network assembly in hypoxic hydrogel compared to non-hypoxic hydrogel, a finding that is consistent with BC1-EVCs (Figure SVII).

Three-dimensional analysis of T1D-EVC hypoxic networks

Vascular network quantification is typically performed on 2-dimensional projections of 3-dimensional confocal z-stacks, limiting information along the z-axis. To have a more comprehensive analysis of the 3D T1D-EVC hypoxic networks formed in hydrogels, we employed 3D image analysis. This approach enabled quantification of the differences between vascular networks formed within non-hypoxic and hypoxic encapsulated control BC1-EVCs (Figure SVII) and T1D-EVCs (Figure 3 and Video SI). Based on the analysis, we observed significant differences between the non-hypoxic and hypoxic hydrogel encapsulated T1D-EVCs, especially along the z-axis. At the same cell encapsulation density and after 3 days in culture, the total combined tube length of the hypoxic hydrogel encapsulated T1D-EVC network is four times longer than its non-hypoxic counterpart (Figure 3C), a slightly larger difference when compared to the control-EVCs (Figure SVIIC). In control-EVC hypoxic hydrogels, the length of tubes ranges from 10 μm to 3600 μm, compared to 10 μm to 900 μm in non-hypoxic condition (Figure SVIID). In T1D-EVC hypoxic hydrogel constructs, the individual continuous tube length ranges from 10 μm to 3000 μm, in comparison to 10 μm to 200 μm in the non-hypoxic hydrogel constructs (Figure 3D). Also, the mean tube thickness of T1D-EVC non-hypoxic hydrogel construct is approximately 1 μm thicker in diameter than the T1D-EVC hypoxic hydrogel construct (Figure 3E), similar to the case of BC1-EVCs (Figure SVIIE). A unique aspect of 3D analysis is that it provides information of the T1D-EVC network in the Z-direction. One astounding effect of hypoxic hydrogels on T1D-EVCs tubulogenesis and network formation is that the network structures span a much larger distance along the Z-axis, approximately six times larger than those in the non-hypoxic hydrogel (Figure 3F). Surprisingly, the BC1-EVC networks in hypoxic hydrogel spanned an average Z-distance that is only slightly larger than the network in the non-hypoxic hydrogel (Figure SVIIF). The total volume covered by T1D-EVC networks in hypoxic hydrogels is 2.5×106 μm3, twice the volume of T1D-EVC networks in non-hypoxic hydrogel (Figure 3G-H), and is consistent with BC1-EVC networks (Figure SVIIG-H). This indicates that T1D-EVCs are responding to low DO levels, which plays a role in stimulating the migratory behavior and ability to form networks in hypoxic hydrogel, similar to the control cells. In non-hypoxic hydrogels, most T1D-EVC tubes form along the X-Y axis, instead of the Z-axis, indicating a lack of stimulation and signaling for migratory behaviors of EVCs.

Figure 3. Three-dimensional analysis of T1D-EVC networks in both non-hypoxic and hypoxic hydrogel.

Figure 3

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Representative day 3 confocal z-stack of (A) non-hypoxic and (B) hypoxic hydrogel encapsulated T1D-EVCs analyzed with Imaris Filament Tracer. Green: Phalloidin. Red lines: T1D-EVC tubes/network that are traced and analyzed. Yellow lines: an example of a continuous T1D-EVC network in the hypoxic hydrogel. Scale bars are 100 μm. Multiple aspects of the analysis based on non-hypoxic and hypoxic hydrogel confocal z-stacks are compared and presented on bar graphs or scatter plots: (C) Total tube length; (D) Mean tube length; (E) Mean tube thickness; (F) Mean Z-distance covered in hydrogel; (G) Total tube volume; and (H) Mean tube volume. Significance levels were set at *p<0.05, **p<0.01, and ***p<0.001.

In vivo functionality of T1D-EVCs in zebrafish

To further assess the functionality of T1D-EVCs we utilized a zebrafish xenograft model.37 T1D-EVCs were injected into 2-day post fertilization (2 dpf) embryos via the duct of Cuvier and their integration with the developing host vasculature was determined. The percentage of vessel incorporation was examined and analyzed at 5-day post-injection using Imaris (See Materials and Methods; Figure SVIII). On average, 50.95±12.23% of the injected T1D-EVCs are incorporated into the developing zebrafish vasculature (Figure 4A-B). We found 71.88% of the incorporation located in the trunk of the embryo and 28.12% in the head region (Figure 4C). These observations demonstrate in vivo functionality of T1D-EVCs, by incorporating into the vessels of the developing zebrafish.

Figure 4. In vivo functionality of T1D-ECs.

Figure 4

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(A) Representative zebrafish embryo showing the integration of T1D-ECs (in red) into the zebrafish host vasculature (in green) 5-day post injection. (B) High-magnification images of (A). T1D-ECs (red) and host vasculature (green). (C) Percentage of T1D-EVCs incorporation into whole zebrafish embryos (n=12) using Imaris analysis (red bar). Percentage of incorporation in the head region (yellow bar) and trunk region (yellow checkered bar).

Discussion

Diabetes patients suffer from serious chronic health complications as a result of prolonged exposure to high blood glucose levels. Since most of these complications are related to dysfunction of ECs and EPCs in these patients, an ideal solution would be an alternative source of adult stem cells (other than the EPCs) for potential autologous vascular therapy. Studies have shown that co-culturing ECs with murine pericytes or human mesenchymal stem cells is important in sustaining engineered blood vessel network in vivo.38-41 In this study, we successfully described the first example of cultured EVC differentiation, consisting of VEcad+ cells (early ECs) and PDGFRβ+ cells (early pericytes), from T1D patient-derived hiPSCs.

In an attempt to increase the number of VEcad+ cells in EVC cultures, we discovered beneficial effects from increasing initial cell seeding density and of ROCK inhibition from both healthy and diabetic hiPSCs. Cell seeding density has been an important factor in stem cell cultures regarding maintaining cells in a differentiated state,42 or in differentiating adult stem cells or iPSCs into a specific cell type.28, 43 For example, it has been shown that the differentiation propensity of hESCs into pancreatic endocrine cells is favored at a high initial seeding density of 5.3 × 104 cells/cm2, which reached 100% confluency 24-hour after seeding.28 This seems to be the case for our EVC differentiation protocol where increased initial seeding density from 5×104 cells/cm2 to 1×105 cells/cm2 provided us with a significant increase of early EC yield. Similar observations apply to our differentiation culture supplemented with a ROCK inhibitor. ROCK inhibition improved the cell survival and cell attachment during seeding,27, 44 in turn, creating an effect that is similar to increasing the initial seeding density by providing a better EC yield during differentiation. It is also plausible that ROCK inhibition plays a role beyond simply increasing plating efficiency and cell survival in culture. For instance, phosphatase and tensin homolog (PTEN) is activated by the Rho/ROCK pathway, which leads to inhibition of the protein kinase B (also known as Akt) pathway which regulates cell proliferation, specifically EC proliferation.45-49 On the other hand, activation of myosin light chain by Rho/ROCK pathway is known to promote myogenic differentiation, including pericyte differentiation.45, 50, 51 Thus, it is possible that suppression of ROCK in our differentiation culture plays a role in inducing EC fates. Ongoing studies are being performed to dissect the underlying mechanism of the ROCK pathway in EC differentiation before drawing any conclusion. Importantly, we found that both T1D-hiPSC lines examined here, which were reprogrammed in two different labs using different approaches,29, 30 show comparable EVC differentiation efficiency and capability to mature to functional ECs, similar to the healthy BC1-hiPSCs.

Since ECs and EPCs isolated from diabetic patients are known to be deficient due to exposure to hyperglycemia,16-18 we investigated if T1D-hiPSCs retain any deficits typical of diabetic ECs. Thus, we addressed the question by assessing whether T1D-derived ECs express mature EC markers and behaved as healthy ECs using functional assays such as Matrigel cord formation and TNFα responsiveness. We found that T1D-ECs expressed mature endothelial markers and displayed behaviors typical of mature and functional ECs, as in the case of BC1-ECs.8

One main advantage of deriving both ECs and pericytes from the same source is that the differentiated early vascular cells are readily available for co-encapsulation in synthetic hydrogels to form stable 3D vascular networks.8, 32 Using two different synthetic hydrogel systems, we demonstrated that the T1D-EVCs are capable for forming 3D networks upon encapsulation, paving the way for future research towards potential translational and therapeutic purposes.

Similar to BC1-hiPSC-derived EVCs, bi-cellular EVC populations derived from T1D-hiPSC formed 3D microvascular networks when embedded in engineered HA-based hydrogel.8 We observed vacuole formation, coalescing of vacuoles to form bigger lumens and sprouting of the T1D-EVCs during the first two days of encapsulation. Eventually, in the next stage of vascular morphogenesis, microvascular networks formed with cell sprouting and tubules joining between neighboring cells. Overall, the kinetics of morphogenesis was found to be similar between the HA-embedded T1D-EVCs and BC1-EVCs as previously described.8 This finding is encouraging as it indicates that T1D-EVCs respond to cues from the surrounding extracellular matrix to form complex 3D vascular networks.

To further validate the functionality of the T1D-EVCs, we employed a novel hypoxic hydrogel system established in our laboratory35. We demonstrated that T1D-EVCs, like endothelial progenitors35 and healthy BC1-EVCs, responded to low DO level in the HI hydrogel, by undergoing vascular morphogenesis to form 3D EVC networks. The observed mean tube length and thickness of T1D-EVCs encapsulated in hypoxic hydrogel are comparable to those of hypoxic hydrogel encapsulated endothelial progenitors35 and BC1-EVCs. In our previous study, we found that accumulation of hypoxia-inducible factors 1α and 2α activates MT1-MMP, which enables vascular morphogenesis of EPCs within the hypoxic hydrogel35. As activation of 3D vascular morphogenesis is similar in EPCs and hiPSC-EVCs,8, 32 it is safe to speculate that hiPSC-EVC morphogenesis in the hypoxic hydrogels follows the same mechanism as in EPCs.

Altogether, these observations demonstrate that T1D-EVCs are sensitive and responsive to a hypoxic microenvironment. In addition, by inducing hypoxia, the hypoxic hydrogel provides a platform for guiding and accelerating 3D EVC network formation in vitro. To our knowledge, this is the first time that the formation of EVC network in response to hypoxic microenvironment has been realized. Overall, this finding demonstrates the ability of T1D-EVCs to respond to a hypoxic matrix to form 3D networks which may lead to a therapeutic tool to treat diabetic vascular complications such as diabetic foot ulcers (DFUs). Endothelial progenitors are limited in function and in number in diabetic patients,52, 53 exacerbating vascular complications such as neovascularization in diabetic wounds. Shortened healing period of DFUs have been demonstrated to decrease chances of amputation.54 Thus, the combined approach of direct and controlled vascular differentiation and delivery of a microvascular bed in hydrogel matrix could potentially promote and accelerate wound healing in complications such as DFUs.

Supplementary Material

Materials and Methods
Supplemental Data
Supplemental Video
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Significance.

Diabetic patients are prone to cardiovascular diseases and wound healing complications due to dysfunctional endothelial progenitor cells under hyperglycemia. HiPSCs serve as a great source for generating patient-specific vascular cells, aiming towards future autologous vascular therapy for diabetics. Our robust differentiation protocol can generate early vascular cells (EVCs), including endothelial cells and pericytes from T1D-hiPSCs. Unlike isolated diabetic endothelial progenitor cells, differentiated ECs can be matured and are functional. When encapsulated in synthetic hydrogels, T1D-EVCs responded to matrix cues and self-assembled to form three-dimensional vascular networks. Moreover, T1D-EVCs responded to hypoxic microenvironments by undergoing vasculogenesis to form complex three-dimensional networks. This is the first example of EVCs derived from T1D-hiPSCs demonstrating both in vitro and in vivo functionalities. The combination of T1D-EVCs and the synthetic hydrogel system establish a platform that could be useful for therapeutic vasculogenesis in diabetic patients.

Acknowledgments

We would like to thank Dr. Tom Shen and Dr. Kyung Min Park and Michael Blatchley for providing AHA and HI hydrogels, respectively; Quinton Smith and Dr. Tom Shen for helpful discussions throughout the course of this work; Lena Washington for zebrafish husbandry; and Daniel Lewis and David White for help with Imaris analysis. We also thank Dr. Julia Wang and Anna Coughlan for critical comments on an earlier draft of this manuscript.

Sources of funding: This work was supported by National Institute of Health grant R01HL107938 and a W.W. Smith Charitable Trust grant (to SG).

Abbreviations

3D

3-dimensional

CVD

cardiovascular diseases

EC

endothelial cell

EPC

endothelial progenitor cell

EVC

early vascular cell

hiPSC

human-induced pluripotent stem cell

HA

hyaluronic acid

HI

hypoxia-inducible

PDGFRβ

platelet-derived growth factor β

ROCK

rho associated coiled coil containing kinase

T1D

type 1 diabetes

T1D-EVC

type 1 diabetes human-induced pluripotent stem cell derived early vascular cell

VEcad

vascular endothelial cadherin

Footnotes

Disclosures: None

References

  • 1.Murphy SL, Xu J, Kochanek KD. Deaths: Final data for 2010. National vital statistics reports : from the Centers for Disease Control and Prevention, National Center for Health Statistics, National Vital Statistics System. 2013;61:1–117. [PubMed] [Google Scholar]
  • 2.Resnick HE, Howard BV. Diabetes and cardiovascular disease. Annual review of medicine. 2002;53:245–267. doi: 10.1146/annurev.med.53.082901.103904. [DOI] [PubMed] [Google Scholar]
  • 3.Wilson PW. Diabetes mellitus and coronary heart disease. American journal of kidney diseases : the official journal of the National Kidney Foundation. 1998;32:S89–100. doi: 10.1053/ajkd.1998.v32.pm9820468. [DOI] [PubMed] [Google Scholar]
  • 4.Kota SK, Meher LK, Jammula S, Kota SK, Krishna SV, Modi KD. Aberrant angiogenesis: The gateway to diabetic complications. Indian journal of endocrinology and metabolism. 2012;16:918–930. doi: 10.4103/2230-8210.102992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Simons M. Angiogenesis, arteriogenesis, and diabetes: Paradigm reassessed? Journal of the American College of Cardiology. 2005;46:835–837. doi: 10.1016/j.jacc.2005.06.008. [DOI] [PubMed] [Google Scholar]
  • 6.Bento CF, Pereira P. Regulation of hypoxia-inducible factor 1 and the loss of the cellular response to hypoxia in diabetes. Diabetologia. 2011;54:1946–1956. doi: 10.1007/s00125-011-2191-8. [DOI] [PubMed] [Google Scholar]
  • 7.Caballero S, Sengupta N, Afzal A, Chang KH, Calzi SL, Guberski DL, Kern TS, Grant MB. Ischemic vascular damage can be repaired by healthy, but not diabetic, endothelial progenitor cells. Diabetes. 2007;56:960–967. doi: 10.2337/db06-1254. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Kusuma S, Shen YI, Hanjaya-Putra D, Mali P, Cheng L, Gerecht S. Self-organized vascular networks from human pluripotent stem cells in a synthetic matrix. Proceedings of the National Academy of Sciences of the United States of America. 2013;110:12601–12606. doi: 10.1073/pnas.1306562110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Seifu DG, Purnama A, Mequanint K, Mantovani D. Small-diameter vascular tissue engineering. Nature reviews Cardiology. 2013;10:410–421. doi: 10.1038/nrcardio.2013.77. [DOI] [PubMed] [Google Scholar]
  • 10.Sun G, Gerecht S. Vascular regeneration: Engineering the stem cell microenvironment. Regenerative medicine. 2009;4:435–447. doi: 10.2217/rme.09.1. [DOI] [PubMed] [Google Scholar]
  • 11.Leeper NJ, Hunter AL, Cooke JP. Stem cell therapy for vascular regeneration: Adult, embryonic, and induced pluripotent stem cells. Circulation. 2010;122:517–526. doi: 10.1161/CIRCULATIONAHA.109.881441. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Cho SW, Moon SH, Lee SH, Kang SW, Kim J, Lim JM, Kim HS, Kim BS, Chung HM. Improvement of postnatal neovascularization by human embryonic stem cell derived endothelial-like cell transplantation in a mouse model of hindlimb ischemia. Circulation. 2007;116:2409–2419. doi: 10.1161/CIRCULATIONAHA.106.687038. [DOI] [PubMed] [Google Scholar]
  • 13.Huang NF, Niiyama H, Peter C, De A, Natkunam Y, Fleissner F, Li Z, Rollins MD, Wu JC, Gambhir SS, Cooke JP. Embryonic stem cell-derived endothelial cells engraft into the ischemic hindlimb and restore perfusion. Arteriosclerosis, thrombosis, and vascular biology. 2010;30:984–991. doi: 10.1161/ATVBAHA.110.202796. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T, Witzenbichler B, Schatteman G, Isner JM. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997;275:964–967. doi: 10.1126/science.275.5302.964. [DOI] [PubMed] [Google Scholar]
  • 15.Li Z, Wilson KD, Smith B, Kraft DL, Jia F, Huang M, Xie X, Robbins RC, Gambhir SS, Weissman IL, Wu JC. Functional and transcriptional characterization of human embryonic stem cell-derived endothelial cells for treatment of myocardial infarction. PloS one. 2009;4:e8443. doi: 10.1371/journal.pone.0008443. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Loomans CJM, van Haperen R, Duijs JM, Verseyden C, de Crom R, Leenen PJM, Drexhage HA, de Boer HC, de Koning EJP, Rabelink TJ, Staal FJT, van Zonneveld AJ. Differentiation of bone marrow-derived endothelial progenitor cells is shifted into a proinflammatory phenotype by hyperglycemia. Molecular Medicine. 2009;15:152–159. doi: 10.2119/molmed.2009.00032. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Menegazzo L, Albiero M, Avogaro A, Fadini GP. Endothelial progenitor cells in diabetes mellitus. Biofactors. 2012;38:194–202. doi: 10.1002/biof.1016. [DOI] [PubMed] [Google Scholar]
  • 18.Tepper OM, Galiano RD, Capla JM, Kalka C, Gagne PJ, Jacobowitz GR, Levine JP, Gurtner GC. Human endothelial progenitor exhibit impaired proliferation, cells from type ii diabetics adhesion, and incorporation into vascular structures. Circulation. 2002;106:2781–2786. doi: 10.1161/01.cir.0000039526.42991.93. [DOI] [PubMed] [Google Scholar]
  • 19.Yu J, Vodyanik MA, Smuga-Otto K, Antosiewicz-Bourget J, Frane JL, Tian S, Nie J, Jonsdottir GA, Ruotti V, Stewart R, Slukvin II, Thomson JA. Induced pluripotent stem cell lines derived from human somatic cells. Science. 2007;318:1917–1920. doi: 10.1126/science.1151526. [DOI] [PubMed] [Google Scholar]
  • 20.Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K, Yamanaka S. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell. 2007;131:861–872. doi: 10.1016/j.cell.2007.11.019. [DOI] [PubMed] [Google Scholar]
  • 21.Mattout A, Biran A, Meshorer E. Global epigenetic changes during somatic cell reprogramming to ips cells. J Mol Cell Biol. 2011;3:341–350. doi: 10.1093/jmcb/mjr028. [DOI] [PubMed] [Google Scholar]
  • 22.Kusuma S, Peijnenburg E, Patel P, Gerecht S. Low oxygen tension enhances endothelial fate of human pluripotent stem cells. Arteriosclerosis, thrombosis, and vascular biology. 2014;34:913–920. doi: 10.1161/ATVBAHA.114.303274. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Kusuma S, Facklam A, Gerecht S. Characterizing human pluripotent-stem-cell-derived vascular cells for tissue engineering applications. Stem cells and development. 2015;24:451–458. doi: 10.1089/scd.2014.0377. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Armulik A, Genove G, Betsholtz C. Pericytes: Developmental, physiological, and pathological perspectives, problems, and promises. Developmental Cell. 2011;21:193–215. doi: 10.1016/j.devcel.2011.07.001. [DOI] [PubMed] [Google Scholar]
  • 25.Chou BK, Mali P, Huang X, Ye Z, Dowey SN, Resar LM, Zou C, Zhang YA, Tong J, Cheng L. Efficient human ips cell derivation by a non-integrating plasmid from blood cells with unique epigenetic and gene expression signatures. Cell Res. 2011;21:518–529. doi: 10.1038/cr.2011.12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Cheng L, Hansen NF, Zhao L, et al. Low incidence of DNA sequence variation in human induced pluripotent stem cells generated by nonintegrating plasmid expression. Cell stem cell. 2012;10:337–344. doi: 10.1016/j.stem.2012.01.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Watanabe K, Ueno M, Kamiya D, Nishiyama A, Matsumura M, Wataya T, Takahashi JB, Nishikawa S, Nishikawa S, Muguruma K, Sasai Y. A rock inhibitor permits survival of dissociated human embryonic stem cells. Nature biotechnology. 2007;25:681–686. doi: 10.1038/nbt1310. [DOI] [PubMed] [Google Scholar]
  • 28.Gage BK, Webber TD, Kieffer TJ. Initial cell seeding density influences pancreatic endocrine development during in vitro differentiation of human embryonic stem cells. PloS one. 2013;8:e82076. doi: 10.1371/journal.pone.0082076. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Johannesson B, Sagi I, Gore A, et al. Comparable frequencies of coding mutations and loss of imprinting in human pluripotent cells derived by nuclear transfer and defined factors. Cell Stem Cell. 2014;15:634–642. doi: 10.1016/j.stem.2014.10.002. [DOI] [PubMed] [Google Scholar]
  • 30.Maehr R, Chen S, Snitow M, Ludwig T, Yagasaki L, Goland R, Leibel RL, Melton DA. Generation of pluripotent stem cells from patients with type 1 diabetes. Proceedings of the National Academy of Sciences of the United States of America. 2009;106:15768–15773. doi: 10.1073/pnas.0906894106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Kusuma S, Macklin B, Gerecht S. Derivation and network formation of vascular cells from human pluripotent stem cells. Methods in molecular biology. 2014;1202:1–9. doi: 10.1007/7651_2013_39. [DOI] [PubMed] [Google Scholar]
  • 32.Hanjaya-Putra D, Bose V, Shen YI, Yee J, Khetan S, Fox-Talbot K, Steenbergen C, Burdick JA, Gerecht S. Controlled activation of morphogenesis to generate a functional human microvasculature in a synthetic matrix. Blood. 2011;118:804–815. doi: 10.1182/blood-2010-12-327338. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Davis GE, Stratman AN, Sacharidou A, Koh W. Molecular basis for endothelial lumen formation and tubulogenesis during vasculogenesis and angiogenic sprouting. International review of cell and molecular biology. 2011;288:101–165. doi: 10.1016/B978-0-12-386041-5.00003-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Capla JM, Grogan RH, Callaghan MJ, Galiano RD, Tepper OM, Ceradini DJ, Gurtner GC. Diabetes impairs endothelial progenitor cell-mediated blood vessel formation in response to hypoxia. Plastic and reconstructive surgery. 2007;119:59–70. doi: 10.1097/01.prs.0000244830.16906.3f. [DOI] [PubMed] [Google Scholar]
  • 35.Park KM, Gerecht S. Hypoxia-inducible hydrogels. Nature communications. 2014;5:4075. doi: 10.1038/ncomms5075. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Kusuma S, Facklam A, Gerecht S. Characterizing human pluripotent-stem-cell-derived vascular cells for tissue engineering applications. Stem cells and development. 2015;24:451–458. doi: 10.1089/scd.2014.0377. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Orlova VV, van den Hil FE, Petrus-Reurer S, Drabsch Y, Ten Dijke P, Mummery CL. Generation, expansion and functional analysis of endothelial cells and pericytes derived from human pluripotent stem cells. Nature protocols. 2014;9:1514–1531. doi: 10.1038/nprot.2014.102. [DOI] [PubMed] [Google Scholar]
  • 38.Au P, Daheron LM, Duda DG, Cohen KS, Tyrrell JA, Lanning RM, Fukumura D, Scadden DT, Jain RK. Differential in vivo potential of endothelial progenitor cells from human umbilical cord blood and adult peripheral blood to form functional long-lasting vessels. Blood. 2008;111:1302–1305. doi: 10.1182/blood-2007-06-094318. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Au P, Tam J, Fukumura D, Jain RK. Bone marrow-derived mesenchymal stem cells facilitate engineering of long-lasting functional vasculature. Blood. 2008;111:4551–4558. doi: 10.1182/blood-2007-10-118273. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Koike N, Fukumura D, Gralla O, Au P, Schechner JS, Jain RK. Tissue engineering: Creation of long-lasting blood vessels. Nature. 2004;428:138–139. doi: 10.1038/428138a. [DOI] [PubMed] [Google Scholar]
  • 41.Wang ZZ, Au P, Chen T, Shao Y, Daheron LM, Bai H, Arzigian M, Fukumura D, Jain RK, Scadden DT. Endothelial cells derived from human embryonic stem cells form durable blood vessels in vivo. Nature biotechnology. 2007;25:317–318. doi: 10.1038/nbt1287. [DOI] [PubMed] [Google Scholar]
  • 42.Holy CE, Shoichet MS, Davies JE. Engineering three-dimensional bone tissue in vitro using biodegradable scaffolds: Investigating initial cell-seeding density and culture period. Journal of biomedical materials research. 2000;51:376–382. doi: 10.1002/1097-4636(20000905)51:3<376::aid-jbm11>3.0.co;2-g. [DOI] [PubMed] [Google Scholar]
  • 43.Ghosh S, Dean A, Walter M, Bao Y, Hu Y, Ruan J, Li R. Cell density-dependent transcriptional activation of endocrine-related genes in human adipose tissue-derived stem cells. Experimental cell research. 2010;316:2087–2098. doi: 10.1016/j.yexcr.2010.04.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Chen G, Hou Z, Gulbranson DR, Thomson JA. Actin-myosin contractility is responsible for the reduced viability of dissociated human embryonic stem cells. Cell Stem Cell. 2010;7:240–248. doi: 10.1016/j.stem.2010.06.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Joo HJ, Choi DK, Lim JS, Park JS, Lee SH, Song S, Shin JH, Lim DS, Kim I, Hwang KC, Koh GY. Rock suppression promotes differentiation and expansion of endothelial cells from embryonic stem cell-derived flk1(+) mesodermal precursor cells. Blood. 2012;120:2733–2744. doi: 10.1182/blood-2012-04-421610. [DOI] [PubMed] [Google Scholar]
  • 46.Li Z, Dong XM, Wang ZL, Liu WZ, Deng N, Ding Y, Tang LY, Hla T, Zeng R, Li L, Wu DQ. Regulation of pten by rho small gtpases. Nature Cell Biology. 2005;7:399–U342. doi: 10.1038/ncb1236. [DOI] [PubMed] [Google Scholar]
  • 47.Yang S, Kim HM. The rhoa-rock-pten pathway as a molecular switch for anchorage dependent cell behavior. Biomaterials. 2012;33:2902–2915. doi: 10.1016/j.biomaterials.2011.12.051. [DOI] [PubMed] [Google Scholar]
  • 48.Song MS, Salmena L, Pandolfi PP. The functions and regulation of the pten tumour suppressor. Nature Reviews Molecular Cell Biology. 2012;13:283–296. doi: 10.1038/nrm3330. [DOI] [PubMed] [Google Scholar]
  • 49.Shiojima I, Walsh K. Role of akt signaling in vascular homeostasis and angiogenesis. Circ Res. 2002;90:1243–1250. doi: 10.1161/01.res.0000022200.71892.9f. [DOI] [PubMed] [Google Scholar]
  • 50.Castellani L, Salvati E, Alema S, Falcone G. Fine regulation of rhoa and rock is required for skeletal muscle differentiation. Journal of Biological Chemistry. 2006;281:15249–15257. doi: 10.1074/jbc.M601390200. [DOI] [PubMed] [Google Scholar]
  • 51.Pagiatakis C, Gordon JW, Ehyai S, McDermott JC. A novel rhoa/rock-cpi-17-mef2c signaling pathway regulates vascular smooth muscle cell gene expression. Journal of Biological Chemistry. 2012;287:8361–8370. doi: 10.1074/jbc.M111.286203. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Liu ZJ, Velazquez OC. Hyperoxia, endothelial progenitor cell mobilization, and diabetic wound healing. Antioxidants & redox signaling. 2008;10:1869–1882. doi: 10.1089/ars.2008.2121. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Loomans CJM, de Koning EJP, Staal FJT, Rookmaaker MB, Verseyden C, de Boer HC, Verhaar MC, Braam B, Rabelink TJ, van Zonneveld AJ. Endothelial progenitor cell dysfunction: A novel concept in the pathogenesis of vascular complications of type 1 diabetes. Diabetes. 2004;53:195–199. doi: 10.2337/diabetes.53.1.195. [DOI] [PubMed] [Google Scholar]
  • 54.Steed DL, Attinger C, Colaizzi T, Crossland M, Franz M, Harkless L, Johnson A, Moosa H, Robson M, Serena T, Sheehan P, Veves A, Wiersma-Bryant L. Guidelines for the treatment of diabetic ulcers. Wound repair and regeneration : official publication of the Wound Healing Society [and] the European Tissue Repair Society. 2006;14:680–692. doi: 10.1111/j.1524-475X.2006.00176.x. [DOI] [PubMed] [Google Scholar]

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