- Record: found
- Abstract: not found
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Human Skin Constructs with Spatially Controlled Vasculature Using Primary and iPSC-Derived Endothelial Cells
Author(s):
Hasan E. Abaci
1 ,
Zongyou Guo
1 ,
Abigail Coffman
1 ,
Brian Gillette
2 ,
Wen-han Lee
2 ,
Samuel K. Sia
2 ,
Angela M. Christiano
1
,
3
Publication date (Electronic):
June 22 2016
Journal:
Advanced Healthcare Materials
Publisher:
Wiley
There is no author summary for this article yet. Authors can add summaries to their articles on ScienceOpen to make them more accessible to a non-specialist audience.
Abstract
<p class="first" id="P1">Vascularization of engineered human skin constructs is crucial
for recapitulation
of systemic drug delivery and for their long-term survival, functionality, and viable
engraftment. In this study, we used the latest microfabrication techniques and established
a novel bioengineering approach to micropattern spatially-controlled and perfusable
vascular networks in 3D human skin equivalents using both primary and induced pluripotent
stem cell (iPSC)-derived endothelial cells (ECs). Using 3D printing technology, we
were able to control the geometry of the micropatterned vascular networks. We verified
that vascularized human skin equivalents (vHSEs) can form a robust epidermis and establish
an endothelial barrier function, which allows for the recapitulation of both topical
and systemic delivery of drugs. In addition, we examined the therapeutic potential
of vHSEs for cutaneous wounds on immunodeficient mice and demonstrated that vHSEs
can both promote and guide neovascularization during wound healing. Overall, this
innovative bioengineering approach could enable
<i>in vitro</i> evaluation of topical and systemic drug delivery, as well as improve
the potential
of engineered skin constructs to be used as a potential therapeutic option for the
treatment of cutaneous wounds.
</p>
Related collections
Most cited references29
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In vitro microvessels for the study of angiogenesis and thrombosis.
Ying Zheng, Junmei Chen, Michael P Craven … (2012)
Microvascular networks support metabolic activity and define microenvironmental conditions within tissues in health and pathology. Recapitulation of functional microvascular structures in vitro could provide a platform for the study of complex vascular phenomena, including angiogenesis and thrombosis. We have engineered living microvascular networks in three-dimensional tissue scaffolds and demonstrated their biofunctionality in vitro. We describe the lithographic technique used to form endothelialized microfluidic vessels within a native collagen matrix; we characterize the morphology, mass transfer processes, and long-term stability of the endothelium; we elucidate the angiogenic activities of the endothelia and differential interactions with perivascular cells seeded in the collagen bulk; and we demonstrate the nonthrombotic nature of the vascular endothelium and its transition to a prothrombotic state during an inflammatory response. The success of these microvascular networks in recapitulating these phenomena points to the broad potential of this platform for the study of cardiovascular biology and pathophysiology.
- Record: found
- Abstract: found
- Article: found
Efficient Differentiation of Human Pluripotent Stem Cells to Endothelial Progenitors via Small-Molecule Activation of WNT Signaling
Xiaojun Lian, Xiaoping Bao, Abraham Al-Ahmad … (2014)
Introduction Human pluripotent stem cells (hPSCs) offer unprecedented opportunities to study the earliest stages of human development in vitro, to model human disease, to perform drug tests in culture, and to develop unlimited new sources of cells for possible therapeutic applications. To realize this potential, it is essential to be able to control hPSC differentiation to somatic lineages with high efficiency and reproducibility in a scalable and inexpensive manner (Ashton et al., 2011; Burridge et al., 2012; Kinney et al., 2014; Murry and Keller, 2008). Functional human endothelial cells differentiated from hPSCs could be beneficial for many potential clinical applications (Burridge et al., 2012; Kaupisch et al., 2012; Levenberg et al., 2002; van der Meer et al., 2013), including engineering new blood vessels, endothelial cell transplantation into the heart for myocardial regeneration (Robey et al., 2008), and induction of angiogenesis for treatment of regional ischemia (Liu et al., 2014). Endothelial cell dysfunction is also associated with many diseases, including Alzheimer’s disease, stroke, multiple sclerosis, and atherosclerosis (Boyle et al., 1997; Weiss et al., 2009). hPSC-derived endothelial progenitors and endothelial cells may provide building blocks for the establishment of in vitro disease models for screening and development of drugs to treat these diseases. Functionality of hPSC-derived endothelial cells has been shown using in vitro cell culture platforms and in vivo animal models (Adams et al., 2013; Kusuma et al., 2013; Orlova et al., 2014; Samuel et al., 2013; Wang et al., 2007). Similar to other somatic cells derived from hPSCs, differentiated CD31+ endothelial cells exhibited functional heterogeneity (Rufaihah et al., 2013). Previously reported studies of hPSC differentiation to endothelial cells have demonstrated that Activin/Nodal/transforming growth factor β (TGF-β), bone morphogenetic protein (BMP), vascular endothelial growth factor (VEGF), and microRNA-21 signaling promote this differentiation (Di Bernardini et al., 2013; James et al., 2010; Kane et al., 2010; Lu et al., 2007; Marchand et al., 2014; Rufaihah et al., 2011; Wang et al., 2004; Zambidis et al., 2005). In addition, mechanical sheer stress also promoted embryonic stem cell-derived endothelial phenotypes (Wolfe and Ahsan, 2013). During murine embryogenesis, hemangioblasts, which can differentiate into multipotent hematopoietic stem cells and endothelial progenitors, are derived from a subpopulation of mesoderm that coexpresses brachyury and KDR (Huber et al., 2004). Similar blast colony-forming cells were also isolated from mouse embryonic stem cell aggregates in the presence of cytokines (Kennedy et al., 1997). When cocultured with OP9 stromal cells, hPSCs differentiated to mesodermal progenitors with the capacity to form blast or hemangioblast colonies in response to fibroblast growth factor 2 (FGF2) (Vodyanik et al., 2010). As another approach, hPSCs cultured as embryoid bodies were exposed to a growth factor cocktail containing activin A, BMP4, FGF2, and VEGF to induce differentiation to CD34+CD31+ endothelial progenitors (Costa et al., 2013; Levenberg et al., 2002; Song et al., 2013). The CD34+CD31+ vascular progenitor population generated endothelial cells and smooth muscle cells in the proper culture environments (Bai et al., 2010). TGF-β signaling enhanced smooth muscle cell differentiation from these endothelial progenitors, whereas the TGF-β signaling inhibitor SB431542 promoted endothelial cell generation and expansion (James et al., 2010). Global gene transcription analysis demonstrated low variability between endothelial cells (ECs) differentiated from multiple lines of human embryonic stem cells (hESCs) and induced pluripotent stem cells (iPSCs) in the presence of these cytokines (White et al., 2013). Although prior studies have demonstrated differentiation of hPSCs to endothelial progenitors, and subsequently to ECs and smooth muscle cells, by applying growth factors from different signaling pathways, it is largely unknown whether these distinct differentiation protocols produce identical endothelial cells and their progenitors, and which developmental signaling mechanisms are necessary and sufficient to specify these differentiation fates. Here, we describe a simple and efficient method for the conversion of hPSCs to CD34+CD31+ endothelial progenitors. Appropriate temporal activation of regulators of WNT signaling alone, in the absence of exogenous FGF2 and VEGF signaling, was sufficient to drive multiple hPSC lines to differentiate to greater than 50% CD34+CD31+ endothelial progenitors. However, endogenous MEK signaling was required for hPSC differentiation to endothelial progenitors because MEK inhibitor treatment substantially diminished the yield of CD34+CD31+ cells. These hPSC-derived endothelial progenitors were further enriched to 99% purity with a single step of CD34-based magnetic separation. Single-cell clonal differentiation assays revealed that CD34+CD31+ endothelial progenitors generated by WNT pathway activation were bipotent and could differentiate to functional endothelial cells and smooth muscle cells. Results GSK3 Inhibition Is Sufficient to Induce hPSC Differentiation to Endothelial Progenitors in the Absence of Exogenous Growth Factors We previously demonstrated that temporal modulation of canonical WNT signaling is sufficient to generate functional cardiomyocytes at high yield and purity (Lian et al., 2012, 2013b). In that study, a virtually pure population of brachyury+ cells was generated within 24 hr after treatment of undifferentiated hPSCs with a GSK3 inhibitor. In order to direct these brachyury+ cells to a cardiomyocyte fate, we then inhibited canonical WNT signaling by either expression of β-catenin small hairpin RNA (shRNA) or treatment with small-molecule inhibitors of WNT signaling at the proper corresponding time points. To assess whether these GSK3 inhibitor-induced brachyury+ mesendoderm progenitors also have the potential to differentiate to endothelial progenitors, we treated 19-9-11 iPSCs with the GSK3 inhibitor CHIR99021 and subsequently cultured the cells in StemPro-34 medium supplemented with VEGF, both of which have been reported to stimulate vascular cell differentiation of hPSCs (Grigoriadis et al., 2010) (Figure 1A). After optimizing the concentrations and timing of application of various components, the differentiation protocol consisted of two phases. Phase I generated brachyury+ cells by treatment of undifferentiated hPSCs with CHIR990211 for 2 days, whereas phase II directed the brachyury+ progenitors to CD34+CD31+ cells by culture in StemPro-34 supplemented with VEGF for 3 days. We found that without CHIR99021 treatment, very few CD34+CD31+ cells were generated from human iPSCs, even in the presence of VEGF (Figure 1B). However, treatment of the human iPSCs with CHIR99021 generated more than 20% CD34+CD31+ cells. Interestingly, VEGF treatment did not significantly increase the yield of CD34+CD31+ cells in this differentiation system (Figure 1B). However, in the presence of sunitinib, which inhibits several receptor tyrosine kinases including VEGFR, very few CD34+CD31+ cells were generated, suggesting that endogenous VEGF signaling is required for endothelial progenitor differentiation stimulated by CHIR99021 treatment (Figure 1C). Because of these observations, we did not include exogenous VEGF in subsequent differentiation experiments. In addition, this differentiation protocol primarily generated CD34+CD31+ cells and produced very few CD34−CD31+ or CD34+CD31− cells (Figure 1B). We further tested the concentration dependence of CHIR99021 for inducing CD31+CD34+ endothelial progenitor differentiation and found that 6–10 μM generated 25% endothelial progenitors (Figure 1D). Similar results were obtained in the H1 hESC line (Figure S1 available online). These CD34+CD31+ endothelial progenitors also expressed CD144 (VE-cadherin), but not CD45, indicating these cells were not in hematopoietic lineages (Figure S1). These results illustrate that induction of WNT signaling via the GSK3 inhibitor CHIR99021 permitted subsequent differentiation of hPSCs to CD34+CD31+ cells, and that exogenous VEGF was not required. Endothelial Progenitor Differentiation Induced by GSK3 Inhibitors in hPSCs Is β-Catenin Dependent Selectivity is a general concern with the use of chemical inhibitors. We tested other GSK3 inhibitors, BIO-acetoxime and CHIR98014, and found that both also effectively induced endothelial progenitor differentiation to a similar extent as CHIR99021 in the absence of VEGF treatment (Figure 2A). These small molecule compounds have distinct chemical structures, reducing the likelihood of shared off-target effects. However, GSK3 inhibition can affect multiple signaling pathways, including the desired activation of canonical WNT signaling by stabilizing β-catenin protein. In order to evaluate the role of β-catenin in GSK3 inhibitor-induced hPSC endothelial differentiation, we generated an iPSC line (19-9-11 ishcat-1) expressing β-catenin shRNA under the control of a tet-regulated inducible promoter. Upon doxycycline (dox) addition, the shRNA efficiently downregulated β-catenin expression (Figure 2B). We used this cell line to examine the stage-specific roles of β-catenin during monolayer endothelial differentiation stimulated by GSK3 inhibition. Undifferentiated 19-9-11 ishcat-1 iPSCs were treated with 6 μM CHIR99021 for 48 hr followed by another 3 days culture in StemPro-34 medium. Dox was added at various time points between day 0 and day 4, and endothelial progenitor differentiation was assessed at day 5 as the percentage of CD34+CD31+cells. We found that WNT/β-catenin signaling was essential for endothelial induction by CHIR99021 because β-catenin knockdown at day 0 did not generate CD34+CD31+ cells. We previously showed that β-catenin is essential for brachyury expression after hPSC treatment with GSK3 inhibitors (Lian et al., 2012). We reasoned that abrogation of the CD34+CD31+ population by β-catenin knockdown at day 0 was due to blocking differentiation of hPSCs to brachyury-expressing mesendoderm progenitors. We profiled the expression of brachyury during the 5 day differentiation and found a virtually pure population of brachyury+ cells generated by day 2 in the absence of dox. The presence of dox treatment, however, abolished brachyury expression (Figure S2). Importantly, depletion of β-catenin expression at time points after brachyury expression was detected also reduced the percentage of CD34+CD31+ cells generated (Figure 2C), indicating β-catenin might also be involved in differentiation of mesendoderm to CD34+CD31+ cells. Because of the important role of FGF2 signaling on brachyury+ cell differentiation from hPSCs (Yu et al., 2011) and the mesodermal origin of endothelial progenitors, we used an MEK inhibitor to study the role of MEK/ERK signaling on CD34+CD31+ cell differentiation. We found that 1 μM of MEK inhibitor PD0325901, which effectively blocked ERK phosphorylation (Figure 2D), substantially diminished CD34+CD31+ cell differentiation, suggesting that endogenous MEK/ERK signaling promotes endothelial progenitor differentiation (Figure 2E). Differentiation Medium and Cell Density Dependence of GSK3 Inhibitor-Induced Endothelial Progenitor Differentiation We have shown that hPSCs can be differentiated to CD34+CD31+ endothelial progenitors by 2 days (phase I) of culture in an optimized defined medium supplemented with GSK3 inhibitors and another 3 days (phase II) in StemPro-34 medium. In order to streamline the differentiation process, we tested whether a single basal medium could support endothelial progenitor differentiation (Figure 3A). We examined 13 different media supplemented with CHIR99021 for 2 days (phase I) followed by 3 additional days in the same basal medium lacking exogenous growth factors or chemical inhibitors (phase II). We found that advanced DMEM/F12 supplemented with ascorbic acid, which we termed LaSR basal medium, was the most efficient for endothelial progenitor differentiation, resulting in 25% CD34+CD31+ cells in 5 days (Figure 3B). These results indicated the importance of basal medium composition in endothelial progenitor differentiation, with some media (e.g., RPMI/B27 and EC medium containing serum) not supporting differentiation to any detectable CD34+CD31+ cells (Figure 3B). On the basis of the knowledge that initial cell density affects hPSC differentiation fates to lineages such as cardiomyocytes, epithelial progenitors, and pancreatic cells (Gage et al., 2013; Lian et al., 2013a; Selekman et al., 2013; Takizawa-Shirasawa et al., 2013), we quantitatively assessed the effect of initial cell seeding density on endothelial progenitor differentiation efficiency. Singularized hPSCs were seeded at densities ranging from 12,000 to 300,000 cells per cm2 and then expanded in mTeSR1 for 3 days before transition to LaSR basal medium supplemented with CHIR99021 for 2 days and another 3 days in LaSR basal medium lacking CHIR99021. The optimal seeding density was 50,000 cells per cm2 at day −3, which resulted in 325,000 cells per cm2 at day 0 (Figure 3C). At day 5, 55% of cells seeded at this density expressed CD34 and CD31 (Figures 3C and 3D). We performed fluorescence-activated cell sorting (FACS) analysis of OCT4 expression of day 0 cells and found that more than 99.5% of the cells were OCT4+, suggesting that hPSCs were pluripotent before addition of CH (Figure 3E). CD34 + CD31 + Endothelial Progenitors Are Multipotent Molecular analysis of endothelial progenitor differentiation from hPSCs revealed dynamic changes in gene expression, with downregulation of the pluripotency markers SOX2 and OCT4, and induction of the primitive streak-like genes T (Nakanishi et al., 2009) and MIXL1 (Davis et al., 2008) in the first 24 hr after CH addition (Figure 4A). Expression of the endothelial progenitor markers CD34 and CD31 was detected at day 4 and increased at day 5 (Figure 4A). Western blot analysis of protein expression revealed decreasing OCT4 expression during the first 2 days of differentiation, and brachyury induction on days 1 and 2 followed by downregulation on day 3. The endothelial marker VE-cadherin was only detected after 5 days of differentiation (Figure 4B). Immunofluorescent analysis revealed the expression of CD34, CD31, and VE-cadherin on day 5 (Figure 4C). Ten days after differentiation without passaging, however, the expression of CD34 diminished, whereas the cells retained expression of CD31+ and VE-cadherin (Figure 4C). To study the multipotent nature of the transient population of CD34+ cells, we first enriched the day 5 differentiation culture to 99% purity of CD34+ cells by magnetic-activated cell sorting (MACS) (Figure 5A). We also enriched the day 5 differentiation cultures of additional hESC lines (H13, H14) and human iPSC lines (19-9-7 and 6-9-9) to more than 99% purity of CD34+ cells (Figures S3A–S3D). The sorted CD34+ putative endothelial progenitors were plated on Matrigel-coated 48-well plates at a density of one cell per well in a combined medium (50% smooth muscle and 50% endothelial medium). After 10 days of culture, the differentiation populations from ten different single-cell clones contained both VE-cadherin+ von Willebrand factor (vWF)+ cells and Calponin+ SMMHC+ (Smooth muscle myosin heavy chain) cells, including some cells expressing both endothelial and smooth muscle markers (Figure 5B; Figures S3E and S3F), thereby demonstrating that single CD34+ endothelial progenitors generated from hPSCs were multipotent. In addition, when the sorted CD34+ clones were cultured in endothelial or smooth muscle medium rather than the mixed medium, they generated relatively pure populations of endothelial cells and smooth muscle cells, respectively (Figures 5C and 5D). The hPSC-derived endothelial progenitors expressed C-KIT (CD117), KDR (VEGFR2), TIE-2, CD31, CD34, CD144, but did not express vWF or intercellular adhesion molecule 1 (ICAM-1) (Figure 6). We also quantified TIE-2, CD117, KDR, and ICAM-1 expression with flow cytometry (Figure S4), and results were consistent with immunostaining experiments. The day 15 endothelial cells expressed CD31, CD144, vWF, ICAM-1, and KDR, but did not express CD117, CD34, or TIE-2 (Figure 6). Characterization of Endothelial Cells Derived from hPSCs To study the intrinsic properties of endothelial cells differentiated from hPSCs by GSK3 inhibition and culture in LaSR basal medium, we cultivated the purified CD34+ cells in endothelial medium on laminin-coated plates and found that the resulting CD34+CD31+vWF+CD144+ endothelial cells were capable of approximately 20 population doublings over 2 months, generating more than 1 million cells from a single endothelial cell (Figure 7A). After 2 months in culture, the cells still expressed CD31 and VE-cadherin based on immunostaining (Figure 7B). Flow cytometry analysis also showed that more than 99% of cells expressed CD31 and VE-cadherin after 60 days in culture (Figure S5). Furthermore, upon treatment with exogenous VEGF, the CD144+ cells formed vascular tubes in Matrigel matrix (Figure 7C) and were capable of acetylated low-density lipoprotein (AcLDL) uptake, indicative of endothelial function (Figure 7D). The endothelium also responds to inflammatory mediators, such as tumor necrosis factor α (TNF-α), by upregulating adhesion molecules including ICAM-1, which has been implicated in the capture of circulating leukocytes. Although unstimulated endothelial cells expressed low levels of ICAM-1, TNF-α treatment greatly increased the expression of ICAM-1 in the hPSC-derived endothelial cells (Figure 7E). To characterize the barrier phenotype of the hPSC-derived endothelial cells, we measured the transendothelial electrical resistance (TEER) and transport of fluorescent 40 kDa dextran across a monolayer of differentiated endothelial cells. VEGF treatment induced a sustained decrease in TEER and an increase in dextran permeability. However, treatment of endothelial cells with the cAMP analog 8-pCPT-2′O-Me-cAMP (o-Me) decreased dextran permeability and increased TEER (Figures 7F and 7G). Discussion Prior reports of differentiating hPSCs to endothelial cells required the addition of expensive growth factors and/or undefined serum to direct endothelial development, and these methods generated heterogeneous cell mixtures typically containing less than 10% endothelial cells. Here, we show that small-molecule activation of canonical WNT signaling via GSK3 inhibition, in the absence of exogenous growth factors, is sufficient to generate high yields of CD34+CD31+ endothelial progenitors from hPSCs. Prior to this study, endothelial progenitors have not been isolated from hPSCs under defined conditions, and the mechanisms by which developmental signaling pathways regulate endothelial progenitor commitment remained largely unknown (Slukvin, 2013). With a doxycyline-inducible shRNA knockdown of the WNT pathway modulator β-catenin, we identified that β-catenin is required for CD34+CD31+ endothelial progenitor generation from hPSCs following GSK3 inhibition. Furthermore, although previous reports required more than 10 days to generate CD34+CD31+ cells, this small molecule differentiation approach accelerates developmental timing and converts hPSCs to greater than 50% CD34+CD31+ cells in 19-9-11 iPSCs in 5 days. This protocol produced more than 99% pure populations of CD34+CD31+ cells from six different hPSC lines following a single magnetic affinity separation. GSK3 inhibition has been shown to be a potent inducer of mesendoderm lineage commitment in hPSCs (Tan et al., 2013). The WNT pathway exhibits crosstalk with a variety of other developmental signaling networks, including TGF-β superfamily (Cai et al., 2013), FGF (Stulberg et al., 2012), Notch (Wang et al., 2013), Hippo (Hergovich and Hemmings, 2010), and retinoic acid signaling (Chanda et al., 2013). The necessity of β-catenin in generating CD34+CD31+ cells indicates a direct role of canonical WNT signaling in endothelial progenitor differentiation, whereas the diminished yield of CD34+CD31+ cells in the presence of MEK and receptor tyrosine kinase inhibitors suggests endogenous pathways also mediate endothelial progenitor differentiation. Thus, canonical WNT signaling is not the only pathway required for endothelial progenitor specification, but it may act as a master regulator to orchestrate signaling that leads to endothelial progenitor differentiation. We have previously reported a method to differentiate hPSCs to a relatively pure population of cardiomyocytes by dynamic modulation of canonical WNT signaling using small-molecule GSK3 inhibitors and Porcupine inhibitors in a RPMI/B27 medium lacking insulin (Lian et al., 2012, 2013a). This protocol uses WNT pathway activation to direct mesendoderm differentiation, followed by WNT pathway inhibition to specify cardiac mesoderm and cardiomyocyte fates. Although the small molecule cardiomyocyte and endothelial progenitor differentiation protocols result in optimal cell yields in distinct defined basal media, both progress through mesodermal progenitors via GSK3 inhibition. These mesodermal progenitors can be directed to spontaneously contracting cardiomyocytes via WNT inhibition, while permitting endogenous canonical WNT signaling yields a population enriched in endothelial progenitors. Together, these results demonstrate the concept of chemically guiding differentiation to distinct cell types via different patterns of modulation of master regulators of cell fates. This paradigm may represent a promising general approach to efficiently produce cells and tissues from stem cell sources in a defined manner. Interestingly, the WNT pathway-induced endothelial progenitor differentiation protocol primarily generates CD34 and CD31 double-positive cells, as opposed to other methods using BMP4, which produce a mixture of CD34+CD31−, CD34+CD31+, CD34−CD31+ cells (Bai et al., 2010; Song et al., 2013). The CD34+CD31+ endothelial progenitors may be more amenable to large-scale expansion. Furthermore, these endothelial progenitors can be further directed to endothelial cells or smooth muscle cells by culture in the appropriate inductive media. Purified CD31+VE-cadherin+vWF+CD34− endothelial cells were expanded for 20 doublings over 2 months and maintained expression of endothelial cell markers (CD31+VE-cadherin+CD34−). These cells exhibited uptake of acetylated low-density lipoprotein and formed tube-like structures when cultured on Matrigel. These endothelial cells also responded to TNF-α treatment by increasing ICAM-1 expression and maintained a dynamic barrier, responding to VEGF and cAMP analog by increasing or decreasing dextran permeability, respectively. Our findings demonstrate efficient and reproducible endothelial progenitor and endothelial differentiation of hPSCs by GSK3 inhibition in the absence of exogenous growth factor stimulation. Experimental Procedures Maintenance of hPSCs Transgene and vector free human iPSCs (19-9-11, 19-9-7, 6-9-9) (Yu et al., 2009) and hESCs (H1, H13, H14) (Thomson et al., 1998) were maintained on Matrigel (BD Biosciences)-coated plates (Corning) in mTeSR1 medium (STEMCELL Technologies) according to a previously published method (Lian et al., 2013a). Endothelial Progenitor Differentiation via Modulation of Canonical WNT Signaling hPSCs maintained on a Matrigel-coated surface in mTeSR1 were dissociated into single cells with Accutase (Life Technologies) at 37°C for 5 min and then seeded onto a Matrigel-coated cell culture dish at 50,000 cell/cm2 in mTeSR1 supplemented with 5 μM ROCK inhibitor Y-27632 (Selleckchem) (day −3) for 24 hr. Cells were then cultured in mTeSR1, changed daily. At day 0, cells were treated with 6–10 μM CHIR99021 (Selleckchem) for 2 days in LaSR basal medium, which consists of Advanced DMEM/F12, 2.5 mM GlutaMAX, and 60 μg/ml ascorbic acid (Sigma, A8960). After 2 days, CHIR99021-containing medium was aspirated and cells were maintained in LaSR basal medium without CHIR99021 for 3–4 additional days. PD0325901 was purchased from Tocris Bioscience. Differentiation of CD34+ Cells to Endothelial Cells Day 5 differentiated populations were dissociated with Accutase for 10 min and purified with an EasySep Magnet kit (STEMCELL Technologies) using a CD34 antibody according to the manufacturer’s instructions. The purified CD34+ cells were plated on collagen-IV-coated dishes (BD BioCoat) in EGM-2 medium (Lonza) and split every 4–5 days with Accutase. Differentiation of CD34+ Cells to Smooth Muscle Cells Day 5 differentiated populations were dissociated with Accutase for 10 min and purified with an EasySep Magnet kit (STEMCELL Technologies) using a CD34 antibody according to the manufacturer’s instructions. The purified CD34+ cells were plated on collagen-IV-coated dishes (BD BioCoat) in SmGM-2 medium (Lonza) and split every 4–5 days with Accutase. Vascular Tube Formation Assay To assess the formation of capillary structures, 1 × 105 endothelial cells in 0.4 ml EGM-2 medium (Lonza) supplemented with 50 ng/ml VEGF (R&D Systems) were plated into one well of 24-well tissue culture plate precoated with 250 μl Matrigel (BD Bioscience). Tube formation was observed by light microscopy after 24 hr of incubation. RT-PCR and Quantitative RT-PCR Total RNA was prepared with the RNeasy mini kit (QIAGEN) and treated with DNase (QIAGEN). One microgram RNA was reverse transcribed into cDNA via Oligo (dT) with Superscript III Reverse Transcriptase (Invitrogen). Real-time quantitative PCR was done in triplicate with iQSYBR Green SuperMix (Bio-Rad). RT-PCR was performed with Gotaq Master Mix (Promega) and then subjected to 2% agarose gel electrophoresis. ACTB was used as an endogenous housekeeping control. PCR primer sequences are provided in the Supplemental Experimental Procedures. Flow Cytometry Cells were dissociated into single cells with Accutase for 10 min and then fixed with 1% paraformaldehyde for 20 min at room temperature and stained with primary and secondary antibodies (Supplemental Experimental Procedures) in PBS plus 0.1% Triton X-100 and 0.5% BSA. Data were collected on a FACSCaliber flow cytometer (Beckton Dickinson) and analyzed using FlowJo. FACS gating was based on the corresponding isotype antibody control. Immunostaining Cells were fixed with 4% paraformaldehyde for 15 min at room temperature and then stained with primary and secondary antibodies (Supplemental Experimental Procedures) in PBS plus 0.4% Triton X-100 and 5% nonfat dry milk (Bio-Rad). Nuclei were stained with Gold antifade Reagent with DAPI (Invitrogen). An epifluorescence microscope (Leica DM IRB) with a QImaging Retiga 4000R camera was used for imaging analysis. Western Blot Analysis Cells were lysed in M-PER Mammalian Protein Extraction Reagent (Pierce) in the presence of Halt Protease and Phosphatase Inhibitor Cocktail (Pierce). Proteins were separated by 10% Tris-Glycine SDS/PAGE (Invitrogen) under denaturing conditions and transferred to a nitrocellulose membrane. After blocking with 5% dried milk in TBST, the membrane was incubated with primary antibody overnight at 4°C. The membrane was then washed, incubated with an anti-mouse/rabbit peroxidase-conjugated secondary antibody (Supplemental Experimental Procedures) at room temperature for 1 hr, and developed by SuperSignal chemiluminescence (Pierce). Monolayer Integrity Purified CD31+ endothelial cells were seeded on 12-well plate size 0.4 μm porous polystyrene membrane inserts (Transwells, Corning) coated with a 0.4 mg/ml collagen IV and 0.1 mg/ml fibronectin mixture at a density of 250,000 cells/insert. Cells were maintained in endothelial cell serum-free medium (SFM) (Life Technologies) for 48 hr prior to addition 0.5 ml and 1.5 ml of endothelial SFM at the apical and basolateral chambers respectively. Barrier tightness was assessed by measurement of transendothelial electrical resistance (TEER) using aEVOM STX2 electrode system (World Precision Instruments) before medium replacement. Baseline resistance values served to normalize subsequent TEER measurements. Cells were treated with either 100 ng/ml vascular endothelial growth factor (VEGF) or 10 μM 8-(4-chlorophenylthio)-2′-O-Me-cAMP (oMe). TEER measurements were acquired every 2 hr, with the exception of oME-treated cells. After 24 hr of treatment, vascular permeability was assessed by replacing the medium in the top chamber with 500 μl of 1 μM 40 kDa fluorescein isothiocyanate-dextran (Sigma-Aldrich) dissolved in endothelium SFM. At 15, 30, 45, and 60 min, 150 μl aliquots were taken from the basolateral chamber and replaced with warm endothelial SFM. Cell permeability was calculated following the Pe clearance method (Calabria et al., 2006; Jones and Shusta, 2007). Statistics Data are presented as mean ± SEM. Statistical significance was determined by Student’s t test (two-tail) between two groups. p < 0.05 was considered statistically significant. Author Contributions X.L. and X.B. conceived, performed, and interpreted most of the experiments and wrote the manuscript; J.L., Y.W., W.D., and K.K.D. assisted in endothelial differentiation and characterization experiments; A.A. performed and interpreted endothelial characterization experiments; E.V.S. provided experimental advice and funding support; S.P.P. conceived experiments, provided experimental advice, wrote the manuscript, and provided funding support.
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In vitro perfused human capillary networks.
Monica Moya, Yu-Hsiang Hsu, Abraham Lee … (2013)
Replicating in vitro the complex in vivo tissue microenvironment has the potential to transform our approach to medicine and also our understanding of biology. In order to accurately model the 3D arrangement and interaction of cells and extracellular matrix, new microphysiological systems must include a vascular supply. The vasculature not only provides the necessary convective transport of oxygen, nutrients, and waste in 3D culture, but also couples and integrates the responses of organ systems. Here we combine tissue engineering and microfluidic technology to create an in vitro 3D metabolically active stroma (∼1 mm(3)) that, for the first time, contains a perfused, living, dynamic, interconnected human capillary network. The range of flow rate (μm/s) and shear rate (s(-1)) within the network was 0-4000 and 0-1000, respectively, and thus included the normal physiological range. Infusion of FITC dextran demonstrated microvessels (15-50 μm) to be largely impermeable to 70 kDa. Our high-throughput biology-directed platform has the potential to impact a broad range of fields that intersect with the microcirculation, including tumor metastasis, drug discovery, vascular disease, and environmental chemical toxicity.
Author and article information
Journal
Title:
Advanced Healthcare Materials
Abbreviated Title:
Adv. Healthcare Mater.
Publisher:
Wiley
ISSN:
21922640
Publication date Created:
July
2016
Publication date
(Print):
July
2016
Publication date
(Electronic):
June
22 2016
Volume: 5
Issue: 14
Pages: 1800-1807
Affiliations
Article
DOI: 10.1002/adhm.201500936
PMC ID: 5031081
PubMed ID: 27333469
SO-VID: 7cd4246d-9781-402b-86cd-436f3728e8ed
Copyright © ©
2016
History
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