Basic Research

A Novel Three–Dimensional Human Peritubular Microvascular System

Giovanni Ligresti, Ryan J. Nagao, Jun Xue, Yoon Jung Choi, Jin Xu, Shuyu Ren, Takahide Aburatani, Susan K. Anderson, James W. MacDonald, Theo K. Bammler, Stephen M. Schwartz, Kimberly A. Muczynski, Jeremy S. Duffield, Jonathan Himmelfarb and Ying Zheng
JASN August 2016, 27 (8) 2370-2381; DOI: https://doi.org/10.1681/ASN.2015070747

Abstract

Human kidney peritubular capillaries are particularly susceptible to injury, resulting in dysregulated angiogenesis, capillary rarefaction and regression, and progressive loss of kidney function. However, little is known about the structure and function of human kidney microvasculature. Here, we isolated, purified, and characterized human kidney peritubular microvascular endothelial cells (HKMECs) and reconstituted a three-dimensional human kidney microvasculature in a flow-directed microphysiologic system. By combining epithelial cell depletion and cell culture in media with high concentrations of vascular endothelial growth factor, we obtained HKMECs of high purity in large quantity. Unlike other endothelial cells, isolated HKMECs depended on high vascular endothelial growth factor concentration for survival and growth and exhibited high tubulogenic but low angiogenic potential. Furthermore, HKMECs had a different transcriptional profile. Under flow, HKMECs formed a thin fenestrated endothelium with a functional permeability barrier. In conclusion, this three-dimensional HKMEC-specific microphysiologic system recapitulates human kidney microvascular structure and function and shows phenotypic characteristics different from those of other microvascular endothelial cells.

The kidneys play an essential role in the body to eliminate harmful substances from blood, including endogenous metabolic waste products, exogenously administered xenobiotics, and environmental toxins. As a major recipient of cardiac output (approximately 25%) and the primary filter of exogenous drugs and toxins, kidneys are highly vascular, and the tubulointerstitium is particularly susceptible to injury, clinically resulting in AKI1 and contributing to the incidence and progression of CKD.2,3

Of the two major components in the kidney tubulointerstitium, the kidney microvasculature has received relatively less attention in human studies. These vessels play a critical role in delivering nutrients to tubular epithelial cells, possess unique transport properties,4–7 and participate in the tubular secretion and reabsorption of solutes.8 Studies over the past decade have shown that kidney peritubular microvessels are highly susceptible to rarefaction after exposure to toxins, xenobiotics, or injury.9,10 After injured, they exhibit limited regenerative capacity, which may contribute to tissue ischemia, tubular dysfunction, inflammation, fibrosis, and the development of CKD.11

The mechanisms underlying the microvascular response to peritubular kidney injury, however, remain unclear, in part because of difficulties resulting from in vivo imaging as well as challenges in isolating human kidney microvascular cells for in vitro study.12 Although glomerular endothelial cells have been successfully isolated and characterized,13,14 little progress has been made on human kidney peritubular microvascular cells. Much of our understanding of kidney capillary formation and maintenance has been extrapolated from the study of other endothelial cells,9,15 which may not capture specific properties of the human kidney peritubular microvasculature. New evidence from genetic fate–mapping studies in mice suggests that the microvascular endothelium of the internal organs may not arise from a single–yolk sac progenitor as was originally thought but rather, from discrete organ–specific mesenchymal cells that appear early in embryogenesis and subsequently, give rise to multiple organ–specific populations, including the endothelium.16 Odd Skipped–Related 1–positive progenitors likely give rise to all cell populations in the kidney, including the microvascular endothelium. This suggests that, rather than the endothelium being imposed on by the organ developing around it, organ-specific characteristics might be intrinsic to the endothelium.17

Another important characteristic of the kidney microvasculature is the constant subjection to high blood flow and transport. Conventional planar cultures of endothelial cells fail to recreate the in vivo physiology of the microvasculature with respect to the three-dimensional (3D) geometry (lumen and axial branching) and the interactions of the endothelium with blood flow and extracellular matrix. To address these challenges, we have recently engineered functional vascular networks on the basis of microfluidic design principles that permit precise control of vascular cell types, branching architecture, lumen diameter, and flow dynamics.18 This approach allows us to now reconstruct human kidney microvessels under physiologic geometry and flow conditions.

In this study, we present new methods to isolate, purify, and expand human kidney peritubular microvascular endothelial cells (HKMECs) and recreate the kidney microvasculature with appropriate geometry and flow. We show that HKMEC-formed microvessels have kidney-specific properties, exemplified by the presence of fenestral diaphragms on the endothelial membrane, low angiogenic potential, and increased sensitivity to flow-induced biophysical changes. These experiments indicate a functioning human kidney microvasculature can be recapitulated in vitro. We discuss the potential application of our system for modeling nephrotoxicity as well as the onset and progression of kidney disease.

Results

Purification, Expansion, and Characterization of HKMECs

HKMECs were purified from both adult nephrectomy specimens and fetal kidneys between 100 and 135 days postconception. The fetal kidneys at that stage contained established nephrons, including glomeruli, tubules, and interstitium (Figure 1A), and have begun to produce urine. Compared with the mature adult kidneys (Figure 1B), fetal kidneys showed signs of ongoing development, exemplified by having a higher cellularity [(1.66±0.04)×106 cells per mm3 in fetal versus (0.61±0.08)×106 cells per mm3 in adults] and smaller glomeruli diameter (73.75±17.5 μm in fetal versus 166.25±22.5 μm in the adult). In the interstitium of fetal kidneys, the microvasculature has established a network around the tubules, and endothelial cells strongly express CD31 (Figure 1A.2). The adult kidneys have fully established tubular structure surrounded with a robust peritubular microvascular network that strongly expresses CD31 (Figure 1B.2), VE Cadherin, and CD34 (Supplemental Figure 1, A and B). This peritubular microvascular endothelium is enveloped by a scattered layer of PDGFRβ+ stroma (Supplemental Figure 1, C and D). In addition, the peritubular microvascular endothelium was distinguished from the glomerular endothelium by low granular expression of vWF and high expression of a plasmalemma protein (PV1) delineating fenestral diaphragms (Figure 1, A.3 and B.3). In adult kidneys, glomerular endothelium did not express PV1, whereas partial staining of PV1 was still present in the fetal kidney glomeruli, suggesting the development stage of the fetal kidneys with immature glomerular capillaries. This is consistent with previous observations that PV1 is present in the glomerular capillaries during early development but no longer expressed in postnatal glomerular endothelium.19

Figure 1.
Figure 1.

HKMECs were characterized in human kidney peritubular microvessels in vivo. (A and B) Histologic images of (A) human fetal and (B) adult kidney tissues containing (A.1 and B.1) established nephron structures. In the interstitial stroma, microvascular endothelial cells strongly expressed (A.2 and B.2) CD31 (red) and (A.3 and B.3) PV1 (red) but not (A.3 and B.3) vWF (green). Blue indicates nuclei. (C and D) From flow cytometric analysis, roughly 1.67% and 0.77% of the total cell population are endothelial cells (CD31+CD45−) in (C) fetal and (D) adult kidneys, respectively. G, glomeruli; S, stroma; T, tubule. Scale bar, 50 μm.

The endothelial cell population (CD31+CD45−) in human kidneys was identified with flow cytometric analysis and accounted for 3.1±1.5% of cells in fetal kidney tissues (Figure 1C) and 2.2±2.1% of cells in adults (Figure 1D). The direct sorting of this population from flow cytometry, however, resulted in limited cell survival, lack of attachment, and impurity when cultured. Our modified enrichment protocol (Figure 2A) showed that depletion of epithelial cells and supplementing with a high concentration of vascular endothelial growth factor (VEGF) in culture media were critical for enhancing endothelial cell growth (Figure 2B) and achieving purified HKMECs (VE Cadherin+/CD31+ CD45− PDGFRβ−) in large quantities.

Figure 2.
Figure 2.

HKMECs were isolated, purified, and characterized in vitro. (A) Summarized procedure for HKMEC enrichment. (B) Flow cytometric analysis of a single-cell suspension of isolated kidney cells after 5 days of culture indicating the proportion of endothelial cells in three distinct culture conditions: without VEGF-A (top panel), with 40 ng/ml VEGF-A (middle panel), and with VEGF-A and prior depletion of epithelial cells from the single-cell suspension (bottom panel). (C) RT-PCR confirmed the endothelial cell expression of PECAM, vWF, VE Cadherin, VEGFR2, TIE2, PDGF-BB, and the microvascular markers CD146 and ROBO4 and the absence of CD45 and E Cadherin. ECs: HKMECs; M: DNA marker; NC: Negative control. (D and E) Isolated cultured HKMECs show uniform morphology with purity >98%, and strongly express (D.1 and E.1) CD31, (D.2 and E.2) PV1, and (D.3 and E.3) VE Cadherin but not (D.4 and E.4) vWF in both (D) fetal and (E) adult kidneys. Scale bar, 50 μm.

Isolated HKMECs showed expression of typical endothelial transcripts by RT-PCR, and expressed genes include PECAM, VE Cadherin, VEGFR2, TIE2, vWF, and PDGF-BB as well as genes restricted to microvascular ECs, including ROBO4 and CD146 (Figure 2C, Supplemental Table 1). The purity was also verified by showing that these cells lack CD45 and E Cadherin expression. Isolated HKMECs from both fetal and adult tissue formed sheets in two-dimensional (2D) culture (Figure 2, D and E) with consistent expression of CD31 (Figure 2, D.1 and E.1) and VE Cadherin (Figure 2, D.3 and E.3) at cell-cell contacts. Expression of Claudin-5 was also found near the junctions between adjacent cells (Supplemental Figure 2); however, this expression followed a sawtooth distribution typically associated with the lack of tight junctions.20 Both fetal and adult endothelial cells showed abundant PV1 (Figure 2, D.2 and E.2) and low vWF expression (Figure 2, D.4 and E.4), respectively. This expression was consistent between the tissue sources, with no apparent difference in the HKMECs with respect to their morphology and surface markers. The consistent PV1 expression was indicative of a peritubular microvascular endothelial cell phenotype. In addition, these HKMECs expressed consistent VEGFR2 throughout the cell (Supplemental Figure 2).

HKMECs Are Highly Tubulogenic but Not Angiogenic

To understand the functional characteristics of HKMECs, we evaluated their capacity to self-assemble into complex tubular branching structures in 48 hours using a tubulogenic assay as previously described.21,22 When exposed to VEGF at 40 ng/ml, HKMECs showed a high degree of tubulogenic activity evident from extensive complex 3D structures with connected networks (Figure 3, A and B). These networks formed an enclosed lumen with an average diameter of roughly 25 μm (Figure 3A, arrows). Each connected segment was highly 3D and composed of hundreds of cells. In contrast, at identical culture conditions, HUVECs formed vacuoles, with isolated tube-like structures comprised of <10 cells. These structures were attenuated and poorly branched, and no complex networks were formed (Figure 3, C and D). The average vessel diameter was approximately 10 μm, significantly smaller (less than one half) than that found in the HKMEC-formed vascular networks (Figure 3E). Additionally, HKMEC-formed vessel networks displayed a vessel density nearly fourfold higher than vessel networks formed with HUVECs (Figure 3F).

Figure 3.
Figure 3.

HKMECs had different tubulogenic and angiogenic characteristics compared with HUVECs. (A and C) Confocal images of vascular tube formation (tubulogenesis assay) for (A) HKMECs and (C) HUVECs: xy plane (left panel) and (A.1, A.2, C.1, and C.2) two yz cross–sectional planes at the dashed lines. Arrows indicate the enclosed lumen. Scale bar, 50 μm. (B and D) 3D reconstruction of vessel tubes in A and C with a 200-μm depth in the z direction, showing the (B) connected microvascular network formed by HKMECs and (D) disconnected tubes formed by HUVECs. (E) Quantification of vessel diameters indicated that HKMECs formed connective networks with an average vessel diameter of approximately 25 μm, significantly larger than in HUVEC networks (approximately 10 μm). ***P<0.001. (F) Quantification of vessel density within HKMEC networks was around fourfold higher than that in HUVEC networks. (G and H) Confocal images of (G) HKMEC and (H) HUVEC monolayers remaining on the surface of a 2 mg/ml collagen gel in the xy plane (left panel) and yz cross-sections at the dashed lines (right panel) after 72 hours of culture. HUVECs readily sprouted into matrix, whereas HKMECs did not. (I) Quantification of the number of sprouts per area for HKMECs and HUVECs.

In a separate experiment, we tested the angiogenic potential of HKMECs compared with HUVECs (Figure 3, G and H). HKMECs formed a confluent monolayer that was quiescent and devoid of any sprouting in the presence of 40 ng/ml VEGF. In contrast, HUVECs readily formed tip cells and invaded the collagen matrix in response to VEGF. Quantification of the invasion frequency revealed a significantly higher number of sprouts per area in HUVECs compared with HKMECs (Figure 3I).

Molecular Characteristics of HKMECs

To gain a better understanding of the molecular characteristics of HKMECs, we compared gene expression by microarray analysis of HKMECs and HUVECs cultured under the same conditions. We identified 1748 significantly differentially expressed genes between HKMECs and HUVECs using criteria of 1.5-fold difference and a 0.05 false discovery rate. In HKMECs, 964 genes are upregulated, whereas 784 genes are downregulated compared with HUVECs. These data are summarized on the basis of fold difference using a list of representative genes (Table 1). Upregulation of genes, including CD34, PV1, ANGPT2, CXCL12, JAM2, and VEGFR2, and downregulation of genes, such as MMP1 and RGS5, were verified by real-time PCR (Figure 4A). In particular, the expression of structural protein PV1 was >10-fold higher in HKMECs than in HUVECs, consistent with the abundance of fenestrae observed when evaluating cell morphology (Figure 2D). ANGPT2 is known to be associated with vessel instability,23 and its upregulation in HKMECs may be correlated to the susceptibility of microvessels within the kidney to injury. The significant downregulation of RGS524 and MMP125 and upregulation of DLL426 support a lack of angiogenic potential for HKMECs, which is consistent with the lack of sprouting seen in response to VEGF (Figure 3, G–I).

View this table:
Table 1.

Selected genes that were significantly upregulated and downregulated in HKMECs compared with HUVECs in microarray studies

Figure 4.
Figure 4.

Molecular signatures were intrinsically different between HKMEC and HUVECs. (A) Real–time quantitative PCR on selective genes (bold) verified similar trends in downregulated genes MMP2 and RGS5 and upregulated genes CD34, ANGPT2, CXCL12, DLL4, JAM2, KDR, PDGFB, and PV1. (B) Ingenuity pathway analysis for angiogenesis function with significantly changed gene expression comparing HKMECs versus HUVECs. Red indicates upregulation, and blue indicates downregulation. (C) Heat map of canonical pathway analysis of HKMECs compared with HUVECs showing the most significantly differentiated genes that belong to specific ontologies.

To better understand the regulation of genes involved in angiogenesis and tubulogenesis functions, we further analyzed differential gene expression using Ingenuity Pathways Analysis. There were 62 significantly upregulated and 116 significantly downregulated genes that pertain to angiogenesis in HKMECs compared with HUVECs (Figure 4B). Within tubulogenesis function, 69 genes were significantly upregulated, and 62 genes were significantly downregulated (Supplemental Figure 3). The upstream analysis revealed that the transcription factors TP53 and SMAD3 were activated in HKMECs, indicating the inhibition of angiogenesis and an upregulation of TGFβ signaling. Furthermore, transcription factors, such as MYC and CCND1, were inhibited in HKMECs, indicating their inhibited cell growth and proliferation potential compared with HUVECs. Through canonical pathway analysis, we also observed 12 of a total of 407 canonical pathways that were significantly upregulated in HKMECs compared with HUVECs (with z score >2) (Figure 4C). These upregulated pathways included Tec Kinase signaling, Integrin signaling, Phospholipase C signaling, thrombin signaling, the role of NFAT, Gαq signaling, NGF signaling, NF-κB signaling, and the Wnt/Ca+ pathway. These data support the hypothesis that isolated HKMECs maintain intrinsic properties that are significantly distinct from HUVECs, despite identical culture conditions for >5 days.

These molecular signatures suggest that HKMECs intrinsically lack regenerative growth capabilities and angiogenic potential, which may be implicated in their susceptibility to and induction in kidney-specific injury.

HKMECs Form Stable Microvessels and Adapt to Flow

To recapitulate the physiologic conditions found in vivo, we engineered 3D microvascular networks in collagen gel using lithographic processes that we described previously.18 HKMECs were seeded in the lumen of this microphysiologic system (MPS) and cultured under gravity-driven flow for 3–14 days to achieve a confluent layer of endothelium on luminal walls (Figure 5, A–C). This endothelium expressed CD31 and VE Cadherin at regions of cell-cell contact (Figure 5, D and E). vWF granules became more abundant in HKMECs comprising an endothelium along microvessels compared with those cultured under static 2D conditions (Figures 2, D and E and 5F). A similar increase of vWF was also observed in HUVECs cultured in a 3D microenvironment, which was also greater than HKMECs when cultured in the same manner (Figure 5F). In addition, HKMECs were more sensitive to shear stress than HUVECs, determined by enhanced alignment with the direction of flow in microvessels (Figure 5G). The same shear-sensitive effect was also consistently found in mouse KMECs (Supplemental Figure 4). The recapitulated kidney microvessels also formed a barrier to the transfer of solutes from the lumen into the matrix. A basement membrane consisting predominantly of collagen 4 was found to be deposited by HKMECs along the vessel walls (Figure 5H).

Figure 5.
Figure 5.

Human kidney peritubular microvessels were reconstructed in vitro. (A and B) Schematic diagram of 3D MPS set up with (A) 3D views and (B) cross-sections. (C) Example of kidney microvessel networks generated in a 3D MPS. Red indicates CD31, and blue indicates nuclei. (D) z-Stack projection of confocal image of engineered human kidney microvessel in xy plane (left panel) and cross-sectional yz plane at the dashed line (right panel). Red indicates CD31, green indicates vWF, and blue indicates nuclei. Scale bar, 50 μm. (E) z-Stack projection of confocal image of human kidney microvessels at a junction of the network. Red indicates VE Cadherin, and blue indicates nuclei. Scale bar, 25 μm. (F) Quantification of the amount of vWF per area for both HKMECs and HUVECs and in both 2D static culture and 3D flow-based microvessel cultures. (G) Quantification of SI (4πA/P2) for HKMECs in 3D microvessels and 2D static cultures and HUVECs in 3D microvessels. (H) Immunofluorescence image of a cryosectioned HKMEC vessel network (thickness of 7 μm). Red indicates CD31, green indicates collagen IV, and blue indicates nuclei. Scale bar, 100 μm. (I) Fluorescence image of 40-kD FITC-dextran perfused through the HKMECs after 1 minute of perfusion. Arrows show good barrier, and stars show focal leakage. Scale bar, 100 μm.

We further measured the vessel permeability or molecular sieving effect of a large molecule, 40-kD FITC-dextran, for both HKMECs and HUVECs microvessels. Perfusion of the molecules was driven by the hydrostatic pressure drop through the vessel. The oncotic pressure drop across the vessel wall was approximately zero given the same culture media, and thus, protein content was saturated on the abluminal and luminal sides of the endothelium. Focal permeation of 40-kD FITC-dextran occurred along the endothelium (Figure 5I), leading to an increased spatially averaged permeability, which represents the increased convective flux of the molecules across the endothelium. We used the distribution of fluorescence intensity during the transient flow of dextran to estimate the averaged permeability coefficient of the kidney endothelium to be K=0.16±0.06 μm s−1, including the focal leaky regions. These permeability coefficients were approximately five times higher than that observed in HUVEC-formed microvessels.

The reconstructed kidney microvessels strongly express the plasmalemma protein, PV1, throughout the endothelium. The expression of PV1 was ubiquitously distributed throughout the cytoskeleton in the cell periphery (Figure 6A), whereas PV1 was not detectable in HUVECs vessels (Supplemental Figure 4). Ultrastructural examination of these engineered kidney microvessels substantiated the immunohistochemistry, because HKMEC endothelium displayed abundant closed fenestratae along the microvessel walls (Figure 5, B–D). HKMECs formed a thin layer along the luminal wall and appeared to be polarized between the lumen and the matrix. Junctions formed at focal contacts of lining HKMECs (red arrows in Figure 5, B and C), numerous fenestrated diaphragms were clearly visible, and all contained a central knob (black arrows in Figure 5, B and D). The average size of fenestrae was 59±11 nm, which is similar to the size of fenestrae in kidney microvessels in vivo (approximately 62–68 nm).27 The ultrastructure of HUVEC-formed microvessels revealed a continuous endothelium lacking fenestrae but rich in caveolae, which may be a result of the high VEGF culture conditions (Supplemental Figure 5, B and C). The presence of abundant fenestrae in HKMECs microvessels increases the total area permissible to transport molecules moving across the vessel wall, which likely contributed to the increase of vessel permeability compared with HUVEC microvessels.

Figure 6.
Figure 6.

Reconstructed human kidney peritubular microvessels were fenestrated in vitro. (A) z-Stack projection of confocal image of engineered human kidney microvessel at a junction of vessel network (left panel) and in a zoomed view (right panel). Red indicates F-actin, green indicates PV1, and blue indicates nuclei. (B–D) Transmission electron microscopy reveals the ultrastructure of HKMECs microvessels containing proper junctions (red arrows) formed at cell-cell contacts between (B and C) two adjacent cells C1 and C2 and (B and D) numerous fenestrae (black arrows) throughout the peripheral regions.

Discussion

There is currently great interest in developing microfluidic organs on chips, including the vasculature, which carry the potential to create predictive models of human disease and interrogate the organ-specific physiologic and pathophysiologic responses to drugs and environmental toxins.28 Within the kidney is a complex interplay between the tubular and vascular units comprising filtration and resorption. Specifically, the peritubular microvessels are highly susceptible to rarefaction after exposure to toxins, xenobiotics, or injury. Characterizing the relevant cells and recreating a representative microvasculature are crucial for the understanding of drug transport, drug-induced nephrotoxicity, and capillary rarefaction in progressive kidney diseases. Although comparatively more attention has been focused on the glomerulus, the peritubular capillaries represent an understudied and important physiologic system to investigate given their critical importance. In this work, we developed methods to isolate, purify, and culture HKMECs. The success of the isolation and subsequent culture were particularly dependent on VEGF concentration and epithelial cell depletion before enrichment and flow cytometric sorting. HKMECs exhibited standard endothelial surface markers, adherens junctions, and displayed an endothelial cell gene expression pattern. In addition, they maintained kidney microvascular-specific structures, including closed fenestrae, characterized by the plasmalemma protein, PV1. HKMECs exhibited marked differences from HUVECs in phenotype, gene expression pattern, and functional assays. Microarray analysis revealed that the intrinsic transcriptional signatures of HKMECs were different from those in HUVECs, with more tubulogenic but less angiogenic potential.

We then reconstructed a 3D human kidney microvascular network to assess the structure and function of human kidney peritubular microvessels under flow. We showed that the HKMECs in 3D MPS altered their alignment along flow-directed pathways and increased granular vWF compared with the same cells in static 2D culture conditions. HKMECs formed comprehensive barriers to large dextran molecules and deposited physiologically observed basement membrane proteins on the vessel walls. Our data indicate that HKMECs in a 3D flow-directed MPS exhibit features highly representative of the kidney microvasculature in vivo, while simultaneously recapitulating standard microvascular properties. Future studies will be required to further define the unique transport properties of these fenestrated endothelia for different size molecules and under varying flow and pressure conditions.

Our data show that the use of organ-specific microvascular endothelial cells in a 3D MPS can provide unique insights into the properties of the peritubular microvasculature, which appear to closely approximate human anatomy and physiology. Because the loss of kidney peritubular capillaries is common to kidney disease progression from almost all inciting mechanisms of injury, this approach provides a high content system for identifying new therapeutic approaches to preventing human kidney failure. In addition, this approach opens new possibilities for mechanistic studies of heterogeneity in interindividual susceptibility to kidney injury.

Concise Methods

Kidney Tissue

Adult kidney tissues were obtained from normal renal cortex of nephrectomies performed for renal masses or transitional cell carcinoma. Fetal human kidneys were obtained after voluntary pregnancy interruptions performed at the University of Washington Medical Center in compliance with Institutional Review Board protocol (IRB447773EA). Informed consents for the use of fetal tissues were obtained from patients.

Endothelial Cell Isolation, Enrichment, and Culture

Kidney tissues were processed mechanically followed by enzymatic dissociation and sieving through a cell strainer to remove glomeruli and large vessels to obtain a single-cell suspension. The removal of glomeruli was confirmed by microscopic examination, showing that almost all glomeruli are present in the cell strainer, and no glomeruli were found in the single-cell suspension. To obtain a sufficient number of cells with high purity, epithelial cells were depleted from the single-cell suspension using a magnetic immunoaffinity column, and the rest of cells was cultured on a gelatin matrix for 72–96 hours in customized endothelial cell proliferation media with high concentration of VEGF. This expanded cell population was then resuspended and subjected to flow-cytometric sorting for a VE Cadherin+/CD31+ CD45− PDGFRβ− subpopulation. After sorting, the purified cells were immediately cultured in conditions favoring endothelial cell growth for up to five passages. The cells were further characterized by morphology, immunostaining, and PCR to verify the purity and for confirmation of the absence of glomeruli endothelial cell contamination. Details are described in Supplemental Material.

Tubulogenesis and Angiogenesis Assays

For tubulogenesis function,22 endothelial cells were mixed with collagen type 1 solution to reach gel density of 2 mg/ml and cell density of 2×106 cells per ml. pipetted into customized 5-mm-diameter wells of precast polydimethylsiloxane (PDMS) dishes, and allowed to gel at 37 °C and 5% CO2 for 30 minutes. Cultures were then kept in endothelial growth media with 40 ng/ml VEGF and allowed to assemble over time. The collagen invasion (angiogenesis) assay was performed as described.22,29 Briefly, 5-mm-diameter wells of PDMS dishes were filled with rat tail collagen to reach a flat surface and allowed to polymerize for 20 minute at 37 °C. After polymerization, endothelial cells were plated on top of each gel at density of 400 cells per mm2 and cultured with complete EBM-2 endothelial medium. The next day, the cultures were rinsed twice with EBM-2 serum free and incubated with 40 ng/ml VEGF in endothelial complete medium. After 48–72 hours, cultures were fixed in 4% formaldehyde solution, stained, and imaged under microscope.

Microvessel Fabrication and Culture

A neutralized liquid collagen solution was prepared at 7.5 mg/ml from 15 mg/ml collagen stock solution for microvessel fabrication as described previously.18 Briefly, collagen was injected into a PEI/glutaraldehyde–treated Plexiglas housing top half that formed a cavity along with an oxygen plasma–sterilized PDMS stamp, thereby forming a negative impression of a microvessel network. Inlet and outlet ports were formed by inserting stainless steel dowel pins before injecting the collagen. The Plexiglas bottom half consisted of a flat layer of collagen compressed by a flat PDMS surface that was gelled on top of a standard coverslip. The collagen was allowed to gel for 30 minutes at 37 °C. After gelation, the PDMS stamp on the top piece was removed, revealing an open microfluidic network. The PDMS on the flat layer of collagen on the Plexiglas bottom half was used to seal the microvessel network. Culture medium was then added to the reservoirs and incubated for 2 hours before cell seeding. To seed the vessels, 10-μl injections of HKMECs or HUVECs from P1 to P5 at a density of 5×106 cells per ml were delivered to the inlet of an aspirated channel. Flow was allowed by replenishing media in the inlet reservoir twice per day.

Imaging and Quantifications

The immunofluorescence images of the cells and tissue sections on slides, 3D gels, or intact microvessels in situ were taken using a Nikon A1R Confocal Microscope (Nikon, Tokyo, Japan). Image stacks were accumulated with a z step between successive optical slices of approximately 2 μm. Cross-sections, projections, and 3D reconstructions were generated from z stacks of images using ImageJ software with orthogonal projection, z projection, and 3D viewer. The ultrastructured images were acquired on ultrathin after–processed vessel sections (70 nm) using a JEOL JEM-1400 Transmission Electron Microscope (JEOL Ltd.) with a Gatan Ultrascan 1000XP Camera (Gatan, Inc., Pleasanton, CA).

Quantification of Vessel Diameter, Density, and Sprout Frequencies

For tubulogenesis assay, image stacks of vascular network in 3D gels were loaded in ImageJ, and the channels were split to obtain the 3D stack of vessel walls. An intensity threshold was selected to filter the single-channel image stack, so that the vessel walls were preserved and well defined. The enclosed vessel lumen was then filled, and a binary stack output was generated after the filtering and hole-filling functions. The vessel density was calculated by summing the nonzero areas in the stacks and normalizing with the total stack volume. The vessel diameter of each stack image was measured for three well defined connected regions along the direction perpendicular to the axial direction of the tubes. Two image stacks were analyzed for each replicate. For angiogenesis assays, the number of sprouts per imaging field was counted and converted to that per mm2. Three images were counted to reach an averaged value for each replicate. For both assays, three to five replicates were processed for analysis.

Quantification of vWF Expression

In 2D and 3D experiments, vWF expression was taken by setting a binary threshold on the vWF immunofluorescence channel. The area of vWF corresponded to the number of pixels counted in the image using ImageJ. This number was divided by the total number of pixels in the region of interest and then, multiplied by 100 to give the percentage of vWF present in a given area of cells. Three images were analyzed for three separate devices. Data for each device were then averaged and used to calculate SEM with n=3.

Quantification of Shape Index of Cells in Microvessels

Shape index (SI) was calculated according to the equation SI=4πA/P2. Each cell was traced and measured using ImageJ to output its perimeter and area. Nine cells were randomly chosen and calculated for each of three separate devices. The SI for each device was averaged and then, used to calculate SEM with n=3.

Quantification of Vessel Permeability

The permeability of kidney microvessels was measured by delivering 40-kD FITC-dextran through the endothelialized microchannel at a continuous flow rate of approximately 5 μl min−1. Fluorescence images were acquired sequentially, and image analysis was carried out in MATLAB with detailed mathematics described previously18 to obtain the permeability of the vessel wall. Six regions of interest were randomly chosen for each image to obtain an averaged value for permeability coefficient. The fluorescence intensity at the center of interchannel space between endothelialized channels was measured with time to extract the permeability coefficient of the vessel wall.

Statistical Analyses

For all quantitative measurements, the entire population was used to calculate statistical significance, whereas mean values with n=3 were used to calculate SEM and graphical confidence intervals. An F test was used to determine variance equality. A two–tailed t test was then performed on each group of interest. On each graph, error bars represent ±2 SEM (a 95% confidence interval). For P values <0.05, a single asterisk over a solid line was used; for P values <0.01, two asterisks were used, and for P values <0.001, three asterisks were used.

Detailed methods of cell isolation, immunostaining, and molecular characterization are described in Supplemental Material.

Disclosures

None.

Acknowledgments

We acknowledge the Microfabrication Facility and the Lynn and Mike Garvey Imaging Core at the University of Washington and the Electron Microscope Facility and Genomics Department at the Fred Hutchinson Cancer Research Institute. We thank Drs. Ed Kelly, Kenneth Thummel, and Danny Shen for helpful discussions.

This project was supported by National Institutes of Health Grants UH2/UH3 TR000504 (to J.H.), P30-ES07033 (to J.W.M. and T.K.B.), DK93493 (to J.S.D.), and DP2DK102258 (to Y.Z.) and American Heart Association Grants 12040023 (to J.S.D.) and 12SDG9230006 (to Y.Z.).

Footnotes

References

  1. ↵
    1. Himmelfarb J,
    2. Ikizler TA
    : Acute kidney injury: Changing lexicography, definitions, and epidemiology. Kidney Int 71: 971–976, 2007pmid:17396117
    OpenUrlCrossRefPubMed
  2. ↵
    1. Coresh J,
    2. Selvin E,
    3. Stevens LA,
    4. Manzi J,
    5. Kusek JW,
    6. Eggers P,
    7. Van Lente F,
    8. Levey AS
    : Prevalence of chronic kidney disease in the United States. JAMA 298: 2038–2047, 2007pmid:17986697
    OpenUrlCrossRefPubMed
  3. ↵
    1. Chawla LS,
    2. Eggers PW,
    3. Star RA,
    4. Kimmel PL
    : Acute kidney injury and chronic kidney disease as interconnected syndromes. N Engl J Med 371: 58–66, 2014pmid:24988558
    OpenUrlCrossRefPubMed
  4. ↵
    1. Brenner BM,
    2. Falchuk KH,
    3. Keimowitz RI,
    4. Berliner RW
    : The relationship between peritubular capillary protein concentration and fluid reabsorption by the renal proximal tubule. J Clin Invest 48: 1519–1531, 1969
    OpenUrlCrossRefPubMed
    1. LeBrie SJ
    : Renal peritubular capillary permeability to macromolecules. Am J Physiol 213: 1225–1232, 1967
    OpenUrlPubMed
    1. Ojteg G,
    2. Nygren K,
    3. Wolgast M
    : Permeability of renal capillaries. II. Transport of neutral and charged protein molecular probes. Acta Physiol Scand 129: 287–294, 1987pmid:3577815
    OpenUrlCrossRefPubMed
  7. ↵
    1. Källskog O,
    2. Wolgast M
    : Driving forces over the peritubular capillary membrane in the rat kidney during antidiuresis and saline expansion. Acta Physiol Scand 89: 116–125, 1973pmid:4761515
    OpenUrlCrossRefPubMed
  8. ↵
    1. Jen K-Y,
    2. Haragsim L,
    3. Laszik ZG
    : Kidney microvasculature in health and disease. Contrib Nephrol 169: 51–72, 2011pmid:21252511
    OpenUrlCrossRefPubMed
  9. ↵
    1. Lin SL,
    2. Chang FC,
    3. Schrimpf C,
    4. Chen YT,
    5. Wu CF,
    6. Wu VC,
    7. Chiang WC,
    8. Kuhnert F,
    9. Kuo CJ,
    10. Chen YM,
    11. Wu KD,
    12. Tsai TJ,
    13. Duffield JS
    : Targeting endothelium-pericyte cross talk by inhibiting VEGF receptor signaling attenuates kidney microvascular rarefaction and fibrosis. Am J Pathol 178: 911–923, 2011pmid:21281822
    OpenUrlCrossRefPubMed
  10. ↵
    1. Basile DP
    : The endothelial cell in ischemic acute kidney injury: Implications for acute and chronic function. Kidney Int 72: 151–156, 2007pmid:17495858
    OpenUrlCrossRefPubMed
  11. ↵
    1. Basile DP
    : Rarefaction of peritubular capillaries following ischemic acute renal failure: A potential factor predisposing to progressive nephropathy. Curr Opin Nephrol Hypertens 13: 1–7, 2004pmid:15090853
    OpenUrlCrossRefPubMed
  12. ↵
    1. Muczynski KA,
    2. Ekle DM,
    3. Coder DM,
    4. Anderson SK
    : Normal human kidney HLA-DR-expressing renal microvascular endothelial cells: Characterization, isolation, and regulation of MHC class II expression. J Am Soc Nephrol 14: 1336–1348, 2003pmid:12707403
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Ballermann BJ
    : Regulation of bovine glomerular endothelial cell growth in vitro. Am J Physiol 256: C182–C189, 1989pmid:2536227
    OpenUrlPubMed
  14. ↵
    1. Marsden PA,
    2. Brock TA,
    3. Ballermann BJ
    : Glomerular endothelial cells respond to calcium-mobilizing agonists with release of EDRF. Am J Physiol 258: F1295–F1303, 1990pmid:2159719
    OpenUrlPubMed
  15. ↵
    1. Kida Y,
    2. Ieronimakis N,
    3. Schrimpf C,
    4. Reyes M,
    5. Duffield JS
    : EphrinB2 reverse signaling protects against capillary rarefaction and fibrosis after kidney injury. J Am Soc Nephrol 24: 559–572, 2013pmid:23492730
    OpenUrlAbstract/FREE Full Text
  16. ↵
    1. Taguchi A,
    2. Kaku Y,
    3. Ohmori T,
    4. Sharmin S,
    5. Ogawa M,
    6. Sasaki H,
    7. Nishinakamura R
    : Redefining the in vivo origin of metanephric nephron progenitors enables generation of complex kidney structures from pluripotent stem cells. Cell Stem Cell 14: 53–67, 2014pmid:24332837
    OpenUrlCrossRefPubMed
  17. ↵
    1. Abrahamson DR,
    2. Robert B,
    3. Hyink DP,
    4. St John PL,
    5. Daniel TO
    : Origins and formation of microvasculature in the developing kidney. Kidney Int Suppl 67: S7–S11, 1998pmid:9736245
    OpenUrlCrossRefPubMed
  18. ↵
    1. Zheng Y,
    2. Chen J,
    3. Craven M,
    4. Choi NW,
    5. Totorica S,
    6. Diaz-Santana A,
    7. Kermani P,
    8. Hempstead B,
    9. Fischbach-Teschl C,
    10. López JA,
    11. Stroock AD
    : In vitro microvessels for the study of angiogenesis and thrombosis. Proc Natl Acad Sci U S A 109: 9342–9347, 2012pmid:22645376
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Ichimura K,
    2. Stan RV,
    3. Kurihara H,
    4. Sakai T
    : Glomerular endothelial cells form diaphragms during development and pathologic conditions. J Am Soc Nephrol 19: 1463–1471, 2008pmid:18480313
    OpenUrlAbstract/FREE Full Text
  20. ↵
    1. Kluger MS,
    2. Clark PR,
    3. Tellides G,
    4. Gerke V,
    5. Pober JS
    : Claudin-5 controls intercellular barriers of human dermal microvascular but not human umbilical vein endothelial cells. Arterioscler Thromb Vasc Biol 33: 489–500, 2013
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Koh W,
    2. Stratman AN,
    3. Sacharidou A,
    4. Davis GE
    : In vitro three dimensional collagen matrix models of endothelial lumen formation during vasculogenesis and angiogenesis . Methods Enzymol 443: 83–101, 2008
    OpenUrlCrossRefPubMed
  22. ↵
    1. Cross VL,
    2. Zheng Y,
    3. Won Choi N,
    4. Verbridge SS,
    5. Sutermaster BA,
    6. Bonassar LJ,
    7. Fischbach C,
    8. Stroock AD
    : Dense type I collagen matrices that support cellular remodeling and microfabrication for studies of tumor angiogenesis and vasculogenesis in vitro. Biomaterials 31: 8596–8607, 2010pmid:20727585
    OpenUrlCrossRefPubMed
  23. ↵
    1. Maisonpierre PC,
    2. Suri C,
    3. Jones PF,
    4. Bartunkova S,
    5. Wiegand SJ,
    6. Radziejewski C,
    7. Compton D,
    8. McClain J,
    9. Aldrich TH,
    10. Papadopoulos N,
    11. Daly TJ,
    12. Davis S,
    13. Sato TN,
    14. Yancopoulos GD
    : Angiopoietin-2, a natural antagonist for Tie2 that disrupts in vivo angiogenesis. Science 277: 55–60, 1997pmid:9204896
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Jin Y,
    2. An X,
    3. Ye Z,
    4. Cully B,
    5. Wu J,
    6. Li J
    : RGS5, a hypoxia-inducible apoptotic stimulator in endothelial cells. J Biol Chem 284: 23436–23443, 2009pmid:19564336
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Moses MA
    : The regulation of neovascularization of matrix metalloproteinases and their inhibitors. Stem Cells 15: 180–189, 1997pmid:9170209
    OpenUrlCrossRefPubMed
  26. ↵
    1. Leslie JD,
    2. Ariza-McNaughton L,
    3. Bermange AL,
    4. McAdow R,
    5. Johnson SL,
    6. Lewis J
    : Endothelial signalling by the Notch ligand Delta-like 4 restricts angiogenesis. Development 134: 839–844, 2007pmid:17251261
    OpenUrlAbstract/FREE Full Text
  27. ↵
    1. Stan RV
    : Endothelial stomatal and fenestral diaphragms in normal vessels and angiogenesis. J Cell Mol Med 11: 621–643, 2007pmid:17760829
    OpenUrlCrossRefPubMed
  28. ↵
    1. Bhatia SN,
    2. Ingber DE
    : Microfluidic organs-on-chips. Nat Biotechnol 32: 760–772, 2014pmid:25093883
    OpenUrlCrossRefPubMed
  29. ↵
    1. Ligresti G,
    2. Aplin AC,
    3. Dunn BE,
    4. Morishita A,
    5. Nicosia RF
    : The acute phase reactant orosomucoid-1 is a bimodal regulator of angiogenesis with time- and context-dependent inhibitory and stimulatory properties. PLoS One 7: e41387, 2012
    OpenUrlCrossRefPubMed
Back to top

In this issue

View Selected Citations (0)
Print
Download PDF
Email Article
Citation Tools
Request Permissions
A Novel Three–Dimensional Human Peritubular Microvascular System
Giovanni Ligresti, Ryan J. Nagao, Jun Xue, Yoon Jung Choi, Jin Xu, Shuyu Ren, Takahide Aburatani, Susan K. Anderson, James W. MacDonald, Theo K. Bammler, Stephen M. Schwartz, Kimberly A. Muczynski, Jeremy S. Duffield, Jonathan Himmelfarb, Ying Zheng
JASN Aug 2016, 27 (8) 2370-2381; DOI: 10.1681/ASN.2015070747
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

Keywords