Endothelial–Pericyte Interactions Regulate Angiogenesis Via VEGFR2 Signaling During Retinal Development and Disease

Abstract

Purpose

Endothelial–pericyte interaction disruption causes vascular dropout and pathological angiogenesis, severely impacting visual function in ocular microvascular diseases. This study examines VEGF receptor 2 (VEGFR2) signaling in endothelial–pericyte interactions, highlighting VEGFR2 as a potential therapeutic target for promoting pericyte coverage and decreasing vascular leakage in diseased retinas.

Method

Cell–cell interactions with VEGFR2 signaling were assessed using isogenic endothelial cells and pericytes from induced pluripotent stem cells. We investigated changes in VEGFR2 signaling resulting from endothelial–pericyte interactions using quantitative Reverse Transcription PCR, western blot analysis, immunofluorescence staining, migration assays, permeability assays, transendothelial electrical resistance measurements, flow cytometry, and three-dimensional collagen gel vascular networks. We validated VEGFR2 as a therapeutic target via intravitreal injection in the oxygen-induced retinopathy mouse model. Treatment effects were evaluated using western blot analysis, immunofluorescence staining, and an FITC–dextran permeability assay to assess protein expression, pericyte recruitment, and retinal vascular function in response to VEGFR2 modulation.

Results

We demonstrate that direct endothelial-pericyte contact, mediated by N-cadherin, downregulates phosphorylated VEGFR2 in endothelial cells, thereby enhancing pericyte migration and promoting endothelial cell barrier function. Intravitreal injection of a VEGFR2 inhibitor in mouse models of the developing retina and oxygen-induced retinopathy increased pericyte recruitment and decreased vascular leakage. The VEGFR2 inhibitor further rescued ischemic retinopathy by enhancing vascularization and tissue growth.

Conclusions

Our findings uncover a novel mechanism by which VEGFR2 signaling is regulated through endothelial–pericyte interactions, promoting pericyte migration and strengthening endothelial barrier function. These results suggest a pathway that could be harnessed to support the growth of functional and mature microvasculature in ocular microvascular diseases and tissue regeneration overall.

The vascular system is responsible for delivering nutrients and oxygen to the body as well as facilitating communication between organs. It develops from the differentiation of endothelial cells (ECs) from angioblasts¹ and is the first functioning physiological system that forms during embryogenesis. After the formation of the initial vasculature, ECs then proliferate, migrate, and sprout to extend vascular networks to support organ function. In humans, the retinal vasculature extends from the optic nerve head, growing radially and penetrating through the retinal layers to form functional vascular networks.² The recruitment of mural cells, such as pericytes to the capillaries and microvasculature and smooth muscle cells to the larger vessels, results in matured and functional blood vessels that are capable of transporting oxygen and nutrients. Vascularization of injured, diseased, or engineered tissue is required for tissue healing and regeneration, and eventual homeostasis.^3–5

The vascular endothelial growth factor (VEGF)/VEGF receptor 2 (VEGFR2) signaling pathway is essential in forming organ vasculature.⁶ VEGFR2 is activated by VEGF-A, -C, and -D⁷ and plays an important role during angiogenesis.⁸ The establishment of ligand-receptor dimerization upon VEGF binding leads to the phosphorylation of the downstream tyrosine residues of VEGFR2 at the catalytic tyrosine kinase domain and the activation of signaling pathways that regulate EC function and vascular formation.⁹ Several phosphorylation sites are responsible for VEGFR2 activation and cell signaling regulation. The phosphorylation of tyrosine 951 promotes Akt activity to affect EC survival and permeability.^10,11 Additionally, phosphorylation of VEGFR2 at the surface of the tip cell mediates tip cell migration and increases glycolysis.^12–14 It has been demonstrated that cancer cells secrete angiopoietin-1 after VEGFR2 inhibition, leading to temporary pericyte recruitment and tumor vascular normalization.¹⁵ In retinopathies, VEGFR2 has been shown to regulate vascular permeability.^10,16 However, it remains unknown if and how VEGFR2 activity in ECs regulates their interactions with pericytes. Here, we sought to examine whether VEGFR2 in ECs modulates pericyte recruitment, thereby impacting endothelial barrier function during retinal development and retinopathy.

Pericyte contact with the capillaries and microvasculature is critical for ECs to maintain barrier function and vascular homeostasis.^17–19 Platelet-derived growth factor-BB (PDGF-BB) secreted by ECs has been shown to regulate pericyte recruitment, proliferation, and migration during vascular development.¹⁸ Pericytes guide vascular remodeling by migrating along sprouting ECs^20,21 and secreting extracellular matrix (ECM) degradation molecules.²² Pericytes also secrete extracellular matrix, which contributes to the vascular basement membrane to support organ function.^23,24 Therefore, for successful tissue regeneration or healing, the revascularization process must include pericytes. However, despite our current understanding of the initial pericyte recruitment, it remains unclear how pericytes respond and modulate the initial shift from angiogenesis to stabilization in ECs.

Diabetic retinopathy (DR) is a microvascular disease characterized by pericyte loss, increased vascular permeability, and EC death, followed by the growth of abnormal blood vessels and vision impairment.²⁵ The available treatment options for revascularizing a damaged retina is limited to intravitreal injections of antiangiogenic agents, corticosteroids, and nonsteroidal anti-inflammatory drugs.^26,27 VEGF is a key driver of DR disease progression²⁸ and is negatively correlated with pericyte recruitment.²⁹ Neutralizing excessive VEGF secretion is used to treat retinopathy, including U.S. Food and Drug Administration (FDA)–approved drugs such as aflibercept, faricimab, and ranibizumab. However, these drugs primarily sequester VEGF-A, preventing their binding to all VEGFRs and impacting a wide range of VEGF downstream signaling, potentially making them less effective while leading to unwanted side effects and therapeutic resistance.^30,31 Furthermore, not all patients respond effectively to these treatments, and methods to reverse vascular damage are yet to be developed.^32,33 Pericyte destruction is the first pathological event observed in mice with DR,³⁴ followed by the switching of ECs to an angiogenic phenotype.³⁵ Therefore, it is important to consider pericytes’ role in modulating angiogenesis through cellular crosstalk. However, the mechanisms of revascularizing damaged tissue or reestablishing EC–pericyte communication in diseases such as DR have remained largely unidentified.^36,37

Here, we set out to determine the role of pericytes in regulating VEGFR2 signaling on ECs, and subsequent vascular stabilization and maturation in both developing and diseased retina. Using isogenic ECs and pericytes derived from human induced pluripotent stem cells (hiPSCs), we showed decreased EC permeability along with upregulation of pericyte recruitment factors after VEGFR2 inhibition. We further examine the hypothesis that, after initial pericyte recruitment by PDGF-BB,³⁸ direct physical contact between pericytes and ECs is required to downregulate VEGFR2 activity in ECs. This finding identified the mechanistic role of pericyte contact in modulating VEGFR2 signaling.

Next, using hiPSC in vitro modeling and in vivo mouse models of the oxygen-induced retinopathy (OIR) and the healthy developing retina, we examine the hypothesis that the downregulation of VEGFR2 activity amplifies pericyte recruitment to the nascent vascular networks, leading to network stabilization. Using a specific VEGFR2 inhibitor, we demonstrate that downregulating VEGFR2 pY951 restores endothelial barrier function and vascular stability by enhancing pericyte recruitment to the developing and diseased retina. This result highlights the potential of blocking VEGFR2 activity to decrease pericyte loss in the treatment of injured and diseased tissue, as well as to vascularize tissue-engineered organs by stabilizing the developing vasculature through increased pericyte recruitment.

Methods

Maintenance and Differentiation of hiPSCs

C1-2 hiPSCs were cultured on a Vitronectin-coated plate in Essential 8 medium (Thermo Fisher Scientific, Waltham, MA, USA) with the media changed daily for 3 days. Endothelial differentiation was started with mesodermal induction when cells were 60 to 80% confluent by adding CHIR99021 (6 µM) (STEMCELL Technologies, Cambridge, MA, USA) to the Essential 6 medium (Thermo Fisher Scientific) with daily media changes. On day 2 of the mesodermal induction, cells were detached using TrypLE Express (Thermo Fisher Scientific) and seeded on a type I collagen coated plate at 2 × 10⁴ cells/cm² with codifferentiation media containing EC growth media (ECGM; PromoCell, Heidelberg, Germany) supplemented with 10 µM SB-431542 (Cayman Chemical Company, Ann Arbor, MI, USA), 50 ng/mL VEGF-A (PeproTech, Cranbury, NJ, USA), and 10 µM Y-27632 (Selleckchem, Houston, TX, USA). After 24 hours, the media was changed to ECGM supplemented with 10 µM SB-431542 (Cayman Chemical Company) and 50 ng/mL VEGF-A (PeproTech), following media change every other day for 6 days.

Isolation of hiPSCs to ECs and Pericytes

On day 8 of the differentiation, CD31⁺ cells were isolated with a magnetic-activated cell sorter (MACS; Miltenyi Biotech, Bergisch Gladbach, Germany). Cells were washed with dPBS (Thermo Fisher Scientific) once and detached using TrypLE Express. After centrifuging, cells were then resuspended in 100 µL MACS buffer containing 0.5 mM EDTA and 0.5% BSA in dPBS. We added 10 µL of phycoerythrin -conjugated anti-human CD31 (BD Biosciences) to the MACS buffer, and cells were incubated at 4°C for 10 minutes. After incubation, cells were washed with MACS buffer twice to remove unbound primary antibodies. Next, cells were incubated with 20 µL of anti-phycoerythrin microbeads (Miltenyi Biotec) and 80 µL of MACS buffer at 4°C for 15 minutes. Cells were washed once with MACS buffer and sorted using the MS MACS separation column (Miltenyi Biotec). The purified CD31⁺ cells were expanded in ECGM supplemented with 10 µM SB-431542 (Cayman Chemical Company), and 50 ng/mL VEGF-A (PeproTech) on a type I collagen-coated plate. CD31⁻ cells were expanded in Pericyte Medium (ScienCell Research Laboratories, Carlsbad, CA, USA) on a type I collagen-coated plate.

VEGFR2 Inhibition

For all two-dimensional in vitro inhibition studies, cells were cultured in ECGM (PromoCell) supplemented with 10 µM SB-431542 (Cayman Chemical Company) and 25 ng/mL VEGF-A (PeproTech). For all three-dimensional (3D) in vitro inhibition studies, cells were cultured in ECGM (PromoCell) supplemented with 50 ng/mL VEGF-A (PeproTech). ZM323881 (Selleckchem) was dissolved in DMSO and added to the cell culture media at 1 µM. Cells were incubated with the inhibitor for 0.5, 1.0, 12.0, 24.0, or 48.0 hours, as indicated. DMSO was used as a vehicle control.

N-Cadherin Inhibition

N-Cadherin inhibition was performed by adding 100 µM ADH-1 (MedChemExpress, Monmouth Junction, NJ, USA) to the EC–pericyte coculture for the final 24 hours of the 96-hour culture period.

Flow Cytometry Analysis

Cells were harvested for analysis using TrypLE (Invitrogen, Waltham, MA, USA) dissociation buffer and collected in 100 µL of 0.1% BSA. Cells were then incubated with primary antibody for 30 minutes on ice. Antibodies are detailed in Supplementary Table S1. Cells were washed three times with 0.1% BSA and passed through a 40-µm cell strainer. Flow analysis was conducted on a BD FACSCanto flow cytometer. Following manufacturer instructions, dead cell populations were gated out with forward-side scatter plots. All analyses were conducted using FlowJo software.

Immunofluorescence Staining and Imaging

Induced EC iECs), hiPSC-derived pericytes (iPericytes), and 3D constructs were fixed in 3.7% paraformaldehyde (Sigma-Aldrich, St., Louis, MO, USA) for 10 minutes or in ice-cold methanol for 5 minutes. Cells fixed with paraformaldehyde were permeabilized with 0.1% Triton X-100 (Sigma-Aldrich). Cells were washed and blocked in 1% BSA for 1 hour at room temperature or overnight at 4°C. Cells were incubated with primary antibodies (Supplementary Table S1) overnight at 4°C. The next day, cells were washed three times with 0.1% TWEEN 20 (Sigma-Aldrich) in PBS. Cells were incubated with secondary antibodies for 1 hour at room temperature. Primary and secondary antibodies are detailed in Supplementary Table S1. Cells were washed three times with 0.1% TWEEN 20 (Sigma-Aldrich) in PBS followed by DAPI staining for 10 minutes at room temperature. Samples were imaged on a Nikon AX-R Confocal microscope using Element software. Adherens junctions were quantified using FIJI/Image J (NIH, Bethesda, MD, USA). We randomly selected 8 to 10 cells per image from triplicates for each experiment condition for manual tracing along cell–cell junctions.

Quantitative Reverse Transcriptase PCR Gene Analysis

Total RNA was extracted using TRIzol reagent (Thermo Fisher Scientific) and purified using the RNeasy Mini Kit (Qiagen, Hilden, Germany). RNA quality and concentration were measured using a nanodrop spectrophotometer. cDNA was generated using GoScript Reverse Transcriptase Random Primers kit (Promega, Madison, WI, USA) per the manufacturer's protocol. The TaqMan Universal PCR Master Mix and Gene Expression Assay were used for the genes of interest. TaqMan PCR was performed using the QuantStudio 3 PCR System. The results were calculated as 2^−ΔΔCT obtained by comparing the cycle threshold between samples as normalized to the endogenous control gene TATA-binding protein. Supplementary Table S1 lists all primers.

Western Blot Protein Analysis

Cells were lysed using RIPA Buffer (Thermo Fisher Scientific) with 1× Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher Scientific). Retinas were collected 24 hours after intravitreal injection to assess VEGFR2 activity. Retinas were lysed using RIPA Buffer (Thermo Fisher Scientific) with 1× Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher Scientific). The concentration of extracted protein was quantified using Pierce BCA Protein Assay Kits (Thermo Fisher Scientific). Twenty to thirty micrograms of protein lysate were loaded into a 4% to 20% Mini-PROTEAN TGX precast protein gel (Bio-Rad, Hercules, CA, USA) with electrophoresis and then transferred to a polyvinylidene fluoride membrane. Membranes were blocked with 5% milk or BSA for 1 hour and incubated in primary antibody (Supplementary Table S1) overnight at 4°C. Primary antibodies were detected by using horseradish peroxidase-linked secondary antibody (Cell Signaling Technologies, Davers, MA, USA). Membranes were visualized with Clarity Western ECL Substrate (Bio-Rad) and imaged using Bio-rad ChemiDoc. Blots were analyzed using imaging J. Primary and secondary antibodies are detailed in Supplementary Table S1.

3D PDGF-BB Pericyte Recruitment Assay

Collagen gels were prepared as described elsewhere in this article, with the addition of PDGF-BB (1 ng/mL, 5 ng/mL, and DMSO as vehicle control). iPericytes (75,000 cells/well) were seeded on the top of polymerized collagen gels in ECGM (PromoCell) and invasion was allowed to occur over 24 hours. After 24 hours, gels were fixed then stained for phalloidin and DAPI. The migration of cells into gels was analyzed for the number of cells invading and the distance from the top using the IMARIS software Spots package.

ELISA Analysis of PDGF-BB

iECs were seeded at 13,000 cells/cm² and cultured for 48 hours in ECGM (PromoCell) supplement with 10 µM SB-431542 (Cayman Chemical Company), and 50 ng/mL VEGF-A (PeproTech) or conditioned media collected on day 8 of EC differentiation. PDGF-BB secretion was quantified by ELISA (ThermoFisher Scientific) following the manufacturer’s protocol. Data were normalized relative to 100,000 cells.

Transwell Migration Assay

iECs were seeded on a type I collagen-coated 24-well plate and cultured in the incubator until confluency. iECs were then treated with VEGFR2 inhibitor or vehicle control for 24 hours. The next day, the basolateral side of the transwell inserts with an 8-µM pore size (Corning 3464; Corning, NY, USA) were coated with type I collagen for 1 hour at room temperature. iPericytes were seeded on the apical side of the Transwell inserts with VEGFR2 inhibitor or vehicle control and cultured for 6 hours in the incubator. iPericytes that did not migrate through the filters were removed with cotton swabs. iPericytes that migrated to the basolateral side of the Transwell insert were fixed with methanol and stained for 20 minutes with 0.2% crystal violet in distilled water. Images were taken using an EVOS microscope. The area of iPericytes coverage was quantified using ImageJ. For cell number quantification, iPericytes were lysed with 10% acetic acid and the optical density was analyzed using a microplate reader at 595 nm. Cell density was quantified by fitting absorbance values into a standard curve. Standard curves were generated by seeding iPericytes at various cell densities into a 96-well plate for 24 hours. Cells were then fixed with methanol, stained with 0.2% crystal violet, and lysed with 10% acetic acid. Optical density was analyzed using a microplate reader (BioTek Synergy, Agilent Technologies, Santa Clara, CA, USA) by measuring the optical density at 595 nm. Three to four wells of transwell inserts were used for each experiment with two fields of view for each sample.

3D Collagen Vascular Network Assay

Collagen gels were prepared as described previously.³⁹ Briefly, Rat Tail Collagen-I (Corning) was diluted in 1× PBS to form an 8 mg/mL working solution. To prepare 1 mL of collagen gel solution, 0.8–2.0 × 10⁶ iECs were resuspended in 210.1 µL Medium 199 (1×) (Gibco, Grand Island, NY, USA), 336.5 µL ECGM supplemented with 50 ng/mL VEGF, 38.4 µL Medium 199 (10×) (Gibco), and 375 µL of collagen-I solution. The pH of the gel solution was adjusted by adding up to 40 µL 1 M NaOH. Then, 62.5 µL of the hydrogel solution was added to each well of a 96 well and polymerized at 3 °C for 30 minutes. ECGM supplemented with 50 ng/mL VEGF-A (PeproTech) was added to the gels after 30 minutes. Constructs were incubated for 48 hours to allow network formation with daily media change. For hypoxia experiments, hydrogel constructs were moved to a hypoxia chamber (1% O₂) at 37°C after 24 hours and incubated for additional 24 hours. After a total of 48 hours of incubation, iPericytes labeled with CellTracker Green (Thermo Fisher Scientific), according to the manufacturer's protocol, were passaged to the hydrogel networks (0.5–1.5 × 10⁵ cells/well) with ECGM supplemented with 50 ng/mL VEGF with or without VEGFR2 inhibitor. The hydrogel constructs were returned to the incubator or hypoxia chamber and media with or without VEGFR2 inhibitor was changed every 24 hours for 2 days. Time lapse imaging was performed using Nikon AX-R Confocal microscope using Element software. For staining, gels were fixed in 4% formaldehyde for 20 minutes, followed by a PBS wash three times. Gels were incubated in 1% Triton-X 100 for 10 minutes and washed with PBS for 30 minutes. Next, gels were blocked in 5% BSA solution for 1 hour at room temperature, followed by primary antibody (Supplementary Table S1) incubation overnight at 4°C. Gels were washed three times with PBS containing 0.1% TWEEN 20. Gels were then incubated with a conjugated phalloidin probe and secondary antibody (Supplementary Table S1) for 2 hours at room temperature. Gels were washed three times with PBS containing 0.1% TWEEN 20 and stored in unsupplemented PBS until imaging (Nikon AX-R confocal microscope). Timelapse imaging was quantified at a depth of 100 µm, originating from the gel surface using TrackMate^40,41 to quantify cell speed along the x, y, and z axes. Images were taken at the center of the gel, total of six to seven gels from each group were used for analysis with total of 134 cells analyzed. iPericytes in the representative images were pseudo-colored green for better visual distinction from the vascular networks labeled in magenta. The total volume of migrated pericytes in the gel was quantified using Nikon NIS-Elements software. A total of four gels from each group were used for analysis with three fields of view for each sample.

Transwell Permeability and Transendothelial Electrical Resistance (TEER) Assay

Cells were seeded on type I collagen-coated 24-well Transwell inserts with a 0.4 µM pore size (Corning 3470). After 2 days, VEGFR2 inhibitor or vehicle control was added to the well for 24 hours. Both assays were conducted 24 hours after the treatment. For the permeability assay, after aspirating the media, cells were incubated with transport buffer containing HBSS (Thermo Fisher Scientific), HEPES (Thermo Fisher Scientific), and 0.1% of platelet poor human serum (Millipore Sigma, Burlington, MA, USA) for 30 minutes at 37°C. Thereafter, 70 kDa FITC-dextran (250 µg/mL) was applied on the apical side of the transwell. Samples were collected after 30 minutes to be analyzed with a plate reader at an excitation of 490 nm and emission of 520 nm. The FITC–dextran concentration was calculated with a standard curve. Relative fluorescence intensity was calculated by normalizing to the average reading of the vehicle control group. For the TEER assay, an EVOM3 system (World Precision Instruments, Sarasota, FL, USA) was used to measure the resistance (Ω). Relative TEER values were calculated by normalizing to average reading of the vehicle control group.

In Vivo Experiments: Sex as a Biological Variable

Both in vivo experiments were performed on newborns, including female and male mice. Therefore, sex was not considered a biological variable.

Retina Development Model and Intravitreal Injection

C57BL/6J (Jackson Laboratory, Bar Harbor, ME, USA) mice were used for all experiments. On postnatal day 8 (P8), female or male mice were intravitreally injected with 0.5 µM of ZM323881 or 4 µM of tivozanib diluted in PBS in one eye using a microinjector. The contralateral eye was injected with PBS as vehicle control. All studies were conducted under the Duke University Institutional Animal Care and Use Committee-approved animal protocol A070-22-04.

ORI Mouse Model and Intravitreal Injection

We followed a well-established OIR protocol.⁴² C57BL/6J (Jackson Laboratory) mice were used for all experiments. Mice were subjected to 75% oxygen from P7 to P12 (BioSpherix ProOx 360). On P12, female or male mice were intravitreally injected with 0.5 µM of ZM323881 or 4 µM of Tivozanib diluted in PBS in one eye by using a microinjector. The contralateral eye was injected with PBS as vehicle control. All studies were conducted under the Duke University Institutional Animal Care and Use Committee-approved animal protocol A070-22-04.

Retinal Vascular Permeability Assay in the OIR Mouse Model

Retinal vascular permeability was measured 5 days after intravitreal injection detailed above. Pups were anesthetized with isoflurane, and 4 kDa FITC–dextran (25 mg/mL) was injected into the retro-orbital venous sinus. Pups were euthanized 30 minutes later; eyes were enucleated and immediately fixed in 4% PFA for 10 minutes. The cornea, lens, and retina were removed from the eye cup using dissection forceps and microscissors under a dissecting scope. Retinas were stained and imaged as detailed elsewhere in this article. For quantification of permeability, FITC–dextran was extracted directly from the retina by incubating the tissue in 200 µL N,N-dimethylformamide (Sigma Aldrich) overnight at 60°C, followed by quantifying the fluorescence intensity using a plate reader.

Retina Flat Mount and Immunofluorescence Staining

Eyes from P17 pups (OIR or healthy) were enucleated and immediately fixed in 4% PFA for 30 minutes. The cornea, leans, and retina were removed from the eye cup using dissection forceps and microscissors under a dissecting scope. The isolated retinas were blocked for 1.5 hours using 10% normal goat serum containing 0.3% Triton X-100 in PBS. The retinas were then incubated with DyLight 594-conjugated Griffonia Simplicifolia Lectin I Isolectin B4 (Vector Laboratories, Burlingame, CA, USA) and Alexa Fluor-conjugated Anti-neural glial antigen-2 (NG2) overnight at 4°C in 5% normal goat serum containing 0.3% Triton X-100 in PBS. Retinas were washed and mounted on the slide using Fluoromount-G (Invitrogen). Images of the samples were obtained using a Nikon AX-R Confocal microscope using Element software. Avascular area and neovascular tufts were quantified by comparing the number of pixels in the area of VO or neovascular tufts with the total number of pixels in the retina. Mice with body weight of less than 5 g on the day of tissue harvest (P17) were excluded from the analysis.

Retinal cross-sections were prepared by fixing the eyes from P17 pups in 4% PFA on ice for 10 minutes and embedding into Optimal Cutting Temperature compound overnight at −20°C. Sections were cut along a direction parallel to the sagittal plane of the eyecup on a cryostat at 8 µm. Immunofluorescence staining was conducted by permeabilizing and blocking the cryosections with 5% normal goat serum containing 0.05% TWEEN 20 in PBS, followed by incubation of DyLight 594-conjugated Griffonia Simplicifolia Lectin I Isolectin B4 (Vector Laboratories) overnight at 4°C in the blocking buffer. Slides were washed and incubated with DAPI for 10 minutes the next day. Samples were mounted and imaged on a Nikon AX-R Confocal microscope using Element software.

Statistics

Biological replicates of in vitro assays are indicated as N and technical replicates are indicated as n. For in vivo experiments, the number of mice in each experiment is indicated as n. Details of replicates for each experiment can be found in the figure legends. All bar graphs represent means ± SD. A two-tailed unpaired or paired Student t tests were used to determine significance (GraphPad Prism 10, La Jolla, CA, USA). Significance levels were set at (ns) P > 0.05, *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, and ****P ≤ 0.0001.

Results

iPericytes Express Characteristic Markers and Responsiveness to PDGF-BB

To better understand EC–pericyte interactions, we used hiPSCs to codifferentiate into a bicellular population of ECs and pericytes.^42–44 We used the glycogen synthase kinase 3 inhibitor CHIR99021 to induce mesoderm, followed by subsequent differentiation into CD31⁺ ECs (iECs) based on an established protocol.^45,46 iPericytes were obtained by culturing the CD31⁻ cells in the pericyte growth media for 5 days (Fig. 1A). Simultaneously obtaining iECs and iPericytes from the same hiPSC line allows for a shared genetic background that is ideal for cell–cell interaction studies.

Figure 1.

Characterization of iPericytes. (A) Schematic illustration of iPericyte differentiation. (B) Flow cytometry analysis of PDGFRβ expression in iPericytes. (C) Immunofluorescence images of pericyte markers at passage 2 post sort: neural glial antigen-2 (NG2), calponin-1, CD13, and SM22a (all in green; nuclei in blue). Scale bar, 50 µm. (D) (i) Schematic illustration of iPericyte recruitment experiment. (ii-iii) Representative confocal images showing side views of hydrogels with pericytes (F-actin in green, nuclei in blue) and corresponding quantification show dose-dependent iPericyte migration in response to PDGF-BB. Bars represent means ± SD; N = 2 (0 ng/mL), N = 3 (1 ng/mL, 5 ng/mL). Scale bar, 50 µm. (E) ELISA analysis of PDGF-BB levels in the media. Illustrations created with BioRender.com. Bars represent means ± SD; Significance levels were set at *P ≤ 0.05 and **P ≤ 0.01.

The derived CD31⁺ population (iECs) was expanded and characterized with immunofluorescence staining for EC markers, including PECAM-1 (CD31), vascular endothelial cadherin (VE-cadherin), and von Willebrand factor (Supplementary Fig. S1). The identity of iPericytes was confirmed by flow cytometry analysis and immunofluorescence staining. The iPericytes expressed pericyte markers, including PDGF receptor β (PDGFRβ), NG2, CD13, calponin-1, and SM22a (Figs. 1B, 1C). Expression was maintained for up to 10 passages (Supplementary Fig. S2A). It is worth noting that PDGFRβ is consistently expressed for extended culture, implying its ability to maintain phenotype through passages (Supplementary Fig. S2B).

Next, we evaluated whether iPericytes have functional PDGFRβ receptors, as PDGF-BB/PDGFRβ signaling is essential for initial pericyte recruitment to ECs.⁴⁷ We simulated pericyte recruitment by seeding iPericytes on the top of type I collagen hydrogels with or without exogenous PDGF-BB in the gel for 24 hours (Fig. 1Di). Notably, iPericytes migrate toward the bottom of the hydrogel containing PDGF-BB in a dose-dependent manner (Fig. 1Dii, iii), demonstrating the functionality of the highly expressed of PDGFRβ in iPericytes.

To confirm that iECs can recruit iPericytes, we examined PDGF-BB secretion. Here, PDGF-BB secretion from iECs cultured in ECGM supplemented with VEGF and TGF-β inhibitor (control) was compared with iECs cultured in codifferentiation conditioned media collected before sorting (ECGM supplemented with VEGF and TGF-β inhibitor). Surprisingly, iECs secrete more PDGF-BB when cultured in the codifferentiation conditioned media (Fig. 1E). Therefore, we determined that the presence of iPericytes leads to an increase in PDGF-BB secretion from iECs.

VEGFR2 Inhibition in iECs Leads to Increased Pericyte Recruitment

In a previous study, we found that iECs are highly angiogenic with abundant expression of VEGFR2.⁴⁵ When vessels are in an angiogenic state, they are highly migratory with increased junctional permeability allowing for extensive remodeling.⁴⁸ Furthermore, VEGFR2 is negatively correlated with pericyte recruitment.⁴⁹ We thus hypothesized that pericyte recruitment and vessel stabilization require downregulation of VEGFR2, which is achieved through direct physical contact between ECs and pericytes.

To investigate whether modulating VEGFR2 signaling could affect the ability of ECs to recruit pericytes, we began by treating iECs with VEGFR2 inhibitor ZM323881. First, we analyzed EC-derived factors that have been reported to be important for pericyte migration, invasion, and proliferation.³⁸ After 48 hours of VEGFR2 inhibitor treatment, qRT-PCR was performed on iECs. We found upregulation of PDGF-BB, heparin-binding EGF-like growth factor, and Endothelin-1 when compared with the group without inhibitor treatment (Fig. 2A).

Figure 2.

Inhibition of VEGFR2 in iECs enhances iPericyte recruitment. (A) RT-qPCR for PDGF-BB, heparin-binding EGF-like growth factor (HB-EGF), and endothelin-1 (EDN1) in iECs treated with VEGFR2 inhibitor ZM323881 for 48 hours. Bars represent means ± SD; N = 3. (B) (i) Schematic illustration of two-dimensional migration assay. (ii) Representative images of migrated iPericytes stained with crystal violet. Scale bar, 300 µm. (iii) Quantification of the percentage of area covered by cells and the number of cells migrated. N = 3, total of 10 samples for each group with 2 fields of view for each sample. (C) (i) Schematic illustration of iPericytes recruitment to 3D vascular networks. (ii) Representative immunofluorescence images of iECs networks (CD31, magenta) and iPericytes (CellTracker, green; nuclei, blue) at 24 hours. Scale bar, 100 µm. (iii) Quantification of average iPericyte migration speed and average migration speed in the z axis; n = 6–7 gels for each group, total of 134 cells were analyzed. (D) (i) Representative immunofluorescence images of iECs networks (UEA1, magenta) and iPericytes under hypoxia (CellTracker, green; nuclei, blue) at 24 hours. Scale bar, 150 µm. (ii) Quantification of iPericyte volume in the gel; four gels from each group with three fields of view for each sample. Illustrations created with BioRender.com. Bars represent means ± SD; Significance levels were set at not significant (ns) P > 0.05, *P ≤ 0.05, **P ≤ 0.01, and ****P ≤ 0.0001.

Next, we asked if we would observe increased pericyte recruitment by inhibiting VEGFR2. Using a transwell migration assay, we found that VEGFR2 inhibition increased iPericyte migration with increased cell coverage and cell number (Fig. 2B).

We then asked if we could enhance pericyte recruitment to 3D vascular networks by inhibiting VEGFR2 signaling. To model pericyte migration to endothelial networks, iECs were encapsulated in the type I collagen hydrogels for 48 hours where they self-assembled into 3D vascular networks. iPericytes were then seeded on the top of the hydrogel-containing vascular networks with or without a VEGFR2 inhibitor to mimic pericyte recruitment during capillary network formation (Fig. 2Ci). We observed increased pericyte recruitment in the inhibitor-treated group where more CellTracker-positive cells were recruited to the vascular networks (Fig. 2Cii). We also examined iPericyte migration speed, and although we found no differences in the average cell migration speed across experimental groups, we observed increased cell migration speed of iPericytes in the z axis in the treatment group compared with the control group (Fig. 2Ciii). These data suggest that VEGFR2 inhibition leads to increased iPericyte migration and recruitment through iEC chemoattraction mechanisms.

Finally, to recapitulate the pathological environment in DR, iPericytes were seeded on top of the hydrogel-containing iEC vascular networks under hypoxic conditions. Here, too, we observed greater pericyte penetration, evident by an increase in the total volume of pericytes in the VEGFR2 inhibitor-treated group compared with the control (Fig. 2D).

Overall, the in vitro results indicate that the inhibition of VEGFR2 enhances pericyte recruitment to the vascular networks, potentially through the upregulation and increased secretion of pericyte recruitment factors, leading to increased EC-pericyte interactions.

Pericytes Modulate Phosphorylated VEGFR2 (pVEGFR2) Activity Via Direct EC Contact

We next sought to determine whether pericytes regulate VEGFR2 expression through soluble factors or through direct contact. We first examined VEGFR2 pY951 because it is one of the major phosphorylation sites of VEGFR2 and has been shown to be the key regulator of EC survival, migration, and permeability during angiogenesis.⁸ We cultured iECs in iPericyte-conditioned media (Fig. 3A) or cocultured iECs with iPericytes (Fig. 3D). Western blotting was conducted to analyze VEGFR2 activity and compared among treatment groups over the course of 4 days. We found that VEGFR2 pY951 levels in iECs did not change along the culture with iPericyte-conditioned media (Figs. 3B, 3C). In contrast, the expression of VEGFR2 pY951 in iECs was decreased along the coculture with iPericytes for 3 to 4 days (Figs. 3E, 3F). The same expression pattern was observed with another VEGFR2 phosphorylation site Y1175, which also participated in regulating vascular permeability (Supplementary Fig. S4). We confirmed the absence of VEGFR2 in both iPericytes and primary human retinal pericytes, regardless of VEGF supplementation in the culture media (Supplementary Fig. S5). These results confirm that measured VEGFR2 activity is exclusive to the ECs.

Figure 3.

Direct EC–pericyte contact is required for VEGFR2 pY951 downregulation, leading to barrier stabilization. (A, D) Schematic illustrations of conditioned media and coculture experiments. (B, E) Representative Western blots for VEGFR2 pY951, VEGFR2, and β-actin of control iEC and after culturing in iPericyte-conditioned media or iEC-pericyte coculture for 2, 3, and 4 days. (C, F) Quantification of western blot for pVEGFR2 pY951 and VEGFR2 normalized to β-actin loading control and vehicle control. Bars represent means ± SD; N = 3. (G) Representative maximum intensity projection of immunofluorescence images of VE-cadherin, and β-catenin in iECs treated with or without VEGFR2 inhibitor for 24 hours. Quantification of junctional intensity of VE-cadherin, and β-catenin. N = 3, with two to five fields of view for each sample. Scale bar, 50 µm. (H) FITC–dextran Transwell permeability assay on iECs treated with or without VEGFR2 inhibitor for 24 hours. Bars represent means ± SD; N = 3. (I) TEER measurements on iECs treated with or without VEGFR2 inhibitor for 24 hours. N = 3 (J) TEER measurements on iECs cocultured with or without iPericytes. N = 3. Illustrations created with BioRender.com. Bars represent means ± SD; Significance levels were set at *P ≤ 0.05, **P ≤ 0.01,***P ≤ 0.001, and ****P ≤ 0.0001.

The establishment of direct EC–pericyte contact was confirmed by N-cadherin staining (Supplementary Fig. S3A). Notably, the downregulation of VGFR2 pY951 was abolished when blocking EC–pericyte interaction using N-cadherin inhibitor ADH-1 (Supplementary Fig. S3B). Interestingly, no significant change in pAkt levels was observed across VEGFR2 inhibitor-treated, coculture, or N-cadherin inhibitor-treated groups (Supplementary Fig. S3B, C). Altogether, these data suggest that pericytes modulate EC VEGFR2 activity only by establishing direct physical contact with ECs, while not impacting cellular processes governed by Akt.

VE-cadherin forms adherens junctions between ECs that are important in regulating vascular permeability.⁵⁰ When ECs are in the angiogenic state, they exhibit increased vascular permeability, allowing sprouting and lumen formation.^51,52 Once new vessels are formed, ECs become quiescent, and vascular permeability decreases. Decreased vascular permeability indicates that the vasculature is stabilized, becoming functional and mature. To examine whether inhibition of VEGFR2 signaling enhances VE-cadherin adherens junctions and barrier function, iECs were treated with VEGFR2 inhibitor. As expected, inhibiting VEGFR2 resulted in the downregulation of VEGFR2 pY951 at both early and later time points (Supplementary Fig. S6). Interestingly, inhibition of VEGFR2 pY951 did not significantly impact VE-cadherin expression levels, but rather increased VE-cadherin and β-catenin localization to the cell membrane of iECs (Supplementary Fig. S6, Fig. 3G). To examine the impact on barrier function, we analyzed cell permeability using FITC–dextran and TEER using EVOM3. We found decreased cell permeability (Fig. 3H) and increased TEER (Fig. 3I) when VEGFR2 is inhibited in iECs. The coculture of pericyte and iECs on the transwell membranes results in an increase in TEER compared with the control group (Fig. 3J). Overall, these results suggest that direct contact with pericytes inhibits VEGFR2, resulting in enhanced vascular barrier function and contributing to the stabilization of the vascular network.

VEGFR2 Inhibition Increases Pericyte Recruitment in the Developing Mouse Retina

Next, we sought to test whether the role of VEGFR2 is critical in EC–pericyte interactions during developmental angiogenesis. To achieve this, healthy pups were used as a developing mouse model to examine how VEGFR2 inhibitors impact pericyte recruitment. It has been established that the retinal vessels and pericytes start to emerge, grow, and penetrate the intermediate and deep layer of the retina from approximately P7 onward.^53,54 Thus, we performed an Intravitreal injection on P8 (Fig. 4A). Staining with pericyte marker neural glial antigen-2, revealed that pericyte coverage is increased in the intermediate and deep plexus in the healthy retina treated with VEGFR2 inhibitor compared with untreated retinas (Fig. 4B). Our findings demonstrate that inhibiting VEGFR2 could strengthen pericyte recruitment to the endothelial vasculature, aiding in vascular stability during developmental angiogenesis. Overall, our data present a previously unknown mechanism that governs pericyte-mediated vascular stabilization, in which the downregulation of VEGFR2 occurs through direct contact between ECs and pericytes mediated by N-cadherin. This interaction results in enhanced pericyte recruitment, ultimately leading to the stabilization of the nascent vasculature (Fig. 4C).

Figure 4.

Pericyte recruitment is increased in the developing mouse retina treated with VEGFR2 inhibitor. (A) Schematic illustration of experimental schedule in the developing mouse. (B) (Left) Representative immunofluorescence images of the healthy mouse retinal tissue at P17 throughout different layers. (Right) Quantification of the presence of pericytes (neural glial antigen-2 [NG2], green) relative to mouse vasculature (Isolectin B4, red) (n = 6). Scale bar, 50 µm. (C) Conclusion schematic showing that downregulation of VEGFR2 through N-cadherin–mediated direct contact between ECs and pericytes enhances pericyte recruitment, thereby leading to the stabilization of the nascent vasculature. Illustrations created with BioRender.com. Bars represent means ± SD. Significance levels were set at (ns) P > 0.05, *P ≤ 0.05, and **P ≤ 0.01.

Inhibition of VEGFR2 Enhances Pericyte Recruitment, Resulting in Improved Vascular Integrity in the OIR Mouse Model

To examine the translation implications of our findings, we sought to investigate if inhibiting VEGFR2 in vivo could restore vascular stability in ischemic retinopathy by enhancing pericyte recruitment. We chose the OIR model because it is currently the most widely used and accepted model for investigating mechanisms and treatment of vascular dropout. The OIR model mimics hypoxia-mediated abnormal blood vessel growth seen in proliferative DR. Notably, although diabetic rodent models such as Akita mice recapitulate important DR features, including microaneurysms, pericyte loss, and vascular leakage, these vascular changes develop slowly, beginning at approximately 6 months of age and peaking at 9 months. However, these models show only mild capillary loss and rarely exhibit retinal hypoxia, limiting their usefulness in mimicking the proliferative stage of DR.^55,56

Intravitreal injection of VEGFR2 inhibitor or vehicle control PBS was performed on P12 when pups were returned to room air after 5 days of exposure to 75% oxygen to induce ischemic retinopathy (Fig. 5A). On P17, we confirmed the downregulation of VEGFR2 pY951 in the inhibitor-injected retinas (Fig. 5B). We found an increase in revascularization through a decrease in pathological neovascularization (NV) and less vaso-obliteration (VO) in the VEGFR2 inhibitor-treated OIR retina when compared with the contralateral eye injected with vehicle control PBS (Fig. 5C). To confirm the specificity of the VEGFR2 inhibition, we examined the efficiency of an FDA-approved pan-VEGFR inhibitor, tivozanib, to treat the OIR mouse. Tivozanib was approved by FDA in 2021 to treat relapsed or refractory advanced renal cell carcinoma.⁵⁷ We found no significant decrease in VO and a significant reduction of NV in the inhibitor-treated retina (Supplementary Fig. S7A). Given the critical role of VEGF in NV in the OIR model,⁵⁸ these data suggest that the downregulation of VEGFR2 pY951 activity ameliorates pathological angiogenesis under hypoxia.

Figure 5.

VEGFR2 inhibition improves retinal vascular integrity in OIR by enhancing pericyte coverage, reducing leakage, and supporting revascularization. (A) Schematic illustration of experimental schedule of OIR mouse model. (B) Representative western blots for VEGFR2 pY951 and β-actin in mouse retina collated 24 hours after ZM323881 injection. (C) (i) Representative immunofluorescence images of the OIR mouse retinal tissue at P17, 5 days after intravitreal injection of VEGFR2 inhibitor (Isolectin B4, red; VO area, blue; NV area, white). (ii–iii) Quantifications of VO area and pathological NV at P17 (n = 15). Scale bar, 300 µm. (D) (Left) Representative immunofluorescence images of the OIR mouse retinal tissue at P17 throughout different layers. (Right) Quantification of the presence of pericytes (neural glial antigen-2 [NG2], green) relative to mouse vasculature (Isolectin B4, red) (n = 11). Scale bar, 50 µm. (E) (i) Representative immunofluorescence images of the OIR mouse retina cross-section at P17. (ii, iii) Quantification of retina thickness and the percentage of vessel coverage (Isolectin B4, red) relative to retina area (DAPI, blue) (n = 3 mice, 8 sections each). Scale bar, 100 µm. (F) (i) Schematic illustration of the experimental schedule of the vascular permeability assay with the OIR mouse model. (ii) Representative immunofluorescence images of the OIR mouse retinal tissue (left). Scale bar, 50 µm. (iii) Relative fluorescence intensity of FITC-dextran extracted from OIR retinas treated with or without VEGFR2 inhibitor. n = 6. Illustrations created with BioRender.com. Bars represent means ± SD. Significance levels were set at not significant (ns) P > 0.05, *P ≤ 0.05, ***P ≤ 0.001, and ****P ≤ 0.0001.

We next assessed the involvement of pericytes in ischemic angiogenesis in the OIR retina treated with or without VEGFR2 inhibitor by staining the retina with the pericyte marker neural glial antigen-2. The spatial development of mouse retinal vasculature is formed starting from the superficial to the intermediate and the deep plexus.⁵³ We thus analyzed pericyte recruitment to the vascular plexus in the three layers. Because the superficial plexus vasculature is fully formed at P17,⁵³ we found no difference in pericyte coverage between the treated and control groups (Fig. 5D, left). However, we found that the percentage of pericyte presence in the intermediate and deep layers of the retina is significantly increased in the VEGFR2 inhibitor-treated eyes compared with untreated controls (Fig. 5D). In contrast, treatment with the pan-VEGFR inhibitor tivozanib increased pericyte recruitment in the superficial and intermediate but not in the deep layer of the retina compared with control (Supplementary Fig. S7B). The difference between the two treatments led us to suspect that pericyte-mediated vascular stabilization is mostly mediated by VEGFR2 rather than other VEGFRs.

Next, we stained the cross-sections of the retina from OIR mice to further investigate whether increased pericyte recruitment results in a healthier retinal structure. We observed a significant increase in retinal thickness in the VEGFR2 inhibitor group (Fig. 5Ei, ii), with increased vessel coverage area when VEGFR2 is downregulated (Fig. 5Eiii), suggesting that augmented pericyte involvement improves retina structure and vascular normalization.

Finally, to determine the impact of VEGFR2 inhibition on vessel functionality, we performed a vascular permeability assay on retinas injected with PBS control or VEGFR2 inhibitor (Fig. 5Fi). After FITC–dextran injection to the retro-orbital venous sinus, we found that retinal vascular leakage was reduced in the VEGFR2 inhibitor-treated retina as indicated by representative fluorescence imaging and direct quantification of the amount of FITC-dextran extracted directly from the retina using N,N-dimethylformamide (Fig. 5Fii, iii). This result again supports the role of VEGFR2 in vascular stabilization during pathological angiogenesis. Overall, these outcomes demonstrate that the inhibition of VEGFR2 leads to a healthier and thicker retina, as evidenced by enhanced pericyte recruitment and decreased vascular permeability in an OIR mouse model.

Taken together, our findings demonstrate that pericyte recruitment to the endothelial vasculature can be strengthened through inhibiting VEGFR2, aiding in vascular stability during pathological and physiological angiogenesis.

Discussion

Pericytes play a critical role in stabilizing microvasculature and in maintaining microvascular homeostasis.^18,37 In contrast, during vascular remodeling, pericyte detachment was observed in the embryonic dorsal skin.⁵⁹ Loss of pericytes is reported in various microvascular diseases, such as DR, Alzheimer's disease, and stroke.⁶⁰ It has also been shown that low pericyte coverage on tumor vessels in cancer patients correlates with cancer metastasis.^37,61 Nonetheless, the mechanism by which pericytes stabilize vasculature in angiogenesis during the development of nascent vessels or during pathological angiogenesis is not fully understood.

Most in vitro studies use primary or immortalized cells from different origins. In this study, isogenic ECs and pericytes derived from the same hiPSC origin were used to analyze cell–cell interactions during vascular network formation. PDGF-BB is secreted by ECs to recruit pericytes to blood vessels.¹⁸ Consistent with this finding, we found that iPericytes migrate in response to PDGF-BB. Furthermore, iECs secrete more PDGF-BB when cultured in conditioned media collected from iEC and iPericyte codifferentiation compared with iECs expansion alone. VEGFR2 has been shown to be highly expressed in angiogenic ECs⁸ and is essential in vasculogenesis and angiogenesis.^45,62 Because VEGFR2 was reported to be negatively correlated with pericyte recruitment,⁴⁹ we postulated that the increase in PDGF-BB secretion is controlled by the interactions between pericytes and VEGFR2. We inhibited VEGFR2 in iECs and observed the upregulation of pericyte recruitment-related genes PDGF-BB, HB-EGF, and EDN1 in iECs. It is worth noting that HB-EGF, in addition to PDGF-BB, has been found to be secreted by ECs to attract mural cells during vascular network formation,⁶³ and EDN1 has been linked to pericyte survival and proliferation.⁶⁴

Although VEGFR2 signaling is primarily activated by VEGFA, it is also influenced and modulated by a broader network of related growth factors and other VEGFRs that contribute to vascular development and remodeling. Placenta growth factor and VEGF-B, which mainly bind to VEGFR1, can indirectly regulate VEGFR2 activity through receptor crosstalk.^65,66 VEGF-C and VEGF-D, which primarily interact with VEGFR3, are also implicated in angiogenesis and lymphangiogenesis.⁶⁷ Additionally, fibroblast growth factors, TGF-β, and angiopoietins act synergistically with VEGFR2 pathways to regulate EC angiogenesis.⁶⁸ This complex interplay of growth factors ensures fine-tuned control of vascular formation, maturation, and response to pathological stimuli beyond the VEGFA–VEGFR2 axis.

We next examined the recruitment of pericytes to ECs and observed increased recruitment when VEGFR2 was inhibited. Our observation aligns with a seminal cancer study, which demonstrated that the blockage of VEGFR2 temporarily facilitates pericyte recruitment to the tumor vasculature.¹⁵ The study concluded that pericyte recruitment is regulated by increased secretion of tumor cell-derived Ang-1. Building on this finding, our work uniquely shows that ECs themselves regulate pericyte recruitment through VEGFR2 modulation, which is particularly relevant in treating DR. When treated with the inhibitor, there was a noticeable increase in iPericyte migration speed toward 3D vascular networks on the z axis. However, there was no significant increase in the overall iPericyte migration speed in all three axes. This finding confirms that increased iPericytes migration during EC VEGFR2 inhibition was due to chemoattractants rather than random migration. In the Transwell migration assay, increased iPericyte migration was observed in the group treated with the VEGFR2 inhibitor. This confirms that pericyte recruitment increases in developing EC tube networks when VEGFR2 activity is reduced.⁶⁹

A previous in vivo study demonstrated that vessels lacking VEGFR2 pY951 expression in the murine embryonic body are mostly covered by pericyte-like cells, whereas pericytes are missing from vessels containing VEGFR2 pY951.⁷⁰ Indeed, we found that pericytes modulate vascular stability through the downregulation of VEGFR2 pY951. Interestingly, we only observed downregulation of pVEGFR2 in the coculture group but not in the conditioned media-treated group. This result agrees with recent studies that demonstrate the formation of direct EC–pericyte cell–cell contact is crucial in vasculogenesis and EC maturation.^71,72 Therefore, we conclude that pericytes can regulate VEGFR2 signaling only by establishing a direct contact with ECs, but not through paracrine signaling. Similar to VEGFR2 pY951, other VEGFR2 phosphorylation sites, including Y1175 and Y801, have been implicated in regulating vascular permeability via the PLCγ–PKC pathway.^73,74 Downregulation of VEGFR2 pY1175 was also observed in the coculture group. Although further studies are needed to elucidate the impact of pY1175 on EC function, this observation further confirmed the role of pericyte in EC stabilization.

VEGFR2 pY951 has also been shown to be responsible for the formation of EC adherens junctions in a knocked-out mouse model.⁷⁵ When we treated iECs with VEGFR2 inhibitor, adherens junction formation as well as barrier function were augmented, suggesting a more stable EC phenotype. Although previous studies have established the role of VEGFR2 pY951 in the regulation of vascular permeability in retinopathies,^10,16 our study provides novel insights into how VEGFR2 signaling influences the crosstalk between ECs and pericytes. We speculate that proteins mediating EC–pericyte cell signaling, such as N-cadherin, which require physical adjacency for their binding, mediate the dephosphorylation of VEGFR2 pY951 through downstream signaling pathways upon their interaction. N-cadherin adherens junctions form between ECs and pericytes and regulate the endothelial barrier by inducing VE-cadherin recruitment to EC junctions.⁷⁶ Indeed, blocking EC–pericyte interactions via N-cadherin inhibition diminished VEGFR2 phosphorylation. Another mediator of EC–pericyte interactions is connexin-43, which forms gap junctions that allow for the exchange of small molecules and ions, which is critical during blood vessel formation.⁷¹ Future experiments examining the involvement of these proteins in VEGFR2 downregulation upon EC–pericyte contact and their role in potential therapeutic resistance are warranted. The Akt pathway plays a vital role in cell survival, proliferation, and other cellular functions. Interestingly, we observed no change in pAkt levels across VEGFR2 inhibitor-treated, cocultured, or N-cadherin inhibitor-treated groups, suggesting that the observed downregulation of pVEGFR2 suppresses angiogenic signaling that promotes vascular stabilization without compromising EC viability.

Depletion of pericytes in the early stage of vascular development results in elevated VEGFR2 expression and increased EC proliferation and sprouting.⁷⁷ We inhibited VEGFR2 in healthy pups and observed enhanced pericyte coverage in the retina. This finding is in agreement with a previous study that showed that pericyte coverage is reinstated in VEGFR2 inhibitor-treated mouse patent blood vessels.⁴⁹ Our results provide a deeper understanding of the mechanism in which this occurs.

Pericyte loss is a common occurrence in ocular microvascular diseases, and no effective treatments have been developed to restore the interaction between ECs and pericytes for the revascularization of damaged tissues. We observed a reduction in pathological NV as well as a decrease in VO after VEGFR2 inhibitor treatment, with reductions comparable with those reported in published studies of aflibercept-treated eyes (Supplementary Fig. S8). This finding suggested that VEGFR2 inhibition can achieve vascular normalization effects similar to those of existing anti-VEGF therapies, but with greater specificity. We also observed increased pericyte coverage in the intermediate and deep plexus of the OIR retina when treated with the VEGFR2 inhibitor. The observed decrease in VO by P17 after VEGFR2 inhibitor treatment likely indicates improved revascularization rather than a prevention of vessel loss. This finding implies that the lower VO in the VEGFR2 inhibitor-treated group could be due to increased recruitment of pericytes to the abnormal vessels. Greater pericyte coverage helps to stabilize vessels, thus promoting healthier vascular remodeling and reducing areas of nonperfusion. Indeed, retinal thickness and vessel density are noticeably increased, and vascular permeability is decreased in the OIR eyes injected with VEGFR2 inhibitor. These findings indicate a sign of vascular normalization and, consequently, a healthier retina compared with untreated OIR eyes. However, it is arguable that reduced VO and NV in the mouse OIR retina observed in our study might be due to the result of specifically downregulating VEGFA–VEGFR2 interactions. However, other factors, including insulin-like growth factor 1,⁷⁸ erythropoietin,⁷⁹ and angiopoietin-2⁸⁰ have been suggested in aiding pathological angiogenesis in OIR mouse model. It is conceivable that pericytes may also contribute to the decrease of NV.

In addition to uncontrolled pathological angiogenesis, the loss of therapeutic efficacy after anti-VEGF treatment is a frequent challenge in managing DR. Acquired resistance and local adverse effects are two major resistance events that happen after anti-VEGF treatment. Unlike in cancer, therapeutic resistance in DR is primarily driven by hyperglycemia, chronic inflammation, and persistent vascular leakage.^81,82 Given the limited role of alternative angiogenic pathways in DR,⁸³ stabilizing leaky pathological vasculature is key to improving therapeutic outcomes.⁸⁴ Our study showed that VEGFR2 inhibition enhanced pericyte recruitment, which results in vascular normalization, and therefore might decrease the chance of resistance often occurring after anti-VEGF treatment. Additional long-term functional validation of retinal rescue, such as an ERG or imaging modalities such as optical coherence tomography, could be performed to demonstrate healing benefits. When considering treatment timing, although pericyte dropout occurs early in DR, we expect VEGFR2 inhibition to be more effective later, when NV begins. We recommend that using a VEGFR2 inhibitor is most appropriate during the proliferative stage of DR, ideally at the first signs of new blood vessel growth. At this stage, abnormal vessels caused by hypoxia become visible, and adjusting VEGFR2 signaling may help to decrease abnormal blood vessel formation while promoting blood vessel stability.

Conclusions

Above all, this research highlights the connection between pericytes and VEGFR2 regulation in ECs during vascular network stabilization in development and disease. Using a translationally relevant model, we show that pericytes must establish direct cell–cell contact with ECs to downregulate VEGFR2 signaling. Then, the downregulation of VEGFR2 leads to increased pericyte recruitment, vessel stabilization, and improved vessel function. We demonstrated that targeting VEGFR2 signaling to enhance pericyte recruitment in retinopathy, which in turn promotes revascularization and reduces vascular leakage, thus offering a therapeutic modality.