Identification of RUNX1 as a Mediator of Aberrant Retinal Angiogenesis

Neovascularization is a pathological feature of proliferative diabetic retinopathy (PDR), wet age-related macular degeneration, retinopathy of prematurity, cancer, and other conditions (1). Antiangiogenic therapies typically target vascular endothelial growth factor (VEGF) and are effective treatments for a number of neovascular ocular diseases and some solid tumors (2). Despite the success of anti-VEGF therapies, considerable interest remains in identifying new therapeutic targets for aberrant angiogenesis as anti-VEGF therapies may acutely trigger hemorrhages and tractional retinal detachments in patients with PDR, whereas extended therapy may lead to tissue atrophy, ischemia, and reperfusion injury (3–6).

A major challenge to understanding neovascularization in PDR is that animal models of diabetes do not develop the proliferative phase of diabetic retinopathy consistently (7). Fibrovascular membranes (FVMs) in patients are surgically removed to relieve retinal traction with associated retinal detachment and remain largely understudied as they are often discarded after ocular surgery. To establish platforms for discovering the mechanisms underlying aberrant angiogenesis, we developed methods for the isolation and characterization of vascular endothelial cells (ECs) from patient-derived PDR FVMs (8).

The Internal Review Board of Massachusetts Eye and Ear (MEE) approved this study. Research protocols adhered to the ARVO Statement on Human Subjects and the tenets of the Declaration of Helsinki. All participants gave informed consent prior to surgery and inclusion in the study. Surgical samples were collected at MEE. Control retinal samples were obtained from cadaver eyes of subjects without a diagnosis of diabetes through an approved internal review board protocol from Massachusetts General Hospital (Supplementary Table 1).

CD31⁺ cells were isolated from FVMs as previously described (8). RNA-sequencing was performed using a HiSEq 2000 (Illumina), aligned to reference genome UCSC hg19/GRCh37 with TopHat, and analyzed using Partek Flow (Partek), CuffLinks, EdgeR, and DESeq2 (9). A mixed-model ANOVA was used with a threshold false discovery rate of <0.05 and fold change ≥ ±2 for significance. Gene ontology was determined using the Database for Annotation, Visualization, and Integrated Discovery (DAVID) (10).

Mouse care and experimental procedures were in accordance with MEE Institutional Animal Care and Use Committee regulations. Oxygen-induced retinopathy (OIR) was induced in wild-type C57BL/6J mice as previously described (11). Intravitreal injections with 1 µL of 75 μmol/L Ro5–3335 Runt-related transcription factor 1 (RUNX1) inhibitor or DMSO were performed on left eyes only at postnatal day (P)13 and P15 under ketamine/xylazine anesthesia. Pups were euthanized at P17, and eyes were collected, fixed in 4% paraformaldehyde, and used for retinal flat mounts (n_vehicle = 7; n_Ro5–3335 = 9). Avascular and neovascular areas were quantified using Photoshop (Adobe Systems) as previously described (12).

Human retinal microvascular ECs (HRMECs) (Cell Systems) and human umbilical vein ECs (HUVECs) (Lonza) were grown at 37°C with 5% CO₂ using endothelial growth media plus antibiotics (Lonza) and 2% FBS (Atlanta Biologicals). For the quantitative RT-PCR (qRT-PCR) gene candidate screen in high glucose, cells were treated for 48 h with endothelial basal media-2 with 2% platelet-poor plasma-derived serum (Alfa Aesar) and d-glucose (Sigma-Aldrich) or osmotic control (mannitol and L-glucose; Sigma-Aldrich).

Small interfering RNA (siRNA) (75 nmol/L) (Integrated DNA Technologies; sequences in Supplementary Table 2) were transfected for 6–8 h using DharmaFECT 1 (GE Life Sciences/Dharmacon) in Opti-MEM (Life Technologies).

FVMs and cells were incubated with mouse anti-RUNX1 (Santa Cruz Biotechnology) followed by biotinylated secondary antibody (Vector Laboratories), horseradish peroxidase–labeled avidin (PerkinElmer), tyramide signal amplification (PerkinElmer), Vector Red chromogenic substrate (Vector Laboratories), and Gill no. 3 hematoxylin counterstain (Sigma-Aldrich). Immunofluorescent staining for CD31 (mouse anti-CD31; DakoCytomation) and Ki67 (rabbit anti-Ki67; Novus Biologicals) was performed as previously described (8). The number of Ki67-positive nuclei was averaged for three fields of view per sample at ×20 original magnification.

Mouse retinal whole mounts were blocked with 1% BSA, 0.1% Triton X-100, and 3% serum in PBlec buffer, incubated with primary (isolectin IB₄ Alexa Fluor 488 Conjugate, Life Technologies; rat anti-CD31, BD Pharmingen; and rabbit anti-RUNX1, LifeSpan BioSciences) and secondary antibodies (donkey anti-rat Alexa Fluor 594 and donkey anti-rabbit Alexa Fluor 647 or goat anti-rabbit Alexa Fluor 594; Life Technologies). Samples were imaged with an Axioskop 2 MOT Plus microscope (Carl Zeiss) or TCS-SP5 confocal microscope (Leica Microsystems).

RNA was extracted using RNeasy Mini Kits (Qiagen). Primers were purchased from Integrated DNA Technologies for 200 candidate genes (Supplementary Table 2). Genes were excluded from further analysis if there was no amplification or amplification was outside the linear range. This resulted in a final list of 101 genes analyzed by qRT-PCR. cDNA was prepared using the iScript cDNA synthesis kit (Bio-Rad Laboratories) and probed using FastStart Universal SYBR Green Master (Hoffmann-La Roche). Fluorescent intensities were normalized to spike-in controls (ERCC RNA Spike-In Mix; Life Technologies), hypoxanthine phosphoribosyltransferase 1, and B₂-microglobulin.

Cells were lysed using RIPA buffer (Cell Signaling Technology). Samples were separated on a 4–15% SDS-PAGE, transferred to polyvinylidene difluoride membranes, blocked with Odyssey Blocking Buffer (LI-COR Biosciences, Lincoln, NE), and probed with mouse anti-RUNX1 (Santa Cruz Biotechnology Inc.), rabbit anti–β-actin (Cell Signaling Technology), IRDye 680RD donkey anti-rabbit, and IRDye 800CW donkey anti-mouse (LI-COR Biosciences) antibodies. Immunoreactive bands were visualized using the Odyssey Infrared Imaging System, and band intensities normalized to β-actin were quantified using Image Studio version 2.1 (LI-COR Biosciences).

Migration was assessed with the scratch-wound assay (13). One scratch was generated per well and imaged on an EVOS imaging system every 3 h for 12 h. Images were analyzed with the TScratch Matlab module (14).

HRMECs were treated overnight with 75 μmol/L Ro5–3335 or DMSO before plating 20,000 cells onto wells precoated with basement membrane extract (Trevigen). Cells were imaged 6 h after plating, and tube formation was quantified using the Angiogenesis Analyzer plugin for ImageJ (National Institutes of Health).

Results are presented as mean ± SEM. The Student t test was performed for comparisons between two groups, and one-way ANOVA (Kruskal-Wallis test) was used for comparisons between multiple groups. A P value <0.05 was considered significant.

Whole transcriptomic profiles were constructed for CD31⁺ cells from FVMs and compared with transcriptomes of CD31⁺ cells from postmortem retinas isolated from individuals without diabetes (Supplementary Fig. 1 and Supplementary Table 1). Postmortem retinas were used to define expression baseline because there is no normal correlate of FVMs. CD31⁺ cells were identified as vascular ECs based on their expression of five vascular endothelial markers: CD93, CD31, KLF4, ESAM, and VEGFR1 (Supplementary Table 3). However, because a single marker, namely CD31, was used for cell isolation, the presence of other cell types cannot be ruled out. Principal component analysis of the transcriptomes demonstrated congruent expression profiles for control samples, whereas profiles of cells from the FVM samples had more variable gene expression patterns (Fig. 1A). The data sets are accessible through the National Center for Biotechnology Information’s Gene Expression Omnibus (GSE94019) (15).

Two hundred genes were identified as differentially expressed in CD31⁺ cells from FVMs involving gene ontology categories related to wound response, vessel development, angiogenesis, and other categories (Fig. 1B and Supplementary Table 4).

Hyperglycemia is a major risk factor for progression to PDR in patients with diabetes (16). High glucose-dependent genes from the candidate gene list were identified in primary HRMEC culture using a qRT-PCR screen. Four out of 101 genes were glucose responsive consistent with transcriptomic data (fold change ± SEM): RUNX1 (2.9 ± 0.2), peptidyl-prolyl cis-trans isomerase F (PPIF) (3.6 ± 0.5), serine/threonine-protein kinase PIM3 (PIM3) (3.9 ± 1.0), and CD44 (3.5 ± 0.4) (Fig. 2A and B). We focused on RUNX1 for further studies because of its documented roles in EC biology (17–19). Exposure of HRMEC and HUVEC cultures to high glucose induced a 30% increase in RUNX1 protein, whereas osmotic controls had no significant effect in HRMEC and HUVECs (Fig. 2C and D). Lastly, we confirmed increased RUNX1 protein expression in vessels from FVMs compared with retina from control subjects without diabetes using immunohistochemical staining (Fig. 2E).

siRNA knockdown of RUNX1 resulted in a 60% decrease in wound closure rate, indicating a significant role of RUNX1 in endothelial cell migration (Fig. 3A). The coefficient of determination (r² value) for the simple linear regression was calculated comparing wound closure (dependent variable) to time (independent variable) (control r² = 0.99; scramble siRNA r² = 0.97; and RUNX1 siRNA r² = 0.97). This experiment was validated with a second siRNA targeting RUNX1 to rule out possible off-target effects (Supplementary Fig. 2B). ECs transfected with RUNX1 siRNA had 40% fewer Ki67-positive cells compared with controls 24 h after transfection, indicating RUNX1 also contributes to retinal EC proliferation (Fig. 3B).

To investigate the potential therapeutic relevance of RUNX1, the small molecule Ro5–3335 RUNX1–core binding factor β inhibitor II was tested in vitro and in vivo (20). Treatment with RUNX1 inhibitor reduced total tube length (18% reduction), nodes (35% reduction), and meshes (46% reduction), supporting RUNX1’s role in vascular morphogenesis (Fig. 3C).

Neovascular tufts of P17 C57BL/6J mice with OIR identified by isolectin B4 and CD31 staining showed increased expression of RUNX1, whereas RUNX1 staining was absent from the underlying normal retinal vasculature, indicating a role for RUNX1 in active pathological angiogenesis (Fig. 4A). In vivo testing of the effects on aberrant angiogenesis of RUNX1 inhibition was conducted by injecting Ro5–3335 intravitreally in mice with OIR. There was no significant change in the extent of vaso-obliteration, and no effect was observed in the contralateral eye (data not shown). Eyes treated with inhibitor exhibited a 50% reduction in neovascularization compared with vehicle-treated eyes (Fig. 4B).

Our analyses of CD31⁺ cells from FVMs of patients with PDR in conjunction with in vitro and in vivo validation assays suggest a role for RUNX1 in the proliferation, migration, and morphogenesis of ECs in vitro as well as in aberrant angiogenesis in vivo. RUNX proteins have pleiotropic functions in vascular development, hematopoiesis, and cancer through direct transcriptional regulation of target genes and complex interactions with fundamental signaling mechanisms including Notch and transforming growth factor-β pathways (21,22). RUNX1 is a major contributor to cancer progression and metastasis, most notably in acute myeloid leukemia, and also plays a role in angiogenesis (18,23–26).

Previous studies in model organisms have shown that glucose levels trigger RUNX1 expression via reactive oxygen species–mediated upregulation of hypoxia-inducible factor 1 and regulate hematopoietic stem cell production by endothelial to hematopoietic transition in a RUNX1-dependent manner (27,28). Consistent with these observations, our results demonstrate that high glucose regulates RUNX1 and implicates RUNX1 in aberrant retinal angiogenesis.

Previous studies of PDR have mainly focused on gene expression in FVMs or biomarkers in the vitreous (29,30). Although FVMs from PDR may be highly informative pathological tissues, they are largely unstudied. The window for processing samples postsurgery is narrow, and intraoperative confounders such as FVM size, location, and involvement of sensory retina further complicate successful sample acquisition. CD31⁺ cells from FVMs do not grow readily in culture, making additional experimentation difficult. Limited sample size may also skew our gene ontology analysis.

We report elevated RUNX1 expression in ECs of patient-derived FVMs from patients with PDR. Also, we demonstrate a role for RUNX1 in EC migration, proliferation, and vascular morphogenesis. Furthermore, we report selectively enhanced expression of RUNX1 in neovascular tufts in an experimental model of OIR and that inhibition of RUNX1 function reduced retinal neovascularization. These findings, including the high glucose–dependent expression of RUNX1, identify a novel pathway of potential therapeutic interest and implicate RUNX1 in aberrant angiogenesis in multiple conditions.

Acknowledgments. The authors thank Dr. Magali Saint-Geniez for insightful discussions and providing mouse retinal samples, Dr. Eric Ng, Mark Graham, and Dr. Arturo Machuca-Parra for helpful discussions, and Dr. Christina Marko for technical contributions (all from Schepens Eye Research Institute, Boston, MA). The authors also thank Kristin Waraska and Andrew Gagne of the Harvard Biopolymers Facility, Harvard Medical School, and the Ocular Genetics Institute core facility, Massachusetts Eye and Ear (Boston, MA) for their insights and work.

Funding. The work performed in the manuscript was supported by National Institutes of Health grants R01-EY-005318 (to P.A.D.), R00-EY-021624 (to J.F.A.-V.), UH2NS100121-01 (to J.F.A.-V.), R21-EY-027061 (to L.A.K.), K12-EY-16335 (to L.A.K.), and Center Core grant P30-EY-003790; American Diabetes Association Innovation Award 7–12-IN-11 (to P.A.D.); Massachusetts Lions Eye Research Fund (to J.F.A.-V.); E. Matilda Ziegler Foundation for the Blind (to L.A.K.); Karl Kirchgessner Foundation (to L.A.K.); Department of Ophthalmology, Harvard Medical School (to J.F.A.-V. and L.A.K.); the Howard Hughes Medical Institute Medical Research Fellows Program (to D.J.O.); and the Vascular Biology Program at Boston Children’s Hospital (to D.R.B.).

Duality of Interest. P.A.D., L.A.K., and J.F.A.-V. have filed a provisional patent application on RUNX1 regulation for the treatment of aberrant angiogenesis and PDR. No other potential conflicts of interest relevant to this article were reported.

Author Contributions. J.D.L. and D.J.O. performed the experiments, analyzed the data, and wrote the manuscript. L.L.W., D.A., C.P.-W., A.V.S., and J.C.-V. performed experiments. D.M., A.O.S.-R., D.E., and D.R.B. provided tissue samples and reviewed and edited the manuscript. T.v.Z., L.S., and X.G. obtained data and reviewed the manuscript. P.A.D., L.A.K., and J.F.A.-V. conceived the project, analyzed the data, and wrote the manuscript. P.A.D., L.A.K., and J.F.A.-V. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.