Transcriptional profiling reveals a critical role for ... - The FASEB Journal

The FASEB Journal article fj.08-123810. Published online January 9, 2009.

The FASEB Journal • Research Communication

Transcriptional profiling reveals a critical role for tyrosine phosphatase VE-PTP in regulation of VEGFR2 activity and endothelial cell morphogenesis Sofie Mellberg,* Anna Dimberg,* Fuad Bahram,* Makoto Hayashi,* Emma Rennel,*,1 Adam Ameur,† Jakub Orzechowski Westholm,† Erik Larsson,‡ Per Lindahl,‡ Michael J. Cross,*,2 and Lena Claesson-Welsh*,3 *Department of Genetics and Pathology, Rudbeck Laboratory, and †Linnaeus Centre for Bioinformatics, Biomedical Centre, Uppsala University, Uppsala, Sweden; and ‡Wallenberg Laboratory of Cardiovascular Research, Sahlgrenska University Hospital, Gothenburg, Sweden ABSTRACT To define molecular events accompanying formation of the 3-dimensional (3D) vascular tube, we have characterized gene expression during vascular endothelial growth factor (VEGF)-induced tubular morphogenesis of endothelial cells. Microarray analyses were performed comparing gene induction in growtharrested, tube-forming endothelial cells harvested from 3D collagen cultures to that in proliferating endothelial cells cultured on fibronectin. Differentially expressed genes were clustered and analyzed for specific endothelial expression through publicly available datasets. We validated the contribution of one of the identified genes, vascular endothelial protein tyrosine phosphatase (VE-PTP), to endothelial morphogenesis. Silencing of VE-PTP expression was accompanied by increased VEGF receptor-2 (VEGFR2) tyrosine phosphorylation and activation of downstream signaling pathways. The increased VEGFR2 activity promoted endothelial cell cycle progression, overcoming the G0/G1 arrest associated with organization into tubular structures in the 3D cultures. Proximity ligation showed close association between VEGFR2 and VE-PTP in resting cells. Activation of VEGFR2 by VEGF led to rapid loss of association, which was resumed with time in parallel with decreased receptor activity. In conclusion, we have identified genes, which may serve critical functions in formation of the vascular tube. One of these, VE-PTP, regulates VEGFR2 activity thereby modulating the VEGF-response during angiogenesis.—Mellberg, S., Dimberg, A., Bahram, F., Hayashi, M., Rennel, E., Ameur, A., Westholm, J. O., Larsson, E., Lindahl, P., Cross, M. J., Claesson-Welsh, L. Transcriptional profiling reveals a critical role for tyrosine phosphatase VE-PTP in regulation of VEGFR2 activity and endothelial cell morphogenesis. FASEB J. 23, 000 – 000 (2009)

Key Words: angiogenesis 䡠 gene expression 䡠 PTPRB 䡠 HPTP␤ 䡠 signal transduction

Angiogenesis, formation of new blood vessels from preexisting capillaries, plays a crucial role in normal 0892-6638/09/0023-0001 © FASEB

physiological development and in pathologies such as cancer and rheumatoid arthritis (1). Angiogenesis is a coordinated process in which endothelial cells become induced to secrete proteases, which digest the basement membrane, allowing migration of endothelial cells into the surrounding tissue to form an angiogenic sprout. The sprout elongates, and the endothelial cells differentiate to form a lumen. Vascular endothelial growth factor (VEGF)-A is a potent angiogenic regulator that binds to 2 related receptor tyrosine kinases: VEGF receptor-1 (VEGFR1) and VEGFR2. VEGFR1 is expressed on hematopoietic and vascular endothelial cells, whereas VEGFR2 is preferentially expressed on vascular endothelial cells (2). It is generally accepted that many of the biological effects of VEGF are mediated by VEGFR2. Homozygous deletion of the murine vegfr2 gene is lethal at d 8.5–9.5 with embryos exhibiting defects in the development of hematopoietic and endothelial precursors (3). Ligand-induced activation of VEGFR2 results in receptor autophosphorylation and activation of intracellular signaling pathways regulating biological responses such as proliferation, migration, survival, and increased vascular permeability (2, 4). Treatment with a neutralizing antibody against VEGF (Avastin/ bevacizumab), in combination with chemotherapy, significantly increases survival of patients with metastatic colorectal cancer (5). These important data suggest that by defining the mechanism of how cells respond to VEGF and other angiogenic factors, it may be possible to specifically target intracellular signaling mechanisms that regulate angiogenesis, as part of anticancer therapy. 1 Current address: Microvascular Research Laboratories, Department of Physiology and Pharmacology, Preclinical Vet School, Southwell St., University of Bristol, Bristol BS2 8EJ, UK. 2 Current address: Department of Pharmacology and Therapeutics, University of Liverpool, Sherrington Bldg., Liverpool L693GE, UK. 3 Correspondence: Department of Genetics and Pathology, The Rudbeck Laboratory, Uppsala University, S-751 85 Uppsala, Sweden. E-mail: [email protected] doi: 10.1096/fj.08-123810

1

We have reported on proteomic screens resulting in the identification of key regulators of VEGF-driven tubular morphogenesis of microvascular endothelial cells, including chloride intracellular channel-4, ␣Bcrystallin, and ninein (6 – 8). Previous gene array studies using human umbilical vein endothelial cells (HUVECs) have started to detail gene regulation during tubular morphogenesis in response to a mixture of growth factors (9 –14). A recent compilation of transcriptional profiling information from publicly available databases allowed identification of 58 gene transcripts with specific expression in the microvasculature (15). The current study was undertaken to specifically identify VEGF-induced signaling pathways and gene expression, which regulates proliferation or tubular morphogenesis of endothelial cells and the kinetics of this regulation. Thereby, we have defined clusters of regulated, endothelial cell-expressed genes that are critical in different phases of the angiogenic process. Of these, we chose to validate the role of vascular endothelial protein tyrosine phosphatase (VE-PTP) in tubular morphogenesis of endothelial cells. We show that VE-PTP silencing was accompanied by increased VEGFR2 activity and signaling indicating that VEGFR2 serves as a substrate for VE-PTP. MATERIALS AND METHODS Cell culture Telomerase-immortalized human microvascular endothelial (TIME) cells (16) (gift from Dr. Martin McMahon, UCSF Comprehensive Cancer Center, University of California, San Francisco, CA, USA) and human dermal microvascular endothelial cells (HDMECs; PromoCell, Heidelberg, Germany) were cultured on gelatin-coated dishes in Endothelial Cell Basal Medium MV 2 (EBM-2, C-22221; PromoCell) with supplemental pack C-39221, containing 5% fetal calf serum (FCS), epidermal growth factor (5 ng/ml), VEGF (0.5 ng/ml), fibroblast growth factor-2 (10 ng/ml), long R3 insulin growth factor-1 (20 ng/ml), hydrocortisone (0.2 ␮g/ml), and ascorbic acid (1 ␮g/ml). TIME cells at passage 15–35 were used. For experimental purposes, cells were serum starved overnight and plated out in EBM-2 medium with 1% FCS without growth factor supplements. Inhibitors of the Src pathway (PP2; 5 ␮M), the phosphoinositide 3⬘ kinase (PI3K) pathway (LY294002; 10 ␮M), and ERK/MEK pathway (U0126; 10 ␮M), all from Calbiochem (La Jolla, CA, USA), were added to cell cultures 1 h before stimulation with or without VEGF for 18 h. Preparation of fibronectin matrix Fibronectin (human, F0895; Sigma-Aldrich, St. Louis, MO, USA) was diluted in EBM-2 medium to 20 ␮g/ml and used to coat dishes at 37°C overnight. Serum-starved cells were seeded at a density of 26,000 cells/cm2, left to adhere for 2 h at 37°C, and then treated with 50 ng/ml VEGF-A165 (henceforth denoted VEGF; Peprotech, Rocky Hill, NJ, USA) in EBM-2 medium, 1% FCS. Tubular morphogenesis in 3-dimensional (3D) collagen gels Collagen type I (Vitrogen; Cohesion Technologies, Palo Alto, CA, USA) was mixed with 0.1 M NaOH and 10⫻ Ham’s F12 2

Vol. 23

May 2009

medium (PromoCell) (8:1:1), and components were added to final concentrations indicated: 0.02 M HEPES, 0.1% w/v bicarbonate, 2 mM Glutamax-I (Gibco, Invitrogen, Carlsbad, CA, USA). The collagen mix polymerized at 37°C overnight. Serum-starved cells were seeded out at 67,600 cells/cm2 and left for 2 h at 37°C. A second layer of collagen was added and left to polymerize for 1 h. Thereafter, cultures were treated with VEGF or fibroblast growth factor 2 (FGF-2), each at 50 ng/ml (Peprotech EC) in EBM-2 medium, 1% FCS, or 10% FCS for up to 24 h. Cells stained for confocal imaging or protein extraction were not overlaid with a second collagen layer; instead medium was aspirated, and new medium with or without VEGF was added 3 h after seeding the cells. Cell cycle distribution by FACS analysis TIME cells were serum starved overnight and cultured on fibronectin or within a collagen gel for 24 h with 50 ng/ml VEGF. Cells on fibronectin were trypsinized. Cells within the collagen matrix were isolated by collagenase digestion (SigmaAldrich) for 15 min at 37°C, followed by passage through a 40 ␮M cell strainer [Falcon, Becton Dickinson (BD) Biosciences, San Jose, CA, USA]. Cells were washed in PBS, and distribution of cells in different phases of the cell cycle was analyzed according to Vindelov (17). Briefly, nuclei were prepared by treating the cells with 0.03 mg/ml trypsin (Sigma-Aldrich) for 10 min at room temperature (RT) and thereafter with 0.08 mg/ml RNase A (Sigma-Aldrich) and trypsin inhibitor 0.5 mg/ml (Sigma-Aldrich) for 10 min at RT, and finally staining with 0.2 mg/ml propidium iodide (Sigma-Aldrich) for 15 min on ice. The stained nuclei were analyzed by the BD FACSDiva program using a flow cytometer (BD LSR II, BD Biosciences) and ModFit software (Verity Software House, Topsham, ME, USA). RNA extraction, cDNA labeling, and microarray hybridization Total RNA was extracted using the RNeasy Mini Protocol (Qiagen, Valencia, CA, USA). Briefly, total RNA was extracted by addition of lysis buffer (1:1 matrix:lysis buffer for cells on collagen). The lysate was passed through a 21⫻ G syringe and then through a QIAshredder column before binding RNA to an RNeasy column. DNase treatment was performed using the On-Column DNase Digestion kit (Qiagen) during RNA isolation. Total RNA quantity and quality was measured using a Bioanalyser with the RNA 600 Nano Chip (Agilent Technologies, Life Sciences & Chemical Analysis, Santa Clara, CA, USA). Fluorescently labeled cDNA was generated using Micromax TSA labeling and detection kit (Perkin Elmer, Life and Analytical Sciences, Boston, MA, USA). Four micrograms of total RNA was used per cDNA reaction together with reference RNA Lucidea Universal ScoreCard (GE Healthcare, Little Chalfont, UK). cDNA cleanup was performed using Microcon YM-30 columns (Millipore, Billerica, MA, USA). Samples from time points 15 min to 24 h were diluted in 60 ␮l hybridization buffer together with cDNA from time point 0 h (reference point) and loaded onto human arrays containing 30,000 human cDNAs (Royal School of Technology/ Wallenberg Consortium North, Stockholm, Sweden) spotted on gamma amino propyl silane-coated slides, UltraGAPS (Corning, Corning, NY, USA). Slides were prehybridized in filtered 1% BSA, 5% sodium citrate-sodium chloride (SSC), and 0.1% sodium dodecyl sulfate at 55°C for 30 min, washed in 2⫻ SSC, 0.2⫻ SSC, and in water before drying by centrifugation. Slides were covered with lifterslip (Erie Scientific Company, Portsmouth, NH, USA). Arrays were washed ac-

The FASEB Journal

MELLBERG ET AL.

cording to the manufacturer’s protocol but with increased volumes of washing buffer and extended washing time periods to reduce nonspecific background. Array slides were dried by centrifugation and scanned using the Gene Pix 4000B scanner with GenePix IV software (Axon Instruments, Molecular Devices, Foster City, CA, USA). PMT settings were set to give a ratio of 1 between the different channels in the dynamic range. Bioinformatics analysis Array data were stored and analyzed within the Linnaeus Centre for Bioinformatics Data Warehouse (18), a system built from the BASE platform (19) for microarray data management. Spot signals were extracted from the median pixel intensities, and data were normalized using the print-tip loess algorithm (20). Spots flagged as “not found” or “bad” were removed, and intensities were merged for reporters printed multiple times on each array. The mean log2-ratios were calculated from replicate arrays (dye-swap experiments) to obtain one single expression level for each reporter. Genes with a log2-ratio ⬎ 1 or ⬍ ⫺1 in at least 2 time points on collagen or fibronectin were kept, which reduced the number of reporters to 1135. Reporters were analyzed using k-means clustering, (k⫽14, euclidean distance; Genesis; Institute for Genomics and Bioinformatics, Graz University of Technology, Graz, Austria). After clustering, each cluster was analyzed for statistical overrepresentation of gene ontology annotations (2-tailed Fisher’s exact test) (21) with respect to the distributions over the whole array.

virus reverse transcriptase (USB Corp., Cleveland, OH, USA). Primers were designed using Primer Express software (Applied Biosystems, Foster City, CA, USA) and purchased from Invitrogen. Real-time PCR was performed on cDNA using 2⫻ SYBR Green PCR Master Mix (Applied Biosystems) and run in triplicates on an ABI Prism 7700 Sequence Detection System instrument (Applied Biosystems) with an initial 10 min at 95°C, followed by 45 cycles at 95°C for 15 s and 60°C for 60 s. The calculated threshold cycle (CT) value for each transcript was normalized against the corresponding ␤-actin CT value. Normalized values were related to time point 0 h and presented as fold change of triplicates ⫾ sd. The following oligonucleotides (Invitrogen) were used for realtime PCR analysis (forward, reverse primers; 5⬘-3⬘): ␤-actin: ATGGATGATGATATCGCCGC, AAGCCGGCCTTGCACAT; VE-PTP: GCGGACCAGGATTCCCTCTA, AACTCCCGGATGGTCC. siRNA TIME cells were transfected with Stealth™ siRNA duplex oligoribonucleotides against human VE-PTP (HSS108847; Invitrogen) or with control siRNA (12935-300; Invitrogen). For quantitative PCR, cells were left in full growth medium (EBM-2 with supplemental pack) for 4 d after transfection, followed by RNA extraction. To quantitate effects of siRNA on tubular morphogenesis, images spanning the entire wells were analyzed using Easy Image Analysis 2000 software (Rainfall, Stockholm, Sweden). Cells forming clusters rather then tubular structures were excluded.

Gene expression profiling Affymetrix microarray datasets covering 61 mouse and 79 human tissues and cell lines were downloaded from the Novartis Foundation (22). Probe sets were annotated against the ENSEMBL genome database using BLAST (23, 24). Multiple probe sets annotated to the same gene were averaged. Real-time PCR analysis One microgram of DNase-treated total RNA was used for cDNA synthesis using dT18 and murine moloney leukemia

Immunoblotting Samples were run on Novex NuPage 3– 8% Tris-Acetate Gels (Invitrogen) and transferred to Hybond-C extra membranes (GE Healthcare). Signals were detected using the enhanced chemiluminescence plus detection reagent (GE Healthcare). Antibodies used were against the following: VEGFR2 (AF357, R&D Systems), pTyr951-VEGFR2 (2471), pTyr1175-VEGFR2 (2478), pTyr1214-VEGFR2 (2477), pThr202/Tyr204-Erk (9101), Erk1/2 (9102), pSer473-Akt (9271), Akt (9272) (all from Cell Signaling, Danvers, MA, USA), pTyr783-PLC-␥ (44-

Figure 1. Morphology of TIME cells on fibronectin and collagen I. TIME cells seeded on fibronectin (FN; panels a– d) or collagen (Coll; panels e–j) were observed at different time points during a 24 h treatment with 50 ng/ml VEGF. Whereas cells on fibronectin responded mitogenically to VEGF, cells in the collagen I cultures fused and formed tubular structures composed of numerous cells in a row. In the absence of VEGF, tubes would initially form but soon regress. Boxed area in panel h is magnified in panels i and j; in j, cells have been stained with rhodamine-coupled phalloidin (red) to visualize actin fibers and with Hoechst 33342 (blue) to visualize nuclei. Scale bar ⫽ 100 ␮m.

VE-PTP IN VEGFR2-INDUCED ENDOTHELIAL MORPHOGENESIS

3

696Z; Biosource, Invitrogen), and ␤-catenin (610153; BD Transduction Laboratories). Secondary antibodies used were antimouse-HRP (NA931V; GE Healthcare), anti-rabbit-HRP (NA934V; GE Healthcare), and anti-goat-HRP (A4174; SigmaAldrich). Immunostaining Cells immunostained for light or fluorescence microscopy were fixed for 30 min in 4% paraformaldehyde (PFA), and permeabilized in 0.2% Triton X-100, 3% BSA. Actin stress fibers were stained with Texas Red-conjugated phalloidin and nuclei visualized using 1 ␮g/ml Hoechst 33342 (Molecular Probes, Invitrogen). For confocal imaging, cells were fixed for 30 min in Zink-fix (0.1 M Tris HCl, pH 7.5; 3 mM CaAc; 23 mM ZnAc; and 37 mM ZnCl2). Permeabilization and blocking were done in 0.2% Triton X-100, 3% BSA (TBS-TB). Cells were incubated with primary antibodies; rabbit anti-human VEGFR2

(2479; Cell Signaling), goat anti-human VE-cadherin (sc6458; Santa Cruz Biotechnology, Santa Cruz, CA, USA), diluted in TBS-TB. The following day, cells were washed in TBS containing 0.1% Tween 20 and stained with fluorescent secondary antibodies, Alexa Fluor 555 donkey anti-goat (A21432; Invitrogen) and Alexa Fluor 488 donkey anti-rabbit IgG (A21206; Invitrogen), diluted in TBS-TB buffer for 1 h at RT, with subsequent washes in TBS with or without 0.1% Tween 20. Nuclei were stained using 1 ␮g/ml Hoechst 33342. Proximity ligation assay (PLA) TIME cells at a density of 20,000 cells/well were seeded into fibronectin-coated 8-well chamber slides. After 24 h, cells were serum starved overnight (EBM-2, 1% FCS) and the following day stimulated with 50 ng/ml VEGF in EBM-2 medium containing 1% FCS for 2–15 min at 37°C. Cultures were fixed in 4% PFA and subjected to PLA (www.olink.com). Briefly, slides were blocked and incubated with primary antibodies; mouse anti-

Figure 2. k-Means clustering and gene ontology (GO) analyses. A) Genes with a log2 ratio ⬎ 1 or ⬍ ⫺1 in at least 2 time points in one of the time series of cells cultured in 3D collagen or on fibronectin were clustered using k-means clustering (k⫽14), to identify genes with different expression patterns on collagen and fibronectin. B) Each cluster was analyzed for statistical overrepresentations of GO annotations for biological process (GO data from Entrez Gene database; U.S. National Center for Biotechnology Information, Bethesda, MD, USA). Cutoff for the 2-sided P value was 0.001.

4

Vol. 23

May 2009

The FASEB Journal

MELLBERG ET AL.

human VEGFR2 (sc-6251; Santa Cruz), rabbit anti-GFP (sc-8334; Santa Cruz), mouse anti-V5 (46-0705; Invitrogen), or rabbit anti-human VE-PTP. The VE-PTP antibody was produced by immunizing rabbits with a peptide raised against the VE-PTP juxtamembrane part in the form of a synthetic peptide. Secondary antibodies (anti-mouse and anti-rabbit) conjugated to unique DNA probes (Olink Bioscience, Uppsala, Sweden) were added. Ligation and circularization of the DNA was followed by a rolling circle amplification step, and reactions were detected by a complementary Cy3-labeled DNA linker (25). Slides were mounted using Vectashield (Vector Laboratories, Burlingame, CA, USA) and evaluated using an LSM 510 META confocal microscope (Carl Zeiss, Oberkochen, Germany). The numbers of red PLA spots/cell were counted by utilizing Blob-Finder 2.5 image analysis software, developed by the Centre for Image Analysis, Uppsala University. Statistical analysis was performed with a confidence interval of ⬍ 0.05, 1-way ANOVA, KruskalWallis test, and Dunn’s multiple comparing test. Values are presented with standard errors of the mean (sem).

RESULTS Molecular profiling of human endothelial cells undergoing tubular morphogenesis TIME cells (16) seeded between 2 layers of a collagen I matrix in the presence of VEGF line up, fuse, and form vessel-like structures, whereas cells cultured on fibronectin grow as a monolayer and respond mitogenically to VEGF (Fig. 1). FACS analyses of cells in collagen showed that 3% of the population was in the dividing phase of the cell cycle (S⫹G2/M), compared to 17% of cells on fibronectin (data not shown). We conclude that tubular morphogenesis is associated with growth arrest. To define molecular events regulating VEGF-driven endothelial cell morphogenesis to form vessel structures, we compared gene induction in the growtharrested, tube-forming endothelial cells harvested from 3D collagen cultures, to that in proliferating endothelial cells cultured on fibronectin. Fluorescently labeled cDNA prepared from time points 15 min, 1, 3, 6, 9, 12, 18, and 24 h of VEGF stimulation were mixed with differently labeled cDNA from time point 0. Hybridization was performed to microarrays containing 30,000 human cDNA clones. Microarray data were normalized and filtered within the Linnaeus Centre for Bioinformatics Data Warehouse (18), a system built from the BASE platform (19) for microarray data management.

in the collagen cultures compared to fibronectin cultures (clusters I–K). The validity of the clustering is indicated by the presence of Spermidine/spermine N1-acetyltransferase (SAT) in cluster A (up-regulated in collagen and down-regulated in fibronectin cultures; Fig. 2). SAT is known to be up-regulated in growth arrested cells (27). See Supplemental Data for a complete list of genes regulated during different stages of VEGF-A-driven endothelial cell tubular morphogenesis. Each cluster was further analyzed regarding gene ontology (21) (Fig. 2). The statistical overrepresentation of ontology annotations for biological process in the clusters was analyzed with a cutoff for the 2-sided P value of 0.001. Cluster C, containing genes highly up-regulated on collagen and relatively unchanged or induced at a lower level on fibronectin, was classified as “angiogenesis,” “blood vessel morphogenesis,” and “vasculature development.” Endothelial cell expression of regulated genes Clusters containing genes differentially expressed between the two time studies were further analyzed with regard to endothelial expression pattern using the Affymetrix microarray datasets provided by the Novartis Foundation (22). This allows profiling of genes with regard to their expression in different organs and during embryo development in mouse and human tissues. As shown in Fig. 3 (see Supplemental Data for a complete list of organs represented in the datasets), the endothelial specific protein VE-cadherin is highly expressed in fetal and adult lung and placenta in mice and humans. VEGFR2 is preferentially expressed in the lung in mice and in adult and fetal lung, cardiomyocytes, placenta, and thyroid in humans. Based on such comparison with the expression patterns of VE-cadherin and VEGFR2, genes in clusters A–C and I–K were screened for endothelial cell-expression pattern. Thereby, genes previously known as preferentially expressed in endothelial cells were iden-

Analysis of gene expression patterns To identify different expression profiles, the array data were clustered by k-means clustering, (k⫽14), using Genesis software (26) (Fig. 2). Clusters with expression patterns that differed between the 2 culture conditions as well as clusters with a very similar pattern were identified. Of particular interest were clusters showing marked up-regulation of genes in the collagen cultures compared to fibronectin cultures (clusters A–C), or, alternatively, clusters showing marked down-regulation

Figure 3. Gene expression profiling of VE-PTP, VEGFR2, and VE-cadherin, using the Novartis microarray data set. See Supplemental Data for complete list of organs analyzed for expression of VE-PTP, VEGFR2, and VE-cadherin.


5

TABLE 1. Endothelial cell expression pattern of differentiation-regulated genes Cluster

A B

C

I

J K

Symbol

— ADCY4

Gene name

Function

Adenylate cyclase, type IV

Membrane-bound, calcium-inhibitable adenylyl cyclase CASKIN2 Cask-interacting protein 2 Calcium/calmodulin-dependent serine protein kinase (CASK) has multiple interactive partners in regulation of trafficking, targeting, and signaling of ion channels RAMP2 Receptor activity-modifying protein 2 precursor Transports calcitonin gene-related peptide type 1 receptor (calcrl) to the plasma membrane; acts as a receptor for adrenomedullin (am) together with calcrl PLVAP Plasmalemma vesicle associated protein A component of diaphragmed endothelial fenestrations, up-regulated in tumor endothelium BGN Biglycan precursor A class I small leucine-rich proteoglycan (SLRP) involved in collagen fiber assembly VE-PTP Vascular endothelial-protein tyrosine phosphatase, Phosphotyrosine phosphatase required for vascular also denoted PTPRB, HPTP␤ development TIE-1 Tyrosine-protein kinase receptor Tie-1 precursor TAL1 T-cell acute lymphocytic leukemia 1 Implicated in the genesis of hemopoietic malignancies; plays an important role in endothelial/hematopoietic differentiation CALCRL Calcitionin gene-related type 1 receptor precursor Receptor for calcitonin-gene-related peptide (cgrp) together with ramp1 and receptor for adrenomedullin together with ramp2 or ramp3 (by similarity); activity of this receptor is mediated by g proteins, which activate adenylyl cyclase FLT-1 Vascular endothelial growth factor receptor 1 Receptor tyrosine kinase binding VEGF, VEGF-B, precursor and PLGF; plays a key role in vascular development and contributes to regulation of vascular permeability; a soluble form sFLT-1 is implicated in preeclampsia and serves an inhibitory role in angiogenesis NOTCH4 Neurogenic locus notch homolog protein 4 Receptor for membrane-bound ligands Jagged1, precursor Jagged2, and Delta1 to regulate cell-fate determination; affects the implementation of differentiation, proliferation, and apoptotic programs; may regulate branching morphogenesis in the developing vascular system RAMP2 Receptor activity-modfying protein 2 precursor Transports the calcitonin gene-related peptide type 1 receptor (calcrl) to the plasma membrane; acts as a receptor for adrenomedullin (am) together with calcrl VE-PTP Vascular endothelial protein tyrosine phosphatase Phosphotyrosine phosphatase required for vascular development CYR61 Cysteine-rich, angiogenic inducer, 61 Promotes cell proliferation, chemotaxis, angiogenesis, and cell adhesion; modulates expression of angiogenic regulators such as VEGF, matrix metalloproteinases, and different integrins ROBO4 Roundabout homolog 4 precursor Endothelial-specific member of the roundabout family of guidance molecules SDPR Serum depriviation response A phosphatidylserine-binding protein, which is a PKC substrate GATA2 Endothelial transcription factor GATA-2 Transcriptional activator, which regulates endothelin-1 gene expression in endothelial cells

Genes in clusters A–C and I–K (Fig. 2) were screened for tissue expression patterns similar to those of the endothelial-specific gene VE-cadherin and VEGFR2. No match was seen for any of the genes in cluster A. TAL1 tissue expression did not match those of VE-cadherin or VEGFR2, but is described to be expressed in endothelial progenitor cells in the bone marrow.

tified, such as TIE-1, TAL1, FLT-1, NOTCH4, and GATA2 (Table 1). Another gene included in the list was RAMP2 (receptor activity-modifying protein 2), which recently was shown to be required for proper vascular development during embryogenesis (28). A number of genes not 6

Vol. 23

May 2009

previously coupled to vascular morphogenesis represented diverse functions. Several of these genes appear to be involved in Ca2⫹-dependent processes (ADCY4, CASKIN2, and indirectly, RAMP2, CALCRL, and SDPR). Another interesting gene product is PLVAP, which is a

The FASEB Journal

MELLBERG ET AL.

Figure 4. Real-time PCR validation of VE-PTP transcript regulation. A) Real-time PCR analysis of VE-PTP transcript levels in TIME cells at time points 0, 3, 9, and 18 h in 3D collagen (diamond) or fibronectin (triangle) cultures were compared to the microarray expression data collected at all indicated time points (bars). Changes in transcript levels are expressed as log2 change relative to time point 0 h. For real-time PCR, average change of triplicates is presented with sd. Array data for VE-PTP show an average of 2 different spots, corresponding to different regions of the VE-PTP transcript. B) Real-time PCR analysis of VE-PTP transcript levels in HDMECs cultured on fibronectin or in 3D collagen for different time points as indicated. Cultures were incubated in the presence of VEGF, FGF-2 (both at 50 ng/ml), or 10% FCS. C) Real-time PCR analysis of VE-PTP transcript levels in TIME cells cultured on fibronectin and treated with Src inhibitor PP2, PI3K inhibitor LY294002, or MEK inhibitor U0126, as indicated in Methods and Materials, in the presence and absence of 50 ng/ml of VEGF.

component of diaphragmed endothelial fenestrations (29). VE-PTP (also denoted PTPRB), showed an expression pattern overlapping with those for VE-cadherin and VEGFR2, with expression in the lung in mice, and in cardiac and skeletal myocytes, adult and fetal lung, and placenta in humans (Fig. 3).

lated cells. We conclude that VEGF-induced VE-PTP expression is negatively regulated by the MEK/ERK pathway in proliferating endothelial cells, which may be activated by both Ras and PLC␥ in response to VEGF (see ref. 2 for a review). VE-PTP depletion inhibits tubular morphogenesis

VE-PTP is induced in endothelial cells undergoing tubular morphogenesis VE-PTP was highly expressed in TIME cells undergoing tubular morphogenesis compared to proliferating cells (Fig. 4A). VE-PTP was represented as two different cDNA fragments on the array. One of these was found in cluster B, and a second in cluster C (Fig. 2). Even though the two clones clustered in different groups, their regulation was very similar. VE-PTP transcript regulation in TIME cells harvested from collagen and fibronectin cultures was analyzed by real-time PCR (Fig. 4A), which revealed ⬃7-fold induction in the tube-forming collagen cultures, whereas induction was considerably less in the proliferating fibronectin cultures. VE-PTP transcript regulation in primary HDMECs, showed induction by VEGF and serum, and to some extent also by FGF-2, during tubular morphogenesis (Fig. 4B). To understand why proliferating endothelial cells did not allow up-regulation of VE-PTP, we analyzed induction in the presence and absence of pharmacological inhibitors of the Src pahway (PP2), the PI3K pathway (LY294002), and the MEK/ERK pathway (U0126). As shown in Fig. 4C, attenuation of MEK activity considerably enhanced the level of VE-PTP produced in VEGF-stimu-

In TIME cells treated with VE-PTP siRNA to silence its expression, VEGF-induced tubular morphogenesis in 3D collagen was arrested (Fig. 5A–D). The loss in ability to organize in tubular structures was accompanied by a distinct VEGFR2 immunostaining (compare Fig. 5C, D with control siRNA transfected cells in Fig. 5G, H), which at least in part colocalized with immunostaining for the Golgi marker Golgin 97 (data not shown). VE-cadherin, which was analyzed in parallel, was still localized to cell-cell junctions in the VE-PTP-depleted cells. Quantification of tube length in the presence of VE-PTP siRNA showed a significant decrease to ⬃20% of that in control siRNA-treated cells (Fig. 5I). Figure 5J shows the efficient reduction in VE-PTP expression by the siRNA treatment, also to 20% of that in control cells. Although silencing of VE-PTP disrupted tubular morphogenesis, there was no sign of increased cellular apoptosis. Instead, the morphogenesis-associated G0/G1 arrest was overcome, as evidenced by a significant increase in the number of cells in the S ⫹ G2/M phase in cells treated with VE-PTP siRNA (Fig. 5K). The loss of growth arrest typical for cells engaged in tubular morphogenesis offers a mechanism for the failure of the VE-PTP-depleted cells to comply with the morphogenic process.


7

Figure 5. siRNA silencing of VE-PTP interferes with tubular morphogenesis. A–H) TIME cells were transfected with VE-PTP siRNA (A–D) or control siRNA (E–H) and subsequently seeded on collagen. Cells in which VE-PTP had been silenced failed to organize in vessel structures, as shown in phalloidinstained cells that outline the cellular cytoskeleton (A, E) and in light microscopy (B, F). Panels C, D and G, H show confocal images of cells fluorescently stained to detect VE-cadherin (red) and VEGFR2 (green). Hoechst 33342 (blue) was used to visualize nuclei. Panels D and H show details at higher magnification. Scale bars ⫽ 100 ␮m (A, B, E, F); 10 ␮m (C, D, G, H). I) Quantification of tube length in the different conditions. J) Quantification by real-time PCR of the levels of VE-PTP transcripts in cells treated with VE-PTP or control siRNA. K) Cell cycle analysis by flow sorting of TIME cells either untreated, or treated with control siRNA or VE-PTP siRNA and seeded in 3D collagen for 24 h in the presence of VEGF.

VE-PTP silencing promotes VEGFR2 activity and signaling To determine if VE-PTP silencing directly affected VEGFR2 activity, we studied the effect on tyrosine phosphorylation of specific VEGFR2 sites (2). As shown in Fig. 6A, a short pulse of VEGF on TIME cell monolayers induced tyrosine phosphorylation of 3 autophosphorylation sites: Y951, Y1175, and Y1214. The degree of phosphorylation increased further in VEGFstimulated cells treated with VE-PTP siRNA with the exception of Y1214. The VEGFR2 expression levels were unchanged in the different conditions. We conclude that VE-PTP silencing was accompanied by increased VEGFR2 tyrosine phosphorylation. Immunoblotting for downstream signal transducers and their tyrosine phosphorylated counterparts 8

Vol. 23

May 2009

showed that PLC␥ and Erk1/2, but not Akt, were activated in TIME cells, in response to the short stimulation with VEGF (Fig. 6B). VE-PTP silencing was accompanied by increased levels of pAkt and pErk already in the unstimulated cells; in the VEGFtreated cells there was a further slight increase in pAkt accumulation in the absence of VE-PTP, compared to control. Similar analyses were performed on TIME cells undergoing VEGF-induced tubular morphogenesis. At 24 h of stimulation, there was increased tyrosine phosphorylation of VEGFR2 at Y951, Y1175, and Y1214 with VEGF, and further increase in phosphorylation at Y951 and Y1175 in cells depleted for VE-PTP (Fig. 6C). VE-PTP-depletion furthermore led to a reproducible but modest increase in VEGFinduced phosphorylation of PLC␥, Akt, and Erk1/2

The FASEB Journal

MELLBERG ET AL.

Figure 6. VEGFR2 activation and signal transduction in VE-PTP-deficient TIME cells. A) TIME cells were either untreated (⫺), or treated with control siRNA (ctrl) or VE-PTP siRNA (VE-PTP) and treated without (basal) or with VEGF for 5 min. Cell lysates were subjected to immunoblotting with antibodies reactive with different phosphorylation sites in VEGFR2. Blotting for VEGFR2 and ␤-catenin shows equal loading of lysates. Figures above lanes indicate quantification of band strengths by densitometric scanning. B) Parallel samples to those in A were immunoblotted for PLC␥, Akt, or Erk1/2 or their tyrosine phosphorylated counterparts. Quantification was done as in A on pPLC␥/PLC␥, pAkt/Akt, and pErk/Erk to allow estimation of activation of the different pathways in the presence and absence of VE-PTP silencing. C) TIME cells treated or not with siRNA as in A were seeded on a 2D collagen gel. Lysates were harvested at time point 0 h (basal) and after 24 h of VEGF stimulation. Samples were processed as in A to detect VEGFR2 tyrosine phosphorylation. D) TIME cells treated or not with siRNA as in A were seeded on a 2D collagen gel and processed as in panel B to detect effects of VE-PTP silencing on downstream signal transducers.

(Fig. 6D). We conclude that both acute and longterm signal transduction via VEGFR2 is enhanced in the absence of VE-PTP. VEGF-regulated complex formation between VE-PTP and VEGFR2 To determine whether VEGFR2 is a substrate for VEPTP, we tested whether the 2 molecules could be coimmunoprecipitated. In agreement with the data by Nawroth et al. (30), we did not detect VEGFR2/VE-PTP complex formation in regular coimmunoprecipitation analysis. We therefore employed new methodology, the PLA, which allows detection of molecular complexes in situ (25). Detection is mediated via rolling circle amplification between adjacent, oligonucleotided-coupled secondary antibodies (see Fig. 7A for schematic outline of the procedure). As shown in Fig. 7B, VE-PTP and VEGFR2 formed complexes in quiescent cells. Stimulation with VEGF induced an initial increase in complex formation, followed by a marked decrease after 5 and 10 min (see Fig. 7C for quantification). At 15 min, the complex formation was back to basal (0 time point) levels. Figure 7D shows that complex formation was not detected when cells were silenced for VE-PTP expression using siRNA. Furthermore, use of irrelevant antibodies detecting antigens not expressed in TIME cells such as the V5 tag or green fluorescent protein (GFP) showed that there was no background VE-PTP IN VEGFR2-INDUCED ENDOTHELIAL MORPHOGENESIS

detection of VEGFR2/VE-PTP complexes (Fig. 7D). Detection of VEGFR2 phosphorylation at Y1175 in a parallel analysis (Fig. 7E) showed an inverse relationship between VE-PTP/VEGFR2 complex formation and VEGFR2 phosphorylation (quantified in Fig. 7F).

DISCUSSION This study identifies a number of genes regulated during different stages of tubular morphogenesis. The 30,000 human cDNA arrays that were utilized represent ⬃25,000 unique genes. Compared to previous studies on gene expression during endothelial tube morphogenesis (9 – 14), our study has the particular aspect that it aims to specifically determine VEGF-regulated changes of genes with preferential expression in endothelial cells, as determined through web-based searches on expression sets in 61 mouse and 79 human tissues. A similar approach was used by Wallgård et al. (15), whose study is focused on endothelial expression per se, rather then gene induction during angiogenesis. Of note, most of the endothelial cell-specific genes identified by us as regulated during tubular morphogenesis (with the exception of PLVAP, BGN, TAL1, FLT-1, CYR61, and GATA2; see Table 1) were also listed by Wallgård et al. Furthermore, certain genes were analyzed by real-time PCR and found to be regulated during tubular morphogenesis. This was true for members of the Delta-like4/Jagged family, which were 9

Figure 7. Complex formation between VE-PTP and VEGFR2 is disrupted by ligand stimulation. A) Schematic outline of the proximity ligation assay showing primary antibodies from different species reactive with the intracellular part of VEGFR2 or VE-PTP. Juxtaposition of secondary, oligonucleotide-ligated antibodies allow a rolling circle amplification detected by a Cy3-labeled probe. B) Detection of VE-PTP/ VEGFR2 complexes (red spots) in situ in TIME cells before and after stimulation with VEGF for different time periods. C) Quantification of the data in panel B. D) Quantification of spots in TIME cells treated with control and VE-PTP siRNA, or detected using irrelevant antibodies against the V5-tag or GFP. E) Immunoblotting to detect tyrosine phosphorylation of VEGFR2 in TIME cells before and after stimulation with VEGF for different time periods. TCL, total cell lysate. F) Quantification shows inverse correlation between level of VEGFR2 tyrosine phosphorylation (pVEGFR2/VEGFR2; red) and VE-PTP/VEGFR2 complex formation (blue).

up-regulated during tubular morphogenesis (data not shown). Recent reports have documented a negative role of Delta-like4 on angiogenic sprouting (31), and therefore up-regulation during tubular morphogenesis may serve to control and modulate the morphogenic process. Certain of the genes shown in Table 1 are functionally linked, which lends further credibility to our experimental strategy. This was true for the calcitonin-gene-related peptide (CALCRL) and receptor activity-modifying protein 2 precursor (RAMP2) (Table 1). CALCRL and RAMP2 together form a receptor for adrenomedullin, a peptide involved in circulatory homeostasis and in the pathophysiology of certain cardiovascular diseases (32). CALCRL/RAMP2 are required for adrenomedullin-induced tubular morphogenesis of HUVECs (33). Gene targeting of adrenomedullin (34) as well as RAMP2 (28) leads to death of the embryo during midgestation, due to 10

Vol. 23

May 2009

vascular fragility, severe edema, and hemorrhage. The phenotypes of these knockout mice agree with a role for adrenomedullin and RAMP2 in formation or stability of the vascular tube. One of the genes that fulfilled the criteria of being regulated during VEGF-induced tubular morphogenesis and preferentially expressed in endothelial cells was the receptor-type phosphatase VE-PTP. Phosphatases constitute an interesting class of enzymes that offer means of both positive and negative regulation of growth factor effects. At least 10 different receptor-type PTPs and 8 nonreceptor-type PTPs are expressed in endothelial cells (35). VE-PTP (alternatively denoted PTPRB and HPTP␤ in humans) is specifically expressed in the vasculature throughout development (36), with a preferential expression in arterial endothelial cells (37). VE-PTP, for which no ligand has been

The FASEB Journal

MELLBERG ET AL.

Figure 8. Schematic outline of dynamic VEPTP/VEGFR2 complex formation and the consequence of reduced VE-PTP expression. A) In resting endothelial cells, VE-PTP and VEGFR2 form a complex (left). This is interrupted by binding of VEGF to VEGFR2, which leads to receptor dimerization, activation (middle; indicated as phosphorylation at the three major sites Y951, Y1175, and Y1214), and signal transduction (pAkt och pErk formation).VE-PTP/VEGFR2 complex formation resumes with time, leading to dephosphorylation and deactivation of VEGFR2 (right). B) In VE-PTP-deficient cells, such as after siRNA treatment, VEGFR2 activation (middle) is exaggerated, leading to increased phosphorylation at the Y951 and Y1175 phosphorylation sites (indicated by larger text; P) and deregulated signaling (right). This in turn leads to increased proliferation and decreased angiogenic sprouting.

described, is structurally related to density enhanced phosphatase (DEP)-1 with extracellular fibronectin type III repeats and a single intracellular phosphatase domain. Like DEP-1, VE-PTP is up-regulated in conjunction with increasing cell density implying a role in contact-inhibition of growth (38). Interestingly, depletion of DEP-1 leads to increased tyrosine phosphorylation of VEGFR2 with consequence for specific downstream Src-dependent signaling pathways (39). Gene targeting of VE-PTP is compatible with early development of the vasculature but angiogenesis is abnormal (37, 40). The veptp⫺/⫺ mice die around embryonic day 10, with a vascular phenotype especially pronounced in the yolk sac, characterized by failure to remodel the primitive plexus. The vascular failure of the veptp⫺/⫺ mice concurs with our data showing that VE-PTP silencing was accompanied by exaggerated VEGFR2 activity and unscheduled signaling, overcoming the growth arrest of endothelial cells engaged in tubular morphogenesis. Using proximity ligation to detect endogenous molecular complexes, we show that VE-PTP is associated with VEGFR2 in resting cells. Strikingly, VEGF-induced increase in phosphorylation level and activation of VEGFR2 coincided with markedly reduced VE-PTP/ VEGFR2 interaction. With time, complex formation resumed, parallel to loss of receptor phosphorylation and activity. These data indicate that activation of the

receptor may require dissociation from VE-PTP (see Fig. 8 for a schematic summary). Our data do not strongly argue for a preferential gain of phosphorylation at particular phosphorylation sites in the absence of VE-PTP, although we noted that Y1214 was relatively unaffected by VE-PTP silencing (see Fig. 6). Whereas the roles of pY951 and pY1214 still need to be addressed in complex models, knock-in of Phe to replace Y1173 in mice (corresponds to Y1175 in humans) leads to embryonic lethality due to lack of endothelial cell development (41; for a review, see ref. 2). Although complex formation was specifically and reproducibly detected by proximity ligation, we did not detect VE-PTP and VEGFR2 complex formation in the classical coimmunoprecipitation assay, in agreement with earlier reports (30). Possibly, the chemical composition of the VE-PTP/VEGFR2 complexes does not withstand the coimmunoprecipitation procedure. Whether other aspects of complex organization is differentially detected between the 2 techniques remains to be studied in more detail. Such aspects would include differences in the tolerated distance between molecules in the complex, or the potential participation of other molecular entities, which may lead to loss of detection in coimmunoprecipitation assays but which may possibly be more amenable to detection by proximity ligation. Substrates for VE-PTP include also the angiopoietin receptor Tie-2 and the adherens junction protein VE-


11

cadherin, which both have been found in complex with VE-PTP (30, 36). It is noteworthy in this context that signal transduction and biological readout downstream of Tie-2 and VE-cadherin is complemented by that of VEGF/VEGFR2. For example, the outcome of Tie-2 activation is dictated by the stimulating ligand (angiopoietin-1 or -2) and by the presence of VEGF in the microenvironment (see ref. 42 for a review). Angiopoietin-2-activation of Tie-2 in the absence of VEGF leads to vessel regression, whereas in the presence of VEGF, angiogenesis is induced. VE-cadherin, on the other hand, is known to be tyrosine phosphorylated in response to VEGF/VEGFR2, leading to loosening of adherence junctions, which is a prerequisite for formation of the endothelial cell sprout (for a review, see ref. 43). Thus, it is likely that VE-PTP is crucial in balancing and fine-tuning the state of activity of several key molecules in the angiogenic process.

10.

11.

12.

13.

14.

This study was supported by grants to M.J.C. (Swedish Cancer Society project 4734-B03-02XBB) and to L.C.-W. (Swedish Cancer Society project 3820-B04-09XAC, Swedish Research Council project K2005-32X-12552-08A, and the Novo Nordisk foundation). A.D. was supported by Svenska La¨karsa¨llskapet fo¨r medicinsk forskning and by a grant to L.C.-W. from the Association for International Cancer Research (04-069).

15.

16.

17.

REFERENCES 1. 2. 3.

4.

5. 6.

7.

8.

9.

12

Folkman, J. (1995) Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat. Med. 1, 27–31 Olsson, A. K., Dimberg, A., Kreuger, J., and Claesson-Welsh, L. (2006) VEGF receptor signalling—in control of vascular function. Nat. Rev. Mol. Cell Biol. 7, 359 –371 Shalaby, F., Rossant, J., Yamaguchi, T. P., Gertsenstein, M., Wu, X. F., Breitman, M. L., and Schuh, A. C. (1995) Failure of blood-island formation and vasculogenesis in Flk-1-deficient mice. Nature 376, 62– 66 Holmes, K., Roberts, O. L., Thomas, A. M., and Cross, M. J. (2007) Vascular endothelial growth factor receptor-2: structure, function, intracellular signalling and therapeutic inhibition. Cell. Signal. 19, 2003–2012 Jain, R. K., Duda, D. G., Clark, J. W., and Loeffler, J. S. (2006) Lessons from phase III clinical trials on anti-VEGF therapy for cancer. Nat. Clin. Pract. Oncol. 3, 24 – 40 Matsumoto, T., Schiller, P., Dieterich, L. C., Bahram, F., Iribe, Y., Hellman, U., Wikner, C., Chan, G., Claesson-Welsh, L., and Dimberg, A. (2008) Ninein is expressed in the cytoplasm of angiogenic tip-cells and regulates tubular morphogenesis of endothelial cells. Arterioscler. Thromb. Vasc. Biol. 28, 2123–2130 Dimberg, A., Rylova, S., Dieterich, L. C., Olsson, A. K., Schiller, P., Wikner, C., Bohman, S., Botling, J., Lukinius, A., Wawrousek, E. F., and Claesson-Welsh, L. (2008) alphaB-crystallin promotes tumor angiogenesis by increasing vascular survival during tube morphogenesis. Blood 111, 2015–2023 Bohman, S., Matsumoto, T., Suh, K., Dimberg, A., Jakobsson, L., Yuspa, S., and Claesson-Welsh, L. (2005) Proteomic analysis of vascular endothelial growth factor-induced endothelial cell differentiation reveals a role for chloride intracellular channel 4 (CLIC4) in tubular morphogenesis. J. Biol. Chem. 280, 42397– 42404 Gerritsen, M. E., Soriano, R., Yang, S., Zlot, C., Ingle, G., Toy, K., and Williams, P. M. (2003) Branching out: a molecular fingerprint of endothelial differentiation into tube-like structures generated by Affymetrix oligonucleotide arrays. Microcirculation 10, 63– 81

Vol. 23

May 2009

18. 19.

20.

21.

22.

23.

24. 25.

Glesne, D. A., Zhang, W., Mandava, S., Ursos, L., Buell, M. E., Makowski, L., and Rodi, D. J. (2006) Subtractive transcriptomics: establishing polarity drives in vitro human endothelial morphogenesis. Cancer Res. 66, 4030 – 4040 Bell, S. E., Mavila, A., Salazar, R., Bayless, K. J., Kanagala, S., Maxwell, S. A., and Davis, G. E. (2001) Differential gene expression during capillary morphogenesis in 3D collagen matrices: regulated expression of genes involved in basement membrane matrix assembly, cell cycle progression, cellular differentiation and Gprotein signaling. J. Cell Sci. 114, 2755–2773 Hahn, C. N., Su, Z. J., Drogemuller, C. J., Tsykin, A., Waterman, S. R., Brautigan, P. J., Yu, S., Kremmidiotis, G., Gardner, A., Solomon, P. J., Goodall, G. J., Vadas, M. A., and Gamble, J. R. (2005) Expression profiling reveals functionally important genes and coordinately regulated signaling pathway genes during in vitro angiogenesis. Physiol. Genomics 22, 57– 69 Ho, M., Yang, E., Matcuk, G., Deng, D., Sampas, N., Tsalenko, A., Tabibiazar, R., Zhang, Y., Chen, M., Talbi, S., Ho, Y. D., Wang, J., Tsao, P. S., Ben-Dor, A., Yakhini, Z., Bruhn, L., and Quertermous, T. (2003) Identification of endothelial cell genes by combined database mining and microarray analysis. Physiol. Genomics 13, 249 –262 Kahn, J., Mehraban, F., Ingle, G., Xin, X., Bryant, J. E., Vehar, G., Schoenfeld, J., Grimaldi, C. J., Peale, F., Draksharapu, A., Lewin, D. A., and Gerritsen, M. E. (2000) Gene expression profiling in an in vitro model of angiogenesis. Am. J. Pathol. 156, 1887–1900 Wallgard, E., Larsson, E., He, L., Hellstrom, M., Armulik, A., Nisancioglu, M. H., Genove, G., Lindahl, P., and Betsholtz, C. (2008) Identification of a core set of 58 gene transcripts with broad and specific expression in the microvasculature. Arterioscler. Thromb. Vasc. Biol. 28, 1469 –1476 Venetsanakos, E., Mirza, A., Fanton, C., Romanov, S. R., Tlsty, T., and McMahon, M. (2002) Induction of tubulogenesis in telomerase-immortalized human microvascular endothelial cells by glioblastoma cells. Exp. Cell. Res. 273, 21–33 Vindelov, L. L., Christensen, I. J., Keiding, N., Spang-Thomsen, M., and Nissen, N. I. (1983) Long-term storage of samples for flow cytometric DNA analysis. Cytometry 3, 317–322 Ameur, A., Yankovski, V., Enroth, S., Spjuth, O., and Komorowski, J. (2006) The LCB data warehouse. Bioinformatics 22, 1024 –1026 Saal, L. H., Troein, C., Vallon-Christersson, J., Gruvberger, S., Borg, A., and Peterson, C. (2002) BioArray Software Environment (BASE): a platform for comprehensive management and analysis of microarray data. Genome Biol. 3, software/ 0003.1– 6 Yang, Y. H., Dudoit, S., Luu, P., Lin, D. M., Peng, V., Ngai, J., and Speed, T. P. (2002) Normalization for cDNA microarray data: a robust composite method addressing single and multiple slide systematic variation. Nucleic Acids Res. 30, e15 Ashburner, M., Ball, C. A., Blake, J. A., Botstein, D., Butler, H., Cherry, J. M., Davis, A. P., Dolinski, K., Dwight, S. S., Eppig, J. T., Harris, M. A., Hill, D. P., Issel-Tarver, L., Kasarskis, A., Lewis, S., Matese, J. C., Richardson, J. E., Ringwald, M., Rubin, G. M., and Sherlock, G. (2000) Gene ontology: tool for the unification of biology. The Gene Ontology Consortium. Nat. Genet. 25, 25–29 Su, A. I., Wiltshire, T., Batalov, S., Lapp, H., Ching, K. A., Block, D., Zhang, J., Soden, R., Hayakawa, M., Kreiman, G., Cooke, M. P., Walker, J. R., and Hogenesch, J. B. (2004) A gene atlas of the mouse and human protein-encoding transcriptomes. Proc. Natl. Acad. Sci. U. S. A. 101, 6062– 6067 Hubbard, T., Barker, D., Birney, E., Cameron, G., Chen, Y., Clark, L., Cox, T., Cuff, J., Curwen, V., Down, T., Durbin, R., Eyras, E., Gilbert, J., Hammond, M., Huminiecki, L., Kasprzyk, A., Lehvaslaiho, H., Lijnzaad, P., Melsopp, C., Mongin, E., Pettett, R., Pocock, M., Potter, S., Rust, A., Schmidt, E., Searle, S., Slater, G., Smith, J., Spooner, W., Stabenau, A., Stalker, J., Stupka, E., UretaVidal, A., Vastrik, I., and Clamp, M. (2002) The Ensembl genome database project. Nucleic Acids Res. 30, 38 – 41 Altschul, S. F., and Lipman, D. J. (1990) Protein database searches for multiple alignments. Proc. Natl. Acad. Sci. U. S. A. 87, 5509 –5513 Soderberg, O., Gullberg, M., Jarvius, M., Ridderstrale, K., Leuchowius, K. J., Jarvius, J., Wester, K., Hydbring, P., Bahram, F.,

The FASEB Journal

MELLBERG ET AL.

26. 27.

28.

29. 30.

31.

32.

33.

34.

Larsson, L. G., and Landegren, U. (2006) Direct observation of individual endogenous protein complexes in situ by proximity ligation. Nat. Methods 3, 995–1000 Sturn, A., Quackenbush, J., and Trajanoski, Z. (2002) Genesis: cluster analysis of microarray data. Bioinformatics 18, 207–208 Porter, C. W., Ganis, B., Libby, P. R., and Bergeron, R. J. (1991) Correlations between polyamine analogue-induced increases in spermidine/spermine N1-acetyltransferase activity, polyamine pool depletion, and growth inhibition in human melanoma cell lines. Cancer Res. 51, 3715–3720 Ichikawa-Shindo, Y., Sakurai, T., Kamiyoshi, A., Kawate, H., Iinuma, N., Yoshizawa, T., Koyama, T., Fukuchi, J., Iimuro, S., Moriyama, N., Kawakami, H., Murata, T., Kangawa, K., Nagai, R., and Shindo, T. (2008) The GPCR modulator protein RAMP2 is essential for angiogenesis and vascular integrity. J. Clin. Invest. 118, 29 –39 Niemela, H., Elima, K., Henttinen, T., Irjala, H., Salmi, M., and Jalkanen, S. (2005) Molecular identification of PAL-E, a widely used endothelial-cell marker. Blood 106, 3405–3409 Nawroth, R., Poell, G., Ranft, A., Kloep, S., Samulowitz, U., Fachinger, G., Golding, M., Shima, D. T., Deutsch, U., and Vestweber, D. (2002) VE-PTP and VE-cadherin ectodomains interact to facilitate regulation of phosphorylation and cell contacts. EMBO J. 21, 4885– 4895 Hellstrom, M., Phng, L. K., Hofmann, J. J., Wallgard, E., Coultas, L., Lindblom, P., Alva, J., Nilsson, A. K., Karlsson, L., Gaiano, N., Yoon, K., Rossant, J., Iruela-Arispe, M. L., Kalen, M., Gerhardt, H., and Betsholtz, C. (2007) Dll4 signalling through Notch1 regulates formation of tip cells during angiogenesis. Nature 445, 776 –780 Ishimitsu, T., Ono, H., Minami, J., and Matsuoka, H. (2006) Pathophysiologic and therapeutic implications of adrenomedullin in cardiovascular disorders. Pharmacol. Ther. 111, 909 –927 Fernandez-Sauze, S., Delfino, C., Mabrouk, K., Dussert, C., Chinot, O., Martin, P. M., Grisoli, F., Ouafik, L., and Boudouresque, F. (2004) Effects of adrenomedullin on endothelial cells in the multistep process of angiogenesis: involvement of CRLR/ RAMP2 and CRLR/RAMP3 receptors. Int. J. Cancer 108, 797– 804 Shindo, T., Kurihara, Y., Nishimatsu, H., Moriyama, N., Kakoki, M., Wang, Y., Imai, Y., Ebihara, A., Kuwaki, T., Ju, K. H., Minamino, N., Kangawa, K., Ishikawa, T., Fukuda, M., Akimoto,


35. 36. 37.

38.

39.

40.

41.

42. 43.

Y., Kawakami, H., Imai, T., Morita, H., Yazaki, Y., Nagai, R., Hirata, Y., and Kurihara, H. (2001) Vascular abnormalities and elevated blood pressure in mice lacking adrenomedullin gene. Circulation 104, 1964 –1971 Kappert, K., Peters, K. G., Bohmer, F. D., and Ostman, A. (2005) Tyrosine phosphatases in vessel wall signaling. Cardiovasc. Res. 65, 587–598 Fachinger, G., Deutsch, U., and Risau, W. (1999) Functional interaction of vascular endothelial-protein-tyrosine phosphatase with the angiopoietin receptor Tie-2. Oncogene 18, 5948 –5953 Baumer, S., Keller, L., Holtmann, A., Funke, R., August, B., Gamp, A., Wolburg, H., Wolburg-Buchholz, K., Deutsch, U., and Vestweber, D. (2006) Vascular endothelial cell-specific phosphotyrosine phosphatase (VE-PTP) activity is required for blood vessel development. Blood 107, 4754 – 4762 Gaits, F., Li, R. Y., Ragab, A., Ragab-Thomas, J. M., and Chap, H. (1995) Increase in receptor-like protein tyrosine phosphatase activity and expression level on density-dependent growth arrest of endothelial cells. Biochem. J. 311(Pt. 1), 97–103 Chabot, C., Spring, K., Gratton, J. P., Elchebly, M., and Royal, I. (2009) New role for the protein tyrosine phosphatase DEP-1 in Akt activation and endothelial cell survival. Mol. Cell Biol. 29, 241–253 Dominguez, M. G., Hughes, V. C., Pan, L., Simmons, M., Daly, C., Anderson, K., Noguera-Troise, I., Murphy, A. J., Valenzuela, D. M., Davis, S., Thurston, G., Yancopoulos, G. D., and Gale, N. W. (2007) Vascular endothelial tyrosine phosphatase (VEPTP)-null mice undergo vasculogenesis but die embryonically because of defects in angiogenesis. Proc. Natl. Acad. Sci. U. S. A. 104, 3243–3248 Sakurai, Y., Ohgimoto, K., Kataoka, Y., Yoshida, N., and Shibuya, M. (2005) Essential role of Flk-1 (VEGF receptor 2) tyrosine residue 1173 in vasculogenesis in mice. Proc. Natl. Acad. Sci. U. S. A. 102, 1076 –1081 Eklund, L., and Olsen, B. R. (2006) Tie receptors and their angiopoietin ligands are context-dependent regulators of vascular remodeling. Exp. Cell Res. 312, 630 – 641 Cavallaro, U., Liebner, S., and Dejana, E. (2006) Endothelial cadherins and tumor angiogenesis. Exp. Cel. Res. 312, 659 – 667 Received for publication October 23, 2008. Accepted for publication December 18, 2008.

13