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The lymphatic vascular system maintains tissue fluid homeostasis, helps mediate afferent immune responses, and promotes cancer metastasis. To address the role microRNAs (miRNAs) play in the development and function of the lymphatic vascular system, we defined the in vitro miRNA expression profiles of primary human lymphatic endothelial cells (LECs) and blood vascular endothelial cells (BVECs) and identified four BVEC signature and two LEC signature miRNAs. Their vascular lineage-specific expression patterns were confirmed in vivo by quantitative real-time PCR and in situ hybridization. Functional characterization of the BVEC signature miRNA miR-31 identified a novel BVEC-specific posttranscriptional regulatory mechanism that inhibits the expression of lymphatic lineage-specific transcripts in vitro. We demonstrate that suppression of lymphatic differentiation is partially mediated via direct repression of PROX1, a transcription factor that functions as a master regulator of lymphatic lineage-specific differentiation. Finally, in vivo studies of Xenopus and zebrafish demonstrated that gain of miR-31 function impaired venous sprouting and lymphatic vascular development, thus highlighting the importance of miR-31 as a negative regulator of lymphatic development. Collectively, our findings identify miR-31 is a potent regulator of vascular lineage-specific differentiation and development in vertebrates.
Vertebrates have developed two parallel but structurally and functionally distinct vascular systems: the blood and lymphatic vascular systems (1, 7). The lymphatic vascular system controls tissue fluid homeostasis, absorbs lipids and fat-soluble vitamins from the intestine, and mediates afferent immune responses by transporting lymphocytes and antigen-presenting cells to regional lymph nodes (1, 7). In addition, malignant cancers can induce lymphatic vessel activation and growth (lymphangiogenesis) within primary tumors and draining lymph nodes, which enhances cancer metastasis to draining lymph nodes and beyond (1, 22). These findings have fueled a surge in studies aimed at defining the molecular characteristics and functional activities of lymphatic vessels and identifying molecules that regulate lymphangiogenesis.
Genomic and proteomic studies have identified novel molecular markers and growth factors for lymphatic vessels (2, 23, 48, 52). Mouse genetic models have characterized the transcription factors PROX1 and SOX18 as master regulators of lymphatic vascular development and differentiation in vivo (12, 56, 65). These studies indicate that SOX18 expression in a subset of cardinal vein endothelial cells initiates lymphatic vascular development by inducing PROX1 expression (12). The resulting lymphatic vascular progenitor cells bud off and migrate away from the cardinal vein and form primitive lymph sacs, which subsequently develop into functional lymphatics (12, 56). PROX1 and SOX18 expression in cultured blood vascular endothelial cells (BVECs) triggers these cells to adopt lymphatic lineage-specific molecular and phenotypic characteristics (12, 26, 48). Conversely, PROX1 knockdown in lymphatic endothelial cells (LECs) inhibits the expression of LEC signature genes and triggers BVEC signature gene expression (44; J. W. Shin et al., unpublished data). Despite these advances, a detailed understanding of the mechanisms controlling lymphatic vascular development and cell type-specific differentiation remains elusive.
A potentially crucial aspect of lymphatic vascular biology has remained unexplored to date, i.e., the role of microRNA (miRNA)-guided posttranscriptional regulation. miRNAs are genomically encoded 19- to 24-nucleotide (nt) noncoding RNAs that regulate the flow of genetic information by limiting protein synthesis (10). This regulation is brought about when mature miRNAs, loaded in the RNA-induced silencing complex, base pair with semicomplementary sites within the 3′ untranslated region (UTR) of target mRNAs. Once base paired with its target, the miRNA represses translation and/or induces mRNA degradation (10). Consequently, miRNAs act as novel and potent regulators of the genome. This notion is underscored by recent studies defining critical roles for miRNAs in embryonic development, cell proliferation, cell cycle progression, differentiation, and apoptosis, as well as their contribution to the etiology of several diseases (10, 46, 68).
Interestingly, functional roles for miRNAs in blood vascular development have recently been defined. Downregulation of the miRNA processing enzymes Dicer and Drosha has been reported to impair angiogenesis (11, 59). Moreover, a few miRNAs have been shown to affect human umbilical vein endothelial cell (HUVEC) migration and proliferation in vitro, regulate nitric oxide synthase expression, promote tumor angiogenesis, control vascular inflammation, and directly contribute to numerous vascular phenotypes (11, 59).
In the study presented here, we identified and addressed the functional relevance of vascular lineage-specific miRNAs. We first defined the miRNA expression profiles of primary human LECs and BVECs and consequently identified four BVEC and two LEC signature miRNAs. Their vascular lineage-specific expression was confirmed in normal tissues by quantitative real-time PCR (qRT-PCR) analysis of ex vivo-isolated murine LECs and BVECs and by in situ hybridization (ISH). Interestingly, our findings have further classified the widely expressed (38) metastasis-associated (63, 64) miRNA miR-31 as a BVEC signature miRNA. In vitro functional analysis of miR-31 demonstrated that this miRNA inhibits lymphatic lineage-specific differentiation in BVECs by repressing lymphatic lineage-specific transcript levels. These effects are, in part, due to direct posttranscriptional repression of Prox1, a master regulator of lymphatic development. Finally, in vivo gain-of-function studies with Xenopus and zebrafish embryos established that overexpression of miR-31 impaired lymphatic development and reduced venous sprouting. Taken together, these findings indicate that miR-31 plays a pivotal role in regulating lineage-specific differentiation within the developing vasculature of vertebrates.
Primary human dermal microvascular LECs and BVECs were isolated from neonatal human foreskins and cultured as previously described (23). LECs and HUVECs were purchased from Cambrex (Verviers, Belgium). IMR91 human dermal fibroblasts (hdFBs) were obtained from the National Institute on Aging, Bethesda, MD. The immortalized human epidermal keratinocyte line HaCaT was provided by Norbert Fusenig, German Cancer Research Center, Heidelberg, Germany (4). Cells, except hdFBs, were propagated in supplemented endothelial cell basal medium (EBM; Cambrex) as previously described (23). hdFBs were propagated in Dulbecco's modified Eagle medium supplemented as described above and transferred 12 h prior to total RNA isolation to EBM supplemented as described above. Primary cells were used at passage 6.
The TaqMan microRNA Assays Human Panel Early Access kit (Applied Biosystems, Foster City, CA), containing 157 individual human TaqMan microRNA assays, was used for qRT-PCR miRNA expression profiling (5). Total RNA was isolated from biological replicates of 80 to 90% confluent 10-cm tissue culture dishes using the mirVana miRNA isolation kit (Ambion, Austin, TX). Reverse transcription reactions were performed using 2 ng of total RNA and the microRNA Reverse Transcription kit (Applied Biosystems). miRNA expression levels of technical duplicates were determined using a 7900HT Fast Real-Time PCR System (Applied Biosystems), and comparative threshold cycle (CT) values were acquired after 40 cycles using SDS 2.2 software (Applied Biosystems). TaqMan microRNA assays (Applied Biosystems) for hsa-miR-31, hsa-miR-137, hsa-miR-99a, hsa-miR-125b, hsa-miR-95, hsa-miR-326, and human RNU48 were used to confirm lineage-specific expression.
For analysis, detection thresholds were set to 0.04 U of fluorescence intensity, and when a miRNA CT value was undetermined in both technical replicates, a CT value of 41 was assigned. Data sets were normalized relative to let-7a and miR-16 using the formula Ave CTNORM = Ave CTmiRNA − (Ave CTlet-7a/miR-16 − 24), where Ave CTlet-7a/miR-16 is the combined average CT value for let-7a and miR-16 from each 96-well plate. RNU48 or sno234 was used to normalize the individual TaqMan microRNA assay data sets using the formula Ave CTNORM = Ave CTmiRNA − (Ave CTRNU48 or sno234 − 25), where Ave CTRNU48 or sno234 is the mean RNU48 or sno234 CT value (n = 3). Relative abundances of LECs and BVECs were calculated from log2 ratios. P values were calculated using a two-tailed Student t test.
Experiments with mice were approved by the Kantonales Veterinäramt Zürich. Colons were excised from sacrificed female FVB mice (12 to 16 weeks old, n = 8; Charles River, Sulzbach, Germany), opened longitudinally, washed in cold phosphate-buffered saline (PBS), and placed in 1 mM dithiothreitol. Mucus was gently removed by scraping. Small tissue pieces were digested with 8 mg/ml collagenase IV (Invitrogen, Carlsbad, CA)-0.5 mg/ml DNase I (Roche, Rotkreuz, Switzerland)-5 mM CaCl2 in PBS at 37°C for 15 min. After passing through a 70-μm cell strainer (BD Biosciences, Franklin Lakes, NJ), the resulting cell suspensions were centrifuged at 500 × g for 10 min and resuspended in 2% fetal bovine serum-supplemented PBS containing 1 mM EDTA.
The antibodies used for FACS sorting were allophycocyanin-conjugated rat anti-mouse CD31 (BD Biosciences Pharmingen, San Diego, CA), fluorescein isothiocyanate-conjugated rat anti-mouse CD45.2 (BD Biosciences), hamster anti-mouse podoplanin (clone 8.1.1; Developmental Studies Hybridoma Bank, Iowa City, IA), and phycoerythrin-conjugated anti-hamster (CALTAG/Invitrogen) and isotype control antibodies. FACS sorting was performed using a FACSAria and the FACSDiva software (BD Biosciences). Cells were lysed by sorting directly into RLT Plus lysis buffer (Qiagen, Hilden, Germany) containing β-mercaptoethanol. Total RNA was extracted from BVECs (CD45− CD31+ podoplanin−) and LECs (CD45− CD31+ podoplanin+) using the RNeasy Plus Micro kit (Qiagen, Hilden, Germany). For miRNA expression analyses, 6 ng of total RNA and TaqMan miRNA assays for mmu-miR-31, mmu-miR-326, hsa-miR-137, hsa-miR-99a, hsa-miR-125b, and mouse sno234 were used.
miR-31 ISH and Lyve-1/CD31 immunofluorescence staining were performed with 20-μm serial frozen colon sections obtained from female FVB mice. ISH for mouse miR-31 was performed using digoxigenin (DIG)-labeled locked nucleic acid (LNA)-modified detection probes (mmu-miR-31 [catalog no. 39153-00], hsa/mmu/rno-U6 [positive control, catalog no. 99002-00], sense miR-159 [negative control, catalog no. 99003-00]; Exiqon, Vedbæk, Denmark) and the formaldehyde-EDC fixation miRNA ISH protocol (47). Briefly, the LNA-modified detection probes were labeled with DIG using the DIG Oligonucleotide Tailing kit (Roche, Basel, Switzerland) according to the manufacturer's instructions. Tissue sections were fixed in 4% formaldehyde-Tris-buffered saline for 10 min and then in EDC solution (47) for 1.5 h. The sections were acetylated in 1% triethanolamine-0.25% acetic anhydride, washed, and prehybridized in hybridization buffer (47) for 1 h at 53°C. The colon tissue sections were hybridized with 4 μM DIG-labeled detection probes overnight at 56°C for miR-31 and 53°C for the controls. Following posthybridization washing and blocking, the slides were probed with alkaline phosphatase-conjugated anti-DIG Fab fragments (Roche). They were then washed in TNT buffer (100 mM Tris-HCl [pH 7.5], 150 mM NaCl, 0.1% Tween 20) and in AP buffer (100 mM Tris-HCl [pH 9.5], 100 mM NaCl, 50 mM MgCl2). Color development was performed with developer solution (AP buffer with 0.175 mg/ml 5-bromo-4-chloro-3-indolylphosphate [BCIP], 0.45 mg/ml Nitro Blue Tetrazolium, and 2 mM levamisol). All incubations and washing steps were performed at room temperature unless otherwise indicated.
Immunofluorescence staining was performed as previously described (25, 35), using a rabbit polyclonal antibody against mouse Lyve-1 (AngioBio, Del Mar, CA), a monoclonal rat antibody against mouse CD31 (BD Biosciences), and corresponding secondary antibodies labeled with Alexa Fluor 488 or Alexa Fluor 594 (Molecular Probes). Sections were examined on an Axioskop2 microscope (Carl Zeiss, Feldbach, Switzerland), and images were captured at magnifications of ×2.5 (Plan-Neofluar 2.5x, 0.075 numerical aperture) and ×20 (Plan-Neofluar 20x, 0.50 Ph2) with an AxioCam MRm digital camera (Zeiss). Bright-field and fluorescent channel image acquisition was accomplished using Axio Vision 4.4 software (Zeiss). Adobe Photoshop CS3 (Adobe Systems, San Jose, CA) was used to adjust image brightness.
All transfections were carried out using the Basic Nucleofector kit for primary mammalian endothelial cells (Amaxa AG, Cologne, Germany). Five hundred thousand LECs were transfected with 2 μM pre-miR-31 or pre-miR-Neg molecules (30) in biological duplicate, and total RNA was isolated using the mirVana isolation kit at 48 h posttransfection. The transcriptome profiles of these cells were defined using the Applied Biosystems Human Genome Survey Microarray v2.0 as previously described (54). Briefly, DIG-UTP-labeled cRNA was generated from 1.5 μg of total RNA using the NanoAmp RT-IVT Labeling kit (Applied Biosystems). Twenty micrograms of cRNA was fragmented and hybridized to the microarrays using the Applied Biosystems Chemiluminescence Detection kit. Signal detection, image acquisition, and initial analyses were performed using the Applied Biosystems 1700 Chemiluminescent Microarray Analyzer.
Raw data were normalized using Quantile normalization available from R/Bioconductor (14). Present calls were defined based on average signal-to-noise ratios of >3 and quality (error) values of <5,000 (54). Feature signal intensities were converted to log2 values. miR-31-repressed genes were identified based on present calls in both pre-miR-Neg arrays with log2 (Pre31/PreNeg) values of ≤−0.59 and P values of <0.05, while miR-31-induced genes were present in both pre-miR-31 arrays and had log2 (Pre31/PreNeg) values of ≥0.59 and P values of <0.05. P values were calculated using empirical Bayes statistics for differential expression (55).
To confirm the microarray data, the mRNA expression levels of selected candidate miR-31-regulated LEC and BVEC signature genes were analyzed in triplicate by qRT-PCR using dually labeled TaqMan Gene Expression Assays for TIMP3 (Assay ID Hs00165951_g1), PPP1R9A (Hs01044146_m1), HOXD10 (Hs00157974_m1), EDNRB (Hs00240752_m1), PROX1 (Hs00160463_m1), NRCAM (Hs00170554_m1), SELE (Hs00950401_m1), ICAM1 (Hs99999152_m1), MMP1 (Hs00899658_m1), RGS4 (Hs00194501_m1), NRG1 (Hs00247620_m1), and LOC554202 (Hs01007340_m1) (all from Applied Biosystems). The probe and primers for LYVE-1 were as previously described (23). Twenty-five nanograms of cDNA generated using the High Capacity cDNA Archive kit (Applied Biosystems) was used. Each reaction was normalized to β-actin expression (54).
To further characterized miR-31 regulation of PROX1, 500,000 LECs were transfected with 2 or 4 μM pre-miR-31 (n = 4) or pre-miR-Neg (n = 4) molecules or 4 μM anti-miR-31 (n = 2) or anti-miR-Neg (n = 2) molecules (6). Total RNA and whole-cell protein lysates were isolated using the mirVana PARIS kit at 48 h posttransfection. qRT-PCR analysis of PROX1 mRNA was performed as described in Results.
Northern blot analyses were performed with 1 μg total RNA. PROX1 mRNA was detected using purified PROX1 3′ UTR [γ-32P]ATP end-labeled probes generated from a NotI-linearized human PROX1 3′ UTR plasmid (YH1551; provided by Young Kwon Hong, University of Southern California, Los Angeles). The membrane was then stripped, and β-actin was detected using [γ-32P]ATP end-labeled human β-actin oligonucleotides (5′-GTGAGGATCTTCATGAGGTAGTCAGTCAGGT-3′).
For Western blotting, 25-μg protein lysate samples were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. Membranes were probed with rabbit polyclonal anti-human PROX1 (RELIATech, Braunschweig, Germany) and mouse monoclonal anti-human β-actin antibodies (Sigma-Aldrich) detected using horseradish peroxidase-conjugated secondary antibodies and standard chemiluminescence (54). QuantityOne software (Bio-Rad) was used to semiquantitatively analyze the amounts of PROX1 protein relative to β-actin in each sample. Pre-miR-31 and pre-miR-Neg PROX1 signal volume averages and standard deviations were calculated from these normalized values (n = 4). P values were calculated using a two-tailed Student t test.
pMIR-Luci/miR31BS and psiCHECK-2/miR31BS contain perfect-match hsa-miR-31 binding sites (miR31BS). pMIR-Luci/miR31BS was generated by cloning annealed miR31BS sense (5′-AGCTTGCTGAGCGGCAAGATGCTGGCATAGCTGA-3′) and antisense (5′-CTAGTCAGCTATGCCAGCATCTTGCCGCTCAGCA-3′) oligonucleotides into the HindIII and SpeI restriction sites of the pMIR-REPORT Luciferase vector (6). psiCHECK-2/miR31BS was constructed by ligating annealed miR31BS/F and miR31BS/R oligonucleotides (see Table S1 in the supplemental material) into the XhoI and NotI sites of psiCHECK-2 (Promega, Dübendorf, Switzerland). PROX1 3′ UTR and coding sequence (CDS) luciferase reporter vectors (see Table S1 in the supplemental material) were constructed by PCR amplification from YH1551 (for the oligonucleotides used, see Table S1 in the supplemental material). PROX1 3′ UTR amplicons were ligated into the XhoI and NotI sites of psiCHECK-2, and the PROX1-CDS amplicon was ligated into the PmeI and NotI sites of psiCHECK-2.
For miR-31 overexpression optimization, 500,000 LECs were cotransfected with 0.7 μg of pMIR-Luci/miR31BS, 0.7 μg of pMIR-REPORT β-galactosidase (β-Gal) control, and 0.02, 0.2, 1, 2, or 4 μM human miR-31 precursor (pre-miR-31) or pre-miR-Neg negative-control molecules (Applied Biosystems). Luciferase and β-Gal activities were monitored 48 h after transfection using the Dual-Light Luciferase and β-Gal Reporter Gene Assay System (Applied Biosystems). β-Gal RLUs (relative light units) were used to normalize luciferase RLUs.
For the PROX1 3′ UTR tethering and PROX1 3′ UTR miR-31 binding site mutagenesis assays, 500,000 LECs or HUVECs were cotransfected in triplicate with 4 μM miR-31 precursor or inhibitor (6) or the corresponding negative controls and 0.7 μg of the psiCHECK-2 constructs containing the miR-31 binding site (miR31BS), the PROX1 3′ UTR fragments (PROX1 FL-F6), the PROX1 CDS, or the PROX1 3′ UTR miR-31 binding site mutants (PROX1 FL-mut or PROX1 F6-mut) (see Table S1 in the supplemental material). Firefly and Renilla luciferase activities were monitored 48 h after transfection using the Dual-Luciferase Reporter Assay System (Promega). PROX1 3′ UTR/CDS-Renilla luciferase RLUs were normalized to firefly luciferase RLUs. Both the dual-light and dual-luciferase assays were performed in triplicate with 20 μl of cell lysate.
Xenopus studies were conducted under protocols approved by the Veterinary Office of the Canton of Zürich, Switzerland. Xenopus laevis eggs were obtained by hormone-induced laying, fertilized in vitro, and prepared for microinjection as previously described (21). Two-cell-stage embryos were unilaterally microinjected with pre-miR-31 or pre-miR-Neg molecules (10 to 100 ng/blastomere) and 0.2 ng β-Gal RNA (lineage tracer). The antisense VEGFC morpholino oligomer (MO) (5′-GTAACGCTCCCTCCAGCAAGTACAT-3′) was purchased from Gene Tools (Philomath, OR), and 5 to 10 ng was unilaterally injected into two-cell-stage embryos. When uninjected embryos reached the embryonic stage indicated, the injected embryos were fixed and processed for ISH. Whole-mount ISH, β-Gal staining, and bleaching of Xenopus embryos were carried out as previously described (29). DIG-labeled probes were transcribed from linearized plasmids encoding Xenopus pecam1 (29), prox1 (GenBank accession no. BU903551), and vegfr3 (28). Images were acquired digitally using AxioVision 4.5 (Zeiss) software and an AxioCam color camera (Zeiss) mounted on a Zeiss Stereo Lumar V12 stereoscopic microscope.
Transgenic TG(fli1a:gfp)y1 (37) and plcg1 t26480 zebrafish lines were maintained at the Hubrecht Institute. Zebrafish experiments were approved by the Animal Experimentation Committee (DEC) of the Royal Netherlands Academy of Arts and Sciences. The plcg1t26480 allele is a W1024X mutation in the plcg1 gene (GenBank accession number AY163168).
MO were ordered from Gene Tools (Philomath, OR). One-cell-stage TG(fli1a:gfp)y1; plcg1t26480 mutant embryos were injected at 40 ng/embryo with an MO targeting dre-miR-31 (5′-TTAACAGCTATGCCAACATCTTGCC-3′) or at 25 ng/embryo with an unrelated control MO (5′-GCATTGACTCTGTAAAACAGACAAT-3′). For miR-31 overexpression, TG(fli1a:gfp)y1; plcg1t26480 mutant embryos were injected with 170 or 340 pg of human pre-miR-31 precursor or pre-miR-Neg control molecules (Applied Biosystems). Venous sprouts were quantified at 48 h postfertilization (hpf), and statistical significance was analyzed using the Student t test. For imaging, embryos were mounted in 0.8% low-melting-point agarose in a dish with a coverslip replacing the bottom. Imaging was performed with a Leica SP2 confocal microscope (Leica Microsystems) using a 20× objective.
The microarray data obtained in this study are accessible at http://www.ncbi.nlm.nih.gov/geo/ under accession no. GSE16908.
Using a TaqMan-based qRT-PCR profiling platform (5), we defined the in vitro expression profiles of 157 human miRNAs in primary LECs and BVECs, as well as two nonendothelial cell types (HaCaT keratinocytes and hdFBs) (see Table S2 in the supplemental material). Following data normalization, expression profiles for each cell type were defined by setting the CT value present call cutoff at 34 (see Table S2 in the supplemental material). Most of the miRNAs analyzed were expressed at comparable levels in both LECs and BVECs (see Fig. S1 in the supplemental material). Nevertheless, based on 2-fold or greater differential expression, 16 candidate LEC and 30 candidate BVEC signature miRNAs were identified (Table (Table1).1). Using a P value cutoff of ≤0.05, two LEC signature miRNAs (miR-95 and miR-326) and four BVEC signature miRNAs (miR-137, miR-31, miR-125b, and miR-99a) were identified (Table (Table1).1). Individual TaqMan miRNA assays confirmed the vascular lineage specificity of these miRNAs. miR-95 and miR-326 expression levels were, on average, 46-fold and 7-fold higher in LECs than in BVECs, respectively (Fig. (Fig.11 A). Conversely, miR-137 expression was 124-fold higher in BVECs than in LECs, miR-31 expression was 48-fold higher in BVECs, and miR-125b and miR99a were 3-fold more abundant in BVECs (Fig. (Fig.1B1B).
We next isolated BVECs and LECs from the colons of eight adult mice by FACS sorting using the leukocyte marker CD45, the panendothelial marker CD31, and the LEC marker podoplanin to differentiate among leukocytes (CD45+ CD31− podoplanin− and CD45+ CD31+ podoplanin−), BVECs (CD45− CD31+ podoplanin−), and LECs (CD45− CD31+ podoplanin+) (18). We obtained 1,500 to 25,000 LECs and 2,500 to 55,000 BVECs, from which total RNA was extracted and used for ex vivo qRT-PCR miRNA profiling. Based on 1.5-fold or greater differential expression, miR-31 was indeed more strongly expressed by BVECs than by LECs in six of the eight mice analyzed (Fig. (Fig.22 A). The degrees of miR-31 differential expression between mouse BVECs and LECs in vivo were less pronounced than those observed between in vitro-cultured human endothelial cells. Nevertheless, statistical analysis of these in vivo data confirmed that the differences in miR-31 expression between mouse BVECs and LECs were statistically significant in six out of the eight mice studied (Fig. (Fig.2A).2A). The LEC signature expression pattern of miR-326 and the BVEC signature classifications of miR-125b and miR-99a were also confirmed, while no major changes were found for miR-137 (see Fig. S2 in the supplemental material). miR-95 is not present in mice.
Low-magnification (×2.5) microscopic analysis of adult mouse colon tissue sections probed for miR-31 expression by ISH revealed strong miR-31 staining throughout the adult mouse colon (Fig. 2B and D). Importantly, immunofluorescence staining of serial sections for the panvascular marker CD31 and the lymphatic marker LYVE-1 revealed that miR-31 preferentially colocalized with blood vessels (CD31+ LYVE1−) present in the submucosa and mesenteric attachments of the colon, as well as the lamina propria (Fig. 2B and D, arrows and arrowheads). In contrast, miR-31 expression was weak or absent in lymphatic vessels (Fig. 2B and D, asterisks). Independent CD31 and LYVE1 immunofluorescence staining and negative-control (sense miR-159) ISH of serial sections of adult colon tissues (Fig. 2H and I), coupled with high-magnification (×20) image analysis of the miR-31 ISH and the CD31/LYVE1 immunofluorescently labeled serial sections demonstrated no preferential staining of either the blood and lymphatic vessels in the sense miR-159-probed sections (Fig. 2H and I). Both vessel types developed equivalent background level signals following ISH processing. In contrast, miR-31 molecules were preferentially associated with colonic blood vessels (CD31+ LYVE1−; arrows), while the lymphatic vessels (CD31+ LYVE1+) developed no miR-31 signals (asterisks) (Fig. 2F and G). Taken together, these in vitro and in vivo data demonstrate that miR-31 transcripts are rare or absent in LECs but enriched in BVECs.
miR-31 is encoded within intron 1 of an uncharacterized gene, LOC554202 (gene ID 554202). This led us to question if miR-31 lineage specificity results from the preferential expression of LOC554202 in BVECs. Indeed, TaqMan-based qRT-PCR analysis demonstrated that LOC554202 transcripts were 54-fold more abundant in BVECs than in LECs (ΔΔCT = 5.78; see Fig. S3A in the supplemental material), which is comparable to the degree of differential expression defined for miR-31 in BVECs versus LECs (ΔΔCT = 5.58). Interestingly, further genome and proteome bioinformatic analyses revealed that the gene is not conserved in vertebrates and the putative gene product has no homology to proteins of known function. Thus, LOC554202 might primarily function as a conduit for miR-31 posttranscriptional regulatory activities in BVECs. Sequence alignments of human LOC554202 with 23 eutherian mammals (Enredo, Pecan, Ortheus) using Ensembl alignment tools (http://www.ensembl.org/index.html) revealed conserved synteny for miR-31 with six of the species queried. Moreover, 13 of the eutherian mammals queried are predicted to contain novel miRNA genes at positions aligned with LOC554202 (data not shown). Further alignment analysis of these 20 miRNA genes demonstrated that these 13 novel miRNA genes are, in fact, miR-31 orthologs (see Fig. S3B in the supplemental material). This highly conserved synteny suggests that the genomic, transcriptional, and epigenetic factors regulating miR-31 expression have remained conserved during mammalian evolution.
The strong differential expression of miR-31 in vitro and in vivo prompted us to further characterize this BVEC signature miRNA by defining the transcriptome profile changes in LECs after miR-31 overexpression. Using a luciferase reporter construct containing a miR-31 binding site (pMIR-Luci/miR31BS) and qRT-PCR, we found that high levels of miR-31 gain of function could be achieved when LECs were transfected with 2 μM pre-miR-31 precursor (30) (see Fig. S4 in the supplemental material). Using ≥1.5-fold differential expression and ≤0.05 P value thresholds, gene microarray analyses of 2 μM pre-miR-31- or pre-miR-Neg-transfected LECs identified 548 miR-31-repressed and 335 miR-31-induced genes (see Tables S3 and S4 in the supplemental material). Comparing the miR-31 targets predicted by TargetScan (17, 39), miRanda (27), miRBase (16), and PicTar (31), 7.2% of the miR-31-repressed genes were predicted targets of miR-31.
In silico biological process analysis by the PANTHER classification system (61, 62) was used to assess whether the observed miR-31-mediated reprogramming of LECs specifically affected lymphatic, blood vascular, and/or endothelial biological functions. Intriguingly, genes involved in cell communication, signal transduction, cell adhesion, apoptosis, and numerous signaling pathways were overrepresented among the miR-31-repressed genes compared to the expected number of genes (see Table S5 in the supplemental material). While these biological processes are common to both endothelial cell types, this enrichment suggests a dramatic reorganization of the LEC surface characteristics, as well as cell signaling network activity, following miR-31 overexpression. The specific effects miR-31 overexpression had on genes involved in vascular lineage-specific differentiation were then identified by comparing the miR-31-regulated genes to the LEC (344 genes) and BVEC (479 genes) signature genes previously identified in vitro (23, 32, 48; Shin et al., unpublished). Interestingly, twice as many LEC signature molecule-encoding transcripts (9.6%) as BVEC signature molecule-encoding transcripts (4.8%) were reduced following miR-31 overexpression (Fig. (Fig.33 A and B; see Table S3 in the supplemental material). Also, approximately four times as many BVEC signature genes (4.6%) as LEC signature genes (1.1%) were induced/stabilized following miR-31 overexpression (Fig. 3A and B; see Table S4 in the supplemental material). Corroborating the microarray data, qRT-PCR experiments confirmed that four of the LEC signature genes (EDNRB, PROX1, PPP1R9A, and HOXD10) and two of the BVEC signature genes (ICAM1 and SELE) tested were significantly less abundant in the pre-miR-31 samples than in the pre-miR-Neg control (Fig. 3C and D). Furthermore, we also validated the upregulation of the miR-31-induced BVEC signature genes MMP1 and RGS4 (Fig. (Fig.3E3E).
Among the miR-31-repressed target genes was PROX1, an essential lymphatic lineage-specific transcription factor (1, 7). qRT-PCR analysis confirmed that transfection of LECs with 4 μM pre-miR-31 resulted in a ≥60% reduction in PROX1 transcripts (Fig. (Fig.44 A), which was further verified by Northern blotting (see Fig. S5 in the supplemental material). Importantly, immunoblotting revealed a consistent decrease in PROX1 protein levels of <40% following miR-31 overexpression (Fig. 4B and C). Similar but less consistent results were observed after transfection with 2 μM pre-miR-31 (see Fig. S6 in the supplemental material). Conversely, loss of miR-31 function in BVECs via transfection of HUVECs with miR-31 inhibitor molecules (anti-miR-31) resulted in a 1.7- to 3.1-fold increase in PROX1 mRNA levels (Fig. (Fig.4D).4D). PROX1 protein remained undetectable in these samples (data not shown).
PROX1 was not predicted to be a target of miR-31 by TargetScan (17, 39), miRanda (27), miRBase (16), or PicTar (31), but human cells express two isoforms of PROX1 mRNA, a 7.9-kb isoform that contains a 5.4-kb 3′ UTR and a 3.1-kb isoform that has a much shorter 602-bp 3′ UTR (57). Only the short 3′ UTR has thus far been used for miRNA binding site predictions. In agreement with previous studies (25, 26, 49), our LECs expressed the longer 7.9-kb isoform of PROX1 (see Fig. S5 in the supplemental material). Therefore, standard nucleic acid sequence alignment techniques (SIM alignment tools) and independent Targetscan 5.0 analyses were used to identify potential miR-31 binding sites in the 5.4-kb 3′ UTR. SIM alignment identified five potential miR-31 recognition sites, and the Targetscan search identified one of these sites, nt 949 to 971, as a 7mer-m8 binding site (Fig. (Fig.5A;5A; see Table S6 in the supplemental material).
To test the functional relevance of these candidate miR-31 binding sites, luciferase reporter genes containing a full-length PROX1 3′ UTR (PROX1 FL), six PROX1 3′ UTR fragments (PROX1 F1 to F6) (Fig. (Fig.5A),5A), or a PROX1 CDS were constructed. The activities of these chimeras were monitored following miR-31 gain of function in LECs (Fig. (Fig.5B)5B) and loss of function in HUVECs (Fig. (Fig.5C).5C). Confirming pre-miR-31 overexpression and anti-miR-31 knockdown activity, miR-31 binding site (miR31BS) luciferase reporter gene activity decreased or increased by <40% after cotransfection with pre-miR-31 or anti-miR-31 molecules, respectively (Fig. 5B and C). The luciferase activities of the PROX1 FL and PROX1 F2 reporter genes, which contain the 7mer-m8 site, decreased significantly (>35% and >45%, respectively) following miR-31 overexpression (Fig. (Fig.5B).5B). Conversely, their activities increased >1.4-fold and 3-fold, respectively, after miR-31 inhibition (Fig. (Fig.5C).5C). While PROX1 3′ UTR F1 and F4 reporter gene activities increased after miR-31 knockdown, reciprocal responses following overexpression were not observed. Together, these findings confirm the direct posttranscriptional regulation of PROX1 by miR-31 and suggested that this regulation is mediated via nt 949 to 971 of the 5.4-kb 3′ UTR.
To validate this, the seed sequence (nt 964 to 971) and 3′ compensatory site interacting nucleotides (nt 954 to 960) of the PROX1 3′ UTR were mutated to match the miR-31 sequence in both the PROX1 FL and PROX1 F2 luciferase reporter plasmids, thus eliminating the predicted PROX1-miR-31 interaction. While the luciferase activities of the wild-type PROX1 FL and PROX1 F2 constructs decreased or increased after miR-31 overexpression or knockdown, respectively (Fig. (Fig.5D),5D), the activities of the mutant full-length and F2 fragment luciferase reporter genes did not change significantly under either condition, thus mapping a bona fide, biologically active miR-31 binding site to nt 949 to 971 of the PROX1 3′ UTR.
Numerous studies have demonstrated that PROX1 transcriptional activities help dictate the molecular characteristics and functional activities of vascular endothelial cells (12, 26, 44, 48; Shin et al., unpublished). Therefore, our characterization of PROX1 as a target of miR-31 posttranscriptional regulation suggested that miR-31 repression of PROX1 should specifically, albeit indirectly, alter the expression PROX1 target genes. Comparing the miR-31-regulated genes identified above with a PROX1 loss-of-function data set generated following lentiviral inhibition of PROX1 in LECs (Shin et al., unpublished) revealed that approximately 20% of the miR-31-repressed and -induced genes were similarly repressed or induced, respectively, following Prox1 knockdown (see Table S3 and S4 in the supplemental material). Intriguingly, <50% of the miR-31-repressed LEC signature genes (see Table S3 in the supplemental material) and 36% of the miR-31-induced BVEC signature genes (see Table S4 in the supplemental material) were similarly differentially expressed following PROX1 depletion from LECs.
During embryogenesis, PROX1 is expressed in a subpopulation of cardinal vein endothelial cells that give rise to the mammalian lymphatic vascular system (1, 7). The BVEC-specific expression of miR-31, together with its ability to posttranscriptionally repress numerous BVEC and LEC signature genes (see Tables S3 and S4 in the supplemental material), including Prox1, suggested that this miRNA might play a role in vascular development. As many of the BVEC and LEC signature genes targeted by miR-31 also play major roles in Xenopus vascular development (9, 20, 45), we reasoned that ectopic expression of miR-31 in early Xenopus embryos might interfere with lymphatic vascular development. To investigate this, two-cell-stage Xenopus embryos were unilaterally microinjected with human pre-miR-31 or pre-miR-Neg molecules. Lymphatic and blood vascular system development was then monitored in stage 39 embryos using whole-mount ISH for specific lymphatic and blood vascular marker genes (28, 29, 45).
No gross developmental defects or externally visible phenotypes were observed following pre-miR-Neg or pre-miR-31 microinjection into Xenopus embryos (Fig. (Fig.6).6). Moreover, prox1 and vegfr3 marker gene analysis demonstrated that lymphatic vascular development progressed normally in 95% and 79% of the pre-miR-Neg control embryos, respectively (Table (Table2).2). The embryos had well-defined and clearly visible lymph hearts, lymph vessels, and punctate patches of LECs in their tails (Fig. 6A and B). In contrast, a dose-dependent increase in lymphatic vascular defects was observed in pre-miR-31-injected embryos. Specifically, vefgr3 ISH demonstrated that the percentage of embryos with lymphatic vascular defects, as monitored by the loss of vegfr3-positive lymphatics sprouting from the lymph hearts, progressively increased from 6.1 to 76.2% as the amount of pre-miR-31 molecules increased from 1 to 50 ng (Table (Table2).2). Generally, lymph hearts were present in these embryos but appeared smaller and less well defined than in control embryos (Fig. 6A and B) or the uninjected side of the pre-miR-31 embryos (data not shown). Moreover, lymphangiogenesis, scored by the presence of vegfr3-expressing lymphatic vessels sprouting from the lymph heart, was either strongly reduced or absent in the presence of excess miR-31 (Fig. 6A and B). These phenotypes were similar to those observed following morpholino inhibition of vegfc, where lymphangiogenesis was disrupted in the lymph heart region of 67 to 100% (n = 3; total number of embryos analyzed = 71) of the injected Xenopus embryos (see Fig. S7 in the supplemental material).
pecam1 expression was used to monitor blood vascular system development in pre-miR-Neg- and pre-miR-31-microinjected Xenopus embryos. In control embryos, all of the major blood vascular structures, such as the posterior cardinal veins and the dorsal aorta, were clearly visible, and angiogenic sprouting of intersomitic veins occurred normally in 73% of the pre-miR-Neg-injected embryos (Fig. (Fig.6C6C and Table Table2).2). By comparison, the percentage of embryos displaying unilateral intersomitic vein growth and/or guidance defects progressively increased from 0% to 76% with increasing amounts of pre-miR-31 injected (Fig. (Fig.6C6C and Table Table22).
Gain-of-function phenotypes can occasionally be attributed to the off-target effects associated with nonphysiological expression levels of a small interfering RNA or miRNA (41, 51). We therefore sought to confirm the Xenopus miR-31 overexpression phenotypes in zebrafish, another highly relevant vertebrate model organism for studying blood and lymphatic vascular development (40). To facilitate the quantification of venous sprouting from the posterior cardinal vein (24, 33, 67), we used phospholipase C gamma 1 (plcg1) mutant embryos in the TG(fli1a:gfp)y1 background (36) [TG(fli1a:gfp)y1; plcg1t26480], in which the venous sprouts contributing to both the blood and lymphatic vasculature are clearly visible. Venous sprouting was quantified at 48 hpf following the injection of 170 or 340 pg of either human pre-miR-31 precursor or pre-miR-Neg control molecules (Table (Table33 and Fig. Fig.77 A). In agreement with our Xenopus studies, a significant dose-dependent reduction in venous sprouting was observed in pre-miR-31-injected embryos compared to embryos injected with 340 pg of pre-miR-Neg molecules or to uninjected controls (Table (Table33 and Fig. Fig.7).7). The increase in venous sprouting observed in pre-miR-Neg-injected embryos (Fig. 7A and C) appears to be a stress response, which we have also observed in a number of unrelated control injections (data not shown). Conversely and importantly, miR-31 overexpression led to a highly significant reduction in venous sprouting and no other developmental defects were observed in these embryos (Table (Table33 and Fig. 7A and D).
Venous sprouting and lymphangiogenesis were also monitored in zebrafish embryos following the injection of increasing concentrations of morpholino oligonucleotides targeting both mature and precursor dre-miR-31. Significant vascular phenotypes could not be specifically attributed to loss-of miR-31 activity in these embryos (data not shown). Taken together, our miR-31 gain-of-function studies with Xenopus and zebrafish embryos indicate that appropriate expression levels of miR-31 during vertebrate embryogenesis are required for normal lymphatic and blood vascular development.
In the study presented here, we first defined the in vitro expression profiles of 157 human miRNAs in primary human LECs and BVECs using a TaqMan-based qRT-PCR profiling platform, whose increased sensitivity facilitated the detection of at least twice as many miRNAs in HUVECs as previously reported (19, 34, 50, 58, 66). We also found that one of the most highly expressed HUVEC miRNAs, miR-126 (19, 34, 50, 58, 66), was >600 times more abundant in both endothelial cell types than in either keratinocytes or fibroblasts. Comparative analysis identified four BVEC and two LEC signature miRNAs. Of the four BVEC signature miRNAs, three were previously reported as highly expressed in HUVECs (19, 34, 50, 58, 66), and a very recent study has demonstrated that tumor necrosis factor treatment augments miR-31 expression in HUVECs (60). Moreover, our miRNA profiling study has further classified the widely expressed (38), metastasis-associated (63, 64) miRNA miR-31 as a BVEC signature miRNA. Finally, in agreement with their LEC-specific expression, neither miR-95 nor miR-326 was detected in the previous studies.
Importantly, further analysis of miR-31, miR-326, miR-125b, and miR-99a in adult mouse tissues confirmed that their vascular lineage-specific expression patterns were maintained in vivo. The degrees of lineage-specific expression differences in vivo were, however, usually less pronounced and more variable than those observed in vitro. This is likely due to the mixed populations of BVECs and LECs isolated from the multiple vessel types present in the colon tissue (capillaries, postcapillary venules, lymphatic capillaries, lymphatic collecting vessels, etc.), which likely exhibit different gene expression patterns. Moreover, their relative contributions to the isolated total RNA might vary, thus contributing to larger variability in miRNA expression patterns. In addition, the ex vivo miRNA expression profiling studies were technically challenging as the whole process took more than 2 h and only a few thousand endothelial cells could be isolated by high-speed cell sorting. Consequently, the smaller amounts of isolated total RNA, reduced RNA quality, and possible gene expression changes incurred during the 2-h isolation procedures likely contributed to the observed differences in in vivo and in vitro miR-31 expression, as well as to the observed interindividual variability in miR-31 expression. Surprisingly, we were unable to confirm the differential expression patterns of miR-137 in vivo. This is likely because miR-137 expression levels were very low in the adult tissues analyzed here, as indicated by the late qRT-PCR detection (CT, >35) and high standard deviations between technical replicates. ISH analysis of chicken embryos revealed that miR-137 is expressed in blood vessels and cardinal veins at stage 25 of embryonic development (8), demonstrating that miR-137 expression is associated with the developing blood vasculature.
The identification of vascular lineage-specific miRNAs suggested that they might regulate fundamental and lineage-specific endothelial cell functions and/or differentiation processes. Indeed, overexpression of the BVEC-specific miRNA miR-31 in LECs induced the preferential degradation of LEC signature genes, including those for the well-characterized lymphatic transcription factors PROX1 and FOXC2. As these lymphatic lineage-specific molecules act as molecular switches, their preferential suppression suggests that BVEC-specific posttranscriptional regulatory mechanisms help maintain BVEC phenotypes by suppressing lymphatic lineage-specific transcription programs. This concept was supported by our findings that ectopic overexpression of miR-31 in LECs preferentially repressed LEC signature gene expression and induced BVEC signature gene expression. In this respect, our identification and validation of PROX1 as a direct miR-31 target are intriguing, as BVEC-specific posttranscriptional regulation of PROX1 could, at least in part, explain these in vitro miR-31-mediated reprogramming events on the molecular level. Indeed, previous studies have demonstrated that PROX1 overexpression in BVECs induces the expression of lymphatic vascular markers and suppresses blood vascular markers (26, 48), whereas PROX1 knockdown in LECs inhibits LEC signature gene expression and triggers BVEC signature gene expression (44; Shin et al., unpublished). Moreover, the overlaps between the miR-31-regulated genes identified here and a PROX1 loss-of-function data set further indicate that transcriptional reprogramming events observed following miR-31 overexpression in LECs were, in part, mediated by miR-31 repression of Prox1. Additional experiments are required to determine which of the miR-31-regulated candidate Prox1 target genes may also be direct targets of miR-31.
While PROX1 was not a predicted target gene of miR-31 (16, 17, 27, 31, 39), our manual miR-31 site prediction analyses of the 5.4-kb PROX1 3′ UTR and subsequent luciferase 3′ UTR tethering assays identified a bona fide miR-31 binding site between nt 949 and 971 of the PROX1 3′ UTR. Interestingly, similar manual miR-31 prediction analyses of the chimpanzee, mouse, rat, chicken, Xenopus, and zebrafish PROX1 3′ UTRs revealed that this site is evolutionarily conserved in vertebrates and identified additional, potentially functional, miR-31 binding sites (see Table S6 in the supplemental material). Taken together, our transcriptome profiling and biochemical studies have revealed a novel, highly conserved, BVEC-specific posttranscriptional regulatory mechanism that suppresses PROX1 expression in the blood vasculature.
Our findings also suggested that miR-31 expression in the developing blood vascular endothelium could regulate the acquisition of lymphatic lineage-specific characteristics and, thus, vascular development in vivo. Multiple miR-31 loss-of-function studies using morpholino oligonucleotides were performed with both wild-type and plcg1 mutant zebrafish embryos. Statistically significant vascular phenotype differences were not observed in zebrafish embryos injected with low-to-moderate amounts (≤10 ng) of MO (data not shown). This suggests that the miR31-mediated regulation of vascular development identified here is redundant. This is not surprising, since miRNAs frequently function cooperatively (3, 15, 17), which in turn complicates the attribution of specific functions to individual miRNAs (53). In contrast, miRNA gain-of-function experiments have proven very informative and have defined important biological functions of several miRNAs (42, 43, 53). For example, overexpression studies with Xenopus embryos have demonstrated that miR-15 and miR-16 restrict the size of Spemann's organizer in vivo by targeting the nodal type II receptor acrvr2a (42). We therefore carried out miR-31 overexpression studies with Xenopus and zebrafish embryos to determine the effect of miR-31 on cells and tissues that normally do not express miR-31, such as the lymphatic vasculature.
Our gain-of-function experiments clearly demonstrated that miR-31 expression is incompatible with normal lymphatic vascular development in Xenopus and, to a lesser extent, zebrafish embryos. The analysis of Xenopus embryos suggests that some aspects of lymphatic vascular development, such as specification of lymph hearts and LECs in the tail, are unaffected by miR-31 overexpression. Lymphangiogenesis and the development of an extensive lymphatic vasculature in the embryonic trunk are, however, clearly reduced and/or disrupted. Furthermore, we demonstrated that these observed lymphatic defects were reminiscent of those observed following MO-mediated inhibition of vegfc. These phenotypic similarities indicate that miR-31 overexpression interferes with an early step in lymphatic development. The identification of evolutionarily conserved miR-31 binding sites in PROX1 3′ UTRs (see Table S6 in the supplemental material) suggests that miR-31 overexpression may directly target and interfere with PROX1 transcripts in vivo. Moreover, the abnormal or disrupted intersomitic vein sprouting seen in Xenopus and zebrafish embryos (data not shown) following miR-31 overexpression implies that miR-31 also regulates BVEC responsiveness to the environmental stimuli directing blood vascular growth and maturation. Interestingly, several genes involved in the Slit/Robo, netrin, and ephrin signaling pathways (see Table S3 in the supplemental material), which provide crucial guidance cues during blood vascular development (1), were repressed following miR-31 gain of function in vitro. In vivo posttranscriptional regulation of any one of these molecules by miR-31 could contribute to the observed blood vascular maturation defects. Taken together, our results indicate that appropriate expression of miR-31 during vertebrate embryogenesis is required for both lymphatic vascular development and blood vascular growth and maturation. Interestingly, our in vivo studies also correlate well with a recent study demonstrating that miR-31 controls the invasive capacity of breast cancer cells (63, 64). Collectively, these studies suggest roles for miR-31 in the regulation of cell migratory behavior during normal embryonic development and under pathological conditions in the adult body.
On the basis of our in vitro studies, we postulate that PROX1 transcripts represent one of the key targets of miR-31. This repression would prevent inappropriate and/or premature transcriptional activation of lymphatic differentiation in the developing blood vasculature. While this notion is an attractive model, it is, however, important to stress that miR-31 targets several other LEC signature genes. It is therefore unlikely that posttranscriptional repression of PROX1 by miR-31 is solely responsible for the vascular developmental defects observed in Xenopus and zebrafish embryos overexpressing miR-31. For example, miR-31-mediated repression of FOXC2, a transcription factor that is required for specification of the lymphatic capillaries versus collecting lymphatic vessels at later stages of embryogenesis (1, 7), may also contribute to the vascular defects observed. Another miR-31 candidate target is RAMP2, a calcitonin receptor-like receptor-associated receptor activity-modifying protein that triggers lymphangiogenesis in response to adrenomedullin signaling (13). Finally, other LEC signature molecules subject to miR-31 regulation, whose lymphatic lineage-specific functions have not yet been characterized, could also enhance the effects miR-31 has on lymphatic and blood vascular development.
The miRNAs profiled in the present study represent approximately 25% of the known human miRNAs. Thus, more comprehensive and global miRNA profiling studies may result in the identification of additional endothelial lineage-specific miRNAs. In summary, we have defined the first vascular lineage-specific miRNAs and identified with miR-31 a novel miRNA-mediated regulatory mechanism that inhibits LEC phenotype acquisition in vitro and vascular development in vivo. From a therapeutic perspective, it remains to be investigated whether the ectopic expression of miR-31 might also inhibit malignant tumor-associated (lymph)angiogenesis, thus preventing tumor growth and cancer metastasis.
This work was supported by National Institutes of Health grant CA69184; Swiss National Science Foundation grant 3100A0-108207; Austrian Science Foundation grant S9408-B11; Cancer League Zürich, Commission of the European Communities, grant LSHC-CT-2005-518178 (M.D.); Swiss National Science Foundation grant 3100A0-101964 (A.W.B.); Netherlands Organization for Scientific Research (NWO) Venigrant (T.K.); and EMBO long-term fellowships ALTF 1104-2007 (D.M.L.P.) and ALTF 52-2007 (T.K.).
We thank Young Kwon Hong for the PROX1 3′ UTR plasmid (YH1551); Salvatore Oliviero, Jay W. Shin, and Ahmad Salameh for sharing the PROX1 lentivirus knockdown microarray data set; Patrick Pedrioli for bioinformatic assistance and critical reading of the manuscript; and Jana Zielinski, Cornelius Fischer, and Jeannette Scholl for expert technical assistance. We also thank the Tübingen 2000 Screen Consortium for identifying the plcg1t26480 allele.
D.M.L.P. and T.K. designed and performed research experiments, analyzed the data, and wrote the manuscript. V.D., G.J., G.V.D.H., and R.E.K. designed and performed research experiments and analyzed the data. D.M., J.W.S., S.L., and P.C. performed research experiments and analyzed the data. M.D., A.W.B., and S.S.-M. designed research experiments, analyzed the data, and wrote the manuscript.
Published ahead of print on 17 May 2010.
†Supplemental material for this article may be found at http://mcb.asm.org/.