Abstract
MicroRNAs contribute to biological robustness by buffering cellular processes from external perturbations. Here we report an unexpected link between DNA damage response and angiogenic signaling that is buffered by two distinct microRNAs. We demonstrate that genotoxic stress-induced miR-494 and miR-99b inhibit the DNA repair machinery by targeting the MRE11a-RAD50-NBN (MRN) complex. Functionally, gain and loss of function experiments show that miR-494 and miR-99b affect telomerase activity, activate p21 and Rb pathways and diminish angiogenic sprouting in vitro and in vivo. Genetic and pharmacological disruption of VEGFR-2 signaling and the MRN complex reveal a surprising co-dependency of these pathways in regulating endothelial senescence and proliferation. Vascular-targeted delivery of miR-494 decreases both growth factor-induced and tumor angiogenesis in mouse models. Mechanistically, disruption of the MRN complex induced CD44, a known driver of senescence and regulator of VEGF signaling in addition to suppressing IL-13 a stimulator of VEGF signaling. Our work identifies a putative miR-facilitated mechanism by which endothelial cells can be insulated against VEGF signaling to facilitate the onset of senescence and highlight the potential of targeting DNA repair to disrupt pathological angiogenesis.
Introduction
Accumulation of DNA damage can overwhelm the repair machinery and lead to senescence (1). Endothelial senescence leads to progressive damage and deterioration of cellular structure and function over time (2) (3). Two major pathways of senescence in endothelial cells are replicative senescence and stress induced premature senescence (SIPS). Replicative senescence is one of the hallmarks of aging and is associated to telomere shortening. Stress-induced premature senescence is triggered by external stimuli, including oxidizing agents and radiation, both of which can induce DNA damage and cell cycle arrest.
Recent studies indicate that DNA repair proteins in endothelial cells (ECs), such as ATM kinase and histone H2AX, have an inherently pro-angiogenic role (4, 5). For example, ATM deficiency kinase decreases tumor angiogenesis while enhancing the anti-angiogenic action of VEGF blockade, suggesting that pathological neoangiogenesis requires ATM. ATM can also function as a redox sensor independent of its DDR function and regulate oxidative stress responses (6) and was recently implicated as a driver of cellular senescence (7). Similarly, global or EC specific deletion of the histone H2AX in mice results in a substantial decrease of pathological angiogenesis in proliferative retinopathy, hindlimb ischemia and tumor angiogenesis. These findings suggest that key regulators of DNA damage repair (DDR) modulate pathological angiogenesis.
Mre11a-RAD50-NBN (MRN) complex acts as a sensor of DNA double strand breaks (DSB), initiating Homologous Recombination (HR) or Non-Homologous-End-Joining (NHEJ) pathway. Once MRN detects DSBs, it tethers the ends, activates and recruits DNA damage response proteins such as ATM (8) (9). Interestingly, MRN is also associated with telomere maintenance, playing a role in the formation and disassociation of the t-loops (10, 11). The MRN complex relationship with aging and cell senescence has been described (12, 13), however its role in ECs and pathological neovascularization is unclear. Similarly, while several miRs have been shown to be involved in EC senescence (2, 14, 15), proliferation, viability and migration (16) (17) (18), our work identifies a novel link between DDR driven induction of miR-494, miR-99b, the MRN complex, EC senescence and tumor angiogenesis. Our results indicate that altering the DDR response in tumor ECs, not only enhances cellular senescence but also has a functional effect on both growth factor induced and pathological angiogenesis leading to decreased tumor growth..
Results
DNA damage induces miRs-494 and miR-99b in vitro and in vivo
We previously reported a seven microRNA signature (miR-103, miR-494, miR-99b, miR-21, miR-224, miR-92a and let-7a) specifically upregulated in ECs after radiation, hydrogen peroxide and cisplatin treatment. Among these, we found that miR-494 and miR-99b are both transcribed rapidly in human umbilical vein endothelial cells (HUVECs) in response to γ-radiation with maximal induction occurring at a lower (2 Gy) dose of radiation. (Figure 1A-B). Consequently, we observed that miR-494 was also upregulated after radiation in a PyMT breast cancer model both at low and high doses at different time points (Figure 1C-D).
Gain and loss of miRs-494 and 99b affect pathological and replicative senescence
To understand the role of these two miRs, we performed different gain and loss of function assays in our endothelial cell model, HUVECs. We observed that the most prominent phenotype with these two miRs was the induction of cellular senescence. Gain of either miR-99b or miR-494 in early passage HUVECs increased senescence associated β-galactosidase levels (SA-β gal). This phenotype was also observed in human microvascular endothelial cells (HMVECs) and normal human lung fibroblasts suggesting this phenotype was not exclusive to venous endothelium (Supplementary Figure 1). In contrast, inhibition of miR-99b or miR-494 decreased SA-β gal levels significantly in HUVECs undergoing pathological senescence induced by a high-dose of radiation (Figure 2B). We asked if these miR inhibitors had an effect on replicative senescence and found that inhibition of either miR-99b or miR-494 in late passage senescent HUVECs decreased their β-gal and increased activation of caspase 3 and 7 (Figures 2C-D). Similarly, gain of miR-494 and miR-99b affected cell cycle progression whereas the inhibition of the miRs decreased the G2 arrest in response to a 10 Gy dose of radiation (Supplementary Figure 2). Interestingly, miR-494 expression decreased BrdU incorporation whereas miR-99b expression increased activation of Caspase-3 & 7 (Supplementary Figure 3). Given the strong influence of DNA damage on cellular senescence, we asked if either of these miRs affected DNA damage. Indeed, ectopic expression of both miR-494 and miR-99b increased histone H2AX phosphorylation, a marker of DNA damage (Figure 2E). Importantly, this increase in DNA damage resulted due to the miRs alone without any additional source of DNA damage. Consistent with these findings, we observed some critical features of senescence - a decrease in telomerase activity (Figure 2G), an increase in the cell cycle regulator p21 as well as a decrease in Rb hyperphosphorylation (Figure 2H). Moreover, Phalloidin staining in HUVECs after 48 hours of miR-494 treatment alsorevealed flattened and multinucleated cells (Figure 2F), another morphological phenotype associated with stress dependent senescence. Taken together, these observations establish that miR-494 and 99b drive EC senescence likely via pathways involved in DNA damage or repair.
miR-494 and miR-99b target the MRN DNA repair complex affecting senescence
To analyze the possible targets of miR-494 and miR-99b responsible for this phenotype, we ectopically expressed these two miRs in HUVECs and analyzed gene expression using a DNA damage array (Supplementary Figure 4). Interestingly, we identified three common targets: MRE11a, RAD50 and NBN. RNA hybrid modeling suggested putative binding sites for both miRs on all three target 3’UTR regions (Supplementary Figures 5-6). Both miR-494 and −99b directly bound MRE11A mRNA, and to a lesser extent, RAD50 and NBN mRNAs, as measured by a miRTrap assay (Clontech) (Figure 3A). Consistent with this finding, transfection of miR mimics decreased MRN RNA levels by qRT-PCR (Figure 3B) and protein levels as measured by a western blot (Figure 3C). We corroborated the interaction of these miRs with their targets by cloning the 3’-UTRs in luciferase reporter vectors, and co-transfecting them in HEK-293T cells. Either of the miRs transfected alone was able to decrease the luciferase activity downstream of the MRN 3’-UTR reporter (Figure 3D). Finally, miR-494 transfected HUVECs showed a decrease in MRE11a nuclear foci number (Figure 3E).
miRs typically regulate several target mRNA transcripts making their functional attribution challenging (19), (20-23). To address this, we used a target protector, a locked nucleic acid (LNA) stabilized oligonucleotide, that binds completely to a specific miR-binding site on a single target mRNA, to rescue it from the negative regulation of the miR. We confirmed that the MRE11A target protector, rescued MRE11A RNA levels even in the miR-494 transfected cells compared to a control target protector oligo (Supplementary Figure 7A). Functionally, the MRE11A target protector significantly restored the telomerase activity and to a lesser extent the senescence associated β-gal levels in miR-494 transfected cells (Supplementary Figure 7B-C). We then sought to determine if disruption of the MRN complex using siRNAs (Supplementary Figure 8) resulted in a similar phenotype. Indeed, silencing each of the components of the MRN complex induced senescence and impaired telomerase activity (Figure 3 F-G). Confirming these observations, we found that siMRN as well as Mirin-1, a small molecule inhibitor specific to MRE11a (24), also enhanced senescence in HMVECs. These data indicate that the miR-494 and 99b both downregulate the MRN machinery and thereby both exacerbate DNA damage and drive senescence.
The MRN pathway cross-talks with angiogenic growth factors
While disruption of DNA damage response (DDR) in ECs can modulate both pathological and developmental angiogenesis via VEGF (4, 5), it is not clear if targeting MRN would also influence VEGF signaling. Our results show that combinatorial disruption of both VEGFR2 and MRE11a with Mirin-1 and Vandetanib, a small molecule inhibitor for VEGFR2 (Figure 4A), led to a decrease in proliferation of HUVECs with a synergistic interaction at the lowest dose (Chou-Talalay Combination Index <0.4). Conversely, disruption of the MRN complex with Mirin-1 affected bFGF mediated pERK signaling and to a lesser extent VEGF induced pERK in HUVECs (Figure 4B). Interestingly, we silenced VEGF-receptor 2 (VEGFR2) and tested whether disruption of the MRN complex induced senescence. Surprisingly, knockdown of VEGFR2 diminished the senescence promoting activities of both the siRNAs targeting the MRN and Mirin-1 (Fig 4C-D). These experiments argue that the disruption of MRN could inhibit angiogenic growth factor signaling. Indeed, siRNA targeting of Mre11a or NBN significantly decreased angiogenic sprouting over a four days in a 3D fibrin matrix.
miRs-494 and 99b drive senescence through cell intrinsic and extrinsic mechanisms
To understand the shared molecular pathways that may drive senescence downstream of miR-494, miR-99b and the MRN complex, we utilized a human senescence gene signature array. We found 6 genes with similar changes across both the miR and siRNA treatment groups (Supplementary Fig 9A). Interestingly, CD44, a known negative regulator of VEGF signaling (25, 26), was the most significantly upregulated gene in the four groups. This increase in gene expression was also reflected by increased cell surface expression of CD44 (Supplementary Figure 9B). We also performed a cytokine expression profiling using a multiplex luminex assay and discovered that miR-494 robustly suppressed IL-13 levels. IL-13 can function as a driver of VEGF signaling (27) or via its shared IL-4 receptor leading to STAT6 phosphorylation and senescence (28). We are currently investigating the relative contributions of these pathways to miR-494 and MRN function in EC senescence.
miR-494 mimic disrupts growth factor driven and pathological angiogenesis
Thus far, our data shows that miR-494 and 99b induce senescence by inhibiting the MRN complex, which diminishes VEGF/bFGF signaling, and sprouting angiogenesis. Next, we addressed whether this miR pathway can be modulated to regulate angiogenesis in vivo. First, we confirmed that the miR mimics miR-494 and 99b also disrupted sprouting angiogenesis in the fibrin bead assay (Figure 5A). Next, we used a growth factor induced angiogenesis model by implanting bFGF containing Matrigel plugs in mice. We treated the mice with either a miR-494 mimic or a control mimic in vascular-targeted 7C1 nanoparticles (29, 30). We found a robust decrease in CD31 staining in miR-494 treated plugs compared to the control miR mimic treated group (Figure 5B). To test the utility of this approach in a pathological angiogenesis, we utilized a highly aggressive, triple negative 4T1 mouse mammary carcinoma model. Treatment of vascularized tumors with miR-494 in the 7C1 nanoparticles resulted in a delayed tumor growth for a few days (Figure 5C). We also observed a trend towards a decrease in lung metastases (Figure 5D). Importantly, we observed the miR-494 treated tumors had a significant decrease in angiogenesis as measured by CD31 staining (Figure 5E). Our in vivo studies indicate that miR-494 is able to decrease both growth factor induced and pathological angiogenesis in vivo.
These results demonstrate that the DNA repair pathway and the MRN complex have an essential functional role in angiogenesis and we propose that we can exploit this DNA repair-VEGF pathway codependency for future antiangiogenic therapies. Genotoxic stress agents induce DNA damage in ECs; however, they also increase expression of miR-494 and miR-99b. These two miRs lead to senescence by targeting the DNA repair complex MRN. Moreover, disruption of MRN decreases proliferation and ERK phosphorylation in response to angiogenic growth factors. In addition, other effects such as CD44 induction and IL-13 suppression can also amplify the induction of senescence and inhibition of angiogenesis.
Discussion
We have postulated that a network of stress-dependent miRs target DNA repair pathways leading to impaired angiogenesis (29). Among this group of stress-induced miRs, miR-494 and miR-99b lead to senescence, likely through MRN disruption. miR-494 has been shown to be highly upregulated in retinoblastoma (31), cardiovascular pathologies such as cardiac injury (32) and in atherosclerotic lesion development (33). miR-494 has also been reported as an angiogenesis inhibitor, supporting our data here (34-36). Recently, Esser et al demonstrated that miR-494 is downregulated in ECs treated with the pro-angiogenic factor BMP4, in opposition to an increase in the pro-angiogenic miR-126 (37). miR-494 has been implicated in tumor senescence and the development of resistance to radiation and chemotherapy (38) (39) (40) (41).
In EC-progenitors, miR-99b affects the expression of PECAM-1 and VE-cadherin (42). miR-99b also modulates β-catenin in endothelial junctions (43). Interestingly, miR-99b expression has been proposed as a prognostic marker to assess the response to VEGFR2 Tyrosine Kinase Inhibitors (TKI). Lukamowicz-Rajska et al described higher expression of miR-99b-5p in clear cell renal cell carcinoma (CCRC) patients with complete response to TKI treatments (44) compared to partial or non-responders. Another example is dermal wound healing, where TK pathways have decreased signaling. Prior studies proposed that miR-99b targets several members of the IGFR1/AKT/mTOR signaling pathway. They demonstrated that miR-99b was downregulated in the early phases of wound healing likely facilitating cell proliferation (45). We propose here that miR-99b expression is leading to senescence through MRN disruption, but that its effect on the VEGFR2/ERK signaling pathway enhances the anti-angiogenic phenotype. Similarly, our observations of miR-99b diminishing a specific VEGF-dependent early-gene signature (46), also suggests that the induction of miR-99b in complete responders can enhance the effect of VEGFR2 TKIs.
We find that miR-494 and miR-99b target MRE11a, NBN and RAD50 to affect EC senescence and sprouting angiogenesis. EC senescence leads to dysfunction in cardiovascular diseases (47, 48). Loss of function mutations in the MRN genes cause inherited genetic disorders that are characterized by elevated sensitivity of patients’ cells to radiation damage (49, 50). Interestingly, there is some evidence that NBN disruption has an anti-angiogenic effect in vivo (51). Our observations also imply that the senescence phenotype maybe mediated by the increase in CD44 expression on the cell surface (52). CD44 is a known regulator of endothelial interactions with leukocytes in addition to its other critical roles in EC junction integrity (53). Induction of CD44 in response to DNA damage by miR disruption of the MRN complex may increase adhesiveness of the endothelium (54, 55). Similarly, there are demonstrated roles for IL-13 in enhancing VEGF signaling (27) or senescence (28) that are plausible downstream of miR-494 regulation of the MRN pathway.
Emerging studies have implicated deficient telomere maintenance as leading to senescence when MRN function is compromised. For example, decreased MRE11a has been shown to increase T-cell aging in arthritis due to compromised telomere maintenance and heterochromatin unraveling (56). Recent structural studies also shed light on how the phosphorylation of NBS influences its interaction with TRF2 and dictates the repair of telomeres (57). RAD50 downregulates the association of TRF1 from telomeres and also contribute to maintain telomere length (58). We saw a decrease in telomerase activity across miRs and siRNAs experiments targeting the MRN complex. Moreover, the MRE11a target protector largely restored this function, implying this was the miR-494 target responsible for the compromised telomerase activity. Therefore, we suspect that the telomere maintenance function of the MRN complex drives the EC senescence downstream of miR-494 and miR-99b induced by DNA damage.
Our observations indicate that the function of these miRs in regulating telomerase activity and EC senescence may depend on the expression of VEGFR2. This is surprising, since expression of VEGFR2 may steer cells into a robust replication program rather than a cell cycle exit characteristic of senescence. However, it is possible that replication stress and DNA damage are enhanced in the presence of these miRs. Similarly, our findings demonstrate a decrease in MRN activity either by miR-99b induction or Mirin-1 affects the VEGFR signaling at the level of pERK. These data argue that the MRN pathway and the VEGFR2 signaling cross-talk functions as a threshold to determine the level of DNA damage in ECs. In the presence of significant DNA damage the MRN pathway is disrupted and the VEGF signaling is diminished thereby decreasing angiogenesis.
It is possible that during physiological aging, increase in miR-494 and miR-99b levels render the ECs less responsive to VEGF levels and therefore, decrease angiogenic responses. We envision that this senescence associated decrease in VEGF sensitivity functions in a feedforward loop and contributes to cardiovascular aging. Data from our mouse models demonstrates the utility of exploiting this pathway to decrease pathological angiogenesis. On the basis of these studies, we propose that the MRN complex is novel anti-angiogenic target.
Materials and Methods
Cell Culture and Reagents
HUVECs and HMVECs (Lonza) were cultured in EGM-2 media (Lonza) supplemented with 10% Fetal Calf Serum (Hyclone). NHLFs (Lonza) were cultured in FBM media (Lonza) supplemented with 10% Fetal Calf Serum (Hyclone). 4T1 cells (ATCC) were culture in McCoy’s media, DMEM or RPMI-1640 supplemented with 10% Fetal Calf Serum and antibiotics. Cells were tested and found negative for mycoplasma contamination before use in the assays described. Mirin-1 and Vandetanib were purchased from Cayman Chemical and Selleckchem respectively. VEGF was purchased from PeproTech, Inc.
miRs/anti-miRs/siRNAs
miR mimics, inhibitors and respective controls were purchased from Life Technologies and Exiqon. For in vivo studies, high-performance liquid chromatography-purified miRs were purchased from Life Technologies in bulk quantities. siRNAs against MRE11a, RAD50 and NBN were purchased from Life Technologies. VEGFR2 Gapmer and MRE11a target protector were purchased from Exiqon.
Vectors/Plasmids
MRE11a Luciferase-3-UTR plasmid was purchased from SwitchGear Genomics. RAD50 and NBN luciferase constructs were generated by cloning the entire 3’UTR regions into pmiR-REPORT vector (Ambion). Luciferase assay reagents were purchased from SwitchGear Genomics and Promega.
Transfection
Cell were transfected at 50-60% confluence using standard forward transfection protocols using RNAimax reagent (Life Technologies) for miRs, siRNAs or gapmers and Lipofectamine 2000 for plasmid or plasmid RNA dual transfections. Typically 50 nM RNA and 1-2 μ g plasmid DNA were used for transfections. Target protectors were transfected at a concentration of 50 nM or equivalent to the miR amounts.
Radiation of Cells/Mice
Cells or mice were irradiation on a Shepherd□137cesium irradiator at a rate of ~166 cGy/min. In tumor-targeted radiation experiments, mice were restrained in a lead shield with flank cut-outs (Brain tree scientific) to minimize exposure to the non-tumor areas.
β-Gal Senescence assay
HUVEC were transfected for 48-72h with the microRNAs or siRNAs. After this time cells were washed with cold PBS and then stained for β-galactosidase activity following manufacturer’s protocol (Senescence Cells Histochemical Staining Kit, Sigma).
Telomerase activity assay
Cells were transfected with microRNAs or siRNAs for 24h. Cells were lysed and processed according to manufacturer’s instructions (Quantitative Telomerase Detection Kit, Allied Biotech). The telomerase activity level in the cell extract was determined through its ability to synthesize telomeric repeats onto an oligonucleotide substrate. The resultant extended product was subsequently amplified by polymerase chain reaction (PCR).
Cell Titer Glo/ Caspase Glo
Cells were transfected in a 6 well plate with miR mimics, siRNAs or inhibitors, and the corresponding controls from Life Technologies as previously described (59)‥ Cell Titer-Glo and Caspase 3/7 Glo were analyzed at 48-96h post treatments, according to manufacturer’s instructions.
Western blot and densitometric analysis
After treatment, cells were washed in phosphate-buffered saline (PBS) and lysed in RIPA buffer (Sigma) supplemented with Complete Protease inhibitor cocktail (ROCHE) and Phosphatase inhibitors cocktail 2 and 3 (Sigma). Lysed cells were harvested by scraping, and proteins were analyzed by Western blot. Equivalent amounts of protein were loaded on a 4-12% gradient SDS-polyacrylamide gel (BioRAD) and transferred for 30 min in a TransBlot turbo (BioRAD) onto Nitrocellulose membranes. Membranes were blocked in 5% milk or 3% BSA and incubated with antibodies as indicated: MRE11a (Cell Signaling, 4847, 1:1000), RAD50 (Cell Signaling, 3427, 1:1000), NBS1 (Cell Signaling, 14956, 1:500), p21 (Cell Signaling, 2947, 1:1000), pRb (Cell Signaling, 9301, 1:500),. β-actin (Sigma, A5316, 1:10,000 1□h RT) was used as a housekeeping control for the total levels of protein loaded. Membranes were washed in TBST and incubated with secondary antibodies from Licor Biosciences. Licor antibodies used were goat anti mouse 925-68020 (1:15,000) and goat anti rabbit 925-32211 (1:15,000). Blots were scanned on the Licor Odyssey scanner according to manufacturer’s instructions.
RNA extraction, RT-PCR, miR Profiling
Total RNA and microRNA were isolated using a miRVana microRNA isolation kit (Ambion). Reverse transcription was performed using TaqMan™ Advanced cDNA Synthesis Kit (Life Tech) according to the manufacturer’s instructions. RT-PCR was performed using multiplexed TaqMan primers (Applied Biosystems). The relative quantification of gene expression was determined using the 2-△△Ct method (35). Using this method, we obtained the fold changes in gene expression normalized to an internal control gene, GAPDH or U6 snRNA, respectively. For target analysis, a 92 gene DNA damage array (Life tech, 4418773) was used and for senescence phenotype profiling, the Human Cellular Senescence array was utilized (SA Biosciences) per manufacturer’s recommendations.
vH2AX staining
100,000 HUVECs were cultured on glass coverslips in 24-well plates and transfected with miRs/siRNAs using RNAimax (Life Technologies). Cells were fixed at different time points with 4% paraformaldehyde for 10 minutes at room temperature, permeabilized with 90% methanol for 10 minutes at 4°C.□□Coverslips were blocked with 1.5% normal goat serum (NGS) and incubated with primary antibody H2AX (Abcam, 11174, 1:500), at a 1:1000 dilution in NGS for 1h, washed and then incubated with secondary antibody for 30 minutes, washed and then mounted on glass slides for confocal imaging.
miR in-situ hybridization
In-situ hybridization was performed on frozen tumor sections as described in our previous studies (60, 61) using a DIG labeled miR-494 Locked Nucleic Acid (LNA) probe (Exiqon). DIG was detected by an anti-DIG HRP antibody (Roche) and amplified using a TSA-Plus Cy3 system (Perkin Elmer).
miR-TRAP/RISC TRAP assay
293 T cells were co-transfected with a plasmid coding for a flag-tagged dominant negative GW418 mutant (Clontech kit #632016) along with a control mimic, miR-99b or miR-494 mimic according to kit instructions. 24h later the RNA protein complexes were crosslinked and the RISC complex was immunoprecipitated using an anti-FLAG antibody and RNA was isolated for quantitative real-time PCR of target genes. The fold enrichment was calculated using pre and post IP controls as well as normalization to the control mimic pull-downs.
3-D Angiogenic Sprouting Assay
Early passage HUVECs were coated on cytodex-3 beads (GE Healthcare) at a density of 10 million cells/40 μl beads and incubated in suspension for 3-4 hours with gentle mixing every hour. They were plated on TC treated 6 well dishes overnight and resuspended in a 2mg/ml fibrin gel with 200,000 human smooth muscle cells. The gel was allowed to polymerize and complete EGM-2 media was added. Sprouts were visualized from days 3-4 via confocal imaging after overnight incubation with FITC labeled Ulex europaeus lectin (Vector labs). Immunofluorescence imaging was performed on a Yokogawa CSU-W1 spinning disk confocal microscope with 20x, 0.45 Plan Fluor objective (Nikon).
Flow cytometry
CD44 expression was analyzed by flow cytometry in HUVECs transfected with miR-494 for 48 hours. Cells were washed in PBS and trypsinized. Cells were incubated in blocking solution (0.5 %BSA/ 10% goat serum) for 30 min and then incubated for 1 hour in primary antibody-PEcy7 conjugated (CD44, BD, 560533) prepared in blocking solution. Then cells were washed 3X with PBS. Cells were analyzed in a CANTO II flow cytometer.
Cell cycle analysis
HUVEC were transfected for 48 hours with microRNAs, inhibitors or siRNAs. Cells were then harvested, washed, and fixed in 70% ice-cold ethanol at 4°C overnight. Then, cells were centrifuged, washed with cold PBS, and re-centrifuged. Cells were then re-suspended in 250□μL PBS and stained with 10□μL propidium iodide (PI, 1 Cmg/mL) and 10CμL RNase A (10□mg/mL) for 30□min at room temperature. DNA content was assessed using flow cytometry (CANTO II) to calculate the percentage of cells in subG1, G0/G1, S, and G2/M phases with FlowJo software.
Immunofluorescence and microscopy
In some experiments, CD31a and MRE11a were visualized using immunofluorescence staining from OCT sections. Slides were fixed with 4% PFA and stained overnight for CD31 488 (BD bioscience 611986 1:200 o/n), MRE11a (Cell Signaling Technology 1:100) and Phalloidin Alexa 647 (1:50). For MRE11a antibody, following day fixed cells were incubated with Goat anti-Rabbit Alexa 488 (1:500) in 5%BSA/TBS for 1 hour. Imaging was performed on a Nikon Spectral C1 confocal microscope (Nikon C1si with EZC1 acquisition software, Nikon Instruments) with Plan Apo 10X/0.45 air, Plan Apo 20X/0.75 air, and Plan Apo 60X/1.40 oil objective lenses (Nikon). Some immunofluorescence imaging was performed on a Yokogawa CSU-W1 spinning disk confocal microscope with 20 0.45 Plan Fluor objective (Nikon). All images were taken with a channel series. Images were analyzed with Image J software for quantitation.
pERK ELISA
We measured the abundance of p-ERK with an ELISA kit (PathScan® Phospho-p44/42 MAPK (Thr202/Tyr204) Sandwich ELISA, Cell Signaling Technology) following manufacturer’s instructions.
In vivo assays
All animal work was approved by the OHSU Institutional Animal Use and Care Committee. Immune-compromised 8-10 week old female nu/nu mice were purchased from Jackson Labs. Growth factor reduced Matrigel (BD) with 400 ng/ml recombinant human bFGF (Millipore) was injected subcutaneously in nu/nu mice. Mice were injected i.v. with 7C1-nanoparticles containing miR-494 or control miR (~1mg/kg, i.v) 3 or 4 days after plugs were implanted. At day 7 mouse tissues were harvested and processed to obtain RNA or frozen in OCT for tissue staining. 4T1 cells (1X10s^4) were implanted into the mammary fat pad #4 of 6-8 week old female Balb/C mice in 100μl Matrigel. Mice were randomized into groups once the average tumor volume reached 150 mm3, approximately 10 days after implantation. Mice were treated with 7C1-nanoparticles containing miR-103 or control miR (0.7 mg/kg, i.v.). Mice were euthanized ~ day 18-20 for analysis of metastatic burden in lungs. Additional experiments were carried out in postpubertal female FVB/n mice (10-12 weeks), which received injections of 50,000 PyMT-derived tumor cells into the right lower mammary fat pad for induction of orthotopic tumors as described in (62).
Statistics
All statistical analysis was performed using Excel (Microsoft) or Prism (GraphPad). Two-tailed Student’s t-test was used to calculate statistical significance. Variance was similar between treatment groups.
Conflict of Interest
C.E.S and S.A. are named inventors on a US patent application that is based on some of the findings reported in this manuscript.
Acknowledgements
We thank Lisa Coussens (OHSU) for help with the PyMT model and sharing archived FFPE tissue sections from (62). We thank Sergio Fazio and Stephen Lloyd (OHSU) for useful discussions. We thank Shushan Rana and Katherine Kelly (OHSU) for comments on the manuscript. We acknowledge the OHSU Advanced Light Microscopy Core, Knight Cancer Institute Flow Cytometry Core and the Gene Profiling Shared Resource for technical help and useful discussions. This work was supported by US NIH grant R00HL112962, R56HL137779 and an innovative research grant from the American Heart Association (17IRG33400218) all to S.A.