Lysophosphatidic Acid Receptor 4 Activation Augments Drug Delivery in Tumors by Tightening Endothelial Cell-Cell Contact

Article

Lysophosphatidic Acid Receptor 4 Activation Augments Drug Delivery in Tumors by Tightening Endothelial Cell-Cell Contact Graphical Abstract

Authors Kazuhiro Takara, Daisuke Eino, Koji Ando, ..., Satoshi Ishii, Haruhiko Kishima, Nobuyuki Takakura

Correspondence [email protected]

In Brief Takara et al. find that lysophosphatidic acid (LPA) promotes fine capillary network formation and improves drug delivery in tumors. LPA controls localization of VE-cadherin in endothelial cells through LPA receptor 4 (LPA4) signaling.

Highlights d

Lysophosphatidic acid (LPA) promotes fine vascular network formation in tumors

d

LPA improves drug delivery into tumors

d

Vascular network formation by LPA is induced by LPA4 through Gi or Ga12/13 activation

d

LPA promotes membrane localization of VE-cadherin in endothelial cells

Takara et al., 2017, Cell Reports 20, 2072–2086 August 29, 2017 ª 2017 The Author(s). http://dx.doi.org/10.1016/j.celrep.2017.07.080

Cell Reports

Article Lysophosphatidic Acid Receptor 4 Activation Augments Drug Delivery in Tumors by Tightening Endothelial Cell-Cell Contact Kazuhiro Takara,1,2,6 Daisuke Eino,1,3,6 Koji Ando,4 Daisuke Yasuda,5 Hisamichi Naito,1 Yohei Tsukada,1 Tomohiro Iba,1 Taku Wakabayashi,1 Fumitaka Muramatsu,1 Hiroyasu Kidoya,1 Shigetomo Fukuhara,4 Naoki Mochizuki,4 Satoshi Ishii,5 Haruhiko Kishima,3 and Nobuyuki Takakura1,7,* 1Department of Signal Transduction, Research Institute for Microbial Diseases, Osaka University, 3-1 Yamada-oka, Suita, Osaka 565-0871, Japan 2Research Unit/Frontier Therapeutic Sciences Sohyaku, Innovative Research Division, Mitsubishi Tanabe Pharma Corporation, 1000 Kamoshida-cho, Aoba-ku, Yokohama 227-0033, Japan 3Department of Neurosurgery, Osaka University Graduate School of Medicine, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan 4Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Osaka 565-8565, Japan 5Department of Immunology, Akita University Graduate School of Medicine, Akita University, Akita 010-8543, Japan 6These authors contributed equally 7Lead Contact *Correspondence: [email protected] http://dx.doi.org/10.1016/j.celrep.2017.07.080

SUMMARY

Vascular normalization in tumors may improve drug delivery and anti-tumor immunity. Angiogenesis inhibitors induce hypoxia, which may facilitate malignant progression; therefore, we investigated other methods to promote vascular maturation. Here, we show that lysophosphatidic acid (LPA) enhances blood flow by promoting fine vascular networks, thereby improving vascular permeability and suppressing tumor growth when combined with anti-cancer drug treatment. Six different G protein-coupled receptors have been identified as LPA receptors (LPA1–6). In studies using mutant mice, we found that LPA4 is involved in vascular network formation. LPA4 activation induces circumferential actin bundling beneath the cell membrane and enhances linear adherens junction formation by VE-cadherin in endothelial cells. Therefore, we conclude that activation of LPA4 is a promising approach for vascular regulation. INTRODUCTION Angiogenesis is a fundamental requirement for tumor growth, and therefore it was widely believed that anti-tumor angiogenesis would be a promising approach to block cancer growth (Bergers and Benjamin, 2003). Accordingly, inhibitors of vascular endothelial growth factor-A (VEGF) and its cognate receptors (VEGFRs) have been developed and utilized clinically. Neutralizing antibody against VEGF (bevacizumab) was the first such approved drug for colon cancer and has been used globally (Ferrara et al., 2004). However, based on the evidence that

combination therapy using angiogenic inhibitors and anti-cancer drugs is more effective than monotherapy with either alone, a different concept has emerged, namely, that angiogenic inhibitors in fact normalize immature and leaky blood vessels in the tumor, resulting in improved drug delivery (Jain, 2001). Despite the anticipated benefits of angiogenic inhibitors in cancer therapy, there has been concern that side effects such as hypertension, lung hemorrhage, and renal dysfunction due to vascular damage can be induced by excessive amounts of these drugs. Moreover, hypoxia resulting from blood vessel regression in the tumor causes malignant conversion of cancer cells that facilitates invasion and metastasis (Ebos et al., 2009; Pa`ez-Ribes et al., 2009). Therefore, sustained normalization of the tumor vasculature by methods other than the use of angiogenic inhibitors may enhance effective drug delivery, resulting in improved anti-tumor effects without stimulating malignant behavior of cells in the tumor microenvironment. Gene modification models suggest that homozygous mice lacking Rgs5, a G protein activator expressed on pericytes, or mice heterozygous for proline hydroxylase (PHD2) for stability of hypoxia-inducible factor 1a generate blood vessels in tumors that appear normal (Hamzah et al., 2008; Mazzone et al., 2009). Mice carrying a Sox17 mutation, a molecule crucial for embryonic endoderm formation, also have mature blood vessels in their tumors (Yang et al., 2013). In these mice, it is commonly observed that sustained tumor vessel normalization improves drug delivery and inhibits tumor metastasis, suggesting a benefit of long-lasting tumor vasculature normalization for suppression of malignant progression. Recently, in addition to angiogenic growth factor proteins, such as VEGF, angiopoietin, and platelet-derived growth factor, critical roles of lipid mediators such as lysophosphatidic acid (LPA) and sphingosine-1-phosphate (S1P) have also been documented in vascular formation and integrity (Kono et al., 2004; Yukiura et al., 2011). LPA is generated via several pathways from

2072 Cell Reports 20, 2072–2086, August 29, 2017 ª 2017 The Author(s). This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

Figure 1. Vascular Network Formation Stimulated by LPA in Tumors (A–C) Thick slices of LLC tumor were stained with anti-CD31 mAb. Tumor-bearing mice were treated with vehicle (A) or LPA (B) daily for 5 days. Right-hand panel shows a higher magnification of the center of the tumor indicated by the dashed box in the left-hand panel. Thickness, 150 mm. Scale bar, 1 mm (left panels) and 200 mm (right panels). (C) Quantification of total vessel length at the tumor center (n = 6 tumors per group).

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lipid precursors, with its production from lysophosphatidylcholine (LPC) by the enzymatic activity of extracellular autotaxin (ATX) having been most extensively analyzed (Aoki et al., 2002). LPA has several functions in cell proliferation, migration, morphological change, and anti-apoptotic effects mediated by its binding to specific receptors (LPA1–6). These are coupled to diverse G proteins such as Gi, G12/13, Gq, and Gs (Yanagida and Ishii, 2011). Knocking out the ATX gene in mice results in embryonic lethality around embryonic day 10.5, accompanied by abnormal blood vessel formation in the yolk sac, placenta, and elsewhere (Tanaka et al., 2006). Moreover, it has been reported that targeted disruption of LPA4 leads to embryonic lethality caused by disorganized blood and lymphatic vessels (Sumida et al., 2010). LPA also has a role in the proliferation of endothelial cells (ECs) (Yin and Watsky, 2005). Therefore, inhibition of LPA and its downstream signaling has gained increased attention as a means to suppress tumor angiogenesis (Su et al., 2013). Contrary to the previous concept, however, here, we show that stimulation via the LPA receptor induces enhanced vascular network formation and improves drug delivery into tumors, resulting in significant anti-tumor effects. RESULTS LPA Promotes Vascular Network Formation in the Tumor When the tumor size reached 30–50 mm3 7–10 days after subcutaneous inoculation of Lewis lung carcinoma (LLC) cells, we initiated treatment by injecting LPA (3 mg/kg) intraperitoneally (i.p.) for 5 days. In the absence of LPA, as expected, blood vessels showed tortuous and discontinuous patterns, especially in the center of the tumor (Figure 1A). In contrast, after LPA administration, larger blood vessels aligned in parallel and fine capillary network formation was induced even in the center of the tumor, furthermore, vessel length was increased in LPA-treated tumors (Figures 1B and 1C). LPA dose-dependently induced vascular network formation, suggesting its direct effects on ECs (Figures S1A and S1B). In order to determine the time course of vascular network formation in these tumors, a single injection of LPA was given to tumor-bearing mice. As shown in Figures 1D and 1E, enhancement of vascular network formation was observed in a time-dependent manner. Expression of the ETS-related gene (ERG), which is normally expressed in ECs (Birdsey et al., 2008, 2015), was observed in elongated ECs in vessels with extended lengths (Figures 1D and 1F). LPA did not increase the number of Ki67-positive nuclei in ERG-positive ECs (Figures S1C and S1D). Pericyte coverage was not significantly altered

over 24 hr (Figures S1E and S1F). These data suggest that LPA does not affect proliferation of ECs, but does induce morphological changes, which lead to vascular network formation in tumors. The vascularization-promoting effect of LPA was not only observed in the LLC tumor, but also in colon26 tumors using the same schedule as described in Figure 1A (Figures S1G and S1H). Next, we examined the intraluminal surface of tumor blood vessels by emission scanning electron microscope and found that ECs did not tightly adhere to each other, inter-endothelial gaps were observed, suggesting loose inter-cellular junctions in the lumen of the vessel. However, fewer such inter-endothelial gaps were observed 24 hr after LPA administration, and the surface of the blood vessels from both LLC and colon26 tumors had become smooth (Figures 1G–1I). These effects were induced in all tumors evaluated in our experiments. These are morphological characteristics of mature ECs and have been designated as changes into a ‘‘phalanx’’ phenotype, suggested to reflect tight inter-endothelial junctions and to inhibit tumor metastasis (Mazzone et al., 2009). Accordingly, we investigated whether LPA-induced vascular surface smoothing inhibits tumor metastasis using a model of lung metastases originating from subcutaneous tumors of Ex-3LL (derived from LLC and exhibiting enhanced lung metastasizing ability) or from the highly metastatic melanoma cell line B16BL/6. Both cell lines were engineered to express EGFP. The number of metastases from LPA-treated tumors was significantly lower than from control tumors (Figures 1J, 1K, S1I, and S1J). The metastasis-inhibitory effect of LPA was absent when B16BL/6-EGFP tumor cells were injected directly into the tail vein (Figures S1K and S1L). Therefore, we conclude that LPA not only induces vascular network formation, but also the maturation of these blood vessels possibly by EC-to-cell tightening, and that this reduces metastasis. LPA Promotes Functional Blood Vessel Formation in Tumors We next investigated whether fine capillary network blood vessels in tumors treated with LPA were indeed functionally normal. To assess this, we injected LPA into 30–50 mm3 tumor-bearing mice and, 24 hr later, we injected a lectin that binds to ECs in order to evaluate blood flow and perfusion (Inai et al., 2004; Morikawa et al., 2002). After 10 min in the circulation, this lectin usually labels almost all ECs in normal tissue (Naito et al., 2012) (Figure S2), but in tumors of untreated mice, approximately half the vessels were unmarked, suggesting disturbed perfusion (Figure 2A). However, after injection of LPA, most blood vessels,

(D–F) Time course of tumor vascular network formation after one injection of LPA. Tumor-bearing mice were injected with LPA 6–24 hr before sacrifice; control mice received nothing. All the tumors were harvested at the same time. (D) Representative images of LLC tumor center costained with anti-ERG (red) and antiCD31 (green) mAb. Thickness, 10 mm. Scale bar, 100 mm. (E) Quantification of total vessel length per field (n = 5 tumors per group). (F) Quantification of vessel length per single endothelial cell (n = 5 tumors per group). (G–I) Scanning electron microscopy of LLC and colon26 tumor vessels 24 hr after treatment with vehicle or LPA. The higher magnifications of the areas indicated by the dashed box in the middle panel are shown in the panel on the right. Scale bars, 10 mm (middle panels) and 5 mm (left- and right-hand panel). An interendothelial gap is detected in the vessel lumen (yellow arrowhead). (H and I) Percentage of inter-endothelial gap-positive vessels of colon26 (H) and LLC (I). 5 to 10 vessels were evaluated in each tumor sample (n = 6 tumors per group). (J and K) Metastatic lung tumor spreading from subcutaneous EX-3LL-GFP (J) tumor treated with vehicle or LPA (K). (J) Serial sections were stained with H&E (left panels) or with immunofluorescent antibodies (right panels) anti-GFP (green) pAb, anti-CD31 (red) mAb, and nuclei (blue). Scale bar, 1 mm. (K) Quantification of metastatic foci in mouse lung sections (n = 6 mice per group). All experiments were repeated at least twice. The error bars indicate mean ± SEM. *p < 0.05, **p < 0.01, and ***p < 0.001. See also Figure S1.

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Figure 2. Improved Blood Vessel Function after LPA Treatment of LLC Tumor-Bearing Mice (A and B) Vessel perfusion of LLC tumors visualized by FITC-conjugated lectin. (A) Representative images stained with anti-CD31 mAb (red). The dashed lines show the tumor boundary. Scale bar, 500 mm. (B) Quantification of perfused blood vessels of the tumor center by ennummerating CD31+lectin+ vessels within total CD31+ vessels (n = 6 tumors per group). (C and D) Penetration of doxorubicin (red) into the LLC tumor center. (C) Sections of tumors from mice treated with vehicle or LPA were stained with anti-CD31 mAb (green). Scale bar, 100 mm. (D) Quantification of doxorubicin area (n = 12 tumors per group). (E and F) Tissue hypoxia at the LLC tumor center. (E) Hypoxia within the LLC tumor was estimated by hypoxyprobe staining (pimonidazole; green). ECs were stained with anti-CD31 mAb (red). Scale bar, 200 mm. (F) Quantification of hypoxic area (n = 5 tumors per group). All experiments were repeated at least twice. The error bars indicate mean ± SEM. **p < 0.01 and ***p < 0.001. See also Figure S2.

including those deep inside the tumor, were positive for lectin (Figures 2A and 2B), indicating that blood flow is improved after injection of LPA. Next, we tested changes of drug delivery using doxorubicin, an anti-cancer drug that auto-fluoresces, which was intravenously injected 20 min before tumor dissection. Greater penetration of the drug into tumors was observed in LPA-treated mice (Figures 2C and 2D). This suggests that LPA treatment effectively improves permeability and facilitates access of chemotherapeutic drugs to the tumor. Additionally, we analyzed whether the tissue hypoxia usually observed in the tumor microenvironment was reduced by LPA. For this, we used pimonidazole HCL as a hypoxyprobe 24 hr post-LPA injection. The data indicate that hypoxia was clearly reduced in the tumors of the LPA-injected group (Figures 2E and 2F). Taking all these data together, we conclude that LPA induces functional vascular network formation in tumors. Combination of LPA with Anti-cancer Drug Treatment Effectively Inhibits Tumor Growth Having shown that LPA induces well-organized vascular network formation in tumors with improved functional vascular permeability, we next investigated the therapeutic effects of combining anti-cancer drug treatment with LPA (Figure 3A). We inoculated LLC, B16BL/6 cells, or colon26 subcutaneously and treated the mice from day 6 with either vehicle or LPA (3 mg/kg) daily until

day 20 with or without fluorouracil (5-FU; 100 mg/kg) on days 7 and 14. As depicted in Figures 3B–3E, 5-FU alone exerted an anti-tumor effect in each tumor. However, the combination of 5-FU with LPA (LPA/ 5-FU) inhibited tumor growth more markedly. LPA alone tended to inhibit the growth of LLC and B16BL/6 tumors, but not of colon26, although these differences did not achieve statistical significance. In order to test the effect of LPA alone, we evaluated proliferation and apoptosis of subcutaneous LLC tumors. LPA did not affect the number of Ki67-positive cells or terminal deoxynucleotidyl transferase (TdT)-mediated 2’-deoxyuridine 5’-triphosphate (dUTP) nick-end labeling (TUNEL)positive cells (Figures S3A–S3D). The number of cells containing cleaved caspase-3 was slightly, but not significantly, increased in LPA-treated tumors (Figures S3E and S3F). These data suggest that LPA alone does not significantly influence tumor growth. Oxaliplatin (L-OHP; 1.5 mg/kg) has a different mode of action than 5-FU and, when used instead of 5-FU in colon26 tumors together with LPA according to the same treatment schedule, the combination effectively inhibited tumor growth relative to L-OHP alone (Figure S3G). Assessing the quantity of remaining cancer cells by histology after therapy, the amount per whole tumor mass was reduced by 5-FU, however, in tumors treated with a combination of 5-FU and LPA, this reduction was much more marked (Figures 3F–3I). The LPA-Induced Vascular Network Formation in Tumors Is Mediated via LPA4 LPA binds to six different G protein-coupled receptors with seven transmembrane domains (LPA1–6). It has been well-established that cancer cells frequently express LPA1 or LPA3

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Figure 3. Anti-tumor Effect of a Combination of LPA with an Anti-cancer Drug (A) Schema of LPA treatment. (B, D, and E) LLC tumor growth (B, n = 6 tumors per group), B16BL/6 (D, n = 6 tumors per group), or colon26 (E, n = 4 tumors per group). The tumor-bearing mice were treated with LPA and/or 5-FU. (C) Gross appearance of dissected LLC tumors. (F) LLC-GFP tumor sections on day 21 treated with LPA and/or 5-FU as indicated. LLC cells expressing EGFP were visualized by staining with anti-GFP pAb. Scale bar, 200 mm. (G) Remaining cancer cells were evaluated for GFP-positive areas (n = 8 tumors per group). (H) H&E staining of LLC-GFP tumor sections. The inset in each panel shows a higher magnification of the dashed line box. Scale bar, 200 mm. (I) Evaluation of nuclear density (n = 8 tumors per group). All experiments were repeated at least twice. The error bars indicate mean ± SEM. *p < 0.05, **p < 0.01, and ***p < 0.001. See also Figure S3.

(Su et al., 2013), however, LPA receptor expression in ECs, especially ECs in tumors, has not been precisely determined. We inoculated LLC cells expressing EGFP to investigate LPA

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receptor expression in LLC tumors. Cells in the tumor tissues on day 14 were sorted into three fractions, namely, cancer cells (identified as EGFP+), ECs (EGFP
CD45
CD31+), and other

Figure 4. Vascular Network Formation by LPA Is Mediated through LPA4 (A) Quantitative real-time PCR analysis for LPA receptors 1–6 expressed in cancer cells, ECs, and PDGFRb-positive stromal cells (n = 6 tumors per group). (B and C) LPA4 (B) or LPA6 (C) was significantly expressed in ECs from LLC tumors (n = 6 tumors per group). (D and E) LLC tumors were developed in wild-type, LPA4 knockout (KO), or LPA6KO mice and were harvested 24 hr after administration of vehicle or LPA. (D) Representative image of tumor center stained with anti-CD31 mAb. Scale bar, 100 mm. (E) Quantification of total vessel length per field (n = 5 tumors per group). (F and G) LLC tumors developed in wild-type or LPA4KO mice treated with vehicle or VPC31144(S) (3 mg/kg) for 5 days. (F) Representative images of anti-CD31 staining. The dashed lines indicate the tumor edge. Scale bar, 500 mm. (G) Quantification of vessel length at the tumor center (n = 5 tumors per group). (H) LLC tumor sections developed in wild-type (WT) or LPA4KO mice were stained with anti-LPA4 (green) and CD31 (red) antibodies. Scale bars, 100 mm. All experiments were repeated at least twice. The error bars indicate mean ± SEM. **p < 0.01 and ***p < 0.001. See also Figure S4.

stromal cells (CD45
CD31
PDGFRb+). We found that LLC cells express LPA1 and LPA5, stromal cells express LPA1, and ECs strongly express LPA4 and LPA6, and LPA1 weakly (Figure 4A). Tumor ECs expressed LPA4 or LPA6 at significantly greater levels than stromal cells (Figures 4B and 4C). ECs isolated from B16BL/6 and colon26 tumors also expressed LPA4 and LPA6 (Figures S4A and S4B). We additionally isolated ECs from lung, liver, and skin of wild-type mice and investigated LPA receptor expression. We found that expression of LPA receptors other than LPA4 and LPA6 differs in different organs,

but that LPA4 and LPA6 are expressed in ECs from all tissues tested (Figures S4C–S4E). In order to identify which LPA receptor is involved in the induction of vascular network formation in tumors, we inoculated LLC cells into wild-type, LPA4
/
(Sumida et al., 2010) or LPA6
/
(Hata et al., 2016) mice and tested the effect of LPA treatment. As shown in Figures 4D and 4E, LPA induced vascular network formation in tumors generated in the LPA6
/

mice to the same degree as in wild-type mice, however, it failed to induce vascular network formation in tumors developing in LPA4
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Figure 5. LPA Promotes EC Membrane Localization of VE-Cadherin (A–C) Localization of VE-cadherin in ECs of LLC tumors. (A) Tumor-bearing mice were treated with vehicle (upper) or LPA (lower) 24 hr before tumor resection. Sections were stained with anti-CD31 (red) and anti-VE-cadherin (green) antibodies, and nuclei were labeled with TOPRO3 (blue). Five tumors per group were

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mice. In addition, improvement of lectin perfusion and dextran extravasation induced by LPA was no longer seen in LPA4
/
mice, but the LPA6
/
mice were not affected (Figures S4F–S4I). This suggests that LPA4 is the receptor responsible for vascular network formation in tumors. Based on this result, we utilized VPC31144(S), an analog of LPA which preferentially binds LPA4 rather than LPA6 (Yanagida et al., 2013), and studied its effects on blood vessel formation. We found that like LPA, VPC31144(S) also induced vascular network formation in wild-type mice, but that this effect was also absent in LPA4
/
mice (Figures 4F and 4G). Moreover, when LLC tumor tissue sections were analyzed, we confirmed that most of the CD31+ ECs were positive for LPA4 at the protein level (Figure 4H). These data confirm that LPA4 is involved in the vascular-promoting effects of LPA. LPA Controls the Cell Membrane Localization of VE-Cadherin Improved vascular permeability and oxygenation suggested that LPA controls vascular EC-cell contact. Therefore, we investigated expression of the adherent junction protein VE-cadherin in the blood vessels of LLC tumors treated as described in Figure 4D. First, we isolated ECs from LLC tumors treated or not treated with LPA. We found that LPA does not enhance VEcadherin mRNA expression (Figure S5A). Next, we performed dual staining of LLC tumor sections with CD31 and VE-cadherin antibodies. In the controls, VE-cadherin was localized in the cytoplasm of ECs and did not merge with EC membrane protein CD31 (Figures 5A and 5B). In contrast, in the LPA-treated tumors, VE-cadherin localized together with CD31 at the cell membrane (Figures 5A and 5C). In order to assess the effect of LPA on the membrane localization of VE-cadherin, we tested commercially available EC lines for LPA receptor expression and selected MS-1, which expressed LPA4 and LPA6 as observed in tumor ECs (Figures S5B and S5C). It is known that VEGF induces internalization of VE-cadherin into the cytoplasm, associated with vascular leakage (Taddei et al., 2008). It has been reported that VEGF-A induces phosphorylation and internalization of VE-cadherin, leading to vascular permeability (Dejana et al., 2008). We hypothesized that LPA affects VE-cadherin phosphorylation in ECs. In MS-1 cells, VE-cadherin was phosphorylated in the absence of VEGF and additional VEGF treatment did not influence this. Moreover, LPA did not affect VE-cadherin phosphorylation in the presence or absence of VEGF (Figure S5D). These results therefore suggest that LPA affects cell-cell junctions of ECs by other mechanisms. It is known that VE-cadherin at the inter-cellular junction lines up with circumferential actin bundles beneath the cell membrane

and promotes adherent junction integrity (Noda et al., 2010). We stimulated confluent MS-1 cells with VEGF or LPA, we found that exposure to VEGF alone resulted in decreased amounts of cortical actin fiber at inter-cellular junctions, relative to controls. However, in MS-1 cells, VE-cadherin is already phosphorylated without additional VEGF, explaining why VE-cadherin expression at cell-cell junctions is not altered by this factor. Under these circumstances, however, LPA did result in dose-dependent increases of cortical actin fiber even when stimulated with VEGF, as well as in unstimulated cells. This was accompanied by increased VE-cadherin expression at inter-cellular junctions (Figures 5D–5F and S5E–S5G). Hyper-permeability of blood vessels in the tumor eventually causes interstitial hypertension, resulting in impaired oxygen and drug delivery. Hence, drug delivery into tumors is improved by the restoration of EC barrier function (Carmeliet and Jain, 2011). Because we had shown that LPA promotes VE-cadherin localization to the EC membrane, we asked whether it also enhances barrier function. To do so, we monitored the electrical impedance of monolayers of MS-1 cells using electrical cellsubstrate impedance sensing (ECIS; http://www.nepagene.jp/ index2.html). As depicted in Figures 5G and S5H, treatment with LPA or VPC31144(S) rapidly increased EC barrier function, with a peak effect approximately 60 min after beginning the treatment. Barrier function was continuously maintained for at least 400 min and was increased in a dose-dependent manner by VPC31144(S) and LPA (Figures 5H and S5I). These data suggest that LPA promotes cortical actin fiber amplification and supports membrane localization of VE-cadherin, which leads to tightening of inter-endothelial junctions. LPA4 Signaling Enhances Barrier Function and Network Formation in ECs We showed that tumor vascular formation was promoted by LPA4. In order to determine whether LPA-induced cortical actin fiber amplification and reinforcement of the inter-cellular junctions between MS-1 cells was mediated by LPA4, we blocked the expression of either LPA4 or LPA6 by small interfering RNAs (siLPA4 or siLPA6). Knockdown efficacy was evaluated by quantitative real-time PCR (Figures S6A and S6B). As depicted in Figures 6A–6C, cortical actin fiber formation and VE-cadherin recruitment induced by LPA was prevented by LPA4, but not by LPA6 knockdown. Next, we investigated the effect of LPA4 signaling on primary ECs. We isolated ECs from normal lung of wild-type mice and cultured them on confluent monolayers of OP9 cells. In the control cultures, ECs aggregate together and form a thick network of branches. In contrast, under VPC31144(S) stimulation, ECs form

analyzed and four vessels per tumor section were randomly chosen from tumor center. Representative images were shown in (A). The dashed white box in the left panel is magnified in the three right-hand panels in each group. Scale bar, 20 mm (left panels) and 10 mm (high-power field). (B and C) The fluorescence intensity profile of VE-cadherin (green line) and CD31 (red line) in the ECs indicated by the white dashed lines in the high-power fields of (A). (D–F) Localization of VE-cadherin (green) and F-actin (red) in MS-1 cells. (D) Nuclei were labeled with TOPRO3 (blue). The cells were treated by VEGF without or with different concentrations of LPA as indicated. The dashed white boxes in the third line are magnified in the lower panels. Scale bar, 50 mm (upper three lines) and 10 mm (lower three lines). (E and F) Quantification of VE-cadherin (E) and F-actin (F) at cell-cell junctions relative to control (n = 16 points per group). (G and H) The relative resistance between cells (Rb) in monolayer-cultured MS-1 cells treated with LPA or different concentrations of VPC31144(S) was measured using the ECIS system. (H) The relative Rb values at 60 min after stimulation as shown in (G) (n = 5–7 experiments per group). The error bars indicate mean ± SEM. *p < 0.05, **p < 0.01, and ***p < 0.001. See also Figure S5.

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Figure 6. Analysis of Cell Signaling through LPA4 for Barrier Function in ECs (A–C) Monolayers of MS-1 cells transfected with siRNA for negative control (siCTR), LPA4 (siLPA4), or LPA6 (siLPA6) and stimulated with 10 mM LPA or vehicle, then stained with anti-VE-cadherin mAb (green) and phalloidin (red). (A) Nuclei were labeled with TOPRO3 (blue). The dashed white boxes in the upper panels are magnified in each of the three lower panels. Scale bar, 50 mm (upper panels) and 10 mm (lower three panels). (B and C) Quantification of VE-cadherin (B) and F-actin (C) at cell-cell junctions relative to control (n = 16 points per group). (D–G) CD31+ ECs were isolated from lung of wild-type mice (D) or LPA4
/
mice (F) and seeded on confluent OP9 cells. Cells were stimulated with vehicle or different concentrations of VPC31144(S) and stained with anti-CD31 mAb. The black dashed boxes in the upper panels are magnified in the lower panels. Scale bar, 200 mm (upper panel) and 50 mm (lower panels). (E and G) Quantification of network vessel length of wild-type mice (E) and LPA4
/
mice (G). Quantification was conducted using low-power fields (n = 6 fields per group). The error bars indicate mean ± SEM. **p < 0.01 and ***p < 0.001. See also Figure S6.

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fine networks of branches in a dose-dependent manner (Figures 6D and 6E). This network formation induced by VPC31144(S) was not observed in cultures of primary ECs from LPA4
/
mice (Figures 6F and 6G). These data suggest that LPA4 regulates vascular network formation. Analysis of Cell Signaling through LPA4 That Is Required for Barrier Function in ECs First, we hypothesized that Gi is the signal transduction candidate, because Akt activation is involved in the stabilization of ECs, and Gi has been suggested to activate Akt through phosphatidylinositol 3-kinase (PI3K) (Ambesi and McKeown-Longo, 2009). Therefore, we utilized the Gi inhibitor pertussis toxin (PTX) (van Corven et al., 1993). PTX itself did not affect barrier function of ECs, however, the VPC31144(S)-mediated increase of barrier function was suppressed by PTX (Figures S7A and S7B). Moreover, VPC31144(S)-induced linear adherence junction formation by VE-cadherin was also perturbed by PTX (Figures 7A–7C). Next, we assessed the involvement of Ga12/13 by silencing GNA13 (siGNA13) (Figure S7C) and found that cortical actin fiber formation induced by VPC31144(S) was inhibited (Figures 7D– 7F). To investigate the involvement of other G proteins, we tested Gs and Gq inhibitors. SQ 22536 and 20 50 -dideoxyadenosine (20 50 DDS) are adenylyl cyclase inhibitors and were used as Gs inhibitors. YM-254890 was used to inhibit Gq. Cortical actin fiber formation induced by VPC31144(S) was observed even under treatment with these inhibitors (Figures S7D–S7F). Thus, we conclude that Gi and Ga12/13 mediate the increased endothelial barrier function resulting from LPA4 signaling. DISCUSSION Here, we show that treatment with LPA converts non-functional blood vessels in tumors into functional vessels with affluent blood flow by inducing fine capillary network formation. Morphological changes of blood vessels caused by LPA are the result of elongation of ECs mediated through LPA4 signaling. LPA4 promotes cortical actin fiber formation of ECs and stabilizes VE-cadherin, which lines up with circumferential actin bundles beneath the cell membrane, which contributes to inter-endothelial tightening, reduces vascular permeability, and facilitates the effects of LPA together with anti-cancer drugs on inhibition of tumor growth. Previously, it was reported that LPA induced vascular permeability in experiments using human umbilical vein ECs (HUVECs) (Yanagida et al., 2009). In contrast, we found that LPA inhibited vascular permeability using MS-1 cells. This difference may be due to the dominant expression of LPA6 rather than LPA4 by HUVECs (data not shown), because we found that the LPA-mediated anti-permeability effect was dependent on LPA4. This suggests that the anti-permeability action of LPA4 outweighs any LPA6-mediated vascular permeability enhancing effect when both LPA4 and LPA6 are expressed by the ECs, as is the case with the MS-1 cell line. The induction of vascular network formation occurs very rapidly after injection of LPA (Figures 1D and 1E). To account for this, it remains possible that LPA may induce proliferation of ECs for network formation. However, considering the usual doubling time of cells and the approximate doubling of vessel

length by 24 hr after LPA injection (Figure 1E), it seems unlikely that the mere proliferation of ECs stimulated by LPA could explain this phenomenon. To prove this, we costained LLC tumors with anti-Ki67 and anti-ERG antibodies and showed that LPA did not promote proliferation of ECs. In vitro, we confirmed that LPA did not affect MS-1 cell growth (data not shown). LPA promoted cortical actin fiber formation of MS-1 in 1 hr. Therefore, one possible mechanism responsible for these findings is that rapid cortical actin fiber amplification induced by LPA leads to rapid elongation of ECs and network formation in 24 hr. On the other hand, LPA did stimulate the migration of MS-1 cells (data not shown). This could also contribute to network formation. It has been previously reported that EC network formation is disrupted by VE-cadherin inhibition (Bach et al., 1998). In our study, we found that LPA promotes membrane localization of VE-cadherin in ECs. Taken together, these data strongly suggest that LPA induces network formation by promoting cortical actin fiber formation and following stabilization of VE-cadherin rather than by stimulating their proliferation. It has also been reported that LPA induces proliferation of cancer cells mainly through LPA1 (Tsujiuchi et al., 2014). Therefore, inhibitors for LPA1 have been developed and their effects on tumor growth in mice investigated (Su et al., 2013). LPA1 is involved in the pathogenesis of pulmonary fibrosis and systemic sclerosis, and several LPA1 inhibitors have been employed in successful phase I and phase II clinical trials for these diseases (Stoddard and Chun, 2015). An effect on tumor growth inhibition would also be expected. We do not exclude a benefit of those drugs for tumor inhibition, however, we showed here that LPA did not increase the number of Ki67-positive cells in LLC tumors, and that treatment with LPA to improve drug delivery may be clinically effective via LPA4 signaling. Contrary to initial expectations, we found that LPA alone did not induce tumor growth and inhibited LLC and B16BL6 tumors to some extent even without the addition of any anti-cancer drugs. It has been suggested that normalization of the tumor vasculature allows the penetration of leukocytes including CD8+ T cells into the tumor (Huang et al., 2012). Moreover, LPA is itself involved in lymphocyte trafficking (Hata et al., 2016). Therefore, it is possible that LPA treatment improves tumor immunity to inhibit tumor growth, and that combinations of LPA with immunotherapy could be useful, but further analysis is required to determine this. Here, we found that LPA induces membrane localization of VE-cadherin in ECs, mediated via LPA4, which is involved in cortical actin formation via Gi and/or Ga12/13. However, how activation of these two G proteins induces membrane localization of VE-cadherin has not been clearly elucidated. As described above, Gi signaling activates Akt via PI3K (Ambesi and McKeown-Longo, 2009). It is well known that phosphorylation of c-Src activated by VEGFR2 induces phosphorylation of VE-cadherin, resulting in internalization of VE-cadherin into the cytoplasm (Lampugnani et al., 2006). LPA did not affect phosphorylation of VE-cadherin in MS-1 cells (Figure S5D). Angiopoietin-1, a ligand for receptor tyrosine kinase Tie2, induces membrane localization of VE-cadherin through the PI3K-Akt pathway, and it has been determined that Tie2 phosphorylation-driven activation of RhoA leads to an association of Src

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Figure 7. Analysis of Cell Signaling through LPA4 for Barrier Function in ECs (A–C) Monolayers of MS-1 cells treated with PTX for Gi inhibition with or without VPC31144(S). MS-1 cells were pretreated with normal conditioned medium supplemented with 100 ng/mL of PTX for 12 hr, followed by serum starvation with PTX for 4 hr. Next, 10 mM VPC31144(S) or vehicle was added to the medium and incubated for 1 hr. (A) The cells were stained with anti-VE-cadherin mAb (green) and phalloidin (red). The nuclei were labeled with TOPRO3 (blue). The dashed white boxes in the upper panels are magnified in each of the three lower panels. Scale bar, 50 mm (upper panels) and 10 mm (lower three panels). (B and C) Quantification of VE-cadherin (B) and F-actin (C) at cell-cell junctions compared to control (n = 16 points per group). (D–F) Monolayers of MS-1 cells treated with siRNAs for negative control (siCTR) or GNA13 (siGNA13), serum-starved, and stimulated with 10 mM VPC31144(S) or vehicle for 1 hr. (D) The cells were stained with anti-VE-cadherin mAb (green) and phalloidin (red). The nuclei were labeled with TOPRO3 (blue). The dashed white boxes in the upper panels are magnified in each of the three lower panels. Scale bar, 50 mm (upper panels) and 10 mm (lower three panels). (E and F) Quantification of VE-cadherin (E) and F-actin (F) at cell-cell junctions compared to control (n = 16 points per group). The error bars indicate mean ± SEM. **p < 0.01 and ***p < 0.001. See also Figure S7.

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with mDia, resulting in sequestration of Src from VEGFR2 and inhibition of VE-cadherin internalization (Gavard et al., 2008). It was reported that Ga12/13 is involved in cortical actin formation of HUVECs via RhoA/ROCK (Yukiura et al., 2015). Therefore, LPA4 activation might be involved in these systems. Further analysis using these inhibitors is required to elucidate the signaling pathway of LPA4 for EC tightening. Previously, signal modification of VEGFR2 by VE-cadherin has been reported; i.e., when VE-cadherin is absent, VEGFR2 phosphorylation is strongly induced, resulting in EC proliferation in an uncontrolled way (Lampugnani et al., 2003). Here, we show that LPA4 activation results in the assembly of circumferential actin bundles beneath the cell membrane, which promotes linear adherence junction formation by VE-cadherin. In the absence of VE-cadherin, EC proliferation stimulated by VEGF is suggested to lead to apoptosis. Hence, uncontrolled VEGFR2 signaling is suppressed by VE-cadherin expression enhanced by LPA and stable and functional blood vessels are induced. Recently, several lines of evidence also indicated the benefits of enhancing maturation of the tumor vasculature by using nonangiogenesis inhibitors, for example, the maturation of blood vessels by increasing the stability of the notch pathway using chloroquine (Maes et al., 2014), normalization of vascular permeability using small amounts of RGD peptide, a synthetic fragment of the integrin binding motif (Wong et al., 2015), or Ang2-binding and Tie2-activating antibody (Park et al., 2016). These agents clearly have a different mode of action from LPA. It is widely accepted that there is a great deal of heterogeneity among cancer cells even in a single lesion, and LPA did not show enhancing effects with anti-cancer drugs when we started treatment of bigger tumors (data not shown). This implies that different angiogenesis disorders may occur simultaneously in the same tumor. Hence, to normalize and mature the blood vessels throughout the whole tumor, a combination of several agents with different mechanisms of action would be desirable. LPA4 agonists are one of these candidates that could be utilized clinically together with other vascular normalization drugs. EXPERIMENTAL PROCEDURES Reagents 1-oleoyl-LPA and VPC31144(S) were purchased from Avanti Polar Lipids (Alabaster, AL) and were dissolved in 50% ethanol at 10 mM, and stocked at
20 C. PTX was purchased from List Biological Laboratories (Campbell, CA, USA). SQ22536 (Wako, Osaka, Japan) was dissolved in DMSO at 50 mM and stored at
20 C. 20 ,50 -dideoxyadenosine (20 50 DDA; Wako) was dissolved in water at 100 mM and stored at
20 C. YM-254890 (Wako) was dissolved in DMSO at 10 mM and stored at 4 C. Mice All experiments were performed in accordance with the guidelines of the Osaka University Committee for Animal and Recombinant DNA Experiments. C57BL/6 and BALB/c mice were purchased from SLC (Shizuoka, Japan). LPA4
/
mice (Sumida et al., 2010) and LPA6
/
mice (Hata et al., 2016) on the C57BL/6 background were produced as described previously. Mice 8 to 12 weeks of age were used for these experiments. Cell Lines LLC (RIKEN Cell Bank, Tsukuba, Japan) was maintained in DMEM (Sigma) supplemented with 10% fetal calf serum (FCS) (Sigma) and 1% penicillin/ streptomycin (P/S; Life Technologies, Tokyo, Japan). Colon26 (mouse colon

carcinoma; RIKEN), B16BL/6 (mouse melanoma, RIKEN), and Ex-3LL (RIKEN) were maintained in RPMI-1640 (Sigma) supplemented with 10% FCS and 1% P/S. MS-1 (mouse pancreatic islet EC line; ATCC) was maintained in DMEM supplemented with 5% FCS and 1% P/S. OP9 (mouse fibroblast; RIKEN) was maintained in MEM (Sigma) supplemented with 10% FCS and 1% P/S. LLC, Ex-3LL, and B16BL/6 cells were stably transfected with the expression vector pEGFP N-1 (Clontech, Palo Alto, CA) using Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA). The transfected cells were cultured in selection medium containing 200 mg/mL G418 (Geneticin; Gibco, Grand Island, NY). Tumors Subcutaneous xenograft models of LLC and B16BL/6 were established by injecting 1.0 3 106 tumor cells into the flanks of C57BL/6 mice, and colon26 was injected into BALB/c mice. Tumor size was then measured with calipers. Tumor volume (V) was calculated according to the formula V = 1/2 3 length 3 width 3 height. For establishment of a model of metastatic lung tumor spreading from subcutaneous Ex-3LL and B16BL/6 tumors, 1 3 10 6 tumor cells were inoculated into C57BL/6 mice subcutaneously. Mice were treated with vehicle or LPA (3 mg/kg, i.p.) daily from day 7 to day 28. On day 21, subcutaneous tumors were resected, and lungs were harvested on day 42. For establishment of the metastatic lung tumor model of B16BL/6 spreading from intravenous injection, mice were pretreated with vehicle or LPA (3 mg/kg, i.p.) for 7 days, followed by injection of 1 3 105 B16BL6-EGFP cells in 200 mL of PBS into the tail vein. Daily administration of vehicle or LPA continued until 14 days post inoculation. Lungs were dissected on day 21. For evaluation of therapeutic effects of LPA and/or 5-FU (Kyowa Hakko Kirin, Tokyo, Japan) on the tumor, LPA (3 mg/kg, i.p.) was administered daily from day 6 to day 20 and/or 5-FU (100 mg/kg) was given i.p. on day 7 and on day 14. Immunohistochemistry and Immunocytochemistry The procedure for tissue preparation and staining was as previously reported (Naito et al., 2012). Briefly, tumors were fixed with 4% paraformaldehyde (PFA) in PBS overnight and were embedded in OCT compound (Sakura Finetek, Tokyo, Japan) and sectioned at 10, 20, 40, or 120 mm. Primary antibodies were rat anti-CD31 monoclonal antibody (mAb) (BD Biosciences, San Diego, CA), hamster anti-CD31 mAb (Millipore, Darmstadt, Germany), rabbit antiGFP polyclonal antibody (pAb) (MBL International, Nagoya, Japan), mouse anti-a-SMA-Cy3 mAb (Sigma), rabbit anti-NG2 pAb (Millipore), rabbit antiERG pAb (Cell Signaling Technology Inc., Danvers, MA), mouse anti-Ki67 mAb (Dako, Santa Clara, CA), rabbit anti-Lyve-1 pAb (Reliatech GmbH, Wolf€ttel, Germany), rabbit anti-cleaved caspase-3 mAb (Cell Signaling Techenbu nology Inc.), and rat anti-VE-cadherin mAb (BD Bioscience), used to stain tumor tissue sections. For immunofluorescence, Alexa Fluor 488-conjugated anti-rat immunoglobulin G (IgG) (Invitrogen), Alexa Fluor 546-conjugated anti-rat IgG (Invitrogen), Alexa Fluor 488-conjugated anti-rabbit IgG (Invitrogen), Alexa Fluor 647-conjugated anti-rabbit IgG (Invitrogen), and Alexa Fluor 488-conjugated anti-hamster IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) or Cy3-conjugated anti-hamster IgG (Jackson ImmunoResearch) were used as the secondary antibodies. For immunohistochemistry, biotinylated anti-rat IgG (Vector Laboratories, Burlingame, CA) was used as the secondary antibody followed by VECTASTAIN ABC (Vector Labs) and developed by 3,3’-diaminobenzidine (DAB) staining (Dako). To obtain a specific antibody against mouse LPA4, a rabbit was immunized with a synthetic peptide (CEVSDQTTNNGGELMLESTF) derived from the C-terminal region of LPA4 (Sigma). TUNEL staining was performed as described in the manufacturer’s protocol (Takara Bio Inc., Shiga, Japan). For quantification of vascular length, pericyte coverage, Ki67 index, cleaved caspase-3, TUNEL, and H&E staining, sections were dissected at 10 mm thickness and more than three images were taken for each section. For evaluation of vascular length and pericyte coverage, images were taken randomly from tumor center, which was defined as 500 mm inside from the tumor boundary. Quantitative measurement of vessel length was accomplished using a semi-automated computational tool (AngioTool; available in the public domain at https:// ccrod.cancer.gov/confluence/display/ROB2/Home) (Zudaire et al., 2011). MS-1 cells were grown to confluence, serum starved, and stimulated with VEGF (100 ng/mL) with or without LPA (0.1–10 mM). The cells were then fixed

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with 4% PFA and permeabilized by 0.1% Triton X-100. Cells were labeled with anti-VE-cadherin antibody (BD Biosciences) and visualized with Alexa Fluor 488-conjugated goat anti-rat IgG. Filamentous actin (F-actin) was stained with tetramethylrhodamine isothiocyanate-conjugated phalloidin (Sigma). Nuclear staining was performed using Hoechst 33342 (Sigma) or TOPRO3 (Invitrogen). F-actin and VE-cadherin at the cell-cell contact point were measured as previously described (Ando et al., 2013). Briefly, the fluorescence intensity at the cell-cell contact points was measured using the line scan function in MetaMorph (Universal Imaging, Philadelphia, USA). The 3-pixel-width lines were randomly drawn and the mean pixel intensity for each position along the lines was determined by the line scan analysis. The mean fluorescence intensity at the points across the cell-cell contacts was scored as the amount of F-actin and VE-cadherin at the cell-cell contact points. Scanning Electron Microscopy Tissues were fixed with 2% formaldehyde and 2.5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4), post-fixed with 1% osmium tetroxide and 0.5% potassium ferrocyanide in the same buffer, dehydrated in graded series of ethanol, substituted with t-butyl alcohol, and freeze dried. After freeze drying, samples were coated with osmium tetroxide and observed with a S-4800 field emission scanning electron microscope (Hitachi High-Technologies Corp., Japan). Assessment of Blood Vessel Function To assess blood perfusion, fluorescein Lycopersicon esculentum (tomato) lectin (0.05 mg/mouse; Vector Labs) was injected intravenously into the tail vein of tumor-bearing mice. 10 min after lectin injection, tumors were dissected. Blood flow was interpreted by the lectin positivity in ECs using 10 mm tissue sections. To evaluate drug delivery, fluorescein isothiocyanate (FITC)-conjugated dextran (MW 70,000; 0.5 mg/body; Sigma) was injected into the tail vein. 1 hr after injection, intracardiac perfusion with PBS was performed and tumors were harvested. To evaluate anti-cancer drug delivery, doxorubicin hydrochloride (1.5 mg/kg; Nippon Kayaku, Tokyo, Japan) was injected intravenously into the tail vein and allowed to circulate for 20 min. Tumors were collected after intracardiac perfusion with PBS. For evaluation of dextran permeability and doxorubicin delivery, more than three images were taken from tumor center. To measure hypoxia in tumor tissues, Hypoxyprobe-1 (60 mg/kg, i.p.; Hypoxyprobe, Burlington, MA, USA) was injected 2 hr before tissues were harvested. Tumor sections were stained using the anti-Hypoxyprobe antibody, following the manufacturer’s instructions. Samples were visualized using a Leica TCS SP5 confocal microscope (Leica Microsystems, Nussloch, Germany) or Leica DM5500B and processed with the Leica Application Suite and Adobe Photoshop CS6 software (Adobe Systems, San Jose, CA, USA). All images shown are representative of more than five independent experiments. Cell Preparation Tissue dissection procedures and preparation of single cell suspensions were as previously reported (Naito et al., 2016). For cancer cells and EC preparations, LLC-GFP tumors were dissected and stained with phycoerythrin (PE)conjugated anti-CD45 mAb and allophycocyanin (APC)-conjugated antiCD31 mAb (BD Biosciences). For tumor stromal cell preparation, LLC tumors were dissected and stained with APC-conjugated anti-CD31 mAb (BD Pharmingen), FITC-conjugated anti-CD45 mAb (BD Pharmingen), and biotin-conjugated anti-PDGFRb mAb (Thermo Fisher Scientific K.K., Yokohama, Japan). Streptavidin-PE (BD Pharmingen) was used as the secondary antibody. GFPpositive cancer cells, CD31+CD45
ECs, and CD31
CD45
PDGFRb+stromal cells were analyzed and sorted using a SORP FACSAria (BD Biosciences). MACS Cell Separation and Primary EC Culture on OP9 Monolayers Single cell suspensions of normal lung were labeled with APC-conjugated anti-mouse CD31 mAb (BD Pharmingen) and anti-APC MicroBeads (Miltenyi Biotec, Bergisch Gladbach, Germany) and isolated using an AutoMACS (Miltenyi Biotec) cell separator. To confirm the purity of ECs, isolated cells were stained with FITC-conjugated anti-CD45 mAb (BD Pharmingen), and the cell population was analyzed by fluorescence-activated cell sorting (FACS). We

2084 Cell Reports 20, 2072–2086, August 29, 2017

confirmed that >90% of the isolated cells were CD31+CD45
ECs. ECs were seeded at 3.0 3 105 per well in 24-well plates prepared with confluent OP9 cells. Cells were cultured in minimum essential medium (MEM; Sigma) supplemented with 10% FCS, 1% P/S, and VEGF (20 ng/mL). On day 7, culture medium was changed to new medium containing 10 mM of VPC31144(S) or vehicle. On day 10, culture medium was removed and cells were fixed with 4% PFA in PBS. Biotinylated anti-mouse CD31 mAb (BD Pharmingen) was applied as the primary antibody followed by VECTASTAIN ABC (Vector Labs) and DAB staining (Dako). Real-Time PCR Analysis RNA was extracted using RNeasy Mini Kits (QIAGEN, Hilden, Germany), and cDNA was generated using reverse transcriptase with the ExScript RT reagent Kit (Takara, Otsu, Japan). Real-time PCR was performed using a Stratagene Mx3000P (Stratagene, La Jolla, CA, USA). The PCR was performed on cDNA using specific primers with the following sequences: 50 -TGG CAA AGT GGA GAT TGT TGC C-30 and 50 -AAG ATG GTG ATG GGC TTC CCG-30 for GAPDH, 50 -CCG CTT CCA TTT CCC TAT TT-30 and 50 -AAA ACC GTG ATG TGC CTC TC-30 for LPA1, 50 -CCA TCA AAG GCT GGT TCC T-30 and 50 -TCC AAG TCA CAG AGG CAG TG-30 for LPA2, 50 - TTC CAC TTT CCC TTC TAC TAC CTG-30 and 50 -TCC ACA GCA ATA ACC AGC AA-30 for LPA3, 50 -GCC CTC TCT GAT TTG CTT TT-30 and 50 -TCC TCC TGG TCC TGA TGGTA-30 for LPA4, 50 -AGC GAT GAA CTG TGG AAG G-30 and 50 -GCA GGA AGA TGA TGA GAT TGG-30 for LPA5, 50 -TGT GCC CTA CAA CAT CAA CC-30 and 50 -TCA CTT CTT CTA ACC GAC CAG-30 for LPA6, and 50 -AAG TCC ACC TTC CTG AAG CA-30 and 50 -CTT CTC TCG GGC ATC TAC CA-30 for GNA13. Expression level of the target gene was normalized to the GAPDH level in each sample. Immunoprecipitation and Western Blotting Confluent MS-1 cells were cultured in 6-well plates and serum-starved in DMEM supplemented with 0.5% BSA, after which they were stimulated with 50 ng/mL of VEGF and/or 10 mM of LPA for 30 min. After washing with icecold PBS containing 500 mM of sodium orthovanadate, cells were extracted in TNT buffer (10 mM Tris [pH 7.4], 150 mM NaCl, 1% NP40, 1% Triton, and 2 mM EDTA) containing Halt Protease and Phosphatase Inhibitor Cocktail (Thermo Scientific). After centrifugation, the supernatants were incubated overnight at 4 C with goat polyclonal anti-VE-cadherin (Santa Cruz Biotechnology, C-19, sc-6458), and subsequently incubated with Protein G Sepharose 4 Fast Flow (GE Healthcare) for 4 hr. Sepharose beads were washed 34 with TNT buffer and eluted by boiling in SDS sample buffer. Analysis by SDS-PAGE was as previously described (Kidoya et al., 2008). Impedance Measurement by ECIS Barrier function of EC junctions was evaluated using ECIS-zQ (Applied Biophysics, Jordan Road, Troy, NY), as described previously (Ando et al., 2013). In brief, we used 8W10E electrodes with a thin gold film surface and coated each well containing gold film electrodes with collagen so that the MS-1 cells could adhere to them. 4 3 104 MS-1 cells were plated on electrodes and cultured overnight. Each well has a substrate area of 0.8 cm2 and electrode area of 1.96 mm2. The cells were then starved in 0.1% BSA-containing DMEM for 4 hr and subjected to ECIS. Resistance between cells (Rb) was measured in real time at 37 C with 5% CO2 using an ECIS-zQ system at 4,000; 16,000; and 64,000 Hz. MS-1 cells were pretreated for 24 hr with 100 ng/mL PTX to inhibit Gi. siRNA Knockdown siRNAs for LPA4, LPA6, GNA13, and negative control siRNA were purchased from Thermo Fisher and were transfected into MS-1 cells using Lipofectamine RNAiMAX (Thermo Fisher Scientific), according to the manufacturer’s instructions. Knockdown efficiency was verified using quantitative real-time PCR as described above. Statistical Analysis All data are presented as mean ± SEM. Comparisons between multiple treatments were made using one-way ANOVA, followed by the Mann-Whitney

U test. Pairwise comparisons between treatments were made using Student’s t test. A p value < 0.05 was considered to be significant.

Ebos, J.M., Lee, C.R., Cruz-Munoz, W., Bjarnason, G.A., Christensen, J.G., and Kerbel, R.S. (2009). Accelerated metastasis after short-term treatment with a potent inhibitor of tumor angiogenesis. Cancer Cell 15, 232–239.

SUPPLEMENTAL INFORMATION

Ferrara, N., Hillan, K.J., Gerber, H.P., and Novotny, W. (2004). Discovery and development of bevacizumab, an anti-VEGF antibody for treating cancer. Nat. Rev. Drug Discov. 3, 391–400.

Supplemental Information includes seven figures and can be found with this article online at http://dx.doi.org/10.1016/j.celrep.2017.07.080. AUTHOR CONTRIBUTIONS Conceptualization, K.T., D.E., and N.T.; Methodology, K.T., D.E., and K.A.; Investigation, K.T., D.E., K.A., D.Y., H.N., Y.T., T.I., T.W., F.M., and H.K.; Resources, S.F., N.M., and S.I.; Writing - Original Draft, K.T., D.E., and N.T.; Supervision, S.F., N.M., S.I., H.K., and N.T.; Funding Acquisition, D.E. and N.T. ACKNOWLEDGMENTS We thank K. Fukuhara, N. Fujimoto, M. Ishida, and Y. Mori for technical assistance. This work was partly supported by the Japan Agency for Medical Research and Development (AMED) Projects for Technological Development, Research Center Network for Realization of Regenerative Medicine and for Development of Innovative Research on Cancer Therapeutics, Japan Society for the Promotion of Science (JSPS) Grants-in-Aid for Scientific Research (A) (15H02545), Grant-in-Aid for Young Scientists (B) (15K19968), Project MEET, Osaka University Graduate School of Medicine, and a grant from Mitsubishi Tanabe Pharma Corporation.

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Received: January 17, 2017 Revised: June 15, 2017 Accepted: July 28, 2017 Published: August 29, 2017

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