Ki16425

LLysophosphatidic acid acts on LPA1 receptor to increase H2O2 during flow-induced dilation in human adipose arteriolesysophosphatidic acid acts on LPA1 receptor to increase H2O2 during flow-induced dilation in human adipose arterioles

Abstract
Nitric oxide (NO) produces arteriolar flow-induced dilation (FID) in healthy subjects but is replaced by mitochondria-derived hydrogen peroxide (mtH2O2) in patients with coronary artery disease (CAD). Lysophosphatidic acid (LPA) is elevated in patients with risk factors for CAD but its functional effect in arterioles is unknown. We tested whether elevated LPA changes the mediator of FID from NO to mtH2O2 in human visceral and subcutaneous adipose arterioles. Arterioles were cannulated on glass micropipettes and pressurized to 60 mmHg. We recorded lumen diameter after graded increases in flow in the presence of either nitric oxide synthase (NOS) inhibition (L-NAME) or H2O2 scavenging (Peg-Cat) ± LPA (10 µM, 30 min), ± LPA1/LPA3 receptor antagonist (Ki16425) or LPA2 receptor antagonist (H2L5186303). We analyzed LPA receptor RNA and protein levels in human arterioles and cultured human endothelial cells. FID was inhibited by L-NAME but not Peg-Cat in untreated vessels. In vessels treated with LPA, FID was of similar magnitude but inhibited by Peg-Cat while L-NAME had no effect. Rotenone attenuated FID in vessels treated with LPA indicating mitochondria as a source of ROS. RNA transcripts from LPA1 and LPA2 but not LPA3 receptor were detected in arterioles. LPA1 but not LPA3 receptor protein was detected by Western blot. Pretreatment of vessels with an LPA1/LPA3, but not LPA2, receptor antagonist prior to LPA preserved NO-mediated dilation. These findings suggest an LPA1 receptor-dependent pathway by which LPA increases arteriolar release of mtH2O2 as a mediator of FMD.

Introduction
Endothelial cells form an interface between circulating blood and surrounding tissue and actively respond to changes in blood flow. The endothelium-dependent, flow-stimulated release of nitric oxide (NO) and other vasodilator factors that act on adjacent vascular smooth muscle cells (VSMC) is a key means by which human resistance arterioles dilate and increase tissue perfusion (flow induced dilation; FID). Endothelial release of NO promotes vascular health by reducing inflammation and proliferation (Kim et al., 2008). Previous studies demonstrated that the magnitude of FID is preserved in arterioles from patients with coronary artery disease (CAD) or multiple risk factors for CAD, but the mediator switches from NO to pro-inflammatory and atherogenic mitochondrial hydrogen peroxide (mtH2O2) (Beyer et al., 2017; Liu et al., 2003), providing a potential pathway underlying the microvasc ular contributions to CAD. We have previously reported intracellular components involved in the transition from NO to H2O2, including PGC-1α (Kadlec et al., 2017) and telomerase (Beyer et al., 2016). The upstream cell-surface activators of this transition, however, remain incompletely characterized. Understanding what factors and receptors act at the endothelial cell surface of arterioles is essential for constructing a complete signaling pathway and identifying novel targets to reduce H2O2 release in the context of CAD.

Lysophosphatidic acid (LPA), a simple phospholipid, is an autacoid found at the cell surface and in the circulation. LPA triggers a plethora of cellular responses ranging from enhanced proliferation, migration and invasiveness to morphological changes, increased permeability or apoptotic cell death in different cells of the vasculature (Lin et al., 2006; Panchatcharam et al., 2008; Siess et al., 1999; van Nieuw Amerongen et al., 2000). Physiological concentrations range from nano- to micro-molar levels (Aoki, 2004) and are controlled by a dynamic balance of synthesis (mainly by secreted autotaxin (ATX) (Albers et al., 2010; Gierse et al., 2010) and degradation (primarily by membrane-bound ecto-activity of lipid phosphate phosphatase 3 (LPP3)) (Panchatcharam et al., 2014; Tanyi et al., 2003). Disruption of these control mechanisms can increase LPA levels by more than 10-fold, as reported in individuals with cancer (Baker et al., 2002), neurological disorders, or risk factors for cardiovascular disease, such as hyperlipidemia (Dohi et al., 2012) or hypertension (Xu et al., 2003). A rise in LPA has also been implicated in the development and progression of atherosclerosis and cardiovascular disease (Bot et al., 2013; Busnelli et al., 2017; Kurano et al., 2015).

These effects are mediated by cognate LPA receptors. Currently, there are six LPA receptors that belong to the G-protein coupled receptor family. Each receptor couples to at least one or more heterotrimeric Gα proteins, such as G12/13, Gq/11, Gi/o, and Gs and can activate a complex array of downstream signaling pathways with physiological and pathological effects on the vasculature. In particular, signaling through LPA1 receptor is involved in many LPA-induced vascular effects, such as increased inflammation or endothelial cell migration (Panetti et al., 2004; Panetti et al., 2000). Additionally, inhibition of LPA1 receptor but not others reduces inflammation in cultured endothelial cells (Rizza et al., 1999). Independent studies have shown that increased LPA, up to 10 µM, decreases endothelial nitric oxide synthase (eNOS) expression, reduces NO bioavailability, and increases reactive oxygen species (ROS) production in cultured cells (Chen et al., 2012; Chen et al., 1995; Shin et al., 1999), all hallmarks of endothelial dysfunctio n. However, LPA’s functional influence on intact human arterioles, and the receptor responsible for these effects, has not been explored. Examining the arteriolar effects of elevated LPA is clinically relevant given the prognostic importance of arteriolar dysfunction (van de Hoef et al., 2014). We hypothesized that exposing arterioles from patients without CAD (non-CAD) to elevated levels of LPA (10 µM) would shift the mediator of FID away from NO to mtH2O2, recapitulating the phenotype observed in arterioles from patients with CAD. Given that LPA signals through cognate LPA receptors, we further examined which receptors are expressed in human arterioles and mediate LPA’s effects. We discovered that elevated levels of LPA in vessels from subjects without CAD increases mtH2O2 during FID in a LPA1 receptor- dependent manner.

Protocols for tissue acquisition and processing were approved by the Institutional Review Board of the Medical College of Wisconsin. Human adipose (subcutaneous and visceral) was obtained as discarded tissue at the time of surgery and immediately placed in 4˚C HEPES buffer [(in mM) NaCl 275, KCL 7.99, MgSO4 4.9, CaCl2 ·2H2O 3.2, KH2PO4 2.35, EDTA 0.07,glucose 11, HEPES acid 20). Patient demographics are collected and stored in a de-identifiedfashion using the Generic Clinical Research Database at the Medical College of Wisconsin. Tissue from subjects without clinically diagnosed CAD and no more than one risk factor (Table 1) were categorized as the non-CAD group. Vessels defined as CAD were obtained from tissue from subjects with clinically diagnosed CAD.Human arterioles were isolated and excess adipose and connective tissue was removed. The vessels were cannulated onto glass micropipettes of matched impedance (diameters of both pipettes per single chamber preparation did not vary more than 2 µm, and was typically kept between 35 µm and 45 µm, depending on the size of an arteriole) filled with Krebs buffer [(in mM) NaCl 118, KCl 4.7, CaCl2 2.5, KH2PO4 1.2, MgSO4 1.2, NaHCO3 20, Na2 EDTA0.026, and dextrose 11, pH 7.4] and secured using suture in an organ chamber also filled with Krebs buffer solution. The arterioles were kept at 37˚C and were pressurized to 60 mmHg in a step-wise manner (30 min at 30 mmHg followed by 30 min at 60 mmHg) using dual-reservo ir system, where each micropipette is connected to its own reservoir.

Once equilibration time was reached, the vessels were constricted with endothelin-1 (ET-1) to 30-60% of the passive diameter. With stable constriction lasting ≥10 min, the vessels were exposed to increasing physiological rates of flow (pressure gradient; 5 to 100 cm H2O) with constant intralumi nal pressure, as described previously (Kuo et al., 1991; Kuo et al., 1990; Miura et al., 2003).Briefly, in this preparation, flow is initiated by simultaneously moving of the reservoirs in equal and opposite direction (one down and one up by equal distance from the starting point) to generate a pressure gradient with shear rates estimated at 5 to 25 dynes/cm2, while preserving intraluminal pressure (Kuo et al., 1990). In some vessels, two studies were performed in sequence separated by a wash (20 mL of Krebs buffer replaced five times) and a 30-minute re- equilibration period prior to the second curve. At the end of each experiment, papaverine (100 µM) was used to assess maximal dilation. At the end of each experiment matched impedance of the pipettes was verified by applying max flow (100 cm H2O) in reverse direction for 5 min and in the presence of papaverine. If vessel diameters were different between flows of opposite direction, vessels were excluded from analysis. Additionally, vessels that did not reach or retain stable constriction for at least 10 min were excluded from the data set, as were arterioles that did not reach 75% of max dilation to papaverine. All treatments were added to the organ bath (Krebs buffer) and constituted ≤1% of the total circulating volume.

Vessels were treated with LPA [Abcam, ab146430; dissolved in Phosphate Buffer Saline (PBS), pH 7.4, final concentration in the organ bath 10 µM] for ≥30 min (depending on experiment performed) prior to constriction. To determine whether the mediator of FID was NO or H2O2, we used the nitric oxide synthase (NOS) inhibitor [Nω-nitro-L- arginine methyl ester (L-NAME), 100 µM] or a scavenger of H2O2 [polyethylene glycol- catalase (Peg-Cat) 500 U/mL], respectively, added to the organ chamber for 30 min prior to constriction. Prior studies have shown these doses to effectively block NO and H2O2 components of FID, respectively (Beyer et al., 2014; Freed et al., 2014). To further localize the subcellular source of H2O2, we performed FID studies under LPA challenge in the presence of rotenone (mitochondria complex I inhibitor; 1 µM). In separate studies, we assess endothelial viability dose-response studies to Ach (acetylcholine; 1 nM to 100 µM) with and without LPA were performed. To assess endothelium- independent, vascular smooth muscle cell (VSMC) reactivity, dose-response studies to SNP (sodium nitroprusside; 1 nM to 100 µM) were performed in the presence and absence of LPA. To determine which LPA receptor is responsible, vessels were pretreated with either Ki16425 (Cayman Chem, 10012659, LPA1/3 receptor antagonist; 10 µM) or H2L5186303 (Cayman Chem, 14663, LPA2 receptor antagonist; 1 µM or 10 µM) 30 min prior to LPA administration. Concentrations of Ki16425and H2L5186303 were chosen based on data published by other groups working with these antagonist in the context of vascular biology (Ruisanchez et al., 2014; Wu et al., 2015).Human umbilical vein endothelial cells (HUVECs) were purchased from ATCC and cultured on 100 mm culture plates in endothelial cell growth medium (EGM®-2; Lonza) suppleme nted with growth factors, cytokines and 5% fetal calf serum. Human cardiac microvasc ular endothelial cells (HMVECs) lysates were a gift from Dr. Julie Freed and prepared as previously described (Freed et al., 2017).

Briefly, cells were purchased from Lonza and cultured in 100 mm culture plates in cell growth medium (EBM-2; Lonza) supplemented with growth factors, cytokines and 10% fetal calf serum. Cells at passage 3-5 with 80-95% confluency were used.Vessels were isolated and connective tissue was carefully removed. Vessels were snap frozen in liquid nitrogen and stored in -80˚C until further processing. Cultured endothelial cells (human umbilical vein endothelial cells (HUVECs) were scraped from culture dishes and centrifuged to form a cell pellet, which was then snap frozen until further processing. 3-5 arterioles from each tissue were frozen and constituted one biological replicate. 4 culture dishes with HUVECs and 3 separate cell lysate preparations of HMVECs were used. For lysate preparations, vessels were thawed and homogenized using a glass homogenizer in cold lysis buffer (50 mM Tris, pH 7.4, 150 mM NaCl, 1% deoxycholic acid, 0.1% SDS, 0.5% NP40) supplemented with protease and phosphatase inhibitor cocktail (Roche) and briefly sonicated. Cell pellets were thawed and briefly sonicated. Total protein concentration was determined using BCA protein assay (Thermo Scientific). Protein samples of known concentration from each tissue (patient) were subjected to 4-20% SDS-PAGE and transferred to nitrocellulose membranes. To detect target receptor protein, antibodies to LPA1 (Abcam, ab166903; 1:1000), LPA3 (LSBio, LS-A1014; 1:1000), LPA3 (Abcam [Abc], ab23692, 1:1000) and GAPDH(Abcam, ab8245, 1:10,000) were used, followed by peroxidase conjugated secondary antibodies and chemiluminescent substrate (Bio-Rad).

Quantitative real-time polymerase chain reactionIsolated and cleaned human adipose vessels were snap frozen in liquid nitrogen and stored in – 80˚C until further processing. Total RNA was extracted from samples using RNAqueousTM- Micro Total RNA Isolation Kit (Ambion, AM1931). 25 ng of RNA was used to synthesize cDNA using High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, 4368814).Quantification of target gene expression was performs using Qiagen QuantiTect Primer Assay for LPA1 (Hs_LPAR1_1_SG, QT00021469), LPA2 (Hs_LPAR2_2_SG, QT01851318), LPA3 (Hs_LPAR3_1_SG, QT00092932), PECAM (Hs_PECAM1_1_SG, QT0008172) andnormalized to 18s (Hs_RRN18S_1_SG, QT00199367). Gene expression analysis was performed using BioRad CFX96 Touch Read-Time PCR Detection System. Each sample was evaluated as an average of a triplicate.Vessels were isolated from tissue and immediately placed in zinc formalin buffer followed by paraffin embedding. Immunohistochemistry was performed by the Children’s Hospital of Wisconsin Histology Core. Sections were cut at 4 µm and placed on poly-L-Lysine coated slides, which were then dried at 45˚C overnight and stored in room temperature prior to IHC testing. Slides were then loaded onto the Leica Bond Max immunostainer. All samples were de-waxed prior to staining on the instrument. Antibodies required antigen retrieval pretreatment as follows: LPA1 receptor H1(20) and LPA3 receptor H1(20) citrate buffer at pH 6 and CD31 H2(10) EDTA retrieval. Antibodies used: LPA1 receptor (Abcam, ab166903, 1:100), LPA3 receptor (LS-Bio, LS-A1014/40949, 1:200) and CD31 (Agilent, DAKO, M0823,1:100). All antibodies were detected and visualized using Leica Bond Polymer Refine Detection System (DS9800) with the addition of a DAB enhancer (AR9432), using the modified versions of the MODF protocol installed by Leica filed service engineers. Slides were removed from immunostaining platform, dehydrated, cleared and coverslipped with synthetic mounting media. Omission of the primary antibody served as negative control.

Slides were scanned with a NanoZoomer HT slide scanner (Hamamatsu, Japan).Fluorescence detection of mitochondrial H2O2 in human microvesselsTo evaluate mitochondrial H2O2 release in human arterioles challenged with LPA (10 µmM, 30 min), Mito Peroxy Yellow 1 (MitoPY1) was used (Dickinson & Chang, 2008). As previously described, vessels were cannulated in a chamber containing HEPES buffer warmed to 37˚C and pH 7.4 (Kadlec et al., 2017). The same buffer containing the fluorescent MitoPY1 probe (10 µM, 1 hour) was used to perfuse vessels intraluminally. Fluorescence was imaged using a krypton/argon lamp fluorescent microscope (model TE 200 Nikon Eclipse) or X-cite FIRE LED microscope (model Olympus IX73) and an excitation/emission wavelength of 488 nm/530-590 nm before administration of LPA, 30 minutes after, and every minute for 5 min once max flow (Pressure Gradient of 100 cmH2O) was applied. Relative fluorescence intens ity (arbitrary units) was measured using ImageJ (NIH) by subtracting background fluoresce ncefrom vessels fluorescence. Percent increase in fluorescence intensity from static values after 5 min of max flow (% increase to flow) was compared between intact control and other treatments vessels. Endothelial contribution to the fluorescence signal was assessed by denuding a subset of vessels. Briefly, vessels were isolated, cleaned and one end was cannulated onto a glass micropipette. Vessels were then equilibrated for 30 min at 37˚C. Once equilibrated, 4 mL of air was passed through the vessel to dislodge the endothelium. After denudation, the loose end of the vessel was cannulated and experimental protocol for fluorescence detection of mitochondrial H2O2 was continued.

Viability of the arterioles was assessed by the following protocol: arterioles were allowed to equilibrate without flow for 5- 10 min, after equilibration 0.3 nM of ET-1 was used to constrict the vessels, the vessels needed to retain constriction for 10 min after which 100 µM of acetylcholine was added followed by 100 µM of papaverine to assess vessels ability to dilate. Vessels that did not constrict to ET-1 or did not dilate to papaverine were excluded from the study. After completion of the protocol, arterioles were removed from the glass cannulas and snap frozen in liquid nitrogen and stored in -80˚C until further processing. Denudation efficiency was assessed by measuring expression of endothelial marker, PECAM, using RT-qPCR.All data are expressed as mean±SEM with statistical significance threshold set at p<0.05. For vascular studies, data are expressed as a percent of maximal dilation from ET-1-induced constriction, where 100% represents full relaxation to the maximal diameter noted throughout the experiment. To compare flow-response relationship, a 2-way ANOVA was used with pressure gradient (or dose-response to SNP or ACH) and treatment as parameters. When a significant difference was observed between curves (P<0.05), responses at individual pressure gradients were compared using a Holm-Sidak multiple comparison test. To compare concentration of SNP required to cause 50% of the maximum response (EC50), a three- parameters nonlinear regression using least squares (ordinary) fit analysis was performed. 1- way ANOVA was used to compare fluorescence (MitoPY1 probe) in microvessels before and after interventions. Wilcoxon test was used to compare PECAM expression in control vs denuded arterioles. All analyses were performed using GraphPad Prism 7.04 (GraphPad Software, San Diego, CA) and SAS 9.4 (SAS, Cary, NC) with statistical significance defined at P<0.05.Key protein targets and ligands in this article are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, the common portal for data from the IUPHAR/BPS Guide to PHARMACOLOGY (Harding et al., 2018), and are permanently archived in the Concise Guide to PHARMACOLOGY 2017/18 (Alexander et al., 2017). Results Discarded adipose tissue (subcutaneous and visceral) was collected from a total of 111 patients, 6 with CAD and 105 categorized as non-CAD. Patient demographics are detailed in Table 1. Mean diameter prior to constriction, after constriction, and concentration of ET-1 needed to achieve desired constriction across all experiments are summarized in Table 2. LPA shifts the mediator of FID from NO to H2O2 in human arterioles To confirm NO as the baseline mediator of FID in human adipose arterioles, vessels were treated with either L-NAME or Peg-Cat in the absence of LPA. Presence of L-NAME blunted peak FID (% maximal dilation to 100 cm H2O: Control 73.7±3.8, n=6 versus L-NAME 13.2±4.8, n=7, p<0.05). Peg-Cat had no effect on the dilation in untreated arterioles (Figure 1A). Treatment of Non-CAD vessels with LPA preserved the magnitude of dilation but shifted the meditator of FID to H2O2 (dilation inhibited by Peg-Cat) (% maximal dilation to 100 cm H2O: control 72.3±4.8, n=10 versus Peg-Cat 26.5±7.3, n=7, p<0.05) (Figure 1B). In contrast to its inhibitory effect on dilation in untreated vessels (Figure 1A), pre-incubation with L- NAME in LPA-treated vessels did not inhibit but instead slightly increased dilation at lower levels of flow (Figure 1B) (% maximal dilation to 20 cm of H2O: control 58.5±6.4, n=10 versus L-NAME 83.3±5.0, n=7, p<0.05). Arterioles treated with LPA also showed reduced dilation to an endothelium-dependent dilator (acetylcholine) with differences in peak value (% maximal dilation to -4 (Log M) ACH, control 87.5±5.1, n=5 vs LPA 19.2±8.09, n=5, p<0.05, Figure 1C) and EC50 (– log EC50: 6.83±0.18 for control and – log EC50: 5.86±0.44 for LPA, p<0.05). To determine whether treatment with LPA affected endothelium- independent arteriolar responses, dose-response curves to sodium nitroprusside (a direct NO donor) were performed in LPA-treated and untreated arterioles. No significant differences were observed in EC50 (– log EC50: 7.03±0.24 for control and – log EC50: 7.03±0.23 for LPA, Figure 1D) or peak values (% maximal dilation to – 4 (Log M) SNP, control 85.3±1.90, n=5 vs LPA 92.2±2.85, n=4, Figure 1D), both with p=NS. Additiona l ly, because two videomicroscopy studies (Control vs. L-NAME or Peg-Cat) were often performed in succession on a single arteriole, we also assessed whether LPA treatment had a time- dependent effect on the overall magnitude of FID and found no differences in these time control experiments (Supplemental Figure 1). Signaling through LPA1 receptor leads to the loss of NO-mediated FID in non-CAD human arterioles We next examined which receptor might be responsible for the LPA-induced shift in the mediator of FID. Figure 2A shows that non-CAD arterioles treated with the LPA1/3 antagonist, Ki16425, maintained NO-mediated FID after the acute LPA challenge (% maximal dilation at 100 cm H2O: Ki16425+LPA 81.1±4.6, n=6 versus Ki16425+LPA+L-NAME 23.8±8.7, n=7, p<0.05). Peg-Cat, which inhibits FID after LPA treatment alone, had no effect on the dilatio n after combined treatment with LPA and Ki16425 (% maximal dilation at 100 cm H2O: Ki16425+LPA 81.9±4.1, n=8 versus Ki16425+LPA+Peg-Cat 79.1±4.0, n=7). Blocking LPA2 (Figure 2B) did not prevent the shift to H2O2-mediated dilation (Peg-Cat abolished FID) (% maximal dilation at 100 cm H2O: H2L5186303+LPA 82.6±4.2, n=6 versus H2L5186303+LPA+Peg-Cat -9.7±2.0, n=6, p<0.05). To ensure that the shift to H2O2-mediated dilation is not caused by the LPA2 receptor antagonist alone, we tested the effects of the antagonist in the absence of LPA and observed no effect on NO-mediated dilatio n (Supplemental Figure 2). H2L5186303 is an LPA2 receptor antagonist when used at a nanomolar concentrations, but at a higher, micromolar, concentrations it can also act as an LPA3 receptor antagonist (Fells et al., 2009). We, therefore, tested a higher concentration of H2L5186303 (10 µM) before LPA challenge. As before, Peg-Cat inhibited the dilation (% maximal dilation at 100 cm H2O: H2L5186303+LPA 80.6±4.6, n=3 versus H2L5186303+LPA+Peg-Cat -26.3±1.2, n=3) while L-NAME had no effect (Supplementa l Figure 3). LPA receptors 1 and 2 but not 3 are expressed in human adipose microvessels To investigate which LPA receptors are responsible for the arteriolar effects of LPA, we examined LPA1-3 receptor expression in intact human adipose arterioles. As shown in Figure 3, transcripts for LPA1 and LPA2 receptors but not LPA3 were abundantly detectable by PCR. The pattern of expression was not different between CAD vs. non-CAD tissue for any of the three receptors (Figure 3A). Because our functional studies using LPA2 receptor antagonist demonstrated no involvement of that receptor in the LPA-induced shift of the mediator of FID, the remaining expression studies focused on LPA1 and LPA3 receptors. Western blot analysis was performed on LPA1 and LPA3 receptors to examine protein levels. Figures 3B show robust detection of LPA1 in both CAD and non-CAD vessels as well as HUVECs and left ventricle of the human heart, which was used as a positive control. LPA3 receptor was not detected in arterioles from patients with or without CAD, and only faint expression was detected in HUVECs (Figure 3C). LPA3 receptor was also detected in human heart samples with some additional non-specific bands. Immunohistochemical studies of human non-CAD arterioles also demonstrated strong staining for LPA1 (Figure 4A) but not LPA3 (Figure 4B). A similar pattern of expression was observed in CAD arterioles, with LPA1 receptor but not LPA3 being expressed the vessels. Sections of human pancreas were used to verify efficiency of the LPA3 receptor antibody (Supplemental Figure 6). Immunohistochemical analyses appeared to localize LPA1 receptor expression predominant ly to VSMCs, with little to no staining in ECs (Figure 4A and B). To further investigate whether LPA receptors are expressed in human ECs, we examined expression of LPA1 and LPA3 receptors in two types of cultured endothelial cells, human umbilical vein endothelial cells (HUVECs) and human microvascular endothelial cells (HMVECs). There was robust expression of LPA1 receptors in HUVECs and HMVECs as shown in Figures 5A and 5B, respectively. LPA3 was detected in HUVECs, albeit at a low level, and no expression of the LPA3 receptor was detected in HMVECs (Figures 5C and 5D, respectively). Two differe nt antibodies (see methods section) targeting different epitopes were used to ensure validity of the expression patterns of LPA3 receptor in arterioles and cultured endothelial cells. In both cases, we found the expression of the receptor to be lower or absent across different preparations. Supplemental Figure 4 shows lower expression of LPA3 receptor in two types of cultured endothelial cells, with robust expression in the heart using Abc antibody, while Supplementa l Figure 5 validates lower expression of LPA3 receptor in non-CAD arterioles and immunohistochemical preparations using LSBio antibody. These data suggest that LPA1 receptors are expressed in the arteriolar medial layer, and to a much lower extent in vascular endothelium, even though expression is robust in culture microvascular endothelial cells. LPA3 receptors are minimally expressed in human coronary arterioles or cultured endothelial cells. We previously showed that the mitochondria are the main source of flow-induced H2O2 release in CAD arterioles (Freed et al., 2014; Liu et al., 2003). To assess whether LPA specifica l ly increases mitochondria-derived H2O2 in non-CAD arterioles, we applied mitoPY1, a fluorescent probe specifically designed to image mitochondrial H2O2 in our cannulated arteriole experiments. Figure 6A shows representative images of the cannulated vessels in the bright field, under static conditions and after 5 minutes of maximal (100 cm H2O) flow, in intact untreated and LPA-treated (10 µmol/L, 30 min) and as well as denuded untreated and LPA-treated arterioles. Quantification of the relative fluorescence intensity is shown in Figure 6B. Untreated, non-CAD vessels show minimal increases in fluorescence as compared to LPA- treated, non-CAD arterioles (% increase from static after 5 min of 100 cm H2O: untreated 9±16, n=6 versus LPA 183±32, n=6). Denuded controls and LPA-treated arterioles showed a trend toward an increase in fluorescence in comparison to intact control vessels (Figure 6B). More importantly, denuded arterioles treated with LPA do not showed a significant increase in in fluorescence signal in response to flow when compared to intact arterioles treated with LPA. Denudation efficiency was analyzed by examining expression of endothelium-specific target protein, PECAM (Supplemental Figure 7). To further test the mitochondria as the source of H2O2, we measured FID in the presence of Rotenone, an electron transport chain complex I inhibitor that has been shown previously to reduce mtH2O2 production and block FID in arterioles from patients with CAD (Liu et al., 2003). Figure 6C shows that rotenone significantly reduces FID after LPA treatment (% maximal dilation to 100 cm of H2O, LPA 73.2±4.77, n=10 vs LPA+rotenone 39.8±7.89, n=7, p<0.05). Rotenone alone has been known not to affect either the overall magnitude of the dilation or the mediator of FID (Beyer et al., 2016; Freed et al., 2014). Discussion The major novel findings of the present study are as follows: first, acute (30 min) exposure of non-CAD human arterioles to 10 µM LPA induces a switch in the mechanism of FID from NO to H2O2, recapitulating the CAD phenotype; second, LPA1 receptor is involved in the LPA- induced shift to H2O2-mediated FID; third, LPA1 and LPA2 receptors but not LPA3 are expressed in human arterioles from patients without CAD; and fourth, mitochondria are the source of LPA-induced H2O2 production. These findings show for the first time that elevated concentrations of LPA act in an LPA1 receptor-dependent manner to induce an arteriolar endothelial phenotype characteristic of that seen in patients with CAD. Elevated LPA is associated with vascular pathology and is known to exert multiple harmful effects on vascular endothelial (Avraamides et al., 2007; Lin et al., 2006; Lin et al., 2007; Panetti, 2002; Panetti et al., 2004; Panetti et al., 2000; Rizza et al., 1999; van Nieuw Amerongen et al., 2000) and smooth muscle (Hayashi et al., 2001; Panchatcharam et al., 2008) integrity. In particular, studies have shown that LPA can lead to increased levels of ROS either independent of (Brault et al., 2007; Staiculescu et al., 2014) or in conjunction with decreased eNOS expression (Chen et al., 2012), effectively decreasing endothelium-dependent dilatio n to bradykinin in porcine arterioles (Chen et al., 2012). We extend these findings and demonstrate that acutely elevated levels of LPA disrupt endothelium-dependent FID in human arterioles by shifting the mechanism away from NO to mtH2O2. There was a noticeable improvement in dilation at lower pressure gradients after L-NAME treatment (Figure 1B), which could potentially be ascribed to elimination of uncoupled eNOS-derived reactive nitrogen/oxygen species, which scavenge the dilatory ROS (Yang et al., 2009) or the fact that quenching any remaining NO releases the block on mtH2O2 (Beyer et al., 2017). LPA significantly reduced dilation to ACH, potentially by increasing intracellular ROS, which can compromise endothelial NO bioavailability. This observation is consistent with others from our lab (Beyer et al., 2016; Durand et al., 2016) where acetylcholine-induced dilation is more sensitive to ROS than FID. LPA did not negatively affect smooth muscle responses to NO as indicated by lack of difference in dilation to SNP between treated and untreated vessels. Additionally, the amount of constrictor agent, ET-1, did not vary between control vs LPA- treated vessel. Presence of LPA and L-NAME, did require more ET-1. Although LPA has been described to affect vessel tone (Tokumura et al., 1981; Tokumura et al., 1995), our data argues against either a direct vasoconstrictor or vasodilator effect of LPA, but rather suggests that LPA-induced production of the dilator H2O2 might be responsible for the higher dose required for constriction since presence of Peg-Cat (H2O2 scavenger) eliminated that effect. To our knowledge, this is also the first study examining the function and expression of LPA1- LPA3 receptors in human resistance-size arterioles. Functional data from vessels treated with LPA2 receptor antagonist indicated that unlike LPA1 and/or LPA3, LPA2 fails to shift the mediator of FID. Instead, LPA1 receptor appears to be mechanistically involved. qPCR analysis showed robust expression of LPA1 and LPA2 but not LPA3, with no difference between arterioles from subjects with and without CAD. Protein analysis from human arterioles identified LPA1 receptor as the dominant isoform with no difference between CAD and non- CAD phenotype. LPA3 receptor protein was detected in human heart, albeit at a lower level than LPA1 receptor. Faint expression of LPA3 receptor protein was also detected in HUVECs. Absence or minimal expression of LPA3 has been observed in endothelial (Lee et al., 2000; Lin et al., 2007; Ruisanchez et al., 2014) and smooth muscle (Dancs et al., 2017) cells from animal models or cultured vascular human cell lines. We confirmed previously published data using human umbilical vein endothelial cells and human microvascular endothelial cells. Immunohistochemistry confirmed robust staining for LPA1 in comparison to LPA3 in arterioles but also localized the signal predominantly to SMC with little to no expression in arteriolar endothelial cells.Lack of pronounced expression of LPA receptors, in particular LPA1 receptor, in arteriolar tunica intima was an unexpected finding. Prior findings in animal models and endothelial cell cultures shows majority of LPA effects being mediated through LPA1 receptor (Brault et al., 2007; Cui, 2011; Dancs et al., 2017; Gobeil et al., 2003; Ruisanchez et al., 2014). Additiona l ly, we and others observe strong protein and RNA expression in cultured endothelial cells (Lin et al., 2007; Ptaszynska et al., 2010; Ren et al., 2011). Collectively, we interpret these findings to indicate that LPA1 are likely present in human arteriolar endothelial cells, albeit at a much lower level of expression than in SMC, yet sufficient to cause an LPA-induced shift in the mediator of FID. An alternative explanation of these findings is that LPA acts on SMC to release a factor that acts on the endothelium in a way that changes the mediator of FID. Future efforts should be centered on evaluating the LPA-mediated bidirectional crosstalk between these two vascular layers as well as cell-specific expression of these receptors in their native vascular environment. The source of ROS production is not identified in most LPA-related vascular studies, but mitochondria and NADPH Oxidases (NOX) are prominent candidates in ECs (Breton-Romero & Lamas, 2014; Panieri & Santoro, 2015; Widlansky & Gutterman, 2011). We tested whether mitochondria were a source of H2O2 under LPA challenge using the mitochondrial complex I inhibitor, rotenone. Rotenone reduced dilation to the same extent as the general H2O2 scavenger, Peg-Cat. A mitochondrial source of H2O2 was further suggested using a mitochondria-specific fluorescent probe. In some settings NOX is the source of LPA- stimulated ROS (Lin et al., 2013). We cannot exclude a role for NOX in FID following LPA treatment, which could occur via ROS–induced ROS release where small amounts of NOX- related ROS stimulate release of mitochondrial ROS or vice versa (Zinkevich et al., 2017; Zinkevich & Gutterman, 2011). In fact, a recent study showed LPA1 and LPA3 receptor- dependent activation of PLC, PKC and eventual stimulation of NOX to generate ROS in cell culture model (Lin et al., 2013). Our current findings align with previously published observations demonstrating that the mitochondria are the main source of the dilator H2O2 in human arterioles affected by CAD (Beyer et al., 2016; Freed et al., 2014; Kadlec et al., 2017). Acute LPA exposure thus establishes a CAD phenotype in non-CAD arterioles. Details of the signaling pathway by which LPA induces the shift to H2O2-mediated FID through LPA1 necessitates further investigation. One compelling possibility is the well-docume nted crosstalk between Rho- and Rac-dependent cytoskeleton rearrangement and ROS signaling. In fact, recent publication from Chandra and colleagues, using real-time measurements, demonstrated that increasing levels of LPA or down-regulating expression of LPP3, one of the main enzymes that degrades LPA, leads to dysfunctional mitochondria with increased production of superoxide in a Rho- and ERK- dependent manner (Chandra et al., 2018). LPA induces changes to the endothelial cytoskeleton in an LPA1 receptor-dependent manner by phosphorylating myosin light chain (Wu et al., 2015), forming stress fibers and RhoA- and Rac-dependent cytoskeletal rearrangements (Siess et al., 1999). Studies examining mitochondrial dynamics identified LPA, amongst other glycerol-based lipids, to regulate mitochondrial fission and fusion (Baba et al., 2014; Frohman, 2015; Ha & Frohman, 2014). Together, these studies suggest that LPA may affect mitochondrial structural dynamics, loss of which results in increase in mitochondria-derived ROS production. Future studies will focus on examining the potential downstream mediators of the observed LPA-induced effects. Preliminary studies from our laboratory examined the ability of arterioles to recover and regain NO-mediated FID by incubating LPA-treated arterioles in fresh media lacking LPA. Even overnight (~18 hours) incubation in fresh media of arterioles treated with LPA for 1 hour did not restore NO as the dilator (Supplemental Figure 8). We therefore suggest that once the LPA- activated signaling cascade is initiated, damage is prolonged, predisposing the vasculature to further damage caused by increased ROS. This observation sheds light on an exciting clinica l opportunity, whereupon assessing the expression of enzymes responsible for both synthesis and degradation of LPA may prove to be a novel preventive or early intervention strategy. In fact, a risk allele leading to decreased expression of LPP3, the enzyme that degrades LPA, has been identified by genome-wide association studies in nearly eighty percent of the population, and shown to be highly predictive of CAD, independent of traditional risk factors (Erbilgin et al., 2013; Schunkert et al., 2011). Therefore, the effects of LPA revealed in this study may affect a large portion of the population and represent an underappreciated pathophysiologica l pathway in cardiovascular disease. We obtain samples that are surgical discards and are therefore unable to control for all patient characteristics. Vessels are isolated from different fat depots, but we have not observed major differences in vascular responses to flow as evident in our previous publications. The mechanism of FID seems to be preserved across these vascular beds, including the phenotypic switch in tissue acquired from patients with clinically diagnosed CAD. The non-CAD arterioles are isolated from individuals whose clinical profiles (medications, demographics, etc) are not fully known. However, we do acquire information on prior diagnosis of CAD and presence of risk factors. By defining our non-CAD cohort strictly as having one or no CV risk factors, we limit the degree to which non-CAD samples are classified falsely. Females represent most of our non-CAD vessels. Our sample sizes are too small to detect sex differences in vascular function within and between the phenotypes but this should be addressed in a larger study. Although we used receptor antagonists well-documented in the literature, it is possible that they may be non-specific or have off-target effects. We addressed such possibilities by testing antagonists for different receptors as well as examining receptor expression. In regard to LPA1 receptor expression, although we and others show expression in cultured endothelial cells, we were unable to detect the receptor in arteriolar ECs using immunohistochemical approach. This suggests at least three possibilities. 1) arteriolar ECs may have LPA1 receptors sufficient to elicit the switch but the immunohistochemical approach is not sufficiently sensitive to detect them; 2) the epitope recognized by the antibody is not accessible in IHC preparations; or 3) LPA may be acting on LPA1 receptor in SMC and stimulate release of a factor that acts on the endothelium in a way that changes the mediator of FID. Additionally, we have made attempts to knockdown LPA1 receptor using siRNA at two different concentrations with incubat ions periods as long as 48 hours (time limit for isolated arterioles). However, these attempts have been unsuccessful. siRNA transfected arterioles do not show decrease in LPA1 receptor protein expression. Lastly, because we did not perform histological analysis on all the arterioles used throughout our studies, we cannot exclude the possibility of denudation in some of our vessels. However, because Ki16425 FID is an endothelium-dependent process (in the presence or absence of LPA), maintained dilation suggests that the endothelium was intact during our studies.