Lysophosphatidic Acid (LPA) Signaling and Cardiovascular PathologySUSAN S. SMYTH, ANPING DONG, JESSICA WHEELER, MANIKANDAN PANCHATCHARAM, and ANDREW J. MORRIS
13.1. INTRODUCTION
Since the initial reports that lysophosphatidic acid (LPA) affected blood pres-sure in mammals (1, 2), a body of literature supports an important role for this bioactive lipid in regulating cardiovascular physiology and pathology. LPA exerts diverse effects on most blood and vascular cells. LPA circulates in plasma primarily bound to serum proteins and lipoprotein particles. However, whether circulating pools of LPA are the key determinant of biologic activity remain uncertain. In this chapter, the pathways for maintenance of plasma LPA are briefly reviewed; current understanding of blood and vascular cells responses to LPA are discussed; and evidence for a role in cardiovascular pathology, including a review of relevant methods, is presented.
13.2. CIRCULATING LPA LEVELS
LPA is present in plasma at levels (≥100 nM) sufficient to elicit biologic effects on G protein-coupled receptors (3), although the contribution of plasma LPA to actions at cell surface receptors is not clear. The source for most extracellular, biologically active LPA is the secreted lysophospholipase D (lysoPLD) (4, 5) autotaxin, a member of the ectonucleotidase pyrophos-phate family encoded by Enpp2 and present in plasma at concentrations of 0.5–1 mg/L (approximately 5–10 μM) (6). Autotaxin generates LPA by
hydrolysis of lysophosphatidylcholine (LPC), which is present in plasma at concentrations of approximately 100 μM. LPC is formed when lecithin-cholesterol acyltransferase (LCAT, also called phosphatidylcholine-sterol O-acyltransferase) converts free cholesterol into cholesteryl esters. However, LPA levels are normal in individuals lacking LCAT (7). Therefore, plasma LPA likely derives from LPC generated from phospholipase A (PLA) action on phosphatidylcholine (PC) present in lipoproteins, circulating microparti-cles, platelets, or other blood and vascular cells (7–9). In rodents, small-molecule autotaxin inhibitors and autotaxin antibodies lower LPA levels (10, 11), and mice genetically engineered to have reduced autotaxin expression also have lower LPA levels (12, 13). Plasma LPA may be generated in an autotaxin-independent manner by PLA2-mediated hydrolysis of phosphatidic acid (PA), although the PLA2 responsible has not been identified, and the contribution of this pathway to plasma LPA levels appears to be fairly small relative to that of autotaxin.
In addition to the requirement for autotaxin, blood platelets and platelet-derived microparticles have been implicated in plasma LPA generation. LPA levels are reduced in rats by antibody-mediated thrombocytopenia or treat-ment of mice with a platelet integrin inhibitor that also induces thrombocyto-penia (7, 14). Platelet-derived microparticles have also been cited as a source for the generation of LPA (9). Isolated, washed platelets produce relatively small amounts of LPA upon agonist stimulation (15, 16), and LPA production by isolated, activated platelets is prevented by autotaxin inhibitors (17). One explanation for these observations is that activated platelets, and potentially their microparticles, provide an LPC source for the generation of LPA by autotaxin. In keeping with this, Tigyi and colleagues recently purified a PLA1 from thrombin-activated platelets that increases the production of LPC and LPA in plasma (8). They identified the activity as acyl-protein thioesterase 1 (APT1, also called lysophospholipase A-I [LYPLA-I]) and proposed that sn-2 LPC generated by APT1 undergoes acyl migration to sn-1 LPC, which serves as a substrate for autotaxin (8).
The working model for platelet LPA production, in which PC is converted to LPC by APT1 and then to LPA by autotaxin, necessitates that autotaxin interacts with cell surfaces. How autotaxin accesses LPC generated during platelet activation and whether LPA is generated along the membrane bilayer or released freely or both are not known. The crystal structure of autotaxin reveals a hydrophobic substrate binding channel leading to the active site (18, 19). The channel is formed at the interface of the somatomedin B-like (SMB) and phosphodiesterase (PDE) domains of the protein and may provide a means to spatially restrict the generation and delivery of LPA. Autotaxin can localize to cell surfaces by binding to activated integrins, as has been observed with platelets (20), lymphocytes (21), and oligodendrocytes (22). The interac-tion of the SMB domain of autotaxin with activated integrins on platelets and other cells may alter autotaxin activity and enable localized production of cell-associated LPA.
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13.2.1. Protocols for LPA Measurement in Plasma
Whole blood should be collected into an anticoagulant solution of EDTA (10 mM) and citrate-theophylline-adenoisne-dipyridamole (CTAD; 10% v/v, BD Biosciences, San Jose, CA). An autotaxin inhibitor can be added to prevent ex vivo LPA production during sample processing. We typically spin the blood at 12,000 g × 5 minutes and immediately freeze the resultant plasma. This protocol results in minimal platelet activation as judged by plasma platelet factor 4 levels. LPA circulates in plasma primarily bound to serum proteins and lipoprotein particles. The distribution of LPA can be monitored by size-exclusion fractionation of plasma. Typically, plasma is chromatographed on a Superdex 75 or similar column (Pharmacia, Kalamazoo, MI). LPA content in fractions is measured by high-performance liquid chromatography (HPLC)/electrospray ionization (ESI)/tandem mass spectrometry and lipoprotein frac-tions identified by immunoblot analysis or cholesterol content.
Studies in which autotaxin inhibitors or LPA is administered to animals indicate rapid metabolism of LPA occurs in circulation and whole blood, suggesting dynamic regulation by synthetic and metabolic/clearance path-ways (11, 23). At present, the mechanism(s) responsible for LPA metabolism and clearance in blood are not known, although a role for the lipid phosphate phosphatase 1 (LPP1) has been suggested (23). Unlike the rapidly metabo-lism observed in whole blood, plasma LPA is relatively stable. Therefore, measuring LPA accumulation in plasma incubated at 37°C for 60–120 minutes under conditions that support autotaxin activity is a simple and reliable method to monitor enzyme activity in plasma. Small-molecule inhibitors can be included in the assay to confirm the autotaxin-dependent nature of LPA production.
13.3. LPA SIGNALING IN BLOOD AND VASCULAR CELLS
Three receptor systems have been implicated in mediating LPA’s signaling effects on cells. These are a family of G protein-coupled receptors (24), peroxi-some proliferator-activating receptor γ (PPARγ) (25), and, more recently, the receptor for advanced glycation end products (RAGE) (26). At least six LPA G protein-coupled receptors, termed LPA1–6, may mediate responses to extracellular LPA. These LPA receptors couple to multiple heterotrimeric G proteins (Gi/o, Gq/11/14, Gs, and G12/13) and initiate various signal transduction pathways. In the vasculature, G13-mediated activation of Rho appears to play an especially prominent role. The contribution of LPA receptors to signaling in blood and vascular cells is described in more detail below. The nuclear receptor PPARγ, which is important in blood and vascular cell signaling, has been proposed as a receptor for LPA (25, 27). This pathway may be acti-vated by intracellular LPA generated by the glycerol-3-phosphate pathway
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because cells overexpressing glycerol-3-phosphate acyltransferase-1 have sixfold elevated levels of intracellular LPA and higher PPARγ activity, whereas the addition of extracellular LPA lacked effect on PPARγ (28). These findings suggest an alternate role in “transcellular” LPA signaling. Recently, RAGE has been implicated as a potential mediator of LPA signaling in the vascula-ture, and the absence of RAGE in mice resulted in a loss of vascular LPA signaling (26). The ability of LPA to bind with high affinity to soluble RAGE fragments suggests a direct role for the receptor in LPA signaling, although this remains to be firmly established.
13.4. REGULATION OF BLOOD PRESSURE
Exogenous administration of LPA alters blood pressure in animals. Intrave-nous injection of LPA transiently elevates arterial blood pressure in mice (29) and rats (2). Local application to porcine cerebral vessels elicits vasoconstric-tion (1). The receptors mediating this response are not known. In response to intravenously administered LPA, we have observed an increase in mean arte-rial pressure in mice lacking LPA1, LPA2, both LPA1 and LPA2 (29), LPA4, and PPARγ in smooth muscle cells (SMC) (unpublished observations), and others have reported that the elevated blood pressure occurs in Lpar3–/– mice.
13.4.1. Protocol for Arterial Pressure Measurements
Systolic blood pressure and heart rate can be measured noninvasively in con-scious mice by tail cuff analysis using methods that either detect cuff pressure (BP-2000, Visitech, Apex, NC) or volume pressure recordings (CODA, Kent Scientific, Torrington, CT). The mice are acclimated to the procedure by daily training for 1 week, and recordings are then made for five consecutive days. To measure acute changes in blood pressure in response to intravenous admin-istration of LPA, mice are anesthetized with isoflurane, placed on a mouse pad that has imbedded EKG electrodes and surface mounted semiconductor tem-perature sensor to distribute heat, and a pressure catheter (1.4 Fr Millar [Houston, TX] catheter) is introduced into the carotid artery. A data acquisi-tion and recording system (PowerLab with LabChart, ADInstruments, Colo-rado Springs, CO) is used to display readings in real time. When injected intravenously into either the jugular or the femoral vein, LPA elicits a tran-sient increase in mean arterial pressure that begins within ≈15 seconds and begins to return to baseline at ≈1 minute and is accompanied by a reflex bra-dycardia. The “gold standard” for blood pressure recordings in rodents is implantation of telemetric pressure transmitters, such as those supplied by Data Sciences International (St. Paul, MN) that record arterial pressure for prolonged periods of time in conscious, free-roaming animals. The drawbacks of implantable telemetric units include requirements for advanced skills in rodent survival surgery and expense of the monitoring equipment.
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13.5. BLOOD VESSEL AND LYMPHATIC FORMATION
The most compelling evidence for the role of LPA in blood vessel formation comes from observations in mice with inherited absence of either autotaxin or LPA receptors. The lack of autotaxin results in embryonic lethality in mice due in part to improper vasculogenesis (12, 13). Vascular endothelial growth factor (VEGF) increases autotaxin expression in angiogenic endothelial cells in a phospholipase C γ (PLCγ)-dependent manner, and autotaxin may regulate endothelial tube remodeling (30). When mixed in Matrigel (BD Biosciences) and implanted subcutaneously, autotaxin elicits angiogenesis in mice (31). Although the mechanisms responsible are not fully understood, in cultured endothelial cells, LPA activates protein kinase D (PKD) and downregulates CD36, a receptor for the antiangiogenic protein thrombospondin (32). Attenu-ated expression of CD36 may blunt the inhibitory effects of thrombospondin and allow angiogenesis to proceed unchecked. LPA promotes endothelial migration largely through Rho-dependent processes that alter the actin cyto-skeleton and extracellular matrix composition (32). Genetic deficiency of the LPA4 receptor, which couples to G13 (33, 34), causes edema and hemorrhage in a subset of embryo and resultant lethality (35). In adult Lpar4-null mice, fewer endothelial-lined vessels form in Matrigel implants, impairments in lym-phatics are also present in Lpar4-null mice, and a role for LPA in lymphangio-genesis has been supported in in vitro systems (36). Thus, autotaxin and LPA via LPA4, and potentially other receptor systems, may regulate both vascular and lympathic vessel formation (35).
13.5.1. Protocol for Matrigel Angiogenesis Assay
Anesthetized mice are shaved to expose the skin on both flanks. Matrigel (200–250 μL or sterile, reduced-growth-factor Matrigel at 10 mg/mL) prepared with purified autotaxin (0.1–1 nM) (31) or basic fibroblast growth factor (bFGF; 250 ng/mL) and heparin (60 U/mL) are injected subcutaneously on the left center of the back and on the right center of the back using a 27 G needle and tuberculin syringe. Typically, the Matrigel plug on one side is used as the matched control for the other (e.g., catalytically inactive autotaxin or heparin only). The mice should be singly housed until fully recovered and the Matrigel plug has solidified. Vessel growth in the plug occurs within 7–14 days and can be visualized by intravenously injecting 200 μL of FITC-Dextran, (50 mg/mL, 2 × 106 m.w., Sigma, St. Louis, MO), either through a lateral tail vein or an exposed jugular vein. Mice are euthanized 10 minutes later. Vessels can also be visualized by immunohistochemical visualization of in the Matrigel implants using antibodies to endothelial and SMC-specific antigens.
13.6. VASCULAR PERMEABILITY
While studies in cultured endothelial cells have yielded variable results with regard to the effects of LPA on endothelial barrier function, in a bleomycin
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model of lung injury, mice lacking LPA1 display reduced fibrosis and vascular leak (37). In this injury model, bleomycin increases LPA levels in alveolar fluid. The protection from lung edema observed in the LPA1-deficient mice could reflect a pathologic role of LPA in promoting endothelial permeability. Some, but not all, studies of isolated endothelial cells support a model in which LPA disrupts endothelial barrier function (see References 38–41; reviewed in Ref-erence 42). In support of barrier destabilizing effects of LPA, topical applica-tion provokes cerebromicrovascular permeability in mice (43). Interestingly, in the bleomycin model, lack of LPA1 did not influence recruitment of inflam-matory cells into damaged lungs, which might be expected to be influenced by endothelial barrier function.
13.6.1. Vascular Permeability Protocol
Evans blue dye (200 μL of a 1% solution in phosphate-buffered saline [PBS]) is injected intravenously. Fifteen minutes later, mice are euthanized and per-fused with PBS containing heparin through the right ventricle at a constant rate to yield a pressure of 25 mmHg. The lungs are dissected, weighed, and the entire right lung scanned using an Odyssey Infrared Imaging System 2.1 (LI-COR Inc., Lincoln, NE) to detect Evans blue. As an alternative method to detect Evan blue, the lung can be immersed in 1 mL 4% formamide and Evans blue extracted by shaking in a water bath at 56°C for 24 hours and quantitated by absorbance at 600 nm. The values for Evans blue are normalized to lung weight. It is important to obtain measurements of both wet and dry lung weight. Vascular leak can be provoked with an inflammatory challenge, for example, bacterial LPS (2 mg/kg, E. coli 0111:B4; Sigma) administered intra-peritoneally 4–6 hours before Evans blue.
13.7. VASCULAR INFLAMMATION
LPA may promote inflammatory responses either indirectly through endothe-lial cells or by direct effects on leukocytes. In addition to altering endothelial barrier properties, LPA may “inflame” endothelial cells by upregulating the expression of endothelial chemoattractants (21, 44, 45), stimulating the secre-tion of the pentraxin-3, an acute phase reactant with both proinflammatory and prothrombotic features (46), and triggering exposure of endothelial adhe-sion receptors (47). The endothelial effects of LPA are mediated by both LPA1 and LPA3 and likely promote endothelial–white blood cell interactions in vivo. Autotaxin expression is upregulated in monocytic-like cells following LPS exposure (48), and localized autotaxin activity regulates lymphocyte traffick-ing (21). It is therefore reasonable to speculate that the generation of LPA during inflammation may have direct cellular consequences. Neutrophils (49, 50), eosinophils (51), and mononuclear phagocytes (48, 52) are responsive to LPA, and LPA triggers platelet–monocyte coaggregate formation in whole
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blood (53). LPA also elicits the release of IL-6 from SMCs (54). In an air pouch model, LPA recruits both neutrophils and monocytes, triggers the release of IL-6 and KC, and enhances TNFα effects in an LPA1- and LPA3-dependent manner (55). Together, these results focus attention on LPA1 and LPA3 as potentially important mediators of vascular inflammation.
13.7.1. Inflammation Models
Two classic experimental models of acute inflammation are peritonitis and dermatitis. Peritonitis is induced by intraperitoneal injection of 1 mL of thio-glycollate (3% w/v). The mice are euthanized at 6 hours after the injection, at which time peritoneal fluid is collected to determine the number of leukocytes that transmigrated in response to the thioglycollate. Dermatitis is induced by topical application of croton oil (10 μL of 2% solution in 4:1 acetone : olive oil). After 6 hours, the mice are euthanized and the tissue removed and fixed in paraformaldehyde for subsequent histologic assessment, which includes leu-kocyte accumulation. To our knowledge, the effects of LPA in eliciting perito-nitis or dermatitis has not been examined. However, as discussed above, LPA does trigger neutrophil and monocyte migration into air pouches (55). In the air pouch model, 3 mL of sterile air is injected subcutaneously twice, 72 hours apart. Four days after the second air injection, 1 mL of PBS containing 0.1% bovine serum albumin and 1–6 μg of LPA (2.3–13.9 μM) is introduced into the pouch. The pouch is flushed with sterile buffer 6 hours later and leukocytes collected.
13.8. ATHEROTHROMBOSIS
Atherosclerosis is a chronic inflammatory condition with the hallmark features of endothelial dysfunction, lipid accumulation, vascular inflammation, and a fibroproliferative response of resident SMCs. The complications of atheroscle-rosis occur either as a consequence of tissue ischemia resulting from obstruc-tions in blood flow by the atheromatous plaque or when plaque ruptures or erodes and causes platelet-dependent thrombus formation, the proximate cause of most heart attacks and many strokes. LPA is present in human ath-eroma, particularly enriched in the lipid-rich core, and levels increase in exper-imental mouse models during progression of atherosclerosis (56, 57). Advanced lesions in Ldlr–/– mice, generated by a combination of diet and collar placement around the carotid artery, contain ≈20-fold higher levels of LPA than uninjured vessels, especially highly unsaturated long-chain acyl-LPA species (58). Several pathways may result in accumulation of LPA in atherosclerotic plaque. LPA is produced during mild oxidation of low-density lipoprotein (LDL) (56) and may also be generated from LPC, which markedly increases in atherosclerotic/injured arteries of Ldlr–/– mice (58). Within days to weeks of feeding male New Zealand white rabbits a diet with 1% cholesterol, serum LPC levels increase.
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The increase in LPC is accompanied by heightened generation of biologically active LPA in serum incubated ex vivo for 24 hours, presumably as a conse-quence of the lysoPLD actions of autotaxin (59).
LPA may serve a prothrombotic function based on its ability to elicit pro-coagulant phosphatidylserine exposure on erythrocytes (60), its platelet-activating effects (61), and its stimulation of tissue factor expression on SMCs (62). These observations have led to the speculation that LPA is an important prothrombotic component of atherosclerotic plaque (56, 63, 64). LPA’s effects on platelets are primarily through G13-dependent signaling, resulting in shape change, fibronectin matrix assembly, and potentiation of other platelet ago-nists (65–68). The platelet LPA response demonstrates an interesting pharma-cology, with alkyl glycerol phosphate (“1-alkyl-LPA”) being a more potent agonist than LPA (69), a pharmacologic profile shared by LPA5 (70). While human platelets and megakaryocytes contain mRNA for several LPA recep-tors, transcript levels for LPAR5 are among the highest for GPCRs in human platelets (71). RNAi targeting of LPAR5 in megakaryocyte lines prevents shape change in response to both LPA and to lipid extracts from atheroscle-rotic plaque (72). Taken together, these observations implicate LPA5 as the LPA-activating receptor on human platelets. Confirmation of LPA5 as the major platelet LPA receptor by use of genetic approaches in mice is ham-pered by the observation that LPA does not elicit a similar stimulatory effect on murine platelets (20), which, interestingly, lack expression of Lpar5 tran-scripts by RNA-seq analysis (73).
Phenotypic modulation of SMCs occurs in response to vascular injury and is a critical component in the development of intimal hyperplasia, a defining feature of atherosclerotic and restenotic lesions (74, 75). Phenotypic modula-tion involves the dedifferentiation, proliferation, and migration of normally quiescent SMC in the vessel wall and takes place when isolated SMCs are cultured in serum. As has been reviewed elsewhere in detail (76), LPA pro-motes all aspects of the phenotypic response in SMCs, including dedifferentia-tion (27, 54, 77–79), proliferation (80–82), and migration (83–87), and appears to be a key component of serum responsible for phenotypic modulation of SMCs (78, 88). LPA also triggers inflammatory and prothrombotic phenotypes in SMCs by upregulating expression of mediators such as IL-6 (54) and tissue factor (62). Thus, the effects of LPA on cultured SMCs would support a caus-ative role for the lipid in the development of intimal hyperplasia in the settings of atherosclerosis and restenosis. Such a role is supported by the discovery that exogenous application of LPA stimulates neointimal formation in both rat and mouse arteries (27, 77, 89). The process does not occur in the presence of an antagonist to PPARγ or in mice lacking PPARγ, although it occurs normally in mice deficient in Lpar1 and Lpar2 (89). In contrast, the application of genetic and pharmacologic tools in mice has suggested roles for LPA1 and LPA3 in vascular remodeling elicited by denudation injury (90) and flow-induced injury (29). The development of intimal hyperplasia likely results from effects of LPA on a combination of the processes described above, including recruitment of
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inflammatory and progenitor cells and stimulation of resident SMCs. In par-ticular, LPA may regulate the ability of CXCL12/SDF to recruit smooth muscle progenitor cells to sites of injury (90). As similar events also occur in athero-sclerotic lesions, these results position LPA as a potential mediator of the development of atherosclerosis. Of particular interest from the standpoint of understanding its proatherosclerotic effects, LPA causes reverse transmigra-tion and neutral lipid uptake in monocyte cells (91). Thus, the accumulation of LPA in atherosclerotic lesions could limit macrophage egress during plaque progression. Indeed, treatment of mice with LPA promotes monocyte adhesion to the endothelium, stimulates perivascular macrophage accumulation, and heightens atherosclerotic plaque burden in Apoe–/– mice in an LPA1- and LPA3-dependent manner (44). Minimally oxidized LDL also triggers monocyte adhe-sion to the endothelium and requires the activity of autotaxin, suggesting that LPC in the minimally oxidized LDL may be converted to LPA along the surface of endothelial cells to mediate proatherosclerotic effects. Finally, a pharmacologic inhibitor of LPA1 and LPA3 reduces the development of ath-erosclerosis in Western diet-fed Apoe–/– mice (44).
13.8.1. Experimental Models for Studying the Development of Intimal Hyperplasia and Atherosclerosis
LPA can be infused in the carotid artery of rats or mice by clamping the origin of the vessel and inserting a PE10 catheter in the external common carotid artery of an anesthetized animal. After washing with buffer, the artery is filled with buffer containing LPA (2.5 μM) for 1 hour at which time the external carotid artery is ligated and the common carotid allowed to reperfuse with circulating blood. At various times after the injury, the common carotid can be isolated for histologic and biochemical analysis. To denude the endothelium from the femoral artery, the mouse is anesthetized with inhaled isoflurane and placed in supine position to expose the femoral vessels near the inguinal liga-ment and distal to the epigastric artery. The distal portion of the femoral artery is encircled with a 9-0 nylon suture; a vascular clamp is placed proximally at the level of the inguinal ligament; and a 0.010-in. (0.25-mm) diameter angio-plasty guidewire (Abbott Vascular, Santa Clara, CA) is introduced into the arterial lumen through an arteriotomy made just distal to the suture. After release of the clamp, the guidewire is advanced to the level of the aortic bifur-cation and immediately pulled back; this process is repeated two additional times to denude the endothelium. Experimental atherosclerosis is typically examined in mice on a proatherosclerotic background of either Apoe–/– or Ldlr–/–. The relative advantages of each of these strains has been previously discussed (92). Typically, mice are placed on a Western diet (D12079B, Research Diets, New Brunswick, NJ) for periods of up to 3 months. Atherosclerotic lesions are evaluated by en face staining of the aorta with Oil Red O or by histologic analysis of serial sections taken through the aorta at the level of the aortic valve or of sections from the brachiocephalic artery.
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13.9. FUTURE DIRECTIONS
From experimental observations made in isolated cells and animal models, a picture emerges in which the bioactive lipid LPA is poised as a key modulator of a variety of pathologic responses in blood and vascular cells. In particular, LPA may serve as a critical nexus between proinflammatory and prothrom-botic events in the vasculature. However, a causative role for LPA in the disease processes, including the development and complications of atheroscle-rosis, remains to be established. To date, the strongest evidence for a role for LPA comes from studies in which exogenous, superphysiologic concentrations of LPA have been provided to cells or administered to animals. Establishing a role for endogenous LPA generated in blood or vessels in response to injury or inflammation as a causative factor in disease progression will require the use of experimental therapeutics and genetic manipulations in animals. The recent advances in understanding LPA receptor biology and the generation of small-molecule drugs that inhibit LPA production or antagonize its signaling effects will undoubtedly translate into improved insight of its role in the car-diovasculature. Ultimately, findings in experimental animal models will require confirmation in humans to determine if pharmacologic targeting of LPA sig-naling improves clinical outcomes or if LPA signaling pathways in individuals predict complications and/or response to therapy.
ACKNOWLEDGMENTS
This research was supported by grants from the NIH (to S.S.S. and A.J.M), a VA Merit Award (S.S.S.), and a scientist development grant from the Ameri-can Heart Association (M.P.). This material is the result of work supported with the resources and use of the facilities at the Lexington VA Medical Center.
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