Phospholipase D/phosphatidic acid signal transduction - Springer Link

Molecular and Cellular Biochemistry 234/235: 99–109, 2002. © 2002 Kluwer Academic Publishers. Printed in the Netherlands.

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Phospholipase D/phosphatidic acid signal transduction: Role and physiological significance in lung Rhett Cummings, Narasimham Parinandi, Lixin Wang, Peter Usatyuk and Viswanathan Natarajan Department of Medicine, Division of Pulmonary and Critical Care Medicine, Johns Hopkins University School of Medicine, Baltimore, MD, USA

Abstract Phospholipase D (PLD), a phospholipid phosphohydrolase, catalyzes the hydrolysis of phosphatidylcholine and other membrane phospholipids to phosphatidic acid (PA) and choline. PLD, ubiquitous in mammals, is a critical enzyme in intracellular signal transduction. PA generated by agonist- or reactive oxygen species (ROS)-mediated activation of the PLD1 and PLD2 isoforms can be subsequently converted to lysoPA (LPA) or diacylglycerol (DAG) by phospholipase A1/A2 or lipid phosphate phosphatases. In pulmonary epithelial and vascular endothelial cells, a wide variety of agonists stimulate PLD and involve Src kinases, p-38 mitogen activated protein kinase, calcium and small G proteins. PA derived from the PLD pathway has secondmessenger functions. In endothelial cells, PA regulates NAD[P]H oxidase activity and barrier function. In airway epithelial cells, sphingosine-1-phosphate and PA-induced IL-8 secretion and ERK1/2 phosphorylation is regulated by PA. PA can be metabolized to LPA and DAG, which function as first- and second-messengers, respectively. Signaling enzymes such as Raf 1, protein kinase Cζ and type I phosphatidylinositol-4-phosphate 5-kinase are also regulated by PA in mammalian cells. Thus, PA and its metabolic products play a central role in modulating endothelial and epithelial cell functions. (Mol Cell Biochem 234/235: 99–109, 2002) Key words: phospholipase D, phosphatidic acid, secretion, barrier function, endothelium, airway epithelium Abbreviations: PA – phosphatidic acid; PC – phosphatidylcholine; LPA – lysophosphatidic acid; PLD – phospholipase D; PLA2 – phospholipase A2; PLA1 – phospholipase A1; PLC – phospholipase C; PIP2 – phosphatidylinositol-4.5-bisphosphate; PI – phosphatidylinositol; PIP – phosphatidylinositol-4-phosphate; DAG – diacylglycerol; MG – monoacylglycerol; PKC – protein kinase C; LPP – lipid phosphate phosphatase; DAGK – diacylglycerol kinase; TPA – 12-0-tetradecanoyl phorbol-13-acetate; ECs – endothelial cells; IP3 – inositol-1,4,5-trisphosphate; IL-8 – interleukin-8; PDGFR – platelet-derived growth factor receptor; ERK – extracellular signal-regulated kinase; JNK – c-Jun N-terminal kinase; GST – glutathione-S-transferase; SMC – smooth muscle cell; MEK – mitogen- and extracellularly-activated protein kinase; MKK – mitogen-activated protein kinase kinase

Introduction Lipid derived second-messengers generated by the action of phospholipases on membrane associated phospholipids play an important role in signal transduction. Phospholipase D (E.C. 3.1.4.4; PLD), ubiquitously present in all mammalian cells and tissues, is recognized as a receptor-regulated signaling enzyme

that modulates many cellular functions. PLD is involved in membrane trafficking, exocytosis, secretion, phagocytosis and cytoskeletal reorganization. In the last decade, considerable progress has been made in understanding the regulation and role of PLD in various cellular functions. This review will focus on the regulation and physiological relevance of PLD activation in vascular endothelium and airway epithelial cells.

Address for offprints: V. Natarajan, Department of Pulmonary and Critical Care Medicine, The Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle, Room 4B.64, Baltimore, MD 21224, USA (E-mail: [email protected])

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Phospholipase D catalyzed hydrolysis of phosphatidylcholine Phospholipase D catalyzes the hydrolysis of phosphatidylcholine (PC), the major membrane phospholipid, to phosphatidic acid (PA) and choline [1, 2]. PA is a second-messenger that can be metabolized to other bioactive lipids, which include lysophosphatidic acid (LPA) and diacylglycerol (DAG) [3]. In mammalian cells, conversion of PA to LPA is catalyzed by phospholipase A1/A2 activity [4], while the formation of DAG from PA is regulated by lipid phosphate phosphatases (LPP) [5]. To date, three isoforms of LPP have been cloned and partially characterized [6]. LPA and DAG generation resulting from PLD activation are potential first- and second-messengers [7, 8]. DAG is a well characterized endogenous activator of protein kinase C (PKC) [9]. LPA binds to specific G-protein coupled receptors, LPA-1, -2 and -3 (former nomenclature: EDG-2, -4, and -7), with high affinity [10]. LPA can also be metabolized to 1- or 2-monoacylglycerol (1or 2-MG) and 2-arachidoylglycerol (2-AG) has recently been identified as a ligand for cannabinoid receptors (Fig. 1) [11, 12]. Furthermore, the choline released from PC via PLD activation can exhibit biological functions. Choline can be metabolized to form phosphocholine, a known regulator of cell proliferation [13]. Thus, hydrolysis of PC by PLD can give rise to PA, LPA, DAG, 2-AG and phosphocholine, all of which regulate key cellular functions. In addition to using

PC as a substrate, other membrane phospholipids such as phosphatidylethanolamine, phosphatidylserine and phosphatidylinositol, can also serve as substrates for PLD in cells [14].

PLD catalyzed transphosphatidylation reaction PLD is a phospholipid esterase and, as is common with other esterases, can catalyze a transphosphatidylation reaction utilizing short-chain primary alcohols as phosphatidyl-group acceptors [14]. Therefore, in the presence of methanol, ethanol, 1-butanol or 1-propanol, the PLD catalyzed transphosphatidylation reaction will generate the acidic lipids – phosphatidylmethanol, phosphatidylethanol, phosphatidylbutanol, or phosphatidylpropanol [15]. This reaction is very specific for primary alcohols as secondary and tertiary alcohols are not acceptors of the phosphatidyl-group. The phosphatidylalcohols generated by PLD, which are not a normal constituent of cells or biological membranes, are metabolically stable relative to PA. Thus, the formation of phosphatidylalcohols serves as a reliable and accurate measure of PLD activation in cultured mammalian cells and tissues (Fig. 2) [14, 15].

PLD isoforms and expression in mammalian cells Two mammalian PLD isoforms, PLD1 and PLD2, have been cloned and share 55% sequence homology [16–18]. The mammal, yeast and plant genes comprise a novel gene family that share limited but significant similarities of highly

Fig. 1. Phospholipase D catalyzed hydrolysis of phosphatidylcholine to phosphatidic acid, lysophosphatidic acid, diacylglycerol and monoacylglycerol.

Fig. 2. Phospholipase D catalyzed formation of phosphatidylalcohols by transphosphatidylation reaction.

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Fig. 3. Domain structure of PLD1 and PLD2 from mammals, yeast, plant, bacteria and virus.

conserved sequence motifs (Fig. 3). The PLD gene superfamily members share two conserved HKD domains (II and IV) that are required for catalytic activity as well as two other conserved regions, denoted I and III. The yeast and mammalian PLD1 and PLD2 also exhibit regions of conserved PX (phox homology) and PH (pleckstrin homology) domains, which are absent in plant and bacterial PLD genes. The PX and PH domains are unique because they are known to bind to D3-phosphorylated phosphoinositides and recruit actin-binding, scaffolding proteins and NADPH oxidase subcomponents (p40 phox and p47 phox) to the cytoskeleton [19, 20]. Mammalian PLD1 cDNA, first cloned from a HeLa cell cDNA library, is a 1074-amino acid (~120 KDa) protein while PLD2 is a 933-amino acid (106 KDa) protein. PLD1 has an additional 116-amino acid ‘loop region’ inserted immediately following the first HKD domain motif [21]. Most of the examined mammalian cells and tissues express PLD1 and PLD2 mRNA at different levels [22]. Quantitative studies on the levels of PLD1 and PLD2 proteins have been very difficult due the sensitivities required to detect the low levels of endogenous expression and lack of high affinity antibodies. In lung endothelial and airway epithelial cells,

expression of native PLD1 and PLD2 isoforms have been observed either in the total cell lysates, or after immunoprecipitation and Western blotting. There are limited studies that describe growth or differentiation factors regulating PLD1 and PLD2 at the transcriptional level. In HL-60 cells, differentiation leads to a dramatic upregulation of gene expression of both isoforms [23]. Vitamin D3 induces PLD1 expression during differentiation of murine keratinocytes [24]. PLD activity is elevated in human breast, renal and colon cancers, suggesting a role in tumorigenesis [25–29].

Agonist-mediated activation of PLD1 and PLD2 Although PLD was first described in plants in 1948 by Hannahan and Chaikoff [30], it took at least another 40 years to recognize external stimulation of PLD in animal tissues and mammalian cells [31, 32]. Bocckino et al. first described an increase in PA secretion in hepatocytes after stimulation with vasopressin [33]. Since then, neurotransmitters, growth factors, hormones, antibodies, bioactive lipids, calcium iono-

102 phores, phorbol esters and reactive oxygen species (ROS) have been shown to induce PLD activation [34–36]. Many of these external agonists act through G-protein coupled receptors or growth factor receptors and signals arising from receptor activation indirectly activates PLD. Most of the agonists that trigger PLD also stimulate phosphatidylinositol-4,5-bisphosphate (PIP2) specific PLC stimulation, leading to an increase in intracellular inositol-1,4,5-trisphosphate (IP3) and DAG. IP3 ligation of receptors present in the endoplasmic reticulum causes an increase in intracellular calcium [37] whereas DAG regulates some isotypes of protein kinase C (PKC) [9]. The phorbol ester, 12-0-tetradecanoyl phorbol 13acetate (TPA), potently and universally activates PLD in virtually all cell types, suggesting that PKC-mediated signaling occurs upstream of PLD [14]. It has been clearly demonstrated that in many cells, both agonist- and TPA-induced PLD activation is attenuated by inhibiting PKC with bisindoylmaleimide and calphostin C [38]. In addition to PKC, several other signaling mediators, including small G proteins, calcium, non-receptor tyrosine kinases and mitogen activated protein (MAP) kinases, have been implicated in PLD1 and PLD2 activation [34, 39, 40]. Outlining the multiple, complex mechanisms by which cell surface receptors regulate PLD is beyond the scope of this review. Figure 4 illustrates mechanisms of PLD activation in mammalian cells.

Reactive oxygen species-induced PLD activation Phagocytic leukocytes and the non-phagocytic vascular endothelium generate ROS that subsequently activate multiple signaling molecules [41, 42]. Studies have clearly demonstrated that hydrogen peroxide (H2O2), vanadate and pervanadate can enhance tyrosine phosphorylation of several proteins in neutrophils [43] and endothelial cells (ECs) [44]. H2O2 has been shown to activate PLD in endothelial cells [45, 46] and NIH3T3 fibroblasts [38]. Diperoxovanadate (DPV), 4-hydroxynonenal, fatty acid hydroperoxide and oxidized LDL produce increased PLD activity in endothelial and smooth muscle cells [48–50]. In neutrophils and HL-60 cells, treatment with formyl-methionyl-leucyl-phenylalanine (fMLP) resulted in increased protein tyrosine phosphorylation and PLD activation [51]. The NAD[P]H-mediated oxidative burst in neutrophils stimulated with fMLP suggests that NAD[P]H oxidase could be involved in enhancing protein tyrosine phosphorylation and PLD activation. ROS-induced PLD activation in ECs is insensitive to PKC inhibitors or down regulation of PKC by TPA [45–47]. Treatment of endothelial cells with tyrosine kinase inhibitors such as genistein, herbimycin or erbstatin partially attenuates ROSmediated PLD activation [40]. Addition of the protein tyrosine phosphatase inhibitors, vanadate, phenylarsine oxide or diamide, has been shown to enhance both tyrosine phosphorylation and PLD activity several-fold in ECs, further supporting the view that tyrosine phosphorylation plays a role in PLD activation [49, 52].

Regulation of PLD by Src kinases

Fig. 4. Schematic diagram of the regulation of agonist-induced phospholipase D activation. Binding of agonists such as thrombin, bradykinin or bioactive lipids to their respective receptor linked to a heterotrimeric Gprotein (G PLC or Gq), activates PLC or PLD respectively. Activation of PLC mediates hydrolysis of PIP2 to generate DAG and IP3. DAG is an endogenous activator of PKC while IP3 releases intracellular calcium from the endoplasmic reticulum. Activation of PKC also modulates PLD, catalyzing the hydrolysis of phosphatidylcholine (PC) to phosphatidic acid (PA), which can be metabolized to DAG. Interaction between the agonist and its receptor can also activate tyrosine kinases that regulate PLD.

Members of the Src family, p60 Src and p56 lyn, have been implicated as regulatory enzymes in thrombin- and IgE-induced PLD activation in platelets and RBL-2H3 mast cells [53–55]. The Src kinase inhibitors, PP-1 and PP-2, markedly attenuate DPV- and TPA-mediated PLD activation in ECs [40]. In addition, the transient expression of a Src dominantnegative mutant partially blocks DPV-induced PLD activation. Although Src kinase has failed to directly phosphorylate PLD1 or PLD2 in vitro, Src immunoprecipitates of control cells have revealed the presence of PLD1 and PLD2, indicating an association of PLD with Src kinase under basal conditions [40]. Exposure of ECs to DPV (5 µM) for 2 min enhances the association of PLD2, but not PLD1, with Src. In addition, the presence of Src has been observed in Western blots of immunoprecipitates of PLD1 and PLD2 isoforms [40]. The involvement of Src kinase in PLD activation has also been described in cells treated with angiotensin II [56], 1,25dihyroxy vitamin D3 [57], prolactin [58] and epinephrine [59]. The specific involvement of G protein-coupled signaling

103 seems to vary among the cell type studied. Angiotensin IImediated PLD activation in vascular smooth muscle cells requires coupling to pertussis toxin-insensitive G proteins [56]. Conversely, pertussis toxin markedly reduced 1,25-dihydroxy vitamin D3-induced PLD activity in CaCo-2 cells [57]. Furthermore, the PLD activation by both of these agonists is blocked by the addition of Rho neutralizing antibodies or C3 exoenzyme [57]. DPV-mediated PLD activation is insensitive to pertussis toxin and C3 exoenzyme in ECs (Natarajan, unpublished data). These observations provide evidence that Src kinase activation can transduce signals leading to PLD activation. Although current data supports the involvement of Src in the regulation of PLD activation, the precise mechanisms are not well understood. Activation of Src by an agonist or oxidant results in enhanced tyrosine phosphorylation of several intracellular proteins, including focal adhesion kinases (FAK) [60, 61], myosin light chain kinase [62], and cortactin [63]. Phosphorylation of FAK and Src results in the formation of complexes of Shc, Grb2 and Sos with a number of other scaffolding proteins [63]. ECs or HL-60 cells exposed to oxidants followed by analysis of PLD1 and PLD2 immunoprecipitates with anti-phosphotyrosine antibodies provide evidence that PLD1 and PLD2 could be tyrosine phosphorylated [40, 64]. In Swiss-3T3 cells, pervanadate enhances tyrosine phosphorylation of PLD which is constitutively associated with the platelet-derived growth factor receptor (PDGFR) [47]. Protein–protein interactions may explain a possible mechanism of PLD activation by Src. A similar protein–protein interaction between PKCα and PLD has been described to account for ATP-independent, but PKC-dependent activation of PLD [65]. Other possible targets of Src could include guanidine exchange factors for small G proteins that, upon tyrosine phosphorylation, translocate to the membrane and increase PLD activity [64].

kinase in vitro, the phosphorylation fails to modulate PLD activity. However, both PLD1 and PLD2 are phosphorylated by DPV in vivo. The mechanisms of p38 MAP kinase-mediated PLD activation remains unclear, but similar to Src, PLD1 and PLD2 interact with p38 MAP kinase as determined by coimmunoprecipitation and glutathione-S-transferase (GST)fusion protein pull down assays [39]. In contrast to ECs, PLD activation by fMLP is upstream to p38 MAP kinase in HL60 cells. In neutrophils, TNFα and GM-CSF activate p38 MAP kinase independently of PLD [66]. The role of ERK in PLD activation varies among different cell types. In neutrophils [67], PC-12 cells [68] and smooth muscle cells (SMCs) [69], ERK activation regulates PLD activity. In rat phenochromocytoma PC-12 cells, the H2O2induced PLD activation and MAP kinase phosphorylation is attenuated by PD 98059 [68]. Similarly, PD 98059 blocks fMLP-induced ERK phosphorylation and PLD stimulation in neutrophils [67]. Studies in rabbit aortic SMC demonstrated that norepinephrine-mediated PLD activation is attenuated by farnesyltransferase inhibitors and PD 98059, suggesting Rac/MAP kinase pathways in the regulation of PLD via a phosphorylation-dependent mechanism [68, 69]. On the other hand, DPV- and vasopressin-mediated PLD activation in ECs and A7r5 rat vascular SMC is not dependent on ERK [70]. The role of JNK in PLD activation in mammalian cell types is unknown. The involvement of Src and MAP kinase on agonist- or oxidant-induced PLD activation is diagramed in Fig. 5.

Regulation of PLD by mitogen activated protein kinases Studies from our laboratory and others have demonstrated that agonists and exogenous oxidants activate the MAP kinases, extracellular signal-regulated kinase (ERK), Jun Nterminal kinase (JNK), and p38 MAP kinase [39, 50]. We have demonstrated that DPV-induced p38 MAP kinase activation regulates PLD in ECs [39]. This conclusion was based on experiments using p38 MAP kinase inhibitors, SB203580 and SB202190, as well as transient transfection with a p38 dominant negative mutant, resulting in the mitigation of PLD activation by DPV but not by TPA. The mitogen and extracellularly activated protein kinase (MEK1/2) inhibitor, PD98059, has no effect on DPV-induced PLD activation. While both PLD1 and PLD2 are phosphorylated by p38 MAP

Fig. 5. Postulated signal transduction pathways of Src and p38 MAP kinase involved in phospholipase D activation. Agonists/oxidants can activate mitogen and extracellularly-activated protein kinase (MEK) via MEK-Kinase or p21 ras/Raf-1 involving scaffolding proteins, Son of sevenless (Sos) and growth factor receptor-bound (Grb2). Several mitogen activated protein kinase kinases (MKKs) have been identified which can phosphorylate either ERK, p38 MAPK or JNK. Oxidants can also activate the Src family of non-receptor kinases which regulate PLD by an unknown mechanism.

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Regulation of PLD1 and PLD2 in vitro Both isoforms of PLD catalyze hydrolysis of PC to PA. In vitro, they are selectively activated by different co-factors [14, 71]. Development of a detergent-free assay system for PLD established a requirement for PIP2 [71] in vitro. This polyphosphoinositide is an allosteric regulator of both PLD1 and PLD2. As mentioned, PLD1 and PLD2 have PX and PH domains that bind PIP2 [21]. Other domains present in both PLD1 and PLD2, ‘KR’ motifs, are rich in basic amino acids and also regulated by PIP2 binding. A number of small G proteins, Arf, Rho, and Cdc42, are co-factors for PLD1 but not for PLD2 [21]. In intact cells, there are several studies demonstrating a role for Arf, Rho, and Cdc 42 in agonist-mediated activation of PLD1 [72, 73]. PLD1 and PLD2 exhibit different sensitivity to detergents. PLD2 is insensitive to Triton X-100 whereas the same detergent inhibits PLD1 [14, 74]. Lysates from a variety of mammalian cells exhibit an oleate activated PLD, most likely PLD2 [74].

Modulation of PLD activity Although considerable progress has been made in the past decade in understanding the regulation of agonist-induced PLD activation, lack of specific inhibitors for PLD1 and PLD2 has slowed the progress of determining specific PLD isoform activation and isoform-specific cellular responses. Current experimental approaches that implicate PLD in a explicit physiologic response employ the use of primary vs. secondary or tertiary short chain alcohols. This approach has been widely utilized, but interpretation of the data requires incubation with low concentrations of alcohols and inclusion of secondary or tertiary alcohol as a negative control. A second experimental approach has been to add exogenous short chain dioctanoyl PA or dicaproyl PA to intact cell systems to evaluate the possible involvement of endogenously generated PA. Occasionally, investigators have used bacterial PLD preparations co-incubated with cells to generate plasmamembrane bound PA. Such studies, although useful, have the problem of possible protease contaminations in the PLD preparation. Thirdly, inhibitors of PA phosphatase such as propanolol have been frequently used to increase the intracellular accumulation of PA as well as distinguish the relative contributions of PA vs. DAG in a given physiological response. However, in some cell types, propanolol can stimulate PLD activity [75]. With the cloning of PLD1 and PLD2, availability of catalytically inactive mutants, and the utilization of antisense oligonucleotides, investigations using molecular reagents will provide for elegant approaches to study the role of PLD in cellular function.

Role of PLD in NAD[P]H oxidase activation Activated polymorphonuclear leukocytes (neutrophils), cells that exhibit an enhanced oxidative burst, have been employed as a model system to study PLD. Phagocytic particles, chemoattractants, and cytokines can induce PLD activation in neutrophils [76]. A correlation between PLD activation, PA accumulation and NAD[P]H oxidase activation in f-Met-Leu-Phe (fMLP) stimulated neutrophils has been observed [77]. In this study, the inclusion of primary alcohols attenuated the NAD[P]H oxidase mediated superoxide production, while propanolol potentiated the response. Addition of PA and DAG to cell lysates from neutrophils stimulated the NAD[P]H oxidase activity in vitro and when co-incubated with a primary alcohol, degranulation was blocked, suggesting an involvement of PLD activation in degranulation [78]. In human pulmonary artery ECs, hyperoxia-induced superoxide production is mediated by NAD[P]H oxidase and partially blocked by 1-butanol, but not by 3-butanol (Parinandi et al., manuscript in preparation). Over expression of PLD1 and PLD2 wild type adenoviral constructs in human pulmonary artery ECs enhances superoxide/ROS generation while catalytically inactive PLD1 (K898R) and PLD2 (K758R) mutants attenuate ROS production (Natarajan et al., unpublished data). Our observations suggest a unique involvement of PLD1 and PLD2 in NAD[P]H oxidase-mediated ROS production in ECs. Tyrosine kinases, PKC, MAP kinases, p-21 activated kinases and PA-dependent kinases may be involved in the activation of phagocytic and non-phagocytic NAD[P]H oxidase [76–80]. Several components of NAD[P]H oxidase are phosphorylated during cell activation and phagocytosis [81–83]. Recent studies show that p47 phox is phosphorylated by PKC ζ in vitro and that PKC ζ regulates fMLP-stimulated NAD[P]H oxidase in intact neutrophils [84–85]. The mechanism by which PA regulates NAD[P]H oxidase in ECs is unknown, but PA-dependent kinases have been demonstrated to phosphorylate p22 phox and p47 phox in cell free preparations from neutrophils [83]. Inhibitors of Src kinase, MAP kinases and PKC attenuate hyperoxia-induced superoxide production in human lung ECs (Parinandi et al., manuscript in preparation). Thus, the phosphorylation of NAD[P]H oxidase subcomponents at serine, threonine or tyrosine residues may regulate production of superoxide. Independent of DAG, PA can activate PKC ζ in neutrophils [86]. Based on these data, it is reasonable to consider that PA-dependent activation of PKC ζ regulates NAD[P]H oxidase by a phosphorylation dependent mechanism. Detailed studies are necessary to define the mechanism(s) of PA-dependent kinase activation of NAD[P]H oxidase and identify phosphorylation sites on the phox components.

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Role of PLD in the synthesis and release of matrix metalloproteases (MMPs) Matrix metalloproteases (MMPs) have been implicated in facilitating tumor metastasis by enhancing the degradation of basement membrane matrix proteins. In human cancer cells, laminin-induced MMP-2 secretion is blocked by primary alcohols and stimulated by exogenous PA [87, 88]. Phorbol ester-mediated secretion of MMP-9 in Swiss 3T3 cells is mitigated by 1-propanol while short chain PA mimic the phorbol ester response [89]. These results propose a PAdependent mechanism in MMP secretion. However, these studies do not distinguish whether PLD mediates transcription, translation or post-translational processes that result in MMP secretion.

Role of PLD in sphingosine-1-phosphate (S1P) induced interleukin 8 (IL-8) secretion Our laboratory has observed that activation of PLD is involved in sphingosine-1-phosphate (S1P) induced IL-8 secretion in the Beas-2B human bronchial epithelial cell line [90]. S1P, a potent bioactive, angiogenic sphingolipid, markedly enhances IL-8 secretion in a dose- and time-dependent fashion. S1P also activates PLD as demonstrated by labeled phosphatidylbutanol formation. The S1P-induced PLD activation and IL-8 release are blocked by pertussis toxin, indicating the involvement of a Gi protein-coupled receptor. By using 1-butanol as an acceptor of PA in the PLD catalyzed transphosphatidylation reaction, the S1P-mediated IL-8 secretion is attenuated by 60%, demonstrating that PLD-derived PA is part of the signaling pathway. This effect was confirmed using the PKC activator, TPA, as a positive control. Furthermore, both S1P- and TPA-induced PLD activation and IL-8 secretion are inhibited by pretreatment with the general PKC inhibitor, bisindolylmaleimide. Other studies in our lab have shown that infection of Beas2B cells with adenoviral constructs containing PLD1 or PLD2 wild type cDNA exhibit elevated S1P-induced PLD activation and enhanced ERK1/2 phosphorylation (Wang et al., manuscript in preparation). We observed an attenuation of S1P-mediated IL-8 secretion by PD98059, a MEK1/2 inhibitor. Interestingly, S1P-induced ERK1/2 phosphorylation is partially inhibited in the presence of 1-butanol. ERK1/2 activation can be dependent on Raf 1 [91], suggesting that in Beas-2B cells, Raf-1 may be a target of PLD-generated PA. Similar experiments by Parinandi et al. [92] demonstrate that exposure of Beas-2B cells to concentrated ambient particle (CAPs), constituents of urban air, increases IL-8 secre-

tion 4–8-fold over a 3 h period. CAPs also activate PLD prior to the increase in IL-8 secretion. The CAP-mediated IL-8 secretion is blocked by 1-butanol but not by 3-butanol, suggesting that this pathway leading to IL-8 secretion is also regulated by PLD-generated PA. Exposure of cells to short chain PA analogs can stimulate the secretion of IL-8 as well [92]. These data reveal a pro-inflammatory function of PLD activation in airway epithelial cells. By modulating IL-8 secretion, the activation of PLD may play a role in pulmonary inflammatory processes mediated by neutrophils.

Role of PLD in vascular endothelial barrier dysfunction The pulmonary vascular endothelium is critical for normal lung function by its action as a semi-selective barrier between plasma and the interstitium. Dysfunction of this barrier, a pathological characteristic of the adult respiratory distress syndrome, results in increases in oxygen requirements due to a decrease in lung compliance and altered gas exchange. ROS and reactive nitrogen intermediates generated in the vasculature by activated neutrophils or vascular cells have been implicated in the pathobiology of barrier dysfunction. The exact mechanisms that regulate ROS-induced EC barrier are unclear. Studies performed in our laboratory and others suggest that ROS-induced permeability alterations in the endothelium are modulated by signal transduction pathways involving calcium, protein kinases and phosphatases [93]. Exposure of ECs to exogenous PA enhanced albumin flux across the monolayer suggesting that intracellularly generated PA by the PLD pathway may exhibit a similar response [94]. ROS can regulate the activity of PKC, tyrosine kinases, non-receptor Src kinases and MAP kinases [50, 94]. ROS can also activate PLA2, PLC and PLD in ECs [41]. Indices of barrier dysfunction, transendothelial electrical resistance (TER) and albumin flux, are respectively decreased and increased in the presence of ROS [95]. The ROS-induced permeability changes in human and bovine pulmonary artery ECs is attenuated by 1-butanol (at a concentration of 0.05–0.1%), but not by 3-butanol (at a concentration of 0.05–0.1%). This effect of PLD-generated PA on barrier dysfunction is observed in ECs transiently transfected with catalytically inactive PLD1 and PLD2 mutants to attenuate ROS-induced TER alteration. Overexpression of wild type PLD1 and 2 cDNA in bovine pulmonary artery ECs enhances H2O2- and phorbol ester-mediated PLD activation and TER changes [95]. These studies clearly define a role for PA generated from PLD1 and PLD2 in the modulation of EC barrier function. The signaling pathways downstream of PLD leading to EC permeability changes have not been clearly defined. However, PA can directly activate PKC ζ, alter the actin cytoskel-

106 eton and modify the actomyosin contractile apparatus [96, 97]. PIP2 and phosphatidylinositol-3,4,5-trisphosphate are other important regulators of the actin cytoskeleton. PA has been shown to activate PI-4-kinase in vitro and type I PIP-5kinase in vivo [98]. Subsequently, the PA-mediated activation of kinases can alter intracellular levels of PIP2, in turn modulating interactions between actin and actin binding proteins such as vinculin and filamin [73]. PIP2 can also stimulate PLD activity [17], thereby amplifying PA production and PA-dependent kinases. Furthermore, PIP2 enhances interaction between proteins that contain PH and PX domains [19, 20].

Other PLD mediated cellular functions PLD activation has been implicated in arachidonic acid release and prostaglandin synthesis [99], stress fiber formation [100], phagocytosis [76] and assembly of very low density lipoproteins [101]. PLD also plays a role in the membrane trafficking of insulin-mediated GLUT4 glucose transporter from intracellular compartments to the plasma membrane [102].

Conclusions The activation of PLD by a wide range of agonists has been recognized as a key component in signal transduction pathways in mammalian cells. The two isoforms of PLD present in mammalian cells and tissues catalyze the hydrolysis of PC to PA and choline. Regulation of the isoforms has several common features as well as distinct differences. PIP2 is a co-factor for both isoforms in vitro and in vivo. PLD1 and PLD2 are regulated by small G proteins, PKC and calcium. PLD1 is inhibited in vitro by detergents whereas PLD2 is not affected. Generation of PA by the PLD pathway is central in several physiological functions such as vesicular trafficking, secretion, phagocytosis and barrier alterations. The PA-dependent kinases, Raf1, PKC ζ and type I PIP-5-kinase, are regulated by the activation of PLD. Whether PA directly activates these kinases or regulates via intermediary proteins remains somewhat unclear. PLD-generated PA can be metabolized to LPA by PLA1/A2 or to DAG by lipid phosphate phosphatases (Fig. 6). LPA, another effective bioactive lipid, transduces signals through G protein-coupled receptors present in multiple mammalian cells. LPA is secreted in high quantities by stimulated platelets, neutrophils, mast cells, adipocytes and certain cancer cells. DAG generated from PA can acti-

Fig. 6. Postulated cross talk between phospholipase C, phospholipase D, diacylglycerol kinase and type 1 PIP-5-kinase signaling pathways. Agonist-mediated activation of PLC results in the hydrolysis of PIP2 to DAG and IP3 which modulate PKC and calcium release, respectively. Activation of PKC and intracellular calcium changes stimulate PLD, hydrolyzing membrane phosphatidylcholine (PC) to phosphatidic acid (PA). PA is a second-messenger and can be further metabolized to LPA or DAG by phospholipase A1/phospholipase A2 or lipid phosphate phosphatase activity, respectively. DAG can be recycled to PA by DAG kinase(s) present in the cell. PA can also activate type 1 PIP-5-kinase to generate PIP2. PIP2 is a co-factor for PLD1 that regulates actin cytoskeletal reorganisation. Thus, activation of PLC and PLD can generate lipid metabolites that modulate signal transduction pathways in mammalian cells.

vate PKC isoforms or recycled to PA by DAG kinase. Hence, agonist-induced PLD activation results in the generation of at least three bioactive lipids, PA, LPA, and DAG, having first- and second-messenger functions in mammalian cells. Furthermore, PIP2-specific phospholipase C (PLC) that is activated very rapidly in response to agonist and plays a critical role in PLD stimulation. The DAG and IP3 generated by PIP2-specific PLC regulate PKC and calcium, key intracellular signaling molecules. PIP2-specific PLC, PC-specific PLD1 and PLD2, PA-dependent PKC ζ and PIP-5-kinase, and DAG-kinase can cross talk via protein–protein interactions and co-operate to produce lipid-derived second-messengers that control various cellular responses (Fig. 6). Furthermore, the fatty acid composition at the sn-1 and sn-2 position of the various lipid metabolites may dictate specificity, intensity and duration of the signals generated intracellularly. Development of specific inhibitors and the use of molecular reagents for PLD, PLC, PIP-5-kinase and DAG kinase in the coming years will help to further elucidate the role of lipid-derived second-messengers mediating specific cellular responses to environmental stimuli.

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Acknowledgements This work was partly supported by NIH grants HL47671, HL57260 and HL58064. We wish to thank Dr. Andrew Morris for providing us with wild type PLD1b, PLD2, and catalytically inactive mutants.

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