Communication

Participation of Reactive Oxygen Species in the Lysophosphatidic Acid-stimulated Mitogenactivated Protein Kinase Kinase Activation Pathway*
(Received for publication, August 31, 1995, and in revised form, October 5, 1995) Quanlu Chen, Nancy Olashaw, and Jie Wu** From the Molecular Oncology Program and the Cell Biology Program, H. Lee Moffitt Cancer Center and Research Institute and the Department of Medical Microbiology and Immunology and the Department of Anatomy, University of South Florida, Tampa, Florida 33612
THE JOURNAL OF BIOLOGICAL CHEMISTRY Vol. 270, No. 48, Issue of December 1, pp. 28499 28502, by The American Society for Biochemistry and Molecular Biology, Inc. Printed in U.S.A.
Recent evidence suggests that reactive oxygen species (ROS) may function as second messengers in intracellular signal transduction pathways. We explored the possibility that ROS were involved in lysophosphatidic acid (LPA)-induced mitogen-activated protein (MAP) kinase signaling pathway in HeLa cells. Antioxidant N-acetylcysteine inhibited the LPA-stimulated MAP kinase kinase activity. Direct exposure of HeLa cells to hydrogen peroxide resulted in a concentration- and time-dependent activation of MAP kinase kinase. Inhibition of catalase with aminotriazole enhanced the effect of LPA on induction of MAP kinase kinase. Further, LPA stimulated ROS production in HeLa cells. These findings suggest that ROS participate in the LPA-elicited MAP kinase signaling pathway.
. Reactive oxygen species (ROS),1 such as superoxide (O2), hydrogen peroxide (H2O2), and hydroxyl radical (OH), are potent microbicidal agents, but excess ROS can also cause oxidative damage to macromolecules of host cell (1). Previous studies have shown that elevated levels of ROS could trigger intracellular signaling transduction pathways that may mediate cellular protective responses (2 4). In addition to their roles in inflammatory and pathological processes, increasing evidence suggests that ROS may function as second messengers in cytokine (interleukin-1 and tumor necrosis factor ) and some growth factor signal transduction pathways that regulate transcription factors such as NF-B and AP-1 (5, 6). Lysophosphatidic acid (LPA) is released by activated platelets and is thought to be responsible for much of the activity in serum that promotes cell growth and adhesion (7, 8). LPA
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. ** To whom correspondence should be addressed: H. Lee Moffitt Cancer Center and Research Institute, MDC Box 44, 12902 Magnolia Dr., Tampa, FL 33612. Tel.: 813-979-6713; Fax: 813-979-3893. 1 The abbreviations used are: ROS, reactive oxygen species; MAP kinase, mitogen-activated protein kinase; MKK1/2, MAP kinase kinase 1 and MAP kinase kinase 2; BSA, bovine serum albumin; LPA, lysophosphatidic acid; DMEM, Dulbeccos modified Eagles medium; PBS, phosphate-buffered saline; FACS, fluorescence-activated cell sorter; EGF, epidermal growth factor.
elicits its biological responses through a putative receptor that is coupled to heterotrimeric G-proteins (9). Several proximal signaling events are known to be evoked by LPA, including phosphoinositide hydrolysis and Ca2 mobilization, release of arachidonic acid, inhibition of adenylate cyclase, and induction of protein tyrosine phosphorylation (10, 11). It is likely that some of these signaling events cross-interact to induce synergistic responses. LPA rapidly activates the mitogen-activated protein (MAP) kinase pathway (1114). MAP kinases are serine/threonineprotein kinases regulated by dual tyrosine and threonine phosphorylation. Three subfamilies of MAP kinases, MAPK, JNK, and HOG, have been cloned (for reviews, see Refs. 1517). The 42-kDa MAP kinase (p42mapk) also called extracellular signalregulated kinase 2 (ERK2)) and 44-kDa MAP kinase (p44mapk, ERK1) are phosphorylated and activated by highly specific MAP kinase kinase 1 and MAP kinase kinase 2 (MKK1/2) (18). For simplicity, p42mapk and p44mapk will be referred to as MAP kinase in this report. MAP kinase has been shown to play a pivotal role in cell proliferation and differentiation (19). It has been shown that the LPA-induced MAP kinase activation is sensitive to pertussis toxin inhibition, indicating a critical role of a pertussis toxin-sensitive Gi-protein (11, 12, 14). However, these data did not exclude the contribution of other signaling events to the LPA-induced MAP kinase activation. LPA triggers a biphasic arachidonic acid release in HeLa cells.2 The first phase of the LPA-induced arachidonic acid release precedes the activation of MAP kinase kinase.2 Arachidonic acid is known to give rise to ROS through its subsequent metabolism (20) and activation of NADPH oxidase (21), prompting us to investigate the possible involvement of ROS in the LPA-stimulated MAP kinase activation pathway. We present evidence here that demonstrates the involvement of ROS in the LPAinduced MAP kinase signaling pathway.

EXPERIMENTAL PROCEDURES

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MaterialsN-Acetylcysteine was from Fluka. H2O2 was from Fisher Scientific. Lysophosphatidic acid (1-oleoyl), aminotriazole, and fatty acid- and globulin-free bovine serum albumin (BSA) were from Sigma. Dihydrorhodamine 123 was from Molecular Probes. The kinase-defective mutant of p42mapk (K52R) was prepared as described previously (22). Cell CultureHeLa cells were grown at 37 C in Dulbeccos modified Eagles medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum in an atmosphere of 7% CO2. Confluent cells were serum-starved for h in phenol red-free DMEM containing 25 mM Hepes and 0.1% BSA (fatty acid- and globulin-free). Serum-starved cells were washed twice with DMEM (phenol red-free, plus 25 mM Hepes) and incubated in DMEM (phenol red-free, plus 25 mM Hepes) for 90 min before use. MAP Kinase Kinase AssayTotal activity of MAP kinase kinase 1 and MAP kinase kinase 2 (MKK1/2) was determined by the following procedure, modified from that described previously (23). After stimulation, cells were chilled on ice and washed with cold phosphate-buffered saline (PBS). Each 6-cm plate of cells was lysed in 0.6 ml of cold lysis buffer (10 mM Tris acetate, 100 mM NaCl, 1 mM dithiothreitol, 40 mM -glycerophosphate, 1.5 mM EGTA, 0.5 mM EDTA, 25 mM NaF, 1 mM sodium pyrophosphate, 0.5 mM sodium orthovanade, 20 mM 4-nitrophenyl phosphate, 1% Triton X-100, 1 mM benzamidine, 2 g/ml aprotinin, 2 g/ml leupeptin, 1 g/ml pepstatin, and 0.1 mg/ml phenylmethylsulfonyl fluoride, pH 7.5 at 25 C). Cell lysates were centrifuged (4 C) at 16,000 g for 15 min. Protein concentrations of cell lysate supernatants were determined. Equal portions (2 g of protein) of cell lysate
Q. Chen and J. Wu, unpublished data.
Involvement of ROS in Lysophosphatidic Acid Signaling
FIG. 1. Inhibition by N-acetylcysteine on MKK1/2 activation. Serum-starved HeLa cells were treated with or without N-acetylcysteine (NAC, 30 mM) for 90 min, and left unstimulated () or stimulated with lysophosphatidic acid (20 M) or EGF (25 ng/ml) for 5 min. MKK1/2 activity was determined using a kinase-defective p42mapk (K52R) as substrate. The kinase reaction was carried out at 30 C for 15 min. Arrow, K52R band. supernatants were mixed with KR kinase assay mixture (20 mM Hepes, pH 7.5, 10 mM MgCl2, 1 mM dithiothreitol, 10 mM 4-nitrophenyl phosphate, 40 M [-32P]ATP (5000 cpm/pmol), and 2.5 g of K52R). The reaction mixtures were incubated at 30 C for 12 min or as indicated in the figure legends. The kinase reaction was stopped by addition of SDS sample buffer and heat denaturation. Phosphorylation of K52R was analyzed by autoradiography and phosphoimaging after electrophoresis on 10% SDS-polyacrylamide gels. Detection of Intracellular H2O2Intracellular levels of H2O2 were analyzed by fluorescence-activated cell sorting (FACS) using dihydrorhodamine 123 as a probe (6, 24). Experiments were performed under dim light. Confluent, serum-deprived HeLa cells were incubated in DMEM (phenol red-free, plus 25 mM Hepes) containing 10 M dihydrorhodamine 123 for 20 min. Cells were treated with aminotriazole (50 mM), LPA (40 M), or BSA, then chilled on ice and washed with cold PBS. Washed cells were detached from culture plates by trypsin digestion. The activity of trypsin was quenched with 0.05% BSA (fatty acidand globulin-free) in PBS. Following PBS wash, cells were fixed in 1% paraformaldehyde. The fluorescence intensities of rhodamine 123 of 10,000 cells from each sample were analyzed by flow cytometry using a FACScan flow cytometer equipped with an air-cooled argon laser (Becton Dickinson).

RESULTS AND DISCUSSION

FIG. 2. Activation of MKK1/2 by H2O2. Serum-deprived HeLa cells were treated without (ATZ) or with 50 mM aminotriazole (ATZ) for 60 min followed by stimulation for 5 min with different concentrations of H2O2 as indicated (A) or with 1 mM H2O2 in the absence of aminotriazole for the indicated time (B). Total activity of MKK1/2 was determined. The phosphorylation of K52R was quantitated with a PhosphorImager (Molecular Dynamics) after SDS-gel electrophoresis. The basal activity (21 pmol/min/mg) of MKK1/2 in unstimulated cells was arbitrarily set as 1 unit.
In all cell types examined, p42mapk is specifically phosphorylated and activated by dual specificity MKK1 and MKK2 (MKK1/2). Total activity of MKK1/2 in the cells represents a valid measurement for the activation state of the MAP kinase pathway in the rapid activation phase (14, 23). As illustrated in Fig. 1, LPA and EGF markedly stimulated the MKK1/2 activity in HeLa cells. To test for the possible involvement of ROS in MKK1/2 activation induced by LPA, we examined the effect of antioxidant N-acetylcysteine on MKK1/2 activation. N-Acetylcysteine directly scavenges ROS and also increases the intracellular levels of reduced glutathione (GSH). GSH is a hydroxyl radical scavenger and a substrate of glutathione peroxidase which degrades H2O2. N-Acetylcysteine has been used extensively to study the role of ROS in signaling pathways (6, 25 27). An inhibition by N-acetylcysteine can be taken as an indication of the involvement of ROS. The LPA-stimulated MKK1/2 activity was inhibited by 82 4% (average of two experiments range) in cells pretreated with N-acetylcysteine (30 mM), suggesting that ROS are involved in the LPA-induced MKK1/2 activation (Fig. 1). A similar result was obtained in Rat-1 cells. A lesser, but statistically significant, attenuation (38 8% in two experiments) by N-acetylcysteine of the EGF-stimulated MKK1/2 activity was also observed (Fig. 1), but was not investigated further in the current study. To verify that the inhibitory effect of N-acetylcysteine is attributable to its ability to scavenge ROS, we examined the effects of two other ROS scavengers, dimethyl sulfoxide and ascorbic acid, on the LPA-stimulated MKK1/2 activity. Di-
methyl sulfoxide is an effective hydroxyl radical scavenger (28). Ascorbic acid blocks free radical chain reaction, but may also directly remove hydroxyl radical (29). HeLa cells were pretreated with ascorbic acid (100 M, 60 min) or dimethyl sulfoxide (4%, 20 min) and stimulated with LPA (10 M, 5 min). The LPA-stimulated MKK1/2 activity was inhibited by 88 7% by dimethyl sulfoxide and 38 1% by ascorbic acid. If ROS are the signaling molecules that mediate the LPAinduced MKK1/2 activation, then an increase in intracellular concentrations of ROS would be expected to mimic the effect of LPA on MKK1/2 activation. H2O2 is the product of superoxide dismutases and several oxidases in the cells. Thus, cells that produce superoxide would also generate H2O2. In contrast to superoxide, H2O2 can diffuse across the membrane and give rise to the highly reactive hydroxyl radical. H2O2 has been widely used to assess the role of ROS in cells. To test whether H2O2 directly added to the cells can activate MKK1/2, HeLa cells were treated with 0.mM H2O2 for 5 min or with 1 mM H2O2 for 2.530 min, and the MKK1/2 activity was determined. Fig. 2 shows that H2O2 caused a concentration- and time-dependent activation of MKK1/2 in HeLa cells. Thus, H2O2 alone is sufficient to induce MKK1/2 activation. The maximal activity of MKK1/2 induced by H2O2 in HeLa cells was detected approximately 5 min after treatment. Thus, the kinetics of H2O2 induction is similar to that of MKK1/2 activation induced by phospholipids and growth factors (13, 30). However, 2 mM H2O2, a concentration of H2O2 that cannot be achieved in HeLa cells by LPA (data not shown), is required to induce MKK1/2 activation to a similar magnitude (approximately 20-fold) as 20 M LPA. Several possibilities exist that may account for the requirement of high concentrations of H2O2. First, the LPA-induced ROS may be generated at a site that is more proximal to the

FIG. 3. Effect of aminotriazole on the lysophosphatidic acidstimulated MKK1/2 activation. MKK1/2 activity was determined in HeLa cells treated with or without a catalase inhibitor aminotriazole (50 mM, 60 min) and stimulated for 5 min with H2O2 (1 mM) or lysophosphatidic acid (20 M). The kinase reaction was carried at 30 C for 10 min. The data represent the average and range of two experiments.
FIG. 4. Relative levels of intracellular H2O2 in HeLa cells. Serum-starved HeLa cells were incubated for 20 min with 10 M dihydrorhodamine 123, followed by a 10-min incubation with either 50 mM aminotriazole plus BSA (B), 50 mM aminotriazole plus 30 M LPA (C), or an equivalent volume of BSA (A). The fluorescence intensities of 10,000 cells were analyzed.
target, whereas the external added H2O2 diffuses indiscriminately. Second, HeLa cells may contain relatively high catalase activity, and, thus, high concentrations of H2O2 are required to offset the catalase activity. In fact, inhibition of catalase by preincubation of HeLa cells with catalase inhibitor aminotriazole (26) prior to the addition of H2O2 resulted in a marked shift of the H2O2 dose-response curve to the left (Fig. 2A). However, even in the presence of aminotriazole, greater than 0.5 mM H2O2 was still required to activate MKK1/2 to a similar extent as that induced by 20 M LPA. Finally, it is likely that one or more signaling events besides production of ROS are critical for the induction of MKK1/2 activation by LPA, and ROS may function as only one of the parallel signaling intermediates. Thus, although H2O2 alone at low concentrations (0.5 mM) has a marginal effect on MKK1/2 activation, H2O2 and the derived radicals may have a greater effect in the presence of other LPA-induced signaling intermediates because of synergism. To further confirm the involvement of ROS, we treated HeLa cells with or without the catalase inhibitor aminotriazole prior to LPA stimulation. If ROS participate in the LPA-stimulated MKK1/2 activation pathway, inhibition of catalase would potentially augment the response to LPA. A 4.5-fold increase in H2O2-induced MKK1/2 activity was observed when HeLa cells were pretreated with aminotriazole (50 mM, 60 min), demonstrating the effectiveness of the catalase inhibitor (Fig. 3, see also Fig. 2A). In cells pretreated with aminotriazole, the LPAstimulated MKK1/2 activity was 1.9-fold that of cells without aminotriazole pretreatment (36.3- and 18.8-fold above basal, respectively) (Fig. 3). Thus, decreasing the catalase activity effectively enhances the cellular response to LPA, indicating the involvement of ROS. For ROS to fulfill the role of signaling intermediates for LPA, LPA must be able to induce the production of ROS. As described above, LPA rapidly liberates arachidonic acid in HeLa cells. Arachidonic acid is known to generate ROS. Tumor necrosis factor and interleukin-1, both of which are known to utilize ROS as signaling intermediates, also stimulate the release of arachidonic acid (31). However, other routes of ROS generation are not excluded. We measured the relative concentrations of H2O2 in HeLa cells using dihydrorhodamine 123 and fluorescence-activated cell sorting (FACS) (6, 24). Dihydrorhodamine 123 is oxidized to membrane-impermeable, fluorescent rhodamine 123 in the presence of H2O2 and possibly ROS derived from it (24). To minimize the loss of H2O2, aminotriazole was also added to the media. Incubation of HeLa cells with aminotriazole resulted in a time-dependent increase in fluorescence intensity (Fig. 4 and data not shown). A small

increase of the fluorescence intensity induced by LPA was detectable at the earliest time (5 min) examined, but more consistent data were obtained if cells were stimulated for 10 min. In two duplicated experiments, cells treated for 10 min with aminotriazole (50 mM) plus BSA had an average 25% increase in mean fluorescence intensity of rhodamine 123 compared with BSA-treated cells (Fig. 4). An additional 22% increase in mean fluorescence intensity was detected in cells treated for 10 min with LPA (30 M) plus aminotriazole (50 mM) (Fig. 4). Thus, LPA is capable of generating ROS in HeLa cells. In summary, data presented in this study show that ROS are involved in the LPA-induced MAP kinase kinase activation and LPA can stimulate the production of ROS in HeLa cells. Additional experiments using a p42mapk immune complex kinase assay (14) showed that N-acetylcysteine partially inhibited the LPA-stimulated p42mapk activation, H2O2 stimulated p42mapk activity in HeLa and NIH 3T3 cells, and aminotriazole enhanced the effect of LPA on p42mapk activation (data not shown). Previous studies have shown that ROS are involved in the tumor necrosis factor -stimulated NF-B activity (5) and the basic fibroblast growth factor-induced c-fos expression (6). Other data that we have obtained showed that N-acetylcysteine also inhibited the LPA-stimulated NF-B and AP-1 DNA binding activities in HeLa cells.3 Thus, ROS appear to function as signaling intermediates of LPA and mediate a branch of the LPA signaling pathways. Our findings lend support to the emerging concept that ROS can function as physiological signaling intermediates. It will be interesting to examine whether ROS also participate in the signaling pathways of other phospholipids, such as platelet-activating factor (32) and sphingosine 1-phosphate (33). Clearly, further investigation of the mechanisms by which LPA increases the intracellular levels of ROS and ROS relay the cellular regulatory signals is warranted.
AcknowledgmentsWe thank Christine OConnell for the FACS analysis and Drs. Warren J. Pledger and W. Douglas Cress for critical reading of the manuscript.

REFERENCES

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5.5. LPA synergizes with different platelet stimuli in inducing platelet aggregation in washed platelets and whole blood....51 5.6. Role of LPA produced by platelets in thrombin- or collagen- stimulated platelet aggregation....55 5.7. LPA- induced platelet-monocyte interaction...56 5.7.1. Mechanism of LPA-induced platelet-monocyte aggregate formation..57 5.7.2. Specific desensitization of the LPA-receptor mediated platelet- monocyte aggregate formation in whole blood...60 6. Discussion.....61 6.1. LPA induced shape change in washed platelets, PRP, and whole blood..61 6.2. Platelet aggregation in washed platelets, PRP, and whole blood..62 6.2.1.Difference in aggregation using different isolation procedures of washed platelets.62 6.2.2. Aggregation in PRP and whole blood...63 6.2.3. Activation by different LPA species...64 6.2.4. LPA-induced aggregation in blood and PRP- independence of the anticoagulant.65 6.2.5. Synegistic interaction of LPA with serotonin, epinephrine and ADP.66 6.2.6. LPA is not a positive feed-back mediator of platelet activation..67 6.2.7. Perspective: Preventing LPA-induced platelet aggregation...68 6.3. LPA- induced platelet-monocyte adhesion in whole blood..68 6.3.1. Perspective: Prevention of LPA-induced platelet-monocyte aggregate formation.70 7. Summary....71 8. Zusammenfassung.....73 9. Reference List.....75 Acknowledgements....88 List of publications....89 Curriculum vitae.....90

Abbreviations

M -AA3P5P ADP AR-C69931 ASS ATP P2Y1-receptor antagonist adenosine- diphosphate P2Y12- receptor antagonist acetyl- salicic acid adenosine- triphosphate micomolar

bovine serum albumin

CD COX-1
cluster or differentiation, a cell surface marker, i.e. CD41 cyclooxygenase 1

DGPP DMSO

diacylglycerolpyrophosphate dimethyl-sulfoxide

EC EDG EDTA

effective concentration endothelial differentiation gene (former description for LPA- /S1P- receptors) ethylene- diamine- tetra- acetic acid

Abbreviations -G-

membrane glycoprotein, i.e. GPIIb-IIIa = integrin IIb3

interleukin 6

LPA LPAAT LPC LPP Lyso-PLD
lysophosphatic acid LPA-acetyltransferase lysophosphatidylcholine lipidphosphate- phosphohydrolase lysophospholipase D (formerly autotaxin)

M MAG mM MMP MRS2179

molar monoacylglycerol millimolar matrix-metalloproteinase P2Y1- receptor antagonist

NATyrPA NASerPA NFB

N-acyl-Tyrosine- phosphoic acid N-acyl-Serine- phosphoric acid a transcriptionsfactor

Abbreviations -O-

Ox-LDL
oxidized- low-density- lipoprotein
P2Y1/12 PA PAF PBS PC PCR PDGF PLA PLD PPP PPAR PRP PS
ADP-receptors phosphatidic acid platelet activating factor phosphate buffered saline phosphatidylcholine polymerase-chain -reaction platelet derived growth factor phospholipase A phospholipase D platelet poor plasma peroxisome proliferator-activated receptor (a transcription factor) platelet rich plasma phosphatidylserine

Intracellular LPA as a precursor in glycerolipid synthesis In the chain of reactions in glycerolipid synthesis LPA formation can be catalyzed by the enzyme, glycerophosphate acyltransferase (GPAT), located in both endoplasmatic reticulum and mitochondria. GPAT acylates glycerol-3 phosphate into LPA as an intermediate product before LPA is then acylated by monoacylglycerolphosphate acyltransferase (MGAT) into phosphatidic acid (PA), the precursor of all glycerolipids (Haldar and Vancura, 1992). An alternative pathway is the reduction of acyl dihydroxyacetone phosphate (acyl DHAP) in peroxysomes contributing to LPA formation in pancreatic islets exposed to high glucose concentrations (Dunlop and Larkins, 1985). Further LPA can be synthesized by the action of monoacylgycerol kinase (MAG-kinase) on monoacylglycerol, as an important precursor of phosphatidylinositol synthesis(Simpson et al., 1991). MAG kinase has also been proposed to be involved in the formation of arachidonoyl-LPA in platelets (Gerrard and Robinson, 1989). However, whether LPA, produced during glycerolipid synthesis, can accumulate and contribute to the extracellular release of phospholipids is not yet resolved.
Extracellular LPA Generation by phospholipid hydrolysis Hydrolysis of fatty acids at the sn-1 position by phospholipase A1 (PLA1) or at the sn-2 position by phospholipase A2 (PLA2) generates LPA from PA. Platelets and to a smaller degree red blood cells, have been identified as sources of LPA in blood (Eichholtz et al., 1993). PA was found to be rapidly generated in thombin-stimulated platelets and could subsequently be converted to LPA via PLA enzymes (Lapetina et al., 1981a, b). LPA generated through this mechanism appears within 15 min following thrombin stimulation and constitutes only a minor portions, estimated 10%, of LPA detected in serum (Aoki et al., 2002; Gaits et al., 1997; Sano et al., 2002). A larger proportion of LPA has been shown to be produced in plasma by platelets releasing PLA1 and PLA2 that generate a de novo pool of lysophospholipids primarily lysophosphatidylcholine (LPC) from Phosphaditylcholine (PC) in plasma and membrane phospholipids (Sano et al.,
2002). LPC can then be further metabolized by the enzyme lysophospholipase D (lyso-PLD) identical to the tumor cell motility stimulating protein known as autotaxin (Tokumura et al., 2002) (See Fig 2.2 below). Autotaxin belongs to the family nucleotide pyrophosphatase/ phosphodiesterases and promotes tumor cell motility, progression, metastasis, and angiogenesis via a pertussis toxin-sensitive mechanism (Lee et al., 2002b; Nam et al., 2001; Stracke et al., 1997). These activities have also been described for LPA (Imamura et al., 1993; Imamura et al., 1996). LPA production due to lyso-PLD activity in human whole blood ex vivo has been described as fairly rapidly and amounts to about 1.2M within 1 h (starting from approximately 130 nM) (Baker et al., 2001). Sano et al have used activated plasma and showed a 20-fold increase within the first hour (Sano et al., 2002). However, this increase is not due to an increase in Lyso-PLD activity (Aoki et al., 2002; Sano et al., 2002), but has been proposed to be caused by PLA1, PLA2, capable of not only generating lysophosphatidylcholine (LPC), but also lysophosphatidylserine (LPS), and lysophosphatidylethanolamine (LPE) from plasma PC, PS, and PE. An important source of LPC, being the most abundant lysophospholipid (125-150M) in plasma, is the enzyme lecithincholesterol acyltransferase (LCAT) (Croset et al., 2000; Tokumura et al., 1999).

Fig 2.3 Schematic representation of domains of plasma gelsolin and their functions. Amino acid positions are numbered as in human plasma gelsolin and segmental boundaries are based on the structural definitions defined by the gelsolin crystal. Actin, PIP2, and Ca2+-binding segments are shown. Gelsolin domains are circled (Sun et al., 1999).
Extracellular plasma gelsolin is present at concentrations of 100-250g/ml (1.2- 2.9 M) and binds LPA with high affinity close to that of LPA receptors (Kd 6 nM) and exceeding that of albumin (Kd 360 nM). Extracellular gelsolin may function as a high affinity plasma carrier (Kd of 32 nM), which protects a portion of circulating LPA from biodegradation and prevents LPA from binding to numerous possible low affinity proteins (Goetzl et al., 2000). Gelsolin concentrations of less than 10% of the plasma concentration were shown to increase LPA activity, whereas 20% or higher concentrations of those in plasma diminished biological effects of LPA in rat cardiac myocytes (Goetzl et al., 2000). In the context of this work, only a potential effect of gelsolin on the outside cell surface may play a role.
2.3.5. Platelet shape change and aggregation induced by LPA
The first report on platelet aggregation induced by LPA was done in 1979 by Schumacher et al., who discovered that in plasma, incubated at 36C for 18-24 hours, a factor developed, which on intravenous injection in cats evoked platelet aggregation followed by an increase in pulmonary vascular resistance (Schumacher et al., 1979). They also provided evidence that PA and LPA, are the active components causing aggregation of human and feline platelets. Of interest is that LPA was unable to induce aggregation in dogs, rabbits, pigs, and rats (Mauco et al., 1978; Schumacher et al., 1979). LPA (Oleoyl-LPA or 18:1) is a potent activator of platelets and induces shape change of human isolated platelets at very low concentrations (EC50 = 18 nM) via remodelling of the actin cytoskeleton, which is dependent on G12/13- mediated activation of the small GTP- binding protein, Rho and subsequently Rho kinase (Bauer et al., 1999; Retzer and Essler, 2000). LPA also stimulates the tyrosine kinase Src during shape change causing subsequent activation of the
tyrosine kinase Syk (Maschberger et al., 2000). Src and Syk, which are most likely Gi activated in platelets, can elicit an increased exposure of fibrinogen binding sites on the integrin IIb3 which is a prerequisite for subsequent aggregation (Bauer et al., 2001). Higher concentrations of LPA were required to induce aggregation in isolated platelets (EC50 = 10M) (Benton et al., 1982) (Maschberger et al., 2000). Aggregation of washed platelets was considerably influenced by the concentration of albumin, and by fibrinogen (Benton et al., 1982). Benton et al showed that 1mg/ml bovine serum albumin (=15M) was able to inhibit the platelet aggregation caused by 3M of LPA. Due to albumin present in plasma much higher concentrations of LPA are needed to induce aggregation in platelet rich plasma (20-100M) (Tokumura et al., 1987). Studies on the effects of a number of LPA analogues have shown a higher potency for highly unsaturated and long chain acyl LPAs (such as C20:4) than for the LPA analogues with a shorter C18 fatty acyl group, such as an oleoyl group (18:1). Ether-linkedLPAs had the strongest aggregating activity (Tokumura et al., 2002). When platelets are stimulated by thrombin they mostly produce acyl- LPA (18:2) and acyl- LPA (20:4) (Sano et al., 2002). Acyl- LPA (20:4) is a more potent platelet activator in inducing shape change than other acyl- LPA species (Rother et al., 2003; Tokumura et al., 2002). Compared to acyl-LPA (16:0) the (20:4) acyl- LPA- species was 6.5-fold more effective in inducing platelet shape change (Rother et al., 2003). However, alkyl- LPA (16:0) has been shown to have the greatest potency in inducing shape change, 18.5-fold more effective than acyl- LPA (16:0) (Rother et al., 2003). The study of Gueguen et al has determined varying extents of platelet aggregation for nine LPA analogues all showing cross-desensitization for the same LPA receptor (Gueguen et al., 1999). Also in the study of Guegen platelets have shown a decreased response to LPA in a medium containing 1.3 mM calcium, contrasting previous observations of Sugiura et al who showed the necessity of calcium for LPA-induced aggregation (Alberghina et al., ; Gueguen et al., 1999; Sugiura et al., 1994). Thus in our studies experiments acyl-LPA (16:0) and alkyl-LPA (16:0) will be analysed for inducing aggregation with regard to calcium dependency.

The supernatant, platelet-poor-plasma, is carefully and completely removed to avoid the generation of thrombin during subsequent washing steps. Platelets are then resuspended in the first wash solution, prewarmed to 37C, with the addition of 1 l/ml prostacyclin, PGImM. A wash volume of 10 ml is normally required for the platelet pellet derived from 50-100 ml of blood. The platelet suspension is transferred to a closed test tube and incubated at 37C for 10 minutes. Then 1 l/ml of PGI2 is added followed by centrifugation for 8 minutes at 2600 rpm. The platelet pellet is then resuspended in the final suspension medium (20 ml), Tyrode buffer containing 2 mM CaCl2, and 0.35 % fatty acid free human albumin at a density of 3x105 platelets/l. 0.02 U/ml of the ADP scavenger apyrase were added. Platelets were kept at 37C throughout all experiments. When required, cytoplasmatic labeling of platelets may be performed by incubation in the first wash slolution with [3H]serotonin (0.2 Ci/ml) according to (Gachet et al., 1995; Ohlmann et al., 2000).
4.3.6. Measuring shape change and aggregation in washed platelets and platelet- rich- plasma (PRP)
All of the experiments measuring shape change and aggregation in washed platelets and PRP were performed in a LABOR aggregometer (Fa. Fresenius, Bad Homburg, Germany) at 37C under stirring (1000 rpm) by the turbidimetric method of Born. 400l aliquots of PRP or platelet suspension were placed into transluscent plastic cuvettes containing magnetic stirring-bars and incubated for 1 min before being placed on the magnetic stirrer base of the aggregometer. After 30 to 90 seconds of stirring the different agonists were added directly to the samples of platelet suspension and platelet shape change or aggregation were recorded in % decrease or increase in light transmission with a fine-needle writer. Light transmission of unstimulated PRP represents 0 % aggregation the light transmission of PPP 100 % aggregation. Platelet shape change was recorded as a reduction in light transmission caused by negative deflection of light.
4.3.7. Aggregation in blood
Whole blood platelet aggregation was measured using the single-platelet counting technique described by Heptinstall et al with the following modifications(Fox SC, 1982). Aliquots (400l) of anticoagulated blood samples were placed into aggregometer cuvettes and incubated 2 min at 37 C. The samples were stirred in the LABOR aggregometer (section 4.2.6) at 300 rpm for 30 sec. The inhibitor/antagonist was added and incubated for 30 sec before adding LPA (1M20M, from a 1 mM stock solution dissolved in albumin buffer (section 4.1.2). Other antagonists except aspirin were added in the same manner than for PRP experiments. Aliquots (15l) were removed at time 0 for baseline count and at set time intervalls therafter. The 15l were placed in 30l of fixing solution. After dilution to 1 ml the samples were centrifuged at a speed set by the Sysmex Platelet Centrifuge PC-800 (Toa Medical Electronics, Japan) and counted using a platelet counter (Sysmex Platelet Counter PL-100, Toa Medical Electronics, Japan). The percentage aggregation calculated as percentage loss of single platelets compared to baseline count. All platelet counts were done in duplicates. The samples were removed at predetermined time intervals in order to obtain a time dependent course of aggregation. The samples needed to remain at least 30 min in fixing solution before platelet count was performed.

4.3.8. Measuring cAMP levels in platelets
In washed platelets according to method 1 cAMP levels were determined with an enzymeimmuno- assay (EIA) Kit (Assay Designs, Inc).
The platelet suspension, adjusted to 1 million cells/l, was incubated with the stable prostacyclin analog, Iloprost (50 nM), for 5 minutes 37 C in order to raise cAMP levels. Platelets were then stirred at 1000 rpm for 30 seconds until the platelet stimulant, LPA (0.M), ADP (5 and 50 M) or epinephrine (10 M) was added. After 2 minutes of stirring the platelets were lysed in a 0.1M HCl (f.c.) solution. All samples were measured in duplicates. After 5 minutes all platelets had been been subject to lysis and the samples were centrifuged at 1000g for 5 minutes. The supernatant was placed into a 64-well microplate pre-coated with an antibody. The principle of measuring cAMP can be seen in the figure below. cAMP in the supernatant from lysed platelets competes with cAMP conjugated to alkaline phosphatase. Alkaline phosphatase has the ability of splitting paranitrophenylphosphate producing a yellow color of the end product. The bound yellow color is inversely proportional to the concentration of cAMP. The measured optical density is used to calculate the concentration of cAMP in pmol/ 106 platelets.

Alkaline Phosphatase

cAMP cAMP

P-Npp*

Alkaline Phosphatase acts as kinase to p-Npp (paranitrophenylphosphate) O N O
Fig 4.2 Principle of the enzyme immono assay (EIA) when measuring camp cAMP in the supernatant from lysed platelets competes with cAMP conjugated to alkaline phosphatase. Alkaline phosphatase splits paranitrophenylphosphate producing a yellow color. The bound yellow color is inversely proportional to the concentration of cAMP.
4.3.9. Quantifying platelet aggregates and platelet-monocyte- aggregates (PMA) and P-selectin expression
4.3.9.1. Preparation of whole blood samples and measuring aggregation FACS analysis 400l aliquots of hirudinized blood as described for blood. The stirrer was set to a speed of 1000 rpm and timed to 5 min. The agonist was added prior to stirring and the reaction stopped after 5 minutes by placing 100l of blood sample in red blood cell (RBC) lysing solution. Centrifuging the lysed samples at 3000 rpm for 5 minutes allowed the removal of RBC debris. Washing the pellet with 1ml PBS, centrifuging at 3000rpm for 5 minutes, removed most of the lysing solution. This step was followed by incubation with antibodies. CD 14-PE binds to a monocyte specific glycoprotein marking all the monoctes. As a second antibody monoclonal CD 41a- FITC binding to GPIIb part of the fibrinogen receptor on platelets was used to stain platelets. The unspecific binding was determined with a FITC-conjugated IgG1 isotype control antibody. All samples were incubated for 15 min at room temperature in the dark followed by resuspension in 600l PBS. 4.3.9.2. Measuring P-selectin expression in platelets isolated by method 4 For P-selectin expression, platelets (1.5 x 108) were incubated for 15 min with 0.5 mg anti-CD62PE antibody, and 0.5 ml Tyrodes buffer containing 2 mM CaCl2 and 0.35 % fatty acid free albumin was added. Mean fluorescence intensity was measured by FACS after collecting 10000 platelet events. The mean fluorescence intensity of isotype-matched IgG1-PE was substracted 4.3.9.3. Principle of FACS analysis The principle of Flow Cytometry is analyzing the light signals generated by particles as they flow through a liquid stream past a light beam. An argon laser beam emitting light at 488 nm constitutes the light source. The argon laser is focused to an elipse of 20m by 60m in size. The smaller height of 20 m avoids the simultaneous analysis of two cells being in the beam at the same time. Two cells closer than 20m together would be counted as one event, thus there is an ideal cell density of under 1 million cells/m. While cells are in the laser beam illumination signals and fluorochrome signals are collected simultaneously. The illumination signals are collected by the first lens positioned in the forward direction of the laser beam and the second lens positioned at a right angle to the laser beam. The light collected head on is not the laser light directly, but it is light that has been slightly refracted by the cell and strikes the forward positioned lens indicating the size of the cell. The laser light striking the forward positioned lens directly is being blocked by a centrally located obscuration bar. Light hitting the forward scatter lens is focused onto a photodiode where it is converted to an electrical current. In a similar manner the sideward positioned lens collects light that has been scattered to wide angles from the original direction of the beam. Irregularities in the cell cause this wide angle scatter indicating the granularity of the cell.

5.2.3. Shape change induced by LPA of washed platelets, PRP and blood is mediated by Rho-kinase activation but independent of ADP receptors P2Y1 and P2Y12
Many platelet stimuli induce shape change independent of additional platelet activators such as ADP and thromboxane A2. To explore whether LPA- induced shape change is independent of released ADP, the ADP receptor antagonists MRS2179 (a P2Y1 receptor inhibitor) and ARC 69931MX (P2Y12 receptor inhibitor) were used. They showed no inhibitory effect on platelet shape change in washed platelets and PRP (results not shown). Shape change induced by LPA showed a similar dose response curve, whether this response was induced in PRP or in blood (Fig 5.8a). This indicates the independence from red blood cells and suggesting the independence from ADP. Indeed, the P2Y1 receptor antagonist, MRS2179, and the P2Y12 receptor antagonist, AR-C69931MX or the combination of both antagonists did not inhibit LPA-induced shape change in whole blood (Fig 5.8b). The control experiments show that as expected- MRS2179 but not AR-C69931MX blocked the ADP-induced shape change in PRP and whole blood (Fig 5.8b). Together these results indicate that LPA induces shape change independent of synergistic interaction with ADP. LPA-induced shape change of washed platelets is mediated by Rho-kinase activation (Retzer and Essler, 2000). Consistent with this result it was observed, that LPA-induced platelet shape change in blood could be inhibited by the Rho-kinase inhibitor Y27632. Also pre-treatment of platelets with aspirin had no effect on LPA-induced shape change, neither in washed platelets, PRP (results not shown), nor in blood (Fig 5.8b). Therefore LPA-induced shape change seems to be independent of thromboxane. In contrast to the donor-dependent variation of LPA-induced aggregation (see section 5.3.1), LPA-induced shape change of PRP and blood was observed in all blood donors tested.
5.2.4. Comparison of 1-acyl-LPA (16:0) and 1-alkyl-LPA (16:0)- induced shape change in PRP and in whole blood
It has been shown recently that 80% acyl-LPA and 20% alkyl-LPA is present in atherosclerotic plaques (Rother et al., 2003). We determined the potency of the two different LPA species, 1alkyl-LPA (16:0) and 1-acyl-LPA in inducing shape change and aggregation. The results identified 1-alkyl-LPA (16:0) to be almost 20- fold more potent than 1-acyl-LPA in inducing shape change of PRP (Fig 5.8a). 1-Alkyl-LPA (16:0) was also more potent than 1-acyl-LPA (16:0) in inducing shape change of washed platelets and in whole blood (data not shown and section 5.3.3).

Results 5.8a

Shape change in PRP % of maximum
alk yl-LPA acyl -LPA ac yl-LPA

Similar than in PRP donor- dependent differences in aggregation could be observed in whole blood. In some donors platelets aggregated after addition of 2,5 M of LPA. Also the maximum aggregation after adding 20 M LPA was double in group A compared to group B (Fig 5.15). The distribution of the LPA- induced aggregation in blood is shown in Fig 5.16. Of 56 donors tested, 21 donors showed a platelet aggregation response to 20 M LPA higher than 50 % with a mean SD of 70% 14 %. Others (25 donors) showed aggregation lower than 50 % with a mean SD of %. Ten donors showed no significant aggregation as compared to control.

0 2.10 20

Group A

Group B

Fig 5.15 Donor dependent differences in aggregation induced by different concentrations of LPA in LPA-sensitive donors Whole blood anticoagulated with hirudin was stirred for 1 minute at 300 rpm at 37C before different LPA concentrations were added. Group A represents donors that showed a maximum aggregation > 50%, group B lower than 50% after adding 20M LPA. Values are mean SD, n=4 for group A, n=5 for group B.

Aggregation, %

Fig 5.16
Distribution of LPA-sensitive donors in response to 20 M of LPA Whole blood anticoagulated with hirudin was stirred for 1 minute at 300 rpm at 37C before LPA was added. Aggregation was measured 90 seconds after the addition of LPA. Each dot represents % aggregation of one donor (n=56). Dotted line indicates the average spontaneous aggregation after stirring 90 seconds.
In order to measure the intra- individual variation of platelet aggregation, four donors were tested repeatedly (3 to 5 times) on different days and it was found that the aggregation response was fairly consistent within the same blood donor (Table 5.1).
Date Control LPA 20 M ADP 5 M Date Control LPA 20 M ADP 5 M Date Control LPA 20 M ADP 5 M Date Control LPA 20 M ADP 5 M

Table 5.1

Donor A 23.04.02 27.03.78
05.08.29.08.05.03.25.04.35 84

06.08.12.08.07.03.82 6

04.09.76 85
Donor B 19.03.02 27.05.84
Donor C 25.02.02 04.03.90
Donor D 18.09.03 02.04.59

platelet pellet and excluded the use of apyrase (Benton et al., 1982; Gueguen et al., 1999). They were the first to show LPA-induced platelet shape change and aggregation in washed platelets. However, they used acid citrate dextrose (ACD) anticoagulated blood, followed by dilution with ACD to PRP in a 1:1 ratio, thus chelating much more of the extracellular Ca2+ and Mg2+, than in our method 3, and thereby possibly preventing platelet pre-activation during the subsequent centrifugation step. Platelets prepared according to method 4 were not pre-activated and showed shape change followed by aggregation in response to LPA. The reaction of platelets to LPA using this method was similar to platelet activation in PRP. The platelets were washed in the presence of prostacyclin (PGI2) and heparin and resuspended in buffer containing apyrase to avoid desensitization of the ADP receptor P2Y1 (Baurand et al., 2000; Cazenave et al.). PGI2 prevents dense granule release during centrifugation. No dense granule secretion as measured by serotonin release occurred until platelets were stimulated by LPA. LPA in this platelet preparation caused irreversible aggregation via ADP release from dense granules. This can be concluded by the inhibition of LPA-induced platelet aggregation by either ADP-receptor antagonist (Fig5.19).
6.2.2. Aggregation in PRP and whole blood
LPA induced platelet aggregation in whole blood at very low concentrations approaching those found in vivo (0.5- 1M) (Fig 5.12) (Baker et al., 2000). In comparison much higher LPA concentrations were required in platelet rich plasma (Fig 5.9). Therefore, we suspected platelet stimulating factors to be present in whole blood. Using different platelet inhibitors for studying the possible mechanism we found that blockade of either the P2Y1 or the P2Y12 receptor completely inhibited LPA-induced platelet aggregation in whole blood, demonstrating a complete dependence on ADP possibly released from red blood cells or from platelets. LPA on its own induced very little dense granule secretion (1-2%) in washed platelets, but LPA together with ADP, once secreted, effectively induced dense granule release. Remarkably, blockade of either the P2Y1 or the P2Y12 receptor reduced LPA-induced dense-granule secretion. Our results indicate that coactivation of the LPA-receptors with either one of the two purinergic ADP receptors is sufficient to induce full platelet response. Both ADP receptors show equal importance in mediating LPA-induced aggregation in whole blood (Fig 5.23) and aggregation and dense granule secretion of washed platelets (method 4) (Fig 5.19). Most likely both ADP receptors are activated simultaneously by released ADP, and the synergism of signaling pathways induced by the P2Y1 receptor (Gq, Ca2+) and the P2Y12 receptor (Gi) is required for platelet aggregation. Indeed, exogenous ADP also induces platelet aggregation in whole blood which is blocked by either P2Y1 or P2Y12 inhibition (Table 5.5). The situation is different for ADPinduced platelet activation in suspensions or PRP, where the P2Y1 receptor has been responsible for the initial activation and the P2Y12 receptor for amplification (Storey et al., 2001).

Summary

7. Summary
Oxidized LDL and platelets play a central role in the pathogenesis of atherosclerosis and ischemic cardiovascular diseases. Lysophosphatidic acid (LPA) is a thrombogenic substance that accumulates in mildly-oxidized LDL and in human atherosclerotic lesions, and is responsible for the initial platelet activation, shape change, induced by mildly-oxidized LDL and extracts of lipid-rich atherosclerotic plaques (Siess et al., 1999 Proc Natl Acad Sci USA 1999). LPA directly induced platelet shape change in blood and platelet-rich plasma (PRP) obtained from all blood donors. Albumin was one of the main inhibiting factors of platelet shape change in plasma. Interestingly LPA, at concentrations slightly above plasma levels, induced platelet shape change and aggregation in blood. 1-alkyl-LPA (16:0) was almost 20-fold more potent than 1-acyl-LPA (16:0). LPA-stimulated platelet aggregation in blood and PRP was donor-dependent. LPA-induced aggregation in blood could be completely blocked by the ADP- scavenging enzyme, apyrase, and antagonists of the platelet ADP-receptors P2Y1 and P2Y12. These substances also inhibited LPA-induced aggregation of platelet-rich plasma, and aggregation and serotonin secretion of washed platelets. These results indicate a central role for ADP-mediated P2Y1 and P2Y12 receptor activation in supporting LPA-induced platelet aggregation and show that LPA synergistically with ADP induces platelet aggregation in blood. Thus antagonists of platelet P2Y1 and P2Y12 receptors, especially in donors highly sensitive to LPA, might be useful in preventing LPA-elicited thrombus formation in patients with cardiovascular diseases. The mechanism of LPA plus ADP-induced aggregation was independent of the Rho/Rho kinase pathway which mediated LPA-induced platelet shape change in blood. LPA, activating G13, but not Gi or Gq synergized also with epinephrine, activating Gi, and serotonin, activating Gq, in amplifying LPA-induced platelet aggregation in washed platelets. LPA/serotonin-induced aggregation was blocked by either ADP-receptor antagonist whereas synergistic aggregation induced by LPA/epinephrine was independent from ADP-receptor antagonists. The latter results demonstrate an additional mechanism for aggregation independent of P2Y1 and P2Y12. Most surprising, LPA-induced platelet aggregation was insensitive to inhibition by aspirin. LPA at low concentrations, starting slightly above plasma level, was also capable of eliciting platelet-monocyte conjugate formation. LPA-induced platelet-monocyte formation was independent of the blood donor. ADP mediated P2Y1 and P2Y12 receptor activation played only a

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Acknowledgement

Acknowledgements
I sincerely thank Prof. Dr. med. W. Siess who has made this work possible with his wealth in ideas. With the constant search for intricate new experimental approaches, he enabled me to gain new results worthy for publication, for which I am very grateful. Thank you also for the many opportunities I had to participate at international scientific conventions. Thank you to Prof. Dr. med. P.C. Weber for providing the laboratory amentities at the Institut fr Prophylaxe und Epidemiologie der Kreislaufkrankheiten. I respectfully thank Dr. C. Gachet at the INSERM U311, EFS- Alsace, Strasbourg, France, for collaborating on the methodology and collegially sharing his results and ideas. Thank you for the invaluable Computer assistance by Pankaj Goyal. Also I thank Dharmendra Pandey for collaborating on some of the results shown in this work. I greatly appreciate the technical assistance by the diligent work of Nicole Wilke. Lastly I owe many hours of thanks to Wolfgang Erl who was committed to assist me on a new method and therefore made the final part of this work possible.

List of publications

Original Journal Publication
Nadine Haserck, Wolfgang Erl, Dharmendra Pandey, Gabor Tigyi, Philippe Ohlmann, Catherine Ravanat, Christian Gachet, and Wolfgang Siess (2003) The plaque lipid lysophosphatidic acid stimulates platelet activation and platelet-monocyte aggregate formatioini in whole bloodInvolvement of the P2Y1 and the P2Y12 receptor. Blood, First Edition Nov 26, 2003