Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M601670200 on October 21, 2006

J. Biol. Chem., Vol. 281, Issue 50, 38418-38429, December 15, 2006
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
281/50/38418    most recent
M601670200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pilquil, C.
Right arrow Articles by Brindley, D. N.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pilquil, C.
Right arrow Articles by Brindley, D. N.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Lipid Phosphate Phosphatase-1 Regulates Lysophosphatidate-induced Fibroblast Migration by Controlling Phospholipase D2-dependent Phosphatidate Generation*Formula

Carlos Pilquil{ddagger}1, Jay Dewald{ddagger}, Anton Cherney{ddagger}, Irina Gorshkova§, Gabor Tigyi, Denis English||, Viswanathan Natarajan§, and David N. Brindley{ddagger}2

From the {ddagger}Signal Transduction Research Group, Department of Biochemistry, University of Alberta, Edmonton, Alberta T6G 2S2, Canada, the Department of Physiology, University of Tennessee Health Science Center, Memphis, Tennessee 38163, the ||Department of Neurosurgery Center of Excellence for Aging and Brain Repair, University of South Florida College of Medicine, MDC-78, Tampa, Florida 36112, and the §Department of Medicine, Section of Pulmonary and Critical Care, University of Chicago, Chicago, Illinois 60637

Received for publication, February 22, 2006 , and in revised form, October 17, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Lysophosphatidate (LPA) stimulates cell migration and division through a family of G-protein-coupled receptors. Lipid phosphate phosphatase-1 (LPP1) regulates the degradation of extracellular LPA as well as the intracellular accumulation of lipid phosphates. Here we show that increasing the catalytic activity of LPP1 decreased the pertussis toxin-sensitive stimulation of fibroblast migration by LPA and an LPA-receptor agonist that could not be dephosphorylated. Conversely, knockdown of endogenous LPP1 activity increased LPA-induced migration. However, LPP1 did not affect PDGF- or endothelin-induced migration of fibroblasts in Transwell chamber and "wound healing" assays. Thus, in addition to degrading exogenous LPA, LPP1 controls signaling downstream of LPA receptors. Consistent with this conclusion, LPP1 expression decreased phospholipase D (PLD) stimulation by LPA and PDGF, and phosphatidate accumulation. This LPP1 effect was upstream of PLD activation in addition to the possible metabolism of phosphatidate to diacylglycerol. PLD2 activation was necessary for LPA-, but not PDGF-induced migration. Increased LPP1 expression also decreased the LPA-, but not the PDGF-induced activation of important proteins involved in fibroblast migration. These included decreased LPA-induced activation of ERK and Rho, and the basal activities of Rac and Cdc42. However, ERK and Rho activation were not downstream targets of LPA-induced PLD2 activity. We conclude that the intracellular actions of LPP1 play important functions in regulating LPA-induced fibroblast migration through PLD2. LPP1 also controls PDGF-induced phosphatidate formation. These results shed new light on the roles of LPP1 in controlling wound healing and the growth and metastasis of tumors.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
It is well established that LPA plays a major physiological and pathological role in controlling cell signaling. LPA is present in biological fluids and activates cells through families of G-protein-coupled receptors (1, 2) that are coupled through Gi, G12/13, or Gq. This leads to intracellular effects such as decreases in cAMP, stimulation of phospholipase D (PLD)3 and Rho effectors leading to stress fiber formation, and activation of phospholipase C, PKC isoforms, ERK, and Ca2+ transients. Extracellular LPA is produced in inflammatory conditions by secretory phospholipase A2 acting on PA (3). LPA is also produced from circulating lysophosphatidylcholine by secreted lysophospholipase D and autotaxin (4, 5). Autotaxin stimulates fibroblast migration and division for wound repair (6), and its expression shows a positive correlation with tumor progression, angiogenesis, and metastasis (4-6). Importantly, LPA mimics many effects of autotaxin. Furthermore, LPA concentrations are high in ascites fluid of patients with ovarian tumors (7-9). LPA promotes ovarian tumor development and protects against chemotherapy (10). These effects highlight our growing knowledge of the importance of LPA in tissue repair, tumor growth, and metastasis.

Consequently, increasing the degradation of extracellular LPA, or attenuating signaling through the LPA receptors can regulate the extracellular actions of LPA. The present work focuses on the effects of the integral membrane protein, lipid phosphate phosphatase-1 (LPP1), which dephosphorylates LPA and other lipid phosphates, and the mechanisms whereby this enzyme modifies fibroblast migration. LPP1 acts partly in the plasma membrane as an ecto-enzyme (11, 12). It was originally proposed that this ecto activity explained how LPP1 overexpression attenuated the effects of exogenous LPA in activating cell division, ERK, phospholipase D, and Ca2+ transients (13, 14). Subsequent studies demonstrate that gonadotropin releasing hormone increases LPP expression in the plasma membranes of ovarian cancer cells and that this explains the anti-proliferative effects of gonadotropin releasing hormone on ovarian carcinomas (15). Overexpression of LPP3 decreases growth, survival, and tumorigenesis of ovarian cancer cells leading to the conclusion that this occurred through the increase in exogenous LPA degradation (16). Ovarian cancer cells express decreased levels of LPP1, which could contribute to increases in extracellular LPA and the promotion of transformation (17). Consistent with this view, overexpressing LPP1 in these cells decreased LPA-induced migration (17). In platelets exogenous LPA increased the expression of ecto-LPP1 activity (18) thus decreasing net LPA production and LPA-induced shape changes and aggregation. Ecto-LPP activities were also concluded to regulate extracellular LPA accumulation that results in the proliferation of pre-adipocytes (19). The simplest explanation for these combined results is that the ecto-LPP activities control extracellular LPA concentrations, thereby regulating LPA receptor activation. Although, partly true, other mechanisms of action have been demonstrated.

In our fibroblast experiments (13, 14) we limited the degradation of exogenous LPA to <10% of the total added and LPP expression still attenuated LPA-induced stimulation of ERK, cell division, PLD, and Ca2+ transients. Alderton et al. (20) showed diminished activation of ERK by extracellular LPA, S1P, PA, and thrombin in cells that overexpressed LPP1, -1A, and -2, but not in cells that overexpressed LPP3. The effect on thrombin signaling indicates that ecto-LPP activity is not responsible for attenuating ERK activation, but that this depended upon decreases in intracellular PA concentrations (20). Subsequent work showed that overexpressing LPP2 and LPP3 increased apoptosis in serum-deprived HEK 293 cells and this correlated with decreased concentrations of PA and S1P in the cells, respectively (21). Also, HEK 293 cells that overexpress LPP3 exhibited greater DAG formation subsequent to PLD stimulation. PLD2 and LPP3 are both present in caveolin-1-enriched microdomains (22). Zhao et al. (23) also concluded that the effects of LPP1 in inhibiting Ca2+ transients and the secretion of the inflammatory cytokine, interleukin-8, occurred downstream of LPA receptor activation. Furthermore, in vivo studies with LPP1 overexpressing transgenic mice failed to detect alterations in circulating LPA levels (24). Fibroblasts from these LPP1 overexpressing mice revealed decreased steady-state levels of PA and a rise in DAG. This work failed to show decreased LPA- and PDGF-induced ERK1/2 activation in fibroblasts from LPP1 overexpressing mice (24). However, decreased ERK activation by S1P and PDGF was observed in fibroblasts prepared from mice that expressed 20 gene copies for Lpp1, but little effect was seen in fibroblast that expressed two gene copies (25). We used the latter fibroblasts in some of our studies on migration. Long et al. (25) postulated that the effects of LPP1 in decreasing PDGF-induced fibroblast migration in a "wound healing" assay were caused by long-term increases in DAG concentrations and desensitization of PKC-mediated cell signaling.

The actions of the LPPs on intracellular PA concentrations can also modify cell signaling because the PA is an important bioactive lipid (see Refs. 11 and 12 for original citations). PA production in neutrophils stimulates NADPH oxidase. PA also stimulates protein kinase C-{zeta}, phosphatidylinositol 4-kinase, and phospholipase C-{gamma} and increases Ras-GTP concentrations (11, 12). It also increases sphingosine kinase-1 activity (26) and inhibits protein phosphatase-1 (27). PA formation in Madin-Darby canine kidney (28) and HIRcB cells (29) activates Raf-1 and ERK. PA also increases cell division through mTOR (30-32) and stimulates stress fiber formation (33, 34). The relative concentrations of LPA and PA in membranes control their curvature and vesicle budding (35, 36). PLD1 and its activator, ARF, are involved in vesicle movement through PA production (37, 38).

The present experiments focused on the effects of LPP1 on fibroblast migration. Knockdown of LPP1 activity increased LPA-induced migration, whereas increasing LPP1 activity decreased this migration. Under our experimental conditions, changing LPP1 expression did not affect PDGF- or endothelin-induced migration. The regulation of LPA-induced migration by LPP1 did not depend upon the degradation of exogenous LPA. The inhibitory effect of LPP1 on migration was downstream of LPA-receptor stimulation through its activation of Rho and control of PA formation by PLD2. The latter activity is necessary for the stimulation of Rat2 fibroblast migration by LPA, but not by PDGF.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Dulbecco's minimum essential medium (DMEM) and trypsin-EDTA were purchased from Invitrogen. Fetal bovine serum and human recombinant PDGF-betabeta were from Medicorp, Inc. (Montréal, PQ, Canada). Fatty acid-free bovine serum albumin, trypsin inhibitor, PA, LPA, GW9662, and human endothelin-1 were from Sigma. [9,10-3H]Palmitate (57 Ci/mmol) was from PerkinElmer Life Sciences, Inc. and [{gamma}-32P]ATP was from GE Healthcare. 1,2-Dioleoyl-sn-glycero-3-phosphobutanol (PB) and sn-1,2-dioleoylglycerol were from Avanti%20Polar%20Lipids">Avanti Polar Lipids Inc. (Alabaster, AL). PD98059 was purchased from EMD Biosciences (Darmstadt, Germany). The {alpha}-hydroxyphosphonate analogue of LPA, wls-31, was a kind gift from Drs. Kevin R. Lynch and Timothy L. Macdonald, University of Virginia Health Sciences Center, Charlottesville, VA. Polyclonal rabbit antibodies for mLPP1 were prepared as described previously (13) and antibodies for PLD2 were a gift from Dr. S. Bourgoin, University of Laval, Québec, PQ, Canada. Rabbit polyclonal antibodies for PLD1 were purchased from BioSource International Inc. (Camarillo, CA). Monoclonal antibodies for Rho-A, -B, and -C, and Rac were from Upstate (Charlottesville, VA). Mouse monoclonal phospho-ERK1/2 antibody was from Cell Signaling Technology Inc. (Beverly, MA). Mouse monoclonal anti-phosphotyrosine (PY20) and PPAR{gamma} and rabbit polyclonal antibodies for PDGFR-beta, Cdc42, and ERK1 were from Santa Cruz Biotechnology. Twenty-four-well Costar Transwell Permeable Supports were from Corning Inc. (Corning, NY). The membranes were precoated with 6 µg of human plasma fibronectin obtained from Sigma. We described previously the preparation of adenoviral vectors for expressing LPP1, the inactive LPP1(R217K) mutant (23), and dominant/negative PLD1 and PLD2 (39).

Fibroblast Culture and Modifying the Expression of LPP1 Activity—We engineered mixed populations of Rat2 fibroblasts that stably overexpress LPP1 by using a retroviral vector and puromycin selection (13). We also employed fibroblasts obtained from transgenic mice that overexpressed 2 copies of the Lpp1 gene (24). Catalytically active LPP1 and the inactive R217K mutant (40) were transiently overexpressed by using adenoviral constructs at 40 multiplicity of infection for 24 h, when new serum-supplemented medium was added and the cells tested 18 h later (23). Knockdown of LPP1 activity was achieved using double-stranded SMARTpool® siRNAs designed to target rat LPP1 from Dharmacon Inc. (Lafayette, CO). About 500,000 Rat2 fibroblasts were plated on 10-cm plates with DMEM and 10% FBS and allowed to double in number over 24 h. Lipofectamine 2000 (Invitrogen) in Opti-MEM (Invitrogen) was used at 1 x 106 cells to 7 µg of Lipofectamine 2000 with a final concentration of 200 nM siRNAs. Transfection was performed in antibiotic-free medium containing serum according to the manufacturer's instructions. After 4.5 h, the transfection media was replaced with DMEM and 10% FBS. Cells were starved in DMEM and 0.4% BSA 24 h after transfection. For each experiment, controls for LPP1 knockdown were performed with functional non-targeting siCONTROL (Dharmacon) siRNAs, which can silence firefly luciferase, and with Lipofectamine 2000 alone. Total LPP activity was measured using PA in Triton X-100 micelles and ecto-LPP activity was determined by measuring the release of 32Pi from 32P-labeled LPA (13).

Detection of mRNA Expression of LPPs—Changes in mRNA concentrations in fibroblasts that were left over from the migration assay were measured by real time reverse transcriptase-PCR using appropriate controls and primers as described previously (41).

Western Blotting—Cell lysates were collected in buffer that contained 50 mM Hepes, 137 mM NaCl, 1 mM MgCl, 1 mM CaCl, 10% glycerol, 0.1% SDS, 1% Triton X-100, 0.5% deoxycholate, 2.5 mM EDTA, 1 mM orthovanadate, 10 mM NaF, and 1% Sigma protease inhibitor mixture. Sample protein (75 µg) was loaded on 10- and 7.5% SDS-PAGE gels for separating and detecting LPP1 and PLD, respectively. The proteins were transferred to nitrocellulose membranes and blocked overnight in Odyssey Blocking Buffer (LI-COR, Lincoln, NB). Rabbit polyclonal antibodies for mLPP1, PLD1, and PLD2 were diluted 1:1000 in the Odyssey Blocking Buffer supplemented with 0.1% Tween 20 and incubated for 2 h at room temperature. Transactivation of the PDGFR by LPA was measured by immunoprecipitating 100 µg of sample protein with mouse monoclonal anti-phosphotyrosine antibody and analyzing the antibody complex from protein A-Sepharose (from GE Healthcare) on a 7.5% SDS-PAGE gel followed by Western blotting with rabbit polyclonal anti-PDGFR-beta diluted 1:1000. For ERK analysis, 30 µg of protein was analyzed using a 7.5% SDS-PAGE gel. The membrane was probed simultaneously with rabbit polyclonal anti-ERK1 antibodies, and mouse monoclonal antibodies (both diluted 1:2000) that recognize phosphorylated ERK1/2. Goat anti-rabbit IgG conjugated to an IRDye800 (Rockland Immunochemica, Gilbertsville, PA) and goat anti-mouse IgG conjugated to Alexa Fluor 680 (Molecular Probes, Eugene, OR) fluorescent markers were diluted 1:10,000 and incubated for 1 h at room temperature. Fluorescence on the Western blot at 700 and 800 nm was scanned with the Odyssey Infrared Imaging System and quantified with Odyssey software. The phosphorylation of ERK was expressed relative to total ERK.

Measurement of PA and DAG Concentrations and PLD Activation—For the measurement of PA formation, 350,000 Rat2 fibroblasts were plated on 10-cm dishes with DMEM and 10% FBS. After 48 h the medium was changed to DMEM that contained 0.4% BSA and 20 µCi of [3H]palmitate. The cells were incubated 18 h when the media was removed and unlabeled DMEM and 0.4% BSA were added to the cells for an additional 2 h. The cells were then treated with agonists for 30 min, and the reactions were stopped by washing twice with ice-cold Hepes-buffered saline. The cells were then collected by scrapping twice in 0.5 ml of methanol. One ml of chloroform and 0.9 ml of 2 M KCl containing 0.2 M HCl were added and the mixtures were vortexed and then centrifuged. The bottom organic phase was isolated and dried under N2 and dissolved in 100 µl of chloroform/methanol (9:1). One half of the sample was used to measure PA levels and the other half was used for the DAG mass assay. For measuring PA concentrations, samples of the lipids were loaded in the middle of plastic-supported Silica Gel 60 thin layer chromatography plates. Plates were developed twice in chloroform/methanol/ammonium hydroxide (65:35:7.5, by volume), and the plates were cut 1 cm above the PA standard. They were then turned upside down, and developed in the reverse direction with chloroform/methanol/acetic acid/acetone/water (50:10:12:20:5, by volume). The mass of DAG in the cells was determined by converting it to PA in the presence of [{gamma}-32P]ATP and DAG kinase (42). The concentration of DAG was determined by using a standard curve prepared with sn-1,2-dioleoylglycerol. The mass of DAG was expressed relative to the total phospholipid content of the cells (42).

For measurement of PLD activity, 73,000 fibroblasts were plated in DMEM and 10% FBS on 3.5-cm dishes. After 48 h, the medium was changed to 1 ml of DMEM that contained 0.4% BSA and 5 µCi of [3H]palmitate. The cells were incubated 18 h when the media was replaced with unlabeled DMEM plus 0.4% BSA, and the cells were incubated for 2 h. The fibroblasts were then preincubated with 30 mM butan-1-ol for 15 min and then treated with agonists for 6 min so that PLD activity could be measured through the formation of PB. Lipids were extracted, and isolated as for PA above. Samples of the lipids including 50 µg of PB internal standard were loaded on glass-supported Silica Gel 60 thin layer chromatography plates and the plates were developed using the upper organic phase of ethyl acetate/iso-octane/acetic acid/water (130:20:30:100, by volume). Lipids were detected by I2 staining and PB and the total phospholipids near the origin were collected by scrapping. 3H labeling was measured by liquid scintillation counting. Relative PLD activity was calculated as the percentage of PB relative to the labeling of total phospholipids.

Fibroblast Migration—For the Transwell assay, about 350,000 cells were plated on 10-cm dishes with DMEM and 10% FBS and no antibiotics. After 24 h the fibroblasts were treated with adenovirus at 40 multiplicity of infection for LPPs and 50 multiplicity of infection for the PLDs or siRNA as indicated. After a further 24 h the cells were starved with DMEM and 0.4% fatty acid-free BSA for 18 h. The cells were collected using trypsin-EDTA and quickly neutralized with trypsin inhibitor. Cells (100,000) in DMEM and 0.4% fatty acid-free BSA were plated inside 6.5-mm diameter Transwell inserts with 8.0-µm pore size polycarbonate filters for 2 h before they were placed in the bottom well. When indicated, 30 mM butan-1-ol or butan-2-ol, 200 nM GW9662, 40 µM PD98059, or 0.1% Me2SO as a vehicle control were added to the Transwell 30 min before the migration started. The bottom wells contained the agonists, DMEM, and 0.4% fatty acid-free BSA supplemented with 0.4% charcoal-treated FBS plus butanol, or inhibitors as indicated. The cells were allowed to migrate for 6 h at which time the top of the filters was cleared by using a wet cotton-tipped applicator followed by a dry one. The migrated cells were then fixed in 5% formaldehyde in Hepes-buffered saline for 30 min. Visualization of the fixed cells on the filters was achieved with a hematoxylin/eosin combination stain, or the nuclei were stained with 1 µg/ml Hoechst 33258 for 2 h. Images of about 6 fields per filter were taken using a digital camera and a fluorescence microscope (Leica DM IRB) and these were magnified 100-fold.

For the wound healing assay, Rat2 fibroblasts were grown to confluence in 24-well plates (Corning) and starved with DMEM and 0.4% fatty acid-free BSA for 18 h. The monolayer was scratched using the end of a 200-µl pipette tip, and the detached cells were removed with Hepes-buffered saline. DMEM and 0.4% fatty acid-free BSA with or without agonists was added to each well as indicated. Images at x25 magnification were taken at different time intervals with a digital camera and a microscope (Leica DM IRB). The size of the image was quantitated by using the graduations on a hemocytometer. This enabled us to estimate the average distance (measured at seven different points) by which the walls of the wound had closed.

Measurement of Rho, Rac, and Cdc42 Activation—The relative concentrations of Rho-GTP and Rac-GTP plus Cdc42-GTP were determined in pull-down assays using Rhotekin Rho binding domain bound to glutathione-agarose (from Upstate) and the GTPase binding domain of PAK1 bound to glutathione-Sepharose 4B (from GE Healthcare), respectively (43, 44). The amounts of bound Rho, Rac, and Cdc42 were then determined by Western blot analysis using mouse monoclonal antibodies for Rho and Rac at 3 µg/ml and 1:2000 dilution, respectively, and polyclonal antibodies for Cdc42 at 1:500 dilution. The results were normalized to the total amount of Rho, Rac, or Cdc42, respectively, which was used for the pull-down assays.

Statistical Analysis—Statistical differences were calculated by analysis of variance followed by a Newman-Keuls post hoc test.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
LPP1 Activity Regulates the Stimulation of Fibroblast Migration by LPA, but not by PDGF or Endothelin—To determine how LPP1 activity alters migration to various agonists, we established mixed populations of Rat2 fibroblasts that stably overexpressed LPP1 mRNA by about 12-fold (41). This previous work also demonstrated that overexpression, or knockdown of LPP1 mRNA, did not change mRNA expression for LPP2 and LPP3. LPP1 overexpressing fibroblasts used in the present studies showed ~4- and 2-fold increases in total LPP and ecto-LPP activities, respectively, compared with wild-type or vector control cells (Fig. 1, A and B). These LPP1 overexpressing fibroblasts showed a severe inhibition of LPA-induced migration in a wound healing assay (Fig. 1, C and D). There was no significant effect on the actions of PDGF and endothelin on migration (Fig. 1D).

The wound healing assay measures mainly the non-directional, chemokinetic movement of fibroblasts because no diffusion gradient of the agonist is established. Therefore, we also employed a Transwell chamber assay to distinguish between chemotaxis (directional migration toward a gradient of chemoattractant) and chemokinesis. The mode of the migration that was stimulated by the three agonists was determined by adding the same concentration of agonist to the top and bottom chambers and comparing the migration to when the agonist was present only in the bottom chamber. PDGF-induced migration was mainly mediated through chemotaxis because adding 20 ng/ml PDGF to the top chamber inhibited migration to the bottom chamber by about 74 ± 7% in three experiments. By contrast, the migration to 0.5 µM LPA and 100 nM endothelin was mainly mediated by chemokinesis. Adding the equivalent concentrations of LPA to the top chambers blocked migration by 43 ± 22% in five experiments. For endothelin there was no significant decrease in migration in two experiments. Our results for LPA and PDGF migration agree with previous work with fibroblasts (45, 46).

The results from the Transwell assay demonstrate that the migration of Rat2 fibroblasts in response to 0.1-10 µM LPA was severely inhibited by increased LPP1 expression (Fig. 2A). We also employed fibroblasts from mice that expressed two gene copies of Lpp1 and these showed about 2-fold increases in total and ecto-LPP activities compared with fibroblasts from control mice (24). These LPP1 overexpressing fibroblasts also showed decreased migration to LPA (Fig. 2B) with the most striking effects at lower LPA concentrations. We also tried to study the effects of LPP1 using S1P as an agonist. However, S1P did not stimulate fibroblast migration in the assays used (results not shown).

To investigate whether the effect of LPP1 on LPA-induced migration depends upon its catalytic activity, Rat2 fibroblasts were infected with adenoviral vectors containing mLPP1-myc or the inactive mLPP1(R217K)-myc (40). Overexpression of the wild-type LPP1 increased total LPP activity by about 4-fold, whereas expression of the mutant did not change LPP activity significantly (results not shown). As a control, we confirmed by Western blotting that the inactive LPP1 mutant was expressed to similar extents compared with active LPP1 (Fig. 2C, inset). Transient expression of wild-type LPP1, but not LPP1(R217K), decreased LPA-induced migration compared with wild-type cells (Fig. 2C).


Figure 1
View larger version (53K):
[in this window]
[in a new window]

  		FIGURE 1.
Increasing LPP1 activity attenuates LPA-induced fibroblast migration in an "in vitro wound healing" assay. A and B show total LPP and ecto-LPP activities, respectively, for wild-type fibroblasts (Rat2) and fibroblasts transduced stably with the empty vector (Vector) or vector containing cDNA for LPP1 (LPP1). The mean ecto-activity for the vector control cells was 295 pmol of LPA dephosphorylated/min per mg of cell protein. Results are mean ± S.D. for at least six independent experiments. C shows the migration into a "wound" that was scratched 36 h previously. D shows the average distance by which the gap between the fibroblasts closed over 24 h when the indicated concentrations of LPA, PDGF, and endothelin-1 (Et-1) were added to the incubation medium. The results for wild-type and LPP1 overexpressing fibroblasts are shown in white and black columns, respectively. The values are mean ± S.D. for five or three independent experiments for LPA and PDGF, respectively. Means and ranges for two experiments are shown for endothelin. Significant differences between LPP1 overexpressing fibroblasts and wild-type or vector control fibroblasts are indicated by: {dagger}, p < 0.001, and, **, p < 0.05.

 
To determine whether the effects of LPP1 overexpression were specific for LPA-induced migration we also measured the responses to PDGF and endothelin. LPP1 overexpression did not significantly alter chemotaxis to PDGF, or chemokinesis with endothelin (Fig. 2, D and E).

To provide further evidence that changing LPP1 activity controls LPA-induced migration, we knocked down endogenous LPP1 expression using a combination of four siRNAs to rat LPP1. mRNA for rat LPP1 was decreased by about 90% compared with treatment with control siRNA or Lipofectamine alone (Fig. 3A), and this was accompanied by a 50% decrease in total LPP activity (Fig. 3B). The lower effect on LPP activities compared with mRNA levels is explained by the fact that the fibroblasts continue to express LPP2 and LPP3. Fibroblasts in which LPP1 activity was knocked down showed increased (p < 0.05) LPA-induced migration compared with cells treated with control siRNA (Fig. 3C). There was no significant effect of the LPP1 knockdown on migration to PDGF (Fig. 3C). Knockdown of LPP1 also increased fibroblast migration in the absence of added agonist in some experiments. This suggests that basal migration could result from endogenously produced LPA. The combined results from the experiments shown in Figs. 1, 2, 3 therefore, establish that changing the activity of LPP1 specifically regulates LPA-induced fibroblast migration.

The Effect of LPP1 on LPA-induced Migration Is Not Mediated through the Degradation of Extracellular LPA—The most direct explanation for the specific effects of LPP1 expression of LPA-induced cell migration is that the ecto-activity of LPP1 decreases the ability of exogenous LPA to stimulate its receptors. However, we limited the degradation of exogenous LPA in the 6-h Transwell assays to less than 20% even at the lowest LPA concentrations. Also, we saw effects of LPP1 expression when we added excess (up to 10 µM) LPA (Fig. 2A). Therefore, the increased ecto-activity of LPP1 could not have changed bulk LPA concentrations in the medium to an extent that could have affected fibroblast migration.

Alternatively, the ecto-activity could enable mono-oleoylglycerol formed from oleoyl-LPA to enter the cell and thereby regulate cell signaling after re-phosphorylation (11, 12). For example, intracellular LPA could signal through internal LPA receptors (47) or the PPAR{gamma} receptors (48, 49) and thereby affect migration. We excluded this ecto-LPP1 effect by adding 10 nM to 5 µM exogenous mono-oleoylglycerol to the fibroblasts and establishing that this had no effect on LPA-induced migration (results not shown). In further experiments we added 200 nM GW9662 to the fibroblasts to inhibit PPAR{gamma} receptor activity. This had no significant effect on LPA- or PDGF-induced migration. Furthermore, the concentration of PPAR{gamma} receptors, as determined by Western blotting, was not significantly different among the wild-type, vector control, and LPP1 expressing fibroblast (results not shown). These combined results demonstrate that changes in PPAR{gamma} receptor activation do not explain the actions of LPP1 in decreasing fibroblast migration under the present 6-h migration assay.


Figure 2
View larger version (27K):
[in this window]
[in a new window]

  		FIGURE 2.
Overexpression of LPP1 decreases fibroblast migration in a Transwell assay in response to LPA, but not PDGF, or endothelin. A, D, and E show the migration of Rat2 fibroblasts (white bars), fibroblasts transduced stably with the empty vector (gray bars), or vector containing cDNA for LPP1 (black bars). B shows the migration of fibroblasts derived from control mice (white bars) or mice that overexpressed LPP1 (black bars). C shows the migration of wild-type fibroblasts (white bars), those that transiently overexpressed LPP-1-myc (black bars), or the inactive mutant, LPP1(R217K)-myc (hatched bars). Results are mean ± S.D. for at least three independent experiments. The inset to C shows a Western blot performed with rabbit anti-mLPP1 antibody that demonstrates the overexpression of mutant LPP1 (2) and wild-type LPP1 (3) compared with non-transfected fibroblasts (1). Significant differences between LPP1 overexpressing fibroblasts and wild-type or vector control fibroblasts in the same treatment groups are indicated by: {dagger}, p < 0.001, and **, p < 0.05.

 
To investigate the role of the ecto-LPP activity more directly we used the {alpha}-hydroxyphosphonate analogue (wls-31) of LPA to promote cell migration. This analogue cannot be dephosphorylated by the LPPs and it is an agonist for LPA1/3 receptors (50, 51). Reverse transcriptase-PCR analysis demonstrated that Rat2 fibroblasts express LPA1, but not LPA2 and LPA3 (results not shown). Fibroblast migration to LPA or wls-31 (results not shown) was inhibited by more than 90% after preincubation of the cells with pertussis toxin indicating the coupling of LPA1 to G{alpha}i (Fig. 4A). Pertussis toxin, as expected, did not alter PDGF-induced migration. Increased expression of LPP1 also decreased the stimulation of migration by wls-31 (Fig. 4B). These results establish that hydrolysis of extracellular LPA is not required for LPP1-induced regulation of LPA-induced migration.

Effects of LPP1 Expression on the Activation of ERK, Rac, and Rho with Respect to Fibroblast Migration—Our conclusion that LPP1 has a signaling effect downstream of LPA1 activation is compatible with work on the control of ERK activation by the LPPs (20, 21, 25). Under the present experimental conditions, LPA-induced ERK phosphorylation was decreased by an average of 62% by stable overexpression of LPP1 (Fig. 5, A and B). However, there was no significant effect of LPP1 overexpression on PDGF-induced ERK activation. We also investigated whether the decreased activation of ERK could contribute to the effect of LPP1 expression on LPA-induced migration by treating wild-type fibroblasts with 40 µM PD98059 to block ERK activation. This treatment inhibited LPA- and PDGF-induced ERK phosphorylation by about 93 and 77%, respectively (Fig. 5, C and D). LPA- and PDGF-induced migrations were decreased by about 57 and 37% in the presence of 40 µM PD98059, respectively (Fig. 5E). These results confirm that ERK activation is required for fibroblast migration. The lack of effect of LPP1 on PDGF-induced migration is compatible with a failure to decrease ERK activation significantly by this agonist in our experimental system.

In other experiments, we determined the effects of LPP1 expression on the activation of the small G-proteins Rho and Rac, which are well known to be involved in cell migration. Total Rho and Rac concentrations were not changed significantly by LPP1 expression (results not shown). Also, basal Rho-GTP concentrations in wild-type, vector control, and LPP1 overexpressing fibroblasts were not significantly different (Fig. 6A). However, the LPA-induced increase in Rho-GTP concentration was significantly attenuated in fibroblasts that overexpressed LPP1. Stimulation of the fibroblasts with PDGF under these experimental conditions produced no significant increase in Rho-GTP in any of the cell lines.

The concentration of Rac-GTP under basal conditions was decreased in the LPP1 overexpressing fibroblasts compared with wild-type fibroblasts and the vector controls (Fig. 6B). Treatment with LPA did not change Rac-GTP concentrations significantly in any of the cell lines under these conditions. Stimulation of the LPP1 overexpressing fibroblasts with PDGF appeared to increase Rac-GTP concentrations in LPP1 overexpressing fibroblasts such that its concentration was not significantly different from the wild-type or vector control fibroblasts (Fig. 6B). Similarly, Cdc42-GTP concentrations under basal conditions were significantly decreased by an average of 44% (p < 0.05) in LPP1 overexpressing fibroblasts compared with wild-type fibroblasts and the vector controls. However, there was no increase in Cdc42-GTP concentrations upon activation with LPA and PDGF in any of the cell lines under these conditions (results not shown).

LPP1 Expression Decreases PLD Activity and PA Accumulation after Stimulating Fibroblasts with either LPA or PDGF, but PLD-dependent PA Formation Is Only Required for LPA-induced Fibroblast Migration—To determine whether LPP1 inhibited fibroblast migration by controlling PA production, we stimulated the fibroblasts with LPA, or PDGF to increase PLD activity. Basal PA concentrations were not significantly different among the three cell lines (Fig. 7A). Overexpression of LPP1 decreased LPA- and PDGF-induced increases in steady-state accumulation of PA by 70-80% (Fig. 7A). LPP1 activity has been implicated in converting PA to DAG and therefore we determined the effects of LPP1 expression on steady-state DAG concentrations. LPA and PDGF both increased the relative mass of DAG in LPP1 overexpressing fibroblasts (p < 0.05), but there was no significant effect of increasing LPP1 activity on DAG accumulation (Fig. 6B). To determine the site of the LPP1 effect on PA we also measured the effect on PLD activation. Overexpression of LPP1 also decreased PLD activation after stimulation with both PDGF and LPA (Fig. 7C). Thus the decreased accumulation of PA can result from both decreased formation from phosphatidylcholine and the action of LPP1 on the PA.


Figure 3
View larger version (15K):
[in this window]
[in a new window]

  		FIGURE 3.
Knockdown of LPP1 activity increases fibroblast migration in a Transwell assay in response to LPA, but not PDGF. Wild-type Rat2 fibroblasts were treated with vehicle, the control siRNA, or siRNA for rat LPP1. A and B show expression of mRNA for LPP1 and total LPP activity, respectively. The LPP activity in the siCONTROL cells was 19.2 nmol of phosphatidate dephosphorylated/min per mg of protein. C shows the migration in a Transwell of the same fibroblasts treated with control siRNA (white bars) or siRNA for rat LPP1 (black bars). Results are mean ± S.D. for cell numbers in about 6 fields that were counted. The results are from one representative experiment, which was reproduced in three further experiments. Significant differences between fibroblasts with LPP1 knockdown and siRNA controls in A and B are indicated by: {dagger}, p < 0.001; **, p < 0.05.

 
These results on PLD activity and PA accumulation on their own does not explain the differential effect of LPP1 in only attenuating LPA-, but not PDGF-induced migration. We, therefore, postulated that PLD activation and PA formation were required for fibroblast migration to LPA, whereas it was not necessary for PDGF-induced migration. This was tested by adding butan-1-ol to the migration assays to block PA formation by PLD. This treatment inhibited the stimulation of migration by LPA, but not to PDGF. As a negative control we used butan-2-ol, which does not participate in the transphosphatidylation reaction of PLD. Butan-2-ol did not alter LPA- or PDGF-induced migration significantly.


Figure 4
View larger version (19K):
[in this window]
[in a new window]

  		FIGURE 4.
LPA-stimulated migration is receptor-mediated, but the inhibitory effects of LPP1 do not result from dephosphorylation of extracellular LPA. In A, wild-type fibroblasts were preincubated for 18 h with 100 nM pertussis toxin as indicated. B shows the stimulation of migration by wls-31 in wild-type fibroblasts (white bars), fibroblasts transduced with empty vector (gray bars), and fibroblasts that stably overexpressed LPP1 (black bars). Results are mean ± S.D. for three independent experiments. Significant differences of the pertussis toxin effect on LPA-treated fibroblasts (A) and LPP1 overexpressing fibroblasts and wild-type or vector control fibroblasts in the same treatment groups (B) are indicated by: *, p < 0.01.

 
To strengthen these results, we also decreased PLD activity by transfecting the fibroblasts with adenoviral vectors that encoded dominant/negative mutants of PLD1 and PLD2. Expression of both mutant PLDs was confirmed by Western blotting (Fig. 8, B and C). The expression of dominant/negative PLD2 decreased LPA-induced migration by about 75%, whereas expression of dominant/negative PLD1 had no significant effect (Fig. 8D). By contrast, there was no significant effect of either PLD mutant on PDGF-induced migration.

We also investigated whether decreased LPA-induced activation of PLD in LPP1 overexpressing fibroblasts is responsible for the lower activation of Rho (Fig. 6A) or ERK (Fig. 5, A and B), which might account for the decreased LPA-induced migration. Dominant/negative PLD1 or PLD2 were expressed separately in wild-type fibroblasts, which were then treated for 10 min with 0.5 µM LPA. There was no significant difference in the activation of Rho or ERK in fibroblasts that expressed dominant/negative PLD1 or PLD2 (supplementary Fig. i). As controls we verified that the mutant PLDs were overexpressed as in Fig. 8, B and C. We also showed that expressing dominant/negative PLD1 and PLD2 under these conditions decreased LPA-induced PLD activity by an average of 38 and 46%, respectively. Thus, the combined inhibition of total PLD activity was about 84%. These results are compatible with previous work on the efficacy of the mutants in blocking LPA-induced PLD activity (39). Taken together, the experiments in Fig. 8 support the essential role of PLD2 activation and PA formation in LPA-induced fibroblast migration and identify a mechanism for the inhibitory effect of LPP1.


Figure 5
View larger version (23K):
[in this window]
[in a new window]

  		FIGURE 5.
Effect of LPP1 expression on ERK activation by LPA and PDGF. A and C show representative Western blots for phospho-ERK1/2 and total ERK1 that were measured simultaneously on the same membrane. B shows the quantitation of the Western blots at 10 min after stimulation of wild-type fibroblasts (white columns), vector control cells (gray columns), and LPP1 overexpressing cells (black columns) with LPA or PDGF. Results are mean ± S.D. for three independent experiments. D shows a representative quantitation of C. The results in E show the effects of 40 µM PD 98059 (hatched columns) on LPA- and PDGF-induced fibroblast migration compared with vehicle (0.1% Me2SO)-treated fibroblasts (white columns). Results are mean ± S.D. for three independent experiments. Significant differences between LPP1 overexpressing fibroblasts and wild-type or vector control fibroblasts in the same treatment groups in B and the PD98059 treatment in E are indicated by: *, p < 0.01, and **, p < 0.05.

 


Figure 6
View larger version (21K):
[in this window]
[in a new window]

  		FIGURE 6.
Effect of LPP1 expression on Rho and Rac activation by LPA and PDGF. Results in A and B show the relative effects of treating wild-type fibroblasts (white columns), vector control cells (gray columns), and LPP1 overexpressing cells (black columns) for 10 min with LPA or PDGF. Results are mean ± S.D. for six independent experiments for Rho activation, and seven independent experiments for Rac activation. Significant differences between LPP1 overexpressing fibroblasts and wild-type, or vector control fibroblasts in the same treatment groups are indicated by: {dagger}, p < 0.001, and **, p < 0.05.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The present work on fibroblast migration employed both a wound healing assay that detects mainly chemokinesis and a Transwell chamber assay that can detect both chemotaxis and chemokinesis. We used a variety of approaches to establish that the catalytic activity of LPP1 regulates LPA-induced chemokinesis in fibroblasts. The inhibition of LPA-induced migration was observed in Rat2 fibroblasts that stably overexpressed LPP1 by retroviral transduction, in fibroblasts that transiently overexpressed LPP1 after adenoviral infection, and in fibroblasts derived from mice that overexpressed LPP1. Overexpression of enzymes can produce ectopic expression and non-physiological results. Therefore, further evidence was provided by knocking down endogenous LPP1 activity and showing that this increased LPA-induced migration. LPP1 is a major contributor to LPP activity in the fibroblasts (41) because increasing, or knocking down its expression significantly altered total LPP activity.


Figure 7
View larger version (18K):
[in this window]
[in a new window]

  		FIGURE 7.
Increased LPP1 activity decreases LPA- and PDGF-induced PA formation and PLD activation. A and B show the steady-state accumulation of PA and DAG relative to phosphatidylcholine, respectively, whereas C shows the relative PLD activity. The cells are indicated as follows: wild-type Rat2 fibroblasts (white bars), fibroblasts transduced stably with the empty vector (gray bars), and fibroblasts containing cDNA for LPP1 (black bars). Results are mean ± S.D. for at least three independent experiments. Significant differences between LPP1 overexpressing fibroblasts and wild-type or vector control fibroblasts in the same treatment groups are indicated by: {dagger}, p < 0.001; *, p < 0.01; and **, p < 0.05.

 
Our next objective was to determine the mechanism by which LPP1 controls LPA-induced fibroblast migration. First, LPPs might have non-catalytic effects on migration. Human LPP3 possesses an extracellular RGD cell adhesion sequence that promotes cell/cell interactions (52). hLPP3 bound to a subset of integrins, and anti-{alpha}vbeta3 and anti-{alpha}5beta1 antibodies blocked binding. In platelets, LPP3 co-localized with {alpha}11bbeta3 (18). hLPP1 has RGN instead of RGD in hLPP3 (53). This sequence could be involved in regulating migration because peptides with RGD and RGN sequences antagonize the binding of T-lymphocytes to fibronectin (54). The equivalent sequence in the mouse and rat LPP1 used in the present work is QGN (53), which probably does not affect cell/cell interactions. The fact that expressing the inactive R217K mutant (40) of mouse LPP1 did not decrease LPA-induced fibroblast migration eliminates a non-catalytic effect of LPP1 in the present work.

The effect of LPP1 on fibroblast migration was specific for LPA because there was no significant effect of LPP1 expression on the stimulation of migration of Rat2 fibroblasts by endothelin, which also stimulates G-protein-coupled receptors, or to PDGF that signals through a receptor tyrosine kinase. This lack of effect of LPP1 activity on PDGF-induced migration differs from that reported by Long et al. (25) who used a wound healing assay with mouse fibroblasts that expressed 20 gene copies of Lpp1. This level of LPP1 expression resulted in decreased ERK activation in response to LPA, PDGF, and S1P (25). However, we used the equivalent fibroblasts that expressed 2 gene copies of Lpp1 and therefore had lower LPP1 activity (24, 25). In these mouse fibroblasts there was no significant LPP1-induced decrease in ERK activation by LPA or PDGF (24, 25). However, LPA-induced migration was attenuated in mouse fibroblasts that expressed two gene copies of Lpp1 (Fig. 2B). Therefore, ERK activation in the latter fibroblasts does not appear to be the target for LPP1 in controlling LPA-induced migration. In Rat2 fibroblasts, overexpression of LPP1 did attenuate ERK activation by LPA, but not PDGF (Fig. 5). This decrease in LPA-induced ERK activation was not explained by inhibition of PLD1 or PLD2. However, our work also showed that when ERK activation was inhibited with PD98059, this resulted in an inhibition of LPA- and PDGF-induced migration of Rat2 fibroblasts.

Another major difference in our studies and those of Long et al. (25) is that we used Transwell chambers that for PDGF determined its predominant effects on chemotaxis rather than chemokinesis, which is determined by the wound healing assays. Again, LPP1 expression failed to decrease PDGF-induced fibroblast migration (Fig. 2, C and D). The effects of LPP1 on LPA-induced migration does not simply relate to a specific blocking of chemokinesis because there was no significant action of LPP1 activity on the chemokinetic effects of endothelin (Figs. 1D and 2E).

The next question was whether LPP1 modified fibroblast migration specifically to LPA by dephosphorylating exogenous LPA and therefore its ability to activate LPA receptors. We eliminated this possibility by ensuring that the total breakdown of LPA in the Transwell assay never exceeded 20% of the exogenous LPA. We also added excess LPA (up to 10 µM) to stimulate migration, but this did not reverse the LPP1-induced inhibition. Therefore, effects of LPP activity on the bulk concentrations of exogenous LPA cannot account for the observed changes in LPA-induced migration. It is possible that a small pool of added LPA has access to LPA receptors and that this is the pool specifically metabolized by LPP1 to control migration. Alternatively, ecto-activity could enable mono-oleoylglycerol formed from oleoyl-LPA to enter the cell and thereby regulate migration after re-phosphorylation (11, 12) and stimulation of internal LPA1 (47) or PPAR{gamma} receptors (48, 49). We excluded this possibility by adding different concentrations of mono-oleoylglycerol to the outside of the fibroblasts and showing that this did not affect LPA-induced migration. We also showed that inhibiting PPAR{gamma} receptor activity did not alter migration and that PPAR{gamma} receptor expression was not altered significantly by increasing LPP1 activity.


Figure 8
View larger version (17K):
[in this window]
[in a new window]

  		FIGURE 8.
PLD2 activity is required for LPA, but not PDGF-induced fibroblast migration. A shows the effects of butan-1-ol on LPA- and PDGF-induced migration of wild-type Rat2 fibroblasts. Butan-2-ol was used as a negative control because, unlike butan-1-ol, it does not prevent the formation of PA by PLD. B and C show Western blots that demonstrate the overexpression of dominant/negative PLD1 and PLD2, respectively, by adenoviral infection. D shows the effects of expressing dominant/negative mutants of PLD1 (gray bars) and PLD2 (black bars) compared with vector controls (white bars) on LPA- and PDGF-induced migration of wild-type Rat2 fibroblasts. Results are mean ± S.D. for four independent experiments. Significant differences between control and treated fibroblasts are indicated by: **, p < 0.05.

 
However, the most compelling result that excludes an effect of ecto-LPP1 activity on migration is that overexpressing LPP1 also decreased migration to the {alpha}-ketophosphonate analogue (wls-31) of LPA. wls-31 cannot be hydrolyzed by LPP1. This compound has agonist activity for LPA1 receptors (50, 51) that are expressed in Rat2 fibroblasts. In our work the effects of LPA and wls-31 on migration were inhibited by pertussis toxin, indicating receptor coupling to G{alpha}i. Because the LPP1 effect depends on catalytic activity, it must be mediated by changing the metabolism of a lipid phosphate formed downstream of LPA1 activation. LPP1 potentially hydrolyzes a variety of intracellular lipid phosphates including LPA, PA, ceramide 1-phosphate, and S1P provided that these substrates have access to the enzyme (11, 12).

We concentrated our attention on the generation of PA through PLD. The LPPs can dephosphorylate PA to DAG following PLD activation (11, 12) and overexpression of different LPPs can decrease intracellular PA accumulation (20, 21, 24, 55, 56). In the present work, overexpressing LPP1 severely decreased PA accumulation after stimulating the fibroblasts with both LPA and PDGF. Long et al. (25) postulated that the long-term effects of increased LPP1 activity in increasing DAG concentrations decreased the expression of PKC{alpha} and that this could account for decreased ERK activation. Equally, DAG-dependent PKCs activate PLD2 as well as PLD1 (57, 58). A decrease in these PKC activities could, therefore, explain the attenuation of PLD1 and PLD2 activation in cells that have increased LPP1 activity. This could represent a negative feedback loop to prevent further DAG formation through the PLD pathways. We are presently investigating this hypothesis. However, in our studies there were no significant effects of LPP1 activity on the accumulation of bulk cell DAG. The fact that DAG accumulation was maintained in LPP1 overexpressing cells despite a severely decreased PA accumulation (Fig. 7A) is therefore remarkable. This maintenance of DAG concentrations in LPP1 overexpressing fibroblasts is compatible with the hypothesis that LPP1 is involved in the conversion of PA to DAG. In addition, DAG could be derived by the stimulation of phospholipase C activities by LPA and PDGF.

Interestingly, our previous work showed that increasing LPP1 blocked PLD activation by LPA (14). Our present work demonstrates for the first time that LPP1 regulates the PDGF-induced activation of PLD. These LPP1 effects on LPA and PDGF signaling are upstream of PLD activation. This is in addition to the role of LPP1 in converting PA to DAG. Our work, therefore, extends the studies on Long et al. (25) in showing that LPP1 has a critical function in regulating signaling by PDGF as well as by LPA. However, this control by LPP1 of PLD activation by PDGF did not regulate PDGF-induced fibroblast migration in our work.

These results could be explained if increased PA formation were essential for LPA-induced migration, but not PDGF-induced migration. We verified this prediction by showing that blocking PLD-induced PA formation with butan-1-ol attenuated migration to LPA, but not to PDGF. We extended this work and established that PLD2, and not PLD1, is required for LPA-induced migration. Neither PLD isoform was required for PDGF-induced fibroblast migration. However, LPP1 could still be an important regulator of other aspects of PDGF-induced cell activation, or in the internalization and cycling of the PDGF receptor.

PLD2 activation is required for LPA-induced transactivation and tyrosine phosphorylation of the PDGFR (23). However, no significant tyrosine phosphorylation of the PDGFR was observed with 0.5 µM LPA, which supported optimum migration (results not shown). Sakai et al. (45) reached the same conclusion concerning the relatively low LPA concentrations that stimulate fibroblast migration compared with much higher concentrations needed for PDGFR transactivation. Furthermore, if LPA-induced migration required activation of the PDGFR it should stimulate mainly chemotaxis, whereas LPA controls mainly chemokinesis.

PLD2-induced PA formation is also required for the stimulation of phosphatidylinositol 4-phosphate 5-kinase I{gamma}beta and the synthesis of phosphatidylinositol 4,5-bisphosphate (59). These effects are required for activation of cell surface integrins that are required for cell adhesion and migration. Significantly, beta1A integrin is required for LPA-induced fibroblast migration (45). The action of LPP1 in decreasing PLD2 activation by LPA could, therefore, result in decreased engagement of integrins with the extracellular matrix resulting in decreased migration.

We previously determined the effects of LPP1 expression on LPA- and PDGF-induced Ca2+ transients. Increasing LPP1 inhibited Ca2+ release in response to LPA, but not to PDGF (14). This could also explain the different effects of LPP1 in inhibiting LPA-, but not PDGF-induced migration because the stimulation of fibroblast migration by both of these agonists depends upon the Ca2+ transient. This conclusion is illustrated by our observation that adding 35 µM BAPTA-AM blocked LPA- and PDGF-induced migration by 83 and 97%, respectively.4

We also investigated the effects of LPP1 expression on activation of the small G-proteins, Rho, Rac, and Cdc42. LPP1 expression significantly decreased Rac-GTP and Cdc42-GTP concentrations in unstimulated fibroblasts and decreased Rac-GTP in those activated with LPA, but not PDGF. Increased LPP1 expression also decreased LPA-induced increases in Rho-GTP concentrations. There was no significant effect of PDGF on Rho activation under these conditions. In previous work, there was high Rho activation in protrusions from mouse embryonic fibroblasts that were migrating non-directionally (as produced with LPA in our work) compared with cells stimulated with PDGF (60), which induces chemotaxis. Our results with LPP1 and the differences in signaling between LPA and PDGF are compatible with those of Kam and Exton (31). Significantly, inhibition of the activation of Rho family GTPases has a greater effect on the activation of PLD by LPA compared with PDGF, which resembles the effects of LPP1 shown in Fig. 7C. Our observation that expression of dominant/negative PLD1 or PLD2 did not alter the LPA-induced activation of Rho is compatible with Rho activation being upstream and not downstream of PLD. Significantly, Rho activation and PA formation are necessary for actin polymerization (33, 34), which is required for cell migration.

Work from animal models strongly implicates the LPPs as regulators of cell migration. Drosophila expresses two Wunen proteins homologous to LPP3. Wunen-1 and -2 decrease migration of primordial germ cells (61-63). Introduction of LPP1 has no effect on an endogenous Drosophila germ cell-specific factor in vivo, whereas LPP3 causes aberrant migration and germ cell death (63). Wunen proteins may act by promoting the uptake of a product that they form from a lipid phosphate (64). Work with mouse embryos from LPP3 knock-out mice showed failure to form chorio-allantoic placenta and yolk sac vasculature. Some embryos showed shortening of the anterior-posterior axis (65) similar to axin deficiency, a critical regulator of Wnt signaling. It was proposed (65) that LPP3 functions as Wnt signaling antagonists. These combined results demonstrate that different LPPs have distinct functions and participate in signaling systems for cell migration in flies and humans.

In conclusion, the present studies demonstrate that changes in the catalytic activity of LPP1 in fibroblasts are inversely related to migration of the fibroblasts in response to LPA. These effects are specific because migration to endothelin or PDGF was not changed significantly by LPP1 expression. The effect of increased LPP1 expression in attenuating LPA-induced fibroblast migration did not depend upon dephosphorylation of extracellular LPA. It was explained by a decreased LPA-induced activation of Rho and accumulation of PA following activation of PLD2. Significantly, LPP1 also attenuated PLD activation by PDGF. Although this did not affect migration, it could be important in controlling other aspects of fibroblast activation. Taken together these observations provide new concepts for understanding how the LPPs can regulate the effects of growth factors in promoting wound healing, tumor progression, and metastasis.


   FOOTNOTES
 
* This work was supported in part by Canadian Institutes of Health Research Grants MOP 10504 and 49491 (to D. N. B.), United States Public Health Serivce Grants NHBLI 61469 and CA-92160 (to G. T.), and National Institutes of Health Grants RO1 HL061751 (to D. E.) and RO1 HL 71152 and HL 079396 (to V. N.). 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. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. i. Back

1 Received a Graduate Student Research Award from the Alberta Heritage Foundation for Medical Research. Back

2 Supported by a Medical Scientist award from the Alberta Heritage Foundation for Medical Research. To whom correspondence should be addressed: 357 Heritage Medical Research Center, Edmonton, Alberta T6G 2S2, Canada. Tel.: 780-492-2078; Fax: 780-492-3383; E-mail: david.brindley{at}ualberta.ca.

3 The abbreviations used are: PLD, phospholipase D; DAG, diacylglycerol; DMEM, Dulbecco's minimum essential medium; LPA, lysophosphatidate; LPP, lipid phosphate phosphatase; PA, phosphatidate; PDGF, platelet-derived growth factor-betabeta; PDGFR, platelet-derived growth factor receptor-beta; PKC, protein kinase C; S1P, sphingosine 1-phosphate; BAPTA, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid; ERK, extracellular signal-regulated kinase; HEK, human embryonic kidney; PB, 1,2-dioleoyl-sn-glycero-3-phosphobutanol; siRNA, small interfering RNA; FBS, fetal bovine serum; BSA, bovine serum albumin. Back

4 A. Ilich and D. N. Brindley, unpublished results. Back


   ACKNOWLEDGMENTS
 
We thank Drs. K. R. Lynch and T. L. Macdonald for the gift of the {alpha}-hydroxyphosphonate analogue and Dr. S. Bourgoin for the PLD2 antibody. We also thank Dr. G. Eitzen for help establishing Rho-GTP and Rac-GTP assays.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Tigyi, G., and Parrill, A. L. (2003) Prog. Lipid Res. 42, 498-526[CrossRef][Medline] [Order article via Infotrieve]
  2. Moolenaar, W. H., van Meeteren, L. A., and Giepmans, B. N. (2004) Bioessays 26, 870-881[CrossRef][Medline] [Order article via Infotrieve]
  3. Fourcade, O., Simon, M. F., Viode, C., Rugani, N., Leballe, F., Ragab, A., Fournie, B., Sarda, L., and Chap, H. (1995) Cell 80, 919-927[CrossRef][Medline] [Order article via Infotrieve]
  4. Tokumura, A. (2002) Biochim. Biophys. Acta 1582, 18-25[Medline] [Order article via Infotrieve]
  5. Umezu-Goto, M., Kishi, Y., Taira, A., Hama, K., Dohmae, N., Takio, K., Yamori, T., Mills, G. B., Inoue, K., Aoki, J., and Arai, H. (2002) J. Cell Biol. 158, 227-233[Abstract/Free Full Text]
  6. Moolenaar, W. H. (2002) J. Cell Biol. 158, 197-199[Abstract/Free Full Text]
  7. Shen, Z., Wu, M., Wiper, D. W., Morton, R. E., Kennedy, A. W., Belinson, J., Markman, M., Casey, G., and Xu, Y. (1997) Clin. Chem. 43, 577-577
  8. Xu, Y., Shen, Z., Wiper, D. W., Wu, M., Morton, R. E., Elson, P., Kennedy, A. W., Belinson, J., Markman, M., and Casey, G. (1998) J. Am. Med. Assoc. 280, 719-723[Abstract/Free Full Text]
  9. Fang, X., Schummer, M., Mao, M., Yu, S., Tabassam, F. H., Swaby, R., Hasegawa, Y., Tanyi, J. L., LaPushin, R., Eder, A., Jaffe, R., Erickson, J., and Mills, G. B. (2002) Biochim. Biophys. Acta 1582, 257-264[Medline] [Order article via Infotrieve]
  10. Frankel, A., and Mills, G. B. (1996) Clin. Cancer Res. 2, 1307-1313[Abstract]
  11. Brindley, D. N. (2004) J. Cell Biochem. 92, 900-912[CrossRef][Medline] [Order article via Infotrieve]
  12. Sigal, Y. J., McDermott, M. I., and Morris, A. J. (2005) Biochem. J. 387, 281-293[CrossRef][Medline] [Order article via Infotrieve]
  13. Jasinska, R., Zhang, Q. X., Pilquil, C., Singh, I., Xu, J., Dewald, J., Dillon, D. A., Berthiaume, L. G., Carman, G. M., Waggoner, D. W., and Brindley, D. N. (1999) Biochem. J. 340, 677-686
  14. Pilquil, C., Singh, I., Zhang, Q., Ling, Z., Buri, K., Stromberg, L. M., Dewald, J., and Brindley, D. N. (2001) Prostaglandins 64, 83-92[Medline] [Order article via Infotrieve]
  15. Imai, A., Furui, T., Tamaya, T., and Mills, G. B. (2000) J. Clin. Endocrinol. Metab. 85, 3370-3375[Abstract/Free Full Text]
  16. Tanyi, J. L., Morris, A. J., Wolf, J. K., Fang, X., Hasegawa, Y., Lapushin, R., Auersperg, N., Sigal, Y. J., Newman, R. A., Felix, E. A., Atkinson, E. N., and Mills, G. B. (2003) Cancer Res. 63, 1073-1082[Abstract/Free Full Text]
  17. Tanyi, J. L., Hasegawa, Y., Lapushin, R., Morris, A. J., Wolf, J. K., Berchuck, A., Lu, K., Smith, D. I., Kalli, K., Hartmann, L. C., McCune, K., Fishman, D., Broaddus, R., Cheng, K. W., Atkinson, E. N., Yamal, J. M., Bast, R. C., Felix, E. A., Newman, R. A., and Mills, G. B. (2003) Clin. Cancer Res. 9, 3534-3545[Abstract/Free Full Text]
  18. Smyth, S. S., Sciorra, V. A., Sigal, Y. J., Pamuklar, Z., Wang, Z., Xu, Y., Prestwich, G. D., and Morris, A. J. (2003) J. Biol. Chem. 278, 43214-43223[Abstract/Free Full Text]
  19. Simon, M. F., Rey, A., Castan-Laurel, I., Gres, S., Sibrac, D., Valet, P., and Saulnier-Blache, J. S. (2002) J. Biol. Chem. 277, 23131-23136[Abstract/Free Full Text]
  20. Alderton, F., Darroch, P., Sambi, B., McKie, A., Ahmed, I. S., Pyne, N., and Pyne, S. (2001) J. Biol. Chem. 276, 13452-13460[Abstract/Free Full Text]
  21. Long, J., Darroch, P., Wan, K. F., Kong, K. C., Ktistakis, N., Pyne, N. J., and Pyne, S. (2005) Biochem. J. 391, 25-32[CrossRef][Medline] [Order article via Infotrieve]
  22. Sciorra, V. A., and Morris, A. J. (1999) Mol. Biol. Cell 10, 3863-3876[Abstract/Free Full Text]
  23. Zhao, Y., Usatyuk, P. V., Cummings, R., Saatian, B., He, D., Watkins, T., Morris, A., Spannhake, E. W., Brindley, D. N., and Natarajan, V. (2005) Biochem. J. 385, 493-502[CrossRef][Medline] [Order article via Infotrieve]
  24. Yue, J., Yokoyama, K., Balazs, L., Baker, D. L., Smalley, D., Pilquil, C., Brindley, D. N., and Tigyi, G. (2004) Cell Signal. 16, 385-399[CrossRef][Medline] [Order article via Infotrieve]
  25. Long, J. S., Yokoyama, K., Tigyi, G., Pyne, N. J., and Pyne, S. (2006) Biochem. J. 394, 495-500[CrossRef][Medline] [Order article via Infotrieve]
  26. Delon, C., Manifava, M., Wood, E., Thompson, D., Krugmann, S., Pyne, S., and Ktistakis, N. T. (2004) J. Biol. Chem. 279, 44763-44774[Abstract/Free Full Text]
  27. Jones, J. A., and Hannun, Y. A. (2002) J. Biol. Chem. 277, 15530-15538[Abstract/Free Full Text]
  28. Ghosh, S., Strum, J. C., Sciorra, V. A., Daniel, L., and Bell, R. M. (1996) J. Biol. Chem. 271, 8472-8480[Abstract/Free Full Text]
  29. Rizzo, M. A., Shome, K., Vasudevan, C., Stolz, D. B., Sung, T. C., Frohman, M. A., Watkins, S. C., and Romero, G. (1999) J. Biol. Chem. 274, 1131-1139[Abstract/Free Full Text]
  30. Fang, Y., Vilella-Bach, M., Bachmann, R., Flanigan, A., and Chen, J. (2001) Science 294, 1942-1945[Abstract/Free Full Text]
  31. Kam, Y., and Exton, J. H. (2004) FASEB J. 18, 311-319[Abstract/Free Full Text]
  32. Avila-Flores, A., Santos, T., Rincon, E., and Merida, I. (2005) J. Biol. Chem. 280, 10091-10099[Abstract/Free Full Text]
  33. Ha, K. S., and Exton, J. H. (1993) J. Cell Biol. 123, 1789-1796[Abstract/Free Full Text]
  34. Cross, M. J., Roberts, S., Ridley, A. J., Hodgkin, M. N., Stewart, A., Claesson-Welsh, L., and Wakelam, M. J. (1996) Curr. Biol. 6, 588-597[CrossRef][Medline] [Order article via Infotrieve]
  35. Zimmerberg, J. (2000) Traffic 1, 366-368[CrossRef][Medline] [Order article via Infotrieve]
  36. Huijbregts, R. P., Topalof, L., and Bankaitis, V. A. (2000) Traffic 1, 195-202[Medline] [Order article via Infotrieve]
  37. Jones, D., Morgan, C., and Cockcroft, S. (1999) Biochim. Biophys. Acta 1439, 229-244[Medline] [Order article via Infotrieve]
  38. Ktistakis, N. T., Brown, A., Waters, G. M., Sternweis, P. C., and Roth, M. G. (1996) J. Cell Biol. 134, 295-306[Abstract/Free Full Text]
  39. Wang, L., Cummings, R., Zhao, Y., Kazlauskas, A., Sham, J. K., Morris, A., Georas, S., Brindley, D. N., and Natarajan, V. (2003) J. Biol. Chem. 278, 39931-39940[Abstract/Free Full Text]
  40. Zhang, Q. X., Pilquil, C. S., Dewald, J., Berthiaume, L. G., and Brindley, D. N. (2000) Biochem. J. 345, 181-184
  41. Morris, K. E., Schang, L. M., and Brindley, D. N. (2006) J. Biol. Chem. 281, 9297-9306[Abstract/Free Full Text]
  42. Martin, A., Duffy, P. A., Liossis, C., Gomez-Munoz, A., O'Brien, L., Stone, J. C., and Brindley, D. N. (1997) Oncogene 14, 1571-1580[CrossRef][Medline] [Order article via Infotrieve]
  43. Ren, X. D., Kiosses, W. B., and Schwartz, M. A. (1999) EMBO J. 18, 578-585[CrossRef][Medline] [Order article via Infotrieve]
  44. Benard, V., Bohl, B. P., and Bokoch, G. M. (1999) J. Biol. Chem. 274, 13198-13204[Abstract/Free Full Text]
  45. Sakai, T., Peyruchaud, O., Fassler, R., and Mosher, D. F. (1998) J. Biol. Chem. 273, 19378-19382[Abstract/Free Full Text]
  46. Anand-Apte, B., Zetter, B. R., Viswanathan, A., Qiu, R. G., Chen, J., Ruggieri, R., and Symons, M. (1997) J. Biol. Chem. 272, 30688-30692[Abstract/Free Full Text]
  47. Gobeil, F., Jr., Bernier, S. G., Vazquez-Tello, A., Brault, S., Beauchamp, M. H., Quiniou, C., Marrache, A. M., Checchin, D., Sennlaub, F., Hou, X., Nader, M., Bkaily, G., Ribeiro-da-Silva, A., Goetzl, E. J., and Chemtob, S. (2003) J. Biol. Chem. 278, 38875-38883[Abstract/Free Full Text]
  48. McIntyre, T. M., Pontsler, A. V., Silva, A. R., St. Hilaire, A., Xu, Y., Hinshaw, J. C., Zimmerman, G. A., Hama, K., Aoki, J., Arai, H., and Prestwich, G. D. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 131-136[Abstract/Free Full Text]
  49. Zhang, C., Baker, D. L., Yasuda, S., Makarova, N., Balazs, L., Johnson, L. R., Marathe, G. K., McIntyre, T. M., Xu, Y., Prestwich, G. D., Byun, H. S., Bittman, R., and Tigyi, G. (2004) J. Exp. Med. 199, 763-774[Abstract/Free Full Text]
  50. Hooks, S. B., Santos, W. L., Im, D. S., Heise, C. E., Macdonald, T. L., and Lynch, K. R. (2001) J. Biol. Chem. 276, 4611-4621[Abstract/Free Full Text]
  51. Heise, C. E., Santos, W. L., Schreihofer, A. M., Heasley, B. H., Mukhin, Y. V., Macdonald, T. L., and Lynch, K. R. (2001) Mol. Pharmacol. 60, 1173-1180[Abstract/Free Full Text]
  52. Humtsoe, J. O., Feng, S., Thakker, G. D., Yang, J., Hong, J., and Wary, K. K. (2003) EMBO J. 22, 1539-1554[CrossRef][Medline] [Order article via Infotrieve]
  53. Waggoner, D. W., Xu, J., Singh, I., Jasinska, R., Zhang, Q. X., and Brindley, D. N. (1999) Biochim. Biophys. Acta 1439, 299-316[Medline] [Order article via Infotrieve]
  54. Preciado-Patt, L., Levartowsky, D., Prass, M., Hershkoviz, R., Lider, O., and Fridkin, M. (1994) Eur. J. Biochem. 223, 35-42[Medline] [Order article via Infotrieve]
  55. Leung, D. W., Tompkins, C. K., and White, T. (1998) DNA Cell Biol. 17, 377-385[Medline] [Order article via Infotrieve]
  56. Garcia-Murillas, I., Pettitt, T., Macdonald, E., Okkenhaug, H., Georgiev, P., Trivedi, D., Hassan, B., Wakelam, M., and Raghu, P. (2006) Neuron 49, 533-546[CrossRef][Medline] [Order article via Infotrieve]
  57. Meacci, E., Nuti, F., Catarzi, S., Vasta, V., Donati, C., Bourgoin, S., Bruni, P., Moss, J., and Vaughan, M. (2003) Biochemistry 42, 284-292[CrossRef][Medline] [Order article via Infotrieve]
  58. Chen, J. S., and Exton, J. H. (2004) J. Biol. Chem. 279, 22076-22083[Abstract/Free Full Text]
  59. Powner, D. J., Payne, R. M., Pettitt, T. R., Giudici, M. L., Irvine, R. F., and Wakelam, M. J. (2005) J. Cell Sci. 118, 2975-2986[Abstract/Free Full Text]
  60. Pertz, O., Hodgson, L., Klemke, R. L., and Hahn, K. M. (2006) Nature 440, 1069-1072[CrossRef][Medline] [Order article via Infotrieve]
  61. Zhang, N., Zhang, J., Purcell, K. J., Cheng, Y., and Howard, K. (1997) Nature 385, 64-67[CrossRef][Medline] [Order article via Infotrieve]
  62. Starz-Gaiano, M., Cho, N. K., Forbes, A., and Lehmann, R. (2001) Development 128, 983-991[Abstract]
  63. Burnett, C., and Howard, K. (2003) EMBO Rep. 4, 793-799[CrossRef][Medline] [Order article via Infotrieve]
  64. Renault, A. D., Sigal, Y. J., Morris, A. J., and Lehmann, R. (2004) Science 305, 1963-1966[Abstract/Free Full Text]
  65. Escalante-Alcalde, D., Hernandez, L., Le Stunff, H., Maeda, R., Lee, H. S., Jr., Gang, C., Sciorra, V. A., Daar, I., Spiegel, S., Morris, A. J., and Stewart,C. L. (2003) Development 130, 4623-4637[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Lipid Res.Home page
D. N. Brindley and C. Pilquil
Lipid phosphate phosphatases and signaling
J. Lipid Res., April 1, 2009; 50(Supplement): S225 - S230.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Biol. CellHome page
Z. Lee, C.-T. Cheng, H. Zhang, M. A. Subler, J. Wu, A. Mukherjee, J. J. Windle, C.-K. Chen, and X. Fang
Role of LPA4/p2y9/GPR23 in Negative Regulation of Cell Motility
Mol. Biol. Cell, December 1, 2008; 19(12): 5435 - 5445.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
I. Gorshkova, D. He, E. Berdyshev, P. Usatuyk, M. Burns, S. Kalari, Y. Zhao, S. Pendyala, J. G. N. Garcia, N. J. Pyne, et al.
Protein Kinase C-{epsilon} Regulates Sphingosine 1-Phosphate-mediated Migration of Human Lung Endothelial Cells through Activation of Phospholipase D2, Protein Kinase C-{zeta}, and Rac1
J. Biol. Chem., April 25, 2008; 283(17): 11794 - 11806.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
Y. Zhao, S. K. Kalari, P. V. Usatyuk, I. Gorshkova, D. He, T. Watkins, D. N. Brindley, C. Sun, R. Bittman, J. G. N. Garcia, et al.
Intracellular Generation of Sphingosine 1-Phosphate in Human Lung Endothelial Cells: ROLE OF LIPID PHOSPHATE PHOSPHATASE-1 AND SPHINGOSINE KINASE 1
J. Biol. Chem., May 11, 2007; 282(19): 14165 - 14177.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
281/50/38418    most recent
M601670200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Pilquil, C.
Right arrow Articles by Brindley, D. N.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Pilquil, C.
Right arrow Articles by Brindley, D. N.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2006 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement