Mechanisms of Regulation of Phospholipase D1 by Protein Kinase C α

It has been suggested that protein-protein interaction is important for protein kinase C (PKC) α to activate phospholipase D1 (PLD1). To determine the site(s) on PKC α that are involved in binding to PLD1, fragments containing the regulatory domain, catalytic domain and C1-C3 domain of PKC (cid:1) and activate PLD1 in vivo and in vitro. A C-terminal 23 amino acid (aa) deletion mutant of PKC inhibitor and a PKC α mutant deficient in kinase activity. The results indicated that phosphorylation was not required for activation of PLD1, but was associated with inactivation. between PLD1 and the two mutants. The results show that both mutants bound negligibly to PLD1 even in the presence of PMA stimulation. The phosphorylation of PLD1 by α was also studied using antibodies to phosphoSer, phosphoThr and phosphoTyr and the Thr


INTRODUCTION
Phospholipase D (PLD) 1 is a ubiquitous enzyme that hydrolyzes phosphatidylcholine to phosphatidic acid (PA) and choline (1). PA can be metabolized to diacylglycerol (DAG) by PA phosphohydrolase. PA and DAG are involved in receptor-mediated intracellular signal transduction, secretion, cytoskeletal reorganization and the respiratory burst (2). To date two isoforms of mammalian PLD (PLD1 and PLD2) have been cloned. These isoforms share about 50% amino acid similarity, but exhibit quite different regulatory properties (2). PLD1 has a low basal activity and responds to protein kinase C (PKC) and to members of the Rho and Arf families of small G proteins (3)(4)(5)(6), while PLD2 exhibits a high basal activity and shows little or no response to PKC, Rho or Arf in vitro (7)(8)(9). The intracellular localization of PLD1 remains ambiguous. Most reports indicate it is localized in the perinuclear region including the Golgi apparatus and some have reported its presence in caveolae (10)(11)(12).
PKCα belongs to the group of conventional group of PKC isoforms which are regulated by Ca 2+ , DAG and phosphatidylserine (PS). It has a regulatory domain, which includes the pseudosubstrate, C1 (phorbol, DAG binding) domain and C2 (calcium, PS binding) domain and a catalytic domain, which includes the C3 (ATP binding) domain and C4 kinase domain . PKCα is mainly located in the cytosol and can translocate to membrane fraction upon stimulation by phorbol esters or certain agonists (13). PKCα is considered to play a major role in PLD1 activation (2). However, the mechanism(s) involved in PLD1 activation is still not clear. Some studies have shown that PKCα activates PLD1 through a phosphorylation-independent mechanism, since PKCα could activate this PLD isoform in vitro without ATP (4,12,14) 7 û-û-û-û-û-û-DQGû-671) were generated by PCR with primers containing the 5' and 3' EcoRI sites. D481E, F663D and F663A mutants were generated using the QuikChange site directed mutagenesis kit from Stratagene. All constructs were sequenced to verify the coding regions and were well expressed in COS-7 cells.
Six well plates were seeded with 2x10 5 cells/well and 10 cm-dishes were seeded with 8x10 5 cells 24 h before transfection with FuGENE6 according to the manufacturer's instructions.
In vivo PLD Assay -After 5 h of transfection, cells in six well plates were serumstarved overnight (0.5% fetal bovine serum in DMEM) in the presence of 1 µCi/ml [ 3 H]myristic acid. PLD activity was assayed by incubating the cells with 0.3% 1-butanol for 20 min and measuring the formation of [ 3 H]PtdBut as a percentage of total labeled lipids as described before (23). Subcellular Fractionation -After transfection and starvation overnight, 10 cm dishes of COS-7 cells were washed once with ice-cold phosphate-buffered saline (PBS) and then harvested using lysis buffer (25 mM Hepes, pH 7.2, 10% glycerol, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol and protease inhibitor mixture). After 10 s sonication for two times, the cell lysate was first centrifuged at 500 xg for 10 min to remove unbroken cells. The supernatant was then spun at 120,000 xg for 45 min at 4 0 C to separate the cytosolic and crude membrane fractions.
In vitro PLD Assay -For in vitro assay, the cells were either untransfected or separately transfected with PLD1 or PKCα or its mutants. The control supernatant or that containing overexpressed PKCα or its mutants was used as the PKC fraction and the crude membranes containing PLD1 were resuspended in lysis buffer and used as PLD1 fraction. The PLD1 activity was measured by the formation of [ 3 H]PtdBut in vitro as described (14). Briefly, phospholipid vesicles generated from phosphatidylethanolamine/PIP 2 /PC(16:1.4:1) containing [palmitoyl-3 H]PC (0.5 µCi/reaction) were used with 1-butanol (0.6%) as substrate. The reaction mixtures were incubated at 37 o C for 30 min and stopped with chloroform/methanol/HCl (50:98:2). The lipids were extracted from the organic phase and resolved by thin layer chromatography. Bands co-migrating with a PtdBut standard were quantitated by liquid scintillation counting.
Immunoprecipitation and Western Blotting -COS-7 cells cultured in 10 cm plates were transfected and starved overnight as described above. The cells were washed once with ice-cold PBS and harvested using immunoprecipitation (IP) buffer containing 25 mM Hepes pH 7.2, 10% glycerol, 1 mM EDTA, 1 mM EGTA, 50 mM KCl, 10 mM NaF, 10 mM Na 4 P 2 O 7 , 1.2 mM Na 3 VO 4 , 1% NP-40 and protease inhibitors mixture. The cell suspensions were sonicated for 10 s and then spun at 120,000 xg for 45 min to pellet the detergent-insoluble fraction. The supernatant was then precleared by mixing it with 1 µg of affinity purified mouse IgG and 20 µl of a 1:1 slurry of protein G beads for 1h at 4 o C. The mixture was then spun and the supernatant was incubated with 2 µl of anti X-press antibody and 20 µl of protein G beads overnight. The immunoprecipitates were washed four times with the IP buffer and then resuspended in SDS sample buffer. The samples were analyzed by SDS-PAGE and transferred to polyvinylildene difluoride membranes (Immobilon-P, Millipore). The blots were then blocked with 1% BSA and incubated with primary antibody and then with horseradish peroxidase-conjugated secondary antibody. The bands were detected using ECL.  In vitro PLD1 assay results (Fig. 1E) were consistent with the in vivo PLD results i.e.

Regulatory, Catalytic and C1-C3 Domains and a C-terminal Deletion
QRQHRIWKH3.&.GRPDLQVDFWLYDWHG3/')LJV)DQG*VKRZWKHUHVXOWVRIELQGLQJWHVWV between PLD1 and PKCα or its domains. Before PMA stimulation, there was slight binding between PLD1 and PKCα, which was greatly increased by PMA. However, none of the domains was able to bind to PLD1 in the presence or absence of PMA. As shown in Fig. 1B, all the domains were well expressed, as was PLD1 (not shown).
Definition of the C-terminal Residues of PKCα Required for Activation and Binding of PLD1. Fig. 1 demonstrated that a C-terminal 23 aa deletion of PKCα resulted in a loss of its ability to bind and activate PLD1. To further define the residue(s) involved, C-terminal 1, 5, 6, 7, 8, 9, 10, 11 aa deletion mutants were made and shown to be well expressed in COS-7 cells (data not shown). Figure 2A shows the effects of the different PKC mutants on endogenous PLD activity in vivo. It is evident that, with deletion of 9 C-terminal residues (¨3.&α still retained its ability to activate PLD. However, a 10 aa deletion (¨FDXVHG3.&α to lose this. Similar results were found with overexpressed PLD1 in COS-7 cells (data not shown). To

Mutation of Phe 663 causes PKCα to lose its Ability to Activate and Bind PLD1. The
above results indicate that Phe 663 , which is 10 aa from the C-terminus is very important for PKCα to activate PLD1. To prove this, two single amino acid mutants (F663D and F663A) were made to test their effects on PLD1. Both mutants were expressed very well in COS-7 cells and translocated from the cytosol to membrane fraction after PMA stimulation (data not shown).
In vivo PLD assays showed that both mutants could not activate endogenous PLD activity in COS-7 cells in the presence or absence of PMA (Fig. 3A). Similar results were obtained when overexpressed PLD1 in vivo (data not shown). In vitro PLD assay results also showed that both mutants lost the ability to activate PLD1 (Fig. 3B). Similar results were obtained with higher

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binding between PLD1 and the two mutants. The results show that both mutants bound negligibly to PLD1 even in the presence of PMA stimulation. The phosphorylation of PLD1 by PKCα was also studied using antibodies to phosphoSer, phosphoThr and phosphoTyr and the results showed that only Thr residues were detectably phosphorylated after PMA stimulation mutants. The data show that PKCα greatly increased PLD1 phosphorylation upon PMA VWLPXODWLRQ7KHûPXWDQWFRXOGVWLOOSKRVSKRU\ODWH3/'XSRQ30$VWLPXODWLRQZKLOe the ûPXWDQWFRPSOHWHO\ORVWWKLVIXQFWLRQ7KHWZR3KH 663 mutants also showed greatly decreased greatly impaired PLD1 phosphorylation (data not shown). As expected from the findings with the deletion mutants, both the F663D and F663A mutants retained general phosphorylating ability (Fig. 1D). These results indicate that PLD1 is phosphorylated by PKCα upon PMA stimulation and indicate that binding is required for this phosphorylation.

Time Course of PLD1 Activity and Thr Phosphorylation after PMA Stimulation.
The above results indicated that binding was required for PKCα to activate PLD1 since all the deletions and mutations that caused PKCα to lose its ability to bind to PLD1 also caused a loss of activation. However, since phosphorylation of PLD1 by PKCα must involve some association between the two enzymes, it was ambiguous whether or not PLD1 phosphorylation was required for its activation. To further study the role of phosphorylation in PLD1 activation, the time courses of PLD1 activation and phosphorylation upon PMA stimulation were studied.
This required a modification of the usual protocol i.e. COS-7 cells were first treated with PMA for 1, 5, 15, 30 and 60 min and then 1-butanol was added and the cells were incubated for another 2 min. The results are shown in Figure 4. It is evident that PLD1 activity rose very rapidly after PMA stimulation, reaching a maximum in about 3-5 minutes. Thereafter the activity decreased to near basal (0.1% PtdBut) in 30 min. Figure 4B shows that PKCα rapidly translocated from the cytosol to membrane fraction within 1 min upon PMA stimulation and its membrane association increased over 30-60 min. Figure 4C shows the time course of PKCα binding with PLD1 after PMA treatment. The binding was also detectable at 1 min and increased during the 1 h experiment. Figure 4D shows that the PLD1 phosphorylation was not evident until 5 min, but then continuously increased during the 1 h incubation. The time course results indicated that the activation of PLD1 by PMA was much faster than its phosphorylation. This suggested that PLD1 activation was independent of its phosphorylation and raised the possibility that phosphorylation actually decreased the activity of the phospholipase.

The PKC Inhibitor Staurosporine and a Kinase-Deficient PKCα Mutant (D481E) both Eliminate PLD1 Phosphorylation by PMA while Blocking the Later Decline in
Activity. To see if phosphorylation of PLD1 does decrease its activity, two kinds of approaches were used: a PKC kinase inhibitor staurosporine and a kinase-deficient PKCα mutant (D481E) (24). Figure 5A shows that staurosporine strongly inhibited PLD1 phosphorylation induced by 3.&.LQWKHSUHVHQFHof PMA. The D481E mutant also showed barely detectable phosphorylation of PLD1 upon PMA stimulation (Fig. 5B), consistent with its lack of kinase activity. The time course experiments (Fig. 5C) showed that the inhibitor partially blocked the peak activation of PLD1 induced by PMA (0-10 min), and also diminished the later decline in activity (10-30 min). Similar results were seen in cells expressing endogenous PKC (data not smaller initial activation of PLD and a slower later decline in PLD activity (Fig. 5C). In an effort to explain the differences in peak PLD activity, the effects of staurosporine and the D481E mutation on the binding of PKCα to PLD1 were tested. Staurosporine caused minimal effect on the binding between PKCα and PLD1 in the absence of PMA, but partially reduced the association in the presence of the phorbol ester (Fig. 5D). Binding to PLD1 was also less with the D481E mutant compared with wild t\SH3.&.)LJ(

DISCUSSION
Binding between PKCα and PLD1 was first observed in COS-7 cells (17) and later in Swiss 3T3 fibroblasts (18). However, these studies did not examine the relationship between the binding and the activation of PLD1. In the present study, we provide much evidence that the activation in vitro (16) 2 , so we first studied the binding between PLD1 and this domain and showed no detectable binding. More PKCα fragments were made, including the catalytic domain and the C1-C3 fragment, but these fragments also failed to bind and activate PLD1. The fragments were shown to be functionally intact by either their membrane translocation in response to PMA or their retention of kinase activity. Thus these data indicated that PKC.ZLWK both the regulatory and catalytic domains intact was required for association and activation of PLD1.
Because PLD is membrane-associated, it is likely that the interaction between PKCα and PLD1 occurs at a membrane locus. Thus the C1 and C2 domains of the regulatory domain, ZKLFKDUHUHTXLUHGIRUPHPEUDQHWDUJHWLQJDQGWUDQVORFDWLRQRI3.&.-31) would seem to be essential for activation of PLD1. By this reasoning, the catalytic domain, which stays in the cytosol irrespective of PMA stimulation (data not shown), should be unable to activate PLD1.
The observation that the ¨PXWDQWZKLFKSRVVHVVHVWKHGRPDLQVIRUPHPEUDQHWDUJHWLQJZDV unable to activate PLD1, indicates that PKCα also requires a residue(s) in the C-terminus in order to bind and activate PLD1.
Previous reports showed that with deletion of up to 11 aa in the C-terminus, PKCα retained kinase activity (32), so we made more C-terminal truncation mutants to study their HIIHFWVRQ3/'DFWLYLW\7KHUHVXOWVVKRZHGWKDWWKHûPXWDQWstill bound to and activated 3/'ZKLOHWKHûPXWDQWGLGQRWDQGWKDWERWKPXWDQWVUHWDLQHGNLQDVHDFWLYLW\,QVXSSRUW of the conclusion that the C-terminal 10 aa position (Phe 663 ) is important for PKCα to activate PLD1, two mutants of this residue (F663D and F663A) also failed to bind and activate PLD1. In agreement with a previous study (32), the F663D, F663A and ¨PXWDQWVUHWDLQHGNLQDVH activity (Fig. 1D). Thus the inability of these mutants to phosphorylate PLD1 (Fig. 3D), reflects their inability to bind to the phospholipase (Fig. 2C and 3C).
The issue of whether or not phosphorylation is needed for PLD1 activation remains controversial. Several studies have shown that ATP is not needed for PKCα to activate PLD in vitro (4,12,(14)(15)(16). Also, Ser/Thr phosphatase treatment dephosphorylates PLD1 in vitro but does not inhibit its activity (17). However, there is also a report showing that PLD1 is phosphorylated during activation by PKCα (22). Trypsin treatment of the phosphorylated enzyme immunoprecipitated from cells treated with PMA, followed by two-dimensional peptide mapping revealed multiple P-peptides (22). Some of these overlapped with P-peptides generated from the phosphorylation of PLD1 by PKCα in vitro. These P-peptides were analyzed by mass spectrometry to reveal phosphorylation of PLD1 at residues Ser 2 , Thr 147 and Ser 561 (22).
Mutation of these to Ala resulted in a partial loss of PMA-stimulated PLD activity in vivo.
However, many other P-peptides were not analyzed, raising the question of what phosphorylation of these other residues would do to PLD1 activity. In addition, the effects of the PXWDWLRQVRQWKHDFWLYDWLRQRI3/'E\3.&.in vitro were not tested.
Our results showed that PLD1 becomes Thr-phosphorylated during PMA treatment cells

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answer the key question of whether or not this phosphorylation is required for PLD1 activation, we carried out time course experiments to measure the real-time PLD1 activity after PMA treatment (Fig. 4A) (32). Once PtdBut is formed in the PLD assay it is only slowly degraded, so that the standard assay, in which 1-butanol is added first and then PMA is added for different time lengths, only reflects the accumulation of PtdBut, not the real-time PLD1 activity. The realtime results indicated that the increase in PLD1 activity was much faster than the phosphorylation increase (Fig. 4A c.f. 4D). After the initial peak of PLD1 activation, there was a slower decrease in activity, consistent with the frequently observed phenomenon that PtdBut accumulation ceases after several minutes of treatment with PMA and some agonists when PLD activity is measured using the conventional assay (for references, see 33). Thus, the initial activity increase was not associated with detectable phosphorylation, whereas the subsequent activity decrease was correlated with increased phosphorylation (Fig. 4A,D). In other words, the results were consistent with PLD1 phosphorylation having an inhibitory effect on activity. PLD1 activity quickly reached its peak at 1 min, and membrane translocation and association of PKCα with PLD1 could be detected at that time (Fig. 4A, B, C). It therefore appears that the initial association of PKCα with PLD1 is sufficient for PLD1 to reach its full activation.
To further explore the relationship between PLD1 activation and phosphorylation, a PKC kinase inhibitor staurosporine and a kinase-deficient PKCα mutant (D481E) were used. As expected, staurosporine strongly inhibited PLD1 phosphorylation upon PMA stimulation (Fig.   5A). The D481E mutant also caused negligible phosphorylation of the enzyme (Fig. 5B) consistent with its lack of kinase activity. When the effects of these agents on the PLD1 activity time course in response to PMA were compared with wild type PKC.DORQHERWKWKHLQKLELWRU and the D481E mutation induced a lower peak of PLD1 activity, but then slowed the subsequent decline in activity (Fig. 5C). The latter results support the conclusion that phosphorylation of PLD1 inhibits its activity. However, the effects of both staurosporine and D481E on the PLD1 peak activity were not consistent with our proposed effect of phosphorylation. To study this, we explored the effects of these agents on the association between PLD1 and PKCα. Figure 5D shows that staurosporine partly decreased the association in the presence of PMA and Figure   In summary, our results indicate that both the regulatory and catalytic domains of PKCα are required for activation of PLD1 and that there is a required residue (Phe 663 ) in the Cterminus. Surprisingly, our data indicate that phosphorylation is not required for the stimulatory action of PKCα on the enzyme in vivo and suggest that phosphorylation is involved in the down regulation of PLD activation that is commonly seen at later times in cells treated with PMA and agonists. DMEM, Dulbecco's modified Eagle'smediuim.

2.
In a later study (28), the regulatory domain was also shown to activate PLD1 in vitro, but