Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2007 Nov 21.
Published in final edited form as: Methods Enzymol. 2007;434:305–315. doi: 10.1016/S0076-6879(07)34017-2

LIPID PHOSPHATE PHOSPHATASES FROM SACCHAROMYCES CEREVISIAE

George M Carman *, Wen-I Wu †
PMCID: PMC2083191  NIHMSID: NIHMS31321  PMID: 17954255

Abstract

DPP1-encoded and LPP1-encoded lipid phosphate phosphatases are integral membrane proteins in the yeast Saccharomyces cerevisiae. They catalyze the Mg2+-independent dephosphorylation of bioactive lipid phosphate molecules such as diacylglycerol pyrophosphate and phosphatidate. These enzymes possess a three-domain lipid phosphatase motif that is localized to the hydrophilic surface of the membrane. The lipid phosphate phosphatase activities of DPP1-encoded and LPP1-encoded enzymes are measured by following the release of water-soluble radioactive inorganic phosphate from chloroform-soluble radioactive lipid phosphate substrate following a chloroform/methanol/water phase partition. The DPP1-encoded enzyme, commonly referred to as diacylglycerol pyrophosphate phosphatase, is purified from wild-type S. cerevisiae membranes by detergent solubilization with Triton X-100 followed by chromatography with DEAE-cellulose (DE53), Affi-Gel blue, hydroxylapatite, and Mono Q. The purification scheme yields an essentially homogeneous enzyme preparation that is stable for several years upon storage at −80°. The properties of the DPP1-encoded and LPP1-encoded lipid phosphate phosphatase enzymes are summarized.

1. INTRODUCTION

Lipid phosphate phosphatase in the yeast Saccharomyces cerevisiae catalyzes the Mg2+-independent dephosphorylation of bioactive lipid phosphate molecules such as diacylglycerol pyrophosphate (DGPP) and phosphatidate (PA) (Furneisen and Carman, 2000; Toke et al., 1998, 1999a; Wu et al., 1996) (reactions 1 and 2):

  1. Diacylglycerol pyrophosphate → Phosphatidate + Pi

  2. Phosphatidate → Diacylglycerol + Pi

Essentially all of the lipid phosphate phosphatase activities in S. cerevisiae are encoded by the DPP1 (Toke et al., 1998) and LPP1 (Toke et al., 1999a) genes, with the former gene being the major contributor of this enzyme (Toke et al., 1999a). The DPP1- and LPP1-encoded enzymes are integral membrane proteins with six transmembrane-spanning regions; they are localized to the vacuole (Han et al., 2001, 2004) and Golgi (Huh et al., 2003) compartments of the cell, respectively. The lipid phosphate phosphatase enzymes possess a three-domain lipid phosphatase motif that is localized to the hydrophilic surface of the membrane (Han et al., 2004; Stukey and Carman, 1997; Toke et al., 1998, 1999a,b). This catalytic motif consists of the consensus sequences KxxxxxxRP (domain 1)—PSGH (domain 2)—SRxxxxxHxxxD (domain 3). The conserved arginine residue in domain 1 and the conserved histidine residues in domains 2 and 3 are essential for catalytic activity (Stukey and Carman, 1997; Toke et al., 1999b; Zhang et al., 2000).

The substrate specificity of the lipid phosphate phosphatases suggests that these enzymes are involved in signaling events rather than in phospholipid synthesis (Brindley et al., 2002; Sciorra and Morris, 2002). The lipid phosphate phosphatase enzymes may play a role in signal transduction by terminating signaling events of lipid phosphates. Because the products of the lipid phosphate phosphatases are also bioactive lipid molecules, they can initiate signal transduction by producing signaling molecules. Thus, the regulation of lipid phosphate phosphatase activities is likely to modulate the balance of the signaling molecules that are substrates and products in their reactions. The DPP1- and LPP1-encoded PA phosphatase activities are not responsible for de novo lipid synthesis; the PAH1-encoded Mg2+-dependent PA phosphatase is the responsible enzyme for this function in S. cerevisiae (Carman and Han, 2006; Han et al., 2006).

The DPP1-encoded enzyme (Toke et al., 1998), commonly referred to as DGPP phosphatase, has been purified to homogeneity by standard protein purification procedures (Wu et al., 1996). We describe here the purification of the enzyme.

2. PREPARATION OF RADIOLABELED SUBSTRATES

[β-32P]DGPP is synthesized enzymatically from PA and [γ-3P]ATP using Catharanthus roseus PA kinase as described by Wu et al. (1996). [32P]PA is synthesized enzymatically from diacylglycerol using Escherichia coli diacylglycerol kinase (Walsh and Bell, 1986) as described by Lin and Carman (1989). The labeled substrates are purified by thin-layer chromatography on potassium oxalate-treated silica gel 60 plates using the solvent system chloroform/acetone/methanol/glacial acetic acid/water (50:15:13:12:4) (Wissing and Behrbohm, 1993).

3. ASSAY METHODS

Lipid phosphate phosphatase activities are measured for 20 min at 30° by following the release of water-soluble [32P]Pi from chloroform-soluble [°-32P]DGPP (10,000 cpm/nmol) or [32P]PA (10,000 cpm/nmol) (Wu et al., 1996). The reaction mixture for DGPP phosphatase activity contains 50 mM citrate buffer (pH 5.0), 0.1 mM DGPP, 2 mM Triton X-100, 10 mM 2-mercaptoethanol, and enzyme protein in a total volume of 0.1 ml. The reaction mixture for PA phosphatase activity contains 50 mM Tris-maleate buffer (pH 6.5), 0.1 mM PA, 1 mM Triton X-100, 2 mM Na2EDTA, 10 mM 2-mercaptoethanol, and enzyme in a total volume of 0.1 ml. The reactions are terminated by the addition of 0.5 ml of 0.1 N HCl in methanol. Chloroform (1 ml) and 1 M MgCl2 (1 ml) are added, the system is mixed, and the phases are separated by 2 min of centrifugation at 100g. Ecoscint H (4 ml) is added to a 0.5-ml sample of the aqueous phase, and radioactivity is determined by scintillation counting. All enzyme assays are conducted in triplicate. A unit of enzymatic activity is defined as the amount of enzyme that catalyzes the formation of 1 µmol of product/min. Specific activity is defined as units per milligram of protein. Protein concentration is determined by the method of Bradford (1976) using bovine serum albumin as the standard.

4. GROWTH OF YEAST

Strain MATa ade5 (Culbertson and Henry, 1975), which shows normal regulation of phospholipid metabolism (Greenberg et al., 1982; Klig et al., 1985, 1988; Poole et al., 1986), is used for purification of the DPP1encoded DGPP phosphatase. Cultures are maintained on YEPD medium (1% yeast extract, 2% peptone, 2% glucose) plates containing 2% Bactoagar. For enzyme purification, cells are grown in YEPD medium at 30° to late exponential phase, harvested by centrifugation, and stored at −80° as described previously (Fischl and Carman, 1983).

5. PURIFICATION PROCEDURE

All steps are performed at 5°.

5.1. Preparation of cell extract

The cell extract is prepared from 200 g (wet weight) of cells by disruption with glass beads with a Bead-Beater (Biospec Products) in buffer A (50 mM Tris-maleate [pH 7.0], 1 mM Na2EDTA, 0.3 M sucrose, 10 mM 2-mercaptoethanol, and 0.5 mM phenylmethanesulfonyl fluoride) (Fischl and Carman, 1983). Unbroken cells and glass beads are removed by centrifugation at 1500g for 5 min.

5.2. Preparation of microsomal membranes

Microsomal membranes are isolated from the cell extract by differential centrifugation (Fischl and Carman, 1983). They are washed, resuspended in buffer B (50 mM Tris-maleate [pH 7.0], 10 mM MgCl2, 10mM 2-mercaptoethanol, 20% glycerol, and 0.5 mM phenylmethanesulfonyl fluoride), and frozen at −80° until used for purification.

5.3. Preparation of Triton X-100 extract

Microsomal membranes are suspended in buffer B containing 1% Triton X-100 at a final protein concentration of 10 mg/ml. The suspension is incubated for 1 h on a rotary shaker at 150 rpm. After the incubation, the suspension is centrifuged at 100,000g for 1.5 h to obtain the Triton X-100 extract (supernatant).

5.4. DE53 (DEAE53-cellulose) chromatography

A DE53 column (2.5 × 20.5 cm) is equilibrated with buffer C (50 mM Tris-maleate [pH 7.0], 10 mM MgCl2, 10mM 2-mercaptoethanol, 20% glycerol, and 1% Triton X-100). The Triton X-100 extract is applied to the column at a flow rate of 60 ml/h. The column is washed with 1 column volume of buffer C followed by elution of DGPP phosphatase activity in 6-ml fractions with 9 column volumes of a linear NaCl gradient (0–0.25 M) in buffer C. The peak of DGPP phosphatase activity elutes from the column at the beginning of the NaCl gradient. The most active fractions containing activity are pooled and used for the next step in the purification scheme.

5.5. Affi-Gel blue chromatography

An Affi-Gel blue column (2.0 × 16 cm) is equilibrated with buffer C. The DE53-purified enzyme is applied to the column at a flow rate of 30 ml/h. The column is washed with 1 column volume of buffer C followed by 2 column volumes of buffer C containing 0.3 M NaCl. DGPP phosphatase activity is eluted from the column in 3.5-ml fractions with 10 column volumes of a linear NaCl gradient (0.3–0.9 M) in buffer C. The peak of DGPP phosphatase activity elutes from the column at a NaCl concentration from 0.3 to 0.4 M. The most active fractions are pooled, and the enzyme preparation is desalted by dialysis against buffer D (10 mM potassium phosphate [pH 7.0], 10 mM MgCl2, 10mM 2-mercaptoethanol, 20% glycerol, and 1% Triton X-100).

5.6. Hydroxylapatite chromatography

A hydroxylapatite column (1.5 × 8.5 cm) is equilibrated with buffer D. The desalted Affi-Gel blue-purified enzyme is applied to the column at a flow rate of 20 ml/h. The column is washed with 1 column volume of buffer D, and DGPP phosphatase activity is eluted from the column in 2-ml fractions with 20 column volumes of a linear potassium phosphate gradient (10×150 mM) in buffer D. Two peaks of DGPP phosphatase activity elute from the column. The first peak (peak I) of activity elutes from the column at the beginning of the gradient, and the second peak (peak II) of activity elutes at a phosphate concentration of about 24 mM. The most active fractions from each peak are pooled and dialyzed against buffer C.

5.7. Mono Q I chromatography

A Mono Q column (0.5 × 5 cm) is equilibrated with buffer C. The hydroxylapatite-purified enzyme from peak I is applied to the column at a flow rate of 24 ml/h. The column is washed with 2 column volumes of buffer C. DGPP phosphatase activity is eluted from the column in 1-ml fractions with 40 column volumes of a linear NaCl gradient (0×0.3 M) in buffer C. The peak of DGPP phosphatase activity elutes from the column at a NaCl concentration of about 0.12 M. Fractions containing activity are pooled and stored at −80°. The purified enzyme is stable for at least 5 years.

5.8. Mono Q II chromatography

A second Mono Q column (0.5 × 5 cm) is equilibrated with buffer C. The hydroxylapatite-purified enzyme from peak II is applied to the column at a flow rate of 24 ml/h. The column is washed with 2 column volumes of buffer C. DGPP phosphatase activity is eluted from the column in 1-ml fractions with 40 column volumes of a linear NaCl gradient (0–0.3 M) in buffer C. The peak of DGPP phosphatase activity elutes from the column at the beginning of the NaCl gradient. Fractions containing activity are pooled and stored at −80°. The purified enzyme is completely stable for at least 5 years.

5.9. Enzyme purity

Mono Q I chromatography of the hydroxylapatite peak I enzyme results in isolation of an apparent homogeneous protein preparation as shown by SDS-PAGE. The minimum subunit molecular mass of the purified protein is 34 kDa. Mono Q II chromatography of the hydroxylapatite peak II DGPP phosphatase results in isolation of a protein preparation that contains a major protein doublet migrating at a molecular mass of 34 kDa. This DGPP phosphatase preparation also contains some minor protein contaminants. Overall, the hydroxylapatite peak I DGPP phosphatase is purified 33,333-fold over the cell extract with an activity yield of 0.73% to a final specific activity of 150 µmol/min/mg (Table 17.1). The hydroxylapatite peak II DGPP phosphatase is purified 24,666-fold over the cell extract to a final specific activity of 111 µmol/min/mg with an activity yield of 4% (Table 17.1).

Table 17.1.

Purification of DPP1-encoded DGPP phosphatase from Saccharomyces cerevisiaea

Purification step Total units (µmol/min) Protein (mg) Specific activity (units/mg) Yield (%) Purification (-fold)
1. Cell extract 49.4 10,957.6 0.0045 100 1
2. Microsomes 27.4 2,312.8 0.0118 55.5 2.6
3. Triton X-100 16.2 803.2 0.0201 32.8 4.5
4. DE53 5.6 15.41 0.363 11.3 80.6
5. Affi-Gel blue 5.5 2.62 2.10 11.1 466.6
6a. Hydroxylapatite 0.51 0.143 3.49 1.0 775.5
6b. Hydroxylapatite 2.36 0.185 12.7 4.7 2,822
7. Mono Q I 0.36 0.0024 150 0.73 33,333
8. Mono Q II 2.0 0.018 111 4.0 24,666
Open in a new tab
a

Data from Wu et al. (1996).

5.10. Identification of DPP1 and LPP1 genes

Protein sequencing analysis has shown that DGPP phosphatase enzymes purified from Mono Q I and from Mono Q II are the same. The differences in chromatographic behavior of the two DGPP phosphatase preparations might be because of a post-translational modification(s). However, studies to address this hypothesis have not been pursued. The N-terminal amino acid sequence (MNR VSFIKTPFNIGAKWRLE) and two internal amino acid sequences (QPVEGLPLDTLFTAK and FPPIDDPLPFKPLMD) derived from the pure DGPP phosphatase enzyme have been used to identify the DPP1 gene in the Saccharomyces Genome Database (Toke et al., 1998).

The LPP1 gene has been identified in the yeast database because its deduced protein sequence shows homology to the DPP1-encoded protein (Toke et al., 1999a). The expression of DPP1 and LPP1 genes on multicopy plasmids in S. cerevisiae results in enrichment of their encoded enzymes of 10- and 13-fold, respectively (Toke et al., 1998, 1999a). The expression of DPP1 and LPP1 in baculovirus-infected Sf9 insect cells provides 500- and 200-fold enrichments of the DPP1- and LPP1-encoded lipid phosphate phosphatase enzymes, respectively, when compared with that found in wild-type yeast cell extracts (Toke et al., 1998, 1999a). Accordingly, the overexpression of DPP1- and LPP1-encoded lipid phosphate phosphatase enzymes either in yeast or in insect cells should facilitate the purification of these enzymes.

6. PROPERTIES OF DPP1- AND LPP1-ENCODED LIPID PHOSPHATE PHOSPHATASES

Properties of the DPP1-encoded lipid phosphate phosphatase have been examined using purified enzyme (Toke et al., 1998, 1999a; Wu et al., 1996), whereas properties of the LPP1-encoded lipid phosphate phosphatase have been examined using membranes isolated from Sf9 insect cells (Furneisen and Carman, 2000). These enzymes do not have a divalent cation requirement for activity (Furneisen and Carman, 2000; Wu et al., 1996); this is consistent with the enzymes being members of the superfamily of enzymes possessing the lipid phosphatase motif (Han et al., 2004; Stukey and Carman, 1997; Toke et al., 1999b). The kinetic properties of the lipid phosphate phosphatase enzymes utilizing DGPP and PA are summarized in Table 17.2. The specificity constant (Vmax/Km) of the DPP1-encoded enzyme for DGPP is 10-fold greater than that of PA (Wu et al., 1996). DGPP is a very potent inhibitor of the PA phosphatase activity of the DPP1-encoded enzyme, whereas PA does not inhibit the DGPP phosphatase activity of the enzyme (Wu et al., 1996). The specificity constant of the LPP1-encoded enzyme for PA is slightly higher (1.2-fold) than that for DGPP (Furneisen and Carman, 2000). DGPP and PA inhibit PA phosphatase and DGPP phosphatase activities, respectively, of the LPP1-encoded enzyme with equal potencies (Furneisen and Carman, 2000). The affinity (reflected in Km value) of the LPP1-encoded enzyme for PA and DGPP as substrates is greater than the affinity of the DPP1-encoded enzyme for these substrates. Purified DPP1-encoded lipid phosphate phosphatase also utilizes lysoPA (Dillon et al., 1996), phosphatidylglycerophosphate (Dillon et al., 1996), sphingoid base phosphates (Dillon et al., 1997), and isoprenoid phosphates (Faulkner et al., 1999) as substrates. The LPP1-encoded lipid phosphate phosphatase will also utilize lysoPA as a substrate (Furneisen and Carman, 2000). Although these enzymes utilize a variety of lipid phosphate substrates, only DGPP and PA have been shown to be substrates in vivo (Toke et al., 1999a).

Table 17.2.

Kinetic constants for DPP1- and LPP1-encoded lipid phosphate phosphatases

DPP1-encoded lipid phosphatasea
 LPP1-encoded lipid phosphataseb
Substrate or inhibitor Vmax (unitsc/mg) Km (mol%) Vmax/Km(units/mg/mol%) Ki (mol%) Vmax (unitsc/mg) Km (mol%) Vmax/Km (units/mg/mol%) Ki(mol%)
    1.1.1.1       1.1.1.2    
DGPP 172 0.55 313 0.35d 0.244 0.07 3.5 0.12d
PA 70 2.2 32 NIe 0.210 0.05 4.2 0.12f
Open in a new tab
a

Data for the DPP1-encoded enzyme were taken from Wu et al. (1996).

b

Data for the LPP1-encoded enzyme were taken from Furneisen and Carman (2000). Because the LPP1-encoded enzyme is not pure, the specificity constants reported for the enzyme cannot be compared with those reported for the DPP1-encoded enzyme.

c

Unit of activity defined as µmol/min.

d

Inhibitor constant with respect to PA as a substrate.

e

NI, not inhibitory.

f

Inhibitor constant with respect to DGPP as a substrate.

DPP1- and LPP1-encoded enzymes differ with respect to their sensitivity to thioreactive compounds. In particular, the LPP1-encoded enzyme is potently inhibited by N-ethylmaleimide (Furneisen and Carman, 2000), whereas the DPP1-encoded enzyme is insensitive to this compound (Wu et al., 1996). This difference in N-ethylmaleimide sensitivity may be based on the fact that the LPP1-encoded enzyme has 10 cysteine residues, whereas the DPP1-encoded enzyme contains only 3.

ACKNOWLEDGMENT

This work was supported in part by United States Public Health Service, National Institutes of Health Grant GM-28140.

REFERENCES

  1. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976;72:248–254. doi: 10.1016/0003-2697(76)90527-3. [DOI] [PubMed] [Google Scholar]
  2. Brindley DN, English D, Pilquil C, Buri K, Ling ZC. Lipid phosphate phosphatases regulate signal transduction through glycerolipids and sphingolipids. Biochim. Biophys. Acta. 2002;1582:33–44. doi: 10.1016/s1388-1981(02)00135-x. [DOI] [PubMed] [Google Scholar]
  3. Carman GM, Han GS. Roles of phosphatidate phosphatase enzymes in lipid metabolism. Trends Biochem. Sci. 2006;31:694–699. doi: 10.1016/j.tibs.2006.10.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Culbertson MR, Henry SA. Inositol requiring mutants of Saccharomyces cerevisiae. Genetics. 1975;80:23–40. doi: 10.1093/genetics/80.1.23. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Dillon DA, Chen X, Zeimetz GM, Wu W-I, Waggoner DW, Dewald J, Brindley DN, Carman GM. Mammalian Mg2+-independent phosphatidate phosphatase (PAP2) displays diacylglycerol pyrophosphate phosphatase activity. J. Biol. Chem. 1997;272:10361–10366. doi: 10.1074/jbc.272.16.10361. [DOI] [PubMed] [Google Scholar]
  6. Dillon DA, Wu W-I, Riedel B, Wissing JB, Dowhan W, Carman GM. The Escherichia coli pgpB gene encodes for a diacylglycerol pyrophosphate phosphatase activity. J. Biol. Chem. 1996;271:30548–30553. doi: 10.1074/jbc.271.48.30548. [DOI] [PubMed] [Google Scholar]
  7. Faulkner AJ, Chen X, Rush J, Horazdovsky B, Waechter CJ, Carman GM, Sternweis PC. The LPP1 and DPP1 gene products account for most of the isoprenoid phosphatase activities in Saccharomyces cerevisiae. J. Biol. Chem. 1999;274:14831–14837. doi: 10.1074/jbc.274.21.14831. [DOI] [PubMed] [Google Scholar]
  8. Fischl AS, Carman GM. Phosphatidylinositol biosynthesis in Saccharomyces cerevisiae: Purification and properties of microsome-associated phosphatidylinositol synthase. J. Bacteriol. 1983;154:304–311. doi: 10.1128/jb.154.1.304-311.1983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Furneisen JM, Carman GM. Enzymological properties of the LPP1-encoded lipid phosphatase from Saccharomyces cerevisiae. Biochim. Biophys. Acta. 2000;1484:71–82. doi: 10.1016/s1388-1981(99)00202-4. [DOI] [PubMed] [Google Scholar]
  10. Greenberg M, Goldwasser P, Henry SA. Characterization of a yeast regulatory mutant constitutive for inositol-1-phosphate synthase. Mol. Gen. Genet. 1982;186:157–163. doi: 10.1007/BF00331845. [DOI] [PubMed] [Google Scholar]
  11. Han G-S, Johnston CN, Carman GM. Vacuole membrane topography of the DPP1-encoded diacylglycerol pyrophosphate phosphatase catalytic site from Saccharomyces cerevisiae. J. Biol. Chem. 2004;279:5338–5345. doi: 10.1074/jbc.M311779200. [DOI] [PubMed] [Google Scholar]
  12. Han G-S, Johnston CN, Chen X, Athenstaedt K, Daum G, Carman GM. Regulation of the Saccharomyces cerevisiaeDPP1-encoded diacylglycerol pyrophosphate phosphatase by zinc. J. Biol. Chem. 2001;276:10126–10133. doi: 10.1074/jbc.M011421200. [DOI] [PubMed] [Google Scholar]
  13. Han G-S, Wu W-I, Carman GM. The Saccharomyces cerevisiae lipin homolog is a Mg2+-dependent phosphatidate phosphatase enzyme. J. Biol. Chem. 2006;281:9210–9218. doi: 10.1074/jbc.M600425200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Huh WK, Falvo JV, Gerke LC, Carroll AS, Howson RW, Weissman JS, O’Shea EK. Global analysis of protein localization in budding yeast. Nature. 2003;425:686–691. doi: 10.1038/nature02026. [DOI] [PubMed] [Google Scholar]
  15. Klig LS, Homann MJ, Carman GM, Henry SA. Coordinate regulation of phospholipid biosynthesis in Saccharomyces cerevisiae: Pleiotropically constitutive opi1 mutant. J. Bacteriol. 1985;162:1135–1141. doi: 10.1128/jb.162.3.1135-1141.1985. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Klig LS, Homann MJ, Kohlwein SD, Kelley MJ, Henry SA, Carman GM. Saccharomyces cerevisiae mutant with a partial defect in the synthesis of CDP-diacylglycerol and altered regulation of phospholipid biosynthesis. J. Bacteriol. 1988;170:1878–1886. doi: 10.1128/jb.170.4.1878-1886.1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Lin Y-P, Carman GM. Purification and characterization of phosphatidate phosphatase from Saccharomyces cerevisiae. J Biol. Chem. 1989;264:8641–8645. [PubMed] [Google Scholar]
  18. Poole MA, Homann MJ, Bae-Lee M, Carman GM. Regulation of phosphatidylserine synthase from Saccharomyces cerevisiae by phospholipid precursors. J. Bacteriol. 1986;168:668–672. doi: 10.1128/jb.168.2.668-672.1986. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Sciorra VA, Morris AJ. Roles for lipid phosphate phosphatases in regulation of cellular signaling. Biochim. Biophys. Acta. 2002;1582:45–51. doi: 10.1016/s1388-1981(02)00136-1. [DOI] [PubMed] [Google Scholar]
  20. Stukey J, Carman GM. Identification of a novel phosphatase sequence motif. Protein Sci. 1997;6:469–472. doi: 10.1002/pro.5560060226. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Toke DA, Bennett WL, Dillon DA, Chen X, Oshiro J, Ostrander DB, Wu W-I, Cremesti A, Voelker DR, Fischl AS, Carman GM. Isolation and characterization of the Saccharomyces cerevisiae DPP1 gene encoding for diacylglycerol pyrophosphate phosphatase. J. Biol. Chem. 1998;273:3278–3284. doi: 10.1074/jbc.273.6.3278. [DOI] [PubMed] [Google Scholar]
  22. Toke DA, Bennett WL, Oshiro J, Wu WI, Voelker DR, Carman GM. Isolation and characterization of the Saccharomyces cerevisiae LPP1 gene encoding a Mg2+-independent phosphatidate phosphatase. J. Biol. Chem. 1999a;273:14331–14338. doi: 10.1074/jbc.273.23.14331. [DOI] [PubMed] [Google Scholar]
  23. Toke DA, McClintick ML, Carman GM. Mutagenesis of the phosphatase sequence motif in diacylglycerol pyrophosphate phosphatase from Saccharomyces cerevisiae. Biochemistry. 1999b;38:14606–14613. doi: 10.1021/bi991472x. [DOI] [PubMed] [Google Scholar]
  24. Walsh JP, Bell RM. sn-1, 2-diacylglycerol kinase of Escherichia coli: Mixed micellar analysis of the phospholipid cofactor requirement and divalent cation dependence. J. Biol. Chem. 1986;261:6239–6247. [PubMed] [Google Scholar]
  25. Wissing JB, Behrbohm H. Diacylglycerol pyrophosphate, a novel phospholipid compound. FEBS Lett. 1993;315:95–99. doi: 10.1016/0014-5793(93)81141-l. [DOI] [PubMed] [Google Scholar]
  26. Wu W-I, Liu Y, Riedel B, Wissing JB, Fischl AS, Carman GM. Purification and characterization of diacylglycerol pyrophosphate phosphatase from Saccharomyces cerevisiae. J. Biol. Chem. 1996;271:1868–1876. doi: 10.1074/jbc.271.4.1868. [DOI] [PubMed] [Google Scholar]
  27. Zhang QX, Pilquil CS, Dewald J, Berthiaume LG, Brindley DN. Identification of structurally important domains of lipid phosphate phosphatase-1: Implications for its sites of action. Biochem. J. 2000;345(Pt 2):181–184. [PMC free article] [PubMed] [Google Scholar]

RESOURCES