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Lysophosphatidic acid (LPA) is a lipid mediator that stimulates cell proliferation and growth and is involved in physiological and pathological processes such as wound healing, platelet activation, angiogenesis and the growth of tumors. Therefore, defining the mechanisms of LPA production and degradation are of interest in understanding the regulation of these processes. Extracellular LPA synthesis is relatively well understood whereas the mechanisms of its degradation are not. One route of LPA degradation is de-phosphorylation. A candidate enzyme is the integral membrane exophosphatase lipid phosphate phosphohydrolase type 1 (LPP1). We report here the development of a mouse wherein the LPP1 gene (Ppap2a) was disrupted. The homozygous mice, which are phenotypically unremarkable, generally lack LPP1 mRNA and multiple tissues exhibit a substantial (35–95%) reduction in LPA phosphatase activity. Compared to wild type littermates, Ppap2atr/tr animals have increased levels of plasma LPA and LPA injected intravenously is metabolized at a four-fold slower rate. Our results demonstrate that LPA is rapidly metabolized in the bloodstream and that LPP1 is an important determinant of this turnover. These results indicate that LPP1 is a catabolic enzyme for LPA in vivo.
Lysophosphatidic acid (LPA)1 (1-acyl-2-lyso-sn-glycero-3-phosphate) has long been known as an intermediate in de novo biosynthesis of all glycerophospholipids. LPA gained credence as a signaling molecule with the discovery in the early 1990’s by Moolenaar and collaborators that LPA promotes cell proliferation (see  for a review). LPA and other biologically active lipid phosphates, notably sphingosine 1-phosphate (S1P), have since been studied extensively and found to have a role as mediators in a variety of physiologic and pathologic processes. The stimulatory effect of LPA on cell proliferation, survival and migration serves as the underlying mechanism of the major biological effects of LPA. The most conspicuous examples are the involvement of LPA in wound healing, platelet activation, vascular remodeling and the progression of some forms of cancer such as ovarian tumors (see  for a review).
The role of LPA and other lipid phosphates as first messengers was firmly established by the realization that many of the biological effects of these molecules are mediated by their interaction with specific, seven-transmembrane domain G protein-coupled receptors (see  for a review). As with any bona fide signaling molecule, specific mechanisms are expected to exist that ensure its timely destruction to prevent the detrimental effects of overstimulation. LPA catabolism is presumed to be mediated by a group of phosphatases termed lipid phosphate phosphohydrolases (LPPs) rather than phosphatidate phosphatase-1 (lipins) that are specific for phosphatidate [4,5].
LPPs (formerly known as phosphatidic acid phosphatases (PAPs) (see [6,7] for reviews) are enzymes that catalyze the hydrolysis of a variety of lipid phosphate mono-esters. The LPPs are Mg2+-independent, NEM-insensitive (LPP1, LPP2 and LPP3 formerly known PAP2A, PAP2B and PAP2C respectively; see  for a review). LPP1-3 are cell surface, N-glycosylated, integral membrane proteins. LPP3 in some cell types also localizes to intracellular membranes . The topology of membrane LPPs is such that the active site amino acids are exofacial, thus these enzymes function as exophosphatases .
The hypothesis that one or more LPP isotypes may function to delimit LPA signaling has been difficult to test. On the one hand, LPP1 and LPP3 are nearly ubiquitous among mammalian cells (see  for a review) and forced expression of these proteins is either toxic to cells or provide information restricted to cells in culture under somewhat artificial conditions [12,13]. On the other hand, the difficulties inherent in the study of integral membrane proteins – a lack of quality antibodies and purified protein – have slowed progress in understanding these enzymes. A chemical biology approach has not been particularly informative in that selective inhibitors have yet to be developed and structure activity relationship of substrates is rather uninformative. That is, the LPP isotypes act on a wide variety of lipid mono-phosphates including LPA, PA, diacylglycerolpyrophosphate, S1P and ceramide 1-phosphate. Using in vitro assays, LPP1 exhibits a preference for glycerol-versus sphingoid base-containing lipids, while LPP3 does not differentiate as well among these substrates (see  for a review). These ecto-activities of LPP1 have two potentially important functions. First, they could regulate circulating concentrations of LPA or S1P and thus the activation of their respective receptors. Decreased LPP activities occur in tumors and this has been proposed as a mechanism that results in increased LPA-induced growth in ovarian cancers . Secondly, the dephosphorylated product formed by the LPP is taken up by the cell and thereby it stimulates cell signaling itself or after re-phosphorylation. [5,6,15,16].
An alternative approach – genetic manipulation – has yielded several studies reporting the consequences of altering the expression of LPP genes. Disruption of the mouse gene encoding LPP3 (Ppap2b) is embryonic lethal (E9.5), apparently as a consequence of aberrant embryonic and extra-embryonic development of the vasculature . By contrast, mice lacking LPP2 (Ppap2c −/−) are viable, fertile and not noticeably different from wild type littermates . Ablation of the Drosophila homologs of the mammalian LPPs (wun and wun2) results in a defect in germ cell migration, suggesting that a spatial pattern of phospholipid hydrolysis is required for the migration and survival of germ cells during fly development . Global forced expression of LPP1 (Ppap2a) in mice resulted in runted animals with fur abnormalities and impaired spermatogenesis, a phenotype that is not readily ascribed to increased extracellular LPA or PA degradation. Interestingly, theses LPP1 transgenic mice had normal levels of plasma LPA .
In this communication we report the generation and characterization of a mouse strain wherein the gene encoding LPP1 (Ppap2a) is disrupted by insertion of an exon-trapping element. Mice with one (Ppap2a+/tr) or both (Ppap2atr/tr) alleles disrupted are viable and fertile. We used these mice as an in vivo model to establish whether LPP1 plays a physiological role in controlling the degradation of circulating LPA. Our results demonstrate that LPP1 performs this function in vivo.
An embryonic stem (ES) cell line (RRD231) reported to harbor a gene trap vector (pGT1Lxf) in the first intron of the Ppap2a gene were obtained from BayGenomics via the Mutant Mouse Regional Resource Center (MMRRC) at Davis, CA. We verified the presence of the trapping element and further established its precise location by sequencing genomic DNA from Ppap2atr/tr mice (see Fig. 1). Chimeric founder mice were generated at the University of Virginia Transgenic Mouse facility using C57BL/6 blastocysts. The ES cells were sv129 strain, animals used in this study were of mixed genetic background (F1N1, F1N5).
Mice were genotyped by PCR using genomic DNA from tail biopsies, liver or brain samples and the following primers (see Fig. 1): 5′-GAGAGTGAGCGAGTGTCTGAGTTTCTGATG-3′ (forward), 5′-AGTACTGGGCATCTCACACCACAT-3′ (reverse wild type allele), 5′-CCTTCAAAGGGAAGGGGTAAAGTGGTAGGG-3′ (reverse trapped allele). Amplification in presence of 20 ng DNA and 4 mM MgCl2 was carried out as follows: at 94 °C for 3 min, followed by 35 cycles at 94 °C (30 sec), 57 °C (1 min), 72 °C (1.5 min); and a final 72 °C step for 10 min.
RNA was extracted from tissues and cDNA was obtained using the SuperScript First-Strand Synthesis System (Invitrogen) using random hexamers according to the manufacturer’s instructions. Amplification was performed in a iCycler iQ System (Bio-Rad, Hercules, CA) at 95 °C for 4 min, followed by 40 cycles at 95 °C, 55 °C, and 72 °C (30 s each step) in 50 μl of a SYBR Green-based medium (iQ SYBR Green Supermix; Bio-Rad) using the following forward and reverse primers: 5′-TGTACTGCATGCTGTTTGTCGCAC-3′, 5′-TGACGTCACTCCAGTGGTGTTTGT-3′ for LPP1; and 5′-ATAAACGATGCTGTGCTCTGTGCG-3′, 5-TTTGCTGTCTTCTCCTCTGCACCT-3′ for LPP3. The levels of mRNA expression were normalized to the expression level of the 18s ribosomal RNA gene measured using a commercial primer kit (QuantumRNA, Classic II; Ambion, Austin, TX). Quantitative gene expression was obtained from cycle differences at appropriate thresholds and the efficiency of amplification as described by Pfaffl . The size and singularity of the RT-PCR products was established by agarose gel electrophoresis and melting point analysis. For the detection of the different LPP1 isoforms, a similar protocol was followed but using the following forward and reverse primers: 5′-ATCCATTTCAGAGGGGCTTT-3′, 5′-AACCTGCCCTCCTTGACTTT-3′ for isoform 1; and 5′-TTCAAGGCATACCCCCTTC-3′, 5′-GGTGGCTATGTAGGGATTGC-3′ for isoform 2. To amplify the cDNA region containing the trap insertion point in intron 2, a similar protocol was used except for annealing at 53 °C and the use of the following primer 5′-TCTGTTCCTCCCGCCACT-3′ in conjunction with the reverse primer used for LPP1 isoform 1.
[32P]-labeled LPA and PA were synthesized as previously described  using E. coli diacylglycerol kinase (Sigma D3065), ATP (Sigma A2383), γ-[32P]ATP (MP Biochemicals 35001; 7,000 Ci/mmol), cardiolipin (Sigma C0563), and 1-monooleoyl-glycerol (Sigma M-7765) or 1,2-dioleoyl-glycerol (Avanti Polar Lipids 800811C). Final purification by thin layer chromatography (TLC) was carried out on Whatman Silica Gel plates (4865-621) using a mobile phase consisting of 1-butanol:acetic acid:H2O 3:1:1. Small aliquots of the labeled compounds dissolved in chloroform were stored in light-protected, chloroform-rinsed glass vials at −20 °C for no more than four days. No radiolysis was observed under these conditions as judged by TLC analysis.
Mice were anesthetized with isofluorane and sacrificed by cervical dislocation. Organs were immediately obtained, frozen in liquid nitrogen and stored at −80 °C. Brain samples consisted of tissue from the cerebral hemispheres. Skeletal muscle samples were obtained from the hind legs. Frozen tissue samples were homogenized using a motor driven Teflon pestle homogenizer in 10 volumes of 20 mM Tris-HCl, pH 7.5, 1 mM EGTA, 1 mM PMSF, 1mM DTT, 10 μg/ml aprotinin (Sigma A6279), 10 μg/ml leupeptin (Sigma L2023). Homogenates were centrifuged at 500 × g and the supernatant fluids were aliquoted and stored at −80 °C (pellets were discarded). The protein content of these preparations was measured according to Bradford  using a commercial Coomassie blue solution (Bio-Rad). LPP activity was measured as the release of [32P]H3PO4 from [32P]-labeled substrates presented as mixed Triton X-100 micelles as previously described  with some modifications. Assays were carried out in 200 μl of a buffer consisting of 20 mM Tris-HCl, pH 7.5, 1 mM MgCl2, 1 mM DTT, 3.2 mM Triton X-100, 100 μM LPA or PA and 50,000 dpm [32P]LPA or [32P]PA. Assays were carried out at 37 °C in presence of 100 μg protein for different times, 5 min to 30 min, according to the specific LPP activity of each tissue so that degradation did not exceed 5% of the total substrate. Different incubation times were preferred over incubation with different amounts of protein to preserve the ratio Triton X-100/protein content. Reactions were started by mixing 100 μl of mixed micelles with 100 μl of a solution containing the remaining components. Activity was constant with time for every tissue. No LPP activity was observed in absence of tissue. Reaction was stopped by adding 200 μl of 1 M HClO4 containing 100 μM phosphoric acid. Samples were then centrifuged for 5 min at 15,000 × g and the supernatant extracted twice with H2O-saturated 1-butanol. Orthophosphate was recovered from the extracted aqueous phase as described  by precipitation with 12.5 mM ammonium molybdate and extraction with isobutanol:benzene 1:1. Radionuclide in the organic phase was measured by liquid scintillation spectrometry. Specific activity of the substrates was determined by measuring the amount of 32P present in known volumes of reaction media. To measure the NEM-sensitive, Mg2+-independent activity, tissue samples containing 100 μg of protein were brought to 80 μl and supplemented with 10 μl of 50 mM NEM. After a 15 min incubation, NEM was neutralized by adding 10 μl of 22 mM DTT (1mM final DTT). This solution was then added to 100 μl of mixed micelles and processed as described above except for the omission of MgCl2 and the addition of 1 mM EGTA and 2 mM EDTA .
Ppap2a+/+ and Ppap2atr/tr mice were anesthetized with isofluorane and sacrificed by cervical dislocation. The brain hemispheres, kidney, liver and spleen were removed, placed in ice cold Buffer A (140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 5 mM glucose, 20 mM Hepes-NaOH, pH 7.4) and sliced into 300 μm thick slices using a Vibratome type slicer (Dosaka DTK 1500E microslicer). Slices were trimmed to approximately 3 mm × 3 mm squares and incubated in 200 μl of Buffer A containing 0.1% Fatty Acid Free bovine serum albumin (FAF-BSA, Sigma A6003), 100 μM LPA and approximately 2 × 105 cpm [32P]LPA for variable times at 37 °C. At the end of the incubation periods, 100 μl aliquots were withdrawn and the presence of [32P]H3PO4 was assayed as described for tissue homogenates. The remaining 100 μl were discarded and the tissue dissolved in 2% SDS and the protein content measured by the bicinchonic acid method using a commercial kit (Pierce 23223). Activity was linear with time. Linearity with protein concentration was tested only for slices of the same thickness and roughly the same size and shape. In particular, we carried out incubations with half, one or two 3 mm × 3 mm slices and found that activity and protein were linearly correlated. For slices of different thickness, activity and protein were not linearly correlated, therefore and to ensure thickness homogeneity similar organs from Ppap2a+/+ and Ppap2atr/tr were mounted together and sliced simultaneously. Splenocytes were obtained by passing the spleens through a nylon cell strainer (100 μm mesh, Falcon 352360) and discarding the capsule. Cells were resuspended in 1 ml of Buffer A and centrifuged at 300 × g for 5 min. The pellet was resuspended in 1 ml of erythrocyte lysis buffer (8.29 g/l NH4Cl, 1 g/l KHCO3, 0.0372 g/l Na2EDTA, pH 7.3) and incubated on ice for 5 min. Lysis was quenched by adding 9 ml of Buffer A and washing the cell preparation twice (at 300 × g for 5 min each). Finally cells were resuspended cells in 1 ml Buffer A and counted in a hemocytometer. Viability, assessed by Trypan Blue exclusion, was > 99%. LPP activity was measured as described above and started by resuspending pellets (300 × g for 5 min) containing 5 × 105 lysed or non-lysed cells (approximately 100 and 20 μg of protein respectively) in 200 μl of reaction medium. In both cases, slices and splenocytes, incubation times were identical for similar tissues from Ppap2a+/+ and Ppap2atr/tr mice and they were adjusted for substrate degradation not to exceed 5%. No release of [32P]H3PO4 was observed in absence of slices or cells.
Plasma was obtained by cardiac puncture using EDTA (~ 5 mM) as anticoagulant. LPA levels were determined by liquid chromatography-tandem mass spectrometry using a hybrid ABI-4000 triple quadrupoleion trap mass spectrometer (Applied Biosystems, Foster City, CA) coupled with an Agilent 1100 liquid chromatography column and C17:0 LPA as an internal standard . LPA species were separated on a Zorbax Eclipse XDB-C8 HPLC column (4.6 × 150 mm, 5 μm) using methanol/water/HCOOH, 79:20:0.5 v/v as solvent A and 99:0.5:0.5 v/v as solvent B, containing in both cases 5 mM NH4COOH. Elution was carried out in solvent A for 1 min. A gradient change to solvent B lasting 1 min was then effected and a final, isocratic elution of LPA was carried out in solvent B for 7 min. LPA species were analyzed in negative ionization mode with declustering potential and collision energy optimized for 17:0, 18:0, 18:1, 18:2 and 20:4 LPA. Multiple reaction monitoring parameters for nine other LPA molecular species were selected with the closest possible approximation to the available LPA standards. The following transitions, indicated as the m/z values and, in parentheses, chain length: number of double bonds, were monitored: 407.0/153.0 (16:1); 409.0/153.1 (16:0); 423.0/153.1 (17:0, an unnatural species used as an internal standard); 431/153.0 (18:3); 433.0/153.0 (18:2); 435.1/152.9 (18:1); 437.0/153.0 (18:0); 455.1/153.0 (20:5); 457.0/153.0 (20:4); 459.1/153.0 (20:3); 461.1/153.0 (20:2); 481.1/153.0 (22:6); 483.1/153.0 (22:5); and 485.1/153.0 (22:4).
A series of Ppap2a+/+ and Ppap2atr/tr mice were anesthetized and exsanguinated by cardiac puncture using EDTA (~5 mM) as anticoagulant. Plasma was prepared as previously described  by centrifuging the blood twice, at 1,000 and 10,000 × g for 3 min each time at 4 °C, and stored at −80 °C. Aliquots of these plasma preparations were used as a vehicle for [32P]LPA to study LPA clearance as described below. Plasma used in clearance experiments was in each case of the same genotype as the animals whose clearance was being investigated.
Blood samples from a series of Ppap2a+/+ and Ppap2atr/tr mice were obtained and anticoagulated with EDTA (~ 5 mM). Half of these samples were used as whole blood and the other half used as plasma (obtained as described above). Blood and plasma samples were supplemented as soon as they were obtained with [32P]LPA dissolved in either plasma (see above) or saline (0.9 % NaCl) containing 0.1% FAF-BSA. Incorporation of [32P]LPA into plasma or saline solution was carried out by first drying in a glass vial an appropriate amount of [32P]LPA dissolved in chloroform and then dissolving the dried [32P]LPA into the appropriate vehicles (plasma or saline). Two hundred μl blood samples and 100 μl plasma samples were supplemented with plasma amounting to 5% of their volume (10 and 5 μl respectively) and containing approximately 105 cpm [32P]LPA. Immediately after the addition of [32P]LPA (“time zero” sample) or after 5, 15 and 30 min incubation at 37 °C, [32P]LPA was measured as follows: plasma samples and plasma obtained from blood samples as described above were immediately acidified with 3.33 volumes of ice-cold 30 mM citric acid, 40 mM Na2HPO4, pH 4.0, and extracted twice with 8.88 volumes of H2O-saturated butanol. Butanol extractions were then combined, dried under a stream of N2, re-dissolved in chloroform and subjected to TLC as described above. Radioautographic analysis of the developed plates showed only one band that comigrates with authentic LPA (Rf = 0.5). LPA was then measured by scrapping off the LPA bands from the developed TLC plates and measuring their 32P content by liquid scintillation spectrometry. The “time zero” plasma samples were also used to assess the quantitative recovery of LPA from plasma by comparing the amount of [32P]LPA added to the plasma samples with the amount of [32P]LPA recovered from TLC plates. Similar to the previously reported 99% recovery figure for this technique , our recovery was not significantly different from 100%.
Mice (24–27 g) were anesthetized with methoxyfluorane and injected through the tail vein with approximately 108 cpm [32P]LPA (6.5 × 10−12 mole) dissolved in 150 μl of either plasma (see above) or sterile saline (0.9 % NaCl) containing 0.1% FAF-BSA. Immediately after injection, a blood sample (3 to 6 drops or approximately 50–100 μl) was taken by retro orbital bleeding using a Micro-Hematocrit capillary tube and EDTA (~5 mM) as anticoagulant. This first sample was obtained approximately 2–3 min after injection and it was considered to be the “time zero” sample. Additional samples were taken at 5 min, 15 min and 30 min. Immediately after they were obtained, blood samples were processed as described above to measure plasma [32P]LPA. Similar to the in vivo experiments, no band other than LPA was observed by radioautography.
Plasma LysoPLD activity was measured according to Umezu-Goto et al.  by mixing 10 μl of plasma with 90 μl of a buffer consisting of 100 mM Tris-HCl, pH 9.0, 500 mM NaCl, 5 mM MgCl2, 30 μM CoCl2, 0.05% Triton X-100 and 1.1 mM LPC (Avanti Polar 148870). After 4 h incubation at 37 ° C, choline in samples was measured colorimetrically at 555 nm by adding 100 μl of 50 mM Tris-HCl, pH 8.0, 5 mM MgCl2, 50 U/ml horseradish peroxidase, 18 U/ml choline oxidase, 5 mM 4-aminoantipyrine, and 3 mM N-ethyl-N-(2-hydroxy-3-sulfopropyl)-m-toluidine. Net production of choline was calculated by subtracting the amount of choline present in parallel samples that were kept frozen. Blood was obtained by retro-orbital bleeding using heparin as anticoagulant. Choline chloride (Fluka 26978) was used as standard.
The University of Virginia’s Animal Care and Use Committee approved all experiments with animals.
We first established the exact location of the exon trap element by sequencing genomic DNA from Ppap2atr/tr mice. As shown in Fig. 1A, insertion occurred in intron 1 as reported by BayGenomics. Mouse LPP1 is expressed as two different isoforms as a result of alternative splicing of exon 2 . However, in the present instance, both variants were expected to be trapped because insertion occurred upstream of either exon 2 (Fig. 1A). Based on the location of the trap element we designed primers to detect by PCR the normal and trapped versions of the gene and were able to identify wild type, heterozygous and homozygous animals or Ppap2a+/+, Ppap2a+/tr and Ppap2atr/tr animals (Fig. 1B).
Ppap2atr/tr and Ppap2a+/tr mice are phenotypically unremarkable, that is their anatomy, behavior, and fertility are not readily distinguished from those of wild type littermates. We examined the gross anatomy of all major organs of these animals and found no differences with wild type animals. Our examination included the male reproductive tract and the prostate in particular because LPP1 mRNA is particularly prominent in prostate and LPP1 is an androgen-induced gene .
We investigated next the levels of LPP1 mRNA expression in different organs of Ppap2atr/tr and Ppap2a+/+ mice by quantitative Real-Time RT-PCR. In these experiments, primers were anchored to exons 3 and 4 (forward primer) and 5 (reverse primer) and therefore results correspond to the combined expression of both LPP1 isoforms which, as depicted in Fig. 1A, are generated by alternative splicing of exon 2. As presented in Fig. 2, Ppap2atr/tr mice exhibited much reduced levels of LPP1 expression in relation to Ppap2a+/+ littermates in all organs studied except the brain. LPP1 mRNA expression in Ppap2atr/tr mice ranged from 1/300 (kidney) to 1/800 (muscle) the levels found in Ppap2a+/+ mice. The unaltered expression of LPP1 mRNA in the brain prompted us to investigate whether these animals lack the trapping element in this organ, but we found no evidence for a wild type allele in Ppap2atr/tr mice in brain genomic DNA (not shown). Further, we found no difference in the expression of LPP1 mRNA isoforms in comparing brains of Ppap2atr/tr and Ppap2a+/+ mice (not shown). Finally, we examined the expression of LPP3 in the same organs and found that, unlike LPP1, levels of LPP3 expression in Ppap2atr/tr and Ppap2a+/+ mice were similar (Fig. 2).
Tissue homogenates from Ppap2atr/tr mice exhibited reduced phosphatase activity in all organs studied except the brain, although that the extent of reduction varied among tissues (Fig. 3A). Using [32P]LPA as a substrate, we observed values ranging from 43% in heart (i.e. activity in Ppap2atr/tr mice was 57% that of Ppap2a+/+ mice) to 91% in the spleen. With [32P]PA as a substrate, values ranged from 31% in the kidney to 88% in the spleen. We found major differences in LPP activity between the brain and the other organs studied. As predicted from LPP1 mRNA quantification, LPA phosphohydrolase activity was not diminished in brain tissue from Ppap2atr/tr mice. We also found that the lipid phosphohydrolase activity from both Ppap2atr/tr and Ppap2a+/+ mice was higher with PA (vs. LPA) as a substrate in brain homogenates as compared to other tissues. In toto, these results lead us to conclude that Ppap2atr/tr mice are not entirely null for LPP1, rather these animals are LPP1 hypomorphs that exhibit severely reduced expression of LPP1 mRNA and reduced LPP activity in all organs studied except the brain.
As mentioned in the Introduction, LPPs such as LPP1 are NEM-insensitive, Mg2+-independent and act as exophosphatases, i.e. degrade lipids presented to the extracellular face of the cell membrane. A second group of lipid phosphohydrolases have opposite characteristics: they are sensitive to NEM, require Mg2+ and are located inside the cells. We carried out a series of experiments to further map to LPP1 the reduction of lipid phosphohydrolase activity observed in Ppap2atr/tr mice. To this end, we investigated first the effects of NEM alkylation and lack of Mg2+ on our assay of tissue phosphohydrolase activity (Fig. 3A), and, second, the exophosphatase activity of intact tissues (Fig. 4). As shown in Fig. 3A, only a small fraction of the total activity was NEM-sensitive. We compared the NEM-sensitive, Mg2+-dependent fractions from similar organs of Ppap2atr/tr and Ppap2a+/+ mice and found that in some cases Ppap2atr/tr mice show an increased amount of NEM-sensitive, Mg2+-dependent activity. This phenomenon is illustrated in Fig. 3B, where the Ppap2atr/tr/Ppap2a+/+ ratio of NEM-sensitive, Mg2+-dependent activities is shown for each organ. In the cases of the kidney, liver and muscle, ratios were much higher than unity: 7.6, 5.9 and 15 respectively. For the other organs studied, values ranged from 0.83 to 1.1.
The topology of LPP1 is such that the active site is at the cell surface, thus we measured exophosphatase activity (Fig. 4). We prepared tissue slices from different organs, incubated them with [32P]LPA and measured the release of [32P]H3PO4. Similar to our previous findings with tissue homogenates, Ppap2atr/tr mice organs, with the exception of the brain, showed a decreased level of exophosphatase activity and, once again, the spleen was the most severely affected organ. Values were 65, 74 and 94 % reduction for the kidney, liver and spleen (Figs. 2 and and3).3). We examined in more detail the exophosphatase activity of the spleen. Splenocytes isolated from Ppap2atr/tr mice exhibit, as expected, a greatly reduced exophosphatase activity (> 99%)
If LPP1 metabolizes LPA in vivo, the reduced levels of LPA phosphohydrolase activity in Ppap2atr/tr mice should result in elevated levels of plasma LPA. We tested this prediction by measuring plasma LPA levels in Ppap2atr/tr and Ppap2a+/+ mice by LC-MS-MS. We found that Ppap2atr/tr mice have higher levels of plasma LPA on average, but there was substantial variation in LPA levels among animals (Fig 5). To determine the source of this variation, we measured plasma LPA levels in a homogeneous population of pure C57BL/6j mice. We found that LPA levels in age- and gender-matched C57BL/6j mice were in agreement with previously reported values (see Discussion) and exhibited low variability (Fig. 5). This result suggests that the observed difference in plasma LPA levels between Ppap2atr/tr and Ppap2a+/+ mice is genuine and the wide range of LPA plasma values is a reflection of a diverse (genetically as well as age and gender) population. To test this idea, we backcrossed the mutant Ppap2a mice against C57BL/6j mice for four additional generations. After ascertaining that mutant and wild type the F1N5 mice retained the differences in LPP1 mRNA and activity delineated above, we measured plasma LPA levels. Although the F1N5 mice had much less variation in plasma LPA levels, the difference between Ppap2atr/tr mice and their wild type littermates remained (Fig. 5). Interestingly, the mole fractions of the LPA species measured was not different in Ppap2atr/tr and Ppap2a+/+ mice (not shown). The rank order of abundance was as follows: 18:2 > 20:4 > 22:6 > 18:0 ~ 16:0 > 18:1 > 20:3 ~ 22:5 > 16:1 ~ 22:4 14:0 ~ 20:5 ~ 20:2 22:3 ~ 22:2. This result suggests that LPP1 is not selective for any of these molecular species.
Next we examined the ability of Ppap2atr/tr and Ppap2a+/+ mice to metabolize LPA by measuring the rate of disappearance of [32P]LPA in blood both ex vivo and in vivo. Fig. 6A documents that the amount of LPA that can be recovered from whole blood incubated at 37°C rapidly diminished as a function of time. The rate of decay was slower for blood obtained from Ppap2atr/tr mice. Half-lives were approximately 5 and 20 min for Ppap2a+/+ and Ppap2atr/tr mice, respectively. Similar experiments measuring the decay in plasma samples revealed that [32P]LPA was stable in plasma up to 30 min (not shown). Fig. 6B shows a similar series of experiments except that in this case [32P]LPA was injected into the bloodstream of mice. In this case decay was faster and, similar to the ex vivo observations, disappearance of [32P]LPA was markedly slower in Ppap2atr/tr mice. Half lives in this case were approximately 3 and 12 min. Fig. 6 also documents that there were no differences in [32P]LPA decay when a less physiologic alternative — saline solution containing FA-free BSA — was used as a vehicle for [32P]LPA instead of plasma. Finally, we explored whether Ppap2atr/tr animals show any abnormality in the production of LPA, likely a compensatory reduction as a consequence of the elevated plasma LPA levels and the reduced rate of LPA degradation. We found no differences between Ppap2a+/+ and Ppap2atr/tr mice when we used a colorimetric method to detect choline produced by the breakdown of LPC into choline and LPA catalyzed by plasma lysophospholipase D. Values were (average ± S.D. of 5 animals): 7.53 ± 0.21 and 7.73 ± 0.23 × 10−13 moles choline/μl plasma/min.
In vitro studies have implicated LPP1 in the phosphohydrolysis of extracellular LPA, but the lack of specific inhibitors has made testing the physiologic relevance of LPP1 in this process difficult. With this paper we document that disruption of the LPP1 encoding gene, Ppap2a, results in mice with substantially reduced NEM-resistant LPA/PA phosphatase activity, moderately elevated plasma LPA levels and slowed clearance of exogenously administered LPA. Our data demonstrate that LPP1 contributes 43–81% of the total tissue LPA phosphohydrolase activity and 65–99% of the exophosphatase activity. We suppose that the residual LPA/PA phosphatase activity is due to other lipid phosphatases such as LPP3. Our results support the contention LPP1 is a physiologically relevant LPA exophosphatase.
Exophosphatase activity was virtually nil in Ppap2atr/tr splenocytes suggesting that LPP1 is the only functioning exo-LPP in these cells. This is a somewhat surprising observation in view of the presence of LPP3 mRNA in splenocytes. Nevertheless, the lack of LPA exophosphatase activity may be exploited to examine the role of LPP1 further.
The unaltered level of LPP1 mRNA expression and enzyme activity in the brains of Ppap2atr/tr mice was unexpected. The possibility that Ppap2atr/tr mice lack the trapping element in brain tissue, i.e. mosaicism, was ruled out by genotyping brain DNA. We reasoned that alternative splicing might be responsible for the failure of the exon trapping element in brain but we were unable to detect differential expression of known LPP1 isoforms . Thus we do not have a plausible explanation as to why the pre-mRNA splicing machinery does not recognize the exon trap splicing acceptor site in brain.
Our LPP1 hypomorphic mice have elevated plasma LPA levels. The range of reported mouse plasma LPA concentrations is 170–625 nM [34, 35, 36, 37]. The lowest concentration detected in our LPP1 mice was 368 nM, while the highest was 2.45 μM (Fig. 5). We considered whether the source this large variation in LPA levels was an artifact of our methods (plasma preparation, LC-MS-MS) or a reflection of the genetic diversity of our LPP1 colony (F1N1, C57BL/6 x sv129 strains). When one of our laboratories (KRL) generated plasma from age- and gender-matched pure-bred C57BL/6j mice and blinded another of our laboratories (VN) to the source of this plasma, our methods yielded values that were similar to previously reported mouse plasma LPA concentrations and, most importantly, exhibited much less variation: 348 ± 98 nM, i.e. 28%. Additionally, the intra-assay variability was found to be slight (Fig. 5). Thus we reasoned that the disparity in plasma LPA levels measured in our F1N1 Ppap2atr/tr mice was due largely to differences in genetic background and, perhaps, in gender and age. When our LPP1 mouse colony was re-derived at F1N5 after additional crossings onto the C57Bl/6j background, the variability in plasma LPA levels was substantially reduced in age-matched mice (Fig. 5). Nonetheless, our data indicate consistently that LPP1-deficient mice exhibit plasma LPA levels significantly higher than control littermates supporting the hypothesis that LPP1 acts as an LPA exophosphatase. Further, our results document that plasma LPA levels can exceed 2 μM in mice without apparent ill-effects.
We also investigated the ability of Ppap2atr/tr mice to catabolize LPA by comparing the rate of disappearance of tracer amounts of [32P]LPA from the blood of Ppap2atr/tr and Ppap2a+/+ animals. We found that [32P]LPA disappears rapidly from the blood of Ppap2a+/+ animals when incubated ex vivo at 37 °C and that this process is even faster in vivo. As Fig. 6 documents, Ppap2atr/tr mice exhibit, both ex vivo and in vivo, a still rapid but distinctly slower rate of LPA disappearance. We have measured only how much LPA remains after injection so it is not possible to ascribe the disappearance of exogenous LPA to any catabolic process. Nevertheless, the mutant mice demonstrate that LPP1 is in part responsible for this process.
In conclusion, we developed an LPP1 hypomorph mouse that enabled us to provide a definitive demonstration that LPP1 plays a role in the extracellular catabolism of LPA in intact animals under physiological circumstances. Further, our results suggest a role of LPP1 in controlling the kinetics of extracellular LPA turnover in vivo. These results provide evidence that LPP1 is an important determinant of LPA turnover and circulating LPA concentrations, which might alter LPA signaling patterns.
The authors thank Dr. Timothy Bender, Department of Microbiology, University of Virginia, for assisting us with the preparation of splenocytes, as well as Gina Wimer and Perry C. Kennedy for their help with mouse manipulations. This work was supported by National Institute of Health grants R01 GM052722 (to K.R.L.), R01 HL079396 (to V.N.), T32 GM007055 (to A.H.S.) and by the Canadian Institutes of Health Research MOP 81137 (to D.N.B.).