Lysophosphatidic acid: a novel growth and survival factor for renal proximal tubular cells

Jerrold S. Levine, Jason S. Koh, Veronica Triaca, and Wilfred Lieberthal

Renal Section, Department of Medicine, Boston Medical Center, Boston, Massachusetts 02118

    ABSTRACT
Top
Abstract
Introduction
Methods
Results
Discussion
References

Lysophosphatidic acid (LPA) is the smallest and structurally simplest of all glycerophospholipids. LPA is a normal constituent of serum and binds with high affinity to albumin while retaining its biological activity. The effects of LPA are pleiotropic and range from mitogenesis to stress fiber formation. In this report, we demonstrate two novel functions for LPA. LPA acts as a survival factor to inhibit apoptosis of primary cultures of mouse renal proximal tubular (MPT) cells. LPA also acts as a potent mitogen for MPT cells. The ability of LPA to act as both a survival factor and a mitogen is mediated by the lipid kinase phosphatidylinositol 3-kinase (PI3K), since these activities were completely blocked by wortmannin or LY-294002, two structurally dissimilar inhibitors of PI3K. The identification of LPA as a proliferative and anti-apoptotic factor suggests a potential role for this lipid mediator during the injury and/or recovery phases following tubular damage.

apoptosis; phosphatidylinositol 3-kinase; phospholipids

    INTRODUCTION
Top
Abstract
Introduction
Methods
Results
Discussion
References

LYSOPHOSPHATIDIC ACID (LPA; 1-acyl-sn-glycerol-3-phosphate), the smallest and structurally simplest of all glycerol-based phospholipids, has long been recognized as a precursor in phospholipid biosynthesis. Recently, LPA has also emerged as an important intercellular signaling molecule with a specific cell surface receptor of its own (9, 14). Although LPA is not present in plasma or freshly isolated blood, it is produced and released by platelets during blood clotting and is therefore a normal constituent of serum (2, 28). The normal concentration of LPA in serum is in the range of 5-20 µM (2). LPA binds with high affinity to serum albumin while retaining its biological activity (27, 28). Like most growth factors or cytokines, LPA has a pleiotropic range of effects (9, 14). These include stimulation or inhibition of cell proliferation (depending upon the cell type), focal adhesion formation, stress fiber formation, smooth muscle cell contraction, and stimulation of tumor cell invasion (9, 14, 20).

In this report, we show that LPA can act as a "survival factor" to inhibit apoptosis of primary cultures of mouse renal proximal tubular (MPT) cells induced by removal of growth factors. Apoptosis refers to an energy-requiring, gene-directed cellular process, which, if activated, results in cell "suicide" (11, 19). The morphological and biochemical characteristics of cells dying by apoptosis differ from those of cells dying by necrosis. Necrotic cells swell and lyse, with cellular material spilling into the extracellular space and inducing an inflammatory response. In apoptotic cells, on the other hand, the nucleus and cytoplasm shrink and fragment into plasma membrane-bound vesicles called apoptotic bodies. Recognition and ingestion by phagocytes of intact membrane-bound apoptotic bodies or even entire apoptotic cells serves to protect tissues from an otherwise harmful exposure to the contents of dying cells (11, 21).

The viability of most if not all cells depends upon so-called "survival factors," which induce intracellular signals that prevent the cell from committing suicide by apoptosis (19). Competition for survival signals provides a simple means to ensure that a balance exists between cell division and cell death. If a given level of a survival factor supports a certain number of cells, then any increase in the number of cells would tend to increase competition and result in increased cell death, thereby returning cell number to its original value. Correspondingly, a decrease in cell number would permit division of cells back to the original number.

We examined the effect of LPA on the survival of MPT cells based on the following observations. We have recently shown that MPT cells cultured in the absence of growth factors undergo apoptosis (unpublished observations). Apoptosis may be inhibited by renal growth factors such as epidermal growth factor (EGF) and insulin-like growth factor-I (IGF-I). However, neither of these factors is as effective in inhibiting apoptosis as calf serum. Since albumin-bound LPA may be responsible for much of the non-cytokine-mediated activity of whole serum (14), we tested the role of LPA on the survival and proliferation of MPT cells. We show, for the first time in any cell type, that LPA alone can act as a "survival factor" to inhibit apoptosis and that LPA is as effective as serum in prolonging the survival of MPT cell monolayers. We also provide novel evidence that LPA is a potent growth factor for MPT cells.

In addition, on the basis of the results of a recent study in which the survival activity of EGF, insulin, nerve growth factor (NGF), and fetal bovine serum was inhibited by wortmannin and LY-294002 (32), two structurally dissimilar inhibitors of phosphatidylinositol 3-kinase (PI3K) (29, 30), we examined the role of PI3K in mediating the effects of LPA. Our data indicate that stimulation of proliferation and inhibition of apoptosis by LPA may be mediated through the enzyme PI3K.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

Reagents

Bisbenzamide (Hoechst dye or H-33342) was obtained from Calbiochem (San Diego, CA). Sodium oleoyl-L-alpha -lysophosphatidic acid (LPA) was obtained from Avanti Polar Lipids (Alabaster, AL). Trypsin-EDTA was obtained from GIBCO-BRL (Grand Island, NY). LY-294002 [2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one] was obtained from Biomol Research Laboratories (Plymouth Meeting, PA). All other culture supplies and reagents, including trypan blue, wortmannin, and delipidated fraction V bovine serum albumin (BSA), were obtained from Sigma.

Primary Culture of MPT Cells

Cells were cultured from collagenase-digested fragments of proximal tubules obtained from the cortices of kidneys of C57Bl/6 mice by a modification of previously described methods (12). Cortical tubules were plated in serum-free, defined culture medium [1:1 mixture of Dulbecco's modified Eagle's medium (DMEM) and Ham's F-12, containing 2 mM glutamine, 15 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 5 µg/ml transferrin, 5 µg/ml insulin, 50 nM hydrocortisone, 500 U/ml penicillin, and 50 µg/ml streptomycin], hereafter denoted as "full medium." Growth factor-free medium is defined as full medium minus insulin and hydrocortisone. MPT cells grew to confluence from tubules over 4-5 days and were studied within 2 wk of achieving confluence. Cell monolayers were previously shown to be of proximal tubular origin by a combination of morphological, biochemical, and transport characteristics (12).

Preparation of LPA

Fatty acid-free BSA was dissolved at 10 mg/ml in calcium- and magnesium-free phosphate-buffered saline (PBS), through which oxygen-free nitrogen had been bubbled for 20 min. LPA was added to achieve a final concentration of 0.5 mg/ml (1.1 mM) and stored under argon at -70°C.

Induction of Apoptosis

Upon reaching confluence, MPT cell monolayers were washed with PBS and incubated in full or growth factor-free medium (experimental wells). Calf serum (10%) or varying concentrations of LPA in the presence or absence of wortmannin or LY-294002 were added to experimental wells. The effects of LPA on MPT cell survival were determined using growth factor-free medium, because we have previously shown that withdrawal of insulin and hydrocortisone induces apoptosis of MPT cells (unpublished observations).

Cell Viability Assay

Cell viability was assessed 10 days after addition of calf serum or LPA. Cell viability was quantitated by counting the number of viable cells, defined as cells that both remained adherent to the culture dish and excluded trypan blue. Nonadherent cells were removed by two washes with ice-cold PBS. Adherent cells were harvested by incubation with 0.05% trypsin/0.53 M tetrasodium EDTA for 10 min at 37°C. Trypsin was neutralized by addition of DMEM containing 10% calf serum. Cells were centrifuged for 5 min at 100 g and resuspended in DMEM. Trypan blue (0.04 g/dl) was added for 10 min, and the number of viable cells excluding trypan blue was counted in a hemocytometer. Cell viability was expressed as the percentage of viable cells in experimental wells compared with that in freshly confluent monolayers.

Immunofluorescent Staining of Cells

The morphology of nuclear chromatin was assessed by staining with H-33342, a supravital DNA dye with an excitation wavelength of 348 nm and emission wavelength of 479 nm (12). H-33342 enters live cells and therefore stains the nuclei of viable cells, as well as cells that have died by apoptosis or necrosis. Apoptotic cells may be distinguished from viable and necrotic cells on the basis of nuclear condensation and fragmentation as well as increased fluorescent intensity of nuclei stained with H-33342.

Adherent MPT cells (harvested as above) and cells that had detached spontaneously from the monolayer were washed once in PBS before staining with H-33342 (1.0 µg/ml) for 10 min at 37°C. Wet preparations of adherent and detached cells were made on glass slides, and each field of cells was photographed twice, under phase-contrast as well as fluorescence microscopy, for visualizing cell morphology and H-33342 nuclear staining in the same cells.

Flow Cytometry

MPT cells were stained with H-33342 using the same technique described for immunofluorescent microscopy and then placed immediately on ice. Flow cytometry was performed on an Epics ESP Flow Cytometer (Coulter Electronics, Hialeah, FLA) with an ultraviolet (UV)-enhanced argon laser. Hoechst fluorescence was accomplished by excitation with <5 mW of UV laser light (351-364 nm multiline) and detected with a 525-nm bandpass optical filter. Data were analyzed by Epic Elite software (Coulter Electronics). A constant number of events was analyzed for each sample (10,000 events/sample).

We used two methods of flow cytometric analysis to distinguish normal cells from apoptotic cells, as follows.

Light scatter measurements. Normal cells are characterized by relatively high forward angle light scatter (a measure of cell size or volume) and relatively low side angle light scatter (a measure of cell granularity). Apoptotic cells, in contrast, are smaller and more granular than normal cells and are characterized by relatively low forward scatter and high side scatter (25).

Intensity of H-33342 fluorescence. A number of investigators have used the difference in intensity of Hoechst fluorescence between normal (faint nuclear fluorescence) and apoptotic cells (bright nuclear fluorescence) to distinguish these populations on flow cytometry (17, 18, 24).

We have analyzed our cells using both methods, i.e., by comparing forward scatter (x-axis) with side scatter (y-axis) as well as by comparing forward scatter (x-axis) versus intensity of H-33342 fluorescence (y-axis). In both cases, cellular debris was gated out on the basis of size (forward scatter) and therefore not included among the 10,000 events/sample.

Thymidine Incorporation

MPT cells were plated at 50,000 cells/well and incubated overnight in growth factor-free medium. The medium was then replaced with fresh growth factor-free medium alone or growth factor-free medium supplemented with 10% calf serum or various concentrations of LPA. After an additional 24 h, 2 µCi of [3H]thymidine (2 Ci/mmol; New England Nuclear, Boston, MA) were added to all wells. After 2 h, the cells were washed three times with DMEM, then incubated with 2.0 ml of ice-cold 5% trichloroacetic acid (TCA) for 1 h at 4°C. The TCA was removed, and MPT cells were washed once with fresh TCA. A quantity of 2.0 ml ice-cold ethanol containing 200 µM potassium acetate was added to each well for 5 min, following which the cells were incubated twice in 2.0 ml of 3:1 mixture of ethanol:ether, for 15 min per incubation. After allowing the monolayers to air dry, cells were solubilized in 1.0 ml of 0.1 N sodium hydroxide. [3H]thymidine counts per minute (cpm) were measured by adding samples to scintillation fluid and counting cpm using a Tri-Carb Liquid Scintillation Analyzer beta counter (model 1600TR; Packard Instrument, Meriden, CT).

Statistics

In each experiment, duplicate wells were examined, and the results of duplicate wells were averaged. All data are expressed as means ± SE. Comparisons between the multiple different graphs were made using a Student's t-test. When more than one comparison was necessary, the Bonferroni correction was used.

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

Prolongation of MPT Monolayer Confluence by LPA

Freshly confluent MPT monolayers cultured in full medium demonstrate the typical cobblestone appearance of epithelial cells and maintain confluence over 2 wk (Fig. 1A). In contrast, monolayers maintained in growth factor-free medium (full medium without insulin and hydrocortisone) fail to maintain confluence and display typical features of apoptosis. Individual cells become small and rounded before detaching from the monolayer. The detachment of apoptotic cells occurs over many days, resulting in a gradual but progressive loss of the cell monolayer (Fig. 1B).


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Fig. 1.   Phase-contrast microscopy of mouse renal proximal tubular (MPT) cells cultured in growth factor-free medium alone or growth factor-free medium supplemented with calf serum, lysophosphatidic acid (LPA), or LPA plus LY-294002. A: freshly confluent cells show the typical "cobblestone" appearance of normal epithelial cells. B: most of the MPT cells cultured in growth factor-free medium for 10 days have detached from the monolayer. Of the remaining cells, many are smaller and more rounded, both characteristic features of apoptotic cells. C and D: in contrast, MPT cell monolayers cultured for 10 days in growth factor-free medium supplemented with either 10% calf serum (C) or 12 µM LPA (D) remain fully viable and confluent. E: addition of the phosphatidylinositol 3-kinase (PI3K) inhibitor LY-294002 (20 µM) blocked the ability of LPA (12 µM) to maintain confluence of MPT monolayers and resulted in cells having the morphological appearance of apoptosis.

Apoptosis may be inhibited by the addition of calf serum to growth factor-free medium (Fig. 1C). In support of the idea that albumin-bound LPA accounts for much of the non-cytokine-mediated biological activity of whole serum, the addition of LPA was as effective as serum in maintaining confluence of MPT monolayers (Fig. 1D). The concentration of LPA used in these experiments (12 µM) is within the range of concentrations normally found in serum (5-20 µM).

We next compared the protective effect of LPA to that of calf serum by counting the number of viable MPT cells remaining in the monolayer after 10 days of culture (Fig. 2). In comparison with freshly cultured MPT cells, only 27 ± 4% of MPT cells were still viable after 8-10 days of incubation in growth factor-free medium. The addition of 10% calf serum or 12 µM LPA to growth factor-free medium increased viability to 86 ± 4% and 77 ± 4%, respectively (n = 6, P < 0.001 in comparison to growth factor-free medium for each condition). The survival effect of LPA was not different from that of calf serum (P > 0.1).


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Fig. 2.   Effect of LPA on cell viability. Mouse proximal tubular (MPT) cell monolayers were cultured for 10 days in growth factor-free medium alone or supplemented with 10% calf serum or 12 µM LPA. The proportion of viable cells (adherent and excluding trypan blue) is expressed as a percent of the absolute number of cells in freshly confluent control monolayers (274 ± 12 × 103 cells/well; n = 6). * P < 0.0001 compared with growth factor-free medium. The survival effects of LPA and 10% calf serum are comparable.

A clear dose-response relationship existed between LPA concentration and MPT cell survival (Fig. 3). Cell survival, after 7-10 days of culture in growth factor-free medium containing LPA (12, 6, or 1 µM; n = 6) was 77 ± 4%, 55 ± 4%, and 35 ± 8%, respectively, compared with only 27 ± 6% for MPT cells cultured in the absence of LPA (P < 0.005, for 12 and 6 µM LPA; P = 0.25 for 1 µM LPA).


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Fig. 3.   Dose response for effect of LPA on cell viability. MPT cell monolayers were cultured for 10 days in growth factor-free medium alone or supplemented with 12, 6, or 1 µM LPA. The proportion of viable cells (adherent and excluding trypan blue) is expressed as a percent of the absolute number of cells in freshly confluent monolayers (305 ± 12 × 103 cells/well; n = 6). * P < 0.001 compared with growth factor-free medium.

Inhibition of Apoptosis by LPA

The increased numbers of viable cells seen with addition of LPA to growth factor-free medium may potentially be the result of two factors, namely, increased production of MPT cells from stimulation of proliferation and/or decreased loss of MPT cells from inhibition of apoptosis. LPA has been shown to stimulate proliferation in a variety of cells (9, 14), but to our knowledge, no studies have yet addressed a role for LPA as a survival factor. As will be shown, both increased proliferation and decreased apoptotic cell death seem to contribute to the protective effect of LPA on MPT cells in culture.

We first determined the effect of LPA on apoptotic loss of MPT cells. We have previously shown that apoptotic MPT cells can be readily distinguished from viable and necrotic cells by morphological criteria (12). In comparison with viable cells, apoptotic cells are smaller and show nuclear condensation and fragmentation upon staining with the cell-permeant supravital DNA dye H-33342. Adherent MPT cells obtained from freshly confluent monolayers (Fig. 4, A and B) demonstrated the faint chromatin staining pattern of normal nuclei. In marked contrast, the majority of nuclei of MPT cells cultured in growth factor-free medium (Fig. 4, C and D) were abnormal. Nuclei from cells at a relatively early stage of apoptosis showed bright H-33342 staining indicative of nuclear condensation but maintained a relatively normal overall morphology. Nuclei from cells at a later stage of apoptosis also stained brightly but had undergone nuclear fragmentation as well. The addition of either calf serum (Fig. 4, E and F) or LPA (Fig. 4, G and H) led to a dramatic decrease in the number of MPT cells demonstrating apoptotic morphology.


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Fig. 4.   Phase-contrast and fluorescence microscopy of H-33342-stained MPT cells from freshly confluent MPT cells (A and B) and MPT cells after culture in growth factor-free medium alone (C and D) or supplemented with calf serum (E and F) or LPA (G and H). After pooling adherent MPT cells with those that had detached from the monolayer, cells were stained with H-33342, mounted onto glass slides as wet preparations, and photographed under phase-contrast (A, C, E, G) and fluorescence (B, D, F, and H) microscopy for visualizing cell morphology and H-33342 nuclear staining in the same cells. H-33342 staining of cells from freshly confluent monolayers (B) shows faint fluorescence of nuclei with a delicate chromatin pattern. This is a normal nuclear morphology. These cells appear normal under phase (A). In contrast, H-33342 staining of cells cultured in growth factor-free medium for 10 days (D) shows nuclei with varying morphology. The majority of nuclei are abnormal and appear as brightly staining, homogeneous masses of varying size. These morphological features are characteristic of apoptosis and result from chromatin condensation and nuclear fragmentation. Cells displaying apoptotic nuclear morphology may also be identified as apoptotic by phase-contrast microscopy on the basis of decreased cell size (C). Supplementation of growth-factor-free medium with either 10% calf serum (F) or 12 mM LPA (H) prevents apoptosis and results in a nuclear H-33342 staining pattern comparable to that seen with freshly confluent cells.

We used flow cytometry to confirm the inhibitory effect of LPA on apoptosis (Fig. 7). MPT cells were separated into viable and apoptotic populations on the basis of size (forward scatter), granularity (side scatter), and intensity of H-33342 nuclear staining. Viable cells were defined in left of Fig. 7 as those having normal size and low granularity and in right of Fig. 7 as those having normal size and faint H-33342 nuclear staining. Apoptotic cells were defined in left of Fig. 7 as those having decreased size and increased granularity and in right of Fig. 7 as those having decreased size and bright H-33342 staining. Electron microscopic examination of sorted populations based upon size and intensity of H-33342 nuclear staining validated these definitions (Figs. 5 and 6). Cells of normal size and faint H-33342 staining were uniformly viable (Figs. 5A and 6A), whereas cells of decreased size and bright H-33342 staining showed predominantly apoptotic morphology (Figs. 5B and 6, B and C). Comparable micrographs were obtained when cells were sorted on the basis of size and granularity.


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Fig. 5.   Electron microscopy (low-power view) of MPT cells sorted by flow cytometry on the basis of forward light scatter (size) and intensity of H-33342 nuclear staining. On low-power view, cells characterized on flow cytometry by normal size and faint H-33342 nuclear staining all demonstrate normal, viable morphology (A, ×4,200 magnification). In contrast, cells characterized on flow cytometry by decreased size and bright H-33342 nuclear staining are predominantly apoptotic (B, ×4,200 magnification).


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Fig. 6.   Electron microscopy (high-power views) of MPT cells sorted by flow cytometry on the basis of forward light scatter (size) and intensity of H-33342 nuclear staining. The MPT cell shown in A (×6,900 magnification) is a normal, viable MPT cell sorted with the population of cells characterized by normal size and faint H-33342 staining on flow cytometry. Representative cells from the population sorted by flow cytometry on the basis of decreased size and bright H-33342 nuclear staining (B, ×6,900 magnification; and C, ×11,800 magnification) are both apoptotic. The apoptotic cell shown in B is slightly smaller than the normal cell (A), whereas the apoptotic cell (C) is about one-half the size of the normal cell (magnification for C is almost 2-fold greater than for A). In addition to being smaller than normal, both apoptotic cells (B and C) demonstrate chromatin condensation and nuclear fragmentation (both are indicated by arrowheads) that are classic morphological features of apoptosis. Other features of apoptosis demonstrated by B and C include morphologically intact plasma membranes that have lost their microvilli and preservation of the structure of mitochondria and other subcellular organelles.

Using these definitions, we found the percentage of viable cells was only 16% when cells were cultured in growth factor-free medium (Fig. 7a). The addition of either calf serum (Fig. 7b) or LPA (Fig. 7c) markedly increased the percentage of viable cells to 60% and 59%, respectively. Combined with Fig. 4, these data demonstrate that the increased survival of MPT monolayers in the presence of LPA is at least in part attributable to inhibition of apoptosis.


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Fig. 7.   Flow cytometric analysis of MPT cells cultured in growth factor-free medium alone (a) and supplemented with serum (b), LPA (c), or LPA plus inhibitors of PI3K (d and e). After pooling adherent MPT cells with those that had detached from the monolayer, cells were stained with H-33342 and sorted by flow cytometry on the basis of size (forward scatter), granularity (side scatter), and log fluorescent intensity of H-33342 staining. Viable cells were defined on left as those having normal size and granularity (region R1) and on right as those having normal size and faint H-33342 staining (bottom right quadrant). Apoptotic cells were defined on left as those having decreased size and increased granularity and on right as those having decreased size and bright H-33342 staining (top left quadrant). The percentage of viable cells after 10 days in growth factor-free medium is ~20% (a). Supplementation with either 10% calf serum (b) or 12 µM LPA (c) improves viability to ~60%. Addition of the PI3K inhibitors, wortmannin (10 nM) (d) or LY-294002 (20 µM) (e), completely blocked the ability of LPA (12 µM) to maintain viability of MPT cells. This flow cytometric analysis is representative of 5 experiments.

Stimulation of Proliferation by LPA

We assessed directly the contribution of LPA-induced proliferation of MPT cells to maintenance of confluence by measuring [3H]thymidine incorporation as an index of DNA synthesis (Fig. 8). MPT cells were plated at a density of 50,000 cells/well and were cultured for 24 h in growth factor-free medium alone or in growth factor-free medium to which 10% calf serum or LPA had been added. [3H]thymidine was added for the final 2 h. [3H]thymidine incorporation by confluent MPT monolayers incubated in growth factor-free medium was 563 ± 56 disintegrations per minute (dpm) per well. In comparison, calf serum increased [3H]thymidine incorporation to 3,002 ± 544 dpm/well (n = 5, P < 0.02). LPA increased [3H]thymidine incorporation in a dose-dependent fashion (Fig. 8). Stimulation was maximal at a concentration of 96 µM (1,346 ± 164 dpm/well, P < 0.01 vs. growth factor-free medium) and was increased at a dose as low as 3 µM (n = 5, P < 0.05 vs. growth factor-free medium).


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Fig. 8.   Dose-response curve for effect of LPA on thymidine uptake. Confluent MPT monolayers were cultured for 24 h in growth factor-free medium alone or supplemented with 10% calf serum and LPA (1 to 96 µM). [3H]thymidine was added for the final 2 h. Data are presented as the percent increment above baseline [3H]thymidine uptake seen for monolayers cultured in growth factor-free medium (n = 5). * P < 0.002 compared with growth factor-free medium. dagger  P < 0.05 compared with 3 µM LPA. ¶ P < 0.05 compared with 12 µM LPA.

Thus, like calf serum, LPA inhibits apoptosis and stimulates proliferation of MPT cells in culture. Therefore, maintenance of MPT cell monolayers by LPA is likely the result of both increased proliferation and decreased apoptotic cell death. It should be noted that both effects of LPA were seen at concentrations as low as 3 µM. As the normal serum concentration of LPA is ~5-20 µM, these data suggest that a significant portion of the effect of 10% serum may be attributable to albumin-bound LPA.

Dependence of LPA-Induced Proliferation and Inhibition of Apoptosis on PI3K

Finally, we determined the role of PI3K in mediating the effects of LPA using wortmannin and LY-294002, two structurally dissimilar inhibitors of PI3K (29, 30). Addition of wortmannin (10 nM) blocked the ability of LPA (12 µM) to maintain confluence of MPT monolayers (Fig. 1E). We confirmed this observation by counting the number of viable MPT cells remaining in the monolayer after 8-10 days of culture (Fig. 9, A and B). Both wortmannin (10 nM) (n = 5) and LY-294002 (20 µM) (n = 6) blocked the ability of LPA (12 µM) to maintain MPT cell survival (Fig. 9, A and B). Since we have shown that increased cell survival is attributable both to increased proliferation and to inhibition of apoptosis, we examined the effects of wortmannin and LY-294002 on each of these processes separately. Inhibition of PI3K by either wortmannin or LY-294002 completely blocked the ability of LPA both to inhibit apoptosis (Fig. 7, d and e) and to stimulate proliferation (Fig. 10, A and B). Thus both LPA-induced proliferation and inhibition of apoptosis appear to be mediated by a PI3K-dependent signaling pathway.


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Fig. 9.   Effect of the PI3K inhibitors wortmannin and LY-294002 on LPA-mediated cell survival. MPT cell monolayers were cultured for 10 days in growth factor-free medium alone or supplemented with 10% calf serum, 12 µM LPA, PI3K inhibitor alone, or 12 µM LPA plus the PI3K inhibitor. Data for wortmannin (W, 10 nM) are shown in A (n = 5), and data for LY-294002 (LY, 20 µM) are shown in B (n = 6). The proportion of viable cells (adherent and excluding trypan blue) is expressed as a percent of the absolute number of cells in freshly confluent control monolayers (A, 301 ± 17 × 103 cells/well; B, 283 ± 2 cells/well). * P < 0.01 compared with growth factor-free medium. dagger P < 0.01 compared with LPA.


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Fig. 10.   Effect of the PI3K inhibitors wortmannin and LY-294002 on LPA-mediated thymidine uptake. MPT cell monolayers were cultured for 10 days in growth factor-free medium alone or supplemented with 10% calf serum, 12 µM LPA, PI3K inhibitor alone, or 12 µM LPA plus the PI3K inhibitor. [3H]thymidine was added for the final 2 h. Data are presented as the absolute [3H]thymidine uptake (dpm/well). Data for wortmannin (W, 20 nM) are shown in A (n = 8), and data for LY-294002 (LY, 20 µM) are shown in B (n = 3). * P < 0.01 compared with growth factor-free medium. dagger P < 0.05 compared with LPA.

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

In this report, we demonstrate that the glycerol-based phospholipid LPA can act as a survival factor to inhibit apoptosis of MPT cells. This represents the first report showing in any cell type that LPA alone, in the absence of any other growth factor, is capable of inhibiting apoptosis. Thus inhibition of apoptosis should be added to the growing list of biological activities attributable to LPA. These activities include such diverse effects as focal adhesion assembly, stress fiber formation, platelet aggregation, smooth muscle cell contraction, and stimulation of tumor cell invasion (9, 14, 20).

We also demonstrate that LPA stimulates the proliferation of MPT cells. Although the proliferative effectsof LPA are well documented in other cell types (9, 14), including vascular smooth muscle cells, fibroblasts, Jurkat T cells, and keratinocytes, ours is the first report to demonstrate that LPA is a trophic factor for renal tubular cells. Although most biological effects of LPA are mediated by nanomolar concentrations of the phospholipid, LPA-stimulated proliferation has been found to require micromolar concentrations (9). This was true in our study as well, in which stimulation of proliferation, as well as inhibition of apoptosis, required micromolar concentrations of LPA. It is unclear why these two biological actions should be seen only with higher concentrations of LPA, but it has been suggested that the reason may relate to loss of activity from degradation or oxidation given the prolonged incubation times needed to study proliferation and apoptosis (9).

On the basis of inhibition studies with wortmannin and LY-294002, we suggest that both stimulation of proliferation and inhibition of apoptosis by LPA are mediated via PI3K activation. We base this conclusion on the following lines of evidence. At the nanomolar concentrations used in this study, wortmannin is an irreversible and noncompetitive inhibitor of PI3K (29). Inhibition appears to be highly specific for PI3K, as wortmannin does not inhibit the function of a wide variety of protein and lipid kinases, including protein kinases A, C, and G and the structurally related phosphatidylinositol 4-kinase (15, 29, 31). Nonspecific effects of wortmannin, such as inhibition of myosin light chain kinase, occur only at 100-fold higher concentrations than those used in this study (26, 29, 31). Moreover, we obtained identical results using LY-294002, a recently developed alternative inhibitor of PI3K. As for wortmannin, the effective concentration of LY-294002 (20 µM) is consistent with its activity as a PI3K inhibitor (30). Since wortmannin and LY-294002 are structurally dissimilar (29, 30), it is unlikely that their effects in our study can be attributed to any nonspecific interactions. It remains possible, however, that wortmannin and LY-294002 also inhibit an as yet uncharacterized member of the PI3K family and that inhibition of this family member, rather than inhibition of PI3K itself, is responsible for the effects of these inhibitors (cf. below). Moreover, our data do not address the issue of whether signaling events downstream of PI3K affect cell proliferation and apoptosis by the same or separate pathways.

Although the mechanism by which receptor tyrosine kinases (such as those for EGF or IGF-I) activate PI3K is understood, the mechanism for G protein-coupled receptors, such as that for LPA, remains unclear (9, 14, 29). PI3K is a heterodimer consisting of a 110-kDa catalytic subunit (p110alpha and p110beta ) and an 85-kDa regulatory subunit (p85alpha and p85beta ). Activation of PI3K by receptor tyrosine kinases occurs following recruitment of PI3K to membrane-bound signaling complexes. Recruitment is mediated by the binding of src-homology 2 domains within the p85 regulatory subunit to specific phosphotyrosine residues. In contrast, induction of PI3K activity by LPA may involve the recently characterized p100gamma isotype of the catalytic subunit (23). The p100gamma isotype does not interact with p85, and, at least in vitro, is activated by directly associating with G proteins.

PI3K has been implicated in a wide range of cellular processes such as mitogenesis, differentiation, membrane ruffling, and vesicular trafficking (10). A role for PI3K in regulation of apoptosis has emerged from a recent study in which inhibition of PI3K activity by wortmannin or LY-294002 blocked the ability of NGF, EGF, insulin, and serum to prevent apoptosis of PC12 cells in culture (32). An attractive downstream target of PI3K that may be responsible for stimulation of proliferation and inhibition of apoptosis is the 70-kDa S6 kinase (pp70S6k) (1). Activation of pp70S6k is an established part of the mitogenic response and leads to the phosphorylation of the S6 polypeptide of the 40S ribosomal subunit. Full activation of pp70S6k depends not only on PI3K but also on phospholipase Cgamma and the rapamycin-inhibitable enzyme, FK506 binding protein-rapamycin-associated protein. In this regard, it is noteworthy that administration of rapamycin enhanced apoptosis in several different cell lines (22). These data are consistent with the view that pp70S6k is a critical target of PI3K in inhibiting apoptosis.

An alternative explanation for the effects of wortmannin and LY-294002 in this study relates to the fact that the double-stranded DNA-dependent protein kinase (DNA-PK) has a PI3K domain and is inhibitable by wortmannin, albeit at concentrations about 100-fold greater than those used in this study (8). Given the role of DNA-PK in repair of double-stranded DNA breaks, it is tempting to speculate whether there exist other members of this family that are inhibitable by lower concentrations of wortmannin and that participate in regulation of apoptosis via recognition and/or repair of damaged DNA.

Although our inhibition studies focus attention on PI3K, stimulation of proliferation and inhibition of apoptosis by LPA may also be viewed in a larger context. Virtually all cell types depend upon so-called "survival factors," which deliver a signal that prevents the cell from committing suicide by apoptosis. Such survival factors frequently derive from paracrine or autocrine secreted growth factors, but in the case of adhesive cells such as MPT cells, survival factors also include adhesive signals from the extracellular matrix or intercellular contact (3, 4, 7). In this regard, activation by LPA of the Ras-related cytoplasmic GTP-binding protein Rho assumes importance (5, 14, 20). LPA-mediated activation of Rho potently induces the assembly of actin stress fibers and the formation of focal adhesion complexes, which are involved in cell-matrix and some cell-cell interactions (5, 20).

Combined with the data presented in this report, several facts induce us to hypothesize that an intimate connection exists between survival factors, cytoskeletal rearrangement, and cell cycle progression. First, many potent survival factors, such as platelet-derived growth factor (PDGF), EGF, and IGF-I, have all been shown to induce profound changes in the actin cytoskeleton, mediated via various members of the Rho family of GTP-binding proteins (5, 20). Second, as shown in a recent study, activation of Rho may itself play a significant role in progression through the G1 phase of the cell cycle (16). Microinjection of Rho into Swiss 3T3 fibroblasts stimulated cell progression through G1, whereas microinjection of a dominant negative form of Rho blocked traversal of G1 by serum-stimulated cells. Third, not only can signals derived from the extracellular matrix act as survival signals in some cells (3), but adhesion is necessary in many of these same anchorage-dependent cells for progression through G1 (4). Indeed, it is noteworthy that adhesion-dependent cell cycle progression and inhibition of apoptosis show similar signaling requirements (4, 13). Finally, in a fibroblast model of c-Myc-induced apoptosis, only those cytokines known to be important for traversal of G1 checkpoints (i.e., PDGF and IGF-I) were able to inhibit apoptosis, even when these cells were growth arrested in the S phase of the cell cycle (6).

In summary, we have provided novel data showing that the glycerol-based phospholipid LPA not only stimulates the proliferation of MPT cells in culture but can also itself act as a survival factor to inhibit apoptosis. Both effects of LPA appear to be mediated via the intracellular lipid kinase PI3K. Given recent interest in the contribution of apoptosis to the pathogenesis of renal diseases resulting from injury to renal tubular epithelial cells (11) and the protective effect of growth factors following experimental acute renal failure, the identification of LPA as a proliferative and anti-apoptotic factor suggests a potential role for this lipid mediator during the injury or recovery phases of tubular damage. Moreover, although activated platelets are the best characterized source of LPA, injured cells may also release LPA (9, 14) and thus provide an autocrine or paracrine survival and proliferative factor following renal injury.

    ACKNOWLEDGEMENTS

We are grateful to John Daley for technical assistance in flow cytometric analysis.

    FOOTNOTES

This work was supported by National Institutes of Health Grants AR/AI-42732 (to J. S. Levine), DK-375105 (W. Lieberthal), and HL-53031 (W. Lieberthal); by American Cancer Society Grant IN97-N (to J. S. Levine); and by a Young Investigator Award from the National Kidney Foundation (to J. S. Levine).

Address for reprint requests: J. S. Levine, Renal Section, E428, Boston Medical Center, 88 East Newton St., Boston, MA 02118 (E-mail: jlevine{at}bu.edu).

Received 2 January 1997; accepted in final form 26 June 1997.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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AJP Renal Physiol 273(4):F575-F585
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