Swelling-Induced Arachidonic Acid Release via the 85-kDa cPLA2 in Human Neuroblastoma Cells

Srisaila Basavappa1, 3, Stine F. Pedersen2, Nanna K. Jørgensen2, J. Clive Ellory1, and Else K. Hoffmann2

1 University Laboratory of Physiology, University of Oxford, Oxford OX1 3PT, United Kingdom; 2 August Krogh Institute, University of Copenhagen, DK-2100 Copenhagen Ø, Denmark; and 3 Children's Hospital and Harvard Medical School, Boston, Massachusetts 02115

    ABSTRACT
Abstract
Introduction
Methods
Results
Discussion
References

Basavappa, Srisaila, Stine F. Pedersen, Nanna K. Jørgensen, J. Clive Ellory, and Else K. Hoffmann. Swelling-induced arachidonic acid release via the 85-kDa cPLA2 in human neuroblastoma cells. J. Neurophysiol. 79: 1441-1449, 1998. Arachidonic acid or its metabolites have been implicated in the regulatory volume decrease (RVD) response after hypotonic cell swelling in some mammalian cells. The present study investigated the role of arachidonic acid (AA) during RVD in the human neuroblastoma cell line CHP-100. During the first nine minutes of hypo-osmotic exposure the rate of 3H-arachidonic acid (3H-AA) release increased to 250 ± 19% (mean ± SE, n = 22) as compared with cells under iso-osmotic conditions. This release was significantly inhibited after preincubation with AACOCF3, an inhibitor of the 85-kDa cytosolic phospholipase A2 (cPLA2). This indicates that a PLA2, most likely the 85-kDa cPLA2 is activated during cell swelling. In contrast, preincubation with U73122, an inhibitor of phospholipase C, did not affect the swelling-induced release of 3H-AA. Swelling-activated efflux of 36Cl and 3H-taurine were inhibited after preincubation with AACOCF3. Thus the swelling-induced activation of cPLA2 may be essential for stimulation of both 36Cl and 3H-taurine efflux during RVD. As the above observation could result from a direct effect of AA or its metabolite leukotriene D4 (LTD4), the effects of these agents were investigated on swelling-induced 36Cl and 3H-taurine effluxes. In the presence of high concentrations of extracellular AA, the swelling-induced efflux of 36Cl and 3H-taurine were inhibited significantly. In contrast, addition of exogenous LTD4 had no significant effect on the swelling-activated 36Cl efflux. Furthermore, exogenous AA increased cytosolic calcium levels as measured in single cells loaded with the calcium sensitive dye Fura-2. On the basis of these results we propose that cell swelling activates phospholipase A2 and that this activation via an increased production of AA or some AA metabolite(s) other than LTD4 is essential for RVD.

    INTRODUCTION
Abstract
Introduction
Methods
Results
Discussion
References

On exposure to hypo-osmotic conditions, most neuronal cells, including the human neuroblastoma cell line CHP-100, initially swell and subsequently undergo regulatory volume decrease (RVD) (Basavappa et al. 1996, 1998). In some neuronal cell types, this characteristic RVD response has, however not been observed (for further discussion see Andrew et al. 1997; Chebabo et al. 1995). In cells that undergo RVD, one principal mechanism utilized is the activation of separate K+ and Cl- conductances. In addition, recent evidence indicates that efflux of amino acids via the swelling-activated anion channel may also play a prominent role during RVD in many cell types including Madin-Darby Canine Kidney cells (Roy and Malo 1992), C6 glioma cells (Strange et al. 1993), a human erytholeukemia cell line (Huang et al. 1996), flounder erythrocytes (Kirk et al. 1992), and most recently in CHP-100 neuroblastoma cells (Basavappa et al. 1996). In contrast, in Ehrlich ascites tumor cells, a separate and distinct organic osmolyte channel has been proposed (Hoffmann and Lambert 1983; Lambert and Hoffmann 1994). The role of second messengers or regulators such as Ca2+ and arachidonic acid during RVD remains relatively unclear, particularly in neuronal cells (for recent reviews see Basavappa and Ellory 1996; Hoffmann and Dunham 1995; Okada 1997; Strange et al. 1996).

Arachidonic acid (AA) and its metabolites such as leukotrienes are involved in many physiological processes (Irvine 1982), including cell volume regulation in several cell types (Hoffmann and Dunham 1995; Lambert et al. 1987; Margalit et al. 1993; Mastrocola et al. 1993; Nilus et al. 1994). In Ehrlich cells, a reduction in extracellular osmolarity has been shown to stimulate phospholipases, which can lead to increased hydrolysis of arachidonic acid from membrane phospholipids (Thoroed et al. 1997). The release of AA is thought to be the rate-limiting step in the production of eicosanoids (Hoffman and Dunham 1995). The released arachidonic acid and/or its metabolites may subsequently regulate the swelling-induced anion conductance (Hoffmann and Dunham 1995).

Phospholipase A2 (PLA2) hydrolyzes membrane phospholipids such as phosphatidyl choline and phosphatidyl ethanolamine to ultimately produce arachidonic acid (Exton 1994). Several isoforms of PLA2 have been identified, including the recently purified 85-kDa cytosolic PLA2 (cPLA2) (Clark et al. 1991; Sharp et al. 1991). Alternatively, AA can be released secondary to the enzymatic action of phospholipase C (PLC), which hydrolyzes membrane phosphatidylinositol 4-phosphate (PIP2) to diacylglycerol (DAG) and inositol 1,4,5-triphosphate (IP3). IP3 in turn catalyzes the release of Ca2+ from intracellular stores, which activates cPLA2. Finally, AA can be released via activation of PLC or phospholipase D (PLD), followed by the action of DAG-lipase and monoacylglycerol (MAG)-lipase (Burgoyne and Morgan 1990).

The release of AA has been demonstrated to be dependent on calcium and/or calmodulin in many cell types (Feinstein and Halenda 1988; Taki and Kanfer 1979; Van den Bosch 1980). This dependency has been correlated with PLA2 activity, because increased levels of intracellular calcium may be a necessary requirement for activation of PLA2. This has been directly demonstrated in platelet membranes, where the activity of PLA2 was dependent on calcium (Nakashima et al. 1989). However, it should be noted that for the 85-kDa cPLA2, the dependency on intracellular calcium concentration ([Ca2+]i) is in the nanomolar range (Clark et al. 1995).


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FIG. 1. Effect of hypo-osmotic stress on 3H-arachidonic acid (3H-AA) release. CHP-100 cells were loaded with 3H-AA for 18 h as described in METHODS. Reducing extracellular osmolarity from 290 to 190 mOsm/kg H2O resulted in a significant increase in rate of 3H-AA release (bullet ; n = 6) as compared with cells exposed to iso-osmotic conditions (open circle ; n = 8). *Significant increase (P < 0.05) when compared with iso-osmotic conditions. Results are shown as cumulative release of radioactivity release relative to total amount of radioactivity as described in METHODS.

Thus the present study investigated the effects of hypo-osmotic stress on AA release in the human neuroblastoma cell line CHP-100 and the effects of exogenous AA on the swelling-activated anion and taurine permeability and intracellular calcium levels. Our results suggest that the swelling-induced activation of the 85-kDa cPLA2 may play a key role in the release of AA in CHP-100 cells during hypo-osmotic stress and in the subsequent activation of Cl- and taurine permeability.

    METHODS
Abstract
Introduction
Methods
Results
Discussion
References

Cell culture

Cells were maintained in culture at 37°C in a 5% CO2 incubator in RPMI 1640 medium (GIBCO, Paisley, UK) supplemented with 10% fetal bovine serum, 100 IU/ml penicillin, and 100 µg/ml streptomycin.

36Cl and 3H-taurine efflux

Anion and taurine permeabilities were evaluated by using 36Cl and 3H-taurine, respectively, by a method modified from Venglarik et al. (1990) and Basavappa et al. (1995). CHP-100 cells grown to 75-90% confluence on 22-mm culture plates (CoStar, Bucks, UK) were incubated with 2 µCi/ml 36Cl (DuPont NEN, Bad Hamburg, Germany) for ~2 h. After two rapid washes, 1 ml of the medium was exchanged at 15, 30, 45, 60, and 120 s with iso-osmotic, hypo-osmotic, or hypo-osmotic solutions containing test agents. In studies with the cPLA2 inhibitor, cells were pretreated for 4 h with 10-µM AACOCF3 (Cascade Biochem, Reading, UK), a trifluromethyl ketone analogue of arachidonic acid (Street et al. 1993). For studies using other test agents, cells were pretreated for 4 min before hypo-osmotic solution. At the end of the study, cells were lysed with 1 ml of 0.1 N NaOH to determine the total amount of radioactivity remaining inside the cells. Ultima Gold (Packard) scintillation fluid was added to the collected samples and vortexed before measurement of radioactivity in a scintillation counter (Packard TRI-CARD 460C Liquid Scintillation System). The total measure of radioactivity was calculated as the sum of radioactivity in all time samples plus the lysed monolayers. Data are represented as counts per minute (cpm) per milliliter as a function of time.

Arachidonic acid (3H-AA) efflux

3H-AA release was measured essentially as described above for 36Cl efflux with the following modifications. Monolayers of CHP-100 cells were loaded for 18 h with 1 µCi/ml 3H-AA (DuPont) in the presence of 0.1% bovine serum albumin (BSA) as described by Stella et al. (1995). The cells usually incorporated 85-90% of the added radioactivity under these conditions. After three rapid washes with iso-osmotic buffer (see Solutions and Reagents) supplemented with 1.0% (wt/vol) BSA to scavenge extracellular fatty acids as described by Thoroed et al. (1997), studies were initiated by adding and removing 1 ml of iso-osmotic buffer at 1-min intervals. After a 2-min period to establish a basal efflux, hypo-osmotic buffer (see Solutions and Reagents) supplemented with 1.0% (wt/vol) BSA was added. Data are presented as the total number of cpm per ml as a function of time after subtraction of background counts. For some studies the data are presented as percentage change in peak release
% peak release = <FR><NU>(peak release − control release)</NU><DE>control release</DE></FR> × 100
where peak release = maximum swelling-induced 3H-AA release and control release = 3H-AA release under iso-osmotic conditions.

Intracellular Ca2+ ([Ca2+]i) measurement in single cells

CHP-100 cells grown on glass coverslips were loaded with the calcium sensitive dye fura-2 AM (Molecular Probes, Leiden, Netherlands) (10 µM) for 30 min at 37°C in a 5% CO2 incubator, followed by three washes with iso-osmotic buffer. Cells were viewed with a Zeiss Axiovert 100 fluorescence microscope, equipped with a ×40/1.4 NA acrostigmat (UV) objective. The instrumental setup was as described by Jorgensen et al. (1996). Briefly, the light source was a 75-W Xenon lamp. Excitation wavelengths of 340 and 380 nm were obtained by BP 340/10 and BP 380/10 filters. A K12 filter and a BPB 380/20 filter were included in the excitation light path to protect the cells against infrared illumination and to adjust the intensity of the excitation light. Emitted light was passed through a BSP 425 dichroic mirror and filtered by a BP 500-530 filter. The fluorescence was viewed with an intensified CCD camera (CCD 72 with a GenIIsys intensifier DAGE-MTI). The images were collected as an average of six frames after 340 and 380 nm excitation, respectively. The ratio of the images obtained after excitation at 340 nm divided by those obtained after excitation at 380 nm was calculated on a pixel-to-pixel basis after background subtraction.


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FIG. 2. Inhibition of swelling-activated 3H-AA release. Cells previously loaded with 3H-arachidonic acid as described in METHODS were pretreated for 4 h with 85-kDa trifluoromethyl ketone analogue of arachidonic acid, AACOCF3 (10 µM), a potent cPLA2 inhibitor. This resulted in a significant (P < 0.05) inhibition of swelling-induced 3H-AA release (black-triangle, n = 3) as compared with cells exposed to hypo-osmotic solution in absence of AACOCF3 (bullet ; n = 6). Inhibition was significant for all points between 3 and 11 min (P < 0.05). open circle , cells exposed to iso-osmotic conditions.

Solutions and reagents

The iso-osmotic buffer contained (in mM) 140 NaCl, 4 KCl, 1 KH2PO4, 2 MgCl2, 1.5 CaCl2, 3 glucose, and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), pH 7.4 with NaOH. The final hypo-osmotic solution contained (in mM) 93 NaCl, 4 KCl, 1 KH2PO4, 2 MgCl2, 1.5 CaCl2, 3 glucose, and 10 HEPES (pH 7.4 with NaOH). For studies with low Ca2+, calcium was omitted from the above solutions and supplemented with 5 mM ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA). The osmolarities of the iso-osmotic and hypo-osmotic solutions, as measured by using a freezing point depression osmometer, were 290 and 190 mOsm/Kg H2O, respectively. All reagents were obtained from Sigma Chemical (St. Louis, MO) unless otherwise specified.

Data analysis

Results are presented as means ± SE, where n refers to number of cells for intracellular Ca2+ and the number of wells for efflux studies. Results in Figs. 1, 2, and 5 are represented as cumulative release of radioactivity (% of total radioactivity per 1 ml/time). Rates of 3H-AA release and 36Cl efflux were calculated by a first-order linear regression. The results in Figs. 3 and 6 are peak values of release relative to the total radioactivity. The results in Fig. 7 are shown as the release of radioactivity during a 1-min time period relative to the total amount of radioactivity. Statistical comparisons were performed by using a Student's t-test with a significance level of P < 0.05. 


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FIG. 5. Effect of AACOCF3 on swelling-activated 36Cl efflux. CHP-100 cells initially loaded with 36Cl as described in METHODS were pretreated with AACOCF3 for 4 h. AACOCF3 pretreatment resulted in a significant (P < 0.05) inhibition of swelling-induced 36Cl efflux (black-triangle; n = 3) as compared with cells exposed to hypo-osmotic solution alone (bullet ; n = 6). open circle , cells exposed to iso-osmotic solution.


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FIG. 3. Effect of phospholipase C (PLC) inhibitor U73122 or ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) on swelling-induced 3H-AA release. Cells were initially loaded with 3H-AA acid, as described in METHODS. After a 15 or 5 min pretreatment with PLC inhibitor, U73122 (100 nM) or EGTA (5 mM), respectively, cells were exposed to hypo-osmotic conditions in the continued presence of these test agents and release of 3H-AA was measured. *Significant (P < 0.05) inhibition as compared with cells exposed to hypo-osmotic solution in absence of EGTA.


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FIG. 6. Effect of extracellular AA on swelling-induced 36Cl efflux. CHP-100 cells loaded with 36Cl as described in METHODS were pretreated with or without 10 µM AA for 5 min and exposed to hypo-osmotic solution in continued presence or absence of 10 µM AA, respectively. Swelling-induced 36Cl efflux (black-triangle; n = 3) was significantly (P < 0.05) inhibited in AA-treated cells, as compared with cells exposed to hypo-osmotic solution alone (bullet ; n = 6). open circle , cells exposed to iso-osmotic solution.


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FIG. 7. Effects of LTD4 on swelling-activated 36Cl efflux. After loading monolayers with 36Cl, cells were pretreated with LTD4 (100 nM) for 5 min. This treatment did not significantly (P > 0.05) alter swelling-induced 36Cl efflux (n = 3) after exposure to hypo-osmotic solution, as compared with cells exposed to hypo-osmotic solution alone.

    RESULTS
Abstract
Introduction
Methods
Results
Discussion
References

3H-AA release

As illustrated in Fig. 1, the rate of arachidonic acid release increased rapidly after exposure to hypo-osmotic solution as compared with control cells. The rate of swelling-induced 3H-AA release was 6.98 ± 0.95%/min as compared with a release of 3.12 ± 0.43%/min under isotonic conditions. This corresponds to an increase of 224 ± 19% (n = 22). The rate of swelling-induced AA release remained constant for approximately eight minutes and gradually decreased for remainder of the study (Fig. 1).


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FIG. 4. Effect of exogenous AA on [Ca2+]i. This representative single-cell Ca2+ recording in Fura-2 loaded cells (see METHODS) shows that the following addition of 50 µM AA to extracellular iso-osmotic solution (down-arrow ) significantly (P < 0.05) increased cytosolic Ca2+ levels as compared with basal levels. Representative of 17 cells in 4 experiments.

One of the principal pathways involved in the release of membrane bound AA is via cleavage of the sn-2 position of glycerophospholipids by PLA2, of which several types have been identified (Clark et al. 1995). Thus the involvement of the recently purified 85-kDa cytosolic PLA2 in the hypo-osmotically induced release of 3H-AA in CHP-100 cells was investigated. For these studies, the cells were pretreated for 4 h with the potent inhibitor of the 85-kDa cPLA2, AACOCF3, a cell permeable trifluoromethyl ketone analog of arachidonic acid (Street et al. 1993). In this arachidonic acid analogue, the COOH group is replaced by COCF3 and the compound exerts its inhibitory action by directly binding to the enzyme (Street et al. 1993). This pretreatment decreased the rate of swelling-induced 3H-AA release to2.46 ± 0.41%/min, which indicates an inhibition of ~65 ± 6% (n = 10; Fig. 2).

Alternatively, AA release may occur via the PLC activated pathway followed by the action of DAG- and MAG-lipase. However, pretreatment of cells for 15 min with the PLC inhibitor U73122 (100 µM) had no significant (P > 0.05) effect on swelling-induced 3H-AA release (n = 6; Fig. 3). Thus these data suggest that swelling-induced 3H-AA release occurs via activation of the 85-kDa cPLA2.

Activation of the 85-kDa cPLA2 is dependent on minor increases in [Ca2+]i either because of calcium entry or release from internal stores (Clark et al. 1995). To evaluate the possible Ca2+ dependency of AA release, cells were pretreated with iso-osmotic buffer containing low Ca2+ and 5 mM EGTA for 5 min (to reduce the concentration of external Ca2+). This treatment with EGTA significantly decreased (P < 0.05) the peak 3H-AA release induced by subsequent cell swelling to that observed in hypo-osmotic control cells (n = 6; Fig. 3). This suggests that influx of calcium may play an important role in the swelling-induced arachidonic release in CHP-100 cells.

[Ca2+]i

Earlier studies in CHP-100 cells have demonstrated that hypo-osmotic stress increases the free intracellular Ca2+ concentration (Basavappa et al. 1995). To determine whether extracellular arachidonic acid effects [Ca2+]i, CHP-100 cells were loaded with fura-2 AM for 30 min and subsequently exposed to 50 µM AA under iso-osmotic conditions. This resulted in a significant increase in [Ca2+]i (n = 17; Fig. 4). The initial rapid rise in intracellular Ca2+ reached a palateau within 150 s of exposure to arachidonic acid and remained stable at this level for remainder of the experiment.

36Cl efflux

Hypo-osmotic stress has been shown to increase anion permeability in CHP-100 cells (Basavappa et al. 1995). The swelling-activated anion permeability may be modulated by a variety of secondary messengers, which appear to be dependent on the cell type (Hoffman and Dunham 1995). The putative role of activation of cPLA2 and increased AA release during hypo-osmotic stress in the regulation of the swelling-activated anion conductance was investigated using monolayers of 36Cl loaded cells pretreated for 4 h with 10 µM AACOCF3. Upon subsequent exposure to hypo-osmotic stress in the absence of added AACOCF3, the swelling-induced 36Cl efflux (n = 12; Fig. 5) was significantly inhibited. Thus inhibition of cPLA2 may block the swelling-activated 36Cl efflux, suggesting that an increase in intracellular AA production during RVD may be involved in the activation of volume-sensitive anion channels.


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FIG. 8. Effect of extracellular AA on swelling-induced 3H-taurine efflux. CHP-100 cells were loaded with 3H-taurine as described in METHODS. In presence of 10 µM AA, swelling-induced 3H-taurine efflux was significantly (P < 0.05) inhibited (square ; n = 6), as compared with cells exposed to hypotonic solution alone. A similar inhibition was observed in cells pretreated with cPLA2 inhibitor AACOCF3 (10 µM) (black-triangle; n = 6) as compared with hypo-osmotic control cells (bullet ; n = 12). open circle , cells exposed to iso-osmotic conditions. Some error bars (SE) are not evident, as they were smaller than size of symbol. Efflux at each time point was expressed as ratio of counts at each time point divided by total number of remaining counts (see METHODS).

In contrast, a high concentration of extracellular AA has been shown to inhibit the volume-sensitive anion channel (Kubo and Okada 1992; Lambert and Hoffmann 1994; Sanchez-Olea et al. 1995). Thus the effects of high concentrations of exogenous AA on the swelling-induced Cl- efflux were investigated in cells pretreated for 4 min with 10 µM AA. The 36Cl efflux induced by subsequent hypo-osmotic exposure was inhibited significantly by 80 ± 5% (n = 24; Fig. 6; P <0.05) in the presence of 10 µM extracellular AA as compared with hypo-osmotic controls. This suggests that high concentrations of exogenous AA may exert its inhibitory action by blocking the swelling-induced anion channel.

Released AA can be either acylated into the cell membrane or metabolized into 1) prostanoids such as prostaglandins via the cyclo-oxygenase pathway, 2) epoxyeicosatrieonic acids such as dihydroxyeicosatrienoic acid (DHETE) via the cytochrome P-450 pathway, or 3) leukotrienes such as LTD4 via the lipoxygenase pathway (Lambert 1994). In Ehrlich acites tumor cells, there is convincing evidence that LTD4 activates swelling-induced anion channels (Hoffmann and Dunham 1995). Thus the effect of LTD4 on the swelling-induced 36Cl efflux was investigated in CHP-100 cells. Pretreatment of monolayers with LTD4 (100 nM) did not significantly alter the swelling-induced 36Cl efflux (n = 9; Fig. 7), as compared with cells exposed to hypo-osmotic conditions in the absence of LTD4.

Taurine efflux

Previous studies in CHP-100 cells have indicated that efflux of taurine plays a significant role during RVD and furthermore that this efflux may occur via the swelling-induced anion channel (Basavappa et al. 1996). We therefore examined the possible role of cPLA2 activation in modulating the swelling-induced 3H-taurine efflux. In cells incubated for 4 h with AACOCF3, a significant inhibition of the 3H-taurine efflux elicited by subsequent hypo-osmotic exposure was observed (n = 6; P < 0.05; Fig. 8). This may further indicate that activation of the 85-kDa cPLA2 is important for swelling-induced anion efflux. In cells pretreated with 10-µM AA and subsequently exposed to hypo-osmotic solution in the continued presence of AA, a significant (P < 0.05) decrease in the swelling-induced 3H-taurine efflux was also observed (n = 6; Fig. 8), as compared with cells exposed to hypo-osmotic stress in the absence of AA. The above data further indicate that high concentrations of exogenous AA may modulate the swelling-induced anion permeability.

    DISCUSSION
Abstract
Introduction
Methods
Results
Discussion
References

Under resting conditions, intracellular arachidonic acid is maintained at very low levels (Irvine 1982). In response to volume perturbation or shear stress, increased AA release has been observed in some cell types (Civan et al. 1994; McManus et al. 1994; Pearce et al. 1996; Thoroed et al. 1997). In the CNS, increased levels of arachidonic acid have been implicated to play a role in many physiological response processes, including cell volume regulation. During ischemia and subsequent to seizure activity, increased levels of AA have been detected in the brain (Bazan 1989; Siesjo et al. 1989). The present study demonstrates that in the human neuroblastoma cell line CHP-100, a reduction in extracellular osmolarity significantly increased AA release and that this release appears to be mediated by the 85-kDa cPLA2. Although several routes of AA release from the cell membrane have been identified, the major precursors are likely to involve glycerophospholipids, phosphatidylcholine, and phosphatidylethandamine.

As discussed previously, the release of AA appears to be mainly catalyzed by PLA2. Two major forms of PLA2 have been identified, the recently purified 85-kDa cPLA2 and the secreted 14-kDa group I and II PLA2 (sPLA2) (Clark et al. 1995; Davidson and Dennis 1990). There is no sequence homology between cPLA2 and sPLA2 (Clark et al. 1995). The 85-kDa cPLA2 has been identified in many diverse tissues including the hippocampus, murine brain, and astrocytoma cell line and is activated by various stimuli (Clark et al. 1995). In the present study, in cells pretreated with the 85-kDa cPLA2 blocker AACOCF3 (Street et al. 1993), a significant reduction in the rate of swelling-induced release of AA was observed.

In contrast, the PLC inhibitor U73122 did not significantly inhibit swelling-induced AA release. Taken together, these results strongly suggest that release of arachidonic acid in CHP-100 cells occurs via cleavage of the sn-2 position of phospholipids by a cPLA2 catalyzed pathway. This is in agreement with a recent study by Pearce et al. (1996) in human endothelial cells where activation of cPLA2 by shear stress (which maybe equivalent to "hypo-osmotic stress") resulted in increased AA release. Hoffmann and Dunham (1995) have discussed several possible mechanisms behind swelling-induced activation of cPLA2. In Ehrlich cells, Thoroed et al. (1997) report that cell swelling activates cPLA2 mediated AA release. They demonstrate that in cells pretreated with AACOCF3, the swelling-induced 36Cl efflux as well as the swelling-induced 14C-taurine efflux were significantly inhibited. This suggests that swelling-induced release of AA via stimulation of the 85-kDa cPLA2 may play an important role in the activation of swelling-induced anion and taurine conductance.

Earlier studies in CHP-100 cells demonstrated that hypo-osmotic exposure resulted in an increase in cytosolic Ca2+ levels (Basavappa et al. 1995). In the present study, significant increases in [Ca2+]i were observed after addition of AA. Exogenous AA has been shown to increase [Ca2+]i in a variety of cell types by either influx or release of internal stores. Recent studies by Tinel et al. (1997) in rat inner medullory collecting duct (IMCD) cells report a similar increase in [Ca2+]i after addition of extracellular AA. Furthermore, these authors suggest that IMCD cells possess arachidonic-acid-sensitive Ca2+ stores, as [Ca2+]i levels were increased with both high and low extracellular Ca2+ containing solutions. The previously observed swelling-induced elevation in [Ca2+]i in CHP-100 cells may be a requirement for swelling-induced AA release, as the hypo-osmotically induced release of AA was significantly decreased in the absence of extracellular Ca2+ and in the presence of EGTA. Interestingly, Clark et al. (1990) report that the Ca2+ dependence of cPLA2 is biphasic, with the initial elevation in cPLA2 activity occurring in the submicromolar range and the secondary increase occurring at a [Ca2+]i between 0.1 and 10 mM. Our previous observations of inhibition of swelling-activated anion channels and taurine efflux with decreased levels of intracellular Ca2+ (Basavappa et al. 1995; Basavappa and Ellory 1996), in combination with the Ca2+ dependence of cPLA2, suggests that inhibition of swelling-induced anion permeability at low [Ca2+]i may be a consequence of decreased AA production.

Three principal pathways are involved in the metabolization of released AA. In particular, LTD4, one of the major products of the lipoxgenase pathway has been implicated in volume regulation in some cell types, including Ehrlich cells (Jorgensen et al. 1996; Lambert and Hoffmann 1994). In CHP-100 cells, in contrast to the observations in Ehrlich cells, addition of LTD4 did not have any significant effects on the swelling-induced anion permeability. Similarly, in NPE cells the leukotrienes A4, D4, and E4 had no effects on RVD (Civan et al. 1994) and in C6 glioma cells, cell swelling did not result in significant production of any of the metabolites of the lipoxygenase pathway (McManus et al. 1994). In contrast, in Ehrlich ascites tumor cells, increased AA release resulting from cell swelling augmented LTD4 production resulting in an increase in Cl- conductance (Lambert et al. 1987). Whether or not other metabolites of the lipoxgenase pathway distinct from LTD4 play a role in the swelling-activated anion conductance in CHP-100 cells is unknown at present. For instance, in human platelets hepoxillin A was suggested to play a role in RVD (Margalit et al. 1993). Finally, release of AA from phosphatidylcholine via activation of cPLA2 will also result in an increase in released lysophosphatidic acid, which could effect [Ca2+]i and anion permeability (Moolenaar 1995).

Exogenous AA, as such, has been previously reported to inhibit ion conductances in several cell types (Anderson and Welsh 1990; Chan and Fishman 1978; Kubo and Okada 1992; Meves 1994). During hypo-osmotic stress, extracellular AA blocked the volume-activated Cl- channels in Ehrlich ascites tumor cells (Lambert and Hoffmann 1994), C6 glioma cells (Jackson and Strange 1993), rat cerebellar astrocytes (Sanchez-Olea et al. 1995), HSG cells (Fatherazi et al. 1994) and Intestine 407 cells (Kubo and Okada 1992). In Intestine 407 cells, added AA reversibly blocked the volume-sensitive anion channels with an IC50 of 8 µM. Similar results were observed in the present study in CHP-100 cells. The swelling-induced 36Cl and 3H-taurine efflux were significantly inhibited with extracellular AA. Thus there is a contrast between the intracellular effects of AA (or its metabolites), which is stimulatory, while addition of high concentrations of exogenous AA inhibits the volume-sensitive anion channel. This opposing effect by extracellular and intracellular AA on ion-channel modulation is not unique. Wieland et al. (1992) in skeletal muscle demonstrated that exogenous AA inhibited sodium currents, while internal application of AA activated sodium currents. They speculate that differential modulation by AA may be a result of direct and indirect pathways. However, a number of AA metabolites can potentially modulate ion channels (Meves 1994). The present studies examined only one metabolite (i.e., LTD4) and additional studies are essential to delineate whether the inhibition is a result of unspecific effects of addition of fatty acids such as membrane fluidity, or effects on PKC or a metabolite of AA. Our present results suggest that modulation of ion permeability by AA is dependent on the exposure site (i.e., intracellular vs. extracellular). To our knowledge this is the first demonstration of differential effects by AA on neuronal anion permeability.

In CHP-100 cells, the swelling-activated Cl- and taurine efflux were suggested to share a common volume-activated anion pathway (Basavappa et al. 1996). The present observations that swelling-activated taurine as well as Cl- efflux were inhibited by exogenous AA and were furthermore, sensitive to pretreatment with AACOCF3, further indicate that taurine and chloride may share a common pathway during hypo-osmotic stress in CHP-100 cells.

Taken together, the present studies suggest that cell swelling results in an increased release of archidonic acid in CHP-100 cells, which is most likely generated by swelling-induced activation of the 85-kDa cPLA2. Inhibition of cPLA2 and hence of AA release inhibits swelling-induced Cl- and taurine efflux. Therefore, AA released by a cPLA2-mediated pathway during cell swelling may significantly contribute to the RVD process in neuroblastoma cells. The AA metabolite LTD4 does not appear to be involved in modulating the swelling-induced anion conductance, but other metabolites of AA may play a putative role. Finally, in contrast to the stimulatory effects of intracellular released AA during RVD, a high concentration of exogenous added AA appears to directly inhibit the swelling-activated anion conductance in CHP-100 cells.

    ACKNOWLEDGEMENTS

  The human neuroblastoma cell line CHP-100 was provided by Dr. A. Wilson and A. Wilson from the Children's Hospital of Philadelphia.

  This work was supported by the European Molecular Biological Organization (short-term fellowship for S. Basavappa), the Wellcome Trust, and the Danish Natural Science Research Council.

    FOOTNOTES

  Address for reprint requests: S. Basavappa, Dept. of Anesthesiology, Vanderbilt University School of Medicine, 504 Oxford House, 1321 21st St., Nashville, TN 37232.

  Received 15 July 1997; accepted in final form 13 November 1997.

    REFERENCES
Abstract
Introduction
Methods
Results
Discussion
References

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