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Biophys J. 2007 Oct 1; 93(7): 2350–2362.
Published online 2007 Jun 1. doi:  10.1529/biophysj.107.104679
PMCID: PMC1965435

Relationship between Membrane Physical Properties and Secretory Phospholipase A2 Hydrolysis Kinetics in S49 Cells during Ionophore-Induced Apoptosis


During apoptosis, changes occur in lymphocyte membranes that render them susceptible to hydrolysis by secretory phospholipase A2 (sPLA2). To study the relevant mechanisms, a simplified model of apoptosis using a calcium ionophore was applied. Kinetic and flow cytometry experiments provided key observations regarding ionophore treatment: the initial rate of hydrolysis was elevated at all enzyme concentrations, the total amount of reaction product was increased fourfold, and adsorption of the enzyme to the membrane surface was unaltered. Analysis of these results suggested that susceptibility during calcium-induced apoptosis is limited by availability of substrate rather than adsorption of enzyme. Fluorescence experiments identified three membrane alterations during apoptosis that might affect substrate access to the sPLA2 active site. First, intercalation of merocyanine 540 into the membrane was improved, suggesting an increase in lipid spacing. Second, laurdan detected increased solvation of the lower headgroup region of the membrane. Third, the rate at which fluorescent lipids could be removed from the membrane by albumin was enhanced, implying greater vertical mobility of phospholipids. Thus, it is proposed that the membranes of apoptotic cells become susceptible to sPLA2 through a reduction in lipid-neighbor interactions that facilitates migration of phospholipids into the enzyme active site.


Apoptosis is a complex process involving numerous alterations to components of the nucleus, cytoplasm, and cell membrane (1,2). Changes occurring at the cell membrane include exposure of phosphatidylserine, ceramide production, microvesicle shedding, membrane blebbing, loss of membrane potential, alterations to cytoskeletal attachments, and loss of membrane integrity (19). One of the earliest membrane events documented thus far is a substantial increase in the action of secretory phospholipase A2 (sPLA2) to hydrolyze plasma membrane lipids (9,10).

sPLA2 has been used extensively for studying enzymatic activity at the water-membrane interface. A key feature of this enzyme is its ability to distinguish between healthy cells, which resist hydrolysis by sPLA2, and damaged, necrotic, or apoptotic cells, which are often susceptible. Studies with artificial bilayers have demonstrated that the extent to which phospholipids are hydrolyzed by the enzyme depends on specific physical properties associated with the membrane (1117). It has been proposed that analogous properties also govern the susceptibility of cell membranes (1821).

A variety of experimental approaches has led to a general concept in which the action of sPLA2 on the membrane surface involves at least two precatalytic steps (11,12,2226). In the first step, sPLA2 adsorbs to the membrane surface, and in the second, an individual phospholipid migrates into the active site of the enzyme (Scheme 1). In previous studies, this two-step hypothesis was examined using human erythrocytes as a simplified model for cell membranes (21). For those investigations, the apoptotic state of the cell was mimicked by treatment with ionomycin, a calcium ionophore. The resulting calcium influx produced changes in membrane structure accompanied by increased susceptibility to hydrolysis by sPLA2. Analysis of these observations generated the conclusion that the ability of sPLA2 to attack cell membranes is determined primarily by the ease of vertical migration of phospholipids into the active site of adsorbed enzyme (i.e., the second step in Scheme 1).

Reaction scheme for interaction between sPLA2 and cell membranes. E, free sPLA2; EB, sPLA2 adsorbed to membrane surface sites (Ms); S, available substrate (membrane phospholipid); equation M1 adsorbed enzyme with substrate bound to the active site; KA, equilibrium ...

This foundational work from various membrane models has thus generated a testable hypothesis relating bilayer structure and sPLA2 activity. The ultimate goal is a complete understanding of the biophysical and molecular principles governing this relationship in physiological and pathological settings such as apoptosis. Accomplishing this goal requires experiments with nucleated cells. Given the complexities associated with both apoptosis and cellular models for biophysical studies, we sought an experimental system in which measured effects would be maximized, the relationship between sPLA2 activity and apoptotic markers would be known, and links to previous mechanistic studies would be plausible. Accordingly, S49 lymphoma cells treated with calcium ionophore (ionomycin) were chosen because they meet these criteria. Ionomycin stimulates rapid apoptosis that appears synchronized for the majority of the cell population resulting in large and reproducible changes in measured events. Moreover, as in human erythrocytes, the drug induces susceptibility to sPLA2 that, in S49 cells, occurs with or before other major apoptotic changes (9,27).

The monomeric aspartate-49 phospholipase A2 from the venom of Agkistrodon piscivorus piscivorus was used as the source of enzyme for several reasons. First, it has been used extensively in previous mechanistic and structural investigations, including studies with erythrocytes and S49 cells (9,11,1521,23,25,27,28). Second, it is easily purified in large quantities, making it amenable to biophysical experiments. Third, it behaves similarly to some of the human isoforms of the enzyme during ionomycin-stimulated apoptosis (27).

Experiments were designed to examine each of the parameters shown in Scheme 1 to assess which is/are involved in distinguishing resistant cells from those treated with ionophore. This assessment was accomplished through measurements and analysis of hydrolysis kinetics, enzyme adsorption, and membrane structure. The results demonstrated that modulation of the second step (KE in Scheme 1) accounts for the increased susceptibility to sPLA2.



The monomeric aspartate-49 phospholipase A2 from the venom of A. p. piscivorus was isolated according to the procedure by Maraganore et al. (29). Ionomycin was purchased from Calbiochem (La Jolla, CA) and dissolved in dimethylsulfoxide (DMSO) for experiments. The probes Oregon Green 488, 2-(6-(7-nitrobenz-2-oxa-1 3-diazol-4-yl)amino)hexanoyl-1-hexadecanoyl-sn-glycero-3 phosphocholine (NBD-PC), laurdan, acrylodan-labeled fatty acid-binding protein (ADIFAB), 1,3-bis-(1-pyrene)propane (bis-pyrene), diphenylhexatriene (DPH), merocyanine 540 (MC540), and propidium iodide were acquired from Invitrogen (Carlsbad, CA). Bovine serum albumin (BSA) was purchased from Sigma (St. Louis, MO). Other reagents were obtained from standard sources. Probes were dissolved in various solvents as follows: ADIFAB in 50 mM KCl and 3 mM NaN3, bis-pyrene and DPH in N,N-dimethylformamide, NBD-PC in ethanol, and all others in DMSO.

Cell culture and experimental protocol

S49 mouse lymphoma cells were grown at 37°C in humidified air containing 10% CO2 and prepared for experiments as described (28). Cell viability averaged 89 ± 6% (standard deviation). For spectral and kinetic experiments, an aliquot of cells (usually between 0.4 × 106 and 3.0 × 106 cells/ml in modified balanced salt solution (MBSS)) was then transferred to a quartz fluorometer sample cell and allowed 5 min to equilibrate in the spectrofluorometer (Fluoromax 3, Horiba Jobin-Yvon, Edison, NJ, or PC-1, ISS, Champaign, IL). Spectral bandpass varied between 4 and 16 nm depending on the intensity of the probe and instrument sensitivity. Data acquisition over time at multiple wavelengths (ADIFAB, laurdan, and bis-pyrene) was accomplished by rapid sluing of fluorometer mirrors under control of instrument software. Temperature and sample homogeneity were maintained as described previously (19). All experiments and incubations were performed at 37°C. For enzyme adsorption experiments, cells were resuspended in MBSS with 20 mM BaCl2 to inhibit sPLA2 hydrolytic activity without preventing adsorption (26,30,31).

In each procedure where ionomycin was added to cell samples (300 nM final), the experiment was repeated with an equivalent volume of DMSO (0.25% v/v final) to control for the effects of the solvent. A control for direct effects of ionomycin was also included in assessments of membrane physical properties (see below).

Membrane hydrolysis

Release of free fatty acids from cell membranes was assayed in real time with the fatty acid binding protein ADIFAB (excitation, 390 nm; emission, 432 nm and 505 nm). After initiating data acquisition for 100 s, ADIFAB (65 nM final) and ionomycin were added. At 700 s, sPLA2 (0.7–70 nM final) was added and the time course continued for an additional 800 s. Raw data were quantified by calculating the generalized polarization (GP) and fitting to an arbitrary function by nonlinear regression (19). In some cases, hydrolysis was assayed by measurement of the rate of propidium iodide (37 μM final) uptake by cells due to the action of sPLA2 as described (28). Previous experiments have demonstrated that assays with ADIFAB and propidium iodide give identical results under conditions at which cells are susceptible to the enzyme's catalytic activity (28). Thus, the propidium iodide assay was used only in experiments in which all samples were treated with ionomycin.

Enzyme adsorption

Adsorption of sPLA2 to the cell surface was assayed by flow cytometry with Oregon Green 488-labeled sPLA2 (L-PLA2). Enzyme labeling was accomplished according to instructions provided with the kit (Invitrogen, Carlsbad, CA). Flow cytometry data were collected using a BD FACSCanto flow cytometer (BD Biosciences, San Jose, CA) with an argon excitation laser (excitation 488 nm, longpass 502 nm, bandpass filter 515–545 nm). Aliquots of cells (suspended in 1 ml MBSS containing 20 mM BaCl2) were treated with ionomycin or DMSO for 10 min. For isotherm experiments, varying amounts of L-PLA2 were then added (0.01–3 μM final), and samples were incubated for an additional 10 min before data acquisition. For competition experiments, samples were mixed with various concentrations of sPLA2 (0.05–1 μM final) for 5 min before adding L-PLA2 (50 nM final), and the incubation was continued for an additional 5 min. Samples were then immediately processed in the flow cytometer.

Probes of membrane structure

In addition to the “vehicle control” (DMSO), a control experiment was designed to verify that calcium entry into the cell, rather than artifacts resulting from direct effects of ionomycin, was responsible for the observed changes in membrane structure. This control involved incubation of the sample for at least 200 s with the calcium chelator ethylenediaminetetraacetic acid (EDTA) (18 mM final; the pH of the EDTA stock was adjusted to 7.6 at 37°C to avoid altering sample pH) before addition of ionomycin. In every case, the results from this “EDTA control” were indistinguishable from those obtained with the vehicle control. Hence, data from both types of controls are pooled in the presentation of results.

Data with MC540 (170 nM final) were acquired as emission spectra (excitation, 540 nm; emission, 550–700 nm). Spectra were obtained before the addition of the probe, after equilibration (5 min), and 10 min after addition of ionomycin or control reagent(s). Spectra were analyzed by subtracting the initial spectrum as background and integrating the intensity from 565 to 615 nm. Effects of experimental treatments were quantified by calculating the difference in integrated intensity before and after treatment. That difference was divided by the integrated intensity before treatment to account for variations in cell number and lamp intensity. Changes in the wavelength of peak intensity were identified from derivatives of the spectra.

Laurdan or bis-pyrene fluorescence emission was acquired at dual wavelengths in real time. After initiating data acquisition, laurdan (50 nM final; excitation, 350 nm; emission 435 and 500 nm) or bis-pyrene (2.2 μM final; excitation, 344 nm; emission 376 and 480 nm) was added at 100 s. Ionomycin or control reagent(s) was/were included after initial equilibration of cells with the probe (laurdan, 10 min; bis-pyrene, 30 min). The laurdan GP and bis-pyrene excimer (480 nm)/monomer (376 nm) ratio were calculated from the data as described (19,32). Background intensity at each wavelength was subtracted from the data before calculations were made. For two-photon excitation microscopy of laurdan fluorescence, the concentration was increased to 250 nM to improve image distinction compared to background.

Data for steady-state anisotropy (DPH; excitation, 350 nm; emission, 452 nm) were acquired using Glan-Thompson polarizers in the vertical and horizontal positions. Data were obtained before the addition of the probe to measure background cell fluorescence, after 20 min equilibration with DPH (240 nM final), and 10 min after addition of ionomycin or control reagent(s). Anisotropy was calculated as described (33).

Phospholipid extraction

The fluorescent phospholipid analog, NBD-PC, was suspended in MBSS immediately before the experiments, as described by Jensen et al. (21). Cell samples were incubated with ionomycin or control reagent(s) (as in previous section) for 5 min. NBD-PC (930 nM final) was then introduced and allowed to incorporate into cell membranes (∼30 min). After this incubation, cells were washed to remove excess probe. Data acquisition (excitation, 485 nm; emission, 535 nm) was then initiated, and BSA (0.7% final) was added for NBD-PC extraction. Apparent extraction-rate constants were obtained by fitting the fluorescence intensity data after BSA addition to a double exponential decay function by nonlinear regression.

Two-photon excitation scanning microscopy

Scanning two-photon excitation microscopy images were obtained using an Axiovert 35 inverted microscope (Zeiss, Thornwood, NY) at 37°C at the Laboratory for Fluorescence Dynamics (University of California, Irvine) as described previously (19,34). Laser emission was 940 nm for NBD-PC and 790 nm for Oregon Green 488. For laurdan (250 nM), dual images were collected simultaneously using a beam-splitter and interference filters (Ealing 490 and Ealing 440) with laser emission of 790 nm.


Hydrolysis kinetics

Addition of sPLA2 to S49 cells previously exposed (10 min) to a control vehicle (DMSO) caused a slight increase in ADIFAB GP followed by a gradual decline, nearly restoring the value to its original level (Fig. 1, curve a). This unusual hydrolysis time course has been shown to represent transient initial hydrolysis followed by reacylation of the lipids and reincorporation into the membrane (27). In contrast, prior exposure of cells to ionomycin resulted in a large increase in ADIFAB GP that quickly reached a stable plateau (Fig. 1, curve b).

Effects of ionomycin treatment on the time course of membrane hydrolysis by sPLA2. S49 cells were incubated with ADIFAB and treated for at least 600 s with DMSO (curve a) or ionomycin (curve b) as explained in Materials and Methods. At the time indicated ...

To further characterize hydrolysis kinetics, the experiment in Fig. 1 was repeated at various enzyme concentrations. Data were analyzed by nonlinear regression (examples are shown in Fig. 1). From these phenomenological fits, two parameters were determined: the initial rate of hydrolysis and maximum hydrolysis product generated. As displayed in Fig. 2, treatment of cells with ionomycin increased both initial and total hydrolysis at all enzyme concentrations. The maximum hydrolysis rate was 0.018 ± 0.004 GP units s−1 for DMSO-treated samples and 0.093 ± 0.011 GP units s−1 for the ionomycin group. The enzyme concentration at which the hydrolysis rate was one-half the maximum was 13.5 ± 7.8 nM and 9.4 ± 3.2 nM, respectively, for the two groups. Two-way analysis of variance indicated that both the effects of enzyme concentration and ionomycin treatment on the initial hydrolysis rate were highly significant (p < 0.0001). A significant interaction (p < 0.0001) between the two variables was also observed, suggesting that the magnitude of the effect of ionomycin depended on enzyme concentration. The ability of ionomycin treatment to increase total hydrolysis product was also significant (p < 0.0001).

sPLA2 concentration dependence of the initial rate of hydrolysis. (A) The experiments of Fig. 1 were repeated at the indicated concentrations of sPLA2 for cells treated with DMSO (squares) or ionomycin (triangles). The initial rate was defined as the ...

Enzyme adsorption

Initial attempts to quantify adsorption of sPLA2 to the cell surface involved a conventional centrifugation assay with fluorescent-labeled enzyme (L-PLA2), as done previously with erythrocytes (21). However, control experiments suggested that the observed association of enzyme with the cells was not reversible. Analysis of microscopy images indicated that this irreversible interaction reflected endocytosis of the labeled enzyme, a phenomenon previously described for nucleated cells (3537).

Flow cytometry experiments were designed to distinguish endocytosis of labeled enzyme from reversible adsorption to the cell surface. Cells were treated with ionomycin or DMSO for 5 min. All samples were then mixed with L-PLA2, and half of the samples were washed to remove free enzyme. Fig. 3 displays histograms of L-PLA2 fluorescence intensity per cell in each of these four groups. Most of the data from DMSO-treated samples appeared to originate from a single population of cells (Fig. 3 A). The intensity of this group was reduced 10-fold (i.e., one log unit along the abscissa) by washing (Fig. 3 C), suggesting that this peak represented reversible adsorption of L-PLA2 to the cell exterior. In unwashed ionomycin-treated samples (Fig. 3 B), at least two distinct populations were identified in the histogram of fluorescence intensity. Cells with low fluorescence intensity (Fig. 3 B, left arrow) accounted for a majority of the sample, yet they only contributed ∼20% of the total fluorescence signal. The fluorescence of this population was also reduced 10-fold by washing, similar to the DMSO-treated cells (Fig. 3 D, left arrow). In contrast, the high intensity staining was less affected by washing (Fig. 3, B and D, right arrow), suggesting that this staining represented internalization of the enzyme (3537). This irreversibility corroborated observations from two-photon microscopy (not shown; see previous paragraph). Given that reversible surface adsorption is the relevant phenomenon for interpreting the hydrolysis kinetics, we focused our analysis on the low-intensity population.

Reversibility of adsorption of L-PLA2 to S49 cells. Data represent histograms for Oregon Green fluorescence intensity obtained by flow cytometry, as explained in Materials and Methods. (A and B) Cells were treated with DMSO (A) or ionomycin (B) for 5 ...

Fig. 4 displays the amount of reversible surface adsorption of various concentrations of L-PLA2. The average apparent equilibrium constant for adsorption of L-PLA2 to DMSO-treated cells was 2.5 × 106 M−1. There was no detectable effect of ionomycin treatment on either the apparent equilibrium constant (1.7 × 106 M−1, p = 0.39) or the maximum number of adsorption sites per cell (p = 0.20).

Concentration dependence of L-PLA2 adsorption to S49 cells. (A) The experiments of Fig. 3, A and B, were repeated at the indicated concentrations of L-PLA2. The amount of adsorbed L-PLA2 was estimated from the fluorescence intensity represented by the ...

To relate these adsorption results to native sPLA2, competition experiments were performed. Unlabeled sPLA2 competed for ∼30–40% of the L-PLA2 adsorption sites at all concentrations within the range tested (0.05–1 μM sPLA2 with 50 nM L-PLA2). This result suggested that more than half of these adsorption sites might not be relevant to hydrolysis by sPLA2. The observation that competition was already maximal when the same concentrations of sPLA2 and L-PLA2 were used (50 nM) indicated that the enzyme adsorbs more tightly when it is not labeled. This inference is consistent with the relatively high potency of the unlabeled enzyme shown in Fig. 2.

To better estimate the apparent adsorption constant for unlabeled sPLA2 (Kapp), we took advantage of control observations (not shown) demonstrating that L-PLA2 is a weak enzyme (possessing ≤1/10 the activity of sPLA2). Accordingly, we repeated the competition experiments using a hydrolysis assay at constant sPLA2 concentration (70 nM final, ET in Eq. 1) in the presence of various amounts of L-PLA2 (0.05–20 μM final, LT in Eq. 1). The initial hydrolysis rate data (dP/dt) were fit to the following equation with α as an arbitrary proportionality constant:

equation M2

We assumed that KL, the adsorption constant for L-PLA2, was the same at sites of hydrolysis as that estimated for the entire cell in Fig. 4. Results shown in Fig. 5 provide convincing evidence for competition. Analysis of the data by nonlinear regression estimated that Kapp equals 2.7 × 107 M−1.

Ability of L-PLA2 to interfere with cell membrane hydrolysis by sPLA2. Cells were treated for 400 s with ionomycin and then subjected to hydrolysis by 70 nM sPLA2. Hydrolysis was assayed using propidium iodide (see Materials and Methods). The relative ...

Physical changes in the cell membrane

Previous experiments with erythrocytes revealed several physical changes in the cell membrane that appeared to explain the basis for increased susceptibility to the action of sPLA2. These included increases in the spacing among bilayer lipids, the average level of lipid order, and the rate at which fluorescent phospholipids could be extracted from the cell membrane (21). Similar experiments have been included here to test the relevance of those findings to nucleated cells.

Phospholipid spacing

Changes in phospholipid spacing were assessed with MC540, a probe that intercalates between the phospholipid heads in the outer leaflet of cell membranes (3840). Treatment of cells with ionomycin resulted in a significant increase in MC540 total fluorescence (Fig. 6 A) and a red-shift in peak wavelength from ∼583 to 586 nm (Fig. 6 B), consistent with increased binding of the probe (41). As shown in Fig. 6, C and D, these effects were reproducible. Control experiments in which cells were treated with ionomycin in the presence of a calcium chelator (EDTA) demonstrated that these effects were due to calcium influx rather than representing artifacts caused directly by the presence of ionomycin in the membrane. Since EDTA controls were indistinguishable from DMSO-treated samples, data from both types of controls were pooled in the presentation of results for MC540 and all other probes of membrane physical properties (see legends to Figs. 6–9).). These spectral changes in MC540 fluorescence suggest that treatment with ionomycin increased the spacing among membrane phospholipids.

Effects of ionomycin treatment on MC540 fluorescence. (A) Representative emission spectra showing the intensity of MC540 bound to S49 cells before (solid curve; labeling details explained in Materials and Methods) and 10 min after ionomycin addition ( ...
Effect of ionomycin on laurdan GP. (A) Cells labeled with laurdan were treated without (curve a, vehicle control) or with (curve b) ionomycin (arrows). (B) For each experiment, 30 points before treatment with ionomycin were averaged and pooled in the ...
Effect of ionomycin on bis-pyrene and DPH fluorescence. (A) Time course of bis-pyrene excimer/monomer ratio; ionomycin was added at the arrow. (B) Cells were labeled with DPH as explained in Materials and Methods, and anisotropy measurements were obtained ...
Effect of ionomycin on the rate of NBD-PC extraction by BSA. (A) Time course of NBD-PC fluorescence in samples treated with ionomycin (gray) or control reagent(s) (black) with t = 0 corresponding to the time of BSA addition. Data points are normalized ...

Membrane order

Three fluorescent probes were used to examine the relationship between susceptibility and membrane order. Laurdan intercalates at the level of the glycerol backbones (42). The shape of laurdan's emission spectrum is highly sensitive to the solvent relaxation effect and is quantified as GP (see Materials and Methods). GP varies between −1.0 and 1.0 and has been interpreted to reflect the degree to which lipid molecules are ordered within the membrane (a value of 1.0 denotes a highly ordered membrane in which no solvent relaxation occurs (32,33)). Bis-pyrene has traditionally been used to evaluate the level of “fluidity” in the membrane, implying the ability of molecules to diffuse laterally along the bilayer plane (43). More recent experiments exploring the phosphatidylcholine/cholesterol phase diagram indicated that the probe is more sensitive to phospholipid acyl chain order (44). Presumably, bis-pyrene resides deep in the membrane, based on the partitioning of the parent compound, pyrene (45). Diphenylhexatriene also locates deep in the membrane (46). The anisotropy of DPH is sensitive to both fluidity (which alters the rotational diffusion rate of the probe) and environmental constraints imposed by the level of chain order (44,47).

Laurdan GP before and after treatment with ionomycin and subsequent controls is illustrated in Fig. 7. In Fig. 7 A, laurdan GP over time is displayed. Ionomycin (added at arrow) caused GP values to decrease until it reached a minimum 10 min after treatment. This effect was significant (Fig. 7 B). Images collected by two-photon microscopy verified that the changes reported in Fig. 7 B were confined to the plasma membrane (Fig. 7, CF). In contrast, no differences in apparent lipid order were identified with either bis-pyrene excimer/monomer ratio or DPH steady-state anisotropy (Fig. 8).

Phospholipid extractability

Changes in the propensity of phospholipids to migrate vertically in the bilayer were estimated by extraction experiments utilizing NBD-PC, a fluorescent phospholipid analog that inserts into lipid bilayers (21,48). We used the protein BSA as a lipid acceptor in these experiments. This protocol allowed us to estimate the rate of extraction. As shown in Fig. 9 A, time courses of NBD-PC fluorescence after BSA addition were fit to a two-phase exponential decay (with t = 0 corresponding to the time of BSA addition), with a rapid initial drop in intensity followed by a gradual decline. An overall rate constant was derived from the regressions and compared for multiple samples (Fig. 9 B). Ionophore treatment generated a reproducible twofold increase in the rate of NBD-PC extraction.

Initial and final fluorescence intensities often differed between DMSO- and ionomycin-treated samples (generally greater in the ionophore group, not shown). For this reason, the data in Fig. 9 A are normalized. We used flow cytometry and two-photon microscopy to address the possibility that intensity differences reflected artifacts (due to the extent of extraction and/or labeling) that might interfere with our analysis. Flow cytometry experiments demonstrated that the amount of fluorophore removed by BSA was comparable between control and treatment groups (89% and 85%, respectively (not shown)). Two-photon microscopy images revealed that it was common for cells to display labeling of internal compartments in addition to the plasma membrane (more prominent in ionomycin-treated cells; example shown in Fig. 9, CE). Upon BSA addition (between Fig. 9, C and D), fluorescence intensity decreased quickly from the perimeter of the cells and more gradually from the interior (Fig. 9 D, 2 min after addition; Fig. 9 E, 4 min after addition). These experiments convinced us that at least the initial rapid decay of fluorescence intensity, dominant in the analysis shown in Fig. 9, A and B, could reasonably be interpreted to represent lipid extraction from the cell membrane.


The purpose of this study was to identify how changes in the cell membrane induced by calcium ionophore treatment might increase the rate of phospholipid hydrolysis by extracellular sPLA2. To this end, the model shown in Scheme 1 was used to analyze the results of the kinetic experiments displayed in Fig. 2. The initial rate of lipid catalysis (dP/dt) is given by

equation M3

where β is proportional to kcat and the total number of adsorption sites for the enzyme (ET) (21). It is assumed that β does not change with experimental treatment because alterations to membrane structure would presumably affect reaction steps preceding the catalytic event (see Jensen et al. (21) and discussion below). For simplicity, β is assigned a value of 1.0 GP units s−1. ST represents the mole fraction of membrane phospholipids available to the enzyme. In previous analyses, it has not been considered separately from KE since the two parameters are completely linked in the model (see Eq. 2 and Jensen et al. (21)). However, the data of Fig. 2 B demonstrate that ST is subject to change upon experimental treatment and provide information to quantify the magnitude of that change. ST is therefore included specifically as a parameter in Eq. 2 and expressed in relative units (i.e., it is assigned a value of 1.0 under control conditions).

As shown in Fig. 10 A, the control data from Fig. 2 A were well fit by Eq. 2. The value of KA (6.7 × 107 M−1) matched favorably the value estimated from competition with L-PLA2 (2.7 × 107 M−1 in Fig. 5). The effect of ionomycin on the initial hydrolysis rate was evaluated by considering ST, KA, and KE individually in the fit to identify the minimum criteria required by the results (parameter values summarized in Table 1). The data of Fig. 2 B indicated that ST increased by a factor of ∼4.4 with ionophore treatment. Accordingly, Eq. 2 was recalculated maintaining all parameter values from the fit of the control data with the exception of ST. The 4.4-fold increase, however, was insufficient to account for the ionomycin data (Fig. 10 B). In fact, a reasonable fit was obtained only with a much larger (8.1-fold) increase in ST, one that cannot be justified based on the data of Fig. 2 B (Fig. 10 C). Since KE and ST are linked in Eq. 3, the dilemma can obviously be resolved by increasing KE by a factor of 1.8 in addition to a fixed 4.4-fold increase in ST (Fig. 10 D). Alternatively, allowing KA to vary instead of KE was inadequate to accommodate the data (Fig. 10 E). The best fit, of course, was obtained when both KA and KE were allowed to float in combination with the prescribed increase in ST (Fig. 10 F). This ideal fit corresponded to a twofold increase in KE (in strong agreement with that of Fig. 10 D) coupled with a reduction in KA by ∼16% (corroborating the result in Fig. 10 E). Thus, we concluded from this analysis that increased susceptibility after ionomycin treatment has little or nothing to do with the strength of adsorption (KA) and can be explained entirely by increases in the accessibility of lipids to the enzyme active site (KE and ST).

Analysis of hydrolysis results in terms of Scheme 1. Initial sPLA2 hydrolysis rates from Fig. 2 A were fit by nonlinear regression to Eq. 2 multiple times with various combinations of constraints. (A) Baseline parameter values were obtained by fitting ...

It is important to note that the differences between control and treatment curves in Fig. 2 A could easily be explained by adjustments to kcat. Variations in kcat could be due to either cofactor availability or enzyme conformational change. Since the same medium was used in all data sets, availability of the cofactor calcium was not an issue. Moreover, if changes in membrane microstructure stabilized the transition to an activated enzyme conformation, that stabilization would be reflected by enhanced adsorption of sPLA2 to the membrane surface. Although multiple conformations of sPLA2 have indeed been observed and may be linked to adsorption and/or substrate binding (49), we have no evidence that adsorption of the enzyme is altered by ionomycin. Therefore, it is unlikely that conformational transitions are affected by ionophore treatment, and our use of a constant value for kcat (i.e., the parameter “β” in Table 1 and Eq. 2) is appropriate.

Parameter values for interaction between sPLA2 and cell membranes

It may seem surprising that the affinity of the enzyme to adsorb to the membrane surface (KA) was not altered by ionomycin treatment (Fig. 4). It is well established that calcium influx promotes exposure of anionic phosphatidylserine on the outer surface of the cell membrane (50,51). In previous studies with ionophore-treated S49 cells, this lipid flip-flop occurred as early as could be detected and corresponded temporally with increased susceptibility to sPLA2 activity (9). Moreover, the presence of anionic lipids greatly promotes sPLA2 adsorption in artificial bilayers and cells (11,5254). Since much of the adsorption of labeled-sPLA2 observed in Fig. 4 appeared unrelated to membrane hydrolysis (for example, binding to heparin sulfate (55)), it could be argued that the methods employed to assess KA were incapable of discerning relevant changes. For this reason, we applied the kinetic analysis detailed in Fig. 10, which substantiated the idea that differential membrane susceptibility is not determined by variation in enzyme adsorption. Furthermore, experiments in human erythrocytes with an inhibitor of phosphatidylserine translocation supported the conclusion that exposure of anionic lipids was not a requirement for enhanced hydrolysis upon ionomycin treatment (9,18). This view is supported by studies indicating that apoptotic stimuli other than ionomycin induced sPLA2 susceptibility before phosphatidylserine exposure in S49 cells (9). Overall, these findings suggest that transbilayer movement of specific phospholipids does not necessarily provide a preferred substrate for the enzyme, nor is flip-flop the primary determinant of membrane vulnerability to sPLA2. Nonetheless, loss of membrane asymmetry could possibly alter other properties of the phospholipid bilayer, which then render the membrane sensitive to sPLA2 activity. For example, exposure of phosphatidylserine could contribute to the increased lipid spacing we observed (56).

A caveat to the conclusion that increased membrane susceptibility to sPLA2 does not result from enhanced enzyme adsorption and/or flip-flop of phosphatidylserine is the fact that the experiments reported here were conducted with enzyme purified from snake venom. Several isoforms of sPLA2 have been identified which differ in their intrinsic catalytic activity (kcat), their affinity for membrane surfaces, and their dependence on the presence of anionic lipids (57). Thus, the degree to which phosphatidylserine exposure and potential differences in KA contribute to the level of membrane hydrolysis under different experimental or physiological conditions may well depend on the isoform in question. Nevertheless, in previous experiments, human groups IIa and V sPLA2 species displayed behavior toward S49 cells that was qualitatively identical to that observed with the snake venom enzyme (27). Therefore, although quantitative differences exist among various sPLA2 types, the concept that substrate availability contributes, at least in part, to the level of membrane susceptibility probably applies to all of them.

Structural evidence suggests that sPLA2 acts at the membrane surface with its active site displaced physically from the position normally occupied by bilayer phospholipids (24). If this is true, substrate availability (ST) represents the number of lipids capable of making a vertical movement from their normal position, which, according to the results (Fig. 2 B), increases during a sustained calcium influx. Moreover, the larger value of KE required to fit the data indicates that this vertical transition also occurs more easily.

We focused our analysis primarily on the initial rate of hydrolysis to avoid concerns about latent hydrolysis of internal bilayers such as the perinuclear membrane (36,37). Of course, measurement of total hydrolysis would include any internal hydrolysis product generated during the 10-min time frame of the experiment (see Fig. 2 B), especially with ionomycin treatment, which promoted internalization of L-PLA2 (see Fig. 3 B). Therefore, it may be unreasonable to include the full measured value of ST in the analysis with Eq. 2. The observation that the majority of total hydrolysis to be accomplished is complete within ∼50 s (Fig. 1) mitigates this concern since this timescale is shorter than that reported for internalization (36,37). Based on the linkage of KE and ST in Eq. 2, overestimation of ST corresponds to underestimation of KE; thus, changes in KE may be greater than we assumed in the analysis shown in Fig. 10 and Table 1. Regardless of the relative contributions of those two parameters to the effect of ionomycin on the initial rate, the argument that the membrane properties have changed to facilitate availability of substrate (either kinetically, KE, or physically, ST) to the active site of the enzyme is still valid. Obviously, the mechanistic basis for increased ST in the presence of ionomycin requires additional investigation.

What could account for the increase in substrate access to the enzyme? The fluorescence of three of the five membrane probes we examined changed reproducibly due to ionophore. For example, MC540 bound more easily to the membrane, and laurdan emission was red-shifted, consistent with increased penetration of water molecules into the bilayer. These two observations suggest that the membrane surface had become more accommodating to intercalation of contaminating molecules from the aqueous phase. This result, together with an enhanced rate of NBD-PC extraction from the outer leaflet of the cell membrane, implies that the strength of interaction among neighboring phospholipids is diminished by calcium influx. With the loss of favorable interactions coupled with increased hydration of the bilayer, the hydrophobic active site of the enzyme would presumably become a more attractive alternative to the membrane, thus enhancing the probability of hydrolysis.

Another possible explanation for the laurdan fluorescence relates to phospholipid chain dynamics. Many studies have suggested that changes in solvent access to laurdan correlate with the level of phospholipid chain order (32,33,5860). The experiments conducted with bis-pyrene and DPH allowed us to examine whether such was the case here, since both probes are sensitive to the order of their environment (44). Interestingly, neither probe detected any change associated with ionomycin treatment. Therefore, the average level of phospholipid chain order in the membrane appeared to remain stable during calcium influx, and our interpretation of laurdan fluorescence relating to hydration changes at the membrane surface appears more plausible.

Initially, it may seem counterintuitive to propose that disruptive alterations have occurred that can be detected readily at the membrane surface without removing constraints on probe motion deep in the bilayer. One possibility is that ionophore treatment has generated local regions of high curvature stress in the membrane causing an increase in headgroup spacing simultaneous with crowding of the chains. Since phospholipid micelles are excellent substrates for sPLA2 (61), such local curvature could explain the various observations in this report. We also note that changes in lipid spacing can occur without accompanying alterations in lipid order and/or lipid fluidity within liquid-ordered phases in both erythrocytes and artificial membranes (44).

This result (alterations at the surface with no change in chain order) contrasts observations from erythrocytes in which treatment with ionomycin actually increased the average lipid order of the membrane (1820). Two distinctions between erythrocytes and lymphocytes are helpful in understanding this difference. First, susceptibility to hydrolysis upon ionophore treatment is about six times greater in lymphocytes than in erythrocytes (compare data in Fig. 2 to Smith et al. (18)). Logically, one would expect that membrane changes relevant to hydrolysis would also be more prominent in S49 cells. Changes observed in laurdan GP and MC540 fluorescence intensity were enhanced by factors of ∼10 (in the opposite direction) and 70, respectively, fulfilling this criterion (compare data in this report with raw data used in Best et al. (20) and Jensen et al. (21)). Second, microscopic images of erythrocytes revealed that ionomycin-induced alterations in lipid order were confined to specific membrane domains (18,20). In general, membrane domains in nucleated cells are much smaller than those of erythrocytes (62), making it unlikely that similar observations of increased order in discrete domains would be possible in lymphocytes, especially against the background of a more pronounced membrane hydration described above. Nevertheless, it is important to note that the data described for erythrocytes led to the same conclusion as that inferred here: that increased susceptibility to enzyme hydrolysis is precipitated by a reduction in the strength of lipid-neighbor interactions.

A major remaining question is the means by which ionophore treatment produces membrane changes responsible for elevated sPLA2 activity. Resolution of this matter will require genetic and biochemical approaches beyond the scope of this study. Nevertheless, possible candidates may be identified from existing reports. The fact that cells rapidly become susceptible to sPLA2 (i.e., within 10 min) during ionomycin-induced apoptosis helps evaluate the possibilities and exclude those events that arise too late (9). One obvious candidate for linking known apoptotic events with membrane sensitivity to sPLA2 is the exposure of phosphatidylserine. As discussed above, this event appears unable to explain the increase in membrane hydrolysis, at least for the snake venom enzyme. Membrane blebbing and loss of integrity can also be eliminated as major contributing factors, because these events occur much later than the onset of susceptibility (9). The images in Figs. 7 and and99 corroborate the observation with respect to blebbing. The sustained loss of membrane potential associated with ionophore- and hormone-induced apoptosis in lymphocytes also appears to occur on a timescale well beyond that relevant to the phospholipase (3,5).

Ceramide accumulation, through de novo synthesis and cleavage of sphingomyelin, is well established as a participant in apoptotic signaling pathways (2). Furthermore, it has been suggested that sphingomyelin inhibits sPLA2, and that its removal could therefore promote increased activity of the enzyme (63,64). The details of ceramide's role in apoptosis vary among cell types (2) and have not yet been characterized in S49 cells. Nonetheless, in prior experiments, treatment of S49 cells with sphingomyelinase did not enhance sPLA2 activity (27).

In addition to blebbing, apoptotic cells sometimes shed smaller pieces of the plasma membrane known as microvesicles. Such is the case with ionomycin-stimulated S49 cells (27). This microvesiculation appears to be facilitated by both phosphatidylserine exposure and cytoskeletal cleavage, and the vesicles released are susceptible to the enzyme (18,63,6568). These observations raise the question of whether the increase in membrane susceptibility reflects direct hydrolysis of the plasma membrane or, instead, degradation of released microvesicles. This issue has been addressed extensively in studies of human erythrocytes and S49 cells. In both cases, the conclusion was reached that although microvesicles are attacked by the enzyme, they represent a minor fraction of the total hydrolysis, with the remainder occurring in membranes associated with the cells (18,27).

Cytoskeletal alterations are more likely candidates for apoptotic events that might alter the membrane's response to sPLA2 (68). Given the sensitivity of various membrane properties to cytoskeletal structure during apoptosis (6), it is reasonable to speculate that the same may be true for the response to sPLA2. For example, membrane tension could be highly important in determining the access of lipids to the enzyme active site (22) and certainly would be affected by the arrangement and number of cytoskeletal attachments. Likewise, cytoskeletal elements may regulate the lateral distribution of membrane lipids. Disruption of domain structure upon cytoskeletal rearrangement could free additional substrate for catalysis that previously was inaccessible to the enzyme (thus, the increase in ST). We are now initiating genetic, pharmacological, and histological studies to explore these interesting possibilities as well as the relationship to other membrane events discussed here.

The long-term goal of this line of research is an understanding of the relationship between cell membrane structure and the differential activity of sPLA2 in physiological settings. To that end, this report applies the model of sPLA2 action developed by multiple researchers (see Scheme 1) to nucleated cells to test hypotheses derived from erythrocyte studies. Erythrocytes were employed in those prior studies because they offer a simplified, quasicellular model devoid of certain experimental complications, such as probes binding to intracellular membranes. Fortunately, we were able to address these complications here using flow cytometry and two-photon microscopy to assist in the interpretation of the data (see Results). The move to lymphocytes was a meaningful step toward a biologically relevant system in which susceptibility to sPLA2 occurs during physiological processes such as apoptosis. Most important, we have demonstrated means by which biophysical techniques and information regarding membrane-protein interactions can be applied to studies of nucleated cells. The next step will be to resolve the timing of the membrane changes described in this report with respect to both the level of sensitivity to sPLA2 and other events during hormone-stimulated apoptosis.


We gratefully acknowledge technical assistance from Summer King, Aubrielle Williamsen, Anne Heiner, Thaothanh Nguyen, Jennifer Nelson, and Chisako McLemore. Two-photon images were acquired at the Laboratory for Fluorescence Dynamics at the University of California, Irvine. We express thanks to the LFD staff, especially Susana Sanchez, Theodore Hazlett, and the director of the facility, Enrico Gratton, for their assistance. Flow cytometry experiments were conducted with the assistance of Sandra Burnett and Nels Nielson at the Research Instrumentation Core Facility at Brigham Young University.

This work was supported by the National Institutes of Health (GM073997).


Editor: Antoinette Killian.


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