Volume regulation in leukocytes: Requirement for an intact cytoskeleton

JOURNAL OF CELLULAR PHYSIOLOGY 163:9&104 (1995)

Volume Regulation in Leukocytes: Requirement for an Intact Cytoskeleton

GREGORY P. DOWNEY,* SERGIO GRINSTEIN, ANDREA SUE-A-QUAN, BARBARA CZABAN, AND CHI KIN CHAN

Departments of Medicine (G.P.D., A.5.-A,-Q., C.K.C.) and Anatomy and Cell Biology (B.C.), The University of Toronto, Toronto, Ontario, Cdnada, MSS 7A8, The Respiratory

Division, the Toronto Hospital, and The Department of Cell Biology (S.G.), Research Institute, Hospital for Sick Children, Toronto, Ontario, Canada.

Animal cells regulate their volume by controlling the flux of ions across their plasma membrane. Recent evidence suggests that ion channels and pumps are physically associated with, and may be regulated by components of the cytoskeleton. To elucidate the role of elements of the cytoskeleton in volume regulation, we studied the effects of cytoskeletal disrupting agents on regulatory volume decrease (RVD) in three different leukocyte types: Jurkat lymphoma cells, HL-60 cells, and human peripheral blood neutrophils. Cell volume was measured in two ways: (i) electronically with a Coulter counter and (ii) by forward light scattering in a flow cytometer. Exposure of all leukocyte types to hypotonic medium (200 mOsm) resulted in an immediate increase in cell volume followed by a regulatory decrease to baseline by 20 min. In the presence of the microtubule disrupting agents, colchicine and nocodazole, RVD was totally inhibited which corre- sponded to loss of microtubules as determined by immunofluorescence. Simi- larly, RVD was inhibited in Jurkat cells incubated with the actin binding agents, cytochalasin B (CB) or D (CD). In contrast, in HL-60 cells and human neutrophils, RVD was unaffected by treatment with either CB or CD. While cytochalasins are generally thought of as microfilament disrupting agents, their primary action is to prevent F-actin polymerization. The extent of ensuing microfilament disruption depends in part on the rate of filament turnover. In an attempt to understand the differential effects of the cytochalasins on RVD, the F-actin content of the different cells was determined by NBD-phallacidin staining and flow cytometry. Pretreat- ment with CB or CD resulted in profound actin disassembly in Jurkat cells (relative fluorescence index RFI: 1 .O control vs. 0.21 * 0.01 for CB and 0.48 * 0.02 for CD). However, the cytochalasins did not induce net disassembly in either HL-60 cells or human neutrophils. To study the effects of an increase in F-actin on volume regulation, neutrophils were treated with the chemoattractant f-Met-Leu- Phe or with an antibody (Ab) to p2 integrins followed by a cross-linking secondary Ab. Despite an increase in F-actin in both circumstances, RVD remained intact. Taken together, these results suggest that both microtubules and microfilaments are important in volume regulation. o 1905 Wiley-Liss, Inc

The regulation of cell volume is a physiological process of fundamental importance to most cell types. In multicellular organisms, anisotonic stresses may be en- countered, for example, during transit of a blood cell through the hypertonic environment of the mammalian kidney (Macy , 1984). Although less widely appreciated, alterations in cell volume occur as part of and may influence the process of activation of leukocytes such as lymphocytes (Grinstein et al., 1985) and neutrophils (Grinstein et al., 1986). Cell volume regulation depends on the tightly controlled flux of ions across the plasma membrane (reviewed by Hoffman and Simonson, 1989). Upon exposure to a hypotonic environment, water flows passively into the cell, resulting in increased volume. In leukocytes, this change in volume initiates a series of regulatory events leading to the loss of intracellular solutes accompanied by osmotically obliged 0 1995 WILEY-LISS, INC.

water, resulting in cell shrinkage. This return towards resting cell volume is termed regulatory volume decrease (RVD). Conversely, when cells are exposed to a hypertonic environment, they initially shrink due to loss of water. This is followed by uptake of solutes from the environment along with osmotically obliged water, inducing a return towards resting volume, a process termed regulatory volume increase (RVI). The transport mechanisms responsible for these volume regula-

Received December 7,1993; accepted September 9,1994. *To whom reprint requestsicorrespondence should be addressed at Clinical Sciences Division, Room 6264 Medical Sciences Building, 1 Kings College Circle, University of Toronto, Toronto, Ontario, Canada, M5S 1A8.

97 ROLE OF THE CYTOSKELETON IN VOLUME REGULATION

tory processes demonstrate great diversity among different cells and organisms. In leukocytes, including lymphocytes (Grinstein et al., 1984) and HL-60 cells (Hallows et al., 19911, RVD is mediated by efflux of K' and C1- through distinct membrane channels. RVI is mediated by activation of the Na+/Hf antiporter that, in concert with activation of the Cl-/HCO,- exchanger, results in net uptake of NaCl (reviewed by Chamberlin and Strange, 1989).

There is evidence that both the sensor and/or effector (i.e., ion channels, exchangers and pumps) mechanisms involved in volume regulation may be influenced by components of the cytoskeleton (reviewed by Mills, 1987). For example, it has been hypothesized that alterations in cell volume are sensed by deformation of the cytoskeletal network and/or that this information may be relayed to the effectors by components of the cytoskeleton (reviewed by Sachs, 1989; Rugolo et al., 1989; Morris, 1990; Cornet et al., 1993a). Ion channels (Christensen and Hoffmann, 1992; Smith et al., 1991; Cantiello et al., 1991; Cornet et al., 1993b) and pumps (Matthews et al., 1992; Grinstein et al., 1993) are closely associated with cytoskeletal elements and their activity may be regulated by this association. However, seemingly discrepant reports exist regarding the effi- cacy of cytoskeletal disrupting agents in the impair- ment of volume regulation. For example, the microfilament disrupting agent cytochalasin B was reported to inhibit RVD in Necturus gallbladder epithelial cells (Foskett and Spring, 1985), Ehrlich ascites tumor cells (Cornet et al., 1993a), and murine peritoneal macro- phages (Galkin and Khodorov, 1988). Conversely, Hal- lows et al. (1991) report that volume regulation in HL-60 cells is unaffected by dihydrocytochalasin B. Discrepancies also exist regarding the effect of microtubule disrupting agents. For instance, Linshaw et al. (1992) reported that RVD was inhibited in rabbit prox- imal renal tubules by the combination of cytochalasins and colchicine. On the other hand, Cornet et al. (1993a1, working with Ehrlich ascites tumor cells and Mills (1987), using MDCK cells, report that colchicine had no effect on volume regulation.

It is possible that genuine differences in the underlying RVD mechanism are responsible for the observed variability in different cell types. Alternatively, while similar mechanisms may be involved, the differences may be attributable to variable efficiency of the cytoskeletal disrupting agents. In this regard, it is note- worthy that cytochalasins, in low micromolar concentrations, do not disrupt existing microfilaments but

Abbreviations

Krebs Ringer's Phosphate Dextrose buffer KRPD PBS phosphate-buffered saline NBD-

RVD regulatory volume decrease RVI regulatory volume increase CB cytochalasin B CD cytochalasin D DFP diisopropylfluorophosphate DMSO dimethylsulfoxide RFI relative fluorescence index ANOVA analysis of variance

phallacidin 7-nitrobenz-2-oxa-l,3-diazole-phallacidin

instead prevent F-actin polymerization (Cooper, 1987). Thus, the ability of these agents to reduce F-actin content depends on the rate of polymerization/ depolymerization, which can vary greatly among cell types (Cooper, 1987). In this report we utilized three cell types to establish a correlation between the ability of cytochalasins to induce net actin depolymerization and their inhibitory effects on RVD. For the sake of completeness, the effect of microtubule disrupters was also tested.

MATERIALS AND METHODS Materials

Dextran T-500 and Percoll were obtained from Phar- macia Chemicals (Montreal, Quebec, Canada). NaC1, KC1, MgCl,, CaCl,, were obtained from Mallinckrodt Inc., Paris, KY. Dextrose, n-formyl-methionyl-leucyl- phenylalanine (fMLP), DMSO, NP-40, PIPES, and HEPES were obtained from Sigma (St. Louis, MO). NBD-phallacidin was obtained from Molecular Probes (Eugene, OR). Formaldehyde and sodium citrate were obtained from Fisher Scientific and lyso-phosphatidyl- choline (lyso PC) from Avanti Polar Lipids (Pelham, AL). The anti-P2 integrin antibody IB4 was a kind gift from Dr. David Chambers, San Diego Regional Cancer Center (San Diego, CA). Rabbit anti-mouse was obtained from Jackson ImmunoResearch Laboratories (West Grove, PA).

Cell culture HL-60 cells and Jurkat lymphoma cells (American

Type Culture Collection) were grown in RPMI 1640 supplemented with L-glutamine, 10% fetal bovine se- rum (Gibco, Grand Island, NY) and penicillin/ streptomycin (Sigma). Cells were passaged biweekly and maintained at a density of between 2.5 x lo5 and 1 x lo6 cells/ml.

Neutrophil isolation Human neutrophils (>98% pure) were isolated from

citrated whole blood obtained by venipuncture, using dextran sedimentation and discontinuous plasma- Percoll gradients as previously described in detail (Haslett et al., 1985). The separation procedure required 2 hr and the cells were used immediately after isolation for the experiments described. The functional integrity and nonactivated state of neutrophils isolated in this manner has been extensively validated in previous publications (Haslett et al., 1985; Downey et al., 1990).

Cell volume measurement Cell volume measurements were performed electron-

ically using a model ZM Coulter counter with a 100 pm diameter orifice interfaced with a Channelizer (Coulter Electronics, Hialeah, FL). The settings were as follows: diameter orifice = 100 pm, current = 1 mA, attenua- tion = 16, and gain = 1. Voltage pulse signals were sorted into channel numbers that were directly proportional to cell volumes. The apparatus was calibrated with polystyrene microspheres (Coulter Electronics). The isoosmotic (295 mOsm) buffer contained (in mM): 140 NaC1, 4 KC1, 1 CaCl,, 1 EGTA, 10 glucose, 10 HEPES, pH 7.4. For measurement of volume changes

DOWNEY ET AL. 98

in response to hypotonic medium, at time = 0 the medium (295 mOsm) containing cells was diluted 33% with distilled (MilliQ, Millipore Canada Ltd., Missis- sauga, Ontario, Canada) water such that the final os- molarity was 197 mOsm. At each time point (defined as the time at which data acquisition was started) cells in the diluted medium were aspirated into the Coulter counter and the mean channel number of the cell distri- bution estimated. The data are presented as the percent of starting volume (V/Vo), as a function of time. For experiments with cytoskeletal disrupting agents, cells were preincubated with the various compounds for 10 min a t 37°C. The water used for dilution contained equimolar concentrations of the drugs so that their concentration remained constant after hypotonicity was induced.

Measurement of forward angle light scatter by flow cytometry was used as a second and independent method of measurement of cell volume (Mullaney and Dean, 1970). The protocol for flow cytometric measurements was essentially as described for the Coulter counter except that cells were aspirated directly into the flow cell of a FacsScan (Becton Dickenson, San Jose, CA) and forward light scatter recorded at each time point indicated. Five thousand cells were collected at each time point and the mean determined using Ly- sis 2 software provided by the manufacturer. The flow cytometer was calibrated prior to each experiment using FluoresBrite beads (calibration grade, Polysciences Inc., Warrington, PA). The data are presented as the percent of starting volume (VNo), as a function of time.

Measurement of F-actin content Neutrophil content of polymerized (F)-actin was de-

termined by NBD-phallacidin staining by the two step method of Howard and Meyer (1984) a s previously reported (Downey et al., 1992). In brief, cells were fixed and permeabilized in one step using a final concentration of 3.7% formaldehyde and 100 pgiml of lysophos- phatidyl choline. Subsequently, the cells were stained with NBD-phallacidin and fluorescence was quantified using a flow cytometer (Coulter Epics). This method has been shown to correlate well with biochemical measurements of F-actin (Howard and Meyer, 1984; Wal- lace et al., 1984; Howard and Wang, 1987). All data are reported as the ratio of the fluorescence intensity of the stimulated cell population to that of the control cells (relative fluorescence index or RFI).

Immunofluorescence detection of microtubules Several different combinations of fixation, extrac-

tion, and staining of cells for microtubules were tried. These included fixation with either 1% paraformaldehyde, 0.3% glutaraldehyde, or 100% methanol; extrac- tion with 100% methanol or either 0.5% NP-40 or 0.5% Triton X-100 in microtubule stabilizing buffer (vide in- fra). The method that gave the best morphology was that modified from Czaban and Forer (1992). Briefly, monolayers of peripheral blood neutrophils were obtained from a few drops of blood obtained by fingerprick from healthy donors deposited on a glass coverslip and allowed to clot in a moist chamber a t 37°C. The clot was gently teased off in warmed Hanks balanced salt solu- tion (HBSS) and the coverslip washed 3 times with HBSS. Cells on the coverslip were preincubated with 10

pM colchicine or 0.1% DMSO (vehicle control) in HBSS for up to 20 min at 37°C and washed twice with microtubule stabilizing buffer (in mM: 5 MgSO,, 10 EGTA, 100 PIPES pH 6.9) a t 4°C. Cells were permeabilized by incubation with microtubule stabilizing buffer containing 0.5% NP-40,5% DMSO, 1 mM PMSF, 1 mM aproti- nin, and 2 mM diisopropylfluorophosphate (DFP) for 90 sec a t 4°C followed by fixation with 1.5% paraformaldehyde for 10 min a t room temperature. Following 2 washes with microtubule stabilizing buffer, cells were incubated with a 1:500 dilution of rat monoclonal anti- tubulin antibody (YOL 1/34, Serotec, Toronto, Canada) for 2 hr a t 4°C in PBS containing 1% BSA, washed three times with PBS, incubated with a 1: 30 dilution of FITC-labelled goat anti-rat antibody (Mandel Scien- tific, Guelph, Ontario, Canada) in PBS containing 1% BSA, washed three times with PBS and mounted using Immumount (Shandon, Pittsburgh, PA). The samples were viewed using a BioRad 600 laser scanning confocal imaging system mounted on a Leitz Metallux-3 microscope using a x 100 (1.32 NA) oil-immersion objec- tive (Leitz). Light for fluorescence stimulation was provided by an air-cooled krypton-argon-ion laser (Bio- Rad). The fluorescent light intensity was detected by a photomultiplier after separation by a 560 nm dichroic barrier. The location of the ventral cell surface (that attached to the substratum) was defined by serial optical sectioning using a computer-controlled variable step-size stage. Optical sections (0.5 km) of the ventral surface were obtained. The confocal aperture was set a t a position between 25% and 35% of its full adjustable range to optimize the efficiency of light collection while maintaining submicrometer depth of field with optical sections. The image size was 768 x 512 pixels with an 8-bit grey level resolution (256 grey levels). Images were acquired using Kallman averaging and photographed on Kodak PlusX film (ASA 125) with a Nikon F-301 camera attached to a high resolution video monitor. Photomicrographs are representative of multiple cells observed on each coverslip from a t least three sep- arate cell preparations done on different days.

RESULTS AND DISCUSSION The magnitude and time course of alterations in cell

volume upon exposure to a hypotonic buffer were studied in freshly isolated human neutrophils and in two human leukocyte cell lines: undifferentiated HL-60 cells and the Jurkat lymphoma cell line. Exposure of all three leukocyte types to hypotonic medium (197 mOsm) resulted in an immediate increase in cell size followed by a decrease towards the original volume (Fig. 1). The latter response is termed regulatory volume decrease (RVD). The magnitude of cell swelling in response to the hypotonic stress varied between cell types: volume increased to 130 5 5.2% of isotonic values in neutrophils, 118 2 4.2% in Jurkat cells and 115 t 6.7% in HL-60 cells. RVD was essentially complete in all cell types by 25 min.

In order to determine the importance of an intact cytoskeleton for volume regulation, the effects of cytoskeletal disrupting agents were studied. We chose initially to study the effects of colchicine and nocodazole, agents known to disrupt microtubules (Bershad- sky and Vasiliev, 1988). Incubation of Jurkat cells, HL-60 cells (data not shown) and peripheral blood neu-

ROLE OF THE CYTOSKELETON IN VOLUME REGULATION 99

A. Neutrophils

Control -t- 1 pM Colchicine + 10 pM Colchicine - 10 pM Lumicolchlcine

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Fig. 1. Colchicine inhibits regulatory volume decrease in Jurkat lymphoma cells and human neutrophils. Cell volume was measured electronically using a Coulter counter as described in Materials and Methods. A Change in cell volume of human neutrophils exposed to 66.6% tonicity. B Change in cell volume of Jurkat lymphoma cells exposed to 66.6% tonicity. Data are expressed as the percent change in volume relative to control cells incubated in a comparable concentration of DMSO (0.1%). Eachvalue represents the mean t S.E.M. of four determinations. Asterisks indicate P < 0.05 with respect to the control, as determined by analysis of variance for repeated measures with correction for multiple comparisons by the method of Scheffe.

trophils with these agents resulted in nearly complete inhibition of RVD (Figs. 1 and 2). Lumicolchicine, an inactive derivative of colchicine, was not inhibitory (Fig. l), confirming the specificity of action of the microtubule disrupting agents. To ensure that these conditions in fact resulted in disruption of microtubules, these structures were visualized by immunofluorescence. Intact microtubules and microtubule organizing centers could be identified in >90% of control cells (Fig. 3a). Treatment with colchicine resulted in disruption of these structures in >SO% of cells observed. Frequently (3040% of cells), an area of collapsed tubulin was apparent in the juxtanuclear region (Fib. 3b). Similar results were obtained with HL-60 and Jurkat cells (data not shown).

We next studied the effects of cytochalasins, fungal toxins known to preclude association of G-actin with the barbed end of actin filaments (Cooper, 1987). In neutrophils and HL-60 cells, RVD was unaffected by treatment with either 5 pM cytochalasin B (CB) or 1 pM cytochalasin D (CD; Fig. 4A and B). In contrast,

6. Jurkat Cells

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Fig. 2. Nocodazole inhibits regulatory volume decrease in Jurkat lymphoma cells and human neutrophils. Cell volume was measured electronically using a Coulter counter as described in Materials and Methods. A: Change in cell volume of human neutrophils exposed to 66.6% tonicity. B: Change in cell volume of Jurkat lymphoma cells exposed to 66.6% tonicity. Data are expressed as the percent change in volume relative to control cells incubated in a comparable concentration of DMSO (0.1%). Each value represents the mean * S.E.M. of four determinations. Asterisks indicate P < 0.05 with respect to the control, as determined by analysis of variance for repeated measures with correction for multiple comparisons by the method of Scheffe.

while Jurkat cells treated with either CB or CD demonstrated the expected increase in volume in response to hypotonic medium, RVD was totally inhibited (Fig. 4C). Two possible explanations for the lack of effect of cytochalasins in neutrophils and HL60 cells, in which RVD remained intact, and in Jurkat cells in which RVD was inhibited, were that either the cytosolic concentration achieved was lower in neutrophils and HL60 cells than in Jurkat cells or that somehow the F-actin in the former was more resistant to the cytochalasin effects. To investigate these possibilities, higher concentrations of cytochalasin B (25 pM: data not shown) and dihydrocytochalasin B (25 pM) were studied. As illustrated in Figure 4A, even this higher concentration did not prevent RVD in neutrophils. Similar results were obtained in HL60 cells (not illustrated).

To elucidate the reasons for the differential susceptibility to cytochalasin-induced inhibition of RVD in Jur- kat lymphoma cells on one hand and HL-60 cells and neutrophils on the other, the effect of cytochalasins on the actin cytoskeleton of these cell types was compared.

100 DOWNEY ET AL

Fig. 3. Colchicine disrupts the microtubule network of human neutrophils. Monolayers of peripheral blood neutrophils were obtained from blood obtained by fingerprick as described in Materials and Methods. Cells on the coverslip were preincubated with 10 KM colchicine or 0.1% DMSO (vehicle control), permeabilized and fixed with 1.5% paraformaldehyde. Microtubules were visualized by immunofluorescence using a rat monoclonal anti-tubulin antibody and a FITC-labelled goat anti-rat secondary antibody. The samples were viewed using a BioRad 600 laser scanning confocal imaging system mounted on a Leitz Metallux microscope. Images were acquired using Kallman averaging and photographed with a Nikon F-301 camera at-

The F-actin content of each cell type was measured by NBD-phallacidin staining and flow cytometry (Howard and Meyer, 1984). Figure 5 illustrates that both CB and CD induced significant F-actin disassembly in Jurkat cells (relative fluorescence index RFI: 1.0 control vs. 0.21 2 0.01 for CB and 0.48 +- 0.02 for CD). In marked contrast, neither CB, CD nor dihydrocytochalasin B produced net F-actin disassembly in either HL-60 cells or peripheral blood neutrophils. Thus, the extent of actin disassembly appeared to correlate with the inhibition of RVD, i.e., inhibition of RVD was apparent only in Jurkat cells where net disassembly of F-actin could be induced. The failure of cytochalasins to induce actin disassembly in neutrophils could not be explained on the basis of a lower affinity of the toxin for the barbed ends of the microfilaments in these cells because at the concentrations used, actin assembly induced by the chemotactic peptide fMLP was completely abrogated (Fig. 5). In additional experiments, we attempted to cause net actin depolymerization in neutrophils and HL-60 cells by the induction of net actin turnover (“cy- cling”) by more prolonged (20 min) exposure of cytochalasin-treated cells to chemotactic agents known to result in actin turnover. This combination of treatments might be expected to allow depolymerization to occur under conditions in which polymerization was blocked resulting in net actin disassembly. However, in neither cell type was a net decrease in F-actin observed under these conditions (Fig. 5).

In the previous experiments, changes in cell volume were measured electronically using the Coulter counter. To ensure the validity of this method and to exclude the possibility that the inability of cytochalasin-treated Jurkat cells to regulate cell volume was not

tached to a high resolution video monitor. a: Optical section (0.5 km) of a control neutrophil illustrating microtubules and the microtubule organizing center. This section is 1.5 pm above the midplane of the cell to maximize the visualization of the microtubule network. The solid arrow indicates the microtubule organizing center. b Optical section (0.5 pm) of a neutrophil that has been incubated with 10 pM colchicine for 20 min at 37°C prior to fixation and staining. This section is through the midplane of the cell. The open arrow illustrates an area of apparently depolymerized tubulin. The negatively stained nuclear lobes are also visible in this section. The bar indicates a distance of 5 pm.

an artifact of this method of measurement, we studied RVD using measurement of forward light scatter by flow cytometry. This method of measurement, under certain conditions, is proportional to the cross-sectional area of the cell (Mullaney and Dean, 1970). Figure 6 illustrates that using measurement of forward light scatter, regulatory volume decrease was intact in neutrophils but absent in Jurkat cells treated with cytochalasin D.

Taken together, the data described above are compat- ible with the notion that a decrease in the amount of F-actin interferes with volume regulation in hypoosmotic media. To explore the effects of an increase in the amount of F-actin on volume regulation, two strategies were employed. I t is known that mechanical stresses on the outside of a cell can be transmitted to the cytoskeleton via interactions of membrane proteins such as adhesion molecules (Wang et al., 1993). Mature human neutrophils express p2 integrins of the CD11ICD18 family and recent studies have demonstrated that spa- tial clustering (“cross-linking”) of these adhesion molecules results in changes in the actin cytoskeleton, in particular actin polymerization (Lofgren et al., 1993). To determine if integrin cross-linking might induce changes in cell volume or modify the ability of the cell to regulate volume in response to an anisotonic stress, RVD was measured in cells after such cross-linking. Incubation of cells with primary anti-CD18 antibody (IB4) followed by cross-linking with rabbit anti-mouse secondary antibody resulted in actin polymerization (Fig. 7) that peaked a t 1 min and was sustained for a t least 10 min. However, p2 integrin cross-linking did not result in any change in cell volume nor did it impair RVD after exposure to hypotonic medium (Fig. 7B). As

ROLE OF THE CYTOSKELETON IN VOLUME REGULATION 101

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Fig. 4. Effects of cytochalasins on regulatory volume decrease in human neutrophils, HL-60 cells and Jurkat lymphoma cells. Cell volume was measured electronically in response to exposure to 66.6% tonicity using a Coulter counter as described in Materials and Meth- ods. A Change in cell volume of human neutrophils pretreated with 5 pM cytochalasin B, 1 p M cytochalasin D, or 25 pM dihydrocytochalasin B for 10 min prior to measurement of RVD. B: Change in cell volume of HL-60 cells pretreated with 5 pM cytochalasin B and or 1 pM cytochalasin D for 10 min prior to measurement of RVD. C:

a second method of induction of increases in F-actin, cells were exposed to fMLP, an alternate stimulus known to induce net actin assembly (Fig. 7A). This strategy is complicated by the fact that exposure to fMLP induces a 20% increase in cell volume by itself (Grinstein et al., 1986). However, the increase in cell volume has reached a peak by 5 min and this volume is sustained for a t least 15 min. Therefore we chose to study the effect of exposure to hypoosmotic media in cells pretreated with fMLP for 5 min. Under these conditions, the magnitude of the volume increase on subsequent exposure to hypotonic medium was slightly en- hanced (as compared to cells that had never been exposed to fMLP) but the subsequent decrease in volume towards resting values was comparable between treated and untreated cells.

Taken together, these data strongly support the notion that functional integrity of both the microtubule and microfilament components of the cytoskeleton is

Change in cell volume of Jurkat lymphoma cells pretreated with 5 pM cytochalasin B and or 1 pM cytochalasin D for 10 min prior to measurement of RVD. Data are expressed as the percent change in volume relative to control cells incubated in a comparable concentration of DMSO (0.1%). Each value represents the mean f S.E.M. offour determinations. Asterisks indicate P < 0.05 with respect to the control, as determined by analysis of variance for repeated measures with correction for multiple comparisons by the method of Scheffe.

essential for effective volume regulation (RVD) after a hypotonic stress in leukocytes. The evidence for the former is that treatment with either nocodazole or colchicine (but not the inactive analog lumicolchicine) impaired RVD in all three leukocyte types (Figs. 1 and 2). The evidence for microfilament involvement is that cytochalasins B and D prevented RVD in Jurkat lymphocytes, an effect that correlated with net disassembly of F-actin as measured by NBD-phallacidin staining. Additionally, leukocytes appeared to differ in their susceptibility to net disassembly of the actin microfilaments by cytochalasins. In particular, neutrophils and HL-60 cells were relatively resistant to the net disassembly of actin microfilaments by cytochalasin B, cytochalasin D, or dihydrocytochalasin B. Regardless of the underlying mechanisms involved, the resistance to microfilament disassembly by the cytochalasins in these two cell types correlated with the inability of the cytochalasin to interfere with RVD. We believe that

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DOWNEY ET AL.

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Fig, 5. Effects of cytochalasins on the F-actin content of human neutrophils, HL-60 cells, and Jurkat lymphoma cells. F-actin content was measured by NBD-phallacidin staining after the treatments as speci- fied. Neutrophils, HL60 cells, and Jurkat cells were incubated with either 5 pM cytochalasin B (cytoB) or 1 pM cytochalasin D (cytoD) for 10 min a t 37°C prior to fixation and staining. Human neutrophils were stimulated with 10-*M fMLP for 1 min (fMLP 10 nM) or pretreated with with 1 pM cytochalasin D for 10 min prior to stimulation with 10-*M fMLP for an additional 1 min (CytoDifMLP 10 nM). Human

this difference in susceptibility to the effects of cytochalasins may explain, a t least in part, the discrepant reports regarding the effects of these agents on volume regulatory processes.

Our results differ slightly from a recent report by Hallows et al. (1991). Specifically, although both studies report that cytochalasins do not interfere with RVD in HL-60 cells, Hallows et al. (1991) found that treatment with dihydrocytochalasin B resulted in net actin disassembly. In our current report, none of the cytochalasins (cytochalasin B, cytochalasin D, or dihydrocytochalasin B) were able to induce net actin disassembly in HL-60 cells or in neutrophils (Fig. 4). As discussed by Cooper (19871, the effects of cytochalasins on actin are complex and are dependent on the dose and type of cytochalasin studied. At low concentrations (equiva- lent to 0.2 pM cytochalasin D), cytochalasins inhibit membrane ruffling by inhibiting growth (or perhaps promoting disassembly) at the barbed ends of microfilaments (Yahara et al., 1982; Cooper, 1987). At higher concentrations (= 10-20 p M cytochalasin D), loss of stress fibers is observed in certain cell types (Yahara et al., 1982) implying that a t these concentrations, cytochalasins might bind to G-actin and, by sequestration of monomers, promote net microfilament disassembly. An alternate but unproved possibility is that cytochalasin actually sever filaments (Hartwig and Stossel, 1979; Cooper, 1987). One potential explanation for the differences between our results and those of Hallows et al. (1991) is the difference in methods of fixation. Our method involving fixation, permeabilization and staining for F-actin differs from that used by Hallows et al.

.. 1 .o 1.5 3.0

RFI

neutrophils and HL60 cells were also pretreated with 25 pM dihydrocytochalasin B for 20 rnin prior to stimulation with lO-*M fMLP for an additional 1 min (DCBlfMLP 10 nM). Data are expressed as the RFI, relative to control neutrophils incubated in a comparable concentration of DMSO (0.1%). Each value represents the mean c S.E.M. of at least four determinations. Asterisks indicate P < 0.05 for the indicated comparisons, as determined by analysis of variance for repeated measures with correction for multiple comparisons by the method of Scheffe.

(1991). Specifically, in our studies, cells were fixed and permeabilized in one step followed by staining with NBD phallacidin. In contrast, in the study by Hallows et al. (1991), cells were first fixed for a more prolonged period (48 h r at 4°C) and then stained without a formal permeabilization step. These small differences may be important since, as described by Watts and Howard (1993), simultaneous fixation and permeabilization may allow the loss of small F-actin oligomers (that are Triton X-100 soluble but still bind NBD phallacidin) while fixation before permeabilization prevents the loss of these oligomers. It is therefore possible that the cytochalasin-induced F-actin disassembly as reported by Hallows et al. (1991) might be confined to this puta- tive subpopulation of short F-actin oligomers which may not be involved in volume regulatory mechanisms.

The precise role of the cytoskeleton in volume regulation remains unknown, but represents an area of in- tense interest (reviewed by Mills, 1987). The cytoskeleton could participate in volume sensing mechanisms or be involved in the effector limb of volume regulatory processes by controlling the activity or the insertion1 retrieval of transport proteins involved in volume regulation. Studies have shown that actin and actin-binding proteins such as spectrin, fodrin, and ankyrin are linked to ion transport proteins including the band 3 anion exchanger (Drenckhahn et al., 1985), the a-sub- unit of the Na'-K'-ATPase (Nelson and Veshmock, 1987), Na+ channels from brain (Srimivasan et al., 19881, and the Na+ channel complex of A6 cells and renal medullary cells (Smith et al., 1991). Moreover, Cantiello et al. (1991) have recently demonstrated that

ROLE OF THE CYTOSKELETON IN VOLUME REGULATION

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140-

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Y 2 120-

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l0OP

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A. F-actin: Neutrophils

Control

1 pJ4 Cytochalasin 0

l o o h 0 5 10 15 20 25

Time (min)

Control p.4 Cytochalasin D

Fig. 6. Changes in cell volume of human neutrophils and Jurkat cells in response to exposure to 66.6% tonicity using forward light scattering on a flow cytometer as described in Materials and Methods. A Change in forward light scatter of human neutrophils pretreated with 0.1% DMSO or 1 pM cytochalasin D for 10 min followed by exposure to 66.6% tonicity. B: Change in forward light scatter of Jurkat lymphoma cells pretreated with 0.1% DMSO or 1 pM cytochalasin D for 10 min followed by exposure to 66.6% tonicity. Data are expressed as the percent change in forward light scatter relative to control cells. Each value represents the mean 2 S.E.M. of four determinations. Asterisks indicate P < 0.05 with respect to the control, as determined by analysis of variance for repeated measures with correction for multiple comparisons by the method of Scheffe.

short actin filaments stimulate the Na+ channel activity of A6 epithelial cells. In an analogous manner, microtubules have also been postulated to be involved in anion transport pathways in murine 5774.1 macro- phages (Melamed et al., 1981) and in regulation of Na.' currents in squid axons (Matsumoto et al., 1984; Murofushi et al., 1983). Clearly, much evidence from diverse sources supports the notion that cytoskeletal elements are involved in the control of ion transport and may participate in the regulation of cell volume.

In summary, the functional integrity of both microtubules and microfilaments, integral components of the cytoskeleton, appears to be required for effective regulatory volume decrease in leukocytes. Differences in the conditions used to disrupt components of the cytoskeleton in a variety of cell types may explain discrepant reports regarding the effects of cytoskeletal disrupting agents on volume regulatory processes. Precise understanding at the molecular level of the mecha-

RFI 2.01 1 ' \

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Fig. 7. Changes in F-actin content and cell volume of human neutrophils: effect of conditions that increase the amount of F-actin on volume regulatory behavior. A: Measurement of F-actin content by NBD- phallacidin staining of fixed and permeabilized cells using flow cytometry. Data are expressed as the relative fluorescence index (RFI) as compared to control. Neutrophils were incubated with 10 kg/ml IB4 (anti-CD18) for 20 min at 4"C, washed and resuspended in fresh KRPD buffer at 37°C followed by the addition of rabbit anti-mouse (RAM) secondary ("cross-1inking":X-link) antibody. Time 0 represents the time of addition of secondary antibody. Alternatively, only the primary antibody (IB4) was added a t time 0. Neutrophils were also treated with 10-'M fMLP at time 0. Each value represents the mean * S.E.M. of at least four determinations. B: Changes in cell volume of human neutrophils in response to the conditions described above ( N L P 10-'M or incubation with IB4 followed by RAM) or in cells treated with either fMLP lO-'M or IB4 and RAM followed by exposure to 66.6% tonicity. Cell volume was measured using the Coulter counter as described in Materials and Methods. Control means that cells were exposed only to hypotonic medium (66% tonicity) but not to either fMLP or IB4 and RAM. Isotonic means that cells were only exposed to the agonist but not to subsequent hypotonicity. For the cells that were pretreated (PreRx) with the fMLP or IB4 and RAM followed by exposure to hypotonicity, the volume is normalized to untreated cells in isotonic buffer. Each value represents the mean f S.E.M. of at least four determinations.

nisms by which cytoskeletal elements participate in volume regulatory processes is still lacking. The fundamental importance of volume regulation mandates fur- ther studies to attempt to elucidate the molecular de- tails of these processes.

ACKNOWLEDGMENTS The authors thank Ms. Sheryl Smith for assistance in

flow cytometry. This work was supported by operating

DOWNEY ET AL. 104

grants from the Medical Research Council of Canada (G.P.D. and S.G.), the Ontario Thoracic Society (G.P.D.), and the National Sanitarium Association (G.P.D.). Dr. Downey is the recipient of a Career Scien- tist Award from the Ontario Ministry of Health. Dr. Grinstein is the recipient of a Howard Hughes Interna- tional Research Scholar Award.

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Volume regulation in leukocytes: Requirement for an intact cytoskeleton