Cell Motility and the Cytoskeleton 45:93–105 (2000) Dynamic Study of Cell Mechanical and Structural Responses to Rapid Changes of Calcium Level Fabienne Richelme, Anne-Marie Benoliel, and Pierre Bongrand* Laboratoire d’Immunologie, INSERM U 387, Hôpital de Sainte-Marguerite, Marseille, France Cell shape control is complex since it may involve multiple cytoskeletal components and metabolic pathways. Here we present a kinetic study of the mechanical and structural responses of cells from the monocytic THP-1 line to a rapid increase of cytosolic calcium level. Cells were exposed to ionomycin in a medium of varying calcium concentration and they were probed at regular intervals for (1) cortical rigidity as determined with micropipette aspiration, and (2) content and distribution of polymerized actin, myosin or ABP-280, as determined with flow cytometry and/or confocal microscopy. An increase of free intracellular calcium level induced: (1) a biphasic deformability change with marked stiffening within a second, and significant softening a minute later; (2) a biphasic change of actin polymerization with initial decrease (within less than a second) and rapid recovery (within a few seconds); (3) a topographical redistribution of microfilaments with an oscillatory behavior of the cortical fraction, while no substantial redistribution of myosin or ABP-280 was detected. It is suggested that a regulation of cell rigidity might be achieved without any structural change by suitable modulation of the lifetime of bridges formed between microfilaments by actin binding proteins. Cell Motil. Cytoskeleton 45:93–105, 2000. r 2000 Wiley-Liss, Inc. Key words: actin; calcium; rheology; kinetic study; monocytic INTRODUCTION Many leukocyte functions are dependent on the cell capacity to deform in response to mechanical forces. Thus, passage through narrow capillary vessels requires the acquisition of an elongated shape [Bagge and Branemark, 1977; Worthen et al., 1989; Doerschuk et al., 1993]. Leukocyte transmigration from blood towards inflamed peripheral tissues involves impressive squeezing between endothelial cells [Marchesi, 1961]. Also, processes such as locomotion or spreading are dependent on both force generation and cell response to the resulting mechanical loads [Adams, 1992]. Further, a cell may need to regulate its deformability. Thus, termination of adhesive interactions may require the decrease of cell stiffness, resulting in the release of a so-called bracing grip [Rees et al., 1977]. It is usually considered that cell mechanical properties are determined by the cytoskeleton, a complex r 2000 Wiley-Liss, Inc. meshwork including three main classes of fibers, i.e., actin microfilaments, microtubules, and intermediate filaments. However, despite remarkable progress in the elucidation of cytoskeletal organization, our understanding of the mechanisms of cell mechanical control remains hampered by at least four major problems: first, a workable description of cell mechanical properties must be achieved. Second, in order to relate these properties to cytoskeletal behaviour, we must determine the mechanical properties of isolated models of the cell cytoskeleton. Third, there is a need to explore the dependence of cytoskeletal organization on parameters related to cell activation, including modification of cytosolic calcium or *Correspondence to: Dr. Pierre Bongrand, Laboratoire d’Immunologie, INSERM U 387, Hôpital de Sainte-Marguerite, BP 29, 13274 Marseille Cedex 09, France. E-mail: email@example.com Received 14 May 1999; accepted 14 October 1999 94 Richelme et al. pH, inositol lipids or phosphorylation/dephosphorylation reactions. Fourth, in view of the interdependence of intracellular biochemical cascades, we must know which properties of the cell interior are modulated by a particular perturbation of a given parameter. It is not an easy task to achieve a quantitative description of cell rheological properties [Mow et al., 1984]. Very diverse methods were devised for this purpose. The deformation of cells subjected to controlled centrifugation was studied [Hiramoto, 1967; Mège et al., 1985; Thoumine et al., 1996]. The displacement of internalized magnetic particles subjected to varying forces was measured [Crick and Hughes, 1950; Valberg and Albertini, 1985]. Magnetic beads were bound to the cell surface through ligand-receptor interaction and subjected to calibrated momenta, for determination of induced rotation [Wang et al., 1993]. Much valuable information was obtained with the cell poker, by measuring the force required for a stylus of micrometer diameter to indent deposited cells [Petersen et al., 1982; Pasternak and Elson, 1985]. Recently, the monitoring of chick fibroblasts adhering to microplates subjected to controlled separation force allowed Thoumine and Ott  to fit cell behavior with a three-element Kelvin viscoelastic model. However, the most widely used method of assaying cell mechanical properties may well consist of monitoring the deformation of individual cells sucked into micropipettes with calibrated pressure [Mitchison and Swann, 1954; Lichtman and Kearney, 1970; SchmidtSchönbein et al., 1981; Evans and Kukan, 1984]. Much information was obtained by varying the pressure and pipette diameter, and observing the kinetics of protrusion formation as well as relaxation after expulsion from the pipette. The problem is that it is difficult to account for all experimental conditions with a single mechanical model. Schmid-Schönbein et al.  achieved satisfactory description of small deformations of blood leukocytes (i.e., protrusion length smaller than 1 µm) with a standard three-element viscoelastic model (with two elastic moduli of 28 and 74 Pa and a viscous component of 13 Pa.s). Large deformations were described by modeling neutrophils as viscous droplets (about 100 Pa.s at 37°C) surrounded by a membrane under tension (about 0.035 mN/m) [Evans and Kukan, 1984; Evans and Yeung, 1989]. Other refinements were suggested such as use of non-Newtonian (deformation-dependent) viscosity [Tsai et al., 1993]. Since most authors studied polymorphonuclear leukocytes, it was probably warranted to neglect the nucleus influence on cell deformation. The second problem consists of identifying the dominant cytoskeletal features that determine cell rheological properties. A comparative study performed on microtubules, microfilaments, and intermediate filaments was consistent with the view that actin microfilaments might be mainly responsible for the resistance of interphasic cells to moderate strain [Janmey et al., 1991]. The measured viscosity of 1 mg/ml F-actin was about 0.6 Pa.s, and it was proportional to the square of the concentration [Janmey et al., 1988]. Since actin concentration may be as high as 20 mg/ml in nonmuscle cells, with more than 50% of molecules in polymerized form [Pollard, 1981; Stossel, 1989], actin alone might account for cell viscosity. Similarly, the elastic modulus of microfilament assemblies is sufficient to account for cell mechanical stability [Janmey et al., 1994]. However, the significance of these conclusions is impaired by the strong dependence of actin mechanical properties on the history of the sample [Xu et al., 1998]. Also, relating microfilament and cell properties is made still more difficult by the existence of a variety of actin binding proteins that may either shorten microfilaments (as was well demonstrated with gelsolin) [Janmey et al., 1988] or cross-link filaments [Hartvig and Shevlin, 1986], resulting in the formation of a gel that might behave as a solid or liquid-like viscoelastic material, depending on the lifetime and force dependence of cross-linking bonds [Sato et al., 1987; Wachsstock et al., 1994]. Third, assuming that the molecular structure of the actin network accounts for cell rheological properties, we need to understand how this structure is regulated. This is quite a difficult challenge: more than 100 actin binding proteins were described, and each of these proteins may perform several separate functions [Theriot and Mitchison, 1993]. Further, the functions of cytoskeletal proteins are regulated by a variety of messengers or control mechanisms including calcium [Yin and Stossel, 1979; Bretscher and Weber, 1980] or hydrogen ions [Sun et al., 1995], cyclic nucleotides [Janmey et al., 1990], phosphorylation/dephosphorylation [Hartwig et al., 1992], or lipids such as phosphatidylinositol [Lassing and Lindberg, 1985]. Also, the activity of a given molecule may be dependent on the balance between several messengers [Aderem, 1992; Lamb et al., 1993]. As a consequence, it is very difficult to predict the effect of a given messenger on the cytoskeleton of intact cells from phenomena observed in model systems. Fourth, exposing a cell to a given stimulus usually results in a cascade of reactions generating many mediators likely to alter cytoskeletal properties [e.g., Snyderman and Pike, 1984; Weisman et al., 1987; Grinstein and Furuya, 1986; Traynor-Kaplan et al., 1988]. Thus, the generation of a single messenger into an intact cell may influence cytoskeletal organization through indirect perturbation of many intracellular parameters. The aim of the present report was first to establish a simple way of measuring the deformability of cells from the human monocytic THP-1 line, then to determine the effect of cytosolic calcium changes on the mechanical Mechanical Cell Response to Calcium Changes 95 properties of whole cells as well as actin organization. In view of the aforementioned difficulties, our strategy consisted of achieving simultaneous control of cytosolic calcium concentration and mechanical properties of intact cells. This included the following points: 1. Real time monitoring of cell mechanical properties was achieved by microscopical observation of the deformation rate of cells sucked into micropipettes with controlled pressure. 2. Preliminary controls suggested that the most significant information was obtained by considering the first few seconds following mechanical stress, which permitted the probing of a welldefined region (i.e., the cortical area), and avoided interpretation problems related to deformationinduced cell activation [Evans and Kukan, 1984; Horoyan et al., 1990]. Also, the potential effect of the cell nucleus was minimized, which may be an important point when cells other than neutrophils are studied. 3. Calcium was manipulated by incubating cells in ionophore-containing solutions of varying calcium concentration. Detergent-mediated cell permeabilization was avoided in order to prevent a possible loss of important regulatory components. 4. Fluorescence labeling of cytoskeletal components was performed to relate cell mechanical alterations to putative changes of microfilament organization. We conclude that calcium rise induces a timedependent decrease of cell mechanical resistance associated with topological redistribution of microfilaments. MATERIALS AND METHODS Cells We used the human monocytic THP-1 cell line [Tsuchiya et al., 1980]. Cells shared some typical properties of mononuclear phagocytes including active phagocytosis of opsonized erythrocytes (not shown) and surface expression of CD11b, CD18, CD32, CD35, CD43, CD45, CD64, and non-polymorphic epitopes of class I histocompatibility molecules, as checked with flow cytometry [Soler et al., 1997] (Sabri et al., unpublished data). Electron microscopical studies recently performed in our laboratory [Soler et al., 1997] revealed that the surface of THP-1 cells was fairly smooth, in contrast with many phagocyte populations. Culture was performed in RPMI 1640 medium (Gibco/Life Technologies, Cergy-Pontoise) supplemented with 20 mM HEPES, 10% fetal calf serum (Biomedia), 2 mM L-glutamine, 50 U/ml penicil- Fig. 1. Experimental study of cell response to mechanical forces. Cells were deposited on the stage of an inverted microscope equipped with a videocamera. The video output was connected to a digitizer mounted on a desk computer. Cells were aspirated into micropipettes connected to a syringe mounted on a syringe holder. Pressure was monitored with a sensor connected to the computer. Pressure and time values were superimposed on live cell images before recording on videotapes for delayed analysis. lin, and 50 µg/ml streptomycin. Incubation was done at 37°C in 5% CO2 atmosphere in 25-ml plastic culture flasks, with 2–3 subcultures a week. Micropipettes Micropipettes were obtained from borosilicate capillary tubes (0.8 mm internal diameter, Clark electromedical instruments, provided by Phymep, Paris) with a programmable micropipette puller (Campden Instruments, model 773) and enlarged to about 5 µm internal diameter with a deFonbrune microforge (Alcatel, Paris). Aspiration Stage Micropipettes were held with a hydraulic micromanipulator (Narishige M0204) and connected to a pressure generator made of a U-tube connected to a 2-ml syringe mounted on a micrometric syringe holder (Fig. 1). The pressure was measured with a sensor (Cole Parmer Instruments, ref. 78300-04) connected to an analog input card (model PC-ADC 12B8V/D, Digimetrix, Perpignan, France) mounted on an IBM compatible desk computer. This allowed real time pressure determination with about 1 Pa resolution. Aspiration was performed on the stage of an Olympus IMT2 inverted microscope equipped with a SIT videocamera (Lhesa, Cergy Pontoise, France, model 4036). We used a 40⫻ dry objective (0.85 numerical aperture), 1.5⫻ magnification facility of the microscope and a 6.7⫻ lens mounted on the camera output (instead of standard 3.3⫻ lens). The output was connected to a 96 Richelme et al. PCVision⫹ digitizer (Imaging Technology, Bedford, MA) mounted on the same computer as the voltage acquisition card. Digitized images were subjected to reverse digitalto-analog conversion and displayed on a monitor with continuous superimposition of time and pressure. A videotape recorder was used for continuous recording and delayed analysis. Experimental Procedure for Deformability Studies Aspiration was performed in saline supplemented with 20 mM HEPES, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 10 mM glucose, and 0.1% bovine albumin. In a typical experiment, a 300 µl aliquot of cell suspension (4 ⫻ 105/ml) was deposited on a glass coverslip (20 ⫻ 60 mm2) set on the microscope stage. A single pipette was used on a given day, in order to optimize the significance of comparisons between different experimental conditions. For each cell, zero pressure was determined by observing the motion of free particles or cells near the pipette mouth, then a negative pressure of 100 Pa (⬃1 cm H2O) was set with the syringe and a nonadherent cell was rapidly aspirated under continuous recording. This pressure was markedly higher than the minimal pressure of ⬃30 Pa required to induce a detectable protrusion of 1 µm length [Richelme et al., 1997]. The pressure was usually reversed 15 sec after aspiration, resulting in rapid expulsion of the cell. Videotapes were subjected to delayed analysis with an image processing software written in our laboratory [André et al., 1990; Sabri et al., 1997]. In most cases, images of the region surrounding the pipette tip were captured with 5/second frequency, and the protrusion length was determined with pixel accuracy (i.e., 0.27 µm). In some cases, the image of a small region surrounding the pipette tip (about 8 ⫻ 12 µm2) was captured with 50 Hz frequency (this is twice the video-rate since the odd and even frames of each interlaced image were recorded separately) [Pierres et al., 1996a] and a series of 256 sequential images was transferred to the computer memory for detailed analysis of the initial step of cell deformation. Fluorescent Labeling and pH or Calcium Monitoring In some experiments, the cell nucleus was labeled by a 10-min incubation at 37°C in culture medium supplemented with 1 µg/ml Hoechst 33342 vital dye (Calbiochem, LaJolla, CA). Observation was performed with the IMT2 DMU filter set (excitation wavelength ⬍ 400 nm). Calcium monitoring was performed as previously described [Kaplanski et al., 1994] by incubating cells for 30 min at 37°C in RPMI 1640 medium containing 3 µM fluo-3-AM and 1/1,000 pluronic (Molecular Probes, Eugene, OR). Cells were then aspirated under microscopi- cal observation with continuous dim visible light and brief exposure to ultraviolet light every 10 sec. Observation was performed with fluorescein filter set (IMT2 DMB). The calcium concentration was calculated with the following formula: [Ca2⫹] ⫽ Kd ⫻ /(1 ⫺ ) (1) where is the ratio between measured cell fluorescence and fluorescence measured after adding ionomycin in calcium rich environment. The dissociation constant of fluo-3/calcium complex was taken as 390 nM (Molecular Probes, Eugene, OR). Formula  is based on the assumption that calcium-free fluo-3 is essentially nonfluorescent (Molecular Probes). Cytosolic pH was studied on whole cell populations [Ohkuma and Poole, 1978]. Cells were labeled by 30-min incubation at 37°C with 10 µM BCECF-AM (Molecular Probes). Calibration curves [Bouvier et al., 1994] were built by incubating cells (106/ml) in buffered solutions (10 mM HEPES, 140 mM KCl) with pH ranging between 6 and 7.8 after a 3-minute incubation with the proton ionophore nigericin (10 µM) before fluorescence determination in a Perkin Elmer LS5 spectrofluorimeter (5 nm slit width, 535 nm emission, 439 and 505 nm excitation). The ratio between fluorescence values at both excitation wavelengths (F505/F439) was then plotted vs. pH. Polymerized actin was revealed [Horoyan et al., 1990] by incubating cells for 20 min at room temperature in presence of 3.7% paraformaldehyde, then for another 20-min period in phosphate buffer containing 10 U/ml bodipy phallacidin (Molecular Probes) and 0.1 mg/ml lysophosphatidylcholine (Sigma, St Louis, MO). Myosin was labeled after permeabilization of paraformaldehyde-fixed cells with 0.2% triton in phosphate buffer for 20 min at room temperature. Cells were then incubated first with 1/10 goat serum in PBS containing 1% BSA (15 min), then with 1/30 mouse anti-myosin (mouse anti-myosin light chain IgM, clone MY-21, ref M-4401 from Sigma) for 60 min at room temperature, and finally with fluorescein-conjugated goat anti-mouse IgM (Sigma, ref F-9259), 1/100 dilution. ABP-280 was labeled with a similar procedure, using mouse IgG1 anti-chicken gizzard filamin (Sigma, clone FIL-2, ref F-1888, 1/20 dilution), then fluoresceinconjugated goat anti-mouse IgG1 antiserum (Sigma, F2772). Cell Calcium Manipulation In some experiments, cells were incubated with 1 µM ionomycin in normal aspiration medium, or in medium supplemented with 1 mM EGTA, or in a 100 nM calcium buffer obtained by mixing 1 mM calcium or EGTA solution. Calcium concentration was checked with a spectrofluorimeter, after adding 1 µM fura-2AM and Mechanical Cell Response to Calcium Changes measuring fluorescence (520 nm emission, 340 and 380 nm excitation, 5 nm slit width), using Grinkiewicz formula [Grynkiewicz et al., 1985] and taking 224 nM as dissociation constant of calcium/fura 2 complex. Ionomycin concentration was selected in order that no lag was detected before ionophore addition and alteration of cytosolic calcium concentration, and this concentration remained constant for at least several minutes. Flow Cytometric Analysis of Actin Polymerization After incubating cells for different periods of time in ionomycin-containing solutions of varying calcium concentration, paraformaldehyde was rapidly added and cells were labeled with bodipy-phallacidin as described. Fluorescence determination was performed with an Epics XL flow cytometer (Coulter, Hialeah, FL). Controls were kept in aspiration buffer without any exposition to ionomycin. Zero time samples were obtained by sequentially adding ionomycin and paraformaldehyde with less than a 1-sec interval. Confocal Microscopy The distribution of cytoskeletal elements was determined as follows: cells were monitored with a confocal laser scanning microscope (CLSM, Leica, Heidelberg, Germany). Under standard conditions, series of 16 parallel sections performed at 1-µm intervals along the z axis were acquired and analysis was performed on the plane closest to the cell center. Images were transferred to an IBM-compatible computer, and a custom-made software was used to determine the dependence of the mean fluorescence intensity on the distance to the cell boundary. This function was chosen instead of the conventional radial distribution of fluorescence (i.e., the dependence of fluorescence intensity on the distance to the cell center) to allow quantitative assessment of the sharpness of the actin subplasmalemmal layer without any dependence on cell shape and deviation from sphericity. RESULTS Mechanical Cell Deformation May Involve Three Distinct Phases In a first step, we observed the deformation exhibited by 19 individual cells when they were subjected to a sucking pressure of 100 Pa with a pipette of ⬃5 µm internal diameter. A typical series of images is depicted in Figure 2. When the protrusion length was plotted vs. time, the curve shape was suggestive of three typical phases (Fig. 3): 1. immediately after cell-to-pipette contact, a protrusion of a few micrometers length appeared within about one tenth of a second (Fig. 3a). 97 2. during a period of about 5 sec, the protrusion length increased with a fairly constant rate of order of 1 µm/sec (Fig. 3b). 3. during the following tens of seconds, the protrusion length still increased with fairly lower velocity. Length of Protrusions Measured After a Few Seconds Aspiration May Be Mostly Representative of the Passive Mechanical Properties of the Cell Cortex Additional experiments were performed to assess the potential influence on protrusion length on several phenomena unrelated to passive mechanical properties of the cytosol. First, cell exposure to mechanical stimulation might trigger biochemical cascades resulting in cytoskeletal alterations and possible active movements [Evans and Kukan, 1984; Horoyan et al., 1990; Zaffran et al., 1993]. This possibility was tested by labeling cells with fluo-3 and monitoring fluorescence during aspiration. The mean cytosolic calcium concentration determined on 21 control cells was 183 nM. When 16 fluo-3-labeled cells were subjected to micropipette aspiration, eleven displayed a substantial rise of cytosolic calcium, without any clearcut intracellular localization, during the first 75 sec. Except in a single case, this increase occurred a few tens of seconds after cell-to-pipette contact. The maximum calcium concentration might be higher than 1 µM (2/16), and in most cases it ranged between 300 and 400 nM (4/16) or 400 and 1,000 nM (5/16). Thus, cell mechanical properties measured 10 sec or more after aspiration might be influenced by uncontrolled biochemical events. A potentially important phenomenon likely to occur during aspiration of cells different from polymorphonuclear granulocytes might be the plugging of the pipette entry by the poorly deformable nucleus. This possibility was tested by monitoring cells with a nucleus made fluorescent by treatment with Hoechst 33342 stain. As exemplified in Figure 4, the nucleus often closely approached the pipette entrance when cells were substantially deformed by the formation of a large protrusion. The significance of deformations measured during the first seconds after the onset of aspiration was then studied. The determination of the initial deformation (e.g., after 1-sec aspiration) was hampered by two phenomena. First, it was difficult to keep the micropipette tip in focus at the moment of contact with the cell, thus making difficult accurate determination of cell boundary. Second, some images strongly suggested that the cell membrane might get separated from underlying cytosolic gel during a short time (Fig. 4a), suggesting that the 98 Richelme et al. Fig. 2. Analysis of cell response to micropipette aspiration. A typical rounded THP-1 cell (A) was sucked into a micropipette. A protrusion was rapidly generated, and it displayed obviously heterogeneous internal structure 1.6 sec after the onset of aspiration (B). The protrusion length was markedly increased 15 sec later (C). A series of images of the pipette tip were recorded with 50 Hz frequency, starting immediately before the onset of aspiration. Sixty-four sequential images are shown (D). Since odd and even lines of each image were separated, a 50% reduction of image height is visible. Cell-to-pipette contact is initiated on the 20th image, and a growing protrusion is clearly apparent on the 22th image. The protrusion was marked with a white line in order to make it more visible. Further, the heterogeneity of protrusion structure is revealed by a double contour that is underlined on the 43th image (6th row, 3rd column) with a black line. Aforementioned images 20, 22, and 43 were surrounded with a dashed boundary. Bar ⫽ 5 µm. protrusion length might not accurately reflect cytoskeletal deformation during the first few seconds (in contrast with later situation) [Richelme et al., 1997]. It was thus concluded that the protrusion length measured 3–5 sec after the onset of aspiration might yield a reproducible and significant account of cytoskeletal mechanical properties. ported by the finding that the protrusion length measured after 5-sec aspiration was not correlated to cell diameter; indeed, correlation coefficient between both parameters was 0.00001 when 26 cells with a diameter ranging between 12 and 20 µm were studied (not shown). Thus, the protrusion length determined under our experimental conditions might be expected to reflect mechanical properties of the cell cortex. Protrusion Length Measured a Few Seconds After the Onset of Low Pressure Aspiration Is Dependent on Local Cell Properties Dependence of Cell Deformation on Applied Pressure It was important to know whether the protrusion length measured under our experimental conditions was mainly dependent on geometric cell properties (e.g., ratio between cell and pipette diameter), or on local behavior of the deformed region. The latter hypothesis was sup- It was important to assess the influence of chosen sucking pressure on the parameters we measured. Thus, deformation kinetics were determined on series of cells subjected to aspiration with a pressure ranging between 100 and 800 Pa. As shown in Figure 5, the protrusion Mechanical Cell Response to Calcium Changes 99 Fig. 4. Nucleus behavior during micropipette aspiration. Cells were processed for intravital fluorescent labeling of their nucleus before micropipette aspiration. They were observed with a combination of fluorescent and visible illumination. A: Before aspiration, the nucleus is separated from the pipette tip by a cytoplasmic layer. B: The nucleus usually went very close to the pipette tip a few seconds after the onset of aspiration. Bar ⫽ 10 µm. Fig. 3. Kinetics of mechanical cell deformation. The kinetics of elongation of the protrusion formed on the cell displayed in Figure 2 during micropipette aspiration is shown. A: Initial elongation kinetics appear with 20-msec resolution, suggesting the occurrence of an initial elastic step (during the first 0.1 sec). B: Variations of protrusion length (black squares, full line) and pressure (black triangles, dashed line). The initial elastic deformation was followed by a rapid (between ⬃1 and 5 sec) and slower (time ⬎5 sec) phases with a fairly constant deformation rate. length was nearly linearly dependent on the pressure when this was comprised of between 100 and 200 Pa, but no significant difference was found between the deformation exhibited by cell populations subjected to sucking pressures of 500 and 800 Pa. Thus, in further experiments, the length of protrusions measured after a 3–5-sec aspiration with a pressure of 100 Pa was considered as representative of the deformability of the submembrane cortical area. Cell Deformability Is Increased by a Prolonged Rise of Cytosolic Calcium Concentration Cells were incubated for about 3 min in medium containing ionomycin and different calcium concentrations before being assayed for deformability. As shown in Figure 6, cells exposed to ionomycin in standard medium exhibited about 30% increase of deformation after a 3-sec aspiration, and this response was not due to the ionophore alone since no deformability change was observed when ionomycin was added in a calcium buffer whose concentration was close to the cytosolic calcium concentration of resting cells. Interestingly, when ionomycin was added 100 Richelme et al. Fig. 5. Effect of applied pressure on the rate of pipette elongation. THP-1 cells were sucked into micropipettes with a pressure of 100 Pa (diamonds, 19 cells), 200 Pa (squares, 7 cells), 500 Pa (triangles, 10 cells), or 800 Pa (crosses, 14 cells) and elongation kinetics was determined. Each point represents a mean value. Vertical bars ⫽ twice the standard error. Fig. 7. Effect of the duration of intracellular calcium increase on measured rigidity. THP-1 cells were incubated with ionomycin and 1 mM Ca2⫹ during a period of time t falling into the range 10 s ⬍ t ⬍ 20 s (squares, 3 cells), 20 s ⬍ t ⬍ 30 s (triangles, 6 cells), 30 s ⬍ t ⬍ 50 s (crosses, 5 cells), 50 s ⬍ t ⬍ 60 s (stars, 6 cells), 60 s ⬍ t ⬍ 80 s (circles, 10 cells), or 300 s ⬍ t (vertical segments, 18 cells). They were then sucked into a micropipette and the length of formed protrusion was determined 3, 6, and 10 s later. Results are expressed as percent of controls (dots, 31 cells). Each point is a mean value. Effect of Calcium Rise on Cell Deformability Is Biphasic Since many intracellular biochemical events can be triggered by calcium changes, it was difficult to know whether a single messenger was responsible for deformability changes. In order to address this possibility, the kinetics of deformability changes was studied. As shown in Figure 7, increasing intracellular calcium resulted in a decrease of cell deformability within 10 sec, and deformability progressively increased during the next minutes. Fig. 6. Effects of intracellular calcium changes on rigidity. THP-1 cells were incubated for 3 min in control medium (diamonds) or medium supplemented with 1 µM ionomycin in presence of a total calcium concentration of about 1 mM (triangles, 18 cells) or 100 nM (squares, 32 cells), or with 1 mM EGTA (crosses, 31 cells). The kinetics of protrusion elongation were determined on individual cells during the first 10 sec following aspiration into a micropipette with a negative pressure of 100 Pa. together with a calcium chelator, a weak increase of deformability was found. Since calcium changes often lead to cytosolic pH fluctuations and since many actin binding proteins are dependent on pH as well as calcium, it was interesting to determine whether intracellular pH was altered by ionomycin treatment. Using BCECF, it was found that no significant pH change occurred 5 min after the addition of ionomycin (not shown). Intracellular Calcium Rise Results in Rapid Changes of F-Actin Content and These Modifications Do Not Match Deformability Changes In view of the hypothesis that the actin cytoskeleton strongly influences cell mechanical properties, it was interesting to know whether calcium-induced deformability changes reflected concomitant alterations of microfilament polymerization. This question was addressed by staining THP1 cells with fluorescent phalloidin at different moments following exposure to ionomycin, then assaying sequential samples with flow cytometry. As shown in Figure 8, a calcium rise resulted in an immediate drop of microfilament labeling followed by a rapid recovery during the following 10–20 sec. This change was not due to the ionophore alone since minimal variations of actin polymerization were found in cells exposed to ionomycin in a medium with a calcium concentration close to cytosolic value. Mechanical Cell Response to Calcium Changes 101 fluorescence micrographs displayed a similar behavior as obtained with flow cytometry (not shown). As shown in Figure 10, the distribution of myosin and ABP-280 was essentially unaltered by calcium rise, with a fairly uniform ABP-280 distribution and a cortical concentration of myosin. In contrast with this finding, microfilament Fig. 8. Effect of intracellular calcium change on actin polymerization. THP-1 cells were incubated in medium containing 1 µM ionomycin and 1 mM CaCl2. They were processed at various moments for determination of polymerized actin with flow cytometry. Each point represents a mean calculated on 6 separate experiments. Vertical bar ⫽ twice the standard error. Intracellular Calcium Rise Results in Rapid Change of Intracellular Distribution of Microfilaments, Not Actin-Binding Proteins Since the cell mechanical properties might be dependent on a particular microfilament population, a mere quantification of F-actin content might not be sufficient to assess relationships between cytoskeletal structure and behavior. Thus, THP-1 cells were fixed at different moments following calcium rise and they were processed for confocal microscopic study of the distribution of microfilaments as well as actin binding proteins ABP-280 and myosin. As shown in Figure 9, while control cells displayed marked concentration of microfilaments in the cortical area, calcium rise might result in marked decrease of peripheral fluorescence. Interestingly, this was probably due to microfilament redistribution rather than actin depolymerization, since the total fluorescence content determined by quantitative analysis of Fig. 9. Quantitative analysis of intracellular fluorescence. THP-1 cells were labeled with bodipy-phallacidin and observed with confocal microscopy. Series of 16 parallel sections separated by 1-µm intervals were recorded and typical images obtained with a control (A) or ionomycin-treated (B) cell are shown. A diametral section was used to determine the variations of the mean fluorescence intensity as a function of distance to the boundary between cell interior and extracellular space. The curve obtained by analysing A is shown in C where fluorescence intensity was plotted vs. distance to the cell boundary. This curve (squares, solid line) is representative of peripheral fluorescence distribution, with a narrow peak at low distance from the cell edge. In contrast, analysis of B yielded a curve (circles, dashed line in C) representative of uniform marker distribution. Bar ⫽ 10 µm. 102 Richelme et al. Fig. 10. Effect of intracellular calcium increase on the distribution of polymerized actin, myosin, and ABP-280. Cells were incubated for various amounts of time in a medium supplemented with 1 µM ionomycin and containing 1 mM Ca2⫹. They were then fixed and polymerized actin (white areas), ABP-280 (hatched areas), or myosin ( grey areas) were stained with fluorescent labels for examination with confocal microscopy. Fluorescence distribution curves were derived from each confocal image as exemplified in Figure 9, and each curve was used to calculate a ‘‘mean width of fluorescence peak,’’ i.e., the width of the region where fluorescence density was higher than half the peak value. The mean width of fluorescent peaks was used as an indicator of fluorescence type (i.e., peripheral vs. homogeneous). Each point represents a mean determined on about ten randomly selected cells. Vertical bar ⫽ twice the standard error. distribution displayed a multiphasic behavior during the first seconds following calcium increase, with a recovery of peripheral concentration a few tens of seconds later. DISCUSSION The aim of our study was first to define a simple method for measuring and describing local cell deformability, second to use this approach to improve our understanding of the influence of free calcium on the mechanical properties of whole cells, and third to relate our findings to microfilament organization. In view of the complexity and highly dynamic features of cell behavior, the basic strategy consisted of achieving time-resolved control of intracellular calcium level, determination of rheological properties, and monitoring of cytoskeletal organization. Our results emphasize the requirement for a kinetic approach in studying cell shape control. Length of the Protrusion Formed After a Few Seconds Aspiration Is a Convenient Index of Cortical Cell Deformability Our results confirm the viscoelastic behaviour of tested cells since applied forces induced a rapid (within the millisecond scale) elastic response followed by con- tinuous deformation suggestive of a viscous material, in accordance with previous reports [Schmid-Schönbein et al., 1981; Evans and Kukan, 1984]. The length of the protrusion induced by 3–5-sec aspiration seemed the most convenient parameter for a semi-quantitative description of cortical cell deformability in view of the following reasons. First, the initial elastic deformation as completed within ⬇1 sec was fairly small, which limited the accuracy of experimental studies. Also, its physiological relevance may be questioned, since more extensive deformations are required to influence important processes such as migration. Finally, the possibility that initial deformation might involve a detachment of the plasma membrane from underlying cytoskeletal elements, as considered by Janmey  and exemplified in Figure 2b, makes the interpretation of initial deformations more difficult. Second, the deformation measured after several tens of seconds is difficult to interpret in view of several problems: (1) parameters unrelated to cytoplasmic rheology, such as nucleus localization (Fig. 4) may influence long-term deformation. This might account for the unexpected finding that cytochalasin treatment decreased the rate of late-phase cell deformation in response to constant forces [Richelme et al., 1997]; (2) the triggering of spontaneous calcium rise during cell aspiration might be indicative of the triggering of intracellular biochemical cascades likely to generate active deformations [Evans and Kukan, 1984; Horoyan et al., 1990]; and (3) as demonstrated by our results, it was important to achieve a rapid measurement of cell deformability in view of the time dependence of studied phenomena. Increasing Intracellular Calcium Results in a Biphasic Alteration of Cell Deformability Calcium was an obvious candidate for a study of the effect of intracellular messengers on mechanical properties since (1) it is known to affect many cytoskeletal components [Schmidt and Hall, 1998]; (2) intracellular calcium changes are often associated to mechanical responses [Sawyer et al., 1985; Kruskal et al., 1986]; and (3) intracellular calcium buffering makes it a suitable candidate for the control of localized intracellular processes [Albritton et al., 1992]. Our conclusion that calcium rise induced a transient stiffening followed by a delayed deformability increase is consistent with the previous report that the disruption of stress fibers by endogenous gelsolin might require a durable increase of cytoplasmic calcium [Kanno and Sasaki, 1989]. This may also account for some discrepancy in reported data: indeed, increasing intracellular calcium increased the rigidity of rat basophilic leukemia cells [Horoyan et al., 1990] or isolated cytoplasmic strands [Adams, 1992], but Mechanical Cell Response to Calcium Changes a similar treatment was occasionally noticed to soften THP-1 cells [Richelme et al., 1996]. Increasing Intracellular Calcium Results in Time-Dependent Changes of Actin Polymerization and Microfilament Distribution Since microfilaments certainly play a dominant role in the determination of cell resistance to deformation [Janmey et al., 1998], it was interesting to look for a correlation between cell rheological changes and cytoskeletal alterations. The moderate long-term increase of actin polymerization is consistent with reports by Sha’afi et al.  and Downey et al. . Interestingly, the latter authors reported that a calcium ionophore might trigger actin polymerization, whereas a calcium rise increased actin depolymerization in permeabilized cells, thus emphasizing the need to study whole cells in order to understand physiological means of regulating cytoskeletal organization. It is worth emphasizing that the very brief decrease of actin polymerization that was detected immediately after addition of ionomycin required very rapid manipulation, with a minimal delay between ionophore addition and cell fixation. As emphasized below, the mere amount of actin polymerization might not account for all of cell rigidity. It was thus warranted to study the effect of calcium changes on the intracellular distribution of microfilaments as well as ABP-280 and myosin, two important actin binding proteins with a potentially important role in cortical microfilament organization and function [Hartwig and Shevlin, 1986; Pasternak et al., 1989; Cunningham et al., 1992]. Indeed, Janmey  proposed the attractive concept that a calcium increase might disrupt microfilaments by activating F-actin severing proteins and inhibiting microfilament-cross-linking species. The main conclusion of our microscopical study is that a rise of intracellular calcium concentration triggered a very rapid (less than a few seconds) displacement of a fraction of cortical microfilaments to inner cell regions. However, no redistribution of myosin or ABP-280 could be detected. Note that more work is required to determine whether the effect of calcium change on cell mechanical properties and cytoskeletal organization was direct or was mediated by downstream cascades such as phosphorylation/dephosphorylation events or phosphatidylinositol release resulting from phospolipase activation. Indeed, intricate links exist between calcium and other activators, as illustrated by possible variations of cell sensitivity to calcium changes [Bradley and Morgan, 1987; Uebata et al., 1997]. Although it is admittedly impossible to control the totality of potential consequences of intracellular calcium changes, it is hoped that our dynamic approach with minimal separation between cell stimulation and 103 mechanical test may allow further dissection of the mechanisms of cytoskeletal controls, through systematic use of available inhibitors of calcium-triggered cascades. Several Mechanisms Might Account for the Relationship Between Cell Structural and Mechanical Properties During the first few seconds following calcium rise, cortical deformability was decreased while a fraction of peripheral microfilaments displayed centripetal displacement. This is consistent with the view that only a subpopulation of these microfilaments might account for cortical rigidity. An attractive hypothesis would be that this subpopulation is interacting with cross-linking proteins such as ABP-280. In this case, it is interesting to determine the structural substratum for cell rigidity changes. Three possible mechanisms for stiffening might be [Ferry, 1980; Wachsstock et al., 1994]: (1) increase of average filament length, (2) increase of filament crosslinking, and (3) increase of the lifetime of cross-links between filaments. The latter mechanism is of particular interest: indeed, if the dissociation of bonds between microfilaments and a particular species of bridging molecules was a limiting step for cell deformation, cells could manage to change their deformations without any visible change of cytoskeletal network if only the off-rate of some class of intermolecular association was modulated by variations of ionic environment or protein phosphorylation. This concept may be tested by recent approaches allowing simultaneous determination of the lifetime and mechanical behavior of intermolecular association [Evans et al., 1991, 1995; Florin et al., 1994; Pierres et al., 1996b]. CONCLUSION The results we describe illustrate the complexity of microfilament regulation by calcium. 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