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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: bongrand@marseille.inserm.fr
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 [1997] 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. [1981] 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 [1] 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 [1995] 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.
[1986] and Downey et al. [1990]. 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 [1994] 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. Indeed, a sharp
increase of intracellular calcium concentration resulted in
(1) marked cortical stiffening within 1 sec, followed by a
significant softening that was delayed by at least 1 min;
(2) decrease of polymerized actin content within 1 sec
followed by complete recovery within about 20 sec; and
(3) oscillatory behaviour of the intracellular distribution
of polymerized actin. These findings suggest that only a
kinetic study of the mechanical and structural changes
induced in a whole cell by well-defined stimuli might
provide valuable insight into the mechanisms of cytoskeletal control. Also, they emphasize the absence of close
relationship between microfilament morphological features and cell mechanical properties. This may be an
incentive to study the effect of metabolic changes on the
lifetime and force dependence of interactions between
actin and actin binding proteins.
104
Richelme et al.
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