Am J Physiol Cell Physiol 302: C527–C538, 2012.
First published November 2, 2011; doi:10.1152/ajpcell.00248.2011.
Kinase activation of ClC-3 accelerates cytoplasmic condensation during
mitotic cell rounding
Vishnu Anand Cuddapah, Christa W. Habela, Stacey Watkins, Lindsay S. Moore, Tia-Tabitha C. Barclay,
and Harald Sontheimer
Department of Neurobiology and Center for Glial Biology in Medicine, University of Alabama at Birmingham, Birmingham,
Alabama
Submitted 15 July 2011; accepted in final form 27 October 2011
glioma; chloride channel
BEFORE DIVIDING INTO TWO DAUGHTER CELLS, mammalian cells
transform shape from flat during interphase to round during
M-phase, typifying a process termed “mitotic cell rounding.”
Mitotic cell rounding is an intrinsic and stereotypic phenomenon associated with cell division (35). The importance of
mitotic cell rounding is manifold: it stabilizes and positions the
mitotic spindle, a curved plasma membrane exposes lipids
important in cell cycle progression, and it ensures that both
daughter cells receive the correct chromosomal set with high
fidelity (30). Therefore mitotic cell rounding seems to be
important in the mechanics as well as signaling of cell division.
A recent study demonstrated that mitotic cell rounding is
primarily driven by two opposing forces: an inward cytoskeletal contraction versus an outward osmotic force (35). Actomyosin cytoskeletal dynamics promoting mitotic cell rounding
have been well studied. Moesin is activated during mitosis and
attaches actin filaments to the plasma membrane, increasing
cortical rigidity and cell rounding (14). Additionally, RhoA, a
small GTPase, activates Rho kinase, leading to increased
actomyosin contractility (19). However, relatively little is
Address for reprint requests and other correspondence: H. Sontheimer, Dept.
of Neurobiology, Univ. of Alabama, 1719 6th Ave. South, CIRC 410, Birmingham, AL 35294 (e-mail: [email protected]).
http://www.ajpcell.org
known about the outward osmotic force contributing to mitotic
cell rounding.
Several studies performed in human glioma cells demonstrate that Cl⫺ channels substantially contribute to this outward
osmotic pressure. M-phase cells have elevated Cl⫺ currents as
compared with interphase cells (9). These elevated Cl⫺ currents in dividing cells are associated with a decrease in cytoplasmic volume, termed “premitotic condensation” (PMC) (9).
As osmotically active Cl⫺ exits dividing cells, water is osmotically driven out of the cytosol, leading to an outward osmotic
pressure and decrease in cellular volume (9). Incubation of
dividing cells in hypertonic media significantly decreases this
outward osmotic pressure and hampers PMC (9). The molecular identity of Cl⫺ channel responsible for this Cl⫺ efflux in
glioma cells was determined to be ClC-3, a voltage-gated Cl⫺
channel/transporter (8). ClC-3 has previously been implicated
in shape and volume changes in a variety of cells, including
glioma cells (26). Additionally, ClC-3 activity facilitates proliferation in a number of cell types including nasopharyngeal
carcinoma cells (39) and smooth muscle cells (25). However,
how ClC-3 becomes activated in dividing cells leading to
premitotic condensation is unknown. Previous reports have
demonstrated that ClC-3 can be regulated by kinases, including
Ca2⫹/calmodulin-dependent protein kinase II (CaMKII) (11).
Using a combination of imaging and biophysical techniques,
we demonstrate that CaMKII activates ClC-3 in dividing cells,
promoting premitotic condensation. ClC-3 and CaMKII
strongly colocalize on the plasma membrane of dividing cells
during mitotic cell rounding. Time-lapse microscopy revealed
that inhibition of CaMKII or ClC-3 impedes premitotic condensation to the same extent. We directly measured ClC-3
activity in dividing cells using whole cell patch-clamp electrophysiology and found that dividing cells had an increased Cl⫺
conductance that was CaMKII dependent. Inhibition of CaMKII impaired PMC selectively in ClC-3-expressing cells, suggesting an important role of CaMKII modulation of ClC-3 in
PMC.
METHODS AND MATERIALS
Cell culture. D54 human glioma cells, derived from a glioblastoma,
are a World Health Organization grade IV cell line. These cells were
gifted by Dr. D. Bigner (Duke University, Durham, NC). Cells were
passaged in Dulbecco’s modified Eagle’s medium (DMEM)-F-12
(Invitrogen), supplemented with 2 mM glutamine and 7% fetal bovine
serum (Hyclone, Logan, UT), and incubated in 10% CO2 at 37°C. All
reagents were purchased from Sigma-Aldrich (St. Louis, MO) unless
otherwise specified.
Time-lapse microscopy. Cells were monitored for spontaneous
divisions as previously described (8). First, D54 cells were stably
transfected with green fluorescent protein (GFP) with the peGFP-N1
0363-6143/12 Copyright © 2012 the American Physiological Society
C527
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Cuddapah VA, Habela CW, Watkins S, Moore LS, Barclay TT,
Sontheimer H. Kinase activation of ClC-3 accelerates cytoplasmic
condensation during mitotic cell rounding. Am J Physiol Cell Physiol 302: C527–C538, 2012. First published November 2, 2011;
doi:10.1152/ajpcell.00248.2011.—“Mitotic cell rounding” describes
the rounding of mammalian cells before dividing into two daughter
cells. This shape change requires coordinated cytoskeletal contraction
and changes in osmotic pressure. While considerable research has
been devoted to understanding mechanisms underlying cytoskeletal
contraction, little is known about how osmotic gradients are involved
in cell division. Here we describe cytoplasmic condensation preceding
cell division, termed “premitotic condensation” (PMC), which involves cells extruding osmotically active Cl⫺ via ClC-3, a voltagegated channel/transporter. This leads to a decrease in cytoplasmic
volume during mitotic cell rounding and cell division. Using a
combination of time-lapse microscopy and biophysical measurements,
we demonstrate that PMC involves the activation of ClC-3 by Ca2⫹/
calmodulin-dependent protein kinase II (CaMKII) in human glioma
cells. Knockdown of endogenous ClC-3 protein expression eliminated
CaMKII-dependent Cl⫺ currents in dividing cells and impeded PMC.
Thus, kinase-dependent changes in Cl⫺ conductance contribute to an
outward osmotic pressure in dividing cells, which facilitates cytoplasmic condensation preceding cell division.
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CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
with a series resistance of ⬎12 M⍀ were omitted because of poor
electrical access. pClamp 9.0 software (Axon Instruments) was used
to acquire and store data on a desktop computer. Standard KCl pipette
solution consisted of 140 mM KCl, 1 mM MgCl2, 0.2 mM CaCl2, 10
mM EGTA, and 10 mM HEPES sodium salt. The pipette solution was
adjusted to an osmolarity of 302–304 mosM and pH 7.2 with Tris
base. The bath solution consisted of 130 mM NaCl, 5 mM KCl, 1 mM
CaCl2, 10.55 mM glucose, and 32.5 mM HEPES acid, and was
adjusted to 308 –312 mosM and pH 7.4 with NaOH. All recordings
were performed in the presence of 2 ␮M paxilline (Santa Cruz
Biotechnology) to block BK channels (38). 5-Nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) was bath applied at 200 ␮M to block
Cl⫺ channels. Cells were incubated in myristoylated autocamtide-2related inhibitory peptide (AIP; Enzo Life Sciences) for 10 min before
recordings were performed.
ClC-3 knockdown. We stably knocked down endogenous ClC-3
expression in D54 human glioma cells as previously described (8). A
pGIPZ-lentiviral small hairpin mir vector containing a hairpin sequence
targeting CLCN3 (Open Biosystems) was transfected with the Amaxa
Biosystems nucleofection technique. After generating clonal populations,
we screened multiple targeting constructs for ClC-3 knockdown by
quantitative real-time (qRT-PCR). Multiple clones with ClC-3 knockdown were selected and probed for protein knockdown with Western
blotting. We then selected a subset of clones to functionally test for cell
migration, which requires ClC-3 activity (4). Experiments in this paper
used a representative clone with the hairpin sequence 5=-TGCTGTTGACAGTGAG CGCCCTACCTCTTTCCAA AGTATAT AGTGAAGCCACAGATGTATATA CTTTGGAAAGAGGTAGGATGCC TACTGCCTCGGA-3=. We used a nontargeting sequence as a negative control:
NT, 5=-TGCTGTTGACAGTGAGCGATCTCGCTTGG GCGA GAG
TAAGTAGTGAAGCC ACAGATGTACTTACTCTCGCCCAAGCGA GAGTG CCTA CTGCCTCGGA-3=. Transfected cells were
maintained on 10 ␮g/ml puromycin.
Western blot analysis. Western blot analysis was performed as
previously described (4). Protein concentrations were determined
using a NanoDrop 1000 spectrophotometer (Thermo Scientific), and
15 ␮g of protein were loaded per well. Sample buffer (6⫻; 60%
glycerol, 300 mM Tris pH 6.8, 12 mM EDTA, 12% SDS, 864 mM
2-mercaptoethanol, 0.05% bromophenol blue) was added to samples,
which were then loaded on 10-well, 10% gradient precast SDSpolyacrylamide gels (Bio-Rad). After gels were run at 120 mV for 75
min, they were transferred to polyvinylidene difluoride paper (Millipore, Bedford, MA) at 200 mA for 120 min. Blots were then blocked
with blocking buffer containing 10% nonfat dried milk and Trisbuffered saline with 0.1% Tween 20. Primary antibodies [rabbit
anti-ClC-3 (Alpha Diagnostic International; 1:1,500), rabbit antiClC-3 (Alomone Labs; 1:1,500), and mouse anti-glyceraldehyde-3phosphate dehydrogenase (Abcam; 1:5,000)] were incubated for 1 h at
room temperature. After washes, blots were incubated in respective
horseradish peroxidase-conjugated secondary antibodies at 1:1,500
(Santa Cruz Biotechnology). After further washes, blots were then
developed with Supersignal West Femto (Thermo Scientific) or Luminol Reagent (Santa Cruz Biotechnology) and imaged on an Eastman Kodak Image Station 4000MM. ClC-3 immunoreactivity was
normalized to glyceraldehyde-3-phosphate dehydrogenase.
Proliferation assay. Cells were placed on six-well plates at a
density of 1 ⫻ 104. On day 0, a well was collected to normalize all
future cell collections. Cell numbers were measured using a Coulter
Counter Multisizer 3 (Beckman Coulter, Miami, FL). Drugs, including 40 ␮M NPPB and 2.5 ␮M KN-93 or vehicle, and media were
changed every other day. Wells were measured in triplicate.
Bromodeoxyuridine incorporation. Cells were cultured on 12-mm
glass coverslips (Macalaster Bicknell, New Haven, CT) in a 24-well
plate for 1–3 days. Cells were then pulsed with 25 ␮M bromodeoxyuridine (BrdU) for 30 h. After 2 washes with PBS, the cells were fixed
at room temperature with 4% paraformaldehyde for 15 min. Cells
were again washed with PBS. DNA was then denatured with 2 M HCl
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(Clontech) plasmid and maintained with 0.25 mg/ml G418 (Invitrogen). Cells were then grown on four-chamber no. 1 German borosilicate glass slides (Nalge Nunc International) and cultured for 1–3 days
before imaging. Images were acquired with an Axiocam MRm camera
mounted on an Axiovert 200M microscope (Carl Zeiss) in 5- to
10-min intervals for 48 h with a ⫻20 Fluar air objective. The stage
was housed in an incubator maintained at 37°C with 5% CO2. We
used a metal halide X-cite 120 light source (EXFO Photonics Solutions) with a FITC filter cube (excitation, 480 ⫾ 15 nm; emission, 535 ⫾
20 nm; Chroma Technologies). Neutral density filters were used to
reduce phototoxicity. The acquired 16-bit TIF images were then
screened for dividing cells. Regions of interest (ROIs) in the cytoplasm of dividing cells were selected in National Institutes of Health
WCIF ImageJ to measure changes in cytoplasmic mean pixel intensity, which is a reflection of cell volume as previously demonstrated
(9). Mean pixel intensities were compiled in Microsoft Excel 2007,
then the time points and fluorescence intensities were normalized. For
each cell, we designated the time point just before cell division as time ⫽
0 and normalized all time points to time ⫽ 0. Therefore, if a cell began
changing volume at ⫺ x minutes, it indicates that x minutes before
division, the cell began to change its volume. Each cell’s fluorescence
intensity (F) was normalized to its own fluorescence intensity at time ⫽ 0
(F0). The normalized averages were then compiled into Origin 6.0
(MicroCal, Northampton, MA). To calculate Fmin, Fmax, and t50,
the data were fit to a sigmoidal curve with a linear X scale
(Boltzmann fit; R2 ⬎ 0.99 for all conditions). The Fmin and Fmax
were the lowest and highest F/F0 intensities for each condition,
respectively.
Immunocytochemistry. Cells were immunolabeled as previously
described (4). D54 cells were grown for 2– 4 days on glass coverslips
(Macalaster Bicknell). After culturing, cells were washed with PBS
and fixed for 10 min with 4% paraformaldehyde. After blocking for 30
min at room temperature with PBS containing 10% normal goat serum
and 0.3% Triton X-100, cells were incubated in primary antibodies
overnight at 4°C. The next day, cells were washed and incubated in
secondary antibodies for 90 min at room temperature. Cells were
washed again and incubated in Hoechst 33342 at 100 ng/ml for 20 min
at room temperature. Cells were then washed and mounted onto glass
slides (Fisher Scientific) with Fluoromont (Sigma).
ClC-3 was labeled with polyclonal rabbit anti-ClC-3 (Alomone
Labs, Jerusalem, Israel; lot no. ACL001AN0702) used at 1:250.
CaMKII was labeled with monoclonal mouse anti-CaMKII (Abcam,
Cambridge, MA) used at 1:250. Autophosphorylated (p)CaMKII was
labeled with polyclonal rabbit anti-pCaMKII (T286; Abcam) used at
1:250. Secondary antibodies used were goat anti-rabbit Alexa 488 and
goat anti-mouse Alexa 546 (Invitrogen) at 1:500. Images were acquired with Slidebook software (version 5.0, Intelligent Imaging
Innovations) using a Hamamatsu IEEE1394 digital charge-coupled
device camera mounted on an Olympus IX81 microscope with an
Olympus scanning disc unit to remove out-of-focus light. A ⫻60 oil
immersion lens (numerical aperture, 1.42) was used to acquire 0.5-␮m
optical sections to generate 10-␮m image stacks. Alexa 488 fluorescence was imaged with a FITC filter set (excitation, 482 ⫾ 17 nm;
emission, 536 ⫾ 20 nm), Alexa 546 fluorescence was imaged with a
tetramethylrhodamine isothiocyanate filter set (excitation, 543 ⫾ 22
nm; emission, 593 ⫾ 20 nm), and Hoechst fluorescence was imaged
with a DAPI filter set (excitation, 387 ⫾ 5.5 nm; emission, 447 ⫾ 30
nm) (Semrock). Images were interpolated, displayed in “three-view,”
and linearly normalized with Slidebook software.
Electrophysiology. Before recordings were performed, cells were
loaded with Hoechst 33342 at 100 ng/ml in cell culture media for 25
min to aid in identification of M-phase cells with condensed DNA. For
whole cell patch-clamp recordings, D54 cells were recorded from
after 1–3 days in culture, with pipette tips ranging from 3 M⍀ to 5
M⍀ from thin-borosilicate glass (TW150F-4, World Precision Instruments, Sarasota, FL). Using an Axopatch 200B amplifier (Axon
Instruments), series resistance was compensated to 80%, and cells
CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
in PBS with 0.3% Triton X to permeabilize the plasma membrane for
30 min at room temperature. Cells were then blocked with 10%
normal goat serum and 0.3% Triton X-100 for 30 min at room
temperature. Mouse anti-BrdU antibody (1:100, Molecular Probes,
Eugene, OR) was then incubated overnight at 4°C. The next day, goat
anti-mouse Alexa 350 (1:500, Invitrogen) was incubated for 2 h at
room temperature. Coverslips were then washed in PBS and mounted
on glass slides.
Data analysis. Electrophysiological data were quantified with
Clampfit (Axon Instruments). Origin 6.0 (MicroCal, Northampton,
MA) or Microsoft Excel was used to compile and graph data.
GraphPad Instat (GraphPad Software) was used to perform statistical
analyses. All data are reported as means ⫾ SE.
RESULTS
by taking advantage of the fact that fluorescence of soluble
GFP is inversely proportional to cell volume, as a constant
number of GFP molecules are dissolved in the cytoplasm (9).
Therefore a decrease in cell volume manifests as an increase in
GFP intensity. Transfected glioma cells were imaged and
monitored for spontaneous cell divisions, and a representative
cell dividing under control conditions is depicted in Fig. 1A.
For each cell, we designated the time point just before cell
division as time ⫽ 0 and normalized all time points to time ⫽
0 for each cell. Therefore, if a cell began changing volume at
⫺x minutes, it indicates that x minutes before division, the cell
began to change its volume. We found that as cells approached
division, the relative intensity of GFP fluorescence increased,
indicating a decrease in cytoplasmic volume (Fig. 1, A and B).
Compiled averages are graphed in Fig. 1B, where the data were
normalized to F0, which is the GFP intensity at time ⫽ 0. Each
cell’s fluorescence intensity (F) was normalized to its own
fluorescence intensity at time ⫽ 0 (F0). As the cells approached
division, the GFP fluorescence intensity (F/F0) rapidly increased, indicating cytoplasmic condensation as expected during PMC. Changes in GFP fluorescence were not attributed to
pH, because as previously reported, no changes in pH were
detected using the ratiometric pH indicator 2=,7=-bis-(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF) in dividing D54
Fig. 1. CaMKII inhibition prolongs premitotic condensation in human glioma cells.
A: representative dividing cells incubated in
vehicle or 10 ␮M autocamtide-2-related inhibitory peptide (AIP), a CaMKII inhibitor.
CaMKII inhibition increases the time needed
for premitotic condensation (PMC). Time (t)
was normalized to t ⫽ 0, which is the last time
point preceding cell division. B: CaMKII inhibition with 10 ␮M AIP increases the total time
needed for PMC. GFP, green fluorescent protein. C: minimum and maximum fluorescence
intensities (Fmin and Fmax) do not significantly
change with CaMKII inhibition. D: CaMKII
inhibition with 10 ␮M AIP increases the time
needed for 50% volume condensation (t50).
Error bars indicate SE; n ⫽ 26 –27. *P ⬍
0.001, unpaired t-test.
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Mitotic cell rounding in glioma cells is associated with
CaMKII-dependent cytoplasmic condensation. Recently, it was
shown that mitotic cell rounding is associated with a decrease
in cytoplasmic volume (8), a process termed premitotic condensation (PMC). In this paper, we hypothesize that CaMKII
regulates Cl⫺ efflux via ClC-3 channels, promoting mitotic cell
rounding and cell division. To test this hypothesis, we transfected D54 human glioma cells, a patient-derived glioma cell
line that is representative of glioblastoma cells (8), with GFP.
We were able to quantitatively assess changes in cell volume
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CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
and CaMKII (red) colocalization as seen in a representative
field-of-view (Fig. 2A). Digital zooms of individual cells
(boxed cells in Fig. 2A) revealed robust colocalization of ClC-3
(green) and CaMKII (red) on the plasma membrane, especially
in dividing M-phase glioma cells with condensed Hoechstlabeled DNA (blue; Fig. 2, B and C). Strong colocalization of
ClC-3 and CaMKII was also seen throughout the 10-␮m
imaging stack as seen in three-view in the x,z (top) and z,y (left)
planes (Fig. 2B). Thus ClC-3 and CaMKII are in a prime
location to interact in mitotic cells. ClC-3 is expressed on the
plasma membrane of dividing glioma cells, as previously
demonstrated by colocalization with phalloidin-labeled cortical
actin (8). Cortical actin provides a cytoskeletal framework for
the plasma membrane and colocalizes with several membranebound proteins. We labeled cortical actin with phalloidin (Fig.
2; red) and found that it colocalizes with a fraction of CaMKII
(Fig. 2; green), suggesting that CaMKII is also present at the
plasma membrane, especially in dividing cells (Fig. 2, D and
F). We then asked whether the CaMKII on the plasma membrane of dividing glioma cells was the activated form of the
kinase. For CaMKII to be active, its autoinhibitory domain
must be separated from the kinase domain. This is achieved
through binding of Ca2⫹/calmodulin to the calmodulin-binding
domain of CaMKII, allowing kinase activity (28). CaMKII can
then autophosphorylate at the Thr286 position (pCaMKII)
making the kinase active, independent of Ca2⫹/calmodulin
binding. We used an antibody specific for phosphorylated
Fig. 2. ClC-3 and CaMKII concentrate on the plasma membrane in dividing glioma cells. The first row contains representative merged images, demonstrating
robust colocalization of CaMKII with ClC-3 (A–C) and phalloidin (D–F). A: a representative field-of-view of glioma cells, with ClC-3 labeled in green, CaMKII
labeled in red, and Hoechst-labeled DNA in blue. Boxed cells are magnified in B and C. B: a representative M-phase dividing cell demonstrating robust
colocalization of ClC-3 and CaMKII, with condensed Hoechst-labeled DNA. C: a representative, interphase nondividing cell. D: a representative field-of-view
of glioma cells, with phalloidin-labeled actin in red, CaMKII labeled in green, and Hoechst-labeled DNA in blue. Boxed cells are magnified in E and F. E: a
representative M-phase dividing cell demonstrating colocalization of CaMKII and phalloidin, suggesting CaMKII expression on the plasma membrane. F: a
representative nondividing cell. All scale bars ⫽ 10 ␮m. n ⫽ 3.
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glioma cells (8). The time taken for glioma cells to decrease
volume by 50% (t50) was 28.4 ⫾ 1.6 min (Fig. 1D). Interestingly, inhibition of CaMKII with 10 ␮M AIP significantly
prolonged the time needed for PMC. AIP (10 ␮M) is specific
for CaMKII and does not block PKC, PKA, or CaMKIV (12).
When dividing glioma cells were incubated in AIP, the rate of
condensation was slower (Fig. 1, A and B). While the minimum
and maximum GFP intensities were not significantly different
(Fmin⫽ 0.4 ⫾ 0.01 for control and 0.44 ⫾ 0.02 for AIP; Fmax⫽
1.02 ⫾ 0.02 for control and 1.01 ⫾ 0.02 for AIP; Fig. 1C), the
time taken for 50% volume condensation was significantly
greater than control (39.6 ⫾ 2.6 min; P ⬍ 0.001; Fig. 1D).
Taken together, these data indicate that cytoplasmic condensation during mitotic cell rounding is facilitated by CaMKII
activity.
ClC-3 and CaMKII strongly colocalize in dividing glioma
cells. We hypothesized that CaMKII facilitated PMC by increasing ClC-3 activity. ClC-3 localizes to the plasma membrane and cytoplasm in human glioma cells (4). Given the
hypothesized activation of ClC-3 by CaMKII in dividing glioma cells, we labeled glioma cells with antibodies targeted
against residues 592– 661 of ClC-3 and residues 305– 410 of
CaMKII to determine whether these proteins were located in
the same subcellular regions in dividing cells. We also labeled
cells with Hoechst to visualize DNA, which condenses during
M-phase. Images acquired as 0.5-␮m optical sections with a
spinning disc confocal microscope demonstrated ClC-3 (green)
CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
chromatin condensation and cell morphology, as previously
described (8). We then washed on 200 ␮M NPPB, a nonspecific anion channel blocker (Fig. 4 column b), to assess NPPBsensitive Cl⫺ channel activity (Fig. 4 column a– b). Interestingly, dividing glioma cells had larger NPPB-sensitive Cl⫺
currents (Fig. 4B) as compared with nondividing control cells
(Fig. 4A). At 100 mV, the condensed, dividing cells have a
current amplitude of 19.4 ⫾ 4.4 pA/pF, which is significantly
greater than nondividing cells (2.6 ⫾ 1.7 pA/pF; P ⬍ 0.01; Fig.
4F). These NPPB-sensitive Cl⫺ currents were slightly outwardly rectifying and inactivated at depolarized potentials,
similar to Cl⫺ currents attributed to ClC-3 (32), and reversed at
⬃0 mV as predicted by symmetric Cl⫺ concentrations in the
pipette and bath solutions. However, when glioma cells were
preincubated in 10 ␮M AIP, Cl⫺ current amplitudes were back
to baseline levels in dividing (7.2 ⫾ 1.7 pA/pF; P ⬍ 0.05; Fig.
4, C and F) and nondividing cells (4.8 ⫾ 2.2 pA/pF; P ⬍ 0.01;
Fig. 4, D and F). This indicates that the larger Cl⫺ currents
seen in dividing cells are CaMKII dependent. As seen in Fig.
4E, dividing cells have approximately threefold elevated
NPPB-sensitive Cl⫺ currents, as compared with nondividing
cells, or cells with inhibited CaMKII activity (corresponding
with Fig. 4, column a– b). Thus dividing cells have enhanced
CaMKII-dependent Cl⫺ currents.
ClC-3 knockdown eliminates CaMKII-dependent Cl⫺ currents in dividing glioma cells. To determine whether CaMKII
specifically enhanced PMC (Fig. 1) and activated Cl⫺ currents
Fig. 3. Activated autophosphorylated
(p)CaMKII concentrates on the plasma membrane in dividing cells. The first row contains
representative merged images. The second row
contains the pCaMKII channel labeled in
green. The third row contains the Hoechstlabeled DNA in blue. A: a representative fieldof-view of glioma cells. Boxed cells are magnified in B and C. B: a representative M-phase
dividing cell demonstrating robust localization
of pCaMKII on the plasma membrane, with
condensed Hoechst-labeled DNA. C: a representative, interphase nondividing cell. All scale
bars ⫽ 10 ␮m. n ⫽ 3.
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Thr286 and found robust binding on the plasma membrane of
dividing glioma cells as compared with nondividing interphase
cells (green; Fig. 3). Dividing and nondividing cells expressed
relatively little CaMKII in the nucleus (Fig. 2), but this CaMKII
appears activated as it bound to pCaMKII antibody (Fig. 3).
Dividing glioma cells with condensed chromatin were round
and had enhanced pCaMKII labeling (green) throughout the
10-␮m optical section in the x,z (top) and z,y (left) planes (Fig.
3B) as compared with nondividing cells (Fig. 3C). These data
cumulatively indicate that activated CaMKII and ClC-3 colocalize in the proper cellular domains where this could facilitate
premitotic condensation.
Dividing glioma cells have large CaMKII-dependent Cl⫺
currents. Given that cytoplasmic condensation is facilitated by
CaMKII, which strongly colocalizes with ClC-3 on the plasma
membrane of dividing cells, we asked whether CaMKII modulated ClC-3 activity in mitotic cells. Mitotic cells have previously been demonstrated to have larger Cl⫺ currents, allowing for enhanced Cl⫺ efflux and obligated osmotic water
release to potentiate cytoplasmic volume condensation (9).
However, how Cl⫺ currents become activated in dividing cells
has been unknown. We performed whole cell patch-clamp
electrophysiology on human glioma cells to assess Cl⫺ channel
activity. We patched onto nondividing and dividing glioma
cells, holding the cells at ⫺40 mV and stepping from ⫺100
mV to ⫹120 mV in 20-mV increments (Fig. 4 column a).
Dividing M-phase cells were identified by Hoechst-labeled
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CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
(Fig. 4) by enhancing ClC-3 activity, we stably knocked down
endogenous ClC-3 expression in glioma cells using stably
expressed short hairpin RNA (shRNA). Glioma cells were
transfected with ClC-3 shRNA in a pGIPZ lentiviral vector.
Clonal populations were then screened for ClC-3 knockdown
with qRT-PCR. Multiple clones with multiple targeting sequences were then isolated, and ClC-3 protein expression and
activity were assessed. As seen in a representative Western
blot, ClC-3 protein expression (⬃120 and 100 kDa bands) was
robustly knocked down in cells transfected with ClC-3 shRNA
as compared with cells transfected with nontargeting shRNA
(Fig. 5A). The smear and multiple adjacent bands in the NT
shRNA lane are due to posttranslational modifications of
ClC-3. As previously reported, the multiple bands of ClC-3,
including the band at 120 kDa, correspond to multiple glycosylation states of ClC-3 (29). Therefore it appears that knockdown of ClC-3 decreased protein expression and the associated
glycosylation. Glyceraldehyde 3-phosphate dehydrogenase
(GAPDH) was used as a loading control (⬃40 kDa band). We
quantified ClC-3 protein knockdown by normalizing ClC-3
band intensity to GAPDH band intensity (relative band intensity; Fig. 5, B and C). Probing with the ClC-3 antibody from
Alomone Labs targeted to an intracellular domain demonstrated decreased band intensity to 0.2 ⫾ 0.05 as normalized to
NT shRNA (Fig. 5B). Probing with a ClC-3 antibody from
Alpha Diagnostic International targeting an extracellular domain demonstrated decreased band intensity to 0.44 ⫾ 0.09 as
compared with control (Fig. 5C).
After knocking down ClC-3 protein expression, we again
performed whole cell patch-clamp electrophysiology 1) to
determine whether the larger Cl⫺ currents in dividing cells
were mediated by ClC-3 and 2) to determine whether the
CaMKII-dependent Cl⫺ currents in dividing cells were medi-
ated by ClC-3. We patched onto ClC-3 knockdown cells and
stepped from ⫺100 mV to ⫹120 mV in 20-mV increments
(Fig. 6 column a). NPPB (200 ␮M) was then washed on to
inhibit Cl⫺ currents (Fig. 6 column b), and the NPPB-sensitive
Cl⫺ currents were subtracted out (Fig. 6 column a-b). After
knocking down ClC-3, there was no elevation of Cl⫺ currents
in dividing M-phase cells as compared with nondividing interphase cells (Fig. 6, A, B, and E). Importantly, this indicates that
the increase in Cl⫺ current activity in dividing cells (Fig. 4) is
specifically mediated by ClC-3. Additionally, CaMKII inhibition did not eliminate Cl⫺ currents after ClC-3 knockdown
(Fig. 6, C–E), indicating that CaMKII specifically acts at ClC-3
to increase Cl⫺ currents in dividing glioma cells. After ClC-3
knockdown there was no significant difference in Cl⫺ current
density at 100 mV in dividing versus nondividing cells (3.9 ⫾
1.5 pA/pF vs. 0.3 ⫾ 1.3 pA/pF, respectively) or in the presence
of 10 ␮M AIP in dividing versus nondividing cells (⫺3.0 ⫾
1.4 pA/pF vs. 0.1 ⫾ 0.9 pA/pF, respectively; Fig. 6F). These
data indicate that ClC-3 is necessary for CaMKII-activated Cl⫺
currents in dividing glioma cells.
CaMKII activation of ClC-3 facilitates premitotic condensation. After finding that CaMKII activates ClC-3 to enhance Cl⫺
currents in dividing glioma cells, we determined whether
CaMKII activation of ClC-3 also facilitates premitotic condensation. Using GFP cells transfected with NT shRNA and ClC-3
shRNA (Fig. 5), we monitored premitotic condensation associated with mitotic cell rounding. As compared with NT
shRNA-transfected cells, ClC-3 knockdown cells took significantly more time to condense cytoplasmic volume preceding
cell division (Fig. 7, A and B). Additionally, incubation of
control cells in AIP, a CaMKII inhibitor, significantly prolonged PMC to the same level as ClC-3 knockdown (Fig. 7, A
and B). Importantly, inhibition of CaMKII in ClC-3 knock-
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Fig. 4. Condensed dividing cells express
elevated CaMKII-dependent Cl⫺ currents.
Human glioma cells were held at ⫺40 mV
and stepped from ⫺100 mV to ⫹120 mV in
20-mV increments. For the representative
traces, the first column a corresponds to
whole cell currents before 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) washon. The second column b corresponds to
whole cell currents after NPPB wash-on.
The third column a– b corresponds to the
NPPB-sensitive subtracted currents. A and
B: representative traces from a nondividing
cell (A) and a dividing glioma cell with
elevated NPPB-sensitive Cl⫺ currents (B). C
and D: incubation in 10 ␮M AIP, a CaMKII
inhibitor, reduces Cl⫺ currents to baseline
levels, even in dividing cells. E: averaged
NPPB-sensitive current density corresponding to a-b. F: analysis of current density at
⫹100 mV demonstrates elevated CaMKIIdependent Cl⫺ currents in dividing cells.
Error bars indicate SE; n ⫽ 5. *P ⬍ 0.05,
Tukey-Kramer multiple-comparisons test.
CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
C533
down cells did not lead to a further prolonging of PMC (Fig. 7,
A and B), indicating that CaMKII inhibition prolongs PMC via
its interaction with CaMKII. While ClC-3 inhibition and/or
CaMKII inhibition did not lead to a significant change in Fmax,
there was a slight change in Fmin indicating that ClC-3 and
CaMKII may function in maintenance of the basal size of
glioma cells (Fig. 7C; for Fmin NT shRNA ⫽ 0.41 ⫾ 0.01, NT
shRNA ⫹ AIP ⫽ 0.36 ⫾ 0.01, ClC-3 shRNA ⫽ 0.40 ⫾ 0.01,
and ClC-3 shRNA ⫹ AIP ⫽ 0.45 ⫾ 0.01; P ⬍ 0.01). Significantly, there was an increase in the time taken for PMC upon
ClC-3 or CaMKII inhibition. The t50 increased from ⫺29.3 ⫾
0.7 min in NT shRNA cells to ⫺38.2 ⫾ 0.7 min in ClC-3
Fig. 6. ClC-3 knockdown significantly reduces CaMKII-dependent Cl⫺ currents in
dividing cells. Human glioma cells with
knocked down ClC-3 expression were held
at ⫺40 mV and stepped from ⫺100 mV to
⫹120 mV in 20-mV increments. For the representative traces, the first column a corresponds to whole cell currents before NPPB
wash-on. The second column b corresponds to
whole cell currents after NPPB wash-on. The
third column a– b corresponds to the NPPBsensitive subtracted currents. A and B: representative traces from nondividing (A) and dividing (B) glioma cells. C and D: incubation in
10 ␮M AIP, a CaMKII inhibitor, does not
further reduce Cl⫺ currents. E: averaged
NPPB-sensitive current density corresponding
to a– b. F: analysis of current density at ⫹100
mV demonstrates that ClC-3 knockdown eliminates CaMKII-dependent Cl⫺ currents in dividing cells. Error bars indicate SE; n ⫽ 5.
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Fig. 5. Endogenous ClC-3 is constitutively knocked
down in human glioma cells. A: after transfection of
human glioma cells with stably expressed short hairpin
RNA (shRNA) against ClC-3, endogenous ClC-3 expression was reduced. In this representative blot probed
with an antibody from Alomone Labs, a band at ⬃100
kDa, corresponding to ClC-3, was reduced in ClC-3
knockdown cells as compared with cells transfected
with nontargeting (NT) shRNA. GAPDH was used as a
loading control. B: quantification of ClC-3 knockdown
as probed with a ClC-3 antibody from Alomone Labs.
C: quantification of ClC-3 knockdown as probed with a
ClC-3 antibody from Alpha Diagnostic International.
Error bars indicate SE; n ⫽ 3.
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CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
knockdown cells, ⫺43.5 ⫾ 1.5 min in control cells incubated
in 10 ␮M AIP, and ⫺37.9 ⫾ 1.2 min in ClC-3 knockdown
cells incubated in AIP (Fig. 7D; P ⬍ 0.001). These data
strongly indicate that CaMKII activates ClC-3 to increase Cl⫺
currents in dividing cells and facilitate premitotic condensation.
ClC-3 and CaMKII inhibition reduces glioma cell proliferation.
Experimentation at the single-cell level demonstrates that
CaMKII activation of ClC-3 facilitates premitotic condensation. Given that CaMKII and ClC-3 inhibition impairs premitotic condensation, we hypothesized that this would also impede glioma cell proliferation. We determined population
growth by counting cells each day over a 6-day period. Cells
were cultured with vehicle, 40 ␮M NPPB to block Cl⫺ channels, 2.5 ␮M KN-93 to block CaMKII, or 40 ␮M NPPB and
2.5 ␮M KN-93. KN-93, a small-molecule inhibitor, was used
instead of AIP, which is a peptide sensitive to degradation
during long incubations at 37°C. Cl⫺ channel inhibition decreased glioma cell number at day 6 to 38.9 ⫾ 5.34 cells as
compared with 62.75 ⫾ 4.67 cells in the control condition.
CaMKII inhibition reduced the number of cells at day 6 to
36.24 ⫾ 5.68, and simultaneous Cl⫺ channel and CaMKII
inhibition reduced the number of cells to 12.8 ⫾ 1.12 (Fig. 8A;
P ⬍ 0.05). Thus, Cl⫺ channels and CaMKII appear to facilitate
glioma cell proliferation. Simultaneous Cl⫺ channel and CaMKII
inhibition decreased proliferation by day 6 to numbers below
that of Cl⫺ channel blockade or CaMKII blockade alone (Fig.
8A; P ⬍ 0.05). This may be due to CaMKII regulation of cell
division independent of ClC-3. CaMKII inhibits p21, leading
to p53 degradation and increased proliferation (17). Increases
in intracellular calcium concentration ([Ca2⫹]i) that activate
CaMKII can also lead to phosphorylation of CDC25C, which
in turn dephosphorylates CDC2 to trigger entry into mitosis
(23, 33). Thus CaMKII may have pleiotropic effects on cell
cycle progression independent of Cl⫺ channels.
Additionally, NPPB is not a specific inhibitor of ClC-3;
NPPB blocks intermediate-conductance K⫹ channels, which
can also regulate proliferation (4a, 37). Therefore, we also
studied the role of ClC-3 in glioma cell proliferation using a
genetic approach (Fig. 5). At day 6, ClC-3 knockdown decreased proliferation from 53.59 ⫾ 3.96 cells in control conditions to 34.46 ⫾ 4.85 cells (Fig. 8B). Additionally, CaMKII
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Fig. 7. ClC-3 knockdown prolongs PMC
to the same extent as CaMKII inhibition.
A: representative dividing cells transfected
with NT or ClC-3 shRNA and incubated in
vehicle or 10 ␮M AIP, a CaMKII inhibitor.
ClC-3 and CaMKII inhibition increases the
time needed for PMC to the same extent and
is not additive. Time was normalized to t ⫽
0, which is the last time point preceding cell
division. Scale bar ⫽ 10 ␮m. B: ClC-3 or
CaMKII inhibition increases the total time
needed for PMC. C: Fmax does not significantly change with CaMKII inhibition, while
Fmin does slightly, but significantly, change.
D: ClC-3 and CaMKII inhibition increases
the time needed for 50% volume condensation (t50). Error bars indicate SE; n ⫽ 22– 67.
*P ⬍ 0.01, Tukey-Kramer multiple-comparisons test.
CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
C535
inhibition with 2.5 ␮M KN-93 significantly decreased glioma
cell proliferation to the same levels as ClC-3 inhibition at day
6 (Fig. 8B; P ⬍ 0.01). This provides further indication that
CaMKII facilitates proliferation by activating ClC-3.
We also measured proliferation by pulsing GFP-expressing
glioma cells with BrdU, a synthetic analog of thymidine, for 30
h. We then fixed the cells (Fig. 8C; green) and probed for BrdU
incorporation (Fig. 8C; blue). In control glioma cells, nearly all
glioma cells labeled for BrdU, indicating that almost all cells
had replicated DNA at least once during the 30-h BrdU pulse
period (Fig. 8C; NT shRNA arrows). However, upon ClC-3
knockdown, there was a significant increase in the number of
cells not incorporating BrdU during the 30-h pulse period (Fig.
8C; ClC-3 shRNA arrows). The number of cells not incorporating BrdU increased from 4.0 ⫾ 1.9% in control conditions
to 20.1 ⫾ 6.2% in ClC-3 knockdown cells (Fig. 8D; P ⬍ 0.05).
However, CaMKII inhibition with 2.5 ␮M KN-93 did not
significantly increases the percentage of nonproliferating
BrdU-negative cells, even after ClC-3 knockdown (Fig. 8D;
NT shRNA ⫹ KN-93 ⫽ 5.8 ⫾ 2.2% and ClC-3 shRNA ⫹
KN-93 ⫽ 17.5 ⫾ 3.8%; P ⬎ 0.05), indicating that the decreases in cell number due to CaMKII inhibition (Fig. 8, A and
B) may partially be attributed to ClC-3-independent regulation.
Interestingly, most of the nonproliferating cells induced by
ClC-3 knockdown appeared larger and flatter than dividing
cells (Fig. 8C). This provides further evidence that ClC-3,
which is activated by CaMKII, facilitates premitotic condensation and glioma cell division.
DISCUSSION
Here we demonstrate that CaMKII activation of ClC-3
facilitates premitotic condensation in human glioma cells.
ClC-3 and CaMKII robustly colocalize on the plasma membrane
of dividing cells, and specifically, auto-activated CaMKII appears
more abundant in M-phase cells versus interphase cells. Inhi-
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Fig. 8. ClC-3 and CaMKII inhibition reduce glioma cell proliferation. A: 40 ␮M NPPB inhibition of Cl⫺ channels and 2.5 ␮M KN-93 inhibition of CaMKII
reduce glioma cell proliferation over 6 days. n ⫽ 4. B: transfection of cells with ClC-3 shRNA and 2.5 ␮M KN-93 inhibition of CaMKII reduce glioma cell
proliferation over 6 days. n ⫽ 4. C and D: ClC-3 knockdown significantly increases the number of cells not incorporating bromodeoxyuridine (BrdU) after a
30-h BrdU pulse period. Error bars indicate SE. Scale bar ⫽ 10 ␮m. *P ⬍ 0.05; Tukey-Kramer multiple-comparisons test and unpaired t-test. Arrows indicate
NT shRNA cells incorporating BrdU and ClC-3 shRNA cells not incorporating BrdU.
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CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
accelerate PMC in glioma cells, it is likely that alternate
channels fluxing osmotically active ions may facilitate PMC
and cell division in other cell types. Additionally, while we
found that inhibition of CaMKII activation of ClC-3 impedes PMC, cells (though fewer in numbers, see Fig. 8)
nevertheless condensed and progressed through M-phase.
This indicates that PMC is integral to cell division, and
dividing cells have redundant mechanisms to ensure PMC
occurs.
Volume condensation in dividing cells may allow for the
bridging of kinases to their substrate proteins. As seen in
Fig. 2, while ClC-3 and CaMKII do colocalize in nondividing cells, the colocalization is enhanced in condensed dividing cells. Thus, condensation of the cytoplasm may
increase macromolecular crowding, leading to enhanced
interaction between kinases and their substrate proteins.
Changes in cellular volume can lead to shifts in actin
polymerization/depolymerization or protein regulation, including activation of membrane ion channels (15). Several
kinases become activated upon cellular condensation, including protein kinase C (16), which can then phosphorylate
and activate the Na⫹/H⫹ exchanger in tumor cells (24).
Additionally, Ca2⫹/calmodulin-dependent protein kinases
phosphorylate NKCC, a Na⫹-K⫹-Cl⫺ cotransporter, enhancing activity after cell shrinkage (22). Thus PMC, by
mechanically shrinking the cytoplasmic space, may increase
macromolecular crowding and interaction of kinases with
their substrate proteins. We found a significant increase in
the number of cells that did not incorporate BrdU and
undergo mitosis after ClC-3 knockdown (Fig. 8). These
nondividing cells were typically larger and flatter than
dividing cells and did not undergo PMC after a 30-h period.
This is in agreement with data from rat C6 glioma cells
demonstrating that there are low and high cell size checkpoints for proliferation (31). Therefore, cells that cannot
undergo PMC, such as after ClC-3 knockdown, may be
senescent and larger. In a number of other cell types,
including fibroblasts, senescent cells are larger and flatter as
compared with dividing cells (1). An understanding of
whether these cells lack mechanisms for mitotic cell rounding and PMC could yield further insight into cytoskeletal
and hydrodynamic changes associated with cell division.
Therapeutically targeting CaMKII activation of ClC-3 in
human glioma. Identifying the mechanisms by which human
glioma cells divide may lead to the development of novel therapeutic targets. Interference with CaMKII activation of ClC-3 in
human glioma cells decreases glioma cell proliferation (Fig. 8), so
further investigation of these proteins may lead to the development of novel therapeutics. Directly inhibiting Cl⫺ channel activity in glioma cells is a promising therapeutic strategy. This has
been achieved with chlorotoxin, a toxin in the venom of Leiurus
quinquestriatus (deathstalker scorpion), which recently completed
Phase II trials for the treatment of glioblastoma multiforme.
Preclinical studies demonstrate that chlorotoxin inhibits Cl⫺ channel activity in glioma cells (21), leading to impaired glioma cell
migration (34). In Phase I studies, chlorotoxin demonstrated
exquisite specificity, binding glioma tissue after intracavitary
injection into patients with recurrent glioma (20). By inhibiting
Cl⫺ channel activity, chlorotoxin decreases glioma cell migration,
and our data suggest that it may also be effective at inhibiting
PMC and glioma cell proliferation.
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bition of CaMKII prolonged cellular condensation preceding
cell division, but only when ClC-3 was expressed. Importantly,
the condensation of glioma cells was associated with enhanced
Cl⫺ currents mediated by CaMKII activation of ClC-3. These
Cl⫺ currents are associated with shape and volume changes in
glioma cells (26) and facilitate cytoplasmic volume decrease
during PMC (8).
A link between mitotic cell rounding and premitotic
condensation. An elegant study recently demonstrated that
mitotic cell rounding is driven by two opposing forces: an
inward contraction of the actomyosin cytoskeleton versus an
outwardly directed osmotic pressure (35). Disruption of either
process perturbed the rounding ability of mitotic cells (35).
Data presented here indicate that CaMKII activation of ClC-3
may be a source for this outward osmotic pressure, at least in
glioma cells. Activation of ClC-3 by CaMKII facilitates cytoplasmic condensation during mitotic cell rounding (Figs. 1 and
7) and is associated with larger Cl⫺ currents (Figs. 4 and 6). As
osmotically active Cl⫺ exits the cell via ClC-3, water is drawn
out of the cell, leading to cytoplasmic volume condensation
(7). Glioma cells accumulate [Cl⫺]i to 100 mM through Na⫹K⫹-Cl⫺ cotransporter 1 (NKCC1) activity, creating an outwardly directed gradient for Cl⫺ movement (6). This gradient
can be used to drive water out of the cell, creating an outward
osmotic pressure and facilitating PMC (7). While we did not
assess the contribution of cytoskeleton contraction to PMC in
glioma cells, our data indicate that PMC is facilitated by efflux of
osmotically active ions in glioma cells, and through a similar
mechanism, different channels/transporters that flux osmotically
active ions may be of importance in other proliferative cells.
Actomyosin contraction and hydrodynamic volume changes
may work in concert to enhance macromolecular crowding.
Considerable attention has been given to the role of cytoskeletal dynamics during mitotic cell rounding. Actomyosin
contraction is essential to generate an inward pressure to
balance the outward osmotic pressure generated in rounding
mitotic cells (35). Additionally, RhoA, a small GTPase,
activates Rho kinase, which phosphorylates myosin II regulatory light chain leading to cortical rigidity and retraction
in mitotic cells (19). Independent of myosin II activity,
moesin proteins link the actin cortex to the plasma membrane, inducing cell rounding and membrane rigidity in
mitotic cells (14). Several other actin-binding proteins such
as L-caldesmon, actin interacting protein 1 (AIP1), and
spectrin also play an integral role in actin dynamics in
dividing cells (5). The importance of mitotic cell rounding
may lie in its ability to form a symmetric mitotic spindle,
assuring that both daughter cells receive the correct set of
chromosomes with high fidelity (30). While these studies
elucidate a mechanism through which cells can attain a
spherical morphology via actomyosin contraction, it does
not account for intracellular contents: do cells maintain the
same cytosolic composition, or do cells modify this intracellular milieu? Here we find that glioma cells condense
cytoplasmic volume by extruding Cl⫺ ions, allowing for the
osmotic release of water and cellular volume decrease.
Thus, given that ⬃70% of a cell is water (18), channels
extruding osmotically active ions contribute to volume condensation and may work in concert with actomyosin contraction to control PMC during mitotic cell rounding. While
the interaction between ClC-3 and CaMKII appears to
CaMKII ACTIVATES ClC-3 IN DIVIDING CELLS
GRANTS
The authors are grateful for the following grants from the National Institutes
of Health (National Institute of Neurological Disorders and Stroke): RO1
NS-31234, RO1 NS-52634, and RO1 NS-36692 (to H. Sontheimer) and F31
NS-73181 (to V. A. Cuddapah).
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the author(s).
AUTHOR CONTRIBUTIONS
Author contributions: V.A.C., C.W.H., and H.S. conception and design of
the research; V.A.C., C.W.H., S.W., L.S.M., and T.-T.C.B. performed the
experiments; V.A.C., C.W.H., S.W., L.S.M., and T.-T.C.B. analyzed the data;
V.A.C., C.W.H., S.W., L.S.M., T.-T.C.B., and H.S. interpreted the results of
the experiments; V.A.C. and L.S.M. prepared the figures; V.A.C. drafted the
manuscript; V.A.C. and H.S. edited and revised the manuscript; V.A.C. and
H.S. approved the final version of the manuscript.
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We demonstrate here that ClC-3 activity in glioma proliferation
is regulated by CaMKII. CaMKII is a Ca2⫹-sensitive kinase;
therefore, CaMKII activation of ClC-3 can be inhibited by identifying upstream Ca2⫹ sources for CaMKII activation. One candidate expressed by gliomas is mutant epidermal growth factor
(EGF) receptor (EGFR). The EGFRvIII mutation, caused by a
deletion of exons 2–7 of the EGFR gene, is the most common and
leads to constitutive EGFR signaling, enhancing proliferation
(10). Given that, in epithelial cells, EGF stimulation leads to
increase in [Ca2⫹]i and subsequent CaMKII activation (36),
EGFRvIII mutations in gliomas may drive constitutive CaMKII
activation. Additionally, transient receptor potential (TRP) canonical 1 (TRPC1) channels play an important role in Ca2⫹ dynamics
and glioma cell division, as inhibition of TRPC1 in glioma cells
impedes store-operated Ca2⫹ entry and proliferation (2). Other
TRP channels, including TRPC3 and TRP vanilloid type 1
(TRPV1), increase [Ca2⫹]i to enhance CaMKII activity (3, 13), so
an understanding of whether TRPC1-mediated Ca2⫹ influx activates ClC-3 via CaMKII could lead to novel mechanistic insight.
During mitosis, several cell types transiently increase [Ca2⫹]i,
including Pt K2 epithelial cells, which spike [Ca2⫹]i during
metaphase for about 20 s (27). Clearly, future studies should
investigate the upstream Ca2⫹ signaling that regulates the CaMKII-ClC-3 interaction during mitosis. These studies may reveal
novel therapeutic targets to improve clinical management of
gliomas.
C537
C538
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