Download Nanosecond electric pulses trigger actin responses in plant cells

Biochemical and Biophysical Research Communications 387 (2009) 590–595
Contents lists available at ScienceDirect
Biochemical and Biophysical Research Communications
journal homepage: www.elsevier.com/locate/ybbrc
Nanosecond electric pulses trigger actin responses in plant cells
Thomas Berghöfer a, Christian Eing a, Bianca Flickinger a, Petra Hohenberger b, Lars H. Wegner a,b,
Wolfgang Frey a, Peter Nick b,*
a
b
Institute for Pulsed Power and Microwave Technology (IHM), Forschungszentrum Karlsruhe GmbH, Karlsruhe Institute of Technology, 76344 Eggenstein-Leopoldshafen, Germany
Botanical Institute I, University of Karlsruhe, Karlsruhe Institute of Technology, Kaiserstr. 2, 76128 Karlsruhe, Germany
a r t i c l e
i n f o
Article history:
Received 9 July 2009
Available online 18 July 2009
Keywords:
Actin
Endoplasmic reticulum
Microtubules
Nanosecond pulsed electrical fields
Nuclear envelope
Tobacco BY-2 cells
a b s t r a c t
We have analyzed the cellular effects of nanosecond pulsed electrical fields on plant cells using fluorescently tagged marker lines in the tobacco cell line BY-2 and confocal laser scanning microscopy. We
observe a disintegration of the cytoskeleton in the cell cortex, followed by contraction of actin filaments
towards the nucleus, and disintegration of the nuclear envelope. These responses are accompanied by
irreversible permeabilization of the plasma membrane manifest as uptake of Trypan Blue. By pretreatment with the actin-stabilizing drug phalloidin, the detachment of transvacuolar actin from the cell
periphery can be suppressed, and this treatment can also suppress the irreversible perforation of the
plasma membrane. We discuss these findings in terms of a model, where nanosecond pulsed electric
fields trigger actin responses that are key events in the plant-specific form of programmed cell death.
Ó 2009 Elsevier Inc. All rights reserved.
Introduction
The transport of biomolecules across the plasma membrane
poses a major challenge to numerous biological applications. Electroporation is now widely used as a central technique to overcome
this barrier, either in order to introduce large molecules into living
cells (for reviews see [1–3]) or to extract cell ingredients [4]. In classical electroporation, the duration of pulsed electric field exposure is
in the range of micro- to even milliseconds, electric field amplitude
is low and this is predicted to create irreversible membrane permeabilization, that allows for bulk flow of macromolecules (for a recent
review see [5]). However, when the cells are exposed to ultra-short
pulses, i.e. electric pulses of high field strength (up to 300 kV cmÿ1),
low risetime (considerably shorter than the time constant for membrane charging) and at the same time very short pulse duration in
the nanosecond range (down to several nanoseconds), the electric
field can penetrate into the cell interior before it is dissipated into
the plasma membrane by subsequent membrane charging due to
charge redistribution at the interphases between membrane and
exterior and the cytoplasmic interface, respectively [6]. This allows
to manipulate intracellular targets, such as membranes of organelles. The biological effects of these nanosecond pulsed electric fields
(nsPEF) differ from those induced by classical electroporation and
Abbreviations: ER, endoplasmic reticulum; CaMV-35S, Cauliflower Mosaic Virus
35S promotor; GFP, green fluorescent protein; nsPEF, nanosecond pulsed electrical
fields.
* Corresponding author. Fax: +49 721 608 4193.
E-mail address: [email protected] (P. Nick).
0006-291X/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved.
doi:10.1016/j.bbrc.2009.07.072
have attracted considerable interest, because they can induce specific cellular responses such as apoptosis [7] that have already been
employed for tumour therapy [8].
Due to this large impact, it is necessary to understand the
underlying cellular and molecular mechanisms. Plasma membrane
charging during exposure of Jurkat cells to nsPEF was demonstrated [9], but experimental data demonstrating pore formation
in the sub-microsecond range as predicted from theoretical studies
of the process [10–12] are scarce.
Moreover, the correlation between membrane pores and the
observed biological processes does not necessarily mean that the
two are physically linked. For instance, intranuclear mRNA-processing complexes have been shown to be disrupted [13]. Interaction of the electric field with these processes is expected to involve
numerous and complex intermediates.
To get further insight into the cellular events, we tested the response of plant cells to nsPEF-treatment. The biology of plant cells
differs specifically from mammalian cells with respect to the organization, topology, and dynamics of internal membranes (for a
recent review see [14]), structure, function, and molecular composition of the nuclear envelope (for a recent review see [15]), and
the mechanisms driving programmed cell death (for a review see
[16]). By making use of the evolutionary divergence between plant
and mammalian cells, the comparison between the responses of
these cells to nsPEFs should therefore help to understand the biological events and, in the long term, to dissect the underlying
molecular mechanisms responsible for the apoptotic response.
As a model, we chose the widely used tobacco cell line Bright
Yellow 2 (BY-2), which has been an excellent model for molecular
T. Berghöfer et al. / Biochemical and Biophysical Research Communications 387 (2009) 590–595
plant cell biology (for review see [17]). Transgenic derivatives of
this cell line expressing fusions of the green fluorescent protein
(GFP) with various organelle markers allow to visualize specific
organelles in living cells. We observed such marker lines for actin
filaments, microtubules, and endoplasmic reticulum/nuclear envelope with confocal laser scanning microscopy, and were able to follow the early cellular events triggered by nsPEFs in plant cells and
to define, for the first time, a specific response of actin filaments.
Materials and methods
Cell material and cultivation. The tobacco (Nicotiana tabacum
L. cv BY-2) cell line [17] was cultivated in liquid medium containing 4.3 g/L Murashige and Skoog salts (Duchefa), 30 g/L Suc,
200 mg/L KH2PO4, 100 mg/L inositol, 1 mg/L thiamine, and
0.2 mg/L 2,4-D, pH 5.8. Cells were subcultured weekly, inoculating
1.5–2 mL of stationary cells into 30 mL of fresh medium in 100-mL
Erlenmeyer flasks. The cell suspensions were incubated at 25 °C in
the dark on an orbital shaker (KS250 basic, IKA Labortechnik) at
150 rpm. Stock BY-2 calli were maintained on media solidified
with 0.8% (w/v) agar and subcultured monthly. Transgenic cells
and calli were maintained on the same media supplemented with
30 mg/L hygromycin. In some experiments, the cell lines were assessed in the absence of selective pressure but without any differences in patterning. All experiments were performed using cells
after 4 day subcultivation. In addition to the non-transformed
BY-2 wild type, transgenic lines were used that expressed GFP in
fusion with markers for actin (actin-binding domain of plant fimbrin [18], tubulin (tobacco tubulin a3 [19]), and the endoplasmic
reticulum (HDEL-GFP, [20]).
nsPEF-treatment and confocal microscopy. To ensure the appropriate conductivity of the cell suspension for the nsPEF-treatment, the
cells were sedimented and the supernatant culture medium
replaced by an equal volume of charging buffer (125 mM KCl,
5 mM CaCl22H2O, 5 mM MgCl26H2O, 150 mM sorbitol, and 1 mM
Tris, pH 7.2). For each experiment, 30 ll of the suspension were
591
transferred between the two titanium electrodes of the microelectrode array (Fig. 1A) that was fixed on a standard microscope slide
and covered with a coverslip. The gap distance between the electrodes was 300 lm with an electrode height of 100 lm. After filling,
the microelectrode array containing the cells was fixed to a receptacle mounted on the linear motion stage of the microscope. The voltage applied to the electrodes was delivered by a square pulse
generator with a risetime of 800 ps, a pulse length of 10 ns and a
maximum output voltage of 1 kV (IPG 2501, HILO-Test, Karlsruhe,
Germany). To monitor the impulse voltage waveforms, a 600 MHz
oscilloscope (WaveRunner 64Xi, LeCroy Corporation, Chestnut
Ridge, NY, USA) in combination with a Philips probe (PM 8931, Philips, The Netherlands) was used. Fig. 1A shows a scheme of the
experimental set-up and a typical output. For all experiments, the
maximum output voltage of the square pulse generator (1 kV) has
been used, resulting in an effective electric field strength of 33 kV/
cm. If not stated otherwise, a single pulse was applied, and the response of the cells to nsPEF was followed by confocal laser scanning
microscopy (TCS SP1, Leica, Bensheim, Germany). Z-stacks were recorded with step-sizes of around 1 lm using a dual wavelength
(ArKr-laser lines 488 and 564 nm, beam splitters LP515 or DD, emission filter LP 515 or OG590, respectively) configuration and a 4frame averaging algorithm. In some experiments, the cells were pretreated with phalloidin (Fluka, Buchs; Switzerland) diluted from an
ethanolic stock solution of 1 mM. To assess membrane permeability,
the cells were mixed with an equal volume of 2.5% w/v Trypan Blue
(Sigma), and then immediately loaded to the gap and subjected to
the pulse treatment.
Results
Response of microtubules to nsPEF
To follow their response to intense electric fields we used a
tobacco BY-2 cell line expressing the tobacco tubulin a3 in fusion
with GFP under control of the CaMV-35S promotor. Upon nsPEF-
Fig. 1. Experimental set-up. (A) Schematic representation of the technical set-up and a typical profile of a voltage pulse applied to the microelectrodes. (B) Topology of BY-2
tobacco cells.
592
T. Berghöfer et al. / Biochemical and Biophysical Research Communications 387 (2009) 590–595
treatment with 1 pulse at 33 kV cmÿ1 we observed that the cortical
microtubules became disordered and were progressively redistributed into mesh-like structures that probably corresponded to cytoplasmic strands (Fig. 2A). This response became detectable from
1 min after the pulse and was fully manifest from 3 min after pulsing. In the cell interior, microtubules surround the nuclear envelope that is ellipsoidal in shape – upon pulsing the nucleus
rounded up rapidly within the first minute after the exposition
(Fig. 2B). At later stages, this could be accompanied by blebbing
of the nuclear envelope (Fig. 2C, white arrow).
Response of the endoplasmic reticulum
The endoplasmic reticulum (ER) is organized in different subdomains that differ with respect to their structural dynamics, visualized by using a cell line expressing the ER-signal peptide HDEL in
fusion with GFP under control of the CaMV-35S promotor
(Fig. 3). Whereas nuclear envelope and transvacuolar ER-strands
are relatively stable, the cortical ER-meshwork below the plasma
membrane is extremely dynamic (Supplementary Movie S1). Upon
nsPEF exposure to 1 pulse of 33 kV cmÿ1 we observed that the nuclear envelope disintegrated such that the GFP signal invaded the
karyoplasm. This response was detectable from 3 to 4 min after
pulsing, i.e. roughly parallel with the blebbing of the nuclear envelope observed in the microtubule marker line.
Response of the actin filaments
The actin cytoskeleton consists of a meshwork in the cell cortex
adjacent to the plasma membrane, and of transvacuolar bundles
that emanate from the nuclear envelope in the cell center that
serve to move (and later to tether) the nucleus to a defined position
prior to mitosis. Both populations were followed using a tobacco
Fig. 3. Response of the endoplasmic reticulum to an electric field pulse of
33 kV cmÿ1 in tobacco BY-2 cells expressing the peptide HDEL in fusion with GFP.
Note the disintegration of the nuclear envelope and the penetration of fluorescence
into the nuclear interior. The images show individual confocal sections of time
series.
BY-2 line, where the actin-binding domain of plant fimbrin was
fused to GFP and driven by the CaMV-35S promotor. The response
of these two major populations of actin to nsPEF exposure (1 pulse
of 33 kV cmÿ1) differs qualitatively: whereas the actin meshwork
Fig. 2. Response of microtubules to an electric field pulse with an amplitude of 33 kV cmÿ1 in BY-2 cells expressing tobacco tubulin a3 in fusion with GFP. (A) Disintegration
and disordering of cortical microtubules. (B) Rounding up of the nuclear rim indicated by the white arrows. (C) Blebbing of the nuclear rim indicated by the white arrows. The
images show individual confocal sections of time series.
T. Berghöfer et al. / Biochemical and Biophysical Research Communications 387 (2009) 590–595
in the cell cortex (Fig. 4, upper row) appears to dissolve, the actin
bundles in the cell center seem to detach from the periphery and to
contract upon the nucleus (Fig. 4A, lower row, Fig. 4B). During later
stages (Fig. 4, lower row, 16 min), this contraction can even detach
the plasma membrane from the cell wall resembling the morphogenetic changes observed in response to plasmolysis. Both actin responses, the cortical dissolution, and the central contraction, are
already manifest within minutes after pulsing with the observation
being limited by the scanning speed of the confocal microscope. To
understand the role of actin disassembly in this response, we pretreated the cells with the actin-stabilizing drug phalloidin. Based
on preliminary studies the phalloidin treatment had been adjusted
to the threshold of visible effects (30 min, 1 lM phalloidin, data
not shown). With this pretreatment, the detachment of actin from
the periphery and the contraction towards the nucleus could be
suppressed.
Slow responses of the membrane permeability
In order to test, whether the electric pulses will lead to an irreversible disintegration of the membrane, we measured the dose–
response for the uptake of Trypan Blue, a dye that cannot permeate the membrane of living cells and is conventionally used for
viability assays. The dose was varied by increasing the number
of pulses of 33 kV cmÿ1. We observed that uptake became detect-
593
able from two pulses (Fig. 4D) and was conspicuous at 10 pulses
(Fig. 4E). Interestingly, a pretreatment with phalloidin (30 min,
1 lM phalloidin) reduced the uptake to a basal level (Fig. 4E,
compare closed versus open squares). Phalloidin treatment without pulse treatment did not alter the uptake of the dye (data
not shown).
Summarizing our results, we observe a rapid disintegration of
cortical actin filaments and cortical microtubules, a rapid detachment of transvacuolar actin strands from the periphery and their
contraction towards the nucleus. These early responses are followed by a slower disintegration of the nuclear envelope (manifest
by blebbing of the microtubular perinuclear rim, and by invasion of
the HDEL-GFP signal into the karyoplasm), and by detachment of
the plasma membrane from the cell wall. These responses are
accompanied by irreversible perforation of the plasma membrane
manifest as uptake of Trypan Blue. By pretreatment with the actin-stabilizing drug phalloidin the detachment of transvacuolar actin from the cell periphery can be suppressed, and this treatment
can also suppress the irreversible perforation of the plasma
membrane.
Discussion
The biological effects of these nanosecond pulsed electric fields
(nsPEF) are of considerable interest, even beyond cell biology, be-
Fig. 4. Response of actin filaments to an electric field pulse of 33 kV cmÿ1 in tobacco BY-2 cells expressing the actin-binding domain of fimbrin in fusion with GFP. (A) Time
series of individual confocal planes in the cortex (upper row) or the center (lower row). (B) Rapid contraction of transvacuolar actin bundles towards the nucleus in control
cell in response to 1 pulse of 33 kV cmÿ1. (C) Reduced response of actin to 1 pulse with an amplitude of 33 kV cmÿ1 after pretreatment with 1 lM of phalloidin over 30 min.
(D) Uptake of Trypan Blue in response to 2 pulses of 33 kV cmÿ1 in tobacco BY-2 cells (time interval 15 min after pulsing). (E) Dose-dependency of Trypan Blue uptake over
number of pulses of 33 kV cmÿ1 in BY-2 (time interval 15 min after pulsing). Open squares: without phalloidin treatment, closed squares: after pretreatment with 1 lM of
phalloidin over 30 min. The data give mean values and standard errors from 52 to 147 individual cells, respectively. (F) Working hypothesis on the cellular responses to
nsPEF-exposition. Explanations in the text.
594
T. Berghöfer et al. / Biochemical and Biophysical Research Communications 387 (2009) 590–595
cause they have been shown to induce cellular responses of medical relevance such as apoptosis [7]. The underlying mechanisms
are far from being understood, though. In the present work, we
have analyzed the response of plant cells to nsPEFs. The comparison of the evolutionary divergent plant and mammalian cells
might help to get insight into the mechanisms underlying the
apoptotic response to nsPEFs. We observe a specific response of
actin filaments, and we further observe that stabilization of actin
by phalloidin can rescue the membrane barrier evident from a
reduced uptake of Trypan Blue. The observed loss of nuclear shape
in response to nsPEFs can also be attributed to the response of
actin. Unlike to its mammalian counterpart, the shape of plant
nuclei is not maintained by an integral lamine network, but by a
cytoskeletal lattice that can even tunnel through the nucleus
[21]. The position of the nucleus is actively controlled during the
cell cycle by both microtubules and actin (for a recent review see
[22]), and this involves the activity of plant-specific KCH-kinesins
that connect to microtubules as well as to actin filaments [23].
Thus, the nucleus is tethered to its position and kept in shape by
balanced tension between the different flanks of the cell. When
the interaction of actin with the plasma membrane and the cell
wall is disrupted by nsPEF, this will result in contraction of actin
filaments towards the nucleus and a loss of nuclear shape (Fig. 4F).
Whereas the bundling of actin and the morphogenetic change of
the nucleus can be explained by cytoskeletal tensegrity, the relationship between actin and membrane permeability (assessed as
uptake of Trypan Blue) is not evident at first glance. However, it
should be noted that in the set-up of this study, the uptake of
the dye did not measure reversible, rapid pore formation during
the nsPEF, but was rather a manifestation of irreversible pores that
persist after the end of the stimulus. Bundling of actin filaments
has been shown to be an evolutionary conserved central element
of apoptosis and programmed cell death that has been observed
in mammalian, as well as in yeast and plant cells (for review see
[24]). The molecular players transmitting the actin signal towards
the apoptotic machinery are far from being understood and seem
to differ partially between the kingdoms. However, a direct interaction of actin filaments with membrane topology has been demonstrated in both directions. For instance, phosphorylation of a
myosin light chain has been found to be necessary and sufficient
for actin-dependent apoptotic membrane blebbing [25]. Conversely, the actin-binding N-WASP/WIP complex can transmit a
signal from membrane curving upon the assembly of F-actin
[26]. In plants, treatment with RGD-peptides (that cannot permeate the membrane, but disrupt the interaction of the cytoskeleton
with the plasma membrane through an unknown integrin-like protein) caused the disruption of cytoplasmic architecture and inhibited deplasmolysis indicating a loss of membrane integrity [27]. In
fact, the so called Hechtian strands, attachment sites of actin filaments at the plasma membrane that become visible during plasmolysis are also disrupted by RGD-peptides [28]. This indicates
that the plasma membrane is not only structured by lipids and
intrinsic membrane proteins, but also by interaction with the adjacent cortical actin filaments (Fig. 4F, right-hand panel). Current
models on irreversible permeabilisation of the plasma membrane
[10–12] have made important contributions to our understanding
of the biological effect of nsPEFs, but our findings suggest that
these models have to be extended by the contribution of the cytoskeleton to membrane permeability. When the interactions between cytoskeleton and plasma membrane are interrupted, as to
be expected during the nsPEF, this will affect the structural integrity of the plasma membrane resulting in irreversible formation of
pores evident from the increased uptake of Trypan Blue. Stabilization of actin by phalloidin would on the one hand improve
membrane-anchoring of microfilaments, and therefore render
structural integrity more resilient to the challenges posed by the
nsPEF. Future work will be dedicated to investigate the role of actin
for membrane identity, and to identify the molecular target within
Hechtian strands that are affected by nsPEFs.
Acknowledgments
This work was supported by the Excellency Programme of the
German Research Council to the University of Karlsruhe (Shared
Research Group 60-1). We acknowledge the following colleagues
for providing BY-2 marker cell lines to this project: Dr. Jan Maisch,
Karlsruhe (FABD2-GFP), Dr. Hasezawa, Tokyo (TuA3-GFP), and Dr.
Jan Petrášek, Prague (HDEL-GFP).
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.bbrc.2009.07.072.
References
[1] J. Teissie, M. Golzio, M.P. Rols, Mechanisms of cell membrane
electropermeabilization: a minireview of our present (lack of ?) knowledge,
BBA 1724 (2005) 270–280.
[2] J.C. Weaver, Y.A. Chizmadzhev, Electroporation, in: C. Polk, E. Postow (Eds.),
Handbook of Biological Effects of Electromagnetic Fields, second ed., CRC Press,
Boca Raton, FL, 1996, pp. 247–274.
[3] U. Zimmermann, G.A. Neil, Electromanipulation of Cells, CRC Press, Boca Raton,
FL, 1996.
[4] M. Sack, C. Schultheiss, H. Bluhm, Triggered Marx generators for the industrialscale electroporation of sugar beets. Industry Applications, IEEE Trans. 41
(2005) 707–714.
[5] C. Chen, S.W. Smye, M.P. Robinson, J.A. Evans, Membrane electroporation
theories: a review, Med. Biol. Eng. Comput. 44 (2006) 5–14.
[6] K.H. Schoenbach, R.P. Joshi, J. Kolb, N. Chen, M. Stacey, P. Blackmore, E.S.
Buescher, S.J. Beebe, Ultrashort electrical pulses open a new gateway into
biological cells, Proc. IEEE 92 (2004) 1122–1137.
[7] S.J. Beebe, P.M. Fox, L.J. Rec, E.L. Willis, K.H. Schoenbach, Nanosecond, highintensity pulsed electric fields induce apoptosis in human cells, FASEB J. 17
(2003) 1493–1495.
[8] R. Nuccitelli, U. Pliquett, X. Chen, W. Ford, J.R. Swanson, S.J. Beebe, J.F. Kolb, K.H.
Schoenbach, Nanosecond pulsed electric fields cause melanomas to selfdestruct, Biochem. Biophys. Res. Commun. 343 (2006) 351–360.
[9] W. Frey, J.A. White, R.O. Price, P.F. Blackmore, R.P. Joshi, R. Nuccitelli, S.J. Beebe,
K.H. Schoenbach, J.F. Kolb, Plasma membrane voltage changes during
nanosecond pulsed electric field exposure, Biophys. J. 90 (2006) 3608–3615.
[10] Z. Vasilkosky, A.T. Esser, T.R. Gowrishankar, J.C. Weaver, Membrane
electroporation: the absolute rate equation and nanosecond time scale pore
creation, Phys. Rev. E 74 (2006) 1–12.
[11] W. Krassowska, P.D. Filev, Modeling electroporation in a single cell, Biophys. J.
92 (2007) 404–417.
[12] K.C. Smith, J.C. Weaver, Active mechanisms are needed to describe cell
responses to submicrosecond, megavolt-per-meter pulses: cell models for
ultrashort pulses, Biophys. J. 95 (2008) 1547–1563.
[13] N. Chen, L.A. Garner, G. Chen, Y. Jing, Y. Deng, R.J. Swanson, J.F. Kolb, S.J. Beebe,
R.P. Joshi, K.H. Schoenbach, Nanosecond electric pulses penetrate the nucleus
and enhance speckle formation, Biochem. Biophys. Res. Commun. 364 (2007)
220–225.
[14] A.Y. Cheung, S.C. De Vries, Membrane trafficking: intracellular highways and
country roads, Plant Physiol. 147 (2008) 1451–1453.
[15] A.C. Schmit, P. Nick, Microtubules and the evolution of mitosis, in: P. Nick
(Ed.), Plant Microtubules – Development and Flexibility, Springer, Berlin,
Heidelberg, New York, 2008, pp. 233–266.
[16] A.M. Jones, Programmed cell death in development and defense, Plant Physiol.
125 (2001) 94–97.
[17] T. Nagata, Y. Nemoto, S. Hasezawa, Tobacco BY-2 cell line as the ‘Hela’ cell in
the cell biology of higher plants, Int. Rev. Cytol. 132 (1992) 1–30.
[18] J. Maisch, J. Fišerová, L. Fischer, P. Nick, Actin-related protein 3 labels actinnucleating sites in tobacco BY-2 cells, J. Exp. Bot. 60 (2009) 603–614.
[19] F. Kumagai, A. Yoneda, T. Tomida, T. Sano, T. Nagata, S. Hasezawa, Fate of
nascent microtubules organized at the M/G1 interphase, as visualized by
synchronized tobacco BY-2 cells stably expressing GFP-tubulin: timesequence observations of the reorganization of cortical microtubules in
living plant cells, Plant Cell Physiol. 42 (2001) 723–732.
[20] J. Petrášek, A. Černá, K. Schwarzerová, M. Elčkner, D.A. Morris, E. Zažímalová,
Do phytotropins inhibit auxin efflux by impairing vesicle traffic?, Plant Physiol
131 (2003) 254–263.
[21] D.A. Collings, C.N. Carter, J.C. Rink, A.C. Scott, S.E. Wyatt, N.S. Allen, Plant nuclei
can contain extensive grooves and invaginations, Plant Cell 12 (2000) 2425–
2439.
T. Berghöfer et al. / Biochemical and Biophysical Research Communications 387 (2009) 590–595
[22] P. Nick, Control of cell axis, in: P. Nick (Ed.), Plant Microtubules – Development
and Flexibility, Springer, Berlin, Heidelberg, New York, 2008, pp. 3–46.
[23] N. Frey, J. Klotz, P. Nick, Dynamic bridges – a calponin-domain kinesin from
rice links actin filaments and microtubules in both cycling and non-cycling
cells. Plant Cell Physiol., 2009 (published online June 27, 2009).
[24] V.E. Franklin-Tong, C.W. Goutay, A role for actin in regulating apoptosis/
programmed cell death: evidence spanning yeast, plants and animals,
Biochem. J. 413 (2008) 389–404.
[25] M.L. Coleman, E.A. Sahai, M. Yeo, M. Bosch, A. Dewar, M.F. Olson, Membrane
blebbing during apoptosis results from caspase-mediated activation of ROCK I,
Nat. Cell Biol. 3 (2001) 339–345.
595
[26] K. Takano, K. Toyooka, Sh. Suetsugu, EFC/F-BAR proteins and the N-WASP–WIP
complex induce membrane curvature-dependent actin polymerization, EMBO
J. 27 (2008) 2817–2828.
[27] J. Zhou, B. Wang, Y. Li, Y. Wang, L. Zhu, Responses of Chrysanthemum
cells to mechanical stimulation require intact microtubules and plasma
membrane-cell wall adhesion, J. Plant Growth Regul. 26 (2007)
55–68.
[28] H. Canut, A. Carrasco, J.P. Galaud, C. Cassan, H. Bouyssou, N. Vita, P.
Ferrara, R. Pont-Lezica, High affinity RGD-binding sites at the plasma
membrane of Arabidopsis thaliana link the cell wall, Plant J. 16 (1998)
63–71.