Download Distinct fluorescent pattern of KAT1::GFP in the plasma membrane of

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European Journal of Cell Biology 86 (2007) 489–500
www.elsevier.de/ejcb
Distinct fluorescent pattern of KAT1::GFP in the plasma membrane of
Vicia faba guard cells
Ulrike Homann1, Tobias Meckel1,2, Jennifer Hewing, Marc-Thorsten Hütt3,
Annette C. Hurst
Institute of Botany, University of Technology Darmstadt, Schnittspahnstrasse 3-5, 64287 Darmstadt, Germany
Received 18 May 2006; received in revised form 15 May 2007; accepted 15 May 2007
Abstract
The organisation of membrane proteins into certain domains of the plasma membrane (PM) has been proposed to
be important for signalling in yeast and animal cells. Here we describe the formation of a very distinct pattern of the
K+ channel KAT1 fused to the green fluorescent protein (KAT1::GFP) when transiently expressed in guard cells of
Vicia faba. Using confocal laser scanning microscopy we observed a radially striped pattern of KAT1::GFP
fluorescence in the PM in about 70% of all transfected guard cells. This characteristic pattern was found to be cell type
and protein specific and independent of the stomatal aperture and the cytoskeleton. Staining of the cell wall of guard
cells with Calcofluor White revealed a great similarity between the arrangement of cellulose microfibrils and the
KAT1::GFP pattern. Furthermore, the radial pattern of KAT1::GFP immediately disappeared when turgor pressure
was strongly decreased by changing from hypotonic to hypertonic conditions. The pattern reappeared within 15 min
upon reestablishment of high turgor pressure in hypotonic solution. Evaluation of the staining pattern by a
mathematical algorithm further confirmed this reversible abolishment of the radial pattern during hypertonic
treatment. We therefore conclude that the radial organisation of KAT1::GFP depends on the close contact between the
PM and cell wall in turgid guard cells. These results offer the first indication for a role of the cell wall in the localisation
of ion channels. We propose a model in which KAT1 is located in the cellulose fibrils intermediate areas of the PM and
discuss the physiological role of this phenomenon.
r 2007 Elsevier GmbH. All rights reserved.
Keywords: Potassium inward rectifier; KAT1; Cell wall; Cellulose fibres; Cytoskeleton; Latrunculin B; Propyzamide; Turgor
pressure; Membrane domains
Corresponding author. School of Biomedical Sciences, University
of Queensland, St. Lucia, QLD 4072, Australia. Tel.: +61 7 3346 1224;
fax: +61 7 3365 1766.
E-mail address: [email protected] (A.C. Hurst).
1
These authors contributed equally to the work.
2
Present address: Institute of Physics, Leiden University, Niels
Bohrweg 2, 2333 CA Leiden, The Netherlands.
3
Present address: School of Engineering and Science, Jacobs
University Bremen, Campus Ring 1, D-28759 Bremen, Germany.
0171-9335/$ - see front matter r 2007 Elsevier GmbH. All rights reserved.
doi:10.1016/j.ejcb.2007.05.003
Introduction
Structural conditions which cause a heterogeneous
distribution of membrane proteins are believed to be
important factors for the regulation of numerous
signalling and transport events, especially at the plasma
membrane (PM). So far a number of different mechanisms that induce a heterogeneous distribution of
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U. Homann et al. / European Journal of Cell Biology 86 (2007) 489–500
membrane proteins have been described mainly in
mammalian cell lines and yeast.
These mechanisms include the lateral separation of
specific membrane lipid species (i.e. mainly cholesterol
and glycosphingolipids) which leads to the formation of
specialised microdomains, so-called lipid rafts. Certain
proteins accumulate in these microdomains (e.g. glycosyl
phosphatidyl inositol (GPI)-anchored proteins) while
others are not affected. This accumulation can be
explained by a slowdown of protein mobility by a factor
of 2 (Dietrich et al., 2002) which results in a
heterogeneous distribution of proteins in the PM. Many
proteins involved in signalling cascades are found in lipid
rafts (Brown and Rose, 1992). This provided the first hint
on the physiological significance of microdomain formation. The slowdown and accumulation of proteins in lipid
rafts is believed to increase the probability of dimer,
multimer and cluster formation which is important for
many signalling events at the PM. Nevertheless, the exact
role of lipid rafts in cellular signalling, trafficking, and
structure has yet to be determined.
Lipid rafts and raft-associated proteins have also been
identified in plants (Borner et al., 2005; Mongrand et al.,
2004). However, like for lipid raft formations in
mammalian and yeast cells, the physiological meaning
of these microdomains remains to be confirmed.
A second factor that was found to limit free diffusion
of membrane proteins is the cytoskeleton. It can
determine the localisation of PM proteins via direct
attachment to the protein or indirectly. A direct connection to the cytoskeleton is for example of particular
importance for the localisation and functioning of the
cellulose synthase complex in the PM of plant cells
(Gardiner et al., 2003). In the case of certain mechanosensitive ion channels the attachment to the cytoskeleton is also proposed to function as a signalling
component in mammalian cells (Barritt and Rychkov,
2005; Ghazi et al., 1998). Apart from the direct
connection to PM proteins it is believed that the actin
cytoskeleton can confine the movement of proteins with
enlarged cytosolic domains by generating ‘‘fenced’’
microenvironments without the direct attachment to
the diffusing components (Ritchie and Kusumi, 2004).
While such a mechanism has so far not been described in
plant cells, a cortical cytoskeleton—the only determinant of this effect—is also present in plant cells (Staiger
and Lloyd, 1991).
The third factor that can contribute to heterogeneous
distribution of PM proteins is the extracellular matrix
(ECM). It is able to directly influence the distribution of
proteins in the PM of eukaryotic cells (Arnold et al.,
2004). For plant cell walls, which can be viewed as the
plant ECM, a role in distribution of PM proteins has
not yet been described.
PM ion channels which play an important role
in signal transduction have been implicated to be
distributed non-homogenously in the PM of plant and
animal cells (Tester, 1990; Deutsch, 2002). In animal
cells lipid rafts have been described as an important
factor for the localisation of ion channels in certain
domains in the PM (Martens et al., 2004). For plant ion
channels mechanisms which determine their localisation
in microdomains have not been identified.
Recently Sutter et al. (2006) demonstrated that the
K+ inward rectifier KAT1 from Arabidopsis thaliana is
localised in clusters in the PM when transiently
expressed in tobacco epidermal cells. The authors
detected KAT1 protein in a ‘moderately’ detergentresistant fraction, indicating its association with lipid
rafts. The KAT1 cluster showed nearly no lateral
mobility. Investigations of the role of SNAREs (soluble
NSF [N-ethylmaleimide-sensitive factor] attachment
protein receptors) on trafficking of KAT1 indicated
that SNAREs are involved in cluster formation and
mobility of KAT1 (Sutter et al., 2006). However, the
mechanism which anchors KAT1 in the PM remains to
be determined.
KAT1 plays an important role in guard cell functioning. We therefore analysed turgid guard cells transiently
expressing KAT1 fused to green fluorescent protein
(GFP). KAT1::GFP was organised in clusters in the PM
similar to what we previously described for guard cell
protoplasts (Hurst et al., 2004). In addition we found a
radial distribution of KAT1::GFP clusters which was
dependent on a close contact between the PM and the
cell wall. In animal cells contacts of the PM with the
ECM are mediated by substrate adhesion molecules
such as fibronectin, vitronectin, collagen, and others, via
the short amino acid sequence Arg-Gly-Asp (RGD)
(D’Souza et al., 1991) that interacts with integrins. The
integrins in turn link the ECM to the cytoskeleton
(Ruoslahti, 1996). For plant cell walls plant biologists
are just beginning to understand how cell wall-tomembrane interactions are established to acquire celland tissue-specific characters and how this affects cell
function and polarity and cell-to-cell interactions. So
far, in plants only few homologues of classical adhesion
molecules, e.g. b-integrin or fibronectin that revealed a
RGD-mediated membrane matrix adhesion have been
identified (Canut et al., 1998; Faik et al., 1998; Gens
et al., 1996; Pellenc et al., 2004). This points to a similar
interaction between the PM and the ECM or cell wall of
animal and plant cells, respectively. In addition a
number of plant PM proteins have been proposed to
directly bind to both the PM and extracellular carbohydrates and may thus anchor the cell to the cell wall (for
review see (Kohorn, 2000)). Among these is the cellulose
synthase complex (Kohorn, 2000), which also requires
cortical microtubule arrays for normal localisation in
the PM (Gardiner et al., 2003). This indicates that a
continuous cytoplasm–cell wall scaffold is essential to
control key events in plant development and growth.
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Our data clearly demonstrate the crucial role of the
guard cell wall in the pattern formation of KAT1. We
propose a model in which KAT1 clusters are located in
the intermediate areas of the cellulose microfibrils and
discuss the physiological role of such a localisation.
Materials and methods
Construction/design of the KAT1::GFP fusion
protein
The kat1 cDNA was amplified by PCR for directed
cloning into pAVA393 (NcoI restriction site) in frame
with the mGFP5 (Haseloff et al., 1997) for expression of
the fusion protein KAT1::GFP under the control of two
P35S promoters, as described in detail by Hurst et al.
(2004). The plasmid was then cloned in Escherichia coli/
DH5a, followed by preparation of plasmid DNA
(Qiagen high speed Midi-Kit, Qiagen, Germany). The
purified vector was used for ballistic bombardment of
guard cells and epidermal cells as described by Hurst et
al. (2004).
The construct GFP::TM23 was kindly provided by
Nadine Paris (University of Rouen, France). It represents the transmembrane domain of human LAMP1
(lysosome-associated membrane protein-1) fused to the
green fluorescent protein in the vector pSGFP6 K
(Brandizzi et al., 2002).
The P-ATPase PMA4 from Nicotiana plumbaginifolia
fused to GFP in the vector pTZ19U-gfp (Lefebvre et al.,
2004) was a kind gift from M. Boutry, (Université
Catholique de Louvain, Belgium). The talin::YFP
construct (Kost et al., 1998) was kindly provided by B.
Kost (University of Heidelberg, Germany), and the
plasmid with TOR1::GFP (Buschmann et al., 2004) by
A. R. Schäffner (GSF Research Center, Germany),
respectively. All fusion constructs were expressed under
P35S.
Transfection of guard cells
Guard cells were transfected via ballistic bombardment as described by Hurst et al. (2004). Briefly, whole
leaves of Vicia faba L. cv. Bunyan were placed upside
down on solid Murashige Skoog Medium and bombarded with 2 mg gold (1 mm particle diameter) coated
with 10 mg DNA according to the manufacturer’s
instructions (BioRad, Munich, Germany), at a pressure
of 650 Psi, a distance of 6 cm, and a vacuum of 25 in Hg.
Confocal microscopy
Confocal microscopic analysis of transfected guard
cells was carried out 16 to 24 h after ballistic bombard-
491
ment as described by Meckel et al. (2004). Briefly,
abaxial epidermal peels from bombarded leaves were
placed in small dishes in a standard buffer solution
consisting of 10 mM MES-KOH (pH 6.1), 45 mM KCl,
and 100 mM CaCl2. Analysis was carried out using a
confocal laser scanning microscope (Leica TCS SP,
Leica Microsystems GmbH, Heidelberg, Germany),
equipped with a 63 water immersion objective (plan
apo, N.A. 1.2). For excitation of mGFP5, the 488-nm
line of a 25-mW Ar/Kr-Ion-Laser was used; emission
was detected at 505–535 nm.The confocal aperture was
adjusted to give optical sections with a full-width at halfmaximum of around 0.68 mm. Images were processed
using the Leica Confocal Software 2.00 (LCS, Leica
Microsystems GmbH, Heidelberg, Germany).
Analysis of guard cells stained with Calcofluor White
For staining of cellulose fibrils Calcofluor White
(Sigma, Munich, Germany) was added to epidermal
peels bathed in standard buffer to a final concentration
of 0.1% (w/v). Epidermal peels were incubated in
Calcofluor White for 5 min and washed 5 times with
standard buffer. Epidermal peels were analysed using
the confocal laser scanning microscope Leica TCS SP2
AOBS (Leica Microsystems GmbH, Heidelberg,
Germany) with a 63 water immersion objective. For
excitation the 405-nm line of a 50-mW Ar-UV-Laser
was used, emission was detected at 415–440 nm.
Cytoskeleton inhibitors
Latrunculin B (Calbiochem, Darmstadt, Germany)
and propyzamide (Sigma, Munich, Germany) were
prepared in dimethyl sulphoxide (DMSO) as 25 and
50 mM stocks, respectively. The toxins were diluted with
standard buffer solution to a final concentration of
10 mM latrunculin B and 50 mM propyzamide. The final
concentration of DMSO was 0.04% (v/v) and 0.05%
(v/v), respectively.
Hyper- and hypotonic treatment of guard cells
Epidermal peels were placed in standard buffer (see
above) which was hypotonic (100 mosmol/kg), ensuring
that cells were fully turgescent. For hypertonic treatment of cells the standard buffer plus sorbitol
(1000 mosmol/kg) was added until plasmolysis was
visible just at the tips of guard cells. The final osmolarity
of the bath solution was between 500 and 800 mosmol/
kg. To restore turgor pressure hypotonic conditions
were induced by replacing the bath solution with
standard buffer.
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Determination of radial distribution versus random
distribution via mathematical methods
The algorithmic challenge in quantifying the degree
of radial organisation in these fluorescence distribution patterns is due to three features of the
data: (i) inspection by eye shows that the degree
of radial organisation varies gradually between the
image sets, (ii) noise content (i.e. randomly excited
pixels in the fluorescence images) varies strongly
within the data and (iii) the image sizes are too
small for an informative spectral analysis. These
circumstances are calling for a quantification of the
pattern to attach a reproducible value to each image
which is independent of the observer. Our method
exploits the fact that the radially organised patterns
and the random spots transform differently under an
Ising algorithm (i.e. a well-defined domain-forming
dynamical process). For all analysis steps we use a
binarised fluorescence image, obtained by substituting
the value of each image point by 1, if it exceeds a
certain threshold and 0 otherwise. In all cases we have
chosen the average fluorescence (without background)
as binarisation threshold. The Ising algorithm is an
early thermodynamic model of ferromagnetism. In
the original model of a ferromagnet the implementation of this algorithm leads to domain formation.
Here it emphasises existing radial patterns. The resulting transformed image is then evaluated using a
standard method from information theory, namely the
mutual information. This process assigns a number
(the information content of the image, as given by
the mutual information) to this transformed image.
The value is related to the radial organisation of
the pattern, due to the different transformation properties of radial stripes and random dots. While
this procedure is sufficient to assess the organisational features of a pattern at fixed noise intensity
of the image, the varying noise content of the data
requires an additional analysis step. We estimate
the noise content of each (binarised) image with the
help of a melting algorithm. The application of
this algorithm gives the percentage of image points
which are more likely to be noise rather than
signal. Consequently, application of our algorithms
to a given fluorescence image yields two numbers,
namely the noise content of the original (binarised)
image and the information content of the
Ising-transformed image as a measure of stripe
formation.
We calibrated the full algorithm on simulated data
where the stripe-to-dot ratio and the noise content have
been varied systematically. This led to the reference
curves shown in Fig. 7G. A more detailed description
and application of the Ising-algorithm in this context is
given by Hütt (2001).
Results
Radial distribution of KAT1::GFP in the PM of
guard cells
Figs. 1A–C show intact turgid guard cells of V. faba
transfected with the K+ channel KAT1 fused to GFP
(KAT1::GFP). The expression of KAT1::GFP resulted
in a distinct staining pattern of the PM which in many
cells appeared as stripes originating from the dorsal side
of the guard cells and radially centred towards their
ventral side (Fig. 1A). For further analysis we categorised the stripe formation as follows: guard cells
showing a clear radial pattern over the whole PM were
classified as type 3 (Fig. 1A); guard cells with less
prominent stripe formation only visible in part of the
PM or displaying shorter stripes not forming a
continuous line from the dorsal to ventral side were
classified as type 2 (Fig. 1B); guard cells without any
clear stripe formation with KAT1::GFP appearing in
large randomly distributed dots in the PM were
classified as type 1 (Fig. 1C). According to this
classification 36% out of the 157 guard cells analysed
were classified as type 3 and 32% each as type 2 and type
1. In addition to staining of the PM, guard cells often
exhibited labelling of intracellular compartments
(mainly ER and nuclear envelope). This most likely
resulted from protein expression under the strong 35S
promoter which can lead to accumulation of the fusion
protein in the ER (Hawes et al., 2001).
A
B
C
D
Fig. 1. Distribution of KAT1::GFP in guard cells and
epidermal cells. Maximum projection of cells expressing
KAT1::GFP (green). (A) Guard cell of stomata displaying a
clear radial staining pattern (type 3). (B) Guard cell of stomata
displaying radial staining pattern only in part of the cell
(type 2). (C) Guard cell displaying random staining pattern
(type 1). (D) Epidermal cell displaying random staining
pattern. Scale bars: 10 mm.
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Distribution pattern of KAT1::GFP is cell type and
protein specific
In addition to guard cells we also analysed the
distribution of KAT1::GFP fluorescence in transfected epidermal cells. Epidermal cells expressing the
KAT1::GFP fusion protein displayed a strikingly
different staining pattern compared to guard cells.
The stripe formation typical of guard cells was never
observed in epidermal cells (n4 50). All epidermal cells
displayed random distribution of KAT1::GFP fluorescence in large dots in the PM (Fig. 1D). This implicates
that the radial distribution of KAT1::GFP is cell type
specific. To examine whether PM proteins in guard cells
are generally distributed in a radial pattern we analysed
the expression of two different PM proteins fused to
GFP. These proteins included the 23 amino acid long
single-pass membrane domain from human LAMP1
(lysosome-associated membrane protein-1; GFP::TM23;
Brandizzi et al., 2002) and the PM ATPase from
Nicotiana plumbaginifolia (PMA4::GFP; Lefebvre
et al., 2004). Fig. 2 shows typical expression patterns
from guard cells transfected with these fusion constructs. Both fusion proteins tested displayed an even
distribution of the fluorescence in the PM without any
formation of radial stripes.
Together these results implicate that the radial
distribution of KAT1::GFP in the PM of transfected
guard cells is cell type and protein specific.
493
and microtubules depolymerise. The distribution pattern of KAT1::GFP in transfected guard cells is at first
glance similar to the distribution of actin filaments and
microtubules in open guard cells indicating that the
cytoskeleton may be involved in the distribution of
KAT1::GFP. This would implicate that the radial
staining pattern can only be found in guard cells of
open stomata.
However, analysis of the relationship between stomatal aperture and stripe formation revealed no correlation
between the type of KAT1::GFP pattern and opening of
the stomatal pore (Fig. 3). Fig. 3 shows the distribution
of the stomatal aperture from type 1, type 2 and type 3
cells. Stomatal aperture was determined by measuring
the inner width of the stomatal pore from maximum
projections of 157 guard cells. All distributions were
fitted with a Gaussian function displaying a peak value
at 9.7, 9.8 and 10.4 mm for type 1, type 2 and type 3,
respectively. Kolmogorov–Smirnov two-sample tests
revealed no significant difference between the three
distributions (a ¼ 0.05). This suggests that the distribution pattern is unrelated to the stomatal aperture and
thus independent of the cytoskeleton.
To further investigate the participation of the
cytoskeleton in the formation of the radial distribution of KAT1::GFP, we applied latrunculin B
type 1
15
type 2
Distribution pattern of KAT1::GFP in guard cells is
independent of stomatal aperture and cytoskeleton
inhibitors
type 3
A
Frequency
10
Investigations of the cytoskeleton in guard cells of
open stomata revealed that both actin filaments and
microtubules are radially distributed (Fukuda et al.,
1998; Kim et al., 1995). In closed stomata the radial
distribution pattern disappears because actin filaments
5
B
0
0
Fig. 2. Distribution of different GFP fusion proteins in guard
cells. (A) Maximum projection of guard cell expressing
GFP::TM23. (B) Maximum projection of guard cell expressing
PMA4::GFP. Scale bars: 10 mm.
4
8
12
Aperture [µm]
16
20
Fig. 3. Stripe formation is independent of stomatal aperture.
Distribution of stomatal aperture of guard cells displaying
either clear radial staining pattern (type 3), radial staining in
parts of cell (type 2) or no radial staining (type 1). Stripe
formation was categorised according to text (see also Fig. 1).
The distributions were each fit to a Gaussian function. The
peak values were 9.7, 9.8 and 10.4 mm for type 1, type 2 and
type 3, respectively. Distributions were not significantly
different (a ¼ 0.05, Kolmogorov–Smirnov two-sample test).
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U. Homann et al. / European Journal of Cell Biology 86 (2007) 489–500
A
C
B
D
Fig. 4. Filamentous structures of actin and microtubules are
destroyed by latrunculin B and propyzamide. (A) Actin
cytoskeleton in guard cells visualised by talin::YFP. (B) Same
guard cell as in (A) after 20 min incubation in 10 mM
latrunculin B. (C) Microtubules in guard cell visualised by
TOR1::GFP. (D) Same guard cell as in (C) after 15 min
incubation in 50 mM propyzamide. Note the complete disappearance of smaller filamentous structures. Scale bars: 10 mm.
(final concentration: 10 mM), an actin-depolymerising
agent and propyzamide (final concentration: 50 mM) as a
microtubule-destabilising drug. The effect of these
inhibitors on the cytoskeleton was tested in guard cells
transfected with talin::YFP (Kost et al., 1998) or
TOR1::GFP (Buschmann et al., 2004) to visualise the
distribution of the actin or microtubules, respectively.
Incubation of guard cells in10 mM latrunculin B for
20 min was sufficient to completely destroy the actin
meshwork (Figs. 4A and B). The radial distribution of
the microtubule marker significantly changed already
after 10 min in 50 mM propyzamide with complete
disappearance of smaller filamentous structures
(Figs. 4C and D). This demonstrates that the arrangement of the cytoskeleton in guard cells is indeed
destroyed by the inhibitors applied.
However, neither the application of latrunculin B nor
the incubation in propyzamide had an appreciable effect
on the radial distribution of KAT1::GFP. The striped
pattern remained unchanged even after prolonged
treatment for 70 and 140 min in propyzamide or
latrunculin B, respectively (Figs. 5B and D). Together
these results demonstrate that the cytoskeleton is not
involved in maintenance of the radial distribution of
KAT1::GFP.
Possible involvement of the cell wall in the radial
distribution of KAT1::GFP
In guard cells not only actin filaments and microtubules but also cellulose fibrils are radially distributed
(Raschke, 1979). Fig. 5 shows Calcofluor White staining
of cellulose fibrils in the cell wall of guard cells and
epidermal cells. Guard cells displayed a parallel and
radial organisation of cellulose fibrils (Fig. 6A). In
epidermal cells such a pattern could not be observed,
A
B
C
D
Fig. 5. Distribution of KAT1::GFP in guard cells is not
affected by cytoskeleton inhibitors. Maximum projection of
guard cells expressing KAT1::GFP. (A) and (C) Guard cells
expressing KAT1::GFP before addition of cytoskeleton
inhibitors. (B) KAT1::GFP-transfected guard cell 70 min after
addition of 50 mM propyzamide. (D) KAT1::GFP-transfected
guard cell 140 min after addition of 10 mM latrunculin B. Scale
bar: 10 mm.
and cellulose fibrils were found to be arranged in a
disordered manner (Fig. 6B). The arrangement of the
cellulose fibrils in guard cells shows the same radial
orientation as KAT1::GFP stripes in the PM, indicating
a causal relation between the two staining patterns.
Further indications for the participation of the cell
wall in formation of the radial distribution of
KAT1::GFP can be derived from images of transfected
guard cell protoplast. In contrast to turgid guard cells
protoplasts lacked the radial fluorescence pattern in the
PM (Hurst et al., 2004).
To further investigate the participation of the cell wall
in the distribution of KAT1::GFP, we incubated
epidermal peels in strong hypertonic solution to
decrease turgor pressure of guard cells. Under this
condition the PM is no longer tightly pressed against the
cell wall. In order to avoid infolding of the PM we
increased the osmolarity of the bath solution until the
PM started to retract from the cell wall only at the cell
tips. At this stage there was no visible detachment of the
PM from the cell wall in the rest of the cell. The changes
in staining pattern from three guard cells before and
after hypertonic treatment are displayed in Figs. 7A–F.
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A
B
495
A
C
E
B
D
F
C
Fig. 6. Radial distribution of cellulose fibrils in guard cells.
Confocal images of cells stained with Calcofluor White.
(A) Partial view from a maximum projection of four
consecutive stacks of a guard cell showing radial distribution
of cellulose fibrils. (B) Partial view from a guard cell (left) and
epidermal cell (right) showing radial and random distribution
of cellulose fibrils, respectively. (C) Overview of the epidermis
utilizing Normarski optics. The selected area corresponds to
image (B). Scale bars: 10 mm.
Hypertonic treatment resulted in an immediate loss of
the radial pattern in the PM (Figs. 7B, D and F).
To quantitatively analyse the systematic features of
the distribution patterns in guard cells we have
developed an image analysis algorithm (see Materials
and methods). The radial organisation seems to exist in
gradually varying degrees, ranging from cases, where it
is a very prominent feature of the image, to cases, where
it is almost completely masked by other features and
hard to distinguish from randomly distributed fluorescent spots. This fact particularly emphasises the need for
a quantitative algorithmic look at these patterns. This
algorithm allows distinguishing cells with a radial
staining from cells with a more random distribution of
KAT1::GFP. Results from analysis of the cells shown in
Figs. 7A–F are represented in Fig. 7G (circles marked
with characters). The reference grid in Fig. 7G has been
obtained by analysing simulated data, where noise
content and the stripe-to-dot ratio have been varied
(see Materials and methods). Note that the length scales
(i.e. the ‘‘thickness’’ of typical stripes) determines the
absolute position of this grid. This additional dependence on other (but constant) features than stripe-todot-ratio and noise has two important consequences for
our interpretation of the data: (1) for a given experimental data set, individual reference grids have to be
simulated and (2) only the relative position in the plane
information content of transformed image
G
0.4
E
(1)
A
0.3
C
B
F
0.2
(2)
D
0.1
0.15
noise intensity
0.2
0.25
Fig. 7. Loss of radial pattern of KAT1::GFP after hypertonic
treatment. (A), (C) and (E) Partial view from maximum
projections of four consecutive stacks of guard cells expressing
KAT1::GFP before hypertonic treatment. (B), (D) and (F)
Partial view from guard cells shown in (A), (C) and (E),
respectively, 3 to 8 min after hypertonic treatment. (G) Result
from mathematical analysis of images shown in (A)–(F);
reference grid has been obtained by analysing simulated data;
values above and below dashed reference line correspond to
stripe and random formation, respectively, of KAT1::GFP;
(1) and (2) indicate increasing noise and dottiness, respectively,
in reference data. Scale bar: 10 mm
for images within a single experiment can be interpreted.
Thus, for the further examples given in Fig. 8 only the
relevant region of the plane is shown without an
additional reference grid.
The guard cells shown in Figs. 7A, C and E are well
above the reference line in this quantitative analysis and,
therefore, represent stripe formation whereas Figs. 7B,
D and F are clearly below this reference line and,
therefore, lack this feature of radial organisation. This
quantitative analysis confirms that loss of turgor is
associated with a decrease in the degree of stripe-like
distribution of KAT1::GFP.
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U. Homann et al. / European Journal of Cell Biology 86 (2007) 489–500
A
B
to the green fluorescent protein (GFP) in V. faba guard
cells, and observed a cell type- and protein-specific
fluorescent pattern. In most guard cells KAT1::GFP was
organised in distinct radial stripes in the PM. Our results
lead to the hypothesis that this staining pattern is due to
interaction of KAT1 with the cell wall of guard cells.
C
D
Distinct KAT1::GFP pattern in the PM of guard
cells does not refer to overexpression artefact
information content
of transformed image
0.45
A
0.4
0.35
C
0.3
B
0.15
0.2
0.25
noise intensity
Fig. 8. Recurrence of radial pattern of KAT1::GFP upon
hypotonic treatment. (A) Partial view from maximum projections of four consecutive stacks of guard cell expressing
KAT1::GFP before hypertonic treatment. (B) Partial view of
guard cell shown in (A) 4 min after hypertonic treatment.
(C) Partial view of guard cell shown in (A) and (B) 15 min after
hypotonic treatment. (D) Result from mathematical analysis
of images shown in (A)–(C); lower information content
corresponds to loss of ordered stripe formation of
KAT1::GFP. Scale bar: 10 mm.
To test for the reversibility of the loss in stripe
formation we replaced the hypertonic solution by the
hypotonic standard bath solution. Figs. 8A–C show part
of the cortical section of a KAT1::GFP-transfected
guard cell before and after hypertonic treatment, and
15 min after replacing the hypertonic solution by the
standard bath solution. The radial fluorescent pattern
which vanished upon loss of turgor (Fig. 8B) started to
recur when high turgor pressure was restored in
standard solution (Fig. 8C). Quantitative analysis of
the images confirmed that the decrease in radial
organisation observed during hypertonic treatment is
reversible (Fig. 8D).
Together the results demonstrate that the formation
of the radial fluorescence distribution in guard cells is
associated with a close contact of the PM to the
surrounding cell wall.
Discussion
In this study we heterologously expressed the K+
inward rectifier KAT1 from Arabidopsis thaliana fused
Ever since GFP in fusion with functional proteins was
applied in plant cell biology, artefacts resulting from
overexpression have been a critical point to take into
account while interpreting such data. This is especially
true for the expression of constructs under a strong
promoter like the 35S promoter used in our studies. At
first sight the observed KAT1::GFP distribution may
thus be explained by a non-physiological association of
fusion proteins into clusters mediated by GFP. However, our results from two other GFP fusion proteins
expressed in guard cells under the same strong promoter
contradict this hypothesis. When fused to GFP neither
the single-pass transmembrane domain TM23 from
LAMP1 nor the functional protein PM-ATPase showed
this striped fluorescent pattern in transfected guard cells.
In addition KAT1::GFP was always found in an evenly
dotted distribution in transfected epidermal cells. GFP
fluorescence intensity per PM area did not differ with
cell type or expressed fusion protein. Hence, varying
protein amounts cannot account for the different
expression patterns observed (for the quantification
procedure see Meckel et al., 2007). We therefore
conclude that the expression pattern of KAT1::GFP is
cell type and protein specific. In addition, guard cell
protoplasts transfected with KAT1::GFP never showed
a striped pattern of KAT1::GFP. Taken together the
results demonstrate that the striped orientation of
KAT1::GFP is clearly not the result of an overexpression artefact, but rather points to a distinct mechanism
that determines the distribution of KAT1 in the PM of
turgid guard cells.
Stomatal aperture and the cytoskeleton are not
involved in pattern formation
From animal cells, a PM-cytoskeleton-ECM continuum is proposed in which various membrane proteins
are functionally included (Gumbiner, 1996). The role of
the cytoskeleton in guard cell function is somewhat
controversial. Marcus et al. (2001) demonstrated a
diurnal cycle in microtubule formation in guard cells
that was dependent on stomata aperture. Furthermore
this cycle could be blocked by application of propyzamide, an inhibitor of microtubule formation (Marcus
et al., 2001). In contrast, Assmann and Baskin (1998)
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found no role for microtubules in guard cell functioning.
Also actin filaments seem to have a role in guard cell
function. Their distribution changes from a more radial
pattern in open stomata to a less organised distribution
in closed stomata (Hwang et al., 1997). Hwang et al.
(1997) also propose that actin is involved in the
regulation of K+ channels in guard cells. Despite
the striking similarity of the KAT1::GFP distribution
to the arrangement of the cytoskeleton (microfilaments
and microtubules) observed in this study, we found no
evidence for the involvement of the cytoskeleton in the
distribution of KAT1. The staining pattern was not
dependent on the stomatal aperture. Additionally
neither the application of propyzamide as an inhibitor
of microtubule formation nor latrunculin B as an
inhibitor of microfilament formation could abolish the
radial pattern of KAT1::GFP. The concentrations of
latrunculin B and propyzamide used in our study were
in the same range of what has been shown to destroy
microfilaments in inner cortex cells of maize root apices
(Baluska et al., 2004) and cortical microtubules in
A. thaliana (Naoi and Hashimoto, 2004), respectively.
Furthermore, sequence predictions from KAT1 do not
reveal an ankyrin motif which would allow direct
binding of actin to KAT1 (Nakamura et al., 1995). We
therefore conclude that neither microtubules nor actin
microfilaments are direct participants in the formation
of the striped pattern of KAT1::GFP in the PM of
guard cells.
Pattern formation is related to a KAT1–cell wall
interaction
Our analysis of guard cells suggests that a tight
contact between the PM and the cell wall is essential for
the striped distribution of KAT1. When turgid guard
cells were incubated in strong hypertonic solution
resulting in a loss of turgor and consequently the PM
being no longer pressed against the cell wall the radial
fluorescent pattern disappeared immediately and dissolved into randomly distributed dots. Restoration of
the turgor pressure by exchange of the hypertonic
solution with hypotonic bath solution led to the
reappearance of the radial pattern. Using a mathematical algorithm the pattern of fluorescence before and
after loss of turgor could clearly be separated by ‘‘grade
of their homogeneity’’ (with the fluorescence being more
ordered before loss of turgor independent of noise and
background). Together with the observation that
KAT1::GFP-expressing guard cell protoplasts displayed
no radial staining pattern (Fig. 1B; Hurst et al., 2004)
this demonstrates that the formation of stripes depends
on a close contact of KAT1 with the cell wall. The radial
arrangement of KAT1 is very similar to the organisation
of cellulose microfibrils in the guard cell wall. In
497
epidermal cells cellulose fibrils displayed no parallel
pattern but showed a rather disordered arrangement.
The observed cell type-specific difference in the fluorescent pattern of KAT1::GFP in epidermal and guard
cells (random versus radial distribution) may thus be the
result from the disordered or radial arrangement of
cellulose fibrils in the respective cell type. We therefore
suggest that KAT1 is directly or indirectly associated
with the cellulose fibrils in the cell wall. The stripes
formed by KAT1::GFP show the same orientation as
the cellulose fibrils but are much broader than single
cellulose fibrils which have a diameter of only about
10 nm. This can be explained by an interaction of several
parallel arranged cellulose fibrils with KAT1 or by the
restriction of KAT1 to the wider gaps found between
bundles of cellulose fibrils.
The observation that about 30% of transfected guard
cells showed no stripe formation supports the hypothesis
that KAT1 is not linked directly to the cellulose fibrils
and that this link is not obligatory. A direct binding of
KAT1 to the cellulose should have led to a radial
distribution in all transfected guard cells because all
fully developed guard cells show a radial distribution of
the cellulose microfibrils. The KAT1–cell wall association is thus most likely mediated by additional protein(s)
which are apparently not active in all cells.
In animal cell PM proteins, such as integrins, link
intracellular proteins to the ECM. Recently, it has been
shown that integrins are also physically and functionally
connected to some classes of ion channels and that this
association is of general importance for cell physiology
(for review see (Arcangeli and Becchetti, 2006)). For the
GIRK K+ channel the link between the channel and
integrins most likely occurs via an RGD motif in the
extracellular loop (McPhee et al., 1998). Other K+
channels do not contain an RGD motif. Some of these
channels may interact with integrins via their N-terminal
domain (Cherubini et al., 2005).
Even though only few homologues of classical
adhesion molecules like integrins have been identified
in plants, for a number of plant PM proteins links to the
cell wall have been suggested on the basis of their
molecular interaction with cell wall carbohydrates.
Among these are the membrane intrinsic cellulose
synthase complex and cell wall-associated kinases
(WAK), and further arabinogalactan proteins (AGP),
which are bound to the outer leaflet of the PM via a GPI
anchor (Kohorn, 2000). Also, the formation of Hechtian
strands during plasmolysis shows that attachment sites
between the PM and the cell wall exist. These attachment sites cannot be restricted to plasmodesmata since
guard cells that in general lack plasmodesmata also
develop Hechtian strands (Oparka et al., 1996). However, the nature of these attachment sites is not clear.
Results from Lang et al. (2004) demonstrated that
attachment sites of Hechtian strands are lost during
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digestion of cellulose which implicates that they are
formed by a tight connection between PM proteins and
cellulose. Likewise, digestion of cellulose diminished the
spatial pattern of KAT1::GFP (Hurst et al., 2004). We
suggest that some of the KAT1 channels may be
contributing to these attachment sites. This is also
supported by the observation that the proposed
KAT1–cell wall association and the formation of
Hechtian strands are both independent of the cytoskeleton (Lang-Pauluzzi, 2000; Lang-Pauluzzi and
Gunning, 2000).
In animal cells there is evidence that the interaction
between ion channels and integrins is accompanied by
the formation of macromolecular complexes that are
located in microdomains in the PM (Cherubini et al.,
2005). Recently, Sutter et al. (2006) analysed the
distribution of KAT1 transiently expressed in tobacco
leaf tissue. In tobacco epidermal cells KAT1 was
localised in small microdomains in the PM similar to
what we observed in epidermal cells of Vicia faba. Using
photoactivatable GFP Sutter et al. (2006) were able to
show that KAT1 was largely immobile in the PM. They
suggest that SNAREs which are known to be involved
in vesicle fusion may also participate in microdomain
formation and mobility of KAT1 in the PM of
epidermal cells. Our results show that the radial
distribution of KAT1 but not the microdomain formation was dependent on a close contact between the PM
and the cell wall. This implies that microdomain
formation and radial distribution of KAT1 are two
separate processes. The protein(s) that mediate the
association of KAT1 with cell wall components yet
remain to be determined.
Possible physiological relevance of the distinct
localisation of KAT1 in the PM
So far we can only speculate on the physiological
importance of the protein- and cell type-specific
distribution of KAT1 in the PM of guard cells. Rapid
freezing methods on plant cells showed that the distance
between the cell membrane and the cell wall is smaller
than previously observed. More recent electron micrographs revealed that the PM is appressed against the cell
wall, thus they are apparently in tight contact (Roberts,
1990). This is particularly true for guard cells as in this
specialised cell type the turgor is extremely high
(4–5 MPa; Franks et al., 2001; Raschke, 1979). Since
the cell wall serves as an external K+ store the tight
contact between the cell wall and the PM may affect the
accessibility of K+ for ion channels in the PM. We
therefore propose the following model for the physiological role of the observed K+ channel distribution in
the PM (Fig. 9). In membrane areas where the PM is
pressed tightly to the cellulose fibrils, the high resistance
cellulose
microfibril
plasma membrane
force
atmospheric
pressure = 0.1 MPa
turgor pressure =
up to 4.5 MPa
R2
R3
R1
R1
Fig. 9. Model for ion accessibility at plasma membrane–cell
wall interface of turgid guard cells. At sites where the PM is
firmly pressed against the cellulose fibrils the resistance for ion
movement (R2) is high; in between cellulose fibrils the
resistance for ion movement (R3) is low and ions can move
freely. R1 corresponds to resistance of the plasma membrane.
for ion flow reduces K+ fluxes, whereas in PM areas
that reach into the intermediate space of the cellulose
fibrils, K+ can move freely and therefore resistance for
ion flow is low. Hence in these areas the ion accessibility
for K+ channels is sufficient for guard cell function. The
K+ inward rectifier in guard cells KAT1 plays a key role
in K+ uptake during stomatal movement. The radial
distribution may reflect the arrangement of KAT1 in the
intermediate areas of cellulose fibrils and the attachment
to the fibrils would assure the stable localisation of the
protein in the PM.
However, electrophysiological measurements on protoplasts show that the cell wall link is not essential for
KAT1 function. Gutknecht et al. (1978) proposed a
turgor sensor in the PM of guard cells. Considering the
tight association of KAT1 with the cell wall found in our
study, KAT1 may participate in turgor sensing. This
underlines the physiological importance of an intimate
interaction between PM and cell wall. The identification
of the molecular nature of the link between KAT1 and
the cell wall is therefore of great importance to further
dissect stomatal functioning.
Acknowledgements
The authors would like to thank Prof. G. Thiel for
discussion and helpful comments on the manuscript.
We are grateful to N. Paris, M. Boutry, B. Kost and
A. R. Schäffner for committing the constructs with the
fusion proteins and Susanne Liebe for assistance with
the CLSM at Leica, Bensheim, Germany. The work was
supported by Grants of the Deutsche Forschungsgemeinschaft to U. Homann (SPP 1108 HO-2046/3-2 and
HO-2046/5-2).
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