The Plant Journal (2001) 26(1), 1±13
Single guard cell recordings in intact plants: light-induced
hyperpolarization of the plasma membrane
M. Rob G. Roelfsema1, Ralf Steinmeyer1, Marten Staal2 and Rainer Hedrich1,*
1
Julius-von-Sachs-Institut fuÈr Biowissenschaften, Lehrstuhl fuÈr Molekulare P¯anzenphysiologie und Biophysik,
UniversitaÈt WuÈrzburg, Julius-von-Sachs-Platz 2, 97082 WuÈrzburg, Germany, and
2
Department of Plant Biology, University of Groningen, PO Box 14, 9750 AA Haren, The Netherlands
Received 16 October 2000; revised 2 January 2001; accepted 9 January 2001.
*For correspondence (fax +49 931 8886157; e-mail [email protected]).
Summary
Guard cells are electrically isolated from other plant cells and therefore offer the unique possibility to
conduct current- and voltage-clamp recordings on single cells in an intact plant. Guard cells in their
natural environment were impaled with double-barreled electrodes and found to exhibit three
physiological states. A minority of cells were classi®ed as far-depolarized cells. These cells exhibited
positive membrane potentials and were dominated by the activity of voltage-dependent anion channels.
All other cells displayed both outward and inward rectifying K+-channel activity. These cells were either
depolarized or hyperpolarized, with average membrane potentials of ±41 mV (SD 16) and ±112 mV (SD
19), respectively. Depolarized guard cells extrude K+ through outward rectifying channels, while K+ is
taken up via inward rectifying channels in hyperpolarized cells. Upon a light/dark transition, guard cells
that were hyperpolarized in the light switched to the depolarized state. The depolarization was
accompanied by a 35 pA decrease in pump current and an increase in the conductance of inward
rectifying channels. Both an increase in pump current and a decrease in the conductance of the inward
recti®er were triggered by blue light, while red light was ineffective. From these studies we conclude
that light modulates plasma membrane transport through large membrane potential changes, reversing
the K+-ef¯ux via outward rectifying channels to a K+-in¯ux via inward rectifying channels.
Keywords: guard cell, intact plant, light-induced, membrane potential, inward rectifying K+-channel.
Introduction
Guard cell ion-transport has been studied with single cells,
using either the patch-clamp technique on protoplasts or
microelectrodes on cells in epidermal strips (Hedrich and
Roelfsema, 1999). The development of these techniques
broke new grounds in the analysis of the biophysical
properties of ion-transport proteins (Hedrich and
Schroeder, 1989). New insights were obtained into the
regulation of ion-transport by physiological factors, such
as hormones (Assmann, 1993). Guard cell responses to
changes in the light conditions, however, received little
attention, although light is a potent promoter for stomatal
opening.
Opening of stomata is induced by blue as well as red
light, but both light qualities act through different signalling pathways (Assmann and Shimazaki, 1999). The red
light effect can be inhibited by 3-(3, 4-dichlorophenyl)-1-1dimethyl-urea (DCMU) (Schwartz and Zeiger, 1984) and
ã 2001 Blackwell Science Ltd
therefore is likely to be mediated by guard cell photosynthesis. In the presence of red light, blue light further
stimulates stomatal opening. The blue light receptors that
have been found in Arabidopsis thaliana appear not to be
involved in this guard cell response (LasceÁve et al., 1999),
while the involvement of zeaxanthin is currently under
debate (Eckert and Kaldenhoff, 2000). Guard cells of the
zeaxanthin free mutant, npq1, were reported to lack a blue
light-speci®c response (Frechilla et al., 1999). However, a
blue light-speci®c stimulation of the transpiration was still
observed with npq1 plants (Eckert and Kaldenhoff, 2000).
Stomatal opening is the result of osmotic swelling of
two guard cells that surround the stomatal pore. The
osmotic swelling is accomplished by the accumulation of
K+-salts within the cell. During stomatal opening, guard
cells of Vicia faba increase their K+-concentration from
values close to 0.1 M to values exceeding 0.5 M
1
2
M. Rob G. Roelfsema et al.
(MacRobbie, 1987). These changes in the K+-concentration
are most likely mediated by K+-selective channels in the
guard cell plasma membrane (Schroeder et al., 1984).
Two types of K+-channels have been identi®ed in the
guard cell plasma membrane. Inward rectifying K+-channels activating at hyperpolarized membrane potentials,
and outward rectifying channels activating at more
depolarized potentials (Blatt, 1988; Blatt, 1992; Schroeder
et al., 1987). The inward channels are suitable for the
uptake of K+, since the negative membrane potential at
which the channel opens can drive an in¯ux of K+.
Likewise, outward channels enable K+-extrusion since
these channels are active at potentials favouring K+-ef¯ux
(Ache et al., 2000; Blatt, 1991; Schroeder, 1988). Although
the K+-uptake may be facilitated by inward K+-channels,
the uptake is energized by H+-ATPases in the plasma
membrane. The H+-ATPases extrude H+, thereby creating a
H+-gradient and an electrical potential difference across
the plasma membrane (Lohse and Hedrich, 1992).
The H+-ATPase is activated by blue light pulses superimposed on a background of red light (Shimazaki et al.,
1986). This activation is mediated via a blue light-dependent phosphorylation of the C-terminus of H+-ATPases
(Kinoshita and Shimazaki, 1999). Blue light-induced stimulation of the H+-ATPase can be measured in the whole-cell
patch-clamp mode, as a change of 3 pA in plasma
membrane current (Assmann et al., 1985; Taylor and
Assmann, 2000). Larger blue light-induced currents up to
20 pA were found, using a slow-whole-cell mode
(Assmann, 1993; Schroeder, 1988). This indicates that the
amplitude of the blue light response depends on diffusible
cytoplasmic components. Although most reports have
focused on the blue light-speci®c stimulation of H+ATPases, red light-induced outward currents have been
measured in guard cells too (Serrano et al., 1988). In these
experiments, a high intensity red light was found to trigger
an outward current of 2 pA.
Guard cells in epidermal strips, impaled with microelectrodes, were found in two physiological states. Cells in the
depolarized state have membrane potentials close to the
K+-Nernst potential and extrude K+, while hyperpolarized
cells have more negative potentials driving K+-uptake
(Roelfsema and Prins, 1998; Thiel et al., 1992). Guard cells
can shift from the depolarized to the hyperpolarized state
and vice versa. These membrane potential changes were
autonomous, rapid and often repetitive and therefore
classi®ed as oscillations (Gradmann et al., 1993).
Recently, these membrane potential changes were linked
to cytoplasmic Ca2+ concentration waves (Grabov and
Blatt, 1998). The phytohormone ABA promotes guard cells
to enter the depolarized state (Blatt and Armstrong, 1993),
while IAA has the opposite effect (Blatt and Thiel, 1994).
Based on these results, light-induced changes from one
state to the other are to be expected. Although light has
been shown to affect the membrane potential (Assmann
et al., 1985; Moody and Zeiger, 1978), light-induced
changes from one to the other state have not been
described so far.
Here, we present the ®rst study in which guard cells
were impaled in their natural environment, the intact plant.
Consequently, the impaled guard cells are still neighboured by viable and turgerescent epidermal and mesophyll cells. Furthermore, the extracellular ion
concentration will closely resemble that of intact leaves.
In this con®guration, we could measure large lightinduced membrane potential changes and determine
their consequence for K+-transport across the plasma
membrane.
Results
Current- and voltage clamp of guard cells in intact plants
Mature guard cells do not possess functional plasmodesmata (Wille and Lucas, 1984) and are electrically isolated
from other cells. Due to this unique property, guard cells
within the intact plant are suitable for current- and voltage
clamp studies. However, before these techniques can be
applied, a proper connection between a reference electrode and the guard cell wall has to be established.
In the present experiments, guard cells in the intact leave
were visualized with a water immersion objective. The
reference electrode was placed in the solution between the
objective and the epidermis. Using blunt electrodes that
are in contact with the guard cell wall via an open stoma, a
potential difference between the guard cell wall and
reference electrode could be measured. This surface
potential was sensitive to changes in the light intensity
and could reach values up to 50 mV in older leaves.
Consequently, we used the youngest fully unfolded leaves
in our experiments, which had an average surface potential of ±0.6 mV (SD 7, n = 7). Switching the microscope
lamp on or off induced transient changes in the surface
potential, with an average of 8 mV (SD 4, n = 7) for light off
and 8 mV (SD 6, n = 7) for light on. Voltage-clamp measurements may be hampered further by the resistance
between the guard cell wall and reference electrode. To get
an estimate of this resistance, blunt double-barreled
electrodes were brought into contact with the guard cell
wall, via open stomata. The series resistance measured
under current clamp was 4.9 MW (SE 0.6, n = 7) in the light
and 4.3 MW (SE 0.7, n = 7) after 10 min in the dark. With
membrane currents up to 2 nA, the applied voltage was
thus maximally overestimated by 10 mV. This overestimation of the clamped voltage, however, does not alter
with changes in the light intensity. When Vicia faba leaves
were mounted on the microscope table, stomata on the
abaxial leaf side were initially open, but closed within
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
Guard cell recordings in intact plants
3
Figure 1. Vicia faba guard cell at the abaxial
side of an intact leaf impaled with a doublebarreled electrode and pressure injected
with lucifer-yellow.
(a) Transmitted light image.
(b)
Fluorescence laser scan image
(excitation 488 nm, emission 515±565 nm).
Note that the ¯uorescent light is emitted
from the body of the cell rather than from a
con®ned region.
the following 15 min. This response may well be due to an
increase in the cytoplasmic Ca2+ concentration, known to
occur in plant cells during mechanical manipulation (Haley
et al., 1995). Thereafter, stomata opened again allowing
the impalement of single guard cells with microelectrodes.
In plant cells, the tip of a microelectrode is normally
located in the cytoplasm rather than in a vacuole (Felle,
1988), a feature of plant cells that also holds for guard cells
in epidermal strips (Blatt et al., 1990; McAinsh et al., 1990).
This subcellular localization was con®rmed for guard cells
in intact plants by pressure injection of lucifer-yellow
(Figure 1). After dye injection, ¯uorescent light was
emitted from the body of the cell, rather than from one
of the vacuoles.
Three states of V. faba guard cells in intact plants
Based on their free-running membrane potential and
plasma membrane conductance, individual guard cells
could be divided into three groups (Figure 2). The plasma
membrane of cells in all three groups depolarized immediately after impalement, while they differed in the magnitude and velocity of the subsequent repolarization
(Figure 2a). Membrane potentials of cells in the ®rst
group slowly repolarized but remained positive (Figure
2a, upper graph). When the plasma membrane of these
cells was clamped from a holding potential of ±140 mV to
more positive values, activation of inward conducting
channels was found (Figure 2b, upper graph). The inward
currents could be separated into an instantaneous and a
slowly activating component. The currents activated at
potentials positive of ±120 mV and reversed at positive
potentials (Figure 2c, upper graph). This voltage dependence is similar to that of anion channels previously
identi®ed in guard cells (Hedrich et al., 1990; Keller et al.,
1989; Linder and Raschke, 1992; Schroeder and Keller,
1992). Out of 117 cells impaled, 10 had positive membrane
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
potentials and conductance properties similar to those in
the upper graphs of Figure 2. These cells were classi®ed as
`far-depolarized' cells.
The majority of cells (66 of 117 cells measured) were
characterized by free-running membrane potentials that
repolarized to moderately negative values (Figure 2a,
middle graph); the mean membrane potential after 5 min
was ±41 mV (SD 16). Clamping the plasma membrane of
these cells from ±100 mV to values positive of ±60 mV
resulted in the activation of outward rectifying channels,
while activation of inward rectifying channels was found at
potentials negative of ±120 mV (Figure 1b, middle graph).
The activation kinetics were similar to those of outwardand inward-rectifying K+-channels, previously identi®ed in
guard cells (Blatt, 1988; Schroeder et al., 1987). The reversal potential, determined by a tail current analysis (data
not shown), was ±56 mV for outward channels (SE 5,
n = 11) and ±73 mV (SE 2, n = 25) for inward channels. A
difference in the reversal potential of both channels was
not expected, since both channels were reported to be
highly K+-selective (Blatt and Gradmann, 1997; Blatt, 1988;
Blatt, 1992). However, the selectivity of outward rectifying
channels in A. thaliana guard cells was pH dependent
(Roelfsema, 1997). In intact V. faba leaves the extracellular
pH ranges from 5.3 to 5.7 (Felle et al., 2000), while the
selectivity of outward K+ channels has been determined at
pH 7.4 (Blatt and Gradmann, 1997; Blatt, 1988). In intact
leaves, the outward rectifying channels therefore may be
less K+-selective than determined previously. The Nernst
potential of K+ is most likely close to the reversal potential
of inward rectifying channels, which do not have a pH
dependent selectivity (Blatt, 1992).
The activation of outward and inward channels was also
apparent in the current-voltage plot (Figure 2c, middle
graph). Note that only small current changes were found
between ±120 and ±60 mV. The current-voltage curve
intersected the zero current axis at ±50 mV, close to the
4
M. Rob G. Roelfsema et al.
Figure 2. Physiological states exhibited by guard cells of Vicia faba in
intact plants. Cells were found to be either far depolarized (upper
graphs), depolarized (middle graphs) or hyperpolarized (lower graphs).
(a) Free-running membrane potential (Em) starting within 10 sec after
impalement of guard cells with a double-barreled electrode, ®lled with
300 mM KCl. In all cells, Em depolarized directly after impalement, but
cells differed in the magnitude of repolarization. Far depolarized cells had
positive Em values. Depolarized cells had Em values that became more
negative, while a further hyperpolarization occurred in hyperpolarized
cells.
(b) Plasma membrane conductance properties. The far depolarized cell
(upper graph) was clamped from a holding potential of ±140 mV to more
positive potentials. Inward currents activate at potentials positive of
±120 mV. Note that inward currents have both instantaneous and slowly
activating components, similar to that of previously identi®ed anion
channels. The depolarized (middle graph) and hyperpolarized (lower
graph) cells were clamped from a holding potential of ±100 mV, to more
negative and positive values. At potentials negative of ±100 mV inward
rectifying channels activated, while outward rectifying channels activated
at potentials positive of ±60 mV. The properties of these channels are
similar to previously identi®ed K+-channels.
(c) Current-voltage relations sampled during the last 100 ms of the test
pulses in (a). Note that in all graphs the potential at which the curve
intersects the zero current axis, closely correlates with Em.
free-running membrane potential of this cell. In accordance with previous reports (Roelfsema and Prins, 1997)
these cells were classi®ed as `depolarized cells'.
A third group of cells (41 of 117 cells measured) reached
more negative membrane potentials; their average membrane potential was ±112 mV (SD 19) (Figure 2a, lower
graph). The plasma membrane conductance of these cells
was similar to that of depolarized cells (Figure 2b, lower
graph). However, the current-voltage curve was shifted to
Figure 3. Light-induced changes in Em. The bar below the graph
indicates the time points at which the microscope lamp was switched on
and off.
(a) Cell showing a transient depolarization after light changes. The cell
remained in the depolarized state throughout the experiment.
(b) Cell switching repetitively from the depolarized state in the dark to
the hyperpolarized state in the light and vice versa. Note a relatively fast
Em change in the light and a much slower Em change in the dark.
more positive current values (Figure 2c, lower graph). As a
result, the current-voltage curve intersected the zero
current axis at ±110 mV, a value slightly negative of the
threshold potential of inward rectifying channels. These
cells were therefore classi®ed as `hyperpolarized cells'.
Light-induced membrane potential changes
After the characterization of the variations in membrane
properties, guard cells were exposed to changes in light
intensity by switching the microscope lamp on and off.
Cells that were depolarized in the light mostly responded
with a transient depolarization of 8 mV (SD 8, n = 16), both
to light on or off signals (Figure 3a). These cells repolarized
to the original membrane potential after 10 min. The
membrane potential change in these cells was probably
related to changes in the surface potential.
Guard cells that were hyperpolarized in the light were
much more light responsive. Switching off the microscope
lamp resulted in a large depolarization in 9 out of the 11
cells tested, the average membrane potential change was
64 mV (SD 12). These cells could repeatedly switch from
the hyperpolarized state in the light to the depolarized
state in the dark and vice versa (Figure 3b). As a result of
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
Guard cell recordings in intact plants
Figure 4. Averaged current voltage-relation of 5 guard cells in the light
(s) and dark (d). Four cells remained in the depolarized state while one
cell switched to the hyerpolarized state in the light. The plasma
membrane conductance was measured in the light, after 15 min in the
dark and again after 15 min in the light. The average current-voltage
relation in the light is compared to that in the dark. Currents were
normalized to values in the light, to the current at ±200 mV for data
obtained at potentials negative of ±100 mV and to the current at 20 mV
for data obtained at potentials positive of ±100 mV. Labels at the right
side are the normalized values, while labels at the left side are
normalized values multiplied with the average inward or outward
current. Error bars represent SE (n = 5).
these membrane potential changes the electrochemical
gradient of K+ reverses, causing a dramatic change in the
K+-¯ux across the plasma membrane. In the light, K+
is taken up through inward rectifying channels, while K+ is
released through outward rectifying channels in the dark.
Out of the 11 cells tested, 2 cells did not depolarize, but
remained hyperpolarized in the dark. This indicates that
factors other than light can also induce hyperpolarized
membrane potentials.
Conductance changes at the plasma membrane
Light-induced conductance changes were measured by
clamping the plasma membrane from a holding potential
of ±100 mV to test potentials ranging from ±200 to 20 mV.
The conductance was measured before, during and after
15 min exposure to darkness. The mean current-voltage
relation in the light was compared to that in the dark.
Figure 4 shows a compilation of current-voltage curves of
5 cells, 4 of which remained depolarized throughout the
experiment. Currents were normalized to those measured
in the light, at ±200 mV for inward currents and 20 mV for
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
5
outward currents. Currents mediated by inward rectifying
channels increased in the dark, while no conductance
changes were apparent at potentials more positive than
±120 mV.
Figure 5(a) shows the free-running membrane potential
of a cell that was hyperpolarized in the light, but slowly
depolarized from ±117 mV to ±55 mV, after a transition to
the dark. Before and during the depolarization, the plasma
membrane was clamped from a holding potential of
±100 mV to test potentials ranging from ±180 to 0 mV
(Figure 5b). A small increase in inward currents was found
after 14 min in the dark, while outward currents remained
unchanged. The same response is displayed in the currentvoltage curves of Figure 5(c) (left graph). When currentvoltage curves were scaled to a smaller current range
(Figure 5c, right graph) an additional change in currents
from ±100 to ±60 mV was apparent. Currents at these
potentials all decreased by 35 pA after the light was
switched off. The free-running membrane potential should
be equal to the zero current potentials of the currentvoltage curve, provided that time-dependent currents
saturate during test pulses of 2 sec. Note that the zero
current potential indeed correlates with the free-running
membrane potential. Similar changes in current-voltage
curves were found for all cells that altered their membrane
potential in response to light/dark transitions. On average,
currents at ±80 mV increased by 35 pA (SE 14, n = 5) in the
light.
The change of steady state currents in Figure 5(c) was
further analysed, extracting time activated currents from
those measured at the end of the capacity compensation
peak. In the dark, currents measured early after the onset
of the clamp (interval a, Figure 6) changed towards more
inward directed values (Figure 6a). In contrast, only small
changes occurred in time-activated currents (interval difference b±a, Figure 6b). This indicates that changes in the
membrane potential are due to a light-dependent response
of fast activating ion-transporters.
Guard cell responses to red and blue light
Changes in the electrical properties of the plasma membrane in response to light/dark transitions could have been
evoked by either blue or red light, since guard cells are
sensitive to both light qualities. The sensitivity to blue and
red light was tested with guard cells that were impaled at a
low intensity red light background, provided by the
microscope lamp. All cells were depolarized after impalement and remained depolarized after the onset of additional red light, introduced from the abaxial side at a
photon ¯ux density of 450 mmol m±2 s±1. The plasma
membrane conductance was measured every 3 min and
guard cells were exposed to additional blue light, only
when inward and outward conductance had remained
6
M. Rob G. Roelfsema et al.
Figure 5. A guard cell in the hyperpolarized
state in the light, slowly depolarizing in the
dark.
(a) Em trace. Dots indicate the periods at
which voltage clamp scans were executed.
Bar shows the time at which the microscope
lamp was switched off.
(b) Current traces of the same cell as in (a)
clamped from a holding potential of
±100 mV to test potentials ranging from
±180 to 0 mV, symbols correspond to those
in (a).
(c) Current voltage relation of the cell
depicted in (a). Symbols correspond to
those below the Em trace. The steady state
currents were sampled during the last
100 ms of a 2s test pulse (as in (b)). In the
right graph the same data are displayed at a
different scale. Note the enhanced inward
current (±180 mV) in the dark (left graph)
and an inward shift of all currents in the
dark (right graph). As a result of the latter
current change, current-voltage curves
intersect the voltage axis at more positive
values, in the dark, these values closely
correlate to the Em trace.
stable over 12 min. The conductance was regarded as
stable when the changes in current during subsequent
pulses were less than 10%.
An additional beam of blue light (photon ¯ux of
200 mmol m±2 s±1) applied from the abaxial side induced
a transient hyperpolarization, in 4 out of 6 cells (Figure 7a).
The plasma membrane was clamped from a holding
potential of ±100 mV to potentials ranging from ±160 to
0 mV (Figure 7b). Blue light had little effect on timedependent outward currents, but inhibited time-dependent
inward currents (Figure 7d). In addition, blue light induced
a change of currents measured early after the onset of the
voltage clamp, relative to the zero current level (Figure 7c).
The change in current was identical for potentials ranging
from ±100 to ±60 mV, indicating that a voltage-independent conductance change underlies this response. As
expected, the potentials at which the steady state current-voltage curves intersect with the zero current axis
(Figure 7e) are the same as found for the free-running
membrane potential, as seen in Figure 7(a).
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
Guard cell recordings in intact plants
7
Figure 6. Change in current-voltage curves
after a transition to the dark, drawn for
conductances activating rapidly (a) and
slowly (b).
Data are from the same cell as those in
Figure 5, which was hyperpolarized in the
light and slowly depolarized in the dark.
Mean values are displayed of two
subsequent test pulses in the light (h) or in
the dark after 1±5 min (d), 7±11 min (j) and
13±17 min (m).
(a) Currents sampled at interval a (90±
110 ms after start of the test pulse). (b)
Time dependent currents, the difference in
current measured at intervals a and b (1850±
1950 ms after the start of the test pulse).
The blue light effect on the plasma membrane
conductance was averaged for 6 cells that were exposed
on 9 occasions to blue light. On average, the outward
rectifying channels remained unaffected by blue light
(Figure 8a). The shift in currents measured early after
onset of the voltage clamp (as shown in Figure 7c),
measured at ±80 mV, increased in amplitude by 60 pA
(Figure 8b). The blue light effect was transient; the
currents returned to their pre-stimulus value 7.5 min
after switching on the blue light. In contrast, a sustained
inhibition of inward rectifying channels was observed
(Figure 8c).
To test whether or not the light responses can be
triggered by red light, guard cells were exposed to
increased intensities of red light. A small beam of red
light, at a photon ¯ux of 450 mmol m±2 s±1, was given on
top of background red light (1.5 mmol m±2 s±1). Under
these conditions, red light could not induce a hyperpolarization in any of the cells measured (n = 5, data not
shown). In agreement with these results, additional red
light neither had an effect on the currents measured at
±80 mV (Figure 8b), nor did it affect the outward or inward
rectifying channels (Figure 8a,c).
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
In the present experiments, guard cells were exposed for
a prolonged period to blue light instead of blue light
pulses that have been used in most other reports.
Nevertheless, transient changes in the membrane potential and plasma membrane current were observed. To
compare our data with blue light-induced proton extrusion, the same light conditions were applied to guard cells
in an epidermal strip of V. faba. The blue light-induced
acidi®cation was measured with a ¯at-tipped pH-electrode
placed in contact with the epidermal strip (Figure 9, inset).
An exponential function was ®tted to the time-dependent
changes in proton concentration, and from its derivative
the rate of proton extrusion in time was calculated (Figure
9). After a lag period of 60 sec, blue light induces a rapid
rise in H+ extrusion, reaching its maximum after 120 sec
and decaying slowly afterwards.
Discussion
The impalement of guard cells within the intact plant
provides a re®ned tool to study guard cells within their
natural environment. The use of this method, however, is
complicated by the presence of a potential difference
8
M. Rob G. Roelfsema et al.
Figure 7. A guard cell in the depolarized
state
responding
with
a
transient
hyperpolarization to blue light.
(a) Em trace of a guard cell that was
depolarized in red light at a photon ¯ux of
450 mmol m±2 s±1
(hatched
bar)
and
subsequently exposed to additional blue
±2 ±1
(open bar). Dots
light 200 mmol m s
indicate the periods at which voltage clamp
scans were executed. Note that additional
blue
light
induces
a
transient
hyperpolarization.
(b) Current traces of the same cell as in (a)
clamped from a holding potential of ±100 to
test potentials ranging from ±160 to 0 mV,
symbols correspond to those in (a). Arrows
indicate the zero current level.
(c) Current voltage relation sampled early
(50±70 ms) after the onset of the clamp
voltage. Data are the mean values of 2
subsequent test pulses measured in the red
light background (j), or after the onset of
blue light for 1±5 min (s) and 13±17 min
(h). Note the shift of all currents negative of
±20 mV relative to the zero current axis.
(d)
Current-voltage relation of time
activated conductances. The difference in
current measured at the start (50±70 ms)
and at the end (1850±1950 ms) of the test
pulses. Symbols are as in (c). Note the
decrease of time activated current at
potentials negative of ±120 mV.
(e) Current voltage relation of steady state
currents, sampled 1850±1950 ms after the
start of the voltage clamp. Symbols
correspond to those below the voltage trace
in (a).
between the guard cell wall and external solution. We
found that this problem can be minimized using young V.
faba leaves, where the surface potential was relatively
small and only small light-induced transients were measured. In our opinion, the advantage of measuring guard
cells within the intact plant justi®es the complications that
arise from the surface potential and the resistance
between the guard cell wall and reference electrode.
Injection of guard cells with ¯uorescent dye revealed
that the tip of the electrode is, in most cases, located in the
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
Guard cell recordings in intact plants
9
Figure 8. Averaged guard cell responses to
red and blue light.
All guard cells were in the depolarized state
at control conditions (m,d) and were
exposed to additional light only when
currents had remained stable for 12 min
(less than 10% change in current during
subsequent test pulses). Additional red light
was given on top of background red light
(n), or blue light was given on top of red
light (s), at t = 0. Effects were measured on
outward currents (a), currents at ±80 mV (b)
and inward (c) currents.
(a) Outward current measured either at 0
or 20 mV, given relative to the current at
t = ±1.5 min. Neither blue nor red light has
an effect on outward currents.
(b) Changes in current at ±80 mV given as
the difference with t = ±1.5 min. Blue light
induces transient outward currents, while
red light has no effect.
(c) Inward currents measured either at ±180
or ±160 mV and given relative to the current
at t = ±1.5 min. Blue light inhibits the inward
currents, while red light has little effect.
Data are the mean value of 6 cells exposed
for 9 times to blue light or 5 cells exposed
for 7 times to additional red light, error bars
represent SE.
cytoplasm. However, the possibility cannot be excluded
that on occasion the electrodes may also have penetrated
the vacuolar membrane. Nevertheless, the conductance
properties of cells in all three physiological states resemble those of ion-channels known to be present in the
plasma membrane only (Figure 2). Vacuolar membranes of
guard cells are dominated by cation channels, with properties quite different from ion-channels in the plasma
membrane (Allen et al., 1998; Hedrich and Neher, 1987).
A minority of the guard cells in intact plants was far
depolarized, with conductance properties similar to those
of previously identi®ed plasma membrane anion channels
(Hedrich et al., 1990; Keller et al., 1989; Linder and Raschke,
1992; Schroeder and Keller, 1992). These channels activate
either slowly or instantaneously, most likely due to the
presence of slow and fast activating anion channels
(Dietrich and Hedrich, 1994). Exposure of these cells to
light/dark transitions did not reveal a light-dependent
regulation of anion channels (data not shown).
Most guard cells were depolarized and displayed the
activity of ion-channels with properties similar to those of
previously identi®ed K+-selective outward- and inward
rectifying channels. Some of these cells displayed lightinduced membrane potential changes that brought about a
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
shift from the depolarized to the hyperpolarized state. A
schematic presentation of the transporters involved in this
response and their current-voltage relation is shown in
Figure 10. In the dark, cells are in the depolarized state and
extrude K+ through outward rectifying channels. This may
be accompanied by the extrusion of Cl± via anion channels,
since anion channels are voltage activated at depolarized
potentials. However, in guard cells that display lightinduced membrane potential changes, the activity of
anion channels was small. Current-voltage relations of
these cells (Figures 5c and 7c,d) lacked a negative slope
from ±80 to ±100 mV, typical for cells with active anion
channels (Figure 2c). This indicates that conductances
other than anion channels may charge balance the K+ef¯ux. These ion-transporters, yet to be identi®ed, could
either facilitate an ef¯ux of anions or an in¯ux of cations. In
the light, H+-ATPases become active and hyperpolarize the
plasma membrane. Inward rectifying K+-channels are
activated, while outward K+ and anion channels are
deactivated. The change in membrane potential reverses
the driving force for K+, and in turn K+ is taken up via
inward rectifying channels.
A typical voltage dependence of the K+-channels in guard
cells is presented in Figure 10(b). The current-voltage
10
M. Rob G. Roelfsema et al.
Figure 9. Blue light-induced proton extrusion by guard cells in an
epidermal strip. (inset) Changes in the proton concentration triggered by
blue light (200 mmol m±2 s±1) applied on top of a red light background
(450 mmol m±2 s±1).
The arrow indicates the time point at which the blue light was turned on.
After a lag period of 60 sec, the changes in proton concentration [H+]
were described with a double exponential function, [H+] = [H+]ini +
[H+]change. (1-e(±t/t1))p. (1-e(±t/t2)), where [H+]ini and [H+]change are the initial
and change in H+ concentration, t1 and t2, time constants and p a free
parameter. The rate of proton extrusion in time was calculated with the
derivative of the function above. After the lag period of 60 sec, the
extrusion rate rapidly rises, peaking after 120 sec and decaying slowly
afterwards.
relation of the K+-channels was calculated from Boltzmann
equations, which were ®tted to data depicted in Figure 3.
Transporters other than K+-channels are represented as a
linear conductance with the same slope as the current
voltage relation of the cell in Figure 5. In the light, the I-V
relation of these transporters is shifted over the whole
voltage range by 35 pA. Such a change in the I-V curve is
typical for the activation of H+-ATPases, since their activity
is virtually constant over the voltage range of ±180 to 50 mV
(Lohse and Hedrich, 1992; Taylor and Assmann, 2000). As a
result of the change in the background I-V-curve, the cells
shift from depolarized to hyperpolarized potentials (Figure
10c).
Light also reduces the activity of inward rectifying K+channels, which is often interpreted as a reduction of the
guard cells capacity to take up K+ (Assmann and
Shimazaki, 1999). However, a decrease in maximum
conductance will cause a small hyperpolarization of the
plasma membrane (Figure 10c), which will in turn activate
inward channels. Overall, the actual conductance of
inward rectifying K+-channels will be little affected by a
change in maximum conductance (Roelfsema and Prins,
1998).
Provided the 35 pA increase of currents is due to the
activation of H+-ATPases, it would drive a H+-extrusion of
1.3 pmol h±1. Given a stochiometry of 1K+ taken up per H+,
this drives an K+-in¯ux of 1.3 pmol h±1. Guard cells of V.
faba take up 1.8±2.5 pmol K+ during stomatal opening
(Allaway and Hsiao, 1973; Humble and Raschke, 1971;
Outlaw and Lowry, 1977), which will thus take about 2 h.
Previously, a K+-uptake rate of 0.7 pmol h±1 was estimated
to occur during stomatal opening in V. faba (Raschke et al.,
1988).
In guard cells depolarized in red light, a beam of blue
light induced an outward current with an average of 60 pA
after 90 sec. In contrast to the effect of white light, the blue
light-induced current change was transient. The same was
found for proton extrusion rates measured in epidermal
strips, indicating that indeed the activation of H+-ATPases
underlies the current change. Stimulation of H+-ATPases
most likely occurs via phosphorylation of the C-terminus
of this protein (Kinoshita and Shimazaki, 1999).
Phosphorylation of the C-terminus, as well as ATPhydrolysis by ATPases, were at a maximum 2.5 min after
a blue light pulse. In the present experiments blue light
was applied for a prolonged period, but the change in
currents at ±80 mV peaks between 1.5 and 4.5 min to
decrease afterwards. This may relate to the nature of blue
light receptors in guard cells. After being excited by blue
light, these receptors recover only slowly (half time of
12 min, Iino et al., 1985). With a red light background, blue
light will simultaneously stimulate all receptors, giving rise
to a maximum response. Subsequently, no receptors are
available any longer and the response will be quenched
until functional receptors reappear.
Blue light also decreased the conductance of the inward
channel, but this response remained stable after 4 min.
This indicates that the light-dependent regulation of the
proton pump and the inward recti®er occur via different
signalling pathways. Further evidence for independent
signalling pathways comes from cells that remain depolarized in white light. These cells did not display a change
in currents measured at ±80 mV, but the conductance of
inward rectifying channels was still inhibited in the light.
A beam of red light, superimposed on a low intensity red
light background, failed to affect the H+-ATPase or the
conductance of the inward recti®er. Under the same
conditions, blue light triggered changes in the activity of
both transporters. This suggests that the response to white
light is due to irradiation in the blue spectrum. However,
the response of the plasma membrane differs for both
light qualities. White light induces a steady hyperpolarization, while blue light causes a response that is maximal
after 2 min and decays afterwards. The origin of this
difference is at present not fully understood. In the present
experiments the CO2 levels were kept low by a steam of
CO2 free air directed on the drop of solution between the
leaf and objective. Low CO2 concentrations are known to
enhance the blue-light sensitivity of stomata (Assmann,
1988; Karlsson, 1986), but a red light response, depending
on photosynthesis, may be inhibited by low CO2. Future
experiments will therefore be directed towards the
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
Guard cell recordings in intact plants
11
Figure 10. Schematic presentation of guard cell ion-transport proteins, their voltage dependence and regulation by light.
(a) Activity of ion-transport proteins in the dark (upper cartoon) and in the light (lower cartoon). In the dark, outward K+-channels are activated due to the
depolarized potential and extrude K+. The K+ extrusion is balanced by the ef¯ux of Cl± via anion channels or by a ion ¯ux via so far unidenti®ed
transporters. In the light, the activation of H+-ATPases hyperpolarizes the plasma membrane and drives K+-uptake via inward K+-channels. At this potential
outward K+ channels and anion channels are deactivated.
(b) Current voltage relation of inward K+-channels, outward K+-channels and background currents. Ion-transporters other than slow activating K+-channels
are represented as a linear conductance, the slope resembles that of current voltage relations in Figure 5. The effect of white light is represented as a shift
along the current axis of 35 pA (striped line). The IV-curves of inward and outward K+-channels were calculated from data in Figure 3. The current voltage
relations were calculated using the following Boltzmann equation; Gch = Gmax/(1 + e(k(V1/2±Vm))), where Gch is the voltage dependent channel conductance
and Gmax the maximal conductance, k a constant and V1/2 the potential of half maximal conductance. For both inward and outward K+-channels Gmax was
set to 10 nS in the light, while Gmax of inward channels was decreased to 7 nS in the dark.
(c) Sum of IV-curves in the dark (solid line) and in the light (striped line). For comparison the IV-curve in the light without a change in the conductance of
inward K+-channels is shown (dotted line).
exploration of interactions between CO2, red-and blue
light, in guard cells.
Experimental procedures
Plant growth and experimental setup
Broad bean (Vicia faba L. cv. GruÈnkernige Hangdown, Gebag,
Hannover, Germany) plants were grown in a green house. Fourto 6-week-old plants were placed next to an upright microscope
(Axioskop 2FS, Carl Zeiss, GoÈttingen, Germany) The adaxial side
of a leaf was mounted on a plexiglas holder in the focal plane
using double-sided adhesive tape. In this experimental con®guration, the abaxial side of the leaf was accessible for impalement
with microelectrodes. For this purpose, a water immersion
objective (Achroplan 40x/0.80 W, Carl Zeiss) was submerged
into a drop of the following solution: 50 mM KCl, 5 mM 2-(NMorpholino) ethane sulphonic acid/Bis-Tris propane pH 6.5 and
0.1 mM CaCl2, which was placed on the leaf surface. Stomata
were shielded for the variable CO2 concentration in the laboratory
ã Blackwell Science Ltd, The Plant Journal, (2001), 26, 1±13
by a ¯ow of CO2-free air directed at the drop of solution. The
reference electrode, a 1 M KCl and 0.1% agarose salt bridge,
connected to an AgCl/Cl half cell, was placed in the solution
between the cuticula and objective. Electrodes were moved
towards an open stomate with a micro manipulator (type 5171,
Eppendorf, Hamburg, Germany). A piezo translator (P-280.30,
Physik Instrumente, Waldbronn, Germany) was used to impale
the electrode into a guard cell. Guard cells under investigation
were 41 mm (SD = 3, n = 60) in length and 14 mm (SD = 1, n = 60)
in diameter.
Electrodes and electrical con®guration
Double-barreled electrodes were pulled from two glass capillaries
(GC100F-10, Clark Electromedical Instruments, Pangbourne,
Reading, UK) that were aligned, heated and twisted 360° on a
customized vertical electrode puller (L/M-3P-A, List Medical
Electronic, Darmstadt, Germany). A ®rst pull was executed on
the vertical electrode puller, while the ®nal pull was carried out on
a horizontal laser puller (P-2000, Sutter Instumant Co., Novato,
CA, USA). The electrodes were ®lled with 300 mM KCl and had
12
M. Rob G. Roelfsema et al.
200 mmol m±2 s±1. Light sources and ®lters were the same as
described previously (Roelfsema et al., 1998).
resistances ranging from 60 to 110 MW. Blunt double-barreled
electrodes, used to measure the surface potential and series
resistance, had a tip resistance ranging from 3 to 14 MW. The
electrodes were connected via a 1 M KCl bridge and AgCl/Cl half
cells to a double microelectrode ampli®er (VF-102, Bio-Logic,
Claix, France) equipped with headstages of 1011 W input impedance.
Voltage step protocols were applied via an ITC±16 interface
(Instrutech, Corp., Elmont, NY, USA) under control of Pulse
software (Heka, Lambrecht, Germany). The test voltages were fed
into a differential ampli®er (CA-100, Bio-Logic, Claix, France)
connected to the VF-102 ampli®er. The data were low pass ®ltered
at 300 Hz with 8-pole Bessel (type 902, Frequency Devices,
Haverhill Ma, USA) and sampled at 1 kHz.
We thank M. Knoblauch and A.J.E. van Bel (University of Giessen,
Giessen, Germany) for their useful suggestions concerning
microscopy and the leaf holder; H.B.A. Prins (University of
Groningen, Haren, The Netherlands) for technical support; D.
Carden (University Giessen, Giessen, Germany) and P. Dietrich
(University of WuÈrzburg) for help with the preparation of the
manuscript.
This
work
was
funded
by
Deutsche
Forschungsgemeinschaft grants to R.H.
Pressure injection
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Light sources
Leaves were illuminated with the microscope lamp (HAL 12 V/
100 W, Carl Zeiss), focused on the adaxial side at an area of 2 cm
in diameter. The photon ¯ux densities were 98 and 4.4 mmol
m±2 s±1 at the adaxial and abaxial side, respectively. Switching the
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Proton extrusion
Epidermal strips were peeled from the abaxial side of leaves and
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placed in a measuring chamber (Roelfsema et al., 1998) ®lled with
50 mM KCl, 0.1 mM CaCl2 and 0.5 mM 2-(N-Morpholino sulphonic
acid/Bis-Tris propane pH 6.5. A combined pH-reference electrode
with a ¯at tip (Ingold Electrodes, Wilmington, MA, USA) was
placed in contact with the epidermal strip. The strip was illuminated at a 45° angle, with red light at a photon ¯ux density of
450 mmol m±2 s±1 and additional blue light at a ¯ux density of
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