Kinetics of the Onset of Stomatal Closure by Jeff Hunter

CO2-Triggered Chloride Release from Guard Cells in Intact Fava Bean Leaves. Kinetics of the Onset of Stomatal Closure1 Stefan M. Hanstein* and Hubert H. Felle Botanisches Institut I, Justus-Liebig-Universita¨t, D–35390 Giessen, Germany The influence of CO2 on Cl⫺ release from guard cells was investigated within the intact leaf by monitoring the Cl⫺ activity in the apoplastic fluid of guard cells with a Cl⫺-sensitive microelectrode. In illuminated leaves adapted to a CO2 concentration within the cuvette of 350 ␮L L⫺1, an increase of 250 ␮L L⫺1 CO2 triggered a transient rise in the apoplastic Cl⫺ activity from 3 to 14 mm within 10 min. This Cl⫺ response was similar to the Cl⫺ efflux evoked by turning off the light, when the substomatal CO2 was kept constant (CO2 clamp). Without CO2 clamp, substomatal CO2 increased by 120 ␮L L⫺1 upon “light off.” The response to an increase in CO2 within the cuvette from 250 to 500 ␮L L⫺1 in dark-adapted leaves was equivalent to the response to an increase from 350 to 600 ␮L L⫺1 in the light. No Cl⫺ efflux was triggered by 2-min CO2 pulses (150–800 ␮L L⫺1). After a switch from 350 ␮L L⫺1 to CO2-free cuvette air, the guard cells were less sensitive to a rise in CO2 and to light off, but the sensitivity to both stimuli partially recovered. Changes in CO2 also caused changes of the guard cell apoplastic voltage, which were generally faster than the observed Cl⫺ responses, and which also promptly occurred when CO2 did not initiate Cl⫺ efflux. The comparatively slow activation of Cl⫺ efflux by CO2 indicates that an intermediate effector derived from CO2 has to accumulate to fully activate plasma membrane anion channels of guard cells.

Stomata close when guard cell turgor drops, a process initiated by plasma membrane depolarization. As soon as the membrane voltage becomes positive of the equilibrium potential (EK), K⫹ leaves the cells accompanied by water due to osmotic forces. This depolarization is caused by anions (Cl⫺ and malate) rapidly moving down their electrochemical gradient, or by a “destimulation” of the plasma membrane proton pump. Although evidence is emerging that the activation of anion channels may be the preponderant event therein, it is clear that during stomatal closure, the activity of the pump must be reduced to be activated again upon stomatal opening. Stomata close when the CO2 concentration within the leaf increases (Willmer and Fricker, 1996). However, the way CO2 is sensed by the leaf, and the signal transduced to the respective transporters, is poorly understood (Assmann, 1999). One reason for our ignorance with this problem may be the difficulty in studying the response of the functional entity (stomata plus substomatal cavity with adjacent cells) to well-controlled substomatal CO2 concentration changes with high-time resolution in a quantitative manner under different physiological conditions. We have developed a technique to monitor ion activities in the guard cell apoplast within the intact leaf (Hanstein and Felle, 1999; Felle et al., 2000). To control the 1 This work was supported by the Deutsche Forschungsgemeinschaft (grant no. Fe213/12–1). * Corresponding author; e-mail stefan.m.hanstein@bot2.bio.unigiessen.de; fax 49 – 641–99 –35119. Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.004283.

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substomatal CO2 concentration, this technique was combined with a minitube cuvette and a newly developed CO2 microsensor that measures the CO2 concentration within an individual substomatal cavity (Hanstein et al., 2001). With this system, it is possible to measure and to control the CO2 concentration inside the leaf and thus any CO2 concentration gradient that may be built up during tests. Because large Cl⫺ fluxes occur before stomata visually start to close, we use the online measurement of Cl⫺ efflux within the cavity (Felle et al., 2000) as an early and reliable indicator of stomatal closure. Monitoring guard cell Cl⫺ efflux from within the substomatal cavity, whereas at the same time the substomatal CO2 concentration can be controlled, offers access to two areas of scientific interest: In the field of guard cell signal transduction, we were able to address the problem of CO2 sensing by guard cells. For instance, the question of whether guard cell anion channels are direct targets of CO2 was addressed. In the field of integration of stomatal responses to different environmental stimuli, the interplay of light and CO2 in the stomatal response to sudden shading was revisited. Sudden shading is accompanied by a rise in substomatal CO2 within seconds (Laisk et al., 1984; Pearcy, 1990). Based on comparative analysis of the steady-state conductance of C3 plants as a function of substomatal CO2 and of the photon flux density in well-watered conditions, the influence of CO2 on stomatal conductance is considered to be slight (Wong et al., 1978; Sharkey and Raschke, 1981; Ball et al., 1987). Differing from this steady-state approach, the current investigation focuses on the trigger po-

CO2-Triggered Cl⫺ Release from Guard Cells

tential of a fast CO2 increase in the physiological range to accelerate the onset of stomatal closure. RESULTS Substomatal CO2 Rapidly Responds to Light off and Light on

Light off is a stomatal closing signal. As CO2 is incorporated into ribulose-1,5-bisphosphate, and the processing thereof is energy dependent, loss of light energy will disturb the dynamic equilibrium of CO2 incorporation and CO2 release (e.g. from respiration). Figure 1 shows a CO2 measurement at a CO2 concentration within the cuvette of 350 ␮L L⫺1. The sensor was first positioned in the air stream above the leaf surface and was then carefully moved toward the leaf surface and into a substomatal cavity. Within the cavity the CO2 concentration was 90 ⫾ 31 ␮L L⫺1 (se, n ⫽ 7) higher compared with the cuvette air. Turning off the light quickly triggered a CO2 increase of 120 ⫾ 20 ␮L L⫺1 (se, n ⫽ 6). Upon light on, the CO2 level before light off was approximately restored. When the leaf surface was flushed with CO2-free air, a substomatal CO2 concentration of 149 ⫾ 34 ␮L L⫺1 (se, n ⫽ 9) was measured. Under these conditions, light off increased substomatal CO2 by 95 ⫾ 30 ␮L L⫺1 (se, n ⫽ 4). Apoplastic Voltage

The apoplastic voltage was monitored with a blunt microelectrode, routinely used as voltage reference to the Cl⫺-selective microelectrode. Because there is a

high electrical resistance between the site of measurement and the bath harboring the cut petiole, initial voltage changes qualitatively represent the inverse of the plasma membrane potential. Thus, the apoplastic voltage can be used as reliable, noninvasive indicator of the direction of membrane potential changes occurring immediately after imposing the test conditions. As shown in Figure 2A, the increase in the CO2 concentration from 350 to 600 ␮L L⫺1 was responded to by an apoplastic depolarization by 8 ⫾ 1 mV within 1.6 ⫾ 0.3 min (se, n ⫽ 6), which reflects a hyperpolarization of the membrane potential; the decrease in CO2 had the inverse effect. The apoplastic voltage also responded with an initial depolarization of comparable extent and velocity to light off. To determine the contribution of the mesophyll to the observed apoplastic voltage changes, the epidermis was removed and the apoplastic voltage was measured at a mesophyll cell (Fig. 2B). It was almost insensitive to CO2 changes, whereas the light off effect was comparable with that measured in the guard cell apoplast (Fig. 2A). Figure 2C shows that the guard cell apoplastic depolarization depends on the CO2 concentration, saturating around 700 ␮L L⫺1. CO2 Triggers Clⴚ Efflux

In Figure 3 the apoplastic Cl⫺ activity is shown to respond continuously to changes in CO2 within the cuvette from 150 to 800 ␮L L⫺1 and back. Short CO2 pulses of 1 or 2 min in duration (measurement of substomatal CO2 see inset) did not or only marginally changed Cl⫺ activity, did not initiate stomatal

Figure 1. Effect of light off on substomatal CO2 concentration measured directly with a CO2 microsensor. The leaf surface was enclosed in a minitube cuvette flushed with air containing 350 ␮L L⫺1 CO2 (see text). The sensor tip was first placed 2 mm above the leaf surface, and was then moved toward a stoma, and was finally positioned within the substomatal cavity. Representative trace of six independent experiments. Plant Physiol. Vol. 130, 2002

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Figure 2. Effect of CO2 changes and of light off on apoplastic voltage (the voltage between the guard cell apoplastic fluid and ground, see text). A, Response of the apoplastic voltage in the vicinity of guard cells to an increase in CO2 within the cuvette compared with the light off effect. Representative kinetics of six independent experiments. B, Apoplastic voltage recorded within the mesophyll without interference from the epidermis. Measurement was performed after locally removing the epidermis. C, Response of the apoplastic voltage in the vicinity of guard cells to different CO2 concentration changes in the cuvette air. The CO2 concentration was switched from 0 ␮L L⫺1 to 100 ␮L L⫺1 for 1 min, and was then switched back to 0 ␮L L⫺1. After 2 min, a concentration of 300 ␮L L⫺1 was applied for 1 min, again followed by a CO2-free period of 2 min, before the next CO2 concentration level was applied.

closure, but they clearly affected the membrane potential (kinetics of membrane potential changes see Fig. 2). Extended exposure to 800 ␮L L⫺1 CO2 resulted in a rapid apoplastic increase in Cl⫺ activity that, after approximately 13 min, peaked around 16 mm, and over about 90 min, spontaneously returned to the starting level of about 2 mm. Stomata were completely closed 45 min upon the rise in CO2. A subsequent decrease in CO2 caused a rapid drop in Cl⫺ to below 1 mm. As indicated by the ovals (visual observation of stomatal aperture), the stomata were still open when Cl⫺ activity peaked; this test demonstrates that Cl⫺ efflux precedes stomatal closure and signals its onset. 942

The Physiological Relevance of the CO2 Trigger

We were interested in the control potential (physiological impact) of the closing signal “elevated CO2” in relation to the closing signal light off. In Figure 4, the substomatal CO2 concentration and the apoplastic Cl⫺ activity are shown responding to different CO2 concentrations in the light and after light off. The switch from 350 to 600 ␮L L⫺1 CO2 within the cuvette is responded to by an increase in apoplastic Cl⫺ activity to a level of 13.8 ⫾ 2.5 mm (se, n ⫽ 5), which is comparable with that shown in Figure 3. It took 1.6 ⫾ 0.9 min until the Cl⫺ activity had changed by more than 10%. A time of 3.8 ⫾ 1.2 min was required until Cl⫺ had changed by more than 50%. Plant Physiol. Vol. 130, 2002

CO2-Triggered Cl⫺ Release from Guard Cells

Figure 3. Response of guard cell apoplastic Cl⫺ activity (pCl) to periods of elevated CO2 of different duration. The CO2 concentration within the cuvette ([CO2]cuv) is given in the bar on the top with shaded areas for periods of high [CO2]cuv. Inset shows the substomatal CO2 change (⌬ci) during the 2-min CO2 pulse, which was directly recorded by a CO2 microsensor within the substomatal cavity. Stomatal apertures were microscopically observed (open ovals, stomata open; closed ovals, stomata closed). The experiment was conducted at a photon flux of 300 ␮mol m⫺2 s⫺1, a temperature of 22°C, and a relative air humidity of 80%. The figure gives an example of three experiments performed with different leaves.

When CO2 was increased by 120 ␮L L⫺1, which is equivalent to the CO2 increase that occurs upon light off, the Cl⫺ increase was about 50% of that induced by 250 ␮L L⫺1. Figure 4 shows that Cl⫺ activity dropped to levels below 1 mm following the reduction in CO2 to 0 ␮L L⫺1. Visual control proved that stomata were open at that point. When light was turned off, substomatal CO2 increased to 300 ␮L L⫺1, but apoplastic Cl⫺ activity responded with a minor transient increase only. Under these conditions, stomata remained open for the next 1 h. Even an increase in CO2 within the cuvette to 250 ␮L L⫺1 (⫽550 ␮L L⫺1 in cavity) could not trigger the typical Cl⫺ efflux to a level above 10 mm. Only when the CO2 within the cuvette was increased to 500 ␮L L⫺1 did the apoplastic Cl⫺ rise to about 16 mm and the stomata close. The kinetics in Figure 5 show that the effectiveness of the light off signal in inducing Cl⫺ efflux increases at a higher CO2 level within the cavity. The protocol shows that, in agreement with Figure 4, the changes in the CO2 concentration within the cuvette from 350 to 600 ␮L L⫺1 and back (control) yielded the expected Cl⫺ response. However, when in contrast to the test shown in Figure 4, the cavity CO2 concentration was experimentally kept constant at 400 ␮L L⫺1 (CO2 clamp), light off alone (without concomitant CO2 increase) triggered Cl⫺ efflux to a level of 23.4 mm Plant Physiol. Vol. 130, 2002

(n ⫽ 2; 22.8 and 24 mm) within 25 min and caused stomatal closure. Adaptation

The Cl⫺ responses to light off and to elevated CO2 differed according to the duration of exposure to low CO2 concentrations. Whereas light off shortly after switching to 0 ␮L L⫺1 (CO2 concentration within the cuvette) only caused a minor rise in apoplastic Cl⫺ to a level of 3.2 mm (n ⫽ 3: 1.3, 2.5, and 5.8 mm), and within the following 1 h did not increase apoplastic Cl⫺ activity or closed the stomata (Fig. 4), a prolonged adaptation to 0 ␮L L⫺1 (⬇150 ␮L L⫺1 cavity CO2) yielded a different response. As shown in Figure 6, light off still did not trigger an immediate Cl⫺ efflux after being exposed to low CO2 for about 100 min, but did so after being about 20 min total darkness. The apoplastic Cl⫺ activity increased to a level of 6.2 mm (n ⫽ 3; 4.5, 6.9, and 7.2 mm). Similar results were obtained for elevated CO2 (Fig. 7): After a 30min adaptation to 0 ␮L L⫺1, the apoplastic Cl⫺ activity remained indifferent to increases from 0 to 225 ␮L L⫺1 or to 450 ␮L L⫺1, and only 900 ␮L L⫺1 triggered the typical Cl⫺ transient (Fig. 7A; n ⫽ 3). However, after prolonged adaptation (70 min) to 0 ␮L L⫺1, the increase from 0 to 225 ␮L L⫺1 triggered the Cl⫺ transient in five out of six leaves (Fig. 7B). These 943

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Figure 4. Response of guard cell apoplastic Cl⫺ activity (pCl) to CO2 changes within the cuvette (bar on the top) in the light (at 300 ␮mol m⫺2 s⫺1) and in darkness. Substomatal CO2 concentration ([CO2]sub) was recorded by a CO2 microsensor directly within the substomatal cavity. Measurements of pCl and of substomatal CO2 were sequentially performed within the same cavity. The data basis is two experiments with different leaves.

experiments demonstrate that for both stimuli, light off and elevated CO2, adaptation shifts the sensing threshold necessary to initiate stomatal closure. A remarkably delayed CO2 response was observed in the case of temporary darkness during the adaptation period to 0 ␮L L⫺1 CO2 within the cuvette (Fig. 6): Approximately 10 min after the end of the intermittent dark phase, the CO2 concentration was raised to 175 ␮L L⫺1, and the Cl⫺ response was delayed by about 30 min. DISCUSSION A Sensitive Parameter to Monitor the Onset of Stomatal Closure

Upon a change in ambient CO2, there is a general pattern of short-term stomatal response: Decreasing the CO2 concentration stimulates stomatal opening, whereas increasing the CO2 concentration triggers closure (Morison, 1987). Because stomatal closure only occurs after ions (anions and K⫹) have started to be released from the guard cells, any method to 944

directly observe the actual movement must be inferior to the measurement of the preceding channel activation, monitored as ion fluxes. The measurement of a pronounced Cl⫺ efflux from guard cells within the intact leaf has been demonstrated following the closing signals light off and abscisic acid (Felle et al., 2000). In the current investigation, we adopted the measurement of apoplastic Cl⫺ activity to monitor the onset of stomatal closure upon a rise in ambient CO2. As expected, the increase in the apoplastic Cl⫺ activity can be measured before the progressive release of osmotica from guard cells visibly affects stomatal aperture (Fig. 3). We do not infer that Cl⫺ efflux was the primary event because there is evidence that the activation of the anion channels is triggered by cytosolic Ca2⫹ elevation (Ward et al., 1995; Webb et al., 1996; Webb and Hetherington, 1997; Allen et al., 1999). Ca2⫹ was not chosen as the measuring parameter because Ca2⫹ as a signaling element is not directly involved in the osmotic or charge balance during stomatal closure, whereas Cl⫺ is and thus reliably signals the onset of stomatal Plant Physiol. Vol. 130, 2002

CO2-Triggered Cl⫺ Release from Guard Cells Figure 5. Response of guard cell apoplastic Cl⫺ activity (pCl) to a CO2 increase by 250 ␮L L⫺1 compared with the CO2-independent light off effect. The CO2 concentration within the cuvette is given in the bar on the top. Upon light off, the substomatal CO2 concentration ([CO2]sub) was held constant by simultaneously lowering the CO2 concentration in the cuvette (CO2 clamp). The data basis is two experiments with different leaves.

closure. The validity of this notion is underscored by the observation that short CO2 pulses from 150 to 800 ␮L L⫺1 (Fig. 3) did not trigger Cl⫺ efflux or induce stomatal closure. As a consequence, measuring apoplastic Cl⫺ activity provides information not only on the kinetics of the upstream signal transduction that activates Cl⫺ efflux, but also on the effectiveness of a given CO2 increase to bring about an aperture change.

Over extended periods, the composition of osmotica within the guard cell may shift between ionic species on the one hand and Suc on the other hand (Talbott and Zeiger, 1996). Of course, whenever such shifts occur, apoplastic Cl⫺ activity is only of limited use in comparing the effectiveness of environmental signals to initiate stomatal closure. The same holds true for long-term shifts between chloride and malate as the major counter ion of potassium. This investi-

Figure 6. Effect of light off (L. off) on guard cell apoplastic Cl⫺ activity (pCl) after adaptation to different CO2 levels (given in the bar on the top). The data basis is three experiments with different leaves. Plant Physiol. Vol. 130, 2002

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Figure 7. Adaptation to low CO2 concentrations. Apoplastic Cl⫺ activity (pCl) responding to a CO2 increase following a prolonged CO2free period. The CO2 concentration within the cuvette is given in the bar on the top. Leaves were exposed to CO2-free cuvette air for 30 min (A) or for 70 min (B) before the CO2 concentration was stepwise increased, as indicated. Data are representative of six leaves investigated after 70 min of adaptation and three leaves investigated after 30 min of adaptation.

gation reports remarkable lag phases upon a rise in CO2 that precede Cl⫺ efflux (Fig. 6: CO2 increase from 0 to 175 ␮L L⫺1) and even insensitivity of the “sensitive parameter” (apoplastic Cl⫺ activity) toward an increase in CO2: When the CO2 concentration in the cuvette drops from 350 ␮L L⫺1 to 0 ␮L L⫺1 (substomatal CO2 concentration in the light about 150 ␮L L⫺1; see Fig. 4), a subsequent rise in CO2 by 225 ␮L L⫺1 does not trigger an increase in apoplastic Cl⫺ activity (Fig. 7A). By further stepwise increase in CO2, it was possible to activate the anion channels, but it is interesting that between the last CO2 step and the onset of Cl⫺ release, several minutes passed (Fig. 7A). Because the apoplastic Cl⫺ activity was in the range of 100 ␮m, any earlier Cl⫺ release from guard cells would have been easily detected. The insensitivity occurring upon CO2 steps from 0 to about 200 ␮L L⫺1 raises the question whether the lack of the Cl⫺ response could be attributed to a replacement of Cl⫺ as a major osmoticum by organic compounds. However, a 30-min treatment with CO2-free air is very unlikely to cause a significant build-up of carbon assimilates like Suc or malate. Moreover, after extended exposure to CO2-free air, the influence of CO2 on the apoplastic Cl⫺ activity did not further decrease, but it increased again (Fig. 7B). As a consequence, we have no argument to underscore that Cl⫺ was replaced by other osmotica during the exposure to CO2-free air. Thus, the data strongly indicate that the CO2 signal transduction chain does not convey the rise in CO2 to guard cell anion channels or that the signal transduction requires a longer time, thereby preventing or delaying stomatal closure. The resulting hints for the mechanism of CO2 signal transduction will be discussed below. 946

Controlling the Substomatal CO2 Concentration by a Minicuvette

The use of microsensors for observing the onset of stomatal closure has implications for the experimental system. A system was required that did not only allow a controlled CO2 concentration and fast concentration changes, but also access of microelectrodes into substomatal cavities. Fast cuvette systems for measuring transient gas exchange have been designed for studying the impact of transient light (sunflecks in forest understories) on photosynthesis (Pearcy, 1990). The use of 2-mm thin, single-sided minicuvettes allowed for response times below 1 s for measuring transient gas exchange (1990). In our approach, a single-sided “leaky“ minicuvette had to be used containing a window for electrode access. The employed tube-shaped minicuvette offered the advantage of efficiently suppressing CO2 exchange between the leaf surface and the electrode window on the opposite side at a low gas supply rate due to the well-defined air flow through the tube (Hanstein et al., 2001). Because the leaf surface enclosed by the leaf window was small, substomatal CO2 could not be measured by conventional infrared gas analyzers (Ball, 1987). Thus, a CO2 microsensor was used to measure CO2 directly within the cuvette or within a single substomatal cavity (Hanstein et al., 2001). It is surprising that measurements with this sensor within the illuminated leaf revealed substomatal levels higher than the CO2 concentration in the cuvette at any concentration tested between 0 and 800 ␮L L⫺1 (Figs. 1, 4, and 5). As this was not due to concentration differences between inside and outside, it must have metabolic sources representing dynamic equiPlant Physiol. Vol. 130, 2002

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libria between CO2 consuming and producing processes, where high respiration rates obviously preponderate. The validity of this observation is underscored by the demonstration that any change in CO2 within the cuvette is closely tracked by substomatal CO2 (e.g. Fig. 4). We consider the elevated substomatal CO2 concentration to be a consequence of the pressure exerted by the sealing cuvette edge, which obviously triggers respiratory processes in the underlying cells and increases CO2 also in the neighboring tissue where the measurement is performed. In fact, after 1 d of acclimation, a substomatal CO2 concentration below 350 ␮L L⫺1 was measured at a CO2 concentration of 350 ␮L L⫺1 in the cuvette (not shown).

Hints for CO2 Signal Transduction Mechanisms

Although the input signal of CO2-dependent regulation of guard cell movements is considered to be the substomatal CO2 (Mott, 1990), the mechanism by which guard cells sense and respond to CO2 signals remains poorly understood. The combination of Cl⫺ and CO2 measurements is a versatile tool in testing current hypotheses on CO2-dependent stomatal control. Upon a fast rise in substomatal CO2 by 650 ␮L L⫺1, it took several minutes until the apoplastic Cl⫺ activity in the direct vicinity of guard cells had increased by 1 mm (calculated from the pCl values in Fig. 3). This slow initial stage of Cl⫺ release indicates that upon contact with increased CO2 concentrations, Cl⫺ channels are not activated immediately, suggesting an indirect influence of CO2 on these channels. Hedrich and Marten (1993) have proposed a positive feedback model in which CO2-stimulated and photosynthetically driven changes in apoplastic malate activate guard cell anion channels (R-type), resulting in anion loss. From the moment of a CO2 concentration change to the arrival of malate in the apoplast at concentrations high enough to stimulate the anion channels (Hedrich et al., 1994), time elapses that would depend on the amount of CO2 added, but also on the CO2 concentration to which the stomata were adapted. This model agrees with our observations of a considerable delay in Cl⫺ efflux that occurs after a switch to CO2-free cuvette air (unless an adaptation process has taken place). The dependence on substomatal CO2 is shown by the data in Figures 4 and 5 where the change from 350 to 600 ␮L L⫺1 CO2 (cuvette) triggered Cl⫺ efflux to an apoplastic level above 10 mm within 10 min in comparison with CO2 changes from 0 to 250 ␮L L⫺1 (Fig. 4), 0 to 175 ␮L L⫺1 (Fig. 6), and 0 to 225 ␮L L⫺1 or 225 to 450 ␮L L⫺1 (Fig. 7A), which did not immediately trigger Cl⫺ fluxes. Brearley et al. (1997) have reported that in epidermal strips activation of anion currents occurs within 2 min upon a rise in CO2 from 350 to 1,000 ␮L L⫺1. This fast response of Cl⫺ currents in the absence of the mesophyll seems to argue against a role for malate in Plant Physiol. Vol. 130, 2002

the activation of guard cell anion channels. However, it does not contradict the more general hypothesis that some CO2 product (not necessarily malate) is required for channel activation, because the rate of formation of a CO2 product is expected to be a function of the CO2 concentration, and the CO2 concentration applied by Brearley et al. (1997) was rather high. Moreover, activation thresholds of guard cell anion channels could be affected by removing the mesophyll tissue from the epidermis due to differences in the apoplastic ionic conditions, a different pressure exerted by the cell wall corset or due to the wounding response of the epidermis itself. We found that the sensitivity of the Cl⫺ efflux to CO2 was approximately the same in the light and in darkness. Due to the lack of reliable malate concentration data from the substomatal cavity, it is difficult to judge whether the malate model would hold in light and in darkness. However, it is clear that because of the long lag phases, the anion channel cannot be the primary CO2 sensor. Minor Cl⫺ responses to CO2 increase that are observed here and there may be related to changes in apoplastic pH (data not shown) or in voltage (see below). Zeiger and Zhu (1998) proposed a model in which an increase in the CO2 level leads, via CO2 fixation, to changes in ATP and NADPH status and thus to an increase in the pH of the chloroplast lumen. Because of the pH sensitivity of the enzymes involved in zeaxanthin formation within the chloroplast lumen, this would lead to a decrease in zeaxanthin levels, which is in fact demonstrated when leaves or epidermal peels were exposed to elevated CO2. Our observation that CO2 sensitivity of Cl⫺ efflux was the same in light and darkness would not favor the idea of different CO2 sensors for light and dark, as suggested by Zeiger and Zhu (1998). Zeaxanthin is supposed to be the photoreceptor for blue light that triggers blue light-dependent stomatal opening (Shrivastava and Zeiger, 1995; Assmann and Shimazaki, 1999). Our data obtained with white light indicate that zeaxanthin does not participate in the transduction of the light off signal: At a low ambient CO2 concentration, which has been shown to increase the zeaxanthin concentration (Zeiger and Zhu, 1998; Zhu et al., 1998), triggering of Cl⫺ efflux by light off was less efficient compared with normal ambient CO2 (Figs. 4 and 6). Is there a role for cytosolic pH? Cl⫺ efflux was shown to be independent of initial changes in the simultaneously recorded apoplastic voltage. Figure 2 shows that the initial response of the apoplastic voltage (qualitatively the inverse of the membrane potential) to an increase in CO2 and to light off is a rapid depolarization (i.e. membrane hyperpolarization) of comparable magnitude. In fact, these voltage changes depend on the CO2 concentration supplied and they saturate at around 700 ␮L L⫺1 (Fig. 2C). Using pH-sensitive microelectrodes, Felle and Bertl 947

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(1986a, 1986b) demonstrated in a variety of green cells, that light off always resulted in an initial cytosolic acidification of about 0.3 pH units. As protons are transport substrate of the plasma membrane H⫹ pump, this doubling of the cytosolic H⫹ activity was responded to by a transient hyperpolarization (equivalent to a depolarization in the apoplast). Here, it is suggested that the apoplastic depolarization (⫽ plasma membrane hyperpolarization) due to CO2 elevation may also be due to cytosolic acidification (proposed by Raschke, 1975). Because the short CO2 pulses (Fig. 3) did not trigger Cl⫺ fluxes, but caused apoplastic depolarizations (and thus presumably also cytosolic acidification), it is unlikely that these early pH changes are involved in CO2 sensing in a causal manner. Because this notion concerns only the primary responses to CO2, it does not collide with Blatt and Armstrong’s (1993) report on the activation of the K⫹out rectifier by cytosolic alkalization. Likewise, apoplastic pH is also not a primary factor in CO2 sensing because it is the result of ion fluxes and not their cause (Felle et al., 2000).

The Potential of CO2 to Initiate Closing Responses upon Light off

The physiological role of substomatal CO2 changes for the stomatal adjustment to varying irradiance has been a matter of extensive debate (Mansfield and Meidner, 1966; Raschke, 1975; Wong et al., 1979; Sharkey and Raschke, 1981; Morison, 1987). The observation that over a wide range of photon flux densities, the substomatal CO2 concentration is stabilized by stomatal adjustment caused Raschke (1975) to assume that stomatal conductance is regulated by the substomatal CO2 concentration in a feedback loop. The assumed feedback loop should also restore substomatal CO2 when the ambient CO2 concentration is changed. However, in many plant species, after stomatal adjustment to increased ambient CO2, substomatal CO2 is at a higher level (Wong et al., 1979; Jarvis and Morison, 1981). After Morison (1987) had raised the question of whether there is a direct CO2 effect on stomata at all, investigations in guard cells of epidermal strips have demonstrated that a rise in CO2 by about 700 ␮L L⫺1 increases cytosolic Ca2⫹ activity (Ward et al., 1995; Webb et al., 1996) and regulates the conductance of plasma membrane ion channels so that the efflux of K⫹ and Cl⫺ is facilitated (Brearley et al., 1997). Our investigation provides data for a comparison between smaller CO2 steps and light off as triggers of guard cell Cl⫺ release in intact leaves. We have demonstrated that stepping up substomatal CO2 within physiologically relevant margins triggers guard cell Cl⫺ efflux. For instance, an increase of 250 ␮L L⫺1 CO2 leads to a Cl⫺ peak within 10 to 12 min (Figs. 4 and 5) and induces stomatal closure. Keeping substomatal CO2 constant by CO2 clamp, light off yields a comparable increase in Cl⫺ 948

activity and a closing response (Fig. 5). On the other hand, when substomatal CO2 is not kept constant, light off causes an increase in CO2 of 120 ␮L L⫺1 (physiological CO2 increase; see Fig. 1), which is roughly one-half of the CO2 step-up shown in Figure 5. From this, it follows that light off and a 2-fold physiological CO2 increase have approximately the same potential to trigger Cl⫺ efflux, i.e. to effectively initiate stomatal closure. The pronounced Cl⫺ efflux upon light off at constant substomatal CO2 (Fig. 5) demonstrates a strong potential of guard cells to initiate stomatal closure independent of CO2. However, the existence of an additional CO2-dependent closing trigger may be required to accelerate stomatal closure under certain environmental conditions: Based on analysis of air trapped in ice cores (Delmas et al., 1980), the atmospheric CO2 level of the past 165,000 years was found to shift between 190 and 280 ␮L L⫺1 (Tolbert, 1997). At a substomatal CO2 concentration of 150 ␮L L⫺1 compared with 440 ␮L L⫺1, we have clearly detected that light off becomes less efficient in triggering Cl⫺ release from guard cells. In most cases, the response was quickly initiated, but the Cl⫺ amplitude remained small (Fig. 4) or the response was delayed (Fig. 6). Under such conditions, selection for accelerated stomatal closing after shading stops photosynthetic carbon acquisition may have driven the evolution of supplementary closing mechanisms. Fluctuation in irradiance is common within plant canopies where leaves are shaded by other leaves of the same or of other plants, and where the shade moves with the sun or as a consequence of wind. In a forest or a crop canopy on clear days, 20% to 80% of the irradiance is received as light flecks with intermittent shading (Pearcy, 1990). It has been reported for trees of a tropical rainforest that stomatal conductance remains high in shade leaves subjected to periodic sunflecks, owing to the fact that the opening response of stomata after the shade-light transition shows a hysteretic nature that predominates the closing response after the subsequent light-shade transition (Pearcy and Calkin, 1983; Assmann et al., 1985; Pearcy, 1990). The CO2-dependent closing trigger may play a major role in terminating this lightinduced hysteretic opening response. Essentially two points emerge from these data. First, we have demonstrated in the intact leaf that upon light off a light-independent CO2 trigger acts in concert with a CO2-independent light off stimulus in promoting Cl⫺ release. The physiological role of the CO2 trigger at low substomatal CO2 concentrations and in fluctuating light deserves further investigation. Second, although some direct influence of CO2 on anion channels cannot be ruled out, the occurrence of pronounced lag phases before the onset of Cl⫺ efflux under conditions of low CO2 supply (Fig. 6: CO2 increase from 0 to 175 ␮L L⫺1) indicates that full channel Plant Physiol. Vol. 130, 2002

CO2-Triggered Cl⫺ Release from Guard Cells

activation requires a strongly time-dependent effector, possibly a product of carboxylation.

MATERIALS AND METHODS

Measuring Conditions Unless stated otherwise, measurements were performed at a photon flux of 300 ␮mol m⫺2 s⫺1, a relative air humidity of 95%, and at a temperature of 22°C. Stomatal aperture in leaves adapted to a CO2 concentration within the cuvette of 350 ␮L L⫺1 was approximately 6 ␮m (measured by a micrometer eyepiece).

Plants and Gas Exposure Fava beans (Vicia faba cultivar Witkiem Major; Nunhems Zaden bv, Haelen, The Netherlands) were grown in a glasshouse at a temperature of 20°C to 30°C in the day, 20°C at night, and at a relative air humidity of 60% ⫾ 5%; during the photoperiod of 13 h d⫺1, natural light was supplemented with 100 ␮mol m⫺2 s⫺1 (SONT-Assimilationsleuchte, Philips, The Netherlands). Plants were grown in 250-mL pots containing a mixture of peat, compost, and sand at a volume ratio of 2:2:1, fertilized with 250 mg of Triabon (BASF, Ludwigshafen, Germany). For the experiments, the 3rd or the 4th fully developed leaf from 4-week-old plants was cut from the stem with a razor blade and was placed immediately in the standard test solution (1 mm KCl and 0.1 mm NaCl and CaCl2 each) on a Plexiglas holder. To control the CO2 concentration at the leaf surface, a small flow-through cuvette was placed on one side of the leaf. Single-sided leaf cuvettes with a small volume have been useful for measuring fast transients in photosynthetic CO2 uptake (Pearcy, 1990). In the current investigation, a small cuvette with a tube-like shape (described by Hanstein et al., 2001) served to perform fast CO2 changes at the leaf surface. The transparent tube was mounted on the leaf with one oval hole (window) enclosing about 12 mm2 of the leaf surface. To achieve a tight fit between the window edge and the leaf, the leaf support, a flexible adhesive compound (plastic-fermit; Nissen and Volk, Hamburg, Germany) had been preformed to match the shape of the window edge. Sealing between the window edge and the leaf was accomplished by a weight fixed to the tube, which exerted a pressure of 4.5 kPa onto the leaf surface lying under the window edge (approximately 40 mm2). The sensors were positioned within the cuvette through a second cuvette window above the enclosed leaf surface. Air with preset CO2 concentrations passed the tube at a rate of 1.5 L min⫺1. Experiments were started 4 h after positioning the cuvette. During this period, the leaves were gradually adapted to a photon flux of 300 ␮mol m⫺2 leaf surface s⫺1 (white light). The light was supplied through a fiber cold-light source (KL 1500; Leica, Wetzlar, Germany), and photon flux was measured by the quantum sensor of an LCA 4 photosynthesis measurement system (ADC, Hoddesdon, Herts, UK).

Sensors The fabrication and measuring principle of the CO2 sensor has been described in detail previously (Hanstein and Felle, 2001). In brief, the sensor is built of two concentrically arranged capillaries, the inner one being a pH-sensitive microelectrode. The tip of this electrode is placed closely behind the tip of the outer capillary (diameter 2 ␮m), the opening of which is plugged with silicone (Dow Corning 3140 RTV Coating; Sasco Semiconductor, Frankfurt, Germany). CO2 diffuses through this plug, reacts with water, and acidifies the tip solution, which is detected by the pH electrode. To accelerate this reaction, carbonic anhydrase is added to the solution behind the plug. A Teflon-coated silver wire (AG-3T; Clark Electromedical Instruments, Reading, UK) leading from the carbonic anhydrase solution to ground serves as reference. Cl⫺-selective microelectrodes were built and used as described recently (Felle et al., 2000). In brief, blunt, heat-polished microcapillaries with a tip diameter of 2 ␮m were internally silanized with 0.2% (w/v) tributylchlorosilane (Fluka Chemical, Ronkonkoma, NY) dissolved in chloroform. Capillaries were backfilled first with a mixture of Cl⫺ sensor cocktail (24902; Fluka), tetrahydrofuran, and polyvinylchloride. After evaporation of the tetrahydrofuran, the remaining gel was topped up with the undiluted sensor cocktail followed by a 0.1 mm KCl buffer solution. CO2 sensor and Cl⫺ electrode were connected to high-impedance amplifiers (FD223; WP Instruments, Sarasota, FL), which simultaneously measured and subtracted the signals coming from the ion-selective electrodes and the voltage reference. The sensors were positioned with micromanipulators from Marzhauser (DC-3K; Wetzlar, Germany).

Plant Physiol. Vol. 130, 2002

ACKNOWLEDGMENT We thank D. Carden (University of Padova, Italy) for help with the preparation of the manuscript. Received February 15, 2002; returned for revision March 25, 2002; accepted June 3, 2002.

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