Transient alkalinization in the leaf apoplast of Vicia faba L. depends on NaCl stress intensity: an in situ ratio imaging study

Authors

  • CHRISTOPH-MARTIN GEILFUS,

    1. Institute of Plant Nutrition and Soil Science, Christian Albrechts University, Hermann-Rodewald-Str. 2, 24118 Kiel, Germany
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  • KARL HERMANN MÜHLING

    Corresponding author
    1. Institute of Plant Nutrition and Soil Science, Christian Albrechts University, Hermann-Rodewald-Str. 2, 24118 Kiel, Germany
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K. H. Mühling. Fax: +49 431 8801625; e-mail: khmuehling@plantnutrition.uni-kiel.de

ABSTRACT

The apoplast is suggested to be involved not only in the response, but also in the perception and transduction of various environmental signals. In this context, apoplastic alkalinization has previously been discussed as a general stress factor caused by abiotic and biotic stress events. In this study, an ion-sensitive fluorescence probe in combination with inverted fluorescence microscopy has been used for in planta monitoring of apoplastic shoot pH during challenging of Vicia faba L. plants by NaCl stress encountered at the roots. We demonstrate that transient increases in leaf apoplastic pH are dependent on the NaCl stress intensity. Moreover, we have visualized spatial pH gradients within the leaf apoplast. Our results indicate that these pH responses are propagated from root to leaf and that this occurs along the apoplast.

Abbreviations
MES

(2-[N-morpholino]ethanesulfonic acid)

INTRODUCTION

Terrestrial plants cannot choose the site in which they grow and thus have to endure a variety of environmental challenges, for example, salinity, drought or low temperatures (Shinozaki & Yamaguchi-Shinozaki 1997; Felle 2001). The apoplast is the first plant compartment that encounters abiotic or biotic stress signals (Gao et al. 2004; Felle et al. 2009) and is suggested to be involved not only in the response, but also in the perception and transduction of various environmental signals, as summarized by Hoson (1998). Information regarding an (ongoing) stress event at the roots might have to be rapidly transferred to the leaves. Such information has to be carried systematically and might be transformed into another signal to reach more distant plant organs such as the leaves (Felle et al. 2005). In this context, apoplastic alkalinization has been discussed as a general stress factor caused by abiotic and biotic stress events (Wilkinson & Davies 1997; Wilkinson 1999; Felle & Hanstein 2002; Felle 2001, 2005; Felle et al. 2005). Nevertheless, whether apoplastic pH is involved in signalling either directly or in crosstalk with plant hormones or calcium is unclear (Hartung & Radin 1989; Gilroy & Trewavas 1994; Roos 2000; Felle 2001; Gao et al. 2004; Monshausen et al. 2011).

In vivo measurements of the apoplastic pH are not straightforward, because of the problem of gaining access to the apoplastic fluid, which has been characterized by Felle & Hanstein (2002) as being an extremely thin film. Apoplastic pH can be measured by using diverse techniques, such as proton-selective electrodes (e.g. Felle 1998), fluorescence dyes (e.g. Hoffmann, Plänker & Mengel 1992; Mühling et al. 1995) or by collecting apoplastic fluid (e.g. Husted & Schjoerring 1995). The limited number of in vivo measurements of apoplastic pH changes available from intact plants under abiotic stress persuaded Gao et al. (2004) to produce transgenic Arabidopsis plants expressing pH indicators in the apoplast. To study extracellular pH dynamics, they targeted pHfluorins as fusion proteins to the apoplast by means of an Arabidopsis chitinase signal sequence (summarized in Minorsky 2004). In addition, Geilfus & Mühling (2011) have recently developed a technique for inserting ion-sensitive fluorescence dyes into the apoplast of intact plants. In combination with microscopy-based imaging, this dye-loading approach enables the high spatial-resolution quantitation of apoplastic pH dynamics over a period of several hours in situ. In the present study, this novel technique has been used for the ratiometric real-time monitoring of apoplastic pH in Vicia faba L. plants challenged by NaCl stress encountered at the roots. Our aim has been to investigate (1) whether the magnitude of a stress-induced pH response depends on the dose of the stress treatment and (2) whether this pH response occurs equally within the different regions of the leaf apoplast.

MATERIALS AND METHODS

Cultivation of plant material

Vicia faba L., minor cv. Scirocco (Saaten-Union GmbH, Isernhagen, Germany) was grown under hydroponic culture conditions in a climate chamber (14/10 h day/night; 20/15 °C; 50/60% humidity). Seeds were soaked in an aerated CaSO4 solution (0.5 mm) for 1 d at 25 °C and subsequently placed into quartz sand moistened with CaSO4 (1 mm). After 7 d of germination, seedlings were transferred to plastic pots containing one-quarter-strength aerated nutrient solution. After 2 d of cultivation, the concentration of nutrients was increased to half strength and, after 4 d of cultivation, to full strength. The nutrient solution had the following composition: 0.1 mm KH2PO4, 1.0 mm K2SO4, 0.2 mm KCl, 2.0 mm Ca(NO3)2, 0.5 mm MgSO4, 60 µm Fe-EDTA, 10 µm H3BO4, 2.0 µm MnSO4, 0.5 µm ZnSO4, 0.2 µm CuSO4, 0.05 µm (NH4)6Mo7O24. The solution was changed every 3rd day to avoid nutrient depletion. After 30 d of plant cultivation, growing Vicia faba L. plants were taken for in vivo pH recording.

Dye loading

For the purpose of the ratiometric in planta measurement of apoplastic pH, the fluorescent indicator was loaded into the apoplast of an intact plant as described in detail by Geilfus & Mühling (2011). Briefly, 25 µm Oregon Green 488-dextran (solubilized in water; Invitrogen GmbH, Darmstadt, Germany) was directly inserted into the apoplast by using a syringe. For this, the opening of the syringe (without needle) was carefully attached onto the surface of the relevant plant organ. By means of gentle pressure, the dye was loaded into the apoplast of the living plant. According to Geilfus & Mühling (2011), any excess of loaded water originating from the dye solution exits the apoplast through the stomata. Following this, measurements are conducted in a water-free apoplast. Images are collected (1) 2 h after loading and (2) not at the dye-loaded area itself but adjacent to the loaded area. The latter is possible because the dye is mobile within the leaf apoplast of the living plant and is carried towards the leaf edges as discussed by Geilfus & Mühling (2011).

Fluorescence microscopy imaging

Fluorescence images were collected as time series with a Leica inverted microscope (DMI6000B; Leica Microsystems, Wetzlar, Germany) connected to a DFC-camera (DFC 360FX; Leica Microsystems) via a 20-fold magnification, 0.4 numerical aperture, dry objective (HCX PL FLUOTAR L, Leica Microsystems). An HXP lamp (HXP Short Arc Lamp; Osram, München, Germany) was used for illumination at excitation wavelengths of 440/20 nm and 495/10 nm. The exposure time was 25 ms for both channels. The dye fluorescence at both excitation channels was collected by using a 535/25 nm-emission band-pass filter (BP 535/25; ET535/25M; Leica Microsystems) and a dichromatic mirror (LP518; dichroit T518DCXR BS, Leica Microsystems). During measurements, intact plants were supplied with aerated nutrient solution.

Ratiometric analysis

As a measure of pH, the fluorescence ratio F495/F440 was obtained by using the pH-sensitive fluophore Oregon Green 488 conjugated to 10 kDa dextran. In combination with the dextran, the dye molecule is not membrane permeable and thus does not enter the symplast. The F440 signal was captured because this fluorescence is almost insensitive to protons, whereas the F495 signal highly depends on protons. Image analysis was carried out by using LAS AF software (version 2.3.5, Leica Microsystems). Ratio images were calculated on a pixel-by-pixel basis as F495/F440. The background noise values were subtracted at each channel. For pseudo-colour display, the ratio was coded by hue on a spectral colour scale ranging from purple (no signal), over blue (lowest pH signal; pH 3.9), to pink (highest pH signal; pH 6.3), with the limits being set by an in situ calibration. Ratios below 1.1 (corresponds to pH ≤ 3.9) and above 3.6 (corresponds to pH ≥ 6.3) were not considered because they proved to lie outside the linear range of the in situ calibration. Quantitative measurements were calculated as the ratio of the mean intensity for user-defined regions of interest (ROIs). The high spatial resolution (Van Hoewyk 2011) allowed the quantification of pH within three apoplastic compartments, viz. the stomatal cavity, the epidermal apoplast and the palisade apoplast (Supporting Information Fig. S1).

In situ and in vitro apoplastic pH calibration

Two different calibration procedures for the conversion of fluorescence ratio data taken from living plants into apoplastic pH values were tested comparatively. For this, 25 µm Oregon Green dye solutions was pH buffered with 100 mm (2-[N-morpholino]ethanesulfonic acid) (MES) in order to obtain pH values ranging from 2.5 to 7.5 (in steps of 0.5 pH units). In situ calibration was carried out by loading the pH-buffered solutions into the leaf apoplast as described previously (see Dye loading section). Images were collected 2 h after the loading of the dye. For in vitro calibration, the pH-buffered solutions of Oregon Green 488 dextran were measured by using the same optical settings as described for the in situ calibration but without loading the solutions into a plant. The Boltzmann fit was chosen for fitting sigmoidal curves to the calibration data as described by Schulte et al. (2006). Fitting was performed by using Origin 7.0 (data not shown; OriginLab Corp., Northhampton, MA, USA).

Confocal laser scanning microscopy (CLSM)

To demonstrate that no Oregon Green 488 dextran had entered the cytosol and the vacuole unintentionally, CLSM imaging via a Leica TCS SP5 confocal laser scanning system (Leica Microsystems) and a Zeiss LSM 510 Axiovert 200M confocal laser scanning system (Carl Zeiss, Jena, Germany) was carried out. For Oregon Green excitation, the 488 nm beam line of the argon laser was chosen (emission bandwith was 498–540 nm for the Leica system and 505–530 nm for the Zeiss system). Chloroplast autofluorescence was excited at 633 nm by a helium-neon laser (emission bandwith was 650 nm–704 nm for the Leica system and 655–719 nm for the Zeiss system). A planapochromatic objective (HC PLAN APO 20.0 × 0.70; Leica Microsystems) and a planneofluar objective (planneofluar 20.0 × 0.5; Carl Zeiss) were used for image collection.

Gas exchange

Gas exchange parameters such as photosynthesis (µmol CO2 m−2 s−1) and intercellular CO2 concentration (µmol COmol−1 air) were measured with an open-flow gas-exchange system (Li-Cor Biosystems GmbH, Bad Homburg, Germany). Leaves were placed across a 2 × 3 cm leaf cuvette. The conditions for the measurements inside the chamber were equal to the outside conditions in the greenhouse: Light was provided by an LED red light source built into the top of the leaf chamber (100 µmol quanta m−2 s−1) and the CO2 concentration was controlled by a Li-Cor LI-6400 CO2 injection system (380 µmol COmol−1).

RESULTS AND DISCUSSION

pH calibration and dye loading into the apoplast of intact plants

For converting fluorescence ratio data taken from living plants into apoplastic pH values, two different calibration procedures were tested comparatively. With regard to in situ calibration, the pH-buffered calibration dye solutions were loaded into the apoplast of living plants by using the dye loading procedure described in the Materials and methods section. In contrast, calibration dye solutions were not loaded into the plant for in vitro calibration. Each of the calibration procedures gave different results, viz. the ratios for the identical pH values that were set by the pH-buffered calibration solution were different (Fig. 1). The shift of the in vitro calibration relative to the equivalent in situ calibration is possibly contributable to the fact that the emitted fluorescence intensities of the dye and the calculated ratios not only depend of the concentration of the quantified ion. Several studies have reported that ratio dyes behave differently in the presence of, for example, proteins (Harootunian et al. 1989; Mühling & Läuchli 2002). Moreover, it was found that the viscosity of the cytoplasm alters ratiometric measurements (Poenie et al. 1986). However, in this study the dye was not in contact with the cytoplasm, but the viscosity of the apoplastic fluid might contribute to the differences between both calibration procedures presented in Fig. 1. As the in situ calibration procedure accounts for optical effects of the specimen on the fluorescent signal intensity caused by, for example, leaf thickness, excitation path length (Tanasugarn et al. 1984; Bright et al. 1989; Gilroy 1997), apoplastic proteins and viscosity of the apoplastic fluid, the in situ calibration seemed to be more suitable for converting ratio data in pH values and thus was chosen for calculation. However, it is also cogitable that the buffer infused into the leaf apoplast does not maintain the desired pH due to an effect of the wall environment or the action of the proton pump on the proton concentration. Against this assumption is, however, that Mühling & Läuchli (2000) were able to buffer light-induced pH changes in the apoplast of Vicia faba plants with 80 mm MES. Moreover, in the recent study pH was buffered with a 100 mm MES buffer. For testing whether 100 mm MES sufficiently buffer changes in the apoplastic pH, the buffer was set to 4.7 and was loaded into the leaf apoplast (Supporting Information Fig. S2). By means of this, the leaf apoplastic pH was set to 4.7. In order to test whether the apoplast is adequately pH buffered, the roots were subsequently challenged by the addition of 75 mm NaCl, because it is known that such a salinization induces a transient leaf apoplastic alkalinization (Wilkinson & Davies 1997; Felle et al. 2005). However, the buffer was able to suppress this transient alkalinization. No pH alterations of the pH-buffered apoplast were detectable in response to the NaCl treatment (Supporting Information Fig. S2). This means that the chosen buffer strength can block expected changes in apoplastic pH induced by salt stress and, thus, is adequate for performing the in situ calibration. As it cannot be excluded that the buffer gets in equilibrium with the steady-state pH environment within the leaf, the given pH values should be viewed as approximate. Nevertheless, this does not detract from the biological meaning of how salt stress of the roots leads to changes in apoplastic pH in the leaves.

Figure 1.

Calibration curves for Oregon Green 488 dextran fluorescence excitation ratio R(490ex/440ex; 525em). Blue curve: in situ calibration; means ± SE. The Boltzmann fit was chosen for fitting sigmoidal curves to in situ calibration data. Fitting (data not shown) resulted in an optimal dynamic range for pH measurements between 3.9 and 6.3 (corresponds to the ratios 1.1 and 3.6). In situ calibration was conducted on six different plants (n = 6 biological replicates), each replicated five times (n = 5 technical replicates) with at least five regions of interest (ROIs) being quantified for each image (n ≥ 5). Red curve: in vitro calibration; means ± SE; (n = 10).

The insertion of the ion indicator into the plant's apoplast represents a potential event of mechanical stress and/or the presence of the dye within the apoplast might have an impact on the apoplastic space of the living plant. To exclude the occurrence of such negative effects, we have previously tested whether parameters such as leaf growth, stomatal conductance and the chlorophyll concentrations within leaves that have been dye loaded changed compared with leaves that have not been loaded with the ion indicator; neither of these parameters is influenced by the dye or the dye loading event (Geilfus & Mühling 2011). In addition, an effect of the dye loading procedure on photosynthesis and on the intercellular CO2 concentration has been tested; neither of these parameters is influenced (Fig. 2). As photosynthesis as a part of the primary metabolism of the plant is not affected (Fig. 2a), we have concluded that the pH measurements do not perturb the plants. The finding that apoplastic CO2 concentrations are identical in dye-loaded leaves and in leaves that have not been loaded (Fig. 2b) demonstrates that no water (originating from the loading step) remains in the apoplast and thus displaces the apoplastic air. This is attributable to the fact that the inserted water is leaving the apoplast over the stomata within a period of 2 to 10 min after dye loading (demonstrated by Geilfus & Mühling 2011). Unaffected apoplastic CO2 concentrations are of major significance for the measurement of apoplastic pH, as apoplastic CO2 fluctuations are known to affect the apoplastic pH immediately and quantitatively (Savchenko et al. 2000).

Figure 2.

Photosynthetic rate (a) and intercellular CO2 concentration (b) as influenced by the dye and the loading procedure. To test whether these parameters were affected, a paripinnate leaf was used. One leaflet was loaded with the dye (light grey), whereas the second was used as control (dark grey). Leaves were tested prior loading and at 2, 3 and 4 h after loading. Neither parameter was influenced by the presence of the dye or the loading procedure. Data are means of four biological replicates ± SE. ns = no statistical significance at P ≤ 0.05.

Leaf apoplastic pH transiently alkalizes in response to NaCl stress encountered by the root system

NaCl stress imposed on roots causes a systematic apoplastic alkalinization that is detectable in the leaves and is thought to be involved in the transmission of information from the site of the trigger (e.g. root) to distant plant tissues (e.g. shoot; Wilkinson & Davies 1997; Felle et al. 2005). In accordance with these findings, our ratiometric in planta quantitation of the leaf apoplastic pH established the occurrence of such an alkalinization when 20 mm NaCl was added to the nutrient solution of faba beans (Vica faba L.; Fig. 3). In response to the initiation of NaCl stress at the root site, leaf pH within the stomatal cavity, the epidermal apoplast and the apoplast surrounding the palisade tissue increased, whereas the pH within the palisade apoplast became the most alkaline. At 60 min after the stress was encountered, the nutrient solution was changed in order to remove the added NaCl. The removal of the stress stimulus out of the nutrient solution was followed by a decrease of the apoplastic pH. However, this pH decrease also occurred when NaCl was not removed as demonstrated in response to the second 20 mm NaCl application (see second transient pH increase within time-course shown in Fig. 3a). To determine that this transient pH increase was induced by the stress treatment and not by the illumination of the specimen by the dye-specific excitation or emission wavelengths (as discussed by Fricker et al. 1994), a time series of excitation at 495 and 440 nm was performed (Geilfus & Mühling 2011). No effects of the illumination regime on the apoplastic pH were detectable. This represents evidence that the transient pH increases as demonstrated in Fig. 3 are solely attributable to the salt treatment. Such an impact on the apoplastic pH is generally possible because the buffering capacity of the apoplast is, at 5 mm H+/pH units, relatively low compared with the 10 times higher buffering capacity of the cytoplasm (Hanstein & Felle 1999; Felle et al. 2008).

Figure 3.

Ratiometric real-time quantitation of leaf apoplastic pH when plants are challenged by the addition of 20 mm NaCl to the root system. (a) pH as recorded at the adaxial face of Vicia faba L. leaves is plotted over time. Black arrows indicate time of addition/removal of 20 mm NaCl stress stimulus in/out of the nutrient solution. Leaf apoplastic pH was discriminated within three apoplastic components, viz. in the stomatal cavity [n = 10 regions of interest (ROIs); green kinetic], in the epidermal apoplast (n = 20 ROI; red kinetic) and in the apoplast surrounding the palisade mesophyll (n = 20 ROI; blue kinetic). Ratiometric images (b–d) show time series of apoplastic leaf pH at (b) 20 min, (c) 50 min and (d) 100 min after measurement had started (time of image acquisition is presented in the upper right corner of the ratio images). Ratios were colour coded on a spectral colour scale (see inset beside d). Representative kinetics of three equivalent recordings of plants gained from independent experiments.

A comparable stress-induced apoplastic alkalinization was previously described by Felle & Hanstein (2002) who measured pH dynamics in Vicia faba L. leaves in response to a 20 mm NaCl treatment by using proton-sensitive microelectrodes. They demonstrated that the alkalinization dropped down to its initial level after the sodium chloride was removed. However, in the present study (Fig. 3), we demonstrated that the alkalinization was transient, although the stress stimulus was still present. This transient nature of the pH increase and the finding that the pH response is triggered at the root site and is detectable in distant leaves are indicative for a role in systemic signalling. Such an interpretation of the function of the previously described pH dynamics agrees with that of Felle et al. (2008) who describe transient apoplastic alkalinizations that are similar in shape and magnitude and are thought to be a pH signal. In detail, the authors demonstrate a transient leaf apoplastic alkalinization in response to the inoculation of Hordeum vulgare with conidia of the powdery mildew fungus Blumeris graminis f. sp. hordei (Felle et al. 2008; first pH peak presented in fig. 2 therein). This pH dynamic is interpreted to be a root-to-shoot signal, as it accompanies a simultaneous decrease in apoplastic Ca2+ (Felle et al. 2004), transient cytoplasmic acidifications and a likewise rapid increase in cytoplasmic Ca2+ activity (Felle et al. 2008). The aim of the apoplastic alkalinizations may be to signal the occurrence of stress events to distant plant organs (Wilkinson 1999). However, the explanation for the drop in pH to its initial level after stress-induced alkalinization fails at this point, as the mechanisms responsible for the pervious stress-induced alkalinization are not fully understood (Bacon, Wilkinson & Davies 1998). Felle & Hanstein (2002) propose that the idea of an NaCl-induced apoplastic alkalinization attributable to the stimulated uptake of the Cl- via a 2H+/Cl- plasma membrane symport is not valid, because salt-induced alkalinization also occurs in the presence of the membrane-impermeable gluconate- used instead of Cl- as the Na+-accompanying anion. As a more likely explanation for apoplastic pH increases in response to increased salt concentration, the authors report a reduction in H+-ATPase activity, as discussed by Hartung & Radin (1989) and Yang et al. (2004), or changes in strong ion concentrations that result in pH changes without reducing proton concentration, according to the concept of strong ions as derived for plants by Gerendás & Schurr (1995).

Magnitude of transient alkalinization depends on doses of NaCl stress treatment

As the application of abiotic stress in terms of 20 mm NaCl to the roots was shown to induce a transient pH response in distant plant organs such as the leaves, we tested whether the magnitude of this response changed with increasing dosages of NaCl. For this, 50 mm NaCl (Fig. 4 and corresponding Supporting Information Video Clip S1) or 75 mm NaCl (Fig. 5 and corresponding Supporting Information Video Clip S2) was added to the nutrient solution. As in the response to the 20 mm NaCl treatment, the pH was transiently alkalized over a period ranging from 40 to 60 min in the three apoplastic compartments, viz., the stomatal cavity, the epidermal apoplast and the palisade apoplast, starting circa 15 to 20 min after stress was initialized. Again, pH was the most alkaline within the apoplast built by the palisade tissue. The pseudo-colour ratios presented in the Supporting Information Video Clips S1 and S2 require additional comment. The ratio images under the most alkaline conditions suggest that fluorescent signals come directly from within the cells. This implies that, in addition to apoplastic pH signals, symplastic pH signals have been quantified erroneously. Such an idea can however be rejected: an XYZ-image stack captured by CLSM had clearly demonstrated that the Oregon Green dextran signals were solely emitted from the apoplast. The mesophyll cells appeared dark, providing evidence that no dye had unintentionally entered the cells (movie S1 presented by Geilfus & Mühling 2011). Another idea is that the dye had entered unintentionally the cytosol but not the vacuole. Following this assumption, the transient alkalinizations presented in Figs 3–5 could also be caused by the uptake of protons from the cytosol into the vacuole. For this, Oregon Green would need to access the cytosol from the apoplastic space across the plasmalemma membrane. However, this assumption can be rejected on the basis of a further confocal laser scanning image (Fig. 6). This image clearly demonstrates that no Oregon Green signals arise from within the cytosol. Assuming the dye had entered the cytosol unintentionally, the space between the cytosolic-located chloroplasts (oval structures presented in pseudo-red) would appear in pseudo-green. However, as indicated by the arrows, this particular space appears black (black = no signal; indicated by arrows in Fig. 6). This proves that no dye had entered the cytosol unintentionally, as otherwise signals would be detectable from between the chloroplast. Moreover, stress-related translocations of the dye across the plasma membrane from the apoplast in the symplast can also be excluded (Supporting Information Fig. S3).

Figure 4.

Ratiometric real-time quantitation of leaf apoplastic pH when plants are challenged by the addition of 50 mm NaCl to the root system. Left ordinate: leaf apoplastic pH as recorded at the adaxial face of Vicia faba L. leaves is plotted over time. Black arrow indicates time of addition of 50 mm NaCl stress stimulus in the nutrient solution. Leaf apoplastic pH was discriminated within three apoplastic components, viz. in the stomatal cavity [n = 10 region of interest (ROI); green kinetic], in the epidermal apoplast (n = 20 ROI; red kinetic) and in the apoplast surrounding the palisade mesophyll (n = 20 ROI; blue kinetic). Representative kinetics of three equivalent recordings of plants gained from independent experiments. The corresponding ratiometric images of this pH kinetic are also presented as a video clip (see Supporting Information Video Clip S1). Right ordinate: pH in nutrient solution as challenged by the addition of NaCl into the nutrient solution (black kinetic): pH in the nutrient solution was not affected by the addition of NaCl.

Figure 5.

Ratiometric real-time quantitation of leaf apoplastic pH when plants are challenged by the addition of 75 mm NaCl to the root system. pH as recorded at the adaxial face of Vicia faba L. leaves is plotted over time. Black arrow indicates time of addition of 75 mm NaCl stress stimulus in the nutrient solution. Leaf apoplastic pH was discriminated within three apoplastic components, viz. in the stomatal cavity [n = 10 regions of interest (ROI); green kinetic], in the epidermal apoplast (n = 20 ROI; red kinetic) and in the apoplast surrounding the palisade mesophyll (n = 20 ROI; blue kinetic). Representative kinetics of three equivalent recordings of plants gained from independent experiments. The corresponding ratiometric images of this pH kinetic are also presented as a video clip (see Supporting Information Video Clip S2).

Figure 6.

Oregon Green 488 dextran did not enter the symplast. Confocal image shows adaxial leaf apoplast of Vicia faba as labelled with Oregon Green 488 dextran. Oregon Green as excited at 488 nm by an argon laser (pseudo-green). Autofluorescence of the chloroplast as excited at 633 by a helium-neon laser (pseudo-red). #, palisade cells appear black as no Oregon Green 488 dextran has entered the cells. Assuming the dye had entered the cytosol unintentionally, the space between the chloroplast would appear in pseudo-green. As indicated by the arrows, this particular space is black. This proves that no dye had entered the cytosol unintentionally, as otherwise signals would be detectable from the cells.

Figures 3a, 4 and 5 clearly demonstrate a relationship between the NaCl concentration of the stress treatment and the pH dynamics: the more NaCl is added to nutrient solution, the stronger the magnitude of the leaf apoplastic alkalization. This is true for all three stress treatment intensities, viz. 20, 50 and 75 mm NaCl, and for all three apoplastic compartments (compared with Table 1). These varying NaCl stress treatments demonstrate (1) that a dependency exists between the magnitude of the apoplastic alkalinization and the severity of the NaCl stress treatments (dose dependency) and (2) that the leaf apoplast does not alkalizes equally over the leaf area but shows spatial differences.

Table 1.  Maximum increase of the leaf apoplastic pH shown as decrease (%) of the corresponding H+ concentration
 Palisade apoplastStomatal apoplastEpidermal apoplast
  1. The maximum decrease of the initial leaf apoplastic H+ concentration (H+ mol L−1) during the transient NaCl-induced alkalinization was calculated on the basis of delogarithmized pH values. The data show the percentage difference between the initial H+ concentration (initial = before NaCl application) and the H+ concentration at most alkaline pH for each apoplastic compartment (palisade-, stomatal-, and epidermal apoplast) and each NaCl-stress treatment intensities (20, 50 and 75 mm).

mm NaCl2060.1949.8849.88
5080.0574.8877.61
7590.0084.1587.41

What does the plant gain from a transient leaf apoplastic alkalinization?

Generally, the previously shown treatments demonstrate that the pH is more alkaline within the apoplast that is formed by the palisade cells compared with the stomatal cavity and the epidermal apoplast (Figs 3a, 4 & 5). Hence the question arises as to the nature of the gain to the plant of a pH increase within the (palisade) apoplast. In this context, the significance of the apoplastic pH for the distribution of abscisic acid (ABA) and, thus, in consequence for the closure of the stomata, needs to be considered. High pH and ABA are known to act in concert as reviewed by Wilkinson (1999): under normal (non-stressed) physiological conditions, ABA sourced from roots, stems or leaves is transported along the transpirational stream and is translocated to the leaves. Within the acid apoplastic milieu, the weak acid ABA is present in its undissociated form (ABAH). The lipophilic ABAH diffuses across the lipid plasma membrane of the cells. When ABAH enters the alkaline cytoplasm, it dissociates to ABA- + H+ and becomes trapped inside the cells because of its lipophobic character. The trapped ABA- can rapidly be catabolized, helping to maintain the concentration gradient for further ABA uptake (Kaiser & Hartung 1981; Gowing, Jones & Davies 1993; Daeter & Hartung 1995; Hartung, Radin & Hendrix 1998; Wilkinson 1999). However, in stressed plants, we have demonstrated that the apoplastic pH increases (Figs 3–5; see also other authors: Wilkinson & Davies 1997; Gao et al. 2004). This may in turn reduce the pH gradient across the plasma membrane for mesophyll and epidermal cells as explained by Wilkinson (1999). Moreover, the increased apoplastic pH causes undissociated weak acids to dissociate to the anionic form, which is then trapped within the alkalized apoplast (Heilmann, Hartung & Gimmler 1980). In consequence, ABA- accumulates in the apoplast and finally reaches the guard cells and causes stomatal closure (Schmidt et al. 1995; Wilkinson & Davies 1997; Jiang & Hartung 2008). In other words, a drought-, salt- or cold stress-induced apoplastic alkalinization indirectly causes the guard cell ABA receptors to become activated and thus the stomata to be closed, enabling the plant to retain water (Wilkinson 1999). In the current study, no ABA concentrations were measured, and thus an effect of the apoplastic alkalinization on the ABA concentration within the apoplast could not be demonstrated. However, taking into account (1) that ABA is synthesized in response to stress and is translocated within the apoplast; (2) that ABA mediates the responsiveness of plants to environmental stresses such as drought, cold or salt; and (3) that an alkalinization has been discussed as a signal in situations in which the plant must communicate the occurrence of a root-sourced stress event to the shoot (Felle 2001; Felle & Hanstein 2002; Becker et al. 2003; Wilkinson & Davies 2008), we consider it likely that the previously described alkalinization (Figs 3–5) acts on the ABA distribution within the plant. Moreover, as the mesophyll tissue is thought to be a major sink for cytosolic-trapped ABA (Wilkinson 1999), strong alkalinization around the palisade cells, as recorded in the Figs 3–5, might have a physiological meaning with respect to avoiding cytosolic ABA trapping.

In the current work, we have studied apoplastic pH dynamics in response to NaCl stress encountered at the roots of the faba beans. As the stress treatment is initiated by the addition of NaCl into the nutrient solution, we need to preclude that the application of the NaCl changes the pH of the nutrient solution itself. This is important because changes in the pH of the hydroponic growth media are known to be rapidly sensed by a so-called pH response system that can speedily alter the global patterns of gene expression accompanying Ca2+ signalling (Lager et al. 2010). Thus, changes in the pH of the external growth media are also likely to affect the pH in the plant apoplast. To exclude such an effect, we have demonstrated that the addition of NaCl to the nutrient solution does not alter the pH of the nutrient solution (Fig. 4, 2nd ordinate). Thus, we can conclude that the pH kinetics previously described is solely attributable to NaCl stress and not to changes in the pH of the nutrient solution.

Propagation of the pH response throughout the plant

Although apoplastic pH is broadly described to act as a stress signal, little is known about the signal translocation from root to shoot. In the current study, the recording of pH dynamics within the internodial stem apoplast has revealed the occurrence of a transient alkalinization in response to 50 mm NaCl stress application (Fig. 7). Interestingly, this alkalinization occurs as early as 10 min after stress application to the roots, whereas the pH response occurs no earlier than at 20 min in more elevated leaves (compared with Figs 3–5). The finding that the transient stress-induced pH response is detectable within the internodial apoplast and that it arrives 10 min earlier than in leaves located above these internodes is indicative of the ideas that (1) the pH response is propagated from root to shoot and (2) this occurs via the apoplast. The more distant the plant organs are from the site of the stress stimulus, the more time the alkalinization needs to occur.

Figure 7.

Ratiometric real-time quantitation of apoplastic pH within the internodes when plants are challenged by the addition of 50 mm NaCl to the root system. pH was plotted over time. Black arrow indicates time of addition of 50 mm NaCl stress stimulus in the nutrient solution. pH was averaged over the complete focused area (630 µm × 475 µm) of the specimen. Representative kinetics of three equivalent recordings of plants gained from independent experiments.

In this study, salt stress of the roots was shown to elicit a transient pH change in the leaves (Figs 3–5). However, Felle et al. (2005) and Felle (2001) discuss that it cannot be explained if the pH change is triggered by the roots and is directly transferred to the leaf apoplast as such or if pH changes are somehow else reacted to through the activity of regulatory forces. One possible idea is that the stress factor itself is transferred and causes pH changes in the leaf (Felle 2005). In this context, Felle et al. (2005) was demonstrating that a large part of 50 mm K+ added to the roots of barley plants reached within minutes the leaf apoplast (30 mm K+ or greater), whereas the kinetics of leaf apoplastic pK and pH changes were highly correlative. As a result, a causal relationship in which the apoplastic pH responds to changes in K+ concentration was suggested. The author was extrapolating this correlative relationship to apoplastic pNa and pH, resulting in the idea that the apoplastic pH would change in a close correlation to the Na+ concentration within the apoplast (Felle 2005). Thus, the pH changes (Figs 3–5) may go along with the translocation of the stress factor Na+ from root to shoot. We have recently tested whether the foliar application of NaCl is already sufficient to elicit a transient apoplastic pH change. For this, 50 µL of a 10 mm NaCl solution was added onto the surface of Vica faba L. leaves. Immediately after salt application, a transient apoplastic alkalinization was recorded (Geilfus & Mühling 2011, therein fig. 4). Magnitude and shape of the pH kinetic were similar to the kinetic recorded in response to the addition of 50 mm NaCl to the roots (Fig. 4). The only difference in the two pH kinetics was the time of the beginning of the alkalization: when NaCl was added to the roots, leaf pH increased at the earliest after 20 min, whereas the pH increased after 1–2 min when NaCl was added directly on the leaf. The occurrence of a transiently increasing pH within 1–2 min after NaCl was applied to the leaves strengthens the idea of a causal relationship in which the apoplastic pH responds to changes in the apoplastic Na+ concentration as, for example, hypothesized by Felle (2005).

Resume of the main conclusions

Here we demonstrate systemic effects of NaCl stress treatments encountered by the roots of Vica faba L. on leaf apoplastic pH. The transient nature of the increase in leaf apoplastic pH is indicative of its role in stress signalling. The data presented here suggest a dependency between the magnitude of the transient leaf apoplastic alkalinization and the NaCl stress treatment (dose dependency). In addition, we have demonstrated that the leaf apoplast does not alkalize equally over the leaf area but shows spatial differences as visualized by fluorescence ratio imaging.

ACKNOWLEDGMENTS

C-M.G. is the grateful recipient of a grant from the Friedrich-Ebert-Stiftung. We thank Dr Christoph Plieth (Center of Biochemistry and Molecular Biology, University of Kiel) for advice on fitting calibration data with a sigmoidal Boltzmann fit. We wish to thank Dr Maria Mulisch and Meike Dibberin (Zentrale Mikroskopie, University of Kiel) and Dr Uwe Bertsch (Institute of immunology, UKSH) for giving advices and helping on CLSM imaging and for providing the microscope.

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