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Slow vacuolar channels in vacuoles from winter and spring varieties of rape (Brassica napus)
ARTICLE IN PRESS Journal of Plant Physiology 165 (2008) 1511—1518
www.elsevier.de/jplph
Slow vacuolar channels in vacuoles from winter and spring varieties of rape (Brassica napus) Halina Dziubinskaa, Maria Filekb,c, Elzbieta Krola, Kazimierz Trebacza, a
Department of Biophysics, Institute of Biology, Maria Curie-Skłodowska University, Akademicka 19, PL-20-033 Lublin, Poland b Institute of Plant Physiology, Polish Academy of Sciences, Niezapominajek 21, PL-30-239 Krako ´w, Poland c Institute of Biology, Pedagogical University, Podbrzezie 3, 30-084 Krako ´w, Poland Received 14 June 2007; received in revised form 9 November 2007; accepted 12 November 2007
KEYWORDS Brassica napus; Generative development; Gibberellic acid; SV channels; Vacuole
Summary Currents passing through slowly activating vacuolar channels (SV) in isolated vacuoles from winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape (Brassica napus) were examined using the patch-clamp technique. Eight-week-long vernalization at 5/2 1C (day/night) was applied to obtain the generative stage of winter rape. SV channels of vacuoles isolated from vegetative (rosette) and generative leaves of both varieties were examined in order to investigate a possible role of these ion channels in rape flowering. Single SV channel conductance measured in a vacuole-attached configuration (natural cell sap) ranged from 60 to 83 pS. Lower values were observed in the generative leaves of both varieties. Unitary conductance measured in excised cytoplasm-out membrane patches did not differ significantly among the experimental variants, with the exception of spring generative vacuoles, where it was significantly lower. There was also no difference in SV current densities measured in the whole-vacuole configuration. Gibberellic acid (GA3) (2 mg/l) caused lowering of macroscopic SV currents by 20%, and had no significant effect on the single channel conductance. We conclude that SV channels play a role in rape vernalization and flowering owing to their multifactor regulation abilities rather than structural changes. & 2008 Elsevier GmbH. All rights reserved.
Introduction Abbreviations: GA3, gibberellic acid; I/V, current/voltage characteristics; SV, slowly activating vacuolar channels. Corresponding author. Tel.: +48 81 537 59 31; fax: +48 81 537 59 01. E-mail address: [email protected] (K. Trebacz).
Numerous factors control flowering time, and diverse signals are integrated to either delay or promote the normally irreversible process of generative development. Many plants survive winter in
0176-1617/$ - see front matter & 2008 Elsevier GmbH. All rights reserved. doi:10.1016/j.jplph.2007.11.007
ARTICLE IN PRESS 1512 the vegetative form and flower in spring. The acceleration of the flowering process as a result of a few months of cold temperature is a process called ‘‘vernalization’’ (Chouard, 1960). Cold acclimation is another process that is initiated by cold sensing. The purpose of cold acclimation is to prepare the plant to withstand low-temperature stress. In keeping with this purpose, cold acclimation is initiated quite rapidly – changes in gene expression occur within minutes of exposure to cold (Thomashow, 2001). In contrast to cold acclimation, vernalization typically requires a long period of cold exposure. Analysis of the vernalization pathway has defined a series of epigenetic regulators crucial for ‘‘cellular-memory’’ of the cold signal, whereas the autonomous pathway appears to function in part through post-transcriptional mechanisms (Henderson et al., 2003, Amasino, 2004). However, the molecular mechanisms underlying vernalization-induced winter annual and biennial plant flowering are still largely unknown (Yong et al., 2003). Winter rape, as other winter plants, requires vernalization for induction of flowering (Gregory and Purvis, 1936). During vernalization, several metabolic changes occur before morphological changes in the shoot apex appear. Winter rape grows as rosettes (vegetative stage) prior to receiving the generative stimulus. In plants with a rosette stage of development and vernalization response, one of the earliest detectable events following thermoinduction of flowering is the rapid increase in the level of endogenous gibberellins. Gibberellins normally play a role in the initiation of flowering in Arabidopsis thaliana (Wilson et al., 1992). In the Brassicaceae family, certain gibberellic acids (GA3) are known to increase in concentration during flowering after thermoinduction (Matzger, 1988). Dahanayake and Galwey (1999) suggested that vernalization responses of rape are probably mediated through gibberellins. Gibberellins, like other plant hormones, interact with specific receptors located in membranes. Such interaction can evoke changes in membrane potential and ion channel activities (Barbier-Brygoo et al., 1991). From our earlier experiments, it was suggested that 2,4-dichlorophenoxyacetic acid (synthetic auxin) caused significant changes in slow vacuolar channel conductance of embryogenic winter wheat cultures (Dziubin ´ska et al., 2003). Mechanosensitive-Ca2+-selective cation channels are strongly influenced by temperature (Ding and Pickard, 1993). Ca2+chelators and channel blockers have been shown to inhibit cold acclimation in alfalfa cultures (Monroy et al., 1993) and coldshock regulation of Arabidopsis genes (Polisensky
H. Dziubinska et al. and Braam, 1996). Thus, the influence of vernalization on changes in ion channel activity can be expected. Slowly activating vacuolar channels (SV) are ubiquitous in higher plants (Barkla and Pantoja, 1996; Allen and Sanders, 1997; Krol and Trebacz, 2000; Pottosin and Scho ¨nknecht, 2007). They exhibit typical slow kinetics of activation at positive transmembrane voltages and no timedependent inactivation. The currents at negative tonoplast potentials are very small, primarily due to low open probability of the SV channels. Thus, the SV channels are strongly rectifying and are suited to pass cations from the cytosol to the vacuole lumen. SV channels are activated by the cytoplasmic calcium ions at the micromolar range (Hedrich and Neher, 1987). The channels are permeable to potassium and to divalent cations, including Ca2+ (Pottosin et al., 2001, 2004). Slow vacuolar channels are modulated by a variety of physical and chemical factors such as calmodulin, phosphorylation, pH, reducing and oxidizing agents, polyamines, Mg2+, heavy metals, and 143-3 proteins (Davies and Sanders, 1995; Bethke and Jones, 1997; Carpaneto et al., 1999; Dobrovinskaya et al., 1999; Pei et al., 1999; Carpaneto et al., 2001; van den Wijngaard et al., 2001; Wherrett et al., 2005). This makes the SV channels important factors in cell signaling. Their roles in a variety of physiological processes, such as stomata movement, salt tolerance, and germination, have been examined (Pei et al., 1999; Ivashkina and Hedrich, 2005; Peiter et al., 2005). However, their detailed physiological role remains unclear. SV is the vacuolar channel that has recently been characterized on the molecular level (Peiter et al., 2005). Thus, rapid progress can be anticipated in understanding their roles in plant cell processes. In this paper, we present a comparison of SV channel parameters from vegetative and generative leaves of winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape. The influence of GA3 on SV channels is also demonstrated. The aim of this study was to examine a possible role of SV channels in vernalization and flowering.
Material and methods Plant material After sterilization and germination on blotting paper, seeds of winter rape cv. ‘Go ´rczan ´ski’ were placed in pots containing a mixture of peat, soil and sand in the ratio 1:1:1, transferred into a conditioned chamber, and kept at a temperature of 17/15 1C day/night (16 h day).
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The plants were grown under these conditions for about 4 weeks, until they reached the stage of 5-leaf rosettes – the vegetative development stage that is optimal for vernalization of winter rape. In this developmental stage, the plants were vernalized for 8 weeks at the temperature of 5/2 1C day/night (16 h day). After vernalization, the winter and spring plants were transferred back to the temperature of 17/15 1C day/night (16 h day) and grown until generative leaves appeared on the extended shoots. Spring rape cv. ‘Młochowski’ was cultured continuously at 17/15 1C day/night (16 h day), because spring plants do not require a cooling period for the induction of flowering. The youngest vegetative and generative leaves of winter and spring rape were used for vacuole isolation and ion-channel activity measurements.
The patch-clamp amplifier EPC-9 (Heka Elektronik, Germany) running under PULSE (Heka Elektronik, Germany) was used for data registration and acquisition. The convention of current and voltage signs was according to Bertl et al. (1992). Membrane potentials in vacuoleattached configurations were calculated by solving second-order regression equations applied to fit the single channel current/voltage characteristics (I/V) data. Considerable equilibration of the isolation (and bath) solution with the vacuolar sap during the isolation procedure and giga-seal establishing was assumed. Liquid junction potentials were calculated using JPCalc software (written by P.H. Barry).
Isolation of vacuoles
In media containing a high concentration of Ca2+ on the cytoplasmic side (2 mM), the current passing through the tonoplast was dominated by SV channel conductance. In the whole-vacuole configuration, membrane potentials between +100 and 100 mV elicited macroscopic currents that displayed a marked rectification and slow activation (Figure 1). An instantaneous current recorded at the onset of voltage pulses was rather a result of non-specific leak. The abundant vacuolar cationselective fast-activating (FV) channel cannot be active at such a high cytosolic Ca2+ concentration. The other known fast-activating K+-selective channel, VK, has so far been found only in guard cells and red beet taproot vacuoles (Allen et al., 1998; Pottosin et al., 2003). Nothing is known about its activity at milimolar [Ca2+]cyt. To analyze SV currents, we subtracted the instantaneous current from the total and plotted the time-dependent current/voltage, (I/V) dependence (Figure 1B). Slowly activating currents showed nearly perfect outward rectification. Activation of SV currents occurred at potentials positive to +20 mV. Records of ion currents passing through macropatches in the cytoplasm-out configuration exhibited similar kinetics (Figure 2). At negative potentials, where the number of open channels is low, it was possible to distinguish single channel openings in a form of square-like transitions (Figure 2, inset). The concept that macroscopic currents are passing through SV channels was confirmed further by records of small membrane patches in the vacuole-attached and cytoplasm-out configurations (Figure 3). The currents consisted of single or multiple channel openings with a higher open probability and a higher number of simultaneously open channels at positive potentials compared with those at negative voltages. It was also possible to observe timedependent activation of single channel ion currents at positive tonoplast potentials.
The vacuoles of rape were isolated by the method described by Trebacz and Scho ¨nknecht (2000). Fragments of leaves were plasmolysed in a medium containing 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2, 500 mM sorbitol. After 1 h incubation in the plasmolysing medium, the leaf fragments were cut with a sharp razor blade perpendicular to the main plane. Incision opened the cell walls of the plasmolysed cells and protoplasts emerged during a stepwise deplasmolysis. Further reduction of the osmolality of the perfusion solution to 300 mOsm/kg caused rupturing of protoplasts and isolation of vacuoles. Most of the vacuoles remained attached to the fragments of cells, which allowed identification of their origin. Patch-clamp experiments Patch-clamp experiments were performed as described by Trebacz and Scho ¨nknecht (2000). Three basic configurations of the patch-clamp method were applied: ‘‘vacuole-attached’’, ‘‘whole-vacuole’’ and ‘‘cytoplasmside-out’’ analogous to ‘‘cell-attached’’, ‘‘whole-cell’’ and ‘‘outside-out’’, respectively (Hamill et al., 1981). An Ag/AgCl reference electrode was connected to the bath solution via a 2% agar bridge filled with 100 mM KCl solution. Microelectrodes were pulled of borosilicate glass capillaries 1.5 mm in outer diameter Kimax 51 (Kimble Products, USA) using a vertical two-step puller PP-830 (Narishige, Japan). The tips were fire-polished by a microforge MF 200-2 (World Precision Instruments, USA). The micropipettes were filled with a filtered solution containing 100 mM KCl, 2 mM CaCl2, 15 mM Mes/Tris, pH 5.85, directly before the experiment. If not otherwise indicated, the bath solution was composed of 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2. Osmolality of all solutions was adjusted with sorbitol to 300 mOsm/kg using a cryoscopic osmometer (Osmomat 030, Gonotec, Germany). A giga-seal was obtained by releasing the positive pressure applied to the patchpipette just before its tip touched a vacuole. Vacuolar ion channels were measured under voltage-clamp conditions.
Results and discussion
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Figure 1. (A) Macroscopic slow vacuolar currents in the tonoplast of rape (winter vegetative) leaves in a whole-vacuole configuration. Bottom: SV currents elicited by a series of voltage steps (top) ranging from 100 to +100 mV. Pipette solution contained 100 mM KCl, 2 mM CaCl2, 15 mM Mes/Tris, pH 5.85. Bath solution consisted of 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2. (B) Time-dependent current – voltage characteristics of slow vacuolar channels of the tonoplast in winter vegetative leaves. Currents passing at the last second of the records were measured and instantaneous current at the onset of the voltage pulses was subtracted. Bars denote standard errors. n ¼ 6 vacuoles.
To examine the difference in the behavior of SV channels between vegetative and generative leaf vacuoles of winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape, we measured single channel activities in small membrane patches in the vacuole-attached configuration and compared single channel I/V curves. Figure 4 shows I/V curves of the four experimental variants. These I/V curves are typical of SV channels from other higher plants (Pottosin et al., 2005; Wherrett et al., 2005), with higher conductance at negative potentials in comparison to that at positive voltages. Unitary conductance of SV channels measured at 100 mV ranging between 60 and 83 pS was relatively small. This value is known to vary between 40 and 280 pS among different plant species and plant tissues (Hedrich and Neher, 1987; Pottosin et al., 1997; Schulz-Lessdorf and Hedrich, 1995). To investigate the voltage dependence of SV channel unitary
conductance, we plotted the appropriate curves (Figure 5). Unitary conductance was extrapolated at 0 mV – close to natural tonoplast potential: 0.170.4 mV in spring vegetative, 0.670.4 mV in winter vegetative, 0.470.3 mV in spring generative, 0.070.3 mV in winter generative. It equaled 73.073.9 pS for spring vegetative vacuoles, 74.374.1 pS for winter vegetative vacuoles, 52.27 3 pS for spring generative and 57.371.9 pS for winter generative vacuoles. Single channel conductance in generative leaves was significantly lower than in vegetative leaf vacuoles. There was also a tendency for the specific conductance of SV channels in the vacuoles of winter plants to be higher than in spring plants. The same approach applied to SV channels in small cytoplasm-out patches yielded the following values of single channel conductance (extrapolated at 0 mV): 58.774 pS for spring vegetative vacuoles,
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Figure 2. Ion currents flowing through an isolated macro-patch of the tonoplast of rape (spring vegetative) leaves in the cytoplasm-out configuration. Inset: record in expanded scales of ion currents passing through single channels at 100 mV. Top: voltage protocol – after a prepulse of 60 mV voltage steps from 100 to +100 mV were applied. Pipette and bath solutions as in Figure 1.
Figure 3. Single SV current recordings from the tonoplast patches of rape (winter vegetative) leaves in the vacuole-attached (A) and cytoplasm-out (B) configurations. Numbers on the left side indicate command potentials. Presented traces begin 50 ms after the onset of voltage pulses. Membrane potential in A: 0.3 mV. Pipette and bath solutions as in Figure 1.
48.375.9 pS for winter vegetative vacuoles, 40.37 4.8 pS for spring generative and 5575.4 pS for winter generative vacuolar patches. There were no statistically significant differences among the groups except for the spring generative vacuoles. The lack of difference in SV channel conductance between vegetative and generative vacuoles in well controlled cytoplasm-out patches with respect to the conductance in the vacuole-attached mode can be explained by differences in the natural vacuolar sap composition in intact vacuoles, and the pipette solution in the cytoplasm-out patches. The comparison of current densities measured in the four experimental variants in the wholevacuole configuration revealed no statistically significant differences among the groups. Figure 6 presents current densities (in A/m2) calculated for +100 mV. Here again, the ion composition of the solutions on both sides of the vacuolar membrane was well defined and differed from natural values. Application of a high Ca2+ concentration (2 mM) in pipette and bath solutions reduced currents passing through the SV channels in comparison to low Ca2+ in the vacuole lumen (Pottosin et al., 2004). Under the experimental conditions differing from optimal, when one might expect current saturation, differences in SV activity among the examined variants should be more expressed. However, the current densities remain relatively constant. The effect of externally added phytohormone GA3 (2 mg/dm3 dissolved in the standard solution) was examined on the single channel level and macroscopic currents in cytoplasm-out macropatches of winter vegetative vacuoles. There was
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Figure 4. Current – voltage characteristics of single SV channels from rape vacuoles. Recordings were obtained in the vacuole-attached configuration. Black circles, spring vegetative (n ¼ 9 vacuoles), open circles, winter vegetative (n ¼ 10 vacuoles), black squares, spring generative (n ¼ 31 vacuoles), and open squares, winter generative vacuoles (n ¼ 13 vacuoles). Bars denote standard errors. Pipette and bath solutions as in Figure 1.
Figure 5. Specific conductance/voltage characteristics of SV channels in rape vacuoles. The data presented in Figure 4 were used to plot the graph. Lines represent linear regression curves. Symbols indicating experimental variants are the same as in Figure 4.
ARTICLE IN PRESS Ion channels in rape vacuoles
Figure 6. Current densities of time-dependent currents recorded in the whole-vacuole configuration at +100 mV. s–v, spring vegetative (n ¼ 5 vacuoles); w–v, winter vegetative (n ¼ 6 vacuoles); s–g, spring generative (n ¼ 3 vacuoles); w–g, winter generative (n ¼ 3 vacuoles). Error bars denote SE. Vacuole diameter was measured under the microscope. Surface area of the vacuoles was calculated assuming a spherical shape.
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Figure 8. Current – voltage characteristics of isolated cytoplasm-out macropatches of winter vegetative rape leaf vacuoles. Open circles – control (n ¼ 6 vacuoles), black circles – GA3 (2 mg/l) treated vacuoles (n ¼ 8 vacuoles).
rape in well-controlled experimental conditions exhibit similar properties. However, the differences between vegetative and generative leaf vacuoles appear in the vacuole-attached configuration, indicating that the vacuolar sap composition, which changes during floral induction, is an important regulating factor affecting SV channel activity.
References
Figure 7. Current – voltage characteristics of isolated cytoplasm-out patches of winter vegetative rape leaf vacuoles. Open circles – control, black circles – GA3 (2 mg/l)-treated vacuoles.
no significant difference in single channel conductance between the respective controls and GA3treated vacuoles (Figure 7). On the other hand, macroscopic currents at positive potentials were significantly lower (80% of the control) in the GA3treated vacuoles (Figure 8). It is known that SV currents exhibit a time-dependent run down. To avoid this effect, we added GA3 to the vacuole isolation medium, which allowed us to perform measurements on the control and GA3-treated vacuoles on a comparable time scale. These data indicate that GA3 added externally leads to a lower probability of SV channels opening. We conclude that SV channels of generative and vegetative leaves of spring and winter varieties of
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ARTICLE IN PRESS 1518 Chouard P. Vernalization and its relations to dormancy. Annu Rev Plant Physiol 1960;11:191–238. Dahanayake SR, Galwey NW. Effects of interactions between low-temperature treatments, gibberellin (GA3) and photoperiod on flowering and stem height of spring rape (Brassica napus var. annua). Ann Bot 1999;84:321–7. Davies JM, Sanders D. ATP, pH and Mg2+ modulate a cation current in Beta vulgaris vacuoles: a possible shunt conductance for the vacuolar H+-ATPase. J Membr Biol 1995;145:75–86. Ding JP, Pickard BG. Mechanosensory calcium-selective cation channels in epidermal cells. Plant J 1993;3: 83–110. Dobrovinskaya OR, Muniz J, Pottosin II. Inhibition of ion channels by polyamines. J Membr Biol 1999;167: 127–40. Dziubin ˛ K. ´ska H, Filek M, Szechyn ´ska-Hebda M, Trebacz Slow vacuolar channels of non-embryogenic and ambryogenic cultures of winter wheat. Acta Physiol Plant 2003;25:179–84. Gregory FG, Purvis ON. Vernalization. Nature 1936; 138:249. Hamill OP, Marty A, Neher E, Sakmann B, Sigworth FJ. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane + patches. Pflugers Arch 1981;391:85–100. Hedrich R, Neher E. Cytoplasmic calcium regulates voltage-dependent ion channels in plant vacuoles. Nature 1987;329:833–5. Henderson IR, Shindo C, Dean C. The need for winter in the switch to flowering. Ann Rev Gen 2003;37:371–92. Ivashkina N, Hedrich R. K+ current through SV-type vacuolar channels are sensitive to elevated luminal sodium levels. Plant J 2005;41:606–14. Krol E, Trebacz K. Ways of ion channel gating in plant cells. Ann Bot 2000;86:449–69. Matzger JD. Localization of the site of perception of thermoinductive temperatures in Thlaspi araense L. Plant Physiol 1988;88:424–8. Monroy AF, Sarhan F, Dhindsa RS. Cold-induced changes in freezing tolerance, protein phosphorylation, and gene expression. Evidence for the role of calcium. Plant Physiol 1993;102:1227–35. Pei ZM, Ward JM, Schroeder JI. Magnesium sensitizes slow vacuolar channels to physiological cytosolic calcium and inhibits fast vacuolar channels in fava bean guard cell vacuoles. Plant Physiol 1999;121:977–86. Peiter E, Maathuis FJM, Mills LN, Knight H, Pelloux J, Hetherington AN, et al. The vacuolar Ca2+-activated
H. Dziubinska et al. channel TPC1 regulates germination and stomatal movement. Nature 2005;434:404–8. Polisensky DH, Braam J. Cold-shock regulation of the Arabidopsis TCH genes and the effects of modulating intracellular calcium levels. Plant Physiol 1996;111: 1271–9. Pottosin II, Tikhonova LI, Hedrich R, Scho ¨nknecht R. Slowly activating vacuolar channels cannot mediate Ca2+-induced Ca2+ release. Plant J 1997:1387–98. Pottosin II, Dobrovinskaya OR, Muniz J. Conduction of monovalent and divalent cations in the slow vacuolar channel. J Membr Biol 2001;181:55–65. Pottosin II, Martines-Esteves M, Dobrovinskaya OR, Muniz J. Potassium-selective channel in the red beet vacuolar membrane. J Exp Bot 2003;54:663–7. Pottosin II, Martines-Esteves M, Dobrovinskaya OR, Muniz J, Scho ¨nknecht R. Mechanism of luminal Ca2+ and Mg2+ action on the vacuolar slowly activating channels. Planta 2004;219:1057–70. Pottosin II, Martines-Esteves M, Dobrovinskaya OR, Muniz J. Regulation of the slow vacuolar channel by luminal potassium: role of surface charge. J Membr Biol 2005;205:103–11. Pottosin II, Scho ¨nknecht G. Vacuolar calcium channels. J Exp Bot 2007;58:1559–69. Schulz-Lessdorf B, Hedrich R. Protons and calcium modulate SV-type channels in the vacuolar-lysosomal compartment-channel interaction with calmodulin inhibitors. Planta 1995;197:655–71. Thomashow MF. So what’s new in the field of plant cold acclimation? Lots!. Plant Physiol 2001;125: 89–93. Trebacz K, Scho ¨nknecht G. Simple method to isolate vacuoles and protoplasts for patch-clamp experiments. Protoplasma 2000;213:39–45. van den Wijngaard PWJ, Bunney TD, Roobeek I, Scho ¨nknecht G, de Boer AH. Slow vacuolar channels from barley mesophyll cells are regulated by 14-3-3 proteins. FEBS Lett 2001;488:100–4. Wherrett T, Shabala S, Pottosin I. Different properties of SV channels in root vacuoles from near isogenic Altolerant and Al-sensitive wheat cultivars. FEBS Lett 2005;579:6890–4. Wilson RN, Heckman JW, Somerville CR. Gibberellin is required for flowering in Arabidopsis thaliana under short days. Plant Physiol 1992;100:403–8. Yong W, Xu Y, Xu W, Wang X, Li N, Wu J, et al. Vernalization-induced flowering in wheat is mediated by a lecitin-like gene VER2. Planta 2003;217: 261–70.
www.elsevier.de/jplph
Slow vacuolar channels in vacuoles from winter and spring varieties of rape (Brassica napus) Halina Dziubinskaa, Maria Filekb,c, Elzbieta Krola, Kazimierz Trebacza, a
Department of Biophysics, Institute of Biology, Maria Curie-Skłodowska University, Akademicka 19, PL-20-033 Lublin, Poland b Institute of Plant Physiology, Polish Academy of Sciences, Niezapominajek 21, PL-30-239 Krako ´w, Poland c Institute of Biology, Pedagogical University, Podbrzezie 3, 30-084 Krako ´w, Poland Received 14 June 2007; received in revised form 9 November 2007; accepted 12 November 2007
KEYWORDS Brassica napus; Generative development; Gibberellic acid; SV channels; Vacuole
Summary Currents passing through slowly activating vacuolar channels (SV) in isolated vacuoles from winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape (Brassica napus) were examined using the patch-clamp technique. Eight-week-long vernalization at 5/2 1C (day/night) was applied to obtain the generative stage of winter rape. SV channels of vacuoles isolated from vegetative (rosette) and generative leaves of both varieties were examined in order to investigate a possible role of these ion channels in rape flowering. Single SV channel conductance measured in a vacuole-attached configuration (natural cell sap) ranged from 60 to 83 pS. Lower values were observed in the generative leaves of both varieties. Unitary conductance measured in excised cytoplasm-out membrane patches did not differ significantly among the experimental variants, with the exception of spring generative vacuoles, where it was significantly lower. There was also no difference in SV current densities measured in the whole-vacuole configuration. Gibberellic acid (GA3) (2 mg/l) caused lowering of macroscopic SV currents by 20%, and had no significant effect on the single channel conductance. We conclude that SV channels play a role in rape vernalization and flowering owing to their multifactor regulation abilities rather than structural changes. & 2008 Elsevier GmbH. All rights reserved.
Introduction Abbreviations: GA3, gibberellic acid; I/V, current/voltage characteristics; SV, slowly activating vacuolar channels. Corresponding author. Tel.: +48 81 537 59 31; fax: +48 81 537 59 01. E-mail address: [email protected] (K. Trebacz).
Numerous factors control flowering time, and diverse signals are integrated to either delay or promote the normally irreversible process of generative development. Many plants survive winter in
0176-1617/$ - see front matter & 2008 Elsevier GmbH. All rights reserved. doi:10.1016/j.jplph.2007.11.007
ARTICLE IN PRESS 1512 the vegetative form and flower in spring. The acceleration of the flowering process as a result of a few months of cold temperature is a process called ‘‘vernalization’’ (Chouard, 1960). Cold acclimation is another process that is initiated by cold sensing. The purpose of cold acclimation is to prepare the plant to withstand low-temperature stress. In keeping with this purpose, cold acclimation is initiated quite rapidly – changes in gene expression occur within minutes of exposure to cold (Thomashow, 2001). In contrast to cold acclimation, vernalization typically requires a long period of cold exposure. Analysis of the vernalization pathway has defined a series of epigenetic regulators crucial for ‘‘cellular-memory’’ of the cold signal, whereas the autonomous pathway appears to function in part through post-transcriptional mechanisms (Henderson et al., 2003, Amasino, 2004). However, the molecular mechanisms underlying vernalization-induced winter annual and biennial plant flowering are still largely unknown (Yong et al., 2003). Winter rape, as other winter plants, requires vernalization for induction of flowering (Gregory and Purvis, 1936). During vernalization, several metabolic changes occur before morphological changes in the shoot apex appear. Winter rape grows as rosettes (vegetative stage) prior to receiving the generative stimulus. In plants with a rosette stage of development and vernalization response, one of the earliest detectable events following thermoinduction of flowering is the rapid increase in the level of endogenous gibberellins. Gibberellins normally play a role in the initiation of flowering in Arabidopsis thaliana (Wilson et al., 1992). In the Brassicaceae family, certain gibberellic acids (GA3) are known to increase in concentration during flowering after thermoinduction (Matzger, 1988). Dahanayake and Galwey (1999) suggested that vernalization responses of rape are probably mediated through gibberellins. Gibberellins, like other plant hormones, interact with specific receptors located in membranes. Such interaction can evoke changes in membrane potential and ion channel activities (Barbier-Brygoo et al., 1991). From our earlier experiments, it was suggested that 2,4-dichlorophenoxyacetic acid (synthetic auxin) caused significant changes in slow vacuolar channel conductance of embryogenic winter wheat cultures (Dziubin ´ska et al., 2003). Mechanosensitive-Ca2+-selective cation channels are strongly influenced by temperature (Ding and Pickard, 1993). Ca2+chelators and channel blockers have been shown to inhibit cold acclimation in alfalfa cultures (Monroy et al., 1993) and coldshock regulation of Arabidopsis genes (Polisensky
H. Dziubinska et al. and Braam, 1996). Thus, the influence of vernalization on changes in ion channel activity can be expected. Slowly activating vacuolar channels (SV) are ubiquitous in higher plants (Barkla and Pantoja, 1996; Allen and Sanders, 1997; Krol and Trebacz, 2000; Pottosin and Scho ¨nknecht, 2007). They exhibit typical slow kinetics of activation at positive transmembrane voltages and no timedependent inactivation. The currents at negative tonoplast potentials are very small, primarily due to low open probability of the SV channels. Thus, the SV channels are strongly rectifying and are suited to pass cations from the cytosol to the vacuole lumen. SV channels are activated by the cytoplasmic calcium ions at the micromolar range (Hedrich and Neher, 1987). The channels are permeable to potassium and to divalent cations, including Ca2+ (Pottosin et al., 2001, 2004). Slow vacuolar channels are modulated by a variety of physical and chemical factors such as calmodulin, phosphorylation, pH, reducing and oxidizing agents, polyamines, Mg2+, heavy metals, and 143-3 proteins (Davies and Sanders, 1995; Bethke and Jones, 1997; Carpaneto et al., 1999; Dobrovinskaya et al., 1999; Pei et al., 1999; Carpaneto et al., 2001; van den Wijngaard et al., 2001; Wherrett et al., 2005). This makes the SV channels important factors in cell signaling. Their roles in a variety of physiological processes, such as stomata movement, salt tolerance, and germination, have been examined (Pei et al., 1999; Ivashkina and Hedrich, 2005; Peiter et al., 2005). However, their detailed physiological role remains unclear. SV is the vacuolar channel that has recently been characterized on the molecular level (Peiter et al., 2005). Thus, rapid progress can be anticipated in understanding their roles in plant cell processes. In this paper, we present a comparison of SV channel parameters from vegetative and generative leaves of winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape. The influence of GA3 on SV channels is also demonstrated. The aim of this study was to examine a possible role of SV channels in vernalization and flowering.
Material and methods Plant material After sterilization and germination on blotting paper, seeds of winter rape cv. ‘Go ´rczan ´ski’ were placed in pots containing a mixture of peat, soil and sand in the ratio 1:1:1, transferred into a conditioned chamber, and kept at a temperature of 17/15 1C day/night (16 h day).
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The plants were grown under these conditions for about 4 weeks, until they reached the stage of 5-leaf rosettes – the vegetative development stage that is optimal for vernalization of winter rape. In this developmental stage, the plants were vernalized for 8 weeks at the temperature of 5/2 1C day/night (16 h day). After vernalization, the winter and spring plants were transferred back to the temperature of 17/15 1C day/night (16 h day) and grown until generative leaves appeared on the extended shoots. Spring rape cv. ‘Młochowski’ was cultured continuously at 17/15 1C day/night (16 h day), because spring plants do not require a cooling period for the induction of flowering. The youngest vegetative and generative leaves of winter and spring rape were used for vacuole isolation and ion-channel activity measurements.
The patch-clamp amplifier EPC-9 (Heka Elektronik, Germany) running under PULSE (Heka Elektronik, Germany) was used for data registration and acquisition. The convention of current and voltage signs was according to Bertl et al. (1992). Membrane potentials in vacuoleattached configurations were calculated by solving second-order regression equations applied to fit the single channel current/voltage characteristics (I/V) data. Considerable equilibration of the isolation (and bath) solution with the vacuolar sap during the isolation procedure and giga-seal establishing was assumed. Liquid junction potentials were calculated using JPCalc software (written by P.H. Barry).
Isolation of vacuoles
In media containing a high concentration of Ca2+ on the cytoplasmic side (2 mM), the current passing through the tonoplast was dominated by SV channel conductance. In the whole-vacuole configuration, membrane potentials between +100 and 100 mV elicited macroscopic currents that displayed a marked rectification and slow activation (Figure 1). An instantaneous current recorded at the onset of voltage pulses was rather a result of non-specific leak. The abundant vacuolar cationselective fast-activating (FV) channel cannot be active at such a high cytosolic Ca2+ concentration. The other known fast-activating K+-selective channel, VK, has so far been found only in guard cells and red beet taproot vacuoles (Allen et al., 1998; Pottosin et al., 2003). Nothing is known about its activity at milimolar [Ca2+]cyt. To analyze SV currents, we subtracted the instantaneous current from the total and plotted the time-dependent current/voltage, (I/V) dependence (Figure 1B). Slowly activating currents showed nearly perfect outward rectification. Activation of SV currents occurred at potentials positive to +20 mV. Records of ion currents passing through macropatches in the cytoplasm-out configuration exhibited similar kinetics (Figure 2). At negative potentials, where the number of open channels is low, it was possible to distinguish single channel openings in a form of square-like transitions (Figure 2, inset). The concept that macroscopic currents are passing through SV channels was confirmed further by records of small membrane patches in the vacuole-attached and cytoplasm-out configurations (Figure 3). The currents consisted of single or multiple channel openings with a higher open probability and a higher number of simultaneously open channels at positive potentials compared with those at negative voltages. It was also possible to observe timedependent activation of single channel ion currents at positive tonoplast potentials.
The vacuoles of rape were isolated by the method described by Trebacz and Scho ¨nknecht (2000). Fragments of leaves were plasmolysed in a medium containing 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2, 500 mM sorbitol. After 1 h incubation in the plasmolysing medium, the leaf fragments were cut with a sharp razor blade perpendicular to the main plane. Incision opened the cell walls of the plasmolysed cells and protoplasts emerged during a stepwise deplasmolysis. Further reduction of the osmolality of the perfusion solution to 300 mOsm/kg caused rupturing of protoplasts and isolation of vacuoles. Most of the vacuoles remained attached to the fragments of cells, which allowed identification of their origin. Patch-clamp experiments Patch-clamp experiments were performed as described by Trebacz and Scho ¨nknecht (2000). Three basic configurations of the patch-clamp method were applied: ‘‘vacuole-attached’’, ‘‘whole-vacuole’’ and ‘‘cytoplasmside-out’’ analogous to ‘‘cell-attached’’, ‘‘whole-cell’’ and ‘‘outside-out’’, respectively (Hamill et al., 1981). An Ag/AgCl reference electrode was connected to the bath solution via a 2% agar bridge filled with 100 mM KCl solution. Microelectrodes were pulled of borosilicate glass capillaries 1.5 mm in outer diameter Kimax 51 (Kimble Products, USA) using a vertical two-step puller PP-830 (Narishige, Japan). The tips were fire-polished by a microforge MF 200-2 (World Precision Instruments, USA). The micropipettes were filled with a filtered solution containing 100 mM KCl, 2 mM CaCl2, 15 mM Mes/Tris, pH 5.85, directly before the experiment. If not otherwise indicated, the bath solution was composed of 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2. Osmolality of all solutions was adjusted with sorbitol to 300 mOsm/kg using a cryoscopic osmometer (Osmomat 030, Gonotec, Germany). A giga-seal was obtained by releasing the positive pressure applied to the patchpipette just before its tip touched a vacuole. Vacuolar ion channels were measured under voltage-clamp conditions.
Results and discussion
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Figure 1. (A) Macroscopic slow vacuolar currents in the tonoplast of rape (winter vegetative) leaves in a whole-vacuole configuration. Bottom: SV currents elicited by a series of voltage steps (top) ranging from 100 to +100 mV. Pipette solution contained 100 mM KCl, 2 mM CaCl2, 15 mM Mes/Tris, pH 5.85. Bath solution consisted of 100 mM KCl, 2 mM CaCl2, 15 mM Hepes/Tris, pH 7.2. (B) Time-dependent current – voltage characteristics of slow vacuolar channels of the tonoplast in winter vegetative leaves. Currents passing at the last second of the records were measured and instantaneous current at the onset of the voltage pulses was subtracted. Bars denote standard errors. n ¼ 6 vacuoles.
To examine the difference in the behavior of SV channels between vegetative and generative leaf vacuoles of winter (Go ´rczan ´ski) and spring (Młochowski) varieties of rape, we measured single channel activities in small membrane patches in the vacuole-attached configuration and compared single channel I/V curves. Figure 4 shows I/V curves of the four experimental variants. These I/V curves are typical of SV channels from other higher plants (Pottosin et al., 2005; Wherrett et al., 2005), with higher conductance at negative potentials in comparison to that at positive voltages. Unitary conductance of SV channels measured at 100 mV ranging between 60 and 83 pS was relatively small. This value is known to vary between 40 and 280 pS among different plant species and plant tissues (Hedrich and Neher, 1987; Pottosin et al., 1997; Schulz-Lessdorf and Hedrich, 1995). To investigate the voltage dependence of SV channel unitary
conductance, we plotted the appropriate curves (Figure 5). Unitary conductance was extrapolated at 0 mV – close to natural tonoplast potential: 0.170.4 mV in spring vegetative, 0.670.4 mV in winter vegetative, 0.470.3 mV in spring generative, 0.070.3 mV in winter generative. It equaled 73.073.9 pS for spring vegetative vacuoles, 74.374.1 pS for winter vegetative vacuoles, 52.27 3 pS for spring generative and 57.371.9 pS for winter generative vacuoles. Single channel conductance in generative leaves was significantly lower than in vegetative leaf vacuoles. There was also a tendency for the specific conductance of SV channels in the vacuoles of winter plants to be higher than in spring plants. The same approach applied to SV channels in small cytoplasm-out patches yielded the following values of single channel conductance (extrapolated at 0 mV): 58.774 pS for spring vegetative vacuoles,
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Figure 2. Ion currents flowing through an isolated macro-patch of the tonoplast of rape (spring vegetative) leaves in the cytoplasm-out configuration. Inset: record in expanded scales of ion currents passing through single channels at 100 mV. Top: voltage protocol – after a prepulse of 60 mV voltage steps from 100 to +100 mV were applied. Pipette and bath solutions as in Figure 1.
Figure 3. Single SV current recordings from the tonoplast patches of rape (winter vegetative) leaves in the vacuole-attached (A) and cytoplasm-out (B) configurations. Numbers on the left side indicate command potentials. Presented traces begin 50 ms after the onset of voltage pulses. Membrane potential in A: 0.3 mV. Pipette and bath solutions as in Figure 1.
48.375.9 pS for winter vegetative vacuoles, 40.37 4.8 pS for spring generative and 5575.4 pS for winter generative vacuolar patches. There were no statistically significant differences among the groups except for the spring generative vacuoles. The lack of difference in SV channel conductance between vegetative and generative vacuoles in well controlled cytoplasm-out patches with respect to the conductance in the vacuole-attached mode can be explained by differences in the natural vacuolar sap composition in intact vacuoles, and the pipette solution in the cytoplasm-out patches. The comparison of current densities measured in the four experimental variants in the wholevacuole configuration revealed no statistically significant differences among the groups. Figure 6 presents current densities (in A/m2) calculated for +100 mV. Here again, the ion composition of the solutions on both sides of the vacuolar membrane was well defined and differed from natural values. Application of a high Ca2+ concentration (2 mM) in pipette and bath solutions reduced currents passing through the SV channels in comparison to low Ca2+ in the vacuole lumen (Pottosin et al., 2004). Under the experimental conditions differing from optimal, when one might expect current saturation, differences in SV activity among the examined variants should be more expressed. However, the current densities remain relatively constant. The effect of externally added phytohormone GA3 (2 mg/dm3 dissolved in the standard solution) was examined on the single channel level and macroscopic currents in cytoplasm-out macropatches of winter vegetative vacuoles. There was
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Figure 4. Current – voltage characteristics of single SV channels from rape vacuoles. Recordings were obtained in the vacuole-attached configuration. Black circles, spring vegetative (n ¼ 9 vacuoles), open circles, winter vegetative (n ¼ 10 vacuoles), black squares, spring generative (n ¼ 31 vacuoles), and open squares, winter generative vacuoles (n ¼ 13 vacuoles). Bars denote standard errors. Pipette and bath solutions as in Figure 1.
Figure 5. Specific conductance/voltage characteristics of SV channels in rape vacuoles. The data presented in Figure 4 were used to plot the graph. Lines represent linear regression curves. Symbols indicating experimental variants are the same as in Figure 4.
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Figure 6. Current densities of time-dependent currents recorded in the whole-vacuole configuration at +100 mV. s–v, spring vegetative (n ¼ 5 vacuoles); w–v, winter vegetative (n ¼ 6 vacuoles); s–g, spring generative (n ¼ 3 vacuoles); w–g, winter generative (n ¼ 3 vacuoles). Error bars denote SE. Vacuole diameter was measured under the microscope. Surface area of the vacuoles was calculated assuming a spherical shape.
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Figure 8. Current – voltage characteristics of isolated cytoplasm-out macropatches of winter vegetative rape leaf vacuoles. Open circles – control (n ¼ 6 vacuoles), black circles – GA3 (2 mg/l) treated vacuoles (n ¼ 8 vacuoles).
rape in well-controlled experimental conditions exhibit similar properties. However, the differences between vegetative and generative leaf vacuoles appear in the vacuole-attached configuration, indicating that the vacuolar sap composition, which changes during floral induction, is an important regulating factor affecting SV channel activity.
References
Figure 7. Current – voltage characteristics of isolated cytoplasm-out patches of winter vegetative rape leaf vacuoles. Open circles – control, black circles – GA3 (2 mg/l)-treated vacuoles.
no significant difference in single channel conductance between the respective controls and GA3treated vacuoles (Figure 7). On the other hand, macroscopic currents at positive potentials were significantly lower (80% of the control) in the GA3treated vacuoles (Figure 8). It is known that SV currents exhibit a time-dependent run down. To avoid this effect, we added GA3 to the vacuole isolation medium, which allowed us to perform measurements on the control and GA3-treated vacuoles on a comparable time scale. These data indicate that GA3 added externally leads to a lower probability of SV channels opening. We conclude that SV channels of generative and vegetative leaves of spring and winter varieties of
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