Author for correspondence: Frans J. M. Maathuis Tel: +44 1904 328652 Email: firstname.lastname@example.org
•Plant two-pore K+ channels (TPKs) have been shown previously to play a role in vacuolar K+ homeostasis. TPK activity is insensitive to membrane voltage, but regulated by cytoplasmic calcium and 14-3-3 proteins. This study reports that membrane stretch and osmotic gradients also alter the activity of TPKs from Arabidopsis, rice and barley, and that this may have a physiological relevance for osmotic homeostasis.
•Mechanosensitivity was studied using patch clamp experiments and TPKs from Arabidopsis, rice and barley. In addition, the capability of TPKs to act as osmosensors was determined. By using protoplast disruption assays and intact plant survival assays, in genotypes that differed in TPK expression, the physiological relevance of TPK-based osmosensing was tested.
•TPKs from all three species showed varying degrees of mechanosensitivity. TPK activity in channels from all three species was sensitive to trans-tonoplast osmotic gradients. TPK osmosensing is likely to proceed via the detection of small perturbations in membrane tension. Intact plant and protoplast assays showed that TPK-based osmosensing is important during exposure to rapid changes in external osmolarity.
•Vacuolar TPK channels can act as intracellular osmosensors and rapidly increase channel activity during hypo-osmotic shock to release vacuolar K+.
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Plants are exposed to many environmental fluctuations. These fluctuations can include periods of water excess (e.g. heavy rainfall or flooding) and water deficiency (e.g. drought or salinization) that can rapidly alternate, and thus require sophisticated mechanisms for turgor adjustment and osmoregulation.
Turgor maintenance is also necessary to endow plants with structure, and thus enables them to carry out important biological processes, such as photosynthesis (Hájek & Beckett, 2008) and expansive growth, whereas dynamic changes in turgor are responsible for organ movement.
Adequate osmoregulation can only proceed if systems are in place that report changes in osmolarity either intracellularly, outside the cell or in both compartments. Such osmosensing can occur via direct and indirect mechanisms: plasma membranes may contain receptor kinases that respond to pressure changes or record mechanical distortion between membrane and cell wall. For example, the plant histidine kinase AtHK1 is believed to form homodimers whose conformation is sensitive to changes in external osmolarity, possibly via the sensing of membrane tension (Tamura et al., 2003; Wohlbach et al., 2008). In bacteria, various members of the mechanosensitive ion channel (Msc) family are essential for the adaptive response to reductions in external osmotic pressure (Wood, 1999). Msc family channels report membrane stretch that results from the hypo-osmotic shock-induced volume change. In addition, specific transporters for the release of osmolytes are activated in response to hypotonic conditions (Wood, 2006). For example, the Escherichia coli proline transporter ProP contains periplasmic loops that respond to reduced osmolarity in this compartment by conformational changes that increase transport activity (Culham et al., 2008). Plants also possess mechanosensitive channels which have been recorded in many plant protoplasts (e.g. Cosgrove & Hedrich, 1991; Spalding & Goldsmith, 1993; Dutta & Robinson, 2004; Zhang et al., 2007). Recently, two members of the mechanosensitive channel of small conductance-like (MSL) family have been shown to express in Arabidopsis root plasma membranes (Haswell et al., 2008) and, although their exact physiological role remains to be revealed, it is hypothesized that they may contribute to turgor regulation (Haswell et al., 2008).
The mechanisms described above are sometimes referred to as ‘indirect’ osmosensors as they do not detect changes in water activity per se, but rather the changes in cell structure that result from hyper- and hypo-osmotic shock. However, proteins may detect changes in osmotic gradients directly, for example via changes in water activity in the surrounding solution or those of kosmotropic (protein conformation-stabilizing) or chaotropic (protein conformation-destabilizing) cosolvents (Wood, 2006). Transmembrane differences in water activity may lead to distortion in the membrane itself, via effects on lipid hydration, van der Waals’ interactions and electrostatic interactions. In each case, these phenomena can provoke conformational changes that lead to protein activation.
In plant cells, turgor exerts itself at the cell wall and changes in turgor pressure will primarily manifest themselves there. The registration of changes in osmotic and turgor pressure may therefore take place at the plasma membrane cell wall boundary, for example via AtHK1. However, in most plant cells, the majority of volume derives from the large lytic vacuole which is the main depository for minerals that provide turgor to plant cells. Changes in turgor and osmolarity inevitably require large trans-tonoplast fluxes of inorganic ions, such as K+ and Cl−, to restore water relations, and vacuoles are therefore essential in the control of osmotic homeostasis.
It is extremely unlikely that plant cell tonoplasts experience significant transmembrane pressure as this would rapidly lead to cell damage. In steady state conditions, even osmotic gradients between cytoplasm and vacuole are likely to be negligible because of the large tonoplast water permeability. However, rapid changes in either external or internal water activity can temporarily create osmotic gradients across the tonoplast that necessitate fast and large fluxes of solutes to dissipate (MacRobbie, 2006). Several studies have shown rapid effects of changes in external osmolarity on vacuolar ion fluxes: In Commelina communis guard cells, hypo-osmotic shock generated a large and rapid transient Rb release from the vacuole (MacRobbie, 2006), whereas, in Arabidopsis thaliana root hair vacuoles, hyperosmotic conditions led to a significant increase in tonoplast conductance (Lew, 2004). There is also evidence that changes in tonoplast tension act as a hysteresis switch for malate loading and unloading in crassulacean acid metabolism (CAM) plants (Luttge, 2000).
Although the intermediate signalling components are unknown in all these examples, it has been suggested that osmosensing at the tonoplast itself is an important aspect of osmotic homeostasis (Alexandre & Lassalles, 1991; Lew, 2004; MacRobbie, 2006). One report describes the presence of mechanosensitive channels at the tonoplast of red beet vacuoles which could fulfil this function (Alexandre & Lassalles, 1991), but both the function and molecular identity of this channel are unknown.
To assess whether vacuolar K+ channels of the two-pore K+ channel (TPK) family can function as osmosensor, the mechanosensitivities of AtTPK1, HvTPK1 and OsTPKa and their activities in response to trans-tonoplast osmotic gradients and to the action of agents that were inserted into the lipid bilayer were studied. It was found that AtTPK1 and its rice and barley homologues show various degrees of mechanosensitive activation, that the activity of these channels responds to trans-tonoplast osmotic gradients and that this property may be important for osmosensing in intact cells.
Materials and Methods
Arabidopsis thaliana (L.) ecotype Columbia (0) wild-type, TPKox and tpk1 mutant seeds (SALK line 146903; Gobert et al., 2007) were surface sterilized and placed on agar plates. Double mutants tpk1:tpc1 were obtained from crosses between homozygous TPK1 loss-of-function mutants (SALK line 146903) and TPC1 loss-of-function mutants (SALK line 145413; Peiter et al., 2005). Growth medium composition was as described previously (Maathuis et al., 1998) and contained 1.25 mM KNO3, 0.5 mM Ca(NO3)2, 0.5 mM MgSO4 and 0.625 mM KH2PO4 as macronutrients. After stratification at 4°C for 2 d, seeds were transferred to a growth cabinet or growth room with the following conditions: 12 h; light intensity, 120 μmol m−2 s−1; day : night temperature, 24 : 20°C; relative humidity, 70–80%.
Transient expression of TPK isoforms
A cDNA for HvTPK1 was obtained from Dr Wieland Fricke (University of Dublin, Ireland; Boscari et al., 2009), whereas a cDNA clone for Nipponbare OsTPKa (Os03g54100) was received from the Rice Genome Resource Center (RGRC, Kome, Japan; Isayenkov et al., 2010) Full-length coding sequences minus stop codon were amplified by PCR using forward and reverse primers containing XhoI and SmaI sites and inserted into the pART7-EYFP vector, as described in Gobert et al. (2007), to produce pART7-TPK vectors encoding C-terminal fusions of TPKs with enhanced yellow fluorescent protein (EYFP) under the control of a cauliflower mosaic virus (CaMV) 35S promoter.
Arabidopsis protoplasts were isolated from true leaves from seedlings that had been grown on plates between 12 and 18 d, and transformed with pART7-TPK-EYFP or pART7-EYFP constructs, as described previously (Gobert et al., 2007). For electrophysiological experimentation, protoplasts derived from tpk1-tpc1 double mutants were used, as these constitute an electrically silent system. For cellular localization of TPK-YFP proteins, epifluorescence was used 24–48 h after transformation.
After transfer of leaf protoplasts to the recording chamber, vacuoles were released by washing protoplasts with a solution containing 10 mM EDTA, 10 mM ethyleneglycol-bis(β-aminoethylether)-N,N′-tetraacetic acid (EGTA), pH 8, with an osmolarity of 350 mOsm. The patch clamp apparatus and methodology were as described in Maathuis et al. (1998). Briefly, the current was recorded using an EPC7 (List, Darmstadt, Germany) amplifier and data acquisition occurred through a CED (Cambridge, UK) A/D converter at 1–3 kHz; data were filtered at 0.5 kHz. CED Patch v5.5 software was used for data analysis. Glass pipettes were pulled from Kimax (Kimble, OH, USA) glass using a List electrode puller. The standard experimental solutions for bath and pipette contained 100 mM KCl, 0.1 mM CaCl2, 5 mM 2-(N-morpholino)ethanesulphonic acid/tris(hydroxymethyl)aminomethane (MES/Tris), pH 6, and sorbitol adjusted to a total osmolarity of 430 mOsm. For low cytoplasmic Ca2+ experiments, solutions contained 100 mM KCl, 1 mM CaCl2, 5 mM MES/Tris, pH 7, and 1.5 mM EGTA. Open probabilities are expressed as Po = (topen/ttotal)/n × 100 where n is the number of channels in the membrane patch (Gobert et al., 2007). Recordings of 60-s duration at a membrane potential of −50 mV were used to determine Po. Open probability data are given as the average ± SD using vacuoles from three to six individual protoplasts. A Boltzman distribution in the form of Po = exp[α(p − p1/2)] was used, where α is the slope, p is the pressure in mmHg and p1/2 is the pressure value where Po is 50%. Negative pressure was applied by mouth and constantly monitored using a manometer.
YFP expression measurements
To assess whether various protoplast preparations showed comparable TPK expression, YFP fluorescence was quantified. Protoplasts were suspended at 2 × 106 ml–1 in the same buffer as used for patch clamp analyses. Protoplasts were placed in a cuvette with stirrer and YFP expression was measured spectrally with excitation at 510 nm and emission at 530 nm using a Horiba Fluoromax-4 spectrofluorometer (Horiba, Stanmore, Middlesex, UK). Fluorescence was measured in three independently transformed batches of protoplasts for soluble YFP, each isoform (AtTPK1, HvTPK1 and OsTPKa) and nontransformed control, 24 h after transformation for TPK fusions and 16 h after transformation with soluble YFP. Data are given as the average ± SD.
For protoplast ion release assays, protoplasts were incubated in 500 mM sorbitol, spun down (2 min, 200 g) and resuspended in 400 mM sorbitol, with the medium conductivity continuously recorded using a Mettler Toledo (Mettler, Giessen, Germany) conductivity meter. Experiments were repeated four times. Protoplast lysis was recorded microscopically by counting the number of intact protoplasts after each step change in medium osmolarity (from 450 to 200 mOsm). Data are given as the average ± SD from five to six independent experiments for each genotype, starting with 100–150 intact protoplasts.
Results and Discussion
Plant TPKs are mechanosensitive
In Arabidopsis, TPK1 has been shown to be involved in general K+ homeostasis, stomatal function and germination (Gobert et al., 2007; Maathuis, 2009). AtTPK1 activity is largely impervious to membrane voltage, but has been shown to be modulated by cytoplasmic Ca2+ and pH (Gobert et al., 2007), and by phosphorylation and 14-3-3 protein binding (Latz et al., 2008). Plant TPKs show structural similarity to mammalian two-pore domain K+ channels of the K2P family (Buckingham et al., 2005). The first cloned member of K2p channels was TWIK (‘tandem P domain, weak inward rectifier K+’), and several subsequently identified members show mechanosensitivity, such as TREK (‘TWIK related K+’) and TRAAK (‘opened by arachidonic acid K+’) channels. The observation that several members of the K2P family are mechanosensitive (Buckingham et al., 2005) motivated us to test whether plant TPKs also respond to mechanical stimuli. Fig. 1(a) shows AtTPK1 activity in an excised patch from a mesophyll vacuolar patch at a holding potential of −50 mV. Increased negative pressure (suction) to the membrane increases channel activity. A similar phenomenon was observed after homologous expression of AtTPK1 in mesophyll protoplasts obtained from Arabidopsis tpk1:tpc1 loss-of-function plants (Fig. 1b). This genotype provides an ideal, electrically silent background, and no mechanosensitive activity was recorded in nontransformed protoplasts (Supporting Information Fig. S1). In the absence of membrane stretch, AtTPK1 activity is readily observed, but this increases in the presence of −40 or −60 mmHg suction. From long-duration current recordings (60 s for each condition), a modest but significant (P <0.05) increase in open probability becomes obvious (Fig. 1c), implying that AtTPK1 is mechanosensitive.
To test whether mechanosensitivity is a generic property of vacuolar TPK channels, cDNA clones for two further TPK isoforms from rice (OsTPKa; Os03g54100; Isayenkov et al., 2010) and barley (HvTPK1; Boscari et al., 2009; Sinnige et al., 2005) were obtained. After translational fusions with YFP, the expression of each TPK was found to be in the main lytic vacuole (Fig. 2a; Isayenkov et al., 2010). When heterologously expressed in tpk1:tpc1 mesophyll protoplasts, single channel currents were obtained for each TPK with very similar characteristics to the previously reported observations for AtTPK1 (Bihler et al., 2005; Gobert et al., 2007) and NtTPK1 (Hamamoto et al., 2008); as shown for AtTPK1 and NtTPK1, OsTPKa and HvTPK1 are highly selective for K+ (data not shown), their activity is voltage independent and channels show intrinsic inward rectification in all cases (Fig. 2b–d). Interestingly, in both rice and barley, TPK mechanosensitivity was obvious, but mostly so in barley TPK1; Fig. 3(a) shows the considerable activation of HvTPK1 even at moderate membrane stretch. As the activity of many TPKs increases when cytoplasmic Ca2+ is increased (e.g. Gobert et al., 2007), mechanosensitivity was also tested for AtTPK1 and HvTPK1 when cytoplasmic Ca2+ concentrations were reduced from 100 to 0.2 μM. In both cases, mechanosensitivity could be observed (Fig. S2), but whether and how cytoplasmic Ca2+ modulates mechanosensitivity remain to be tested.
In Fig. 3(b), typical examples of the relationship between membrane stretch and channel activation are given for all three TPK channels. Using a Boltzmann equation to fit the data allows a quantitative comparison to be made between the isoforms, showing that HvTPK1 has a half-activation value of −23 ± 6.4 mmHg (n = 4), whereas that for OsTPKa is −45 ± 8.1 mmHg (n =3). By contrast, AtTPK1 current saturation was always precluded by the loss of the membrane patch and data could not be fitted to a Boltzmann equation; however, it is apparent from Fig. 3 that this isoform required much greater forces to increase channel activation. Mechanosensitivity was also observed when cytoplasmic Ca2+ concentrations were reduced from 100 to 0.2 μM (Fig. S2).
It appears that all vacuolar TPKs may exhibit a varying degree of mechanosensitivity. How forces in the membrane translate into channel opening remains to be studied. In K2p TREK channels, it has been suggested that a cluster of positive residues in the C-terminus promotes C-terminus membrane interaction, which, in turn, greatly stimulates channel open probability (Chemin et al., 2005). The protein membrane–lipid interaction itself is stimulated by signalling moieties, such as phosphatidylinositol 4,5-bisphosphate (PIP2). No obvious cluster of positive residues is apparent in the C-terminus of plant TPK channels, and only a detailed structural study will reveal whether models that apply to K2p channels are pertinent for plant TPKs.
Plant TPKs respond to osmotic gradients
Changes in turgor could exert considerable force on the membrane and, as such, create tension forces. However, this is unlikely to occur across the tonoplast in vivo. By contrast, (transient) large perturbations in osmotic equilibrium between cytoplasm and vacuolar lumen may occur, especially in response to sudden hypo-osmotic shock (MacRobbie, 2006). It was therefore investigated whether TPK activity was also capable of responding to osmotic gradients by applying changes to the bath osmolarity with 50 or 100 mOsm in either direction relative to the pipette solution. Fig. 4 shows example traces of HvTPK1 activity with symmetrical osmolarity and with the luminal osmolarity reduced or increased by 100 mOsm. A clear increase in channel activity is apparent, particularly when the luminal osmolarity is increased. The cumulative open time of individual patches increased by up to three-fold when a gradient of 100 mOsm was applied (Fig. 4b). A similar but less pronounced activation was observed for OsTPKa in the presence of an osmotic gradient of 100 mOsm (data not shown).
In all, the data show that vacuolar TPK channels respond to trans-tonoplast osmotic gradients, but more so when the luminal osmolarity is increased with respect to the cytoplasmic compartment, i.e. when hypo-osmotic conditions apply. This is in contrast with the yeast vacuolar cation channel YVC1 which is primarily activated by hyperosmotic shock. This channel may contribute to the vacuole-derived Ca2+ signal during hypertonic osmotic stress (Zhou et al., 2003).
Osmosensing may proceed via intrinsic tonoplast properties
The precise mechanism by which living organisms detect osmotic gradients is not clear. Several hypotheses have been proposed which include changes in membrane proteins and/or membrane lipids, for example as a result of altered hydration or membrane fluidity, which, in turn, translates into (local) membrane deformation (Wood, 1999). To assess whether the osmosensing capacity of TPKs is based on such subtle changes in membrane curvature, picric acid and chlorpromazine, compounds that increase or decrease respectively the membrane curvature by inserting asymmetrically in the membrane, were applied to excised patches with HvTPK1 activity. The ‘crenator’ picric acid (trinitrophenol) mimics cell swelling, whereas the ‘cup former’ chlorpromazine does the opposite (Wolfs et al., 2003). Fig. 5 shows examples of recordings when either compound was added to the bath solution. Picric acid (Fig. 5a) caused an increase in channel activity, as is evident from the extra open levels that appear. By contrast, chlorpromazine reduced channel activity (Fig. 5b). The changes in TPK activity are quantified for three independent experiments in Fig. 5(c).
Crenators and cup formers can have many nonspecific effects, particularly on protein–lipid interactions. However, the stimulating effect of picric acid, but not chlorpromazine, suggests that conditions in which cell swelling occurs (hypotonic shock) are those that activate TPKs. In all, such a mechanism could serve as a ‘safety valve’ for the rapid and sustained release of K+ that would be required in these conditions. For example, flooding after a period of drought could constitute a large hypo-osmotic shock for peripheral root cells, as would heavy precipitation in a semiarid saline environment.
Does TPK-mediated osmosensing have physiological relevance?
In Arabidopsis, loss of function in TPK1 led to weak phenotypes, the most pronounced being a delay in stomatal closure in response to abscisic acid (Gobert et al., 2007). No phenotypes associated with drought and/or osmotic stress were reported in long-term growth experiments. However, the role of osmosensing mechanisms is likely to be more relevant during rapid changes in osmotic gradients. Therefore, rapid hypotonic shocks were applied to protoplasts to determine the resistance to osmotic shock at the cellular level. Fig. 6(a) shows the increase in medium conductance after 30, 60 or 120 s in response to a step change in medium osmolarity from 500 to 400 mOsm. It is evident that protoplasts derived from control plants (Attpk1 loss-of-function mutants, transiently transformed with YFP; Gobert et al., 2007) released considerably fewer ions relative to those that were transiently transformed with AtTPK1-YFP. Cells expressing HvTPK1-YFP released ions even more rapidly than those transformed with AtTPK1. To test whether these properties affected cell survival during osmotic stress, protoplast lysis after hypotonic shock was recorded. Protoplasts were exposed to increasing hypo-osmotic shock to determine LD50 values, that is, the osmolarity at which 50% of protoplasts had lysed. Fig. 6(b) shows that protoplasts expressing AtTPK1 were more tolerant to withstand hypo-osmotic shock than were control cells. An even higher degree of tolerance was seen when HvTPK1 was transiently expressed instead of AtTPK1. The YFP signals from AtTPK1 and HvTPK1 were always comparable (Fig. S3), pointing to similar expression levels. In addition, patch clamp experiments showed very similar K+ conduction properties for AtTPK1 and HvTPK1 (Fig. 2), but a greater mechanosensitivity for HvTPK1 (Fig. 3). The latter indicates that the improved osmotolerance is not, or not only, dependent on the K+ transport function of the respective TPKs, as these are virtually identical between AtTPK1 and HvTPK1. Indeed, the larger mechanosensitivity of HvTPK1 may render it more suitable as an osmosensor and, as such, endow protoplasts with more osmotolerance.
Many of the environmental stresses that plants can suffer include osmotic components. Examples include drought stress, temperature stress and salinity. Many genes and gene products have been identified that may be involved in the later stages of the reaction(s) to these stresses, but what happens in the initial, sensing, phase is not clear. This work shows that the vacuolar TPK channels are not only mechanosensitive, but also osmosensitive, and therefore could function as cellular osmosensors during rapid changes in external osmotic pressure (e.g. MacRobbie, 2006). For example, flooding with fresh water of a salt marsh or heavy precipitation after a drought can create hypotonic conditions that require the rapid release of cellular salts. The initial solute release across the plasma membrane can create temporary trans-tonoplast osmotic gradients sensed by TPKs. The activation of osmosensitive TPKs would ensure the rapid release of K+ from the vacuole, the main cellular depository of water and osmotica.
Part of the described research was supported by Biotechnology and Biological Sciences Research Council (BBSRC) funding to F.J.M.M. I thank Dr Wieland Fricke (University of Dublin, Ireland) for his kind gift of the HvTPK1 cDNA.