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Keywords:

  • AtTPK/KCO family;
  • BiFC;
  • expression pattern;
  • tonoplast;
  • FRET

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The Arabidopsis thaliana K+ channel family of AtTPK/KCO proteins consists of six members including a ‘single-pore’ (Kir-type) and five ‘tandem-pore’ channels. AtTPK4 is currently the only ion channel of this family for which a function has been demonstrated in planta. The protein is located at the plasma membrane forming a voltage-independent K+ channel that is blocked by extracellular calcium ions. In contrast, AtTPK1 is a tonoplast-localized protein, that establishes a K+-selective, voltage-independent ion channel activated by cytosolic calcium when expressed in a heterologous system, i.e. yeast. Here, we provide evidence that other AtTPK/KCO channel subunits, i.e. AtTPK2, AtTPK3, AtTPK5 and AtKCO3, are also targeted to the vacuolar membrane, opening the possibility that they interact at the target membrane to form heteromeric ion channels. However, when testing the cellular expression patterns of AtTPK/KCO genes we observed distinct expression domains that overlap in only a few tissues of the Arabidopsis plant, making it unlikely that different channel subunits interact to form heteromeric channels. This conclusion was substantiated by in planta expression of combinations of selected tonoplast AtTPK/KCO proteins. Fluorescence resonance energy transfer assays indicate that protein interaction occurs between identical channel subunits (most efficiently between AtTPK1 or AtKCO3) but not between different channel subunits. The finding could be confirmed by bimolecular fluorescence complementation assays. We conclude that tonoplast-located AtTPK/KCO subunits form homomeric ion channels in vivo.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

In Arabidopsis thaliana, the AtTPK/KCO family combines five tandem-pore K+ channels (TPKs) and one Kir-type K+ channel (KCO3), as deduced from their amino acid sequences (Becker et al., 2004; Mäser et al., 2001; for the nomenclature of AtTPK/KCO channels; see also Experimental procedures). Additionally, 14-3-3-binding motifs and putative Ca2+-binding EF-hands are present in the N- and C-termini, respectively, of five AtTPK/KCO proteins. AtTPK4, however, does not contain these additional regulatory and protein interaction motifs.

AtTPK/KCO genes are expressed at a low level in different plant tissues and cell types. Real-time RT-PCR studies have demonstrated the presence of AtTPK/KCO transcripts in roots, leaves, mesophyll cells, guard cells, flower stalks, hypocotyls and shoots (Deeken et al., 2003; Philippar et al., 2004; Schönknecht et al., 2002) and indicated pollen-specific expression of AtTPK4 (Becker et al., 2004). Cellular distribution of promoter activity in different cell types of Arabidopsis including the root cortex, vascular tissue, mesophyll cells, guard cells and pollen grains was shown for AtTPK1 employing promoter-reporter gene (GUS) fusions (Czempinski et al., 2002).

So far, only a few studies testing the functional characteristics of AtTPK/KCO proteins have been reported. AtTPK1 was shown to be a vacuolar-membrane protein (Czempinski et al., 2002; Schönknecht et al., 2002), that, after heterologous expression in yeast (Saccharomyces cerevisiae) cells, forms a K+-selective, voltage-independent channel activated by cytosolic calcium (Bihler et al., 2005). AtTPK4 was analysed after expression in heterologous systems (Xenopus oocytes and yeasts), and its function in plant cells was studied using Arabidopsis tpk4 knock-out lines, revealing that it is targeted to the plasma membrane where it establishes a voltage-independent K+ channel. Hyperpolarizing as well as depolarizing membrane voltages elicited instantaneous K+ currents, which were blocked by extracellular calcium and cytoplasmic protons (Becker et al., 2004). Thus, K+ channels of the AtTPK/KCO family fulfil their function in different cellular compartments.

Electrophysiological studies have identified several cation and K+ currents over the vacuolar membrane (reviewed in Martinoia et al., 2000). The main K+ currents are carried by the slow activating vacuolar (SV) channel, the fast vacuolar (FV) channel and the vacuolar K (VK) channel. However, the genes coding for the respective channel proteins have mostly not been identified. An exception is the TPC1 gene, which has recently been shown to code for the SV channel protein of plant vacuoles (Peiter et al., 2005). Also, proteomic approaches focusing on the plant tonoplast have not resulted in the identification of many vacuolar channel proteins to date (Carter et al., 2004; Sazuka et al., 2004; Shimaoka et al., 2004; Szponarski et al., 2004); most likely because of the low expression level of channel proteins they were not identified in protein mixtures.

K+ channels form multimeric protein complexes. Active channels can be composed of α subunits alone, or of α subunits and regulatory β subunits, where four so-called P-domains are involved in the formation of the K+-selective pore (Long et al., 2005; MacKinnon, 1991). K+ channel multimerization can occur via interaction of C-termini, as was demonstrated for the plant Shaker channels (Daram et al., 1997; Dreyer et al., 1997, 2004; Ehrhardt et al., 1997), or via disulphide bonds between two α subunits, as was shown for the animal tandem-pore K+ channel (TPK) TWIK-1 (Lesage and Lazdunski, 2000; Lesage et al., 1996). Non-invasive methods based on fluorescence resonance energy transfer (FRET) have become available to observe channel multimer compositions in living cells, and have also been used in plant cells (Immink et al., 2002). Fluorescence resonance energy transfer occurs between a donor fluorescent molecule and a neighbouring fluorophore, the acceptor, when both fluorescent molecules are in close proximity (1–10 nm) (Gadella et al., 1999). Physical protein–protein interactions between fusion proteins with donor and acceptor fluorescent dyes provide the conditions to determine FRET. Multimerization of channel subunits can thus be detected in native cellular environments like the target membrane. Fluorescence resonance energy transfer analysis has been used in plant cells to study cytosolic and nuclear proteins (Immink et al., 2002) and membrane-bound proteins (Bhat et al., 2005; Kluge et al., 2004; Russinova et al., 2004; Seidel et al., 2004) with fluorescence lifetime imaging microscopy, fluorescence spectral imaging microscopy or acceptor photobleaching (APB). Interactions of proteins in plant cells can also be investigated by bimolecular fluorescence complementation (BiFC) detecting yellow fluorescence protein (YFP) fluorescence after the restoration of fluorophore activity by formation of protein complexes (Bracha-Drori et al., 2004; Walter et al., 2004). The BiFC assay is applicable to the analysis of a wide range of protein interactions provided that the two YFP fragments are flexible and close enough to associate with each other when tethered to the protein complex, otherwise false negatives can be obtained with this method. The restoration of the fluorophore from the split fluorescent proteins is slow (t1/2 = 50 min) and essentially irreversible (Hu et al., 2002). With these characteristics the BiFC assay is sufficiently sensitive to enable detection of transient and weak complexes. However, the same characteristics may in some cases limit the applicability of BiFC because it reflects the association between the interaction partners at the time of formation of the complex and does not reveal subsequent shifts in the equilibrium among alternative interaction partners or the dynamics of protein complexes. The BiFC technique has also been used for the visualization of multiple protein interactions within the same cell by using a large number of different green fluorescence protein (GFP) variants (multicolour BiFC assay; Hu and Kerppola, 2003).

The subunit composition of the AtTPK/KCO channels was not examined in previous studies. The present work extends the use of FRET and BiFC analysis to the investigation of vacuolar K+ channel composition. We report on further analysis of the expression pattern of members of the AtTPK/KCO gene family, the subcellular localization of the encoded proteins and the identification of AtTPK/KCO homomers of vacuolar members of the protein family in plant cells. Our data demonstrate that AtTPK/KCO proteins form homomeric vacuolar ion channels in plant cells.

Results and Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Transcript abundance of AtTPK/KCO genes in Arabidopsis organs

Generally, the level of expression of AtTPK/KCO genes is low and can often not be reliably tested using standard Northern blot experiments. Therefore, gene expression patterns of AtTPK/KCO genes were previously analysed using real-time RT-PCR (Deeken et al., 2003; Philippar et al., 2004; Schönknecht et al., 2002). In these experiments, whole seedlings or organs of mature plants such as flowers, siliques, stems, rosette leaves and roots were used. Within the gene family, AtTPK1 exhibits the highest transcript level in all tissues analysed, followed by AtTPK3 and AtTPK5. Very low transcript abundance was observed for genes AtTPK2, AtTPK4 and AtKCO3. Overall, AtTPK/KCO expression showed relatively poor organ specificity. However, both real-time RT-PCR and promoter–reporter gene studies (see below) revealed overlapping expression patterns for some AtTPK/KCO genes, i.e. expression of AtTPK1 and AtTPK3 in root tips, of AtTPK1, AtTPK5 and AtKCO3 in vascular tissues and of AtTPK1, AtTPK3 and AtTPK4 in pollen. We previously reported strong expression of AtTPK4, encoding a plasma membrane two-pore K+ channel, in pollen (Becker et al., 2004).

Expression patterns of AtTPK/KCO genes can also be inferred from microarray (Affymetrix GeneChip) hybridization data available through the Genevestigator interface (http://www.genevestigator.eth2.ch; Zimmermann et al., 2004). Microarray hybridization data indicated that the transcript abundance of AtTPK/KCO genes was low in Arabidopsis organs, mirroring the expression data obtained by real-time RT-PCR. However, no information was available for AtTPK4 because this gene is not represented by the ATH1 gene chip.

A detailed interpretation of AtTPK/KCO expression data employing the Gene Atlas, Gene Chronologer and Response Viewer tools of the Genevestigator package revealed the presence of AtTPK/KCO transcripts throughout all plant developmental stages and in different Arabidopsis organs like roots, stems, leaves and flowers. As detected in real-time RT-PCR, AtTPK1 exhibits the highest transcript level within the gene family in these tissues. Furthermore, enhanced levels of AtTPK3 transcript were recorded in petals, stamen, seeds and senescent leaves; AtTPK5 mRNA shows higher abundance in petals and senescent leaves. In these tissues, AtTPK3 and AtTPK5 transcript abundance mostly exceeded that of the AtTPK1 gene. AtTPK2 transcripts were detected at an elevated level in RNA from stamen (eight chips analysed) and pollen (two chips analysed). Levels of AtTPK/KCO transcript were not strongly affected by a number of applied biotic or abiotic stresses; however, the level of AtTPK5 transcript increased with abscisic acid treatment (about twofold) and with nitrogen starvation (about 2.5-fold), and declined during cold stress (about twofold). Expression of AtTPK3 was induced by methyl jasmonate treatment (about 2.4-fold) in microarray experiments.

Promoter–reporter gene studies

To test the transcriptional activities of AtTPK/KCO promoters we isolated 5′ upstream regulatory regions (1–2.5 kb in length) and fused them to the β-glucuronidase reporter gene. After transformation of A. thaliana with the respective gene constructs, the distribution of GUS staining was analysed in transgenic plants. Previously we have shown in promoter–GUS studies that AtTPK1 is active in different tissues and cell types such as the mesophyll cells and guard cells of leaves, the vascular tissue of leaves and roots, anthers and pollen grains, embryos, and roots and hypocotyls of small seedlings (Czempinski et al., 2002).

Transgenic plants harbouring the fusion construct AtTPK3–GUS exhibited GUS activity in the hypocotyl and root tips of young seedlings (Figure 1a), in first developing rosette leaves (Figure 1b,c), and in anthers (pollen grains; Figure 1d). AtTPK4–GUS expression in transgenic plants confers GUS activity mainly in pollen grains (Becker et al., 2004). Detailed analysis of AtTPK4 promoter activity in transgenic lines revealed the absence of GUS activity during very early stages of flower development (Figure 1e). However, GUS staining was detected at a stage of flower development when buds were still closed (Figure 1f). β-Glucuronidase activity was present in the tapetum (Figure 1g), and later on in fully developed pollen grains (Figure 1h,i). Both AtTPK5 and AtKCO3 promoters drove GUS expression in the vascular tissues of leaves, roots, flower tissues and stems (Figure 1j–s) as well as in hydathodes (Figure 1k,p). In both cases GUS activity in stems was higher in apical regions than in basal regions. Cross sections of stained tissues indicated GUS staining in phloem cells, in parenchyma cells of the interfascicular tissue and in developing xylem cells of stems (Figure 1n,s), leaves, flowers and roots (data not shown). In a few cases GUS activity was detectable in pericycle and endodermis cells of roots of AtKCO3–GUS transgenic lines (data not shown).

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Figure 1. AtTPK/KCO–GUS expression analysis in different Arabidopsis tissues. AtTPK3–GUS expression in 3-day-old plantlets (a), in 20-day-old plantlets (b) and in 5-week-old plants (c,d) raised on soil. AtTPK4–GUS staining of flowers at different stages of floral development: (e) stages 1–8 with immature pollen; (f,g) stages 9–12; (h,i) stages 13–14 with mature pollen. A cross-section through anthers is shown in (i). AtTPK5–GUS expression in 3-day-old plantlets (j) and 5-week-old plants: (k) hydathode; (l) lateral root; (m) flower; (n) stem cross-section. AtKCO3–GUS activity in 3-day-old plantlets (o) and 5-week-old plants: (p) hydathode; (q) root; (r) flower; (s) stem cross-section. X, xylem; P, phloem.

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It is notable that the AtKCO3 and AtTPK5 genes, as demonstrated here employing promoter–reporter gene constructs, have an almost identical expression pattern. Slight effects in response to external factors were only observed in the case of AtTPK5 expression (see above). Transgenic plants harbouring an AtTPK2 promoter–GUS fusion construct were generated and tested (T1 generation). No GUS activity could be detected in plant tissues after plants were grown under normal growth conditions.

AtTPK/KCO proteins locate to the tonoplast in plant cells

To facilitate the investigation of the subcellular localization of AtTPKs and AtKCO3, we prepared transgenic plant material expressing translational AtTPK/KCO–GFP, –CFP (cyan fluorescence protein) or –YFP fusion proteins under the control of the cauliflower mosaic virus (CaMV) 35S promoter. Arabidopsis and tobacco (BY2) protoplasts expressing the GFP, CFP or YFP fusion proteins were screened via direct imaging of fluorescence in living cells.

Images obtained by confocal laser scanning fluorescence microscopy were evaluated from several hundred Arabidopsis protoplasts exhibiting both high and low levels of expression of fusion proteins. The images shown here represent the typical distribution of fluorescence in transformed protoplasts. No detectable signal was present in non-transgenic protoplasts (data not shown). The confocal images shown in Figure 2 indicate that signals from AtTPK2, AtTPK3, AtTPK5 and AtKCO3 fusion proteins originated from vacuolar membranes of both the central vacuole and smaller vacuoles present throughout the cells. In some cases fusion proteins also labelled smaller spherical structures, so-called ‘bulbs’, that are generally regarded as invaginations of the tonoplast (Reisen et al., 2005; Saito et al., 2002; Uemura et al., 2002). Bulbs were labelled most frequently in protoplasts expressing AtKCO3 fusion proteins. Fluorescence of fusion proteins was in all cases excluded from the plasma membrane. In order to release vacuoles from protoplasts we performed osmotic lysis for protoplasts expressing AtTPK5 and AtKCO3 fusion proteins. Green fluorescent protein fluorescence was visible from isolated vacuoles (Figure 2e,f). As control proteins for tonoplast targeting (Figure 2g,h) we used AtNRAMP3-GFP (Thomine et al., 2003) and AtTPK1-GFP (Czempinski et al., 2002). No change was observed in the subcellular localization when different vacuolar AtTPK/KCO fusion proteins were co-expressed in Arabidopsis protoplasts, indicating faithful targeting in the presence of other channel proteins belonging to the same family. AtTPK1 and AtKCO3, AtTPK5 and AtKCO3 (Figure 3) and AtTPK1 and AtTPK5 (not shown) labelled the same vacuoles (central and smaller vacuoles). However, accumulation of AtTPK3 fusion proteins in additional internal membranes was visible in several experiments (Figure 2b).

image

Figure 2.  Subcellular localization of AtTPK/KCO fusion proteins. (a) AtTPK2–green fluorescence protein (GFP): GFP and bright-field image. (b) AtTPK3–GFP: GFP and bright-field image. (c) AtTPK5–cyan fluorescence protein (CFP): CFP and bright-field image. (d) AtKCO3–YFP: YFP and bright-field image. (e) AtTPK5–GFP: released vacuole after osmotic lysis of transformed protoplast, GFP and bright-field image. (f) AtKCO3–GFP: released vacuole after osmotic lysis of transformed protoplast, GFP and bright-field image. All four fusion proteins localize to the tonoplast after transient expression in Arabidopsis cell culture protoplasts. Controls: (g) AtNRAMP3–GFP, GFP and bright-field image, and (h) AtTPK1-GFP, GFP and bright-field image.

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Figure 3.  Co-expression of different AtTPK/KCO fusion proteins. (a) AtTPK1–yellow fluorescence protein (YFP) and AtKCO3–cyan fluorescence protein (CFP). (b) AtTPK5–YFP and AtKCO3–CFP. In (a) and (b) fusion proteins co-localize in the vacuolar membrane after transient expression in Arabidopsis cell culture protoplasts (left, YFP fluorescence; middle, CFP fluorescence; right, bright-field).

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In contrast to the proteins studied here, AtTPK4 has previously been shown to be targeted to the plasma membrane (Becker et al., 2004). Obvious structural differences between AtTPK4 and other AtTPK/KCO subunits are the presence of 14-3-3-binding motifs in N-termini, and Ca2+-binding EF-hand motifs in C-termini of the vacuolar proteins. Recently, it has been shown for animal two-pore channels that the interaction with 14-3-3 protein enables forward trafficking of the channel subunits to target membranes (summarized in Plant et al., 2005). In plants, 14-3-3 protein interacts with HvKCO1 (HvTPK1) from barley, as demonstrated by surface plasmon resonance (Sinnige et al., 2005), and with AtTPK1, as shown by two-hybrid assays (our unpublished data). Whether 14-3-3 protein interaction is also necessary for trafficking and targeting of AtTPK/KCO channel proteins is unclear at present. In the case of alpha tonoplast intrinsic protein (αTIP) the C-terminus of the protein is known to be important for targeting and maintaining enhanced stability of the protein (Höfte and Chrispeels, 1992). Recent studies elucidated that αTIP utilizes a Golgi-independent pathway to protein storage vacuoles (PSVs; Park et al., 2004) and identified (At)SRC2 as a protein interacting with a targeting motif in the cytoplasmic C-terminus of αTIP specific for this endoplasmic reticulum (ER)-to-vacuole pathway (Oufattole et al., 2005). However, no general targeting sequence for tonoplast proteins is known so far (Vitale and Raikhel, 1999). Because members of the AtTPK/KCO family serve functions in either the vacuolar or plasma membrane they might be ideal candidates for studying tonoplast targeting processes in detail.

Channel multimer composition of tonoplast AtTPK/KCO proteins

Results from promoter–reporter studies indicate an almost identical expression pattern for AtTPK5 and AtKCO3; variation for AtTPK5 is possible in response to external factors (see microarray hybridization data above). AtTPK1 shows the highest transcript level within the gene family in several plant tissues and throughout all plant developmental stages, potentially allowing protein association with other members of the protein family. In addition AtTPK1, AtTPK5 and AtKCO3 were clearly detected in the tonoplast. We therefore selected AtTPK1, AtTPK5 and AtKCO3 for analyses of channel subunit associations.

To investigate whether subunits of vacuolar AtTPK/KCO channels form homo- or heteromeric complexes we studied protein–protein interactions in transgenic cells using in vivo FRET and BiFC. C-terminal YFP and CFP fusions of each channel subunit were co-expressed in onion epidermal cells, which provide a natural spatial fixation on slide discs during microscopic observation that is essential for FRET analysis using acceptor bleaching. As in Arabidopsis protoplasts, fusion proteins of AtTPK1, AtTPK5 and AtKCO3 target to the tonoplast and label the large central vacuole and smaller vacuoles in the cells. Fluorescence resonance energy transfer APB was applied to record FRET as it prevents problems associated with variable expression levels. For each pair of constructs tested, data were recorded from at least 16 sample sites in at least 10 transformed cells (for AtTPK5/AtKCO3); usually 30–45 sample sites in 20–30 transformed cells were analysed. Fluorescence resonance energy transfer APB efficiency was calculated after acceptor bleaching of either the whole cell area or of distinct parts of the vacuolar membrane (regions of interest, ROIs). Figure 4(a) shows a confocal image of a cell co-expressing AtTPK1–CFP and AtTPK1–YFP with indicated areas of analysis (i.e. ROI). Mean FRET APB efficiencies of 18.2 ± 5.25% were obtained for energy transfer between AtTPK1 subunits (average ± SD; Figure 4b). Similar FRET signals were obtained for AtKCO3 homomeric subunit interactions (18.2 ± 7.91%). Energy transfer between AtTPK5 subunits was less prominent (10.6 ± 4.03%), indicating a less favourable orientation of the CFP/YFP fluorophores for energy transfer, presumably caused by a larger distance of the subunit molecules or C-termini in a possible AtTPK5 dimer. Importantly, energy transfer between heteromeric combinations was clearly reduced to background level (AtTPK1/AtKCO3, 3.58 ± 3.62%; AtTPK1/AtTPK5, 2.73 ± 3.03%; AtTPK5/AtKCO3, 1.5 ± 1.76%). Mean FRET APB efficiency was calculated for channel subunit combinations with swapped donor and acceptor fluorophores. Similar low-level FRET signals were recorded in control experiments with AtTPK1–CFP/freeYFP (2.61 ± 3.17%) and free YFP/CFP (2.08 ± 2.91%; Figure 4b). The background FRET signal in cells expressing YFP and CFP is probably due to dimer formation of the two fluorophores at very high protein concentrations (Zacharias et al., 2002). This tendency of all forms of GFP to dimerize could also influence the dimerization of investigated membrane proteins (Zacharias, 2003; Zacharias et al., 2002). However, it was shown that a mutation in YFP (A206K) virtually eliminated the homoaffinity of the fluorescence protein and the mutated protein was successfully used in FRET experiments in living cells (Zacharias et al., 2002). To exclude false-positive FRET signals we studied the subunit interaction for AtTPK1 fused to mutated YFP(A206K) and CFP(A206K), respectively. We performed site-directed mutagenesis in AtTPK1 constructs to include ‘non-sticky’ YFP [AtTPK1–YFP(A206K)] and CFP [AtTPK1–CFP(A206K)] in our studies. As shown in Figure 4(b) mean APB efficiencies of 14.62 ± 6.44% were obtained for energy transfer between AtTPK1 subunits fused to ‘non-sticky’ YFP and CFP. Control experiments with AtTPK1–CFP(A206K)/freeYFP resulted in low energy transfer levels (1.98 ± 2.71). This result indicates that FRET occurs due to protein interaction between AtTPK1 subunits and does not result from interactions between fluorescence proteins with high homoaffinity.

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Figure 4.  Fluorescence resonance energy transfer (FRET) analysis of selected combinations of vacuolar AtTPK/KCO proteins. (a) Acceptor photobleaching analysis on a cell co-expressing AtTPK1–cyan fluorescence protein (left) and AtTPK1–yellow fluorescence protein (right). ROI1 indicates the bleached region. ROI2–5 were used for FRET APB efficiency measurements. (b) A FRET APB analysis of the interaction between fluorescently tagged AtTPK/KCO proteins. Mean FRET APB efficiencies ± SD of 30–45 sample sites are shown.

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Taking into account that typical FRET APB efficiencies for protein interaction pairs range from 10–30% (Bhat et al., 2005) clear subunit interaction could be detected only when homomeric combinations of the AtTPK/KCO channel proteins were co-expressed. Co-expression of heteromeric subunit combinations strongly decreased FRET APB efficiency.

To prove the dimerization of vacuolar AtTPK/KCO subunits we additionally performed BiFC assays. The fluorescent complex which is formed by two non-fluorescent fragments of the YFP is restored through the association of interacting proteins, e.g. AtTPK/KCO subunits, fused to these fragments. As a positive control for our studies we used constructs enabling the interaction of bZIP transcription factors in the nucleus (Walter et al., 2004; data not shown). We transiently transformed protoplasts from tobacco and Arabidopsis suspension cells with homomeric and heteromeric AtTPK/KCO subunit combinations. Yellow fluorescence protein signals could be obtained when AtTPK1, AtTPK5 and AtKCO3 subunits were co-transformed as homomers (Figure 5). No fluorescence or only background fluorescence could be observed in control protoplasts which were transformed with single plasmids or with any combination of empty vectors (data not shown).

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Figure 5.  A Bimolecular fluorescence complementation analysis shows homomerization of AtTPK1, AtTPK5 and AtKCO3 channel subunits. Yellow fluorescence protein (YFP) fluorescence and bright-field images of Arabidopsis cell culture protoplasts (a,b) and tobacco BY2 protoplasts (c) co-transformed with (a) AtTPK1–YFPN-terminus and AtTPK1–YFPC-terminus, (b) AtTPK5–YFPN-terminus and AtTPK5–YFPC-terminus, and (c) AtKCO3–YFPN-terminus and AtKCO3–YFPC-terminus.

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Homodimerization-induced YFP fluorescence appeared in the vacuolar membrane confirming the observation that AtTPK1, AtTPK5 and AtKCO3 homomers locate to the tonoplast.

Expression of heteromeric combinations of the selected proteins resulted in diffuse YFP fluorescence in non-target compartments (ER, cytosol) of protoplasts (data not shown). The intensity of YFP fluorescence varied strongly, ranging from a level close to background to a level seen in protoplasts expressing AtTPK/KCO homomers. We never observed YFP fluorescence clearly associated with the tonoplast even after 48 h of incubation, indicating that no interaction of heteromers occurs at the target membrane. It has been reported that YFP fragments tend to establish non-specific interactions at high levels of protein expression (Walter et al., 2004). It has been estimated that fluorescence complementation can occur when YFP fragments are separated by an average distance of more than 10 nm as long as there is sufficient flexibility to allow association of the fragments (Hu et al., 2002). Fluorescence detected in heteromeric AtTPK/KCO combinations could therefore result from an initial association and subsequent permanent coupling via the restored YFP and partial degradation of such incorrect multimers by the cellular quality control system.

Although identification of positive homomer interaction in BiFC analyses is clearly detectable, no quantitative information about complex formation can be obtained. As we also used protoplasts from Arabidopsis suspension cells (in addition to tobacco BY2 cells), unmodified interaction partners (AtTPK/KCO channels) can compete with bimolecular interaction complexes.

Both FRET and BiFC analyses strongly support the existence of AtTPK/KCO channels as homomers, theoretically as dimers (AtTPK1 and AtTPK5) or as tetramers (AtKCO3), in the tonoplast. In our experimental conditions only homomeric combinations allowed the detection of FRET between the channel subunits or restoration of YFP fluorescence at the tonoplast (in BiFC experiments). Similar FRET APB efficiencies were detected for AtTPK1 and AtKCO3 homomeric combinations, whereas the energy transfer efficiency was somewhat lower between AtTPK5 subunits, possibly reflecting a larger distance between AtTPK5 channel subunits or their C-termini. Also the BiFC analysis clearly detected AtTPK5 homomers. Homomeric complexes of either tonoplast AtTPK1 (Bihler et al., 2005) or plasma membrane AtTPK4 subunits (Becker et al., 2004), respectively, have been proven to be functionally active in electrophysiological studies.

The overall interpretation of these results has to take into account that for FRET in a two-dimensional space, like a biological membrane, the orientation of the fluorescent proteins could be more relevant than for FRET occurring in a three-dimensional space, such as the cytosol (Zacharias, 2003 and references therein). Furthermore, for the BiFC analysis it is expected that orientation effects are less problematic for homomer complementation than for heteromer complementation in a membrane. In heteromer associations the separation between the fluorescent proteins (fused to the membrane proteins) and the membrane (i.e. the relative positioning of the split fluorescent protein) could be different, resulting in less or no YFP restoration even if heteromers exist (false negatives). Also for FRET, an offset in height of fluorescent proteins relative to the membrane in a heteromer, which is absent for the homomer, can result in a reduced FRET efficiency. Yet, this offset should be very substantial in order to explain our negative heteromer results. If so, our analysis proves the existence of homomeric AtTPK/KCO combinations, but does not fully exclude that heteromers still exist.

Conclusions

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

A substantial prerequisite for functional analysis of channel proteins in reverse genetic and transgenic approaches is knowledge about the expression pattern, the subcellular localization and possible channel multimerization. Expression studies on AtTPK/KCO genes revealed distinct expression domains that overlap in only a few tissues of Arabidopsis. Intense localization studies elucidated the vacuolar membrane as the final subcellular target of five AtTPK/KCO proteins, whereas AtTPK4 is a pollen-specific, plasma membrane two-pore K+ channel (Becker et al., 2004). Moreover, in vivo interaction studies of fluorescently labelled channel subunits of vacuolar AtTPK/KCO proteins demonstrated the existence of AtTPK/KCO homomers, but not heteromers in plant cells.

Although our data strongly favour the formation of homomeric channels in planta, we cannot fully disregard the possibility that heteromeric channel subunit combinations occur in plant cells at developmental stages or under physiological conditions that were not tested here. However, the inability to form efficiently heteromeric channels at the target membrane (i.e. the tonoplast) even when co-expressed under the control of the strong CaMV 35S promoter, leads us to conclude that this is an unlikely event for the tested AtTPK/KCO proteins under in vivo conditions.

Closely related K+ channel subunits have the ability to co-assemble. Different subunits can associate, leading to heteromeric channels with often different properties compared to homomeric channels. This provides a mechanism to create additional diversity in K+ channel activity within individual cells. The ability to form homomeric and heteromeric channel complexes is known for tandem-pore channels from the animal KCNK family (Berg et al., 2004; Kang et al., 2003; Lesage et al., 1996), Kir-type channels in tetramers (Jan and Jan, 1997; Zhu et al., 2003) and plant Shaker channels (reviewed in Chérel, 2004; Dreyer et al., 2002). The presence of only homomeric AtTPK/KCO channels in plant cells would reduce the spectrum of AtTPK/KCO-mediated K+ currents but would also indicate that each AtTPK/KCO protein has a distinct function. Further, this observation suggests that the regulation of such K+ currents in plant cells mainly results from an interaction with regulatory proteins or other molecules (e.g. 14-3-3 proteins, Ca2+). So far, regulatory effects of cytosolic Ca2+ have been shown for homomeric AtTPK1 channels after heterologous expression in yeast (Bihler et al., 2005).

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Nomenclature of AtTPK/KCO channels

The original nomenclature of the protein family was (according to Mäser et al., 2001) AtKCO1 to AtKCO6. Because of different functional properties of some of the family members (AtKCO1, AtKCO4), Becker et al. (2004) suggested renaming the channel proteins (including the renumbering of AtTPK3, which was previously AtKCO6). This new nomenclature has already been used in subsequent publications (Bihler et al., 2005; Harada and Leigh, 2006).

Plant material

Arabidopsis thaliana (L.) Heynh. cv. Col-0 wild-type and transgenic plants (see below) were grown in half-concentrated Murashige–Skoog (MS) medium supplemented with 1% sucrose and solidified with 0.7% agarose under a 16-h day (140 μE, 22°C)/8-h (22°C) night regime, or in soil (Einheitserde Typ GS90, Gebrüder Patzer, Simtal-Jossa, Germany; 16-h fluorescent light, 60, 120 or 180 μE, 20°C, 60% relative humidity/8-h dark, 16°C, 75% relative humidity). Transgenic A. thaliana plants were generated by vacuum infiltration with Agrobacterium tumefaciens strain GV3101. Kanamycin- or hygromycin-resistant plants (T0) were identified and grown from seeds. Experiments were conducted with T1 or T2 plants, and two to four independent lines each of the T2 generation were selected for detailed analysis of GUS activity. Cell cultures were grown at 22°C with shaking in darkness.

Onion cell bombardment

Particle bombardment was used to introduce the AtTPK/KCO–YFP/CFP fusion plasmids into onion epidermal cells with a Biolistic PDS-1000/He system (Bio-Rad, Munich, Germany). Onions were obtained from a local supermarket and the epidermal pieces were placed on agar plates containing 1× MS. Gold particles (Bio-Rad, 0.6 μm) were coated with the respective plasmid DNA. Gold aliquots (2.5 mg; 20 μl 99.8% ethanol) were incubated with 5 μg of DNA, 20 μl of 0.1 m sperimidine and 50 μl of 2.5 m CaCl2, washed and resuspended in 50 μl 99.8% ethanol. For one shot 1 μg DNA per plasmid, coated on 500 μg gold was used. The onion epidermal pieces were bombarded using 900 p.s.i. rupture discs under vacuum. After bombardment the onion epidermal pieces were kept in darkness at 22°C for 48 h on media plates.

Protoplast transformation

The fusion constructs were introduced into A. thaliana protoplasts prepared from cell culture. The materials were incubated with enzyme solution (1% cellulase R-10; Duchefa Biochemie B.V., Haarlem, the Netherlands; 0.1% pectolyase, Kyowa Chemical Industry, Tokyo, Japan; and 0.1% BSA) at 26°C in darkness with gentle shaking (60 rpm) for 4–5 h. Purification of the resulting protoplasts was done using a Percoll or sucrose gradient. For transformation, a polyethylene glycol-mediated procedure was used (Lee et al., 2001). Bimolecular fluorescence complementation constructs were introduced into tobacco BY2 protoplasts and Arabidopsis protoplasts as described (Dreyer et al., 2004). Protoplasts were analysed for fluorescence 40–48 h after transformation. Vacuoles were isolated according to Czempinski et al. (2002).

Plasmids and constructs

Standard PCR reactions and sub-cloning procedures were used to engineer AtKCO3 and AtTPK sequences into clones described below. Complementary DNA clones were isolated from cDNA preparations from seedlings or flowers of A. thaliana (Col-0 ecotype). Complementary DNAs were cloned into plasmid pCR-Script (Stratagene, Heidelberg, Germany) or pCR2.1 (Invitrogen, Karlsruhe, Germany). Sequence determination was done for each construct. C-terminal GFP, YFP and CFP fusion constructs were created using pA7-GFP (see Supplementary Appendix S1), pEYFP-N1 and pECFP-N1 (Clontech, Heidelberg, Germany) and sub-cloning of YFP and CFP fusion constructs into pA7 (Von Schaewen, 1989). Bimolecular fluorescence complementation constructs have been generated using the pUC-SPYNE and pUC-SPYCE (Walter et al., 2004). AtTPK/KCO-GUS fusion constructs were generated using plant expression vectors pBI101.2 (AtTPK2, AtTPK3, AtTPK4) and pCAMBIA 1305.1 (AtTPK5, AtKCO3). 5′ Regions (1.0–2.5 kbp) containing the promoter and parts of the first translated exon of AtTPK/KCO genes were transcriptionally fused to the GUS gene. The AtTPK4–GUS construct was described in Becker et al. (2004). Details about cloning strategies are given in Supplementary Appendix S1.

Site-directed mutagenesis

Fusion constructs pA7-AtTPK1–YFP and pA7-AtTPK1–CFP were mutated to alter amino acid codon 206 [thereby introducing an alanine (A) to lysine (K) mutation] in YFP and CFP coding sequences, respectively, using the primers YFP(A206K).seq (5′-C TAC CTG AGC TAC CAG TCC AAGCTT AGC AAA GAC CCC AAC GAG-3′); YFP(A206K).rev (5′-GAG CAA CCC CAG AAA CGA TTCGAA CCT GAC CAT CGA GTC CAT C-3′); CFP(A206K).seq (5′-C TAC CTG AGC ACC CAG TCC AAGCTT AGC AAA GAC CCC AAC GAG-3′) and CFP(A206K).rev (5′-GAG CAA CCC CAG AAA CGA TTCGAA CCT GAC CCA CGA GTC CAT C-3′) (altered nucleotides indicated in bold). A silent mutation in L207 of both the YFP and CFP coding sequences inserted a HindIII restriction site in the mutagenized sequences (underlined). Mutagenesis was performed using the FlipFlop Site-Directed Mutagenesis Kit (Bioline, Luckenwalde, Germany). Sequence determination was done for each construct.

Analysis of GUS expression in transgenic plants

T1 or T2 seedlings or plants of transgenic A. thaliana lines (10 to 40 independent lines in T1; three different lines in T2) were used for histochemical analysis. Staining for GUS activity was done as described (Plesch et al., 2000). Plant tissues were incubated in staining solution at 37°C for 2–24 h.

Microscopy

Cells were settled onto clean glass slides covered with MS medium under glass cover slips. Images were obtained by conventional microscopy using a confocal laser scanning microscope (Leica DM IRBE inverse microscope with Leica TCS SP laser scanning unit) using a 10×/63× water objective lens. Fluorescence was observed using emission filter sets of 470–505 nm (CFP), 500–540 nm (GFP) and 525–650 nm (YFP). Excitation was achieved with an Ar/HeNe laser at 458 nm (CFP), 488 nm (GFP) and 514 nm (YFP). Acceptor photobleaching experiments were performed using the Leica application package ‘FRET acceptor bleaching’. Cells were bleached in the acceptor YFP channel by scanning a ROI using the 514-nm laser at 100% to decrease the signal intensity to 50%. Calculation of the FRET efficiency on selected membrane pieces was done as described by Bhat et al. (2005). Images were arranged using adobe photoshop (Adobe Systems, Mountain View, CA, USA).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The BiFC vectors pUC-SPYNE and pUC-SPYCE were kindly provided by K. Harter (Tübingen, Germany). The 35S-NRAMP3-GFP (in psmGFP327) for transient expression was kindly provided by S. Thomine (Gif-sur-Yvette, France). We thank Eike Kamann for expert technical assistance during molecular analyses and Karin Koehl and her colleagues from the MPI-MP Green Team for expert plant care. This work was supported by a grant of the Deutsche Forschungsgemeinschaft (CZ87/1-2) to KC, and by the European Union (NICIP; EU CT-2002-00245) to KC and BMR. CV receives financial support through the International PhD Programme ‘Integrative Plant Science’ (IPP-IPS) funded by the DAAD (Deutscher Akademischer Austauschdienst) and the DFG (Deutsche Forschungsgemeinschaft; DAAD Az. D/04/01336). BMR thanks the Fonds der Chemischen Industrie for financial support (No. 0164389). We would like to thank the anonymous reviewers for their helpful comments on the FRET/BiFC analyses.

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  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results and Discussion
  5. Conclusions
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Appendix S1. Cloning details of GFP-, YFP-, CFP-fusion constructs

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TPJ_2868_sm_AppendixS1.rtf21KSupporting info item

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