TY - JOUR
AU - Dietz, Karl-Josef
AB - Abstract The plant vacuolar H+-ATPase takes part in acidifying compartments of the endomembrane system including the secretory pathway and the vacuoles. The structural variability of the V-ATPase complex as well as its presence in different compartments and tissues involves multiple isoforms of V-ATPase subunits. Furthermore, a versatile regulation is essential to allow for organelle- and tissue-specific fine tuning. In this study, results from V-ATPase complex disassembly with a chaotropic reagent, immunodetection and in vivo fluorescence resonance energy transfer (FRET) analyses point to a regulatory mechanism in plants, which depends on energization and involves the stability of the peripheral stalks as well. Lowering of cellular ATP by feeding 2-deoxyglucose resulted in structural alterations within the V-ATPase, as monitored by changes in FRET efficiency between subunits VHA-E and VHA-C. Potassium iodide-mediated disassembly revealed a reduced stability of V-ATPase after 2-deoxyglucose treatment of the cells, but neither the complete V1-sector nor VHA-C was released from the membrane in response to 2-deoxyglucose treatment, precluding a reversible dissociation mechanism like in yeast. These data suggest the existence of a regulatory mechanism of plant V-ATPase by modification of the peri-pheral stator structure that is linked to the cellular energization state. This mechanism is distinct from reversible dissociation as reported for the yeast V-ATPase, but might represent an evolutionary precursor of reversible dissociation. Introduction The plant vacuolar-type proton-translocating ATPase (V-ATPase) acidifies organelles along the secretory and the endocytotic pathway. Eight different subunits (VHA-A–VHA-H) form the cytosolic V1-sector and five subunits (VHA-a, VHA-c, VHA-c′′, VHA-d and VHA-e) form the membrane-integral sector V0 (Sze et al. 2002; Supplementary Fig. S1). V1 is dominated by its hexameric head with three copies each of VHA-A and VHA-B. Coordinated ATP hydrolysis within the three VHA-A copies is transduced into rotation of the central stalk (VHA-D/VHA-F) and finally drives rotation of the V0 proteolipid ring formed by six copies of VHA-c and VHA-c′′. Each proteolipid subunit bears a proton-binding site. Peripheral stalk subunits stabilize and anchor the head structure to the membrane-integral sector V0. The cytosolic N-terminal domain of VHA-a functions as a membrane anchor of the peripheral stalk (Grabe et al. 2000) whereas the core complex of the peripheral stalk is formed by at least two VHA-E/VHA-G heterodimers (Féthière et al. 2005, Seidel et al. 2005, Ohira et al. 2006, Kitagawa et al. 2007, Zhang et al. 2008). The additional peripheral stalk subunits VHA-C and VHA-H are hypothesized to be of regulatory importance (Sagermann et al. 2001). Distinct subcellular and organ-specific requirements for regulation of V-ATPase function resulted in the evolution of small gene families coding for plant V-ATPase subunits (Sze et al. 2002, Hanitzsch et al. 2007). In Arabidopsis thaliana, the V-ATPase consists of 13 distinct subunits encoded by 28 genes. A linkage between glycolysis and V-ATPase regulation was reported for the yeast V-ATPase, where aldolase enables a molecular switch via an interaction with VHA-B, VHA-E and VHA-a (Lu et al. 2004, Lu et al. 2007). Likewise V-ATPase and aldolase co-immunoprecipitate in rice (Konishi et al. 2004, Konishi et al. 2005). High glucose availability enhances V-ATPase activity, whereas lack of glucose results in reversible inhibition and the regulation involves major structural alterations within the yeast complex. This mechanism was termed ‘reversible dissociation’ (Kane 1995). Parra and Kane (1998) subsequently showed that the level of extracellular glucose supplementation regulates V-ATPase activity in vivo by affecting the extent of association between the V1 and V0 domains. This paper addresses the potential contribution of reversible dissociation to V-ATPase regulation in plants. To answer these questions, fluorescence resonance energy transfer (FRET), co-expression and pull-down assays as well as chaotropic disassembly tests were conducted. Based on the results, we suggest a new regulatory mechanism for the plant V-ATPase which involves alterations in peripheral stalk stability, depends on the cellular energization state and is modulated by VHA-E isoforms. Results Regulation of V-ATPase by reversible dissociation is thought to be linked to peripheral stator stability and should occur in a specific metabolic context. Therefore, the first part of the study addresses the interaction specificity of VHA subunits belonging to the peripheral stator, namely VHA-E, -G and -C. Interaction analysis of VHA-E and VHA-G isoforms Three isogenes each encode VHA-E and VHA-G in A. thaliana. Thus, nine potential combinations exist for heterodimer formation between VHA-E and VHA-G. It was unknown whether the different VHA-E isoforms are capable of forming heterodimers with each of the VHA-G isoforms. Untagged AtVHA-E was heterologously co-expressed with hexahistidine (His6)-tagged VHA-G in Escherichia coli under control of the T7 promotor and purified via Ni-NTA affinity chromatography. Eluted and dialysed proteins were probed with a specific antibody against VHA-E by Western blotting, which resulted in protein band detection at about 27 kDa. This corresponded to the expected size of untagged subunit AtVHA-E (Fig. 1). Elution fractions from control experiments obtained by loading the Ni-NTA column exclusively with isoforms of AtVHA-G did not contain the 27 kDa band. The respective band was also absent when VHA-E1 or -E2 was loaded, but was visible for AtVHA-E3. Here a non-conserved sequence of four histidine residues probably interacted with the Ni-NTA matrix. The results from co-expression analysis revealed a direct interaction between VHA-E and VHA-G. Apparently, the interaction lacked a significant isoform-specific preference (Supplementary Table S1). Fig. 1 View largeDownload slide Isoform-dependent ex vivo interaction between untagged AtVHA-E and His6-tagged AtVHA-G. Escherichia coli cells co-expressing the constructs were lysed and the proteins purified by Ni-NTA affinity chromatography. The eluates obtained with imidazole were separated by SDS–PAGE and VHA-E protein was visualized with anti-AtVHA-E antibody. Control experiments with E. coli exclusively expressing either AtVHA-E1 or AtVHA-E2 or single VHA-G isofoms gave no signal (not shown). The blot shows representative images from four experiments. Fig. 1 View largeDownload slide Isoform-dependent ex vivo interaction between untagged AtVHA-E and His6-tagged AtVHA-G. Escherichia coli cells co-expressing the constructs were lysed and the proteins purified by Ni-NTA affinity chromatography. The eluates obtained with imidazole were separated by SDS–PAGE and VHA-E protein was visualized with anti-AtVHA-E antibody. Control experiments with E. coli exclusively expressing either AtVHA-E1 or AtVHA-E2 or single VHA-G isofoms gave no signal (not shown). The blot shows representative images from four experiments. In order to scrutinize the relevance for the living plant cell of the interaction seen in co-expression experiments, protoplasts were co-transfected with VHA-E and VHA-G fused to cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP), respectively. The occurrence of interactions was tested by FRET measurements. The results were confirmed with swapped fluorophores. To this end, one set of experiments was performed with CFP-tagged VHA-E and YFP-tagged VHA-G and a second set involved YFP-tagged VHA-E and CFP-tagged VHA-G. The measured high FRET efficiencies corresponded to very short distances between AtVHA-E and -G isoforms and proved the presence of both proteins in close proximity in the same complex. FRET efficiencies between CFP and YFP ranged from about 22.5% up to 48.8% when testing the nine possible combinations (Table 1). The highest FRET efficiencies were observed between VHA-E1 and the three VHA-G isoforms in comparison with VHA-E2 and E3. The highest FRET efficiency of 49% was measured between VHA-E1 and VHA-G3. A FRET efficiency of 38.2% was recorded between the pollen-specific isoforms AtVHA-E2 and -G3. Interestingly the FRET efficiencies between VHA-G1 and the VHA-E-isoforms were significantly higher than between VHA-G2 and the VHA-E isoforms (P < 0.05). Regardless of the variation, the data demonstrated that any VHA-E isoform can form heterodimers with each VHA-G isoform. Table 1 Dimerization of VHA-E isoforms   VHA-E1  VHA-E2  VHA-E3  VHA-G1  VHA-G2  VHA-G3  VHA-E1  15 ± 1  16 ± 1  15 ± 2  39 ± 2  33 ± 2  49 ± 2  VHA-E2  16 ± 1  23 ± 1  12 ± 2  36 ± 2  22 ± 2  38 ± 2  VHA-E3  15 ± 2  12 ± 2  22 ± 2  34 ± 2  28 ± 2  32 ± 2    VHA-E1  VHA-E2  VHA-E3  VHA-G1  VHA-G2  VHA-G3  VHA-E1  15 ± 1  16 ± 1  15 ± 2  39 ± 2  33 ± 2  49 ± 2  VHA-E2  16 ± 1  23 ± 1  12 ± 2  36 ± 2  22 ± 2  38 ± 2  VHA-E3  15 ± 2  12 ± 2  22 ± 2  34 ± 2  28 ± 2  32 ± 2  FRET efficiencies of VHA-E isoforms in the combinations E1/E1, E1/E2, E1/E3, E2/E2, E2/E3 and E3/E3. Mean average values are shown, n > 100, and calculated FRET efficiencies of isoforms between AtVHA-E and AtVHA-G. The combinations of CFP- or YFP-tagged proteins E1/G1 (n = 93), E1/G2 (n = 109), E1/G3 (n = 138), E2/G1 (n = 110), E2/G2 (n = 116), E2/G3 (n = 135), E3/G1 (n = 81), E3/G2 (n = 84) and E3/G3 (n = 120) were calculated. Data were confirmed with swapped fluorophores and represent the mean averages ± SE. View Large Furthermore, additional FRET measurements between CFP- and YFP-tagged VHA-E isoforms revealed that each VHA-E is able to form dimers with itself as well as with other VHA-E isoforms (Table 1). Chaotropic disassembly of V-ATPase To study V-ATPase complex stability, isolated vacuoles were treated with increasing concentrations of the chaotropic salt potassium iodide (KI) (Adachi et al. 1990). The initial VHA-C content of vacuoles isolated from A. thaliana protoplasts was similar in glucose- and deoxyglucose-treated protoplasts (Fig. 2A). Deoxyglucose serves as a substrate for hexokinase. Its phosphorylation at the expense of ATP de-energizes the cell and inhibits glycolysis. Vacuolar membranes were sedimented and the proteins separated by SDS–PAGE. VHA-C content was visualized by Western blot analysis using the Det3 antibody (Schumacher et al. 1999), and band intensities were quantified densitometrically. KI at a concentration of 0.3 M destabilized the V-ATPase in the samples treated with MgATP or isolated from glucose- and deoxyglucose-treated cells. In a converse manner, AMP-PNP efficiently protected V-ATPase from disassembly. Even in the presence of 1 M KI, >70% of the initial VHA-C content remained associated with the membrane in the presence of AMP-PNP. The presence of MgATP or pre-treatment of the cells with glucose and deoxyglucose, respectively, resulted in a decrease of membrane-bound VHA-C in the presence of 0.3 M KI to about 60% (Fig. 2B). At higher KI concentrations in the range of 0.4–1 M, the densitometric analysis revealed a more stable membrane association of AtVHA-C with vacuolar membrane from control cells not incubated with deoxyglucose (Fig. 2B). In particular the membrane association of VHA-C from deoxyglucose-treated cells in comparison with glucose-treated cells was significantly lower at 0.4 and 1 M KI. VHA-C contents of tonoplasts isolated from 2-deoxyglucose-pre-incubated protoplasts strongly declined at a concentration of 0.4 M KI. Here the signal intensity at 0.4 M KI resembled the VHA-C contents of glucose-treated cells at much higher concentrations of 1 M KI (Fig. 2B). The results revealed a significant loss of complex stability upon deoxyglucose treatment. Fig. 2 View largeDownload slide Association of VHA-C with tonoplasts isolated from glucose- or deoxyglucose-pre-treated protoplasts. (A) Initial VHA-C content of glucose and deoxyglucose-treated cells. VHA-C content was analyzed by Western blotting. A representative image is shown. (B) Isolated protoplasts were incubated with 5 mM glucose except for deoxyglucose-treated cells; here glucose was replaced by 50 mM 2-deoxyglucose. Subsequently, vacuoles were isolated and incubated with different amounts of KI, and the membranes were sedimented and analyzed by Western blotting with anti-Det3 antibody. In addition, chaotropic disassembly was performed in the presence of AMP-PNP and MgATP, respectively. Data represent means of three densitometric analyses ± SD. Asterisks mark significant differences (P < 0.01) at a given concentration of KI. Fig. 2 View largeDownload slide Association of VHA-C with tonoplasts isolated from glucose- or deoxyglucose-pre-treated protoplasts. (A) Initial VHA-C content of glucose and deoxyglucose-treated cells. VHA-C content was analyzed by Western blotting. A representative image is shown. (B) Isolated protoplasts were incubated with 5 mM glucose except for deoxyglucose-treated cells; here glucose was replaced by 50 mM 2-deoxyglucose. Subsequently, vacuoles were isolated and incubated with different amounts of KI, and the membranes were sedimented and analyzed by Western blotting with anti-Det3 antibody. In addition, chaotropic disassembly was performed in the presence of AMP-PNP and MgATP, respectively. Data represent means of three densitometric analyses ± SD. Asterisks mark significant differences (P < 0.01) at a given concentration of KI. In order to address the physiological significance of such a regulatory mechanism based on enzyme stability, plants were subjected to heat stress or an elongated dark period (Fig. 3). Heat stress leads to enhanced synthesis and activity of ATP-dependent chaperones, e.g. of Hsp101 (Gurley 2000, Hong and Vierling 2001), and consequently results in intensive ATP consumption under heat stress. The extended dark period was applied to the plants, because the diurnal starch metabolism in Arabidopsis is strictly regulated so that the starch pool built up during the day is mostly mobilized during the night (Zeeman et al. 2007). Extended dark periods lead to carbohydrate deficiency and may impair glycolysis and the energy state. Fig. 3 View largeDownload slide Association of VHA-C with tonoplast membranes isolated from stressed plants. Plants were heat treated at 42°C for 1 h or maintained in darkness for 42 h. Isolated tonoplasts were incubated with 0.4 M KI and, as a control, in the absence of KI. The membranes were sedimented and analyzed by Western blotting with Det3 antibody (Schumacher et al. 1999). The blot shows a representative image from four experiments. Densitometric analyses were performed and the data were normalized to the corresponding control. Fig. 3 View largeDownload slide Association of VHA-C with tonoplast membranes isolated from stressed plants. Plants were heat treated at 42°C for 1 h or maintained in darkness for 42 h. Isolated tonoplasts were incubated with 0.4 M KI and, as a control, in the absence of KI. The membranes were sedimented and analyzed by Western blotting with Det3 antibody (Schumacher et al. 1999). The blot shows a representative image from four experiments. Densitometric analyses were performed and the data were normalized to the corresponding control. Both conditions were chosen due to their reduced energization level of the plant cell and hence were expected to result in reduced V-ATPase stability. Vacuolar membranes were isolated from control and stressed plants. The VHA-C content in the membrane fraction of control plants was set to 100%. Initially the membrane-associated fraction of VHA-C from stressed plants (dark, 125 ± 19%; heat stress, 123 ± 18%) slightly exceeded the membrane-associated VHA-C content from control plants. Western blots of isolated and 0.4 M KI-treated membranes from heat-stressed plants revealed a decrease of VHA-C associated with the membrane fraction. The VHA-C content dropped to 31 ± 9% of the control. In contrast, dark treatment resulted in an approximately 2-fold increased VHA-C content (194 ± 28%) in comparison with vacuolar membranes from control plants. While heat stress significantly impaired V-ATPase stability, the dark treatment surprisingly increased the V-ATPase stability as indicated by the improved membrane association of the peripheral subunit VHA-C after treatment with 0.4 M KI (Fig. 3). V0–V1 association in vivo The possibility of conditional occurrence of energy-dependent V-ATPase dissociation and the impact of the energization state on the overall structure in vivo were investigated by FRET analysis. To this end, the catalytic subunit VHA-A was fused to CFP and the membrane-integral VHA-a to YFP. Spatial changes monitored by altered FRET between the CFP and YFP tags attached to VHA-A and VHA-a would indicate structural alterations within the complex. In the absence of deoxyglucose the FRET efficiency was 45.5 ± 0.9% (mean ± SE; n = 223, Fig. 4A). In the presence of deoxyglucose FRET efficiency decreased to 39.6 ± 0.9% with high significance (P < 0.01) (mean ± SE; n = 163, Fig. 4). Despite the lack of knowledge on fluorophore orientation, this decline tentatively corresponds to a lateral movement of <1 nm. When protoplasts were incubated at 25°C instead of 4°C prior to FRET measurements, the obtained FRET efficiency was 47.0 ± 1.1% (mean ± SE; n = 118). In the case of hydrogen peroxide treatment, the FRET efficiency was 46.9 ± 1.2% (mean ± SE; n = 103). Thus, increasing the incubation temperature or application of oxidative conditions had no significant effect on V-ATPase assembly or structure (P = 0.305 and P = 0.346, respectively). Fig. 4 View largeDownload slide V-ATPase arrangement between various VHA subunits dependent on the metabolic state as revealed by in vivo FRET analysis. Energy transfer was measured for the FRET pairs E1-CFP/C-YFP (ncont = 149, ndeoxy = 82), E2-CFP/C-YFP (ncont = 147, ndeoxy = 100), E3-CFP/C-YFP (ncon = 137, ndeoxy = 89), E1-CFP/A-YFP (ncon = 76, ndeoxy = 105) and A-CFP/a-YFP (ncon = 223, ndeoxy = 163) under control conditions (5 mM glucose) or after treatment with 15 mM 2-deoxyglucose for 30 min. (A) The FRET pair VHA-E2/VHA-C showed significantly lower transfer efficiency than VHA-E1/VHA-C and VHA-E3/VHA-C (P < 0.01). Data represent means ± SE; * indicates significance of difference with P < 0.05. (B) Representative images for low (cytosolic CFP/YFP), medium (VHA-E1/VHA-C) and high (VHA-E1/VHA-G1) FRET efficiency. The images ‘FRET’ represent the raw data; ‘FRETcor’ denominates FRET images where CFP cross-talk and YFP direct excitation were subtracted. Quenching of CFP fluorescence and an increase of YFP emission due to energy transfer is visible in particular for the FRET-pair VHA-E1/VHA-G1. Fig. 4 View largeDownload slide V-ATPase arrangement between various VHA subunits dependent on the metabolic state as revealed by in vivo FRET analysis. Energy transfer was measured for the FRET pairs E1-CFP/C-YFP (ncont = 149, ndeoxy = 82), E2-CFP/C-YFP (ncont = 147, ndeoxy = 100), E3-CFP/C-YFP (ncon = 137, ndeoxy = 89), E1-CFP/A-YFP (ncon = 76, ndeoxy = 105) and A-CFP/a-YFP (ncon = 223, ndeoxy = 163) under control conditions (5 mM glucose) or after treatment with 15 mM 2-deoxyglucose for 30 min. (A) The FRET pair VHA-E2/VHA-C showed significantly lower transfer efficiency than VHA-E1/VHA-C and VHA-E3/VHA-C (P < 0.01). Data represent means ± SE; * indicates significance of difference with P < 0.05. (B) Representative images for low (cytosolic CFP/YFP), medium (VHA-E1/VHA-C) and high (VHA-E1/VHA-G1) FRET efficiency. The images ‘FRET’ represent the raw data; ‘FRETcor’ denominates FRET images where CFP cross-talk and YFP direct excitation were subtracted. Quenching of CFP fluorescence and an increase of YFP emission due to energy transfer is visible in particular for the FRET-pair VHA-E1/VHA-G1. Protein–protein interactions between the single copy VHA-C subunit and the VHA-E isoforms The in vitro formation of a subcomplex consisting of the subunits VHA-E, VHA-G and VHA-C has been shown for the yeast V-ATPase (Fethière et al. 2005). Subunit VHA-C is involved in reversible dissociation of V-ATPase in yeast (Smardon and Kane 2007). To investigate the potential occurrence of isoform-specific interactions, recombinant His6-tagged AtVHA-C and His6/Strep-tagged AtVHA-E1, -E2 and -E3 proteins were generated, dialyzed, loaded on an affinity column with strep-tactin–Sepharose and eluted with desthiobiotin. Negative controls lacking either subunit AtVHA-E (Fig. 5D) or AtVHA-C (Fig. 5E) demonstrated the inability of His-tagged VHA-C to bind to strep-tactin (Fig. 5D) or showed a single band of 35 kDa corresponding to His6/Strep-tagged subunit VHA-E (Fig. 5E). The His6-tagged 2-Cys peroxiredoxin was used as an unrelated control and did not co-purify with VHA-E (Fig. 5F, G). Thus co-purification of VHA-C depended on a specific interaction between Strep-tagged VHA-E and VHA-C (Fig. 5A–C). Western blot analyses of VHA-E and VHA-C combinations resulted in two bands with apparent sizes of 35 and 45 kDa, corresponding to the size of His6/Strep-tagged AtVHA-E and His6-tagged AtVHA-C, respectively, and revealed the absence of isoform specificity in vitro (Fig. 5). In the next experiments, interaction specificity was studied between VHA-C and VHA-E isoforms in living plant cells (Fig. 4B). Analysis of in vivo FRET between VHA-C–YFP and VHA-E isoforms fused to CFP showed isoform-specific energy transfer efficiency. FRET efficiencies for the combinations of VHA-E1/VHA-C and VHA-E3/VHA-C ranged around 20% whereas the FRET efficiency between VHA-E2 and VHA-C decreased significantly (P < 0.01, Fig. 4A) and was similar to those measured between freely diffusing cytosolic CFP and YFP (4.6 ± 1.3%) and between VHA-E1–CFP and cytosolic YFP (10.2 ± 2.0%) representing negative controls (P < 0.05). Fig. 5 View largeDownload slide In vitro protein–protein interaction between AtVHA-C and isoforms of AtVHA-E. Each image shows an immunoblot of chromatographic fractions obtained with protein combinations or single proteins as controls: (A) His6-VHA-C/pASK-VHA-E1, (B) His6-VHA-C/pASK-VHA-E2, (C) His6-VHA-C/pASK-VHA-E3, each co-purified via strep-tactin affinity chromatography. Image (D) shows an immunoblot of strep-tactin-purified His6-VHA-C and (E) of pASK-VHA-E1, respectively. VHA-C and VHA-E were detected with an anti-His antibody. Image (F) shows the failure of co-purification of 2-Cys peroxi-redoxin with VHA-E and served as a negative control for unspecific binding. Fig. 5 View largeDownload slide In vitro protein–protein interaction between AtVHA-C and isoforms of AtVHA-E. Each image shows an immunoblot of chromatographic fractions obtained with protein combinations or single proteins as controls: (A) His6-VHA-C/pASK-VHA-E1, (B) His6-VHA-C/pASK-VHA-E2, (C) His6-VHA-C/pASK-VHA-E3, each co-purified via strep-tactin affinity chromatography. Image (D) shows an immunoblot of strep-tactin-purified His6-VHA-C and (E) of pASK-VHA-E1, respectively. VHA-C and VHA-E were detected with an anti-His antibody. Image (F) shows the failure of co-purification of 2-Cys peroxi-redoxin with VHA-E and served as a negative control for unspecific binding. Incubation of co-transfected protoplasts with the glucose analog 2-deoxyglucose induced structural alterations within the complex as indicated by a significant decrease in energy transfer between VHA-E1/VHA-C and VHA-E3/VHA-C (P < 0.01). After 30 min FRET efficiency was as low as was observed for the VHA-E2/VHA-C pair or the negative controls (Fig. 4A). Deoxyglucose treatment had no effect on the low FRET value measured for the VHA-E2/VHA-C pair. As an additional control, FRET efficiency was also measured between AtVHA-E1–CFP and AtVHA-A–YFP. The FRET efficiency did not differ between deoxyglucose-treated or untreated cells (Fig. 4A). Vacuoles were isolated from transfected protoplasts in order to confirm the association of the chimeric proteins of VHA-C and isoforms of VHA-E with the tonoplast. Both the CFP and the YFP fluorescence were localized at the tonoplast (Supplementary Fig. S2). Vacuolar pH measurement and ATP content of A. thaliana mesophyll protoplasts The effect of 2-deoxyglucose treatment on vacuolar pH was monitored as an indicator for the V-ATPase H+ pumping activity in vivo. Isolated protoplasts were treated with 6-carboxyfluorescein diacetate (6-CFDA). Inside the cells 6-CFDA is hydrolyzed and the product, 6-carboxyfluorescein (6-CF), accumulates in the vacuoles. Fluorescence emission of 6-CF sensitively responds to pH changes which can be recorded from single cells by confocal microscopy using the excitation wavelengths 458 and 488 nm (Seidel et al. 2005). The pH value of leaf vacuoles was determined to be 5.8 and served as reference. The vacuolar pH of protoplasts was similar to the pH in leaves (Fig. 6). The pH of the surrounding W5 medium was 5.7. Increasing the surrounding pH to 7.3 resulted in a marginal but insignificant decrease of the vacuolar pH by 0.02 units. As an additional test, protoplasts were loaded with 6-CFDA and the effect on vacuolar pH with 50 nM of the specific V-ATPase inhibitor bafilomycin was analyzed during 15 min incubation. The 488 nm/458 nm ratio of 6-CF fluorescence emission increased significantly from 0.39 to 0.57, corresponding to an increase in pH of about 0.1 units. The data demonstrate on the one hand the sensitivity of the measurement and on the other hand that inhibition of V-ATPase immediately affects the vacuolar pH. The ratio of emission intensities depends on bulk phase pH. The vacuolar pH of deoxyglucose-treated cells was significantly higher (P < 0.05) than that of vacuoles in cells which had been incubated with glucose (Fig. 6). The pH increased significantly by 0.15 units as a consequence of incubation with deoxyglucose for 30 min (Fig. 6). It is concluded that cellular de-energization by deoxyglucose impaired vacuolar acidification. The vacuolar pH was also quantified in protoplasts transiently transfected with the three different VHA-E isoforms. Most striking was the significant increase of vacuolar pH by 0.4 units in the presence of VHA-E2, whereas VHA-E1 and VHA-E3 enabled vacuolar acidification similar to leaf tissue, untransfected protoplasts or protoplasts expressing YFP (Fig. 6). The maintenance of vacuolar acidity in protoplasts similar to leaves demonstrates the functionality of the VHA complexes bearing fluorescent fusion proteins. In addition, the collapse of vacuolar pH in protoplasts expressing VHA-E2 tentatively indicates a regulatory function of VHA-E. Fig. 6 View largeDownload slide Vacuolar pH of A. thaliana mesophyll protoplasts dependent on the metabolic state. Leaves and isolated protoplasts were loaded with 6-carboxyfluorescein diacetate (6-CFDA) for 1–3 h. Fluorescence from 6-carboxyfluorescein (6-CF) released in the vacuole was detected using the excitation wavelengths 458 and 488 nm (B, C). (A) The ratio E488/E458 nm indicates the vacuolar pH. Data were normalized on the leaf pH value and represent the means ± SE; * indicates significance of difference with P < 0.05. Asterisks mark significant differences (P < 0.01) in comparison with the control of glucose-treated cells. Fig. 6 View largeDownload slide Vacuolar pH of A. thaliana mesophyll protoplasts dependent on the metabolic state. Leaves and isolated protoplasts were loaded with 6-carboxyfluorescein diacetate (6-CFDA) for 1–3 h. Fluorescence from 6-carboxyfluorescein (6-CF) released in the vacuole was detected using the excitation wavelengths 458 and 488 nm (B, C). (A) The ratio E488/E458 nm indicates the vacuolar pH. Data were normalized on the leaf pH value and represent the means ± SE; * indicates significance of difference with P < 0.05. Asterisks mark significant differences (P < 0.01) in comparison with the control of glucose-treated cells. The ATP content of protoplasts was quantified by a coupled optical test with glucose-6-phosphate dehydrogenase. The deoxyglucose-treated protoplasts were characterized by a significantly reduced ATP content (ATPDOGlc = 100 ± 18 nmol mg−1 pheophytin; mean ± SE) as compared with glucose-fed cells (ATPGlc = 133 ± 17 nmol mg−1 pheophytin; mean ± SE). These data confirmed that deoxyglucose treatment resulted in a reduced energization of the cell. The leaf ATP content of heat-stressed plants (39 nmol ATP mg−1 pheophytin) and of plants that were incubated in the dark (31 nmol ATP mg−1 pheophytin) decreased in comparison with the control (42 nmol ATP mg−1 pheophytin). However, the ATP content of whole leaf tissue does not necessarily reflect the cytosolic ATP content (Gardeström and Wigge 1988). Discussion The V-ATPase represents a highly abundant primary transporter in eukaryotic cells with both housekeeping and stress-related functions (Lüttge et al. 2001). V-ATPase activity is regulated at several levels: in addition to transcriptional control (Hanitzsch et al. 2007), different biochemical modifications of V-ATPases are known to contribute to the regulation of ATP hydrolysis, proton transport and coupling of both functions. The V-ATPase also contributed to vacuolar acidification of protoplasts as shown in this study. However, V-ATPase is considered as a major energy sink in plant cells and thus its activity must be tightly controlled as suggested for the yeast and the Manduca sexta V-ATPase (Graf et al. 1996, Kane and Parra 2000). Several subunits are targets for protein phosphorylation. For example, VHA-C is a substrate of WNK8 kinases, and phosphorylated VHA-A interacts with 14-3-3 proteins in a light-dependent manner (Hong-Hermesdorf et al. 2006, Klychnikov et al. 2007). Although the substrate of the barley kinase CDPK1 is unkown, its activity was demonstrated to stimulate the V-ATPase (McCubbin et al. 2004). Coordinated stimulation of vacuolar pyrophosphatase (V-PPase) and V-ATPase represents an example for regulatory interplay between transporters at the tonoplast (Guo et al. 2006). Interdependent activity was also observed between Ca2+-ATPase, Ca2+/H+-antiporter and V-ATPase (Cheng et al. 2003). However, salinity results exclusively in enhanced V-ATPase expression in Suaeda salsa, whereas V-PPase is mostly unaffected (Wang et al. 2001). Another post-translational mechanism of V-ATPase regulation is given by reversible oxidation of VHA subunits so that oxidation of VHA-A leads to reversible inhibition of ATP hydrolysis in Zea mays and Hordeum vulgare (Hager and Lanz 1989, Dietz et al. 1998, Tavakoli et al. 2001). This work focused on the impact of the cellular energization state on V-ATPase structure and activity as a novel regulatory mechanism for plant V-ATPases. Structural implication of the energy state VHA-C was suggested to link VHA-E to the membrane-integral sector V0 (Li and Zhang 2004) and to act as a flexible stator subunit in regulating the coupling between ATP hydrolysis and proton transport (Drory et al. 2004). In this context, the coupling ratio of ATP hydrolysis and proton transport sensitively responds to nucleotide availability (Landolt-Marticorena et al. 1999). ATP binding to VHA-C was demonstrated by Armbrüster et al. (2005). ATP binding induces a structural change within VHA-C in vitro (Armbrüster et al. 2005). However, the functional implications of the altered V-ATPase structure have not been analyzed in vitro or in living cells, neither has the role of VHA-E isoforms. The results from this work demonstrate the dependency of V-ATPase stability on ATP availability. Decreased cellular energization following deoxyglucose treatment facilitated the release of VHA-C under chaotropic conditions (Fig. 2). It may be hypothesized that changes within VHA-C weaken the peripheral stator structure which in turn may regulate V-ATPase activity. This hypothesis is in agreement with the reduced ATP content of deoxyglucose-treated cells and is further supported by the in vivo measurements. Cells treated with deoxyglucose were characterized by a less acidic vacuolar lumen, indicating V-ATPase inhibition under these conditions (Fig. 6). On the other hand, the FRET efficiency between VHA-E and VHA-C dropped significantly when deoxyglucose was added and affected the overall structure of the V-ATPase, as demonstrated by decreased FRET efficiency between VHA-A–CFP and VHA-a–YFP (Fig. 4). The latter observations are in agreement with electron microscopic images from the Kalanchoë V-ATPase complex, where the peri-pheral stalk structure appeared diffuse and the V1 head tilted relative to V0 in the absence of nucleotides (Domgall et al. 2002). Interestingly, the pollen-specific isoform VHA-E2 (Strompen et al. 2005, Hanitzsch et al. 2007) showed a reduced FRET efficiency with VHA-C even in the presence of glucose (Fig. 4). It may be assumed that this isoform initially forms an inactive complex which becomes activated during subsequent development of the pollen. Comparison of V-ATPase regulation between A. thaliana and other species Complex destabilization was identified as an essential feature of V-ATPase regulation in starving yeast (Kane 1995). The mechanism depends on glucose and ATP availability (Kane 1995, Parra and Kane 1998). According to this model, the cytosolic sector V1 and VHA-C are released from the membrane upon glucose starvation. This mechanism inhibits ATP hydrolysis and proton transport under conditions of low phosphorylation potential and thus a low energization level (Kane 1995). Reassembly occurs within minutes after glucose repletion (Kane 1999). The dissociation is triggered by weakening the interaction between glycolytic aldolase and VHA-E (Lu et al. 2004). Hence the aldolase links V-ATPase activity directly to early reaction steps in glycolysis, but further glucose metabolism is required for reassembling the V-ATPase (Parra and Kane 1998). A similar mechanism was described for insect and mammalian V-ATPase (Sumner et al. 1995, Trombetta et al. 2003, Sautin et al. 2005, Lafourcade et al. 2008). Although aldolase and V-ATPase were co-immunoprecipitated in plants (Konishi et al. 2004, Konishi et al. 2005) a further characterization of this interaction is unavailable. The A. thaliana V-ATPase was more resistant to chaotropic conditions than reported for the yeast V-ATPase. The yeast V-ATPase complex loses its V1 subunits VHA-A, -B and -C in the presence of 0.3 M KI (Adachi et al. 1990). The oat V-ATPase of vacuolar vesicles is severly affected in the presence of MgATP even at concentrations of 0.1 M KI (Ward et al. 1992). In contrast, the A. thaliana V-ATPase of isolated vacuoles tolerated up to 0.6 M KI until about half of the VHA-C was released from the complex. MgATP had a stabilizing effect (Fig. 2). Under these conditions, the yeast and the oat V-ATPase are completely disassembled. In addition, the cold inactivation of mammalian V-ATPase is enhanced by addition of MgATP (Moriyama and Nelson 1989). Two V-ATPase isocomplexes are present in yeast that differ in their incorporated VHA-a isoforms Vph1p and Stv1p and display distinct catalytic properties. In contrast to yeast, the plant V-ATPase is characterized by several subunits that are encoded by small gene families. VHA-E and VHA-G each are encoded by three isogenes in A. thaliana, resulting in nine possible combinations of peripheral stalks and hence nine potential isoenzymes solely based on variation of these two subunits. This probably indicates a high flexibility of stator formation and adaptability of plant V-ATPase as indicated by the altered enzymatic activity of complexes that bear VHA-E2 (Fig. 6). Physiological relevance In contrast to, for example, yeast, complete inhibition of V-ATPase activity in response to low energization might be less important in photoautotrophic tissue. Interestingly, V-ATPases from heat-stressed plants as well as from plants grown in extended darkness differed in their stability. Survival during heat stress requires redirection of cell energy to repair mechanisms that, for example, involve ATP-dependent chaperone activity (Macario and Conway de Macario 2007). Reorganization of energy metabolism in leaf cells under heat stress also involves inactivation of the reductive pentose phosphate cycle by inhibition of Rubisco activase (Crafts-Brandner and Salvucci 2000). Likewise, down-regulation of V-ATPase activity as a major ATP consumer may be needed under these conditions in order to maximize energy delivery to cell repair mechanisms. The increased sensitivity of VHA-C to KI-mediated release from the heat-treated tonoplast supports this hypothesis. The extended dark treatment was expected to impair the V-ATPase stability as well. However, the treatment resulted in an increased stability (Fig. 3). Based on our hypothesis, this should reflect an increase of cytosolic ATP. In contrast, the measurements of total ATP revealed a reduced ATP content in dark-treated leaves. This can be explained by the observations made by Stitt et al. (1982). They reported that short-term incubation in the dark leads to an increase in cytosolic ATP content at the expense of mitochondrial and plastidic ATP and a decrease of total ATP concomitant with the increase of cytosolic ATP subsequent to dark exposure. A similar effect might explain the observed stability of the enzyme seen in this work. On the other hand, extended darkness might necessitate different cell responses for cell survival which involves increased stability, for example to establish lytic vacuoles, where high activity of V-ATPase is required to maintain cell metabolism (Frigerio et al. 2008). Solute mobilization from storage vacuoles might also depend on high V-ATPase activity. In any case, the observed V-ATPase modification in response to heat stress as well as extended darkness underlines the importance of controlling V-ATPase function during stress acclimation of plants. Conclusion This study shows that the regulatory alteration in peripheral stalk stability of plant V-ATPase is distinct from reversible dissociation in yeast, although VHA-C was identified as a key player in both mechanisms and both processes are ATP dependent. Recently, Diepholz et al. (2008) observed less dense structure of the yeast VHA-E/-G/-C subcomplex in solution than in the holocomplex, and suggested a conformational change during regulatory disassembly. In A. thaliana, structural alterations were observed, but obviously the complex remained fully assembled. The weakening of the peripheral stalk structure is a prerequisite for reversible dissociation. Thus the plant mechanism might correspond to an evolutionarily ancestral mechanism, which evolved prior to reversible dissociation in yeast, insects and mammals, or to a mechanism modified in the plant lineage as suggested by the differences between the oat and the A.thaliana V-ATPase. Materials and Methods Plant material Arabidopsis thaliana Col-0 was grown on soil in a growth chamber with constant humidity of 55% and a day/night cycle of 10/14 h. The light intensity was 240 μmol quanta m−2 s−1 with a temperature of 21°C in light and 18°C in darkness. Plants were harvested at an age of 4–6 weeks. Plants were heat stressed for 1 h at 42°C and incubated in the dark for an extended period of 42 h. C-terminal YFP and CFP constructs for in vivo analysis Full-length cDNA sequences of subunit AtVHA-A (At1g78900), AtVHA-C (At1g12840), the three isoforms of AtVHA-E (At4g11150, At3g08560 and At1g64200) and AtVHA-G (At3g01390, At4g23710 and At4g25950) were obtained from the TIGR database (http://www.tigr.org). The open reading frames were amplified from A. thaliana cDNA without the stop codon using gene-specific oligonucleotide primer pairs introducing flanking restriction sides for BamHI and AgeI, respectively (see Supplementary Table S2). All amplified products and the vectors 35S-YFP-NosT and 35S-CFP-NosT (Seidel et al. 2005) were digested with the restriction enzymes BamHI and AgeI [New England Biolabs (NEB)] using NEB-buffer1 and purified (Promega, Wizard®Plus DNA purification kit). For AtVHA-G2 AgeI was used as the restriction enzyme. Digested PCR products of AtVHA-A, AtVHA-C, each isoform of AtVHA-G (G1, G2 and G3) and AtVHA-E (E1, E2 and E3) were cloned as C-terminal fusions with 35S-YFP-NosT and with 35S-CFP-NosT, respectively. All constructs were sequenced. Cloning and expression for in vitro protein analysis The pDsRed2-C1 vector DNA (Biosciences Clontech) was restricted with AgeI and HindIII to excise the dsRED fluorescent protein-coding sequence. The T7 promotor was amplified by PCR from pCR T7/NT-TOPO-Vector (Invitrogen) (see Supplementary Table S2), digested with the restriction enzymes AgeI and HindIII and cloned into the pDsRed2-C1 vector, resulting in the new expression vector T7-KAN. Using the restriction sites KpnI and BamHI, the full-length sequences of the three AtVHA-E isoforms were cloned into this vector, resulting in the constructs T7-KAN-VHA-E1, -E2 and -E3. In addition, the coding sequences of the three AtVHA-E isoforms without the C-terminal stop codons were separately cloned into the pASK-IBA43 vector (IBA, Göttingen) using isoform-specific primers with the restriction sites SacII and NcoI, respectively. Full-length sequences of two AtVHA-G isoforms (VHA-G1 and VHA-G3) were amplified using gene-specific oligonucleotide primers containing the recognition sequences for BamHI and EcoRI, respectively. The primers for isoform G2 introduced restriction sites for AgeI and EcoRI, respectively. Following digestion of the vector His6–YFP (Seefeldt et al. 2008) with BamHI and EcoRI or AgeI and EcoRI, respectively, the PCR products were ligated with the linearized vector, resulting in His-G1, -G2 and -G3. The cDNA of subunit AtVHA-C was cloned in a similar way, with the restriction sites AgeI and BamHI resulting in the construct His6-C. Protein expression in E. coli strain BL21 Heterologous expression in E. coli strain BL21 was driven by the inducible T7 (T7-KAN, His-YFP) or TET promoter (pASK-IBA43). Bacteria were grown to OD600 = 0.6. Expression was induced by 0.5 mM isopropyl-β-d-thiogalactopyranoside (IPTG) or 0.2 μM anhydrotetracycline, respectively. After 4 h shaking at 37°C, bacterial cells were sedimented at 4°C and 5,000 × g. For heterologous co-expression, each VHA-E isoform was co-transformed with one of the three VHA-G isoforms. Expression of both proteins was ensured by kanamycin and ampicillin selection. Purification of recombinant protein His-tagged proteins were purified using Ni-NTA affinity chromatography. Bacterial sediments were resuspended in lysis buffer (50 mM NaH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8), supplemented with 0.5 mM lysozyme, incubated for 1 h at room temperature and centrifuged at 13,000 × g and 4°C for 40 min. The resulting supernatant was incubated for at least 1 h with 1 ml of Ni-NTA on a platform shaker at 4°C and then transferred to a PD-10 column (GE Healthcare). After settling, the Ni-NTA material was washed with 100 ml of washing buffer I (50 mM NaH2PO4, 300 mM NaCl, 20 mM imidazole, pH 8), 50 ml of washing buffer II [50 mM NaH2PO4, 300 mM NaCl, 50 mM imidazole, 20% (v/v) glycerol, pH 8] and finally eluted with 6–10 ml of elution buffer (50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8). The elution fractions were dialyzed against 40 mM potassium phosphate buffer (pH 7) overnight. In vitro protein–protein interaction studies of AtVHA-C and isoforms of AtVHA-E The purified proteins AtVHA-E1 and AtVHA-C were mixed in a ratio of 1 : 1, loaded onto a strep-tactin–Sepharose column and incubated for 30 min. The same procedure was performed with AtVHA-E2/AtVHA-C, AtVHA-E3/AtVHA-C, AtVHA-E1/2-Cys peroxiredoxin from Synechococcus elongatus PCC7942 (Stork et al. 2009) as well as solely VHA-C and VHA-E as negative controls. The columns were washed with 150 ml of 40 mM potassium phosphate buffer (pH 7) and proteins were eluted with 6 ml of elution buffer [100 mM Tris–HCl (pH 8), 150 mM NaCl, 1 mM EDTA, 2.5 mM desthiobiotin]. Elution fractions were analyzed by SDS–PAGE. Isolation of vacuolar membranes A 20 g aliquot of plant material was homogenized in 12 ml of homogenization buffer [250 mM sucrose, 50 mM Tris–HCl (pH 8), 8 mM EDTA], 0.8 ml of 1 M dithiothreitol (DTT), 24 μl of 0.1 M phenylmethylsulfonyl fluoride (PMSF), and the protease inhibitor cocktail Complete® (Roche) was added according to the manufacturer's protocol. The homogenate was filtered through 100 μm nylon net and diluted to a final volume of 100 ml with homogenization buffer. The suspension was centrifuged at 4°C and 7,000 r.p.m. for 10 min. Pellets were resuspended in 2.5 ml of resuspension buffer [250 mM sucrose, 5 mM PIPES-KOH (pH 7.2), 0.5 mM DTT], loaded onto 4 ml of sucrose buffer [35% sucrose, 5 mM PIPES-KOH (pH 7.2), 0.5 mM DTT] and centrifuged at 55,000 r.p.m. and 4°C for 90 min. Finally the interphase containing vacuolar membranes was subjected to SDS–PAGE and Western blot analysis. SDS–PAGE and Western blot Proteins were separated by reducing discontinous SDS–PAGE with 12.5% (w/v) acrylamide for the separating gel and 6% for the stacking gel. The samples were solubilized in SDS–PAGE sample buffer containing 125 mM Tris–HCl (pH 6.8), 2.5% (w/v) SDS, 10% glycerol and bromophenol blue (0.02%, w/v). For co-expression, Western blot analysis was performed with a primary rabbit antibody directed against VHA-E (Tavakoli et al. 2001) and the secondary antibody was coupled to alkaline phosphatase. Bands were detected after color development in carbonate buffer (100 mM NaHCO3, pH 9.8, 1 mM MgCl2) with 5-bromo-4-chloro-3-indolylphosphate and p-nitro-blue-tetrazolium chloride as substrates. To analyze VHA-E–VHA-C interactions, Western blots were performed with the primary mouse antibody directed against the poly-histidine tag (anti-His) (GE Healthcare). The secondary antibody was peroxidase conjugated and visualization was performed by chemiluminescence detection (Fisher Scientific). Detection of subunit AtVHA-C in vacuolar membrane fractions was done using the anti-Det3 antibody (Schumacher et al. 1999) as primary antibody and anti-rabbit-AP (alkaline phosphatase) IgG as secondary antibody. Isolation and transfection of mesophyll protoplasts The isolation and the polyethylene glycol-mediated transfection of A. thaliana protoplasts were performed as described before (Seidel et al. 2004). 2-Deoxyglucose as metabolic effector was added 30 min before analysis at concentrations as indicated. FRET measurements The constructs of 35S-A-CFP, 35S-A-YFP, 35S-a-YFP (Seidel et al. 2004), 35S-E-CFP (isoforms 1–3), 35S-E-YFP (isoforms 1–3), 35S-G-CFP (isoforms 1–3), 35S-G-YFP (isoforms 1–3) and 35S-C-YFP were used for co-transfection of A. thaliana mesophyll protoplasts. All isoform combinations of 35S-E-CFP/35S-E-YFP, 35S-E-CFP/35S-G-YFP, 35S-G-CFP/35S-E-YFP, 35S-E-CFP/35S-C-YFP, 35S-E-YFP/35S-C-CFP as well as 35S-E1-CFP/35S-A-YFP and 35S-A-CFP/35S-a-YFP were analysed using a confocal laser scanning microscope (Leica SP2). CFP and YFP were excited sequentially with the 458 and 514 nm lines of an argon ion laser, respectively. CFP emission was detected in the range of 470–510 nm and YFP in the range of 530–600 nm. FRET measurements and microscope settings were according to Seidel et al. (2005). The quantification of FRET efficiencies considered the direct excitation of YFP (correction factor ‘dirExYFP’ according to reference measurements) detected in the FRET channel (1) and the cross-talk of CFP into the FRET signal (0.61). The calculated FRET efficiency was normalized to the sum of emission intensities in both channels (2):   (1)  (2) pH measurement in A. thaliana mesophyll protoplasts Isolated protoplasts as well as total leaves were incubated with 5 μM 6-CFDA for 1–3 h. In vacuoles, the non-fluorescent 6-CFDA is hydrolyzed to the fluorescent 6-CF (Preston et al. 1989). The excitation spectrum of 6-CF exhibits two maxima that represent the protonated and deprotonated state of the fluorophore. 6-CF was excited sequentially with the 458 and 488 nm lines of an argon ion laser. The emission was separated using the band pass filter RSP 500 (Leica) and detected in the range of 500–530 nm according to Seidel et al. (2005). The emission intensities were set in relation to the detected fluorescence of 6-CF in leaf vacuoles. Isolation of intact vacuoles Isolated protoplasts were incubated with either 50 mM 2-deoxyglucose, 5 mM glucose, a combination of 50 mM 2-deoxyglucose and 1 mM AMP-PNP or 5 mM glucose and 1 mM MgATP, respectively, and lysed in lysis medium [200 mM sorbitol, 10% (w/v) Ficoll®, 20 mM HEPES-KOH (pH 8), 20 mM EDTA (pH 8), 0.1% (w/v) bovine serum albumin (BSA), 1 mM DTT] at 37°C for 30 min. Intact vacuoles were isolated by density gradient centrifugation. The gradient consisted of lysis medium, betaine medium [400 mM betaine, 30 mM K-gluconate, 20 mM HEPES-KOH (pH 7.2), 0.1% (w/v) BSA, 1 mM DTT] and a mixture of lysis and betaine medium in a ratio of 1 : 3. Peripheral stalk stability was analyzed by chaotropic disassembly. Vacuoles were incubated with increasing amounts of KI (Parra and Kane, 1996) on ice for 60 min and tonoplasts were sedimented at 21,000 × g and 4°C. Chaotropic disassembly was further performed in the presence of 1 mM MgATP and 1 mM AMP-PNP, respectively. ATP measurement Isolated protoplasts were incubated with 5 mM glucose or 50 mM 2-deoxyglucose for 30 min, sedimented and resuspended in 1 ml of 10% HClO4. After centrifugation (4°C, 13,000 × g, 5 min) the supernatant was adjusted to pH 7 using K2CO3. Extracts were mixed with buffer (100 mM Tris, pH 8.5, 5 mM MgCl2) in the ratio 1 : 8 and supplemented with 200 μM NADP+. Absorption was recorded at 340 nm, and reactions were started when the baseline was stable. Addition of 1.5 U of glucose-6-phosphate-dehydrogenase started the reduction of NADP+ proportionally to the amount of glucose-6-phosphate. A fructose-6-phosphate-dependent increase in NADPH/H+ absorption was achieved by addition of phosphoglucose-isomerase. 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TI - The Cellular Energization State Affects Peripheral Stalk Stability of Plant Vacuolar H+-ATPase and Impairs Vacuolar Acidification
JF - Plant and Cell Physiology
DO - 10.1093/pcp/pcr044
DA - 2011-04-06
UR - https://www.deepdyve.com/lp/oxford-university-press/the-cellular-energization-state-affects-peripheral-stalk-stability-of-0cC0GSFW0p
SP - 946
EP - 956
VL - 52
IS - 5
DP - DeepDyve
ER -