Acidic Residues Necessary for Pyrophosphate-energized Pumping and Inhibition of the Vacuolar H+-pyrophosphatase by N,N'-Dicyclohexylcarbodiimide*

(Received for publication, April 10, 1997, and in revised form, June 5, 1997)

Rui-Guang Zhen , Eugene J. Kim and Philip A. Rea Dagger

From the Plant Science Institute, Department of Biology, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6018

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES


ABSTRACT

On the basis of a revised topological model of the vacuolar H+-pyrophosphatase (V-PPase; EC 3.6.1.1) derived from the analysis of four published sequences using two structure-predicting programs, TopPred II and MEMSAT, eight acidic amino acid residues located near or within transmembrane alpha -helices were identified. The codons specifying these amino acids in the cDNA encoding the V-PPase from Arabidopsis thaliana were singly mutated to examine their involvement in pyrophosphate (PPi) hydrolysis and PPi-dependent H+ translocation and the functional significance of the similarities between the sequences encompassing Glu229 (227-245) of the V-PPase and the N,N'-dicyclohexylcarbodiimide (DCCD)-binding transmembrane alpha -helix of the c-subunits of F-ATPases (Nyren, P., Sakai-Nore, Y., and Strid, A. (1993) Plant Cell Physiol. 34, 375-378). Three functional classes were identified after heterologous expression of mutated enzyme in Saccharomyces cerevisiae. Class I (E119Q, E229Q, D573N, E667Q, and E751Q) mutants exhibited PPi hydrolytic and H+ translocation activities and DCCD sensitivities similar to wild type. The one class II mutant obtained (E427Q) was preferentially impaired for H+ translocation over PPi hydrolysis but retained sensitivity to DCCD. Class III (E305Q and D504N) mutants exhibited a near complete abolition of both PPi hydrolysis and H+ translocation and residual activities with decreased DCCD sensitivity. In none of the mutants was diminished insertion of the V-PPase into the membrane or an increase in the background conductance of the membrane to H+ evident. The decoupled character of E427Q mutants and the enhancement of H+ pumping in E427D mutants by comparison with wild type, in conjunction with the retention of DCCD inhibitability in both E427Q and E427D mutants, implicate a role for Glu427 in DCCD-insensitive H+ translocation by the V-PPase. The proportionate diminution of PPi hydrolytic and H+ translocation activity and conservation of wild type DCCD sensitivity in E229Q mutants refute the notion that Glu229 is the residue whose covalent modification by DCCD is responsible for the abolition of PPi-dependent H+ translocation. Instead, the diminished sensitivity of the residual activities of E305Q and D504N mutants, but not E305D or D504E mutants, to inhibition by DCCD is consistent with the involvement of acidic residues at these positions in inhibitory DCCD binding. The results are discussed with regard to the possible involvement of Glu427 in coupling PPi hydrolysis with transmembrane H+ translocation and earlier interpretations of the susceptibility of the V-PPase to inhibition by carbodiimides.


INTRODUCTION

The membranes constituting the vacuolysosomal complex of plant cells are unusual in possessing an H+ translocating inorganic pyrophosphatase (V-PPase1; EC 3.6.1.1) (2). The V-PPase bears no systematic resemblance to soluble PPases at the sequence level (3, 4) and is considered to belong to a fourth class of H+-phosphohydrolase distinct from the F-, P- and V-ATPases (4). Moreover, unlike the V-ATPase, which is ubiquitous in the membranes bounding the acidic intracellular compartments of all eukaryotic cells, the V-PPase appears to be restricted to plants and a few species of phototrophic bacteria (2, 5). Notwithstanding the intrinsic evolutionary interest of this phenomenon, it poses a problem: the lack of sequence-divergent homologs from phylogenically remote organisms. Because all published V-PPase sequences are from the same group of organisms, vascular plants, and exhibit greater than 85% sequence identity at the amino acid level (6), most attempts to identify conserved amino acid residues of potential mechanistic significance by sequence alignment procedures have been unproductive. Crucial, therefore, has been the development of methods for the expression of functional pump in the yeast, Saccharomyces cerevisiae (7, 8). When constructs of the yeast-Escherichia coli shuttle vector pYES2, containing the entire open reading frame of the cDNA (AVP; Ref. 9) encoding the Mr 66,000 substrate-binding subunit2 of the V-PPase from Arabidopsis thaliana are employed to transform S. cerevisiae, endomembrane-associated enzyme active in PPi-dependent H+ translocation is generated (7). Since the heterologously expressed pump is indistinguishable from the native plant enzyme, thereby establishing the sufficiency of AVP for the elaboration of active V-PPase in S. cerevisiae, approaches based on site-directed mutagenesis, epitope tagging, and expression of fusion proteins are now applicable to investigations of the membrane organization and catalytic mechanism of the V-PPase.

By the parallel application of mutational and protein chemical methods, we have demonstrated a specific requirement for a cytosolically oriented Cys residue at position 634 for inhibition of the V-PPase by maleimides and the dispensability of all conserved Cys residues, including Cys634, for catalysis (8, 10). Our current studies of the V-PPase are directed at elucidating the involvement of acidic (Asp, Glu) residues located near or within hydrophobic spans in substrate turnover and/or H+ translocation.

Two factors prompted investigation of these acidic residues. The first was the need to gain insight into the identity and location of acidic residues with the potential for undergoing cycles of protonation and deprotonation within the hydrophobic core of the membrane. On the basis of analyses of other H+ pumps and H+-coupled transporters, acidic residues associated with transmembrane spans might be expected to directly participate in H+ uptake, translocation, and release by the V-PPase. The second factor was the observations of Nyren et al. (1), who noted that the sequences encompassed by positions 227-245 of the V-PPase from Arabidopsis bear a resemblance to the C-terminal regions of the c-subunits of F-ATPases. The C-terminal sequence flanking Glu229 in AVP is 71, 65, and 67% similar (35, 47, and 39% identical) to Rhodospirillum rubrum c-subunit (positions 58-74), Pisum sativum chloroplast subunit III (positions 61-77), and P. sativum mitochondrial subunit 9 (positions 55-72) (Fig. 1). Since the c-peptide, the most highly conserved subunit of the H+-conductive Fo sector of F-ATPases, binds the hydrophobic carboxyl reagent, N,N'-dicyclohexylcarbodiimide (DCCD) at an acidic residue located in the middle of the second of the two transmembrane alpha helices of this polypeptide, to abolish H+ translocation, Nyren et al. (1) have proposed that the sequence flanking Glu229 of the V-PPase may assume an analogous function. Specifically, in view of the sensitivity of the V-PPase to inhibition by DCCD (11), it has been suggested that Glu229 is the residue whose covalent modification by this carbodiimide is responsible for the inhibition of PPi-dependent H+ translocation.


Fig. 1. Comparison of the deduced amino acid sequence of AVP (9) with the amino acid sequences of the c-peptides of representative F-ATPases. The F-ATPase c-peptide sequences shown are R. rubrum subunit c (38), P. sativum chloroplast subunit III (39), and P. sativum mitochondrial subunit 9 (40). The C-terminal sequences of the c-subunits and residues 227-245 of AVP are aligned. Identities and conservative substitutions are indicated by white and shaded boxes, respectively. The DCCD-reactive Glu residues of the F-ATPase subunits are shown in bold type. All characterized V-PPases contain the consensus sequence LFE(A/S)ITGYGLGGSSMALF (6).
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Here we present (i) a revised topological model of the V-PPase, which, unlike those reported previously (9, 12), is derived from the concerted application of multiple computer-based structural criteria to the deduced sequences of the polypeptides specified by the cDNAs from several independent sources; and (ii) the single substitution of most of the conserved Asp and Glu residues inferred to be located near or within transmembrane spans on the basis of this topological model. The results of modeling are consistent with a structure for the V-PPase incorporating 15 transmembrane spans, while the results of mutagenesis demonstrate that Glu229 is unlikely to play a role in H+ translocation or inhibition of the V-PPase by DCCD. Instead, the characteristics of the mutants, combined with the inferred topology of the V-PPase, are better accommodated by a scheme in which membrane-embedded residues Glu305 and Asp504 contribute to DCCD binding, whereas Glu427, which is located at the interface between a transmembrane span and its adjoining cytosolic loop, is required for coupling PPi hydrolysis with H+ translocation.


MATERIALS AND METHODS

Heterologous Expression and Mutagenesis of V-PPase

The cDNA encoding the V-PPase from A. thaliana (AVP; Ref. 9) was heterologously expressed in vacuolar protease-deficient S. cerevisiae haploid strain BJ5459 (MATa, ura3-52, trp1, lys2-801, leu2Delta 1, his3-Delta 200, pep4::HIS3, prbDelta 1.6R, can1, GAL) (8, 13). Transformation of BJ5459 with yeast-E. coli shuttle vector pYES2, containing the entire open reading frame of AVP inserted between the GAL1 promoter and CYC1 termination sequences (pYES2-AVP; Ref. 7), isolation of the Ura+ transformants and growth of the cells for the preparation of membranes were performed as described (8). E. coli DH5alpha and CJ236 (dut- ung-) were employed for the amplification of pYES2-AVP and the generation of single-stranded, uracilated template for site-directed mutagenesis, respectively.

Mutagenesis was performed directly on pYES2-AVP vector (8). In all cases the mutagenic oligonucleotides were designed to singly substitute each conserved Asp or Glu codon with an Asn or Gln codon on the basis of the cDNA sequence of AVP (9). The sequences of the eight oligonucleotides (positions of conserved Asp or Glu codons shown in bold type and positions of degeneracy shown in brackets) were: Glu119 right-arrow Gln, CGGCTCTGTT[C]AGGGATTCAGCAC; Glu229 right-arrow Gln, TCTTTTT[C]AGGCTATTACTGG; Glu305 right-arrow Gln, GGATCATATGCT[C]AAGCATCATGCGC; Glu427 right-arrow Gln, GTTTCGTCA-CT[C]AGTACTACACTAG; Asp504 right-arrow Asn, GGCAATT[A]ATGCTTATGGTCCC; Asp573 right-arrow Asn, CCACACCGTA[A]ATGTTTTGACC; Glu667 right-arrow Gln, CTTTGGAGTT[C]AGACCCTC-TCTGG; Glu751 right-arrow Gln, CATGGCTGTT[C]AGTCTCTTGTC. At four positions (Glu229, Glu305, Glu427, and Asp504), mutants in which the Asp codons were replaced by Glu codons or vice versa were also generated. The sequences of the four oligonucleotides used for this purpose were: Glu229 right-arrow Asp, TCTTTTTGA[C]GCTATTACTGG; Glu305 right-arrow Asp, GGATCATATGC-TGA[T/C]GCATCATGCGC; Glu427 right-arrow Asp, GTTTCGTCACTGA[C]TACTACACTAG; Asp504 right-arrow Glu, GGCAATTGA[G]GCTTATGGTCCC.

Uracilated single-stranded template DNA was isolated from pYES2-AVP-transformed E. coli CJ236, and site-directed mutations were introduced by second strand synthesis from the template using mutant oligonucleotides (14, 15). In all cases, mutagenesis was confirmed by sequencing the target region before yeast transformation. In selected cases, when a pronounced alteration of V-PPase function was observed, the sequence of the target region of the AVP insert of pYES2-AVP was determined after extraction of the vector from the yeast transformants.

Preparation of Vacuolar Membrane-enriched Vesicles

Yeast vacuolar membrane-enriched vesicles were prepared as described (8).

Reaction of V-PPase with N,N'-Dicyclohexylcarbodiimide

The standard mixture for reaction with DCCD contained 30 mM Tris-Mes buffer (pH 8.0), the indicated concentrations of ligands (Mg2+ as MgSO4, K+ as KCl, PPi as Tris-PPi) and membrane protein (9.7-10.7 µg/ml). Reaction was initiated by the addition of DCCD (0-500 µM dissolved in ethanol), and the samples were incubated at 37 °C for the times indicated. After terminating the reaction by the addition of Mg2+ (1.3 mM), the samples were cooled on ice before assaying aliquots for V-PPase activity. Control samples were treated in an identical manner after the addition of equal volumes of ethanol. All stock DCCD solutions were prepared fresh daily.

Measurement of V-PPase Activity and Protein

PPi hydrolytic activity was measured as the rate of liberation of Pi from PPi at 37 °C in reaction media containing 0.3 mM Tris-PPi, 1.3 mM MgSO4, 100 mM KCl, 1 mM NaF, 5 µM gramicidin-D, 1 mM Tris-EGTA, and 30 mM Tris-Mes (pH 8.0) (8). Since yeast-soluble PPase, unlike the V-PPase, is exquisitely sensitive to inhibition by fluoride (Kiapp (soluble PPase) = 20 µM; Kiapp (V-PPase) = 3.4 mM) (16), inclusion of 1 mM NaF in the assay media effectively abolishes the contribution of the former to total hydrolysis (8).

PPi- and ATP-dependent H+ translocation was assayed fluorimetrically using acridine orange (2.5 µM) as transmembrane pH difference indicator in assay media containing vacuolar membrane-enriched vesicles (200 µg), 100 mM KCl, 0.4 M glycerol, 1 mM Tris-EGTA, and 5 mM Tris-HCl (pH 8.0). Reaction was initiated by the addition of Tris-PPi (1.0 mM) to media containing MgSO4 (1.3 mM) in the case of V-PPase-mediated H+ translocation or by the addition of MgSO4 (3 mM) to media containing Tris-ATP (3 mM) in the case of V-ATPase-mediated H+ translocation. The decrease in fluorescence was measured at excitation and emission wavelengths of 495 and 540 nm, respectively (8). The initial rate of H+ translocation and steady state pH gradient were estimated as Delta F%/mg/min (at time zero) and Delta F%/mg (after 5-10 min), where Delta F% = percentage decrease in fluorescence as described (17). Coupling ratio (the ratio of the rate of H+ pumping to the rate of PPi hydrolysis) was estimated as (Delta F%/min)/(µmol of PPi hydrolyzed/min). Protein was estimated by a modification of the method of Peterson (18).

Western Analyses

For Western analyses of the heterologously expressed V-PPase, membrane samples were delipidated by extraction with acetone:ethanol (1:1; -20 °C) (19), dissolved in denaturation buffer, and subjected to one-dimensional SDS-polyacrylamide gel electrophoresis on 11% (w/v) slab gels in a Bio-Rad minigel apparatus (7). The electrophoresed samples were electrotransferred to 0.45-µm nitrocellulose filters in standard Towbin buffer (20), containing 10% (v/v) methanol for 30 min at a current density of 2.5 mA/cm2 in a Millipore semi-dry blotting apparatus. After reversible staining of the transferred protein bands with Ponceau-S, the filters were processed for reaction with antibody (PABHK1) raised against synthetic peptide with the sequence HKAAVIGDTIGDPLK, corresponding to positions 720-734 of AVP (9). Immunoreactive bands were visualized by successive incubations of the membrane filters with horseradish peroxidase-conjugated sheep anti-rabbit immunoglobulin G and buffer containing 0.03% (w/v) H2O2, 0.5 mg/ml diaminobenzidine, and 0.03% (w/v) NiCl2 (21).

Graphical Analysis

Data were fitted by nonlinear least squares analysis (22) using the Ultrafit nonlinear curve-fitting package from BioSoft (Ferguson, MO).

Computer Programs for Modeling V-PPase Topology

Two programs were employed to model the overall topology of the V-PPase: TopPred II and MEMSAT (membrane structure and topology). The TopPred II program, developed by Manuel G. Claros and Gunnar von Heijne (Karolinska Institute, Stockholm, Sweden) for Macintosh computers is a public domain software package for predicting the topology of both prokaryotic and eukaryotic membrane proteins by the concerted application of hydropathy analyses, the "positive-inside" rule and "charge-difference" rule (23). The MEMSAT program, developed by Jones et al. (24) for IBM PCs, is based on expectation maximization. From the distributions of amino acids compiled from membrane proteins, or portions thereof, of defined topology, the log-likelihood ratios (si) for domain classes are calculated for each of the 20 amino acids according to the expression si = ln (qi/pi) where pi is the relative frequency of occurrence of amino acid i in all the sequences in the data set and qi is the relative frequency of occurrence of the same amino acid in a particular domain. These si values or propensities are then used to equate a given sequence with a given topology.

The deduced amino acid sequences of the V-PPases encoded by the cDNAs isolated from A. thaliana (AVP, GenBankTM accession no. M81892) (9), Beta vulgaris (BVP1, L32792; BVP2, L32791) (25), and Hordeum vulgare (HVP, D13472) (12) were processed in parallel using both programs.


RESULTS

Revised Topological Model

A revised topological model of the V-PPase was derived from the deduced sequences of the polypeptides encoded by four cDNAs: AVP from A. thaliana (9), BVP1 and BVP2 from B. vulgaris (25), and HVP from H. vulgare (12). The model shown in Fig. 2 was the only one of the three predicted by the TopPred II and MEMSAT programs of Claros and von Heijne and Jones et al. (24) capable of accommodating a cytosolic orientation for both the C terminus and the hydrophilic loop containing the N-ethylmaleimide (NEM)-reactive cysteine, Cys634, inferred from the characteristics of apoaequorin fusions (26) and the results of peptide mapping and Cys mutagenesis, respectively (8, 10).


Fig. 2. Revised topological model of V-PPase from A. thaliana. The model was based on predictions made using the TopPred II and MEMSAT programs of Claros and von Heijne and Jones et al. (24), respectively, and the probable cytosolic disposition of C-terminal AVP-apoaequorin fusions (26). The same overall topology was predicted for all four of the V-PPase sequences analyzed: those from A. thaliana (AVP; Ref. 9), B. vulgaris (BVP1 and BVP2; Ref. 25), and H. vulgare (HVP; Ref. 12). Boxed sequences, transmembrane spans predicted by TopPred II; hatched sequences, transmembrane spans predicted by MEMSAT. The conserved Asp and Glu residues that were singly substituted in this study are shown in white against a black background.
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Examination of the structure of the Mr 66,000 subunit of the V-PPase by TopPred II consisted of three main stages. (i) The first stage was the construction of hydrophobicity profiles using a trapezoid sliding window (27). Depending on the height and width of the hydrophobicity maxima and the preset "upper cutoff" and "lower cutoff" values for the computed hydrophobicity indices, spans were categorized as either "certain" or "putative." (ii) The second stage was enumeration of the difference in representation of positively charged amino acid residues between the two sides of the membrane and tests of the adherence of any given model to the positive-inside rule, with the bias in favor of Arg and Lys residues in hydrophilic loops with a cytosolic disposition in most polytopic membrane proteins (28). (iii) The third stage was application of the charge-difference rule (29), wherein the net charge difference between the 15 N-terminal and the 15 C-terminal residues flanking the most N-terminal transmembrane span is computed. Transmembrane orientation is correlated with the disposition of charged residues in the immediate vicinity of the first membrane span. The segment C-terminal to the first span is generally positively charged with respect to the N-terminal flanking regions in membrane proteins possessing a luminally oriented N terminus (29).

Deployment of the MEMSAT program entailed analysis of segments of the sequence of the V-PPase in terms of their likelihood of being located within a particular topological element. Based on statistical analysis of the distribution of amino acids in membrane proteins, the MEMSAT program ranks amino acids according to their propensities for being associated with each of five types of topological element: two classes of hydrophilic loop, designated cytoplasmic (inside) loop (Li) and luminal (outside) loop (Lo), and three classes of transmembrane helix domain, designated helix inside (Hi), helix middle (Hm), and helix outside (Ho) (24).

The consensus structure consistent with the predictions from both programs, the disposition of Cys634, and the C-terminal apoaequorin fusion data was a 15-span model containing a luminally localized N terminus and cytoplasmically localized C terminus (Fig. 2). While MEMSAT ranked a 16-span model highest, with the additional span encompassing residues 743-761, two models containing 14 and 15 spans ranked just below this model. In the 14-span model, the two lowest scoring transmembrane spans in the 16-span model (V and VI) were excluded, thus preserving the orientation of the N and C termini and the remaining C-terminal spans. In the 15-span model, the last transmembrane span in the 16-span model (XVI) was excluded, thus transferring the C terminus from the luminal to the cytosolic face of the membrane while preserving the orientation of all of the other spans.

All three models were consistent with a cytosolic disposition for Cys634 (8, 10), but only one, the 15-span model, was compatible with the finding that fusion of apoaequorin with the C terminus of AVP generates a vacuolar membrane-associated polypeptide capable of sensing cytosolic Ca2+ in transgenic A. thaliana plants (26). Assuming that the fusion of apoaequorin with the C terminus does not, itself, change the topology of the V-PPase, these data constrain the C terminus to the cytosolic face of the membrane and exclude the 16- and 14-span models.

Notable is the basic equivalence between the predictions deriving from TopPred II and MEMSAT. Whereas TopPred II constrains the length of transmembrane spans at a specific value (21 amino acid residues in this study) and is based on the assumption that all spans are perpendicular to the phospholipid bilayer, MEMSAT selects the best fit within a user-defined range of minimum and maximum lengths (17-25 amino acids residues in this study), thereby diminishing bias in favor of any one angle of intersection. Nevertheless, the margins of 11 of the 15 spans predicted by the two programs differed by no more than 4 amino acid residues and the average length of the spans (21.5 by MEMSAT versus a fixed value of 21 for TopPred II) were virtually identical. Of the spans predicted by MEMSAT, only two, spans IX (408-425) and XV (671-687), were shorter than the 20 amino acids required to traverse the entire bilayer, but in both cases the counterpart helices predicted by TopPred II included all 17 of these residues. Accordingly, when the MEMSAT settings were altered to increase the minimum span length from 17 to 19 residues, the overall topology of the V-PPase was unchanged; spans IX and XV were simply lengthened.

The three transmembrane spans (V (230-252), VI (292-316), X (452-472)) not identified in the original 13-span model (9) were neglected because of the proximity of adjacent maxima in the hydrophobicity profiles (span X) and the use of window sizes so broad as to obscure hydrophobic segments adjacent to regions of extreme hydrophilicity (spans V and VI). TopPred II, by contrast, permitted better resolution of the neighboring transmembrane spans by the application of a narrower sliding window (11 amino acid residues).

Two of the three transmembrane spans overlooked in the original 13-span model, helices V and VI, were the least likely in terms of their hydrophobicity and expectation maximization scores. Their MEMSAT scores (339 and 449, respectively) were significantly higher than the default cutoff value of 100 but markedly lower than the average score of 2713 for the other transmembrane spans. However, because the low scores of these helices were largely attributable to Arg246 (in helix V) and Asp298 (in helix VI), each residue of which was predicted to be displaced by seven positions from the cytosolic face of the membrane according to MEMSAT, and therefore appropriately positioned for mutual electrostatic screening, both spans were retained in the model.

The orientation of the N terminus was deduced from the charge-difference rule (29). Examination of the N-terminal residues immediately adjacent to the first transmembrane span (positions 14-34) revealed no positively charged residues and two negatively charged residues (Glu9, Glu13), giving a net charge of -2. The corresponding regions of BVP1, BVP2 (25), and HVP (12) had the same charge, attributable to Glu13 in all three sequences, plus Glu9 in HVP and Asp9 in BVP1 and BVP2. Of the 15 amino acid residues on the C-terminal side of the first span of AVP, two (Arg36, Lys38) were positively charged and one (Asp42) was negatively charged, giving a net charge of +1. BVP1 and BVP2 had three positive charges and one negative charge (net charge +2), and HVP had six positive charges and no negative charges (net charge +6) in this region. A net charge difference of at least 3 (-2 for the N-terminal 15 residues before the first span versus +1 for the C-terminal 15 residues after the span for AVP, 4 for BVP1 and BVP2, 8 for HVP) in all four cases was consistent with a luminal orientation for the N terminus.

The cytosolically disposed loops of the 15-span model for AVP contain a significantly greater number of Arg and Lys residues than the luminally oriented loops (83% versus 17%, respectively). Further, the majority of the residues located in hydrophilic loops are cytosolically oriented (79.7%, 45.6% of total), in accord with the inside-positive rule (28) and with the expectation that the overall distribution of hydrophilic loops would be biased toward the side of the membrane responsible for catalysis and ligand binding.

Three Classes of Mutant

According to the 15-span model (Fig. 2), a total of eight conserved acidic amino acid residues (Glu119, Glu229, Glu305, Glu427, Asp504, Asp573, Glu667, and Glu751) were tentatively identified as being near or within putative transmembrane spans. To examine their involvement in PPi hydrolysis, H+ translocation, and DCCD inhibition, these residues were singly substituted. In all cases acidic residues were replaced with their corresponding amides; in those cases where acid right-arrow amide substitutions had an influence on V-PPase activity, enzyme containing structurally conservative Asp right-arrow Glu or Glu right-arrow Asp substitutions was also generated.

Three classes of V-PPase mutant were distinguishable on the basis of their hydrolytic and pumping activities after heterologous expression in S. cerevisiae strain BJ5459: (i) those exhibiting rates of PPi hydrolysis and PPi-dependent H+ translocation similar to wild type, (ii) those exhibiting selective impairment of H+ translocation, and (iii) those exhibiting gross impairment of both PPi hydrolysis and PPi-dependent H+ translocation (Table I and Fig. 3).

Table I. Effects of single substitutions of Asp and Glu residues on activity of heterologously expressed V-PPase

Vacuolar membrane-enriched vesicles were prepared from S. cerevisiae BJ5459 pYES2-AVP transformants in which the codons specifying the amino acids indicated here and in Fig. 2 had been mutated. Substitutions involving no change in charge are underlined.

Class Substitution H+ translocation
PPi hydrolysis Coupling ratio
Initial rate Steady state

 Delta F%/mg/min  Delta F%/mg µmol/mg/min  Delta F%/µmol PPi hydrolyzed
Wild type 164 209 1.0 158
I E119Q 125 185 1.3 94
E229Q 23 41 0.2 99
E229D 279 254 1.5 186
D573N 253 215 0.9 278
E667Q 58 129 0.5 119
E751Q 108 153 1.2 94
II E427Q 10 22 0.5 19
E427D 221 235 1.0 226
III E305Q NDa ND 0.1 NAb
E305D ND ND 0.3 NA
D504N ND ND 0.1 NA
D504E ND ND 0.0 NA

a ND, not detectable.
b NA, not applicable.


Fig. 3. PPi-dependent H+ translocation by vacuolar membrane-enriched vesicles prepared from pYES2-AVP-transformed S. cerevisiae BJ5459 cells expressing either wild type or mutated V-PPase. Membrane vesicles (200 µg of membrane protein) were assayed for H+ translocation with the fluorescent Delta pH indicator, acridine orange, in a total reaction volume of 0.9 ml containing 1.3 mM MgSO4, 0.4 M glycerol, 100 mM KCl, 1 mM Tris-EGTA, and 5 mM Tris-HCl (pH 8.0). Intravesicular acidification was initiated at the times indicated by the addition of 1 mM Tris-PPi.
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In all class I mutants except one (D573N), PPi hydrolytic activity and the rate and extent of PPi-dependent H+ translocation were diminished proportionately. Glu right-arrow Gln substitutions at positions 119, 229, 667, and 751 generated enzyme with at least 15% of wild type PPi hydrolytic and PPi-dependent H+ pumping activity and coupling ratios, enumerated as Delta F%/µmol of PPi hydrolyzed, similar to wild type (Type I). In the case of position 573, an Asp right-arrow Asn substitution resulted in an approximately 1.8-fold increase in coupling ratio, which, since the rate of PPi hydrolysis was unaffected, was almost exclusively attributable to an increase in the rate of PPi-dependent H+ translocation (Table I and Fig. 3). When Glu229 was replaced with Asp, rather than Gln, PPi hydrolytic activity and PPi-dependent H+ translocation were increased coordinately to values 1.4- and 1.7-fold greater, respectively, than wild type (Table I and Fig. 3).

The one class II mutant obtained, E427Q, retained 50% of wild type hydrolytic activity and less than 5% pumping activity and exhibited a coupling ratio of 12% of wild type (Table I). A conservative Glu right-arrow Asp substitution at this position restored wild type PPi hydrolytic activity and generated enzyme with a pump capacity 1.4-fold greater than wild type (Table I and Fig. 3).

The remaining two acid right-arrow amide mutants, E305Q and D504N, fell into class III. Both were completely deficient in PPi-dependent H+ translocation and mediated PPi hydrolysis at less than 10% of the wild type rate (Table I and Fig. 3). In contrast to what was found with class I and class II mutants, Glu right-arrow Asp and Asp right-arrow Glu substitutions at positions 305 and 504, respectively, did not restore PPi-dependent H+ translocation (Table I and Fig. 3). While replacement of the E305Q substitution with an E305D substitution increased PPi hydrolytic activity from 5% to 26% of wild type, the equivalent substitution, to Glu instead of Asn, at position 504 had the converse effect, decreasing activity from 10% to less than 1% of wild type (Table I).

Maintenance of Membrane Integrity and Efficiency of V-PPase Insertion

In none of the mutants was there an increase in membrane H+ conductance of sufficient magnitude to account for the effects of these substitutions. Vacuolar membrane-enriched vesicles harboring any of the V-PPase mutants achieved similar rates and extents of MgATP-dependent H+ translocation by the endogenous, chromosomally coded V-ATPase associated with this fraction as membranes containing wild type V-PPase (Fig. 4A), indicating that the background conductance of the membrane to H+ was unaltered by mutagenesis of the V-PPase. Moreover, the possibility of a substrate (Mg2PPi) elicited increase in H+ conductance associated with the decoupled mutant, E427Q, was excluded by the finding that the rate of MgATP-dependent intravesicular acidification of membranes containing this form of the enzyme was the same regardless of whether or not PPi was added to the V-ATPase assay medium (Fig. 4B). Similarly, the decreased PPi hydrolytic and/or H+ pumping activities of some of the mutants were not explicable in terms of a decrease in the amounts of intact membrane-associated V-PPase through premature maturation, changes in expression level, or expression of polypeptide with decreased stability. Vacuolar membrane-enriched vesicles from cells expressing either wild type or mutated V-PPase contained similar levels of PABHK1-reactive, Mr 66,000 (AVP-specific) polypeptide, irrespective of the type or position of the substitution, and in none of the membrane samples was there an indication of a change in the electrophoretic mobility of the PABHK1-reactive band (Fig. 5).


Fig. 4. ATP-dependent H+ translocation by vacuolar membrane-enriched vesicles prepared from pYES2-AVP-transformed S. cerevisiae BJ5459 cells expressing either wild type or mutated V-PPase.  A, ATP-dependent intravesicular acidification by membranes containing wild type or E229Q, E305Q, E305D, D504N or D504E mutated V-PPase. PPi was not included in the assay media. B, ATP-dependent intravesicular acidification by membranes containing E427Q-mutated V-PPase measured in the presence or absence of PPi. Membrane vesicles (200 µg of membrane protein) were assayed for H+ translocation as described in Fig. 3, except that ATP-dependent pumping was measured in reaction media containing 3 mM Mg2+ and 3 mM Tris-ATP. Tris-PPi (1 mM) was added when indicated.
[View Larger Version of this Image (11K GIF file)]


Fig. 5. SDS-polyacrylamide gel electrophoresis and Western analysis of wild type and mutated V-PPase after heterologous expression in S. cerevisiae BJ5459. Delipidated vacuolar membrane-enriched vesicles (5 µg of membrane protein) prepared from pYES2-AVP-transformed cells were electrophoresed, electrotransferred to nitrocellulose filters, and probed with V-PPase peptide-specific antibody, PABHK1. All of the bands migrated at Mr 66,000. Also shown are the PPi hydrolytic activities of the corresponding membrane samples before delipidation.
[View Larger Version of this Image (27K GIF file)]

Kinetics of Inhibition by DCCD

If DCCD inhibits the V-PPase through its interaction with a carboxyl group located in a transmembrane span, introduction of acid right-arrow amide substitutions at these positions in the heterologously expressed enzyme would be predicted to confer decreased sensitivity to this reagent.

As a prelude to optimizing the conditions for reaction with DCCD, the ligand requirements for inhibition were investigated. Of the ligands tested, Mg2+ was the only one that influenced the susceptibility of the wild type heterologously expressed enzyme to inhibition by DCCD (Fig. 6). In contrast to the requirements for protection of the V-PPase from inhibition by maleimides (8, 10), free PPi and K+ did not influence the inhibitory action of DCCD, and substrate, Mg2+ + PPi, did not confer any greater protection than Mg2+ alone. The I50 values for inhibition by DCCD were 150, 130, 120, and greater than 500 µM for membranes incubated in the absence of ligands and those incubated in the presence of 0.3 mM PPi, 50 mM K+, and 1.3 mM Mg2+, respectively (Fig. 6).


Fig. 6. Effects of PPi, K+, and Mg2+ on the kinetics of inhibition of the heterologously expressed wild type V-PPase by DCCD. Membranes (4 µg of membrane protein) were reacted with DCCD for 60 min, in 30 mM Tris-Mes (pH 8.0) containing either no ligand (bullet ), 50 mM KCl (open circle ), 0.3 mM Tris-PPi (black-triangle), or 1.3 mM MgSO4 (square ) at 37 °C. Values shown are mean PPi hydrolytic activities ± S.E. (n = 3).
[View Larger Version of this Image (17K GIF file)]

The kinetics of inhibition of wild type V-PPase by DCCD were consistent with a scheme in which the modification of two reactive sites on the enzyme is necessary for inactivation and Mg2+ confers protection by binding to a high affinity site. The time dependence of inhibition by DCCD was described by the integrated second order rate equation 1/A = 1/Ao + kt such that a plot of the reciprocal of V-PPase activity (A) at time t approximated a straight line of slope k and ordinate intercept 1/Ao, where Ao is activity at time zero and k is the rate constant (Fig. 7A). Mg2+ decreased the second order rate constant for inactivation by DCCD as a hyperbolic function of Mg2+ concentration to yield an apparent affinity constant of 10-15 µM (Fig. 7B).


Fig. 7. Time dependence of Mg2+-protectable inhibition of heterologously expressed wild type V-PPase by DCCD (200 µM). A, plot of reciprocal activity against time for vacuolar membrane-enriched vesicles treated with DCCD in media containing Mg2+ at concentrations of 1 (bullet ), 2 (open circle ), 5 (black-triangle), 10 (triangle ), 20 (black-down-triangle ), 30 (down-triangle), 50 (black-square), or 100 µM (square ). The data were fitted to the integrated second order rate equation 1/A = 1/Ao + kt, where Ao is activity at time zero, A is activity at time t, and k is a second order rate constant. B, plot of second order rate constants, estimated from the data in A, against Mg2+ concentration. Concentration of Mg2+ required for 50% diminution of k = 10-15 µM.
[View Larger Version of this Image (18K GIF file)]

A screen of all eight acid right-arrow amide mutants for Mg2+-protectable inhibition by DCCD revealed that only two (E305Q and D504N) were markedly less sensitive to DCCD (Fig. 8 and Table II). While acid right-arrow amide substitutions at positions 119, 229, 427, 573, 667, and 751 had little or no effect on DCCD inhibitability versus wild type, the corresponding substitutions at positions 305 and 504 diminished the inhibitability of the residual activity by more than 3- and 4-fold, respectively (Fig. 8). In none of the mutants, with the exception of E427Q, which showed an approximately 2-fold decrease, was Mg2+ protectability affected. In agreement with a requirement for acidic residues at positions 305 and 504 for inhibition by DCCD, replacement of the E305Q and D504N substitutions with E305D and D504E substitutions, respectively, restored wild type DCCD inhibitability (Fig. 8 and Table II).


Fig. 8. Sensitivity of representative V-PPase mutants to inhibition by DCCD in the presence or absence of Mg2+. Vacuolar membrane-enriched vesicles (4 µg of membrane protein) containing heterologously expressed E229Q-, E305Q-, E305D-, E427Q-, D504N-, or D504E-mutated V-PPase were treated with the indicated concentrations of DCCD in the absence (bullet ) or presence of Mg2+ (1.3 mM) (open circle ). The results of single concentration measurements of inhibition by DCCD in the absence or presence of Mg2+ for these and the other mutants generated (E119Q, E229D, E427D, D573N, E667Q, and E751Q) are summarized in Table II.
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Table II. Sensitivity of wild type and mutated V-PPase to Mg2+-protectable inhibition by DCCD

Vacuolar membrane-enriched vesicles were treated with the indicated concentrations of DCCD (µM) in reaction buffer with and without Mg2+ (1.3 mM) for 60 min, and PPi hydrolytic activity was measured. The values shown are activities expressed as a percentage of the activity of membrane vesicles treated identically with 0 µM DCCD. WT, wild type.

[DCCD] Mg2+ WT E119Q E229Q E229D E305Q E305D E427Q E427D D504N D504E D573N E667Q E751Q

0  - 100 100 100 100 100 100 100 100 100 100 100 100 100
100 + 92 80 86 89 102 92 77 93 88 107 93 72 103
100  - 47 35 46 32 75 54 33 40 78 49 39 60 55
250 + 106 62 91 81 93 104 60 66 87 97 71 71 66
250  - 13 9 20 14 63 25 10 10 69 36 7 21 21


DISCUSSION

Both of the predictive methods applied to the substrate-binding subunit of the V-PPase, TopPred II and MEMSAT, yielded a tentative model containing two transmembrane spans in addition to the 13 proposed previously (9) and an opposed orientation for the N and C termini (luminal N terminus; cytosolic C terminus). Although tests of the validity of the new model will depend eventually on the results of direct, independent, and complementary structural studies, we consider it a significant improvement over those proposed previously. The outcomes from both programs were remarkably similar for all four of the sequences analyzed, with only minor differences in the lengths and locations of a few of the transmembrane spans. The results of both were compatible with the available, albeit limited, biochemical data on the localization of Cys634 (8, 10) and the characteristics of AVP-apoaequorin fusions (26). By contrast, the 12-span model of the V-PPase proposed by Tanaka et al. (12), on the basis of hydropathy analyses of the deduced sequence of HVP alone, places Cys634 on the wrong side (on the luminal face) of the membrane and contains a highly improbable transmembrane span (span VI in their model) incorporating 30-40 amino acid residues.

It was by inspection of the revised topological model of the V-PPase that the eight acidic residues located within or near transmembrane spans were identified as targets for substitution. Although six other acidic residues with a similar disposition were evident from the model (Fig. 2), these were either not conserved in all four of the sequences analyzed (Glu13, Glu298, Asp324) or located in relatively hydrophilic environments (Glu225, Asp351, Glu645) and considered less likely to participate in transmembrane H+ translocation and/or DCCD binding. Three main conclusions, discussed below, derive from the results of substituting these acidic residues.

Glu229 Is Not Essential for PPi Hydrolysis or H+ Translocation

E229Q mutated V-PPase shows some impairment of PPi hydrolysis and H+ translocation but, since both processes are diminished in parallel, the diminution of coupling ratio is small. Moreover, in direct opposition to the imputed role of Glu229 in inhibition by DCCD, E229Q-substituted V-PPase is no less sensitive to inhibition by DCCD than wild type or E229D mutated enzyme, indicating that an acidic residue at this position is not required for inhibition by this reagent. On this basis, and in contrast to the speculations in Ref. 1, it is unlikely that the alignments between the V-PPase sequences C-terminal to Glu229 and the second DCCD-reactive transmembrane span of the c-peptides of F-ATPases (Fig. 1) signifies a functional equivalence. Similarly, Asp573 and Glu residues 119, 667, and 751 do not appear to be critical for PPi hydrolysis, PPi-dependent H+ translocation, or inhibition by DCCD. Substitution of these residues by their corresponding amides exerts little or no effect on PPi hydrolytic activity, H+ pumping, coupling ratio, or DCCD inhibitability.

Glu305 and Asp504 Are Critical for Catalysis and Contribute to DCCD Binding

E305Q and D505N mutants exhibit less than 10% wild type PPi hydrolytic activity, no detectable PPi-dependent H+ translocation, and residual activities markedly less sensitive to inhibition by DCCD than that of wild type enzyme. These characteristics, together with the recovery of DCCD inhibitability shown by E305D and D504E mutants, are consistent with the involvement of acidic residues at these positions in inhibition by DCCD. It is unlikely, however, that Glu305 and Asp504 are the sole residues involved in inactivation of the V-PPase by this carbodiimide. Doubly mutated enzyme containing acid right-arrow amide substitutions at both of these positions is no less sensitive than either single mutant to DCCD, implying the participation of residues other than Glu305 and Asp504. The finding that structurally conservative Asp right-arrow Glu or Glu right-arrow Asp substitutions cause a decrease in hydrolytic activity in the case of Glu504 and only a minor increase in activity in the case of Asp305 suggests that the steric constraints for catalysis are more stringent than for DCCD binding. Evidently, a difference of one methyl group in the carboxyl side chain is sufficient to severely impair catalytic function while leaving DCCD inhibitability unaffected.

Mg2+ appears to protect the V-PPase from inhibition by DCCD by interacting with a high affinity binding site. In agreement with the results from earlier studies, the kinetics of protection against DCCD inhibition are consistent with an affinity constant for Mg2+ of 10-15 µM. The steady state kinetics of substrate hydrolysis by the V-PPase approximate a scheme in which the binding of two Mg2+ ions, in addition to those associated with the substrate, dimagnesium pyrophosphate (Mg2PPi), is required for activity: a high affinity binding site with a binding constant of about 25 µM and a lower affinity site with a binding constant of 0.25-0.46 mM (30, 31). Protection of the V-PPase from inhibition by the water-soluble carboxyl-selective reagent, 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDAC), exhibits an Mg2+ concentration dependence consistent with a Km for free Mg2+ of 20-23 µM (32). However, despite their kinetic equivalence, it is improbable that these results denote a direct interaction between Mg2+ and the residues or domains that undergo covalent modification in all cases. Although the notion of direct protection is consistent with the finding that high affinity Mg2+ protection of the V-PPase not only applies to the carboxyl reagents, DCCD and EDAC, but also NEM, a sulfhydryl reagent, and phenylglyoxal, a reagent that preferentially reacts with guanidino side chains (31, 32) and the fact that one cytosolic loop, that between transmembrane spans XIII and XIV, containing the Cys residue (Cys634) responsible for substrate-protectable inhibition of the V-PPase by NEM (8, 10), also contains Arg, Asp, and Glu residues, it does not reconcile two critical findings. It fails to explain why inhibition by DCCD is second order while EDAC, NEM, and phenylglyoxal inhibit the enzyme with first order kinetics (32)3; it cannot account for the diminution of Mg2+-protectable DCCD inhibitability consequent on substituting Glu305 and Asp504, neither of which are located in this hydrophilic loop, with their corresponding amides. We therefore propose that Mg2+ exerts its high affinity effects indirectly through conformational coupling rather than through direct screening of the residues that would otherwise be covalently modified. By implication, Glu305 and Asp504, although contributing to DCCD binding, may not themselves bind Mg2+.

Another explanation compatible with the effects of mutagenesis on Mg2+-protectable inhibition of the V-PPase by DCCD is that neither Glu305 nor Asp504 directly participate in DCCD binding or that either Glu305 or Asp504 does but not the other. For instance, by neutralizing the beta - or gamma -carboxyl groups on the side chains of these residues, acid right-arrow amide substitutions at one or both of these positions may simulate the electrostatic screening action of Mg2+ binding and thereby diminish the sensitivity of the V-PPase to inhibition by DCCD in the absence of Mg2+, while at the same time impairing overall catalytic activity.

Glu427 Is Required for Efficient Coupling

E427Q mutants, although still active in PPi hydrolysis, mediate H+ translocation at less than 6% of the wild type rate to yield an 8-9-fold diminished coupling ratio. While it may be premature to conclude that these results demonstrate a direct role for Glu427 in H+ transfer, since its substitution by Gln might cause a structural change that indirectly effects enzyme function, the large recovery of wild type H+ pumping versus the modest increase in PPi hydrolytic activity by a Glu right-arrow Asp substitution nonetheless implies an important role for an acidic residue at this position for H+ translocation per se. Whereas the capacity for PPi hydrolysis is increased by only 2-fold when the E427Q substitution is replaced with an E427D substitution, the rate of H+ translocation is increased by more than 20-fold.

If Glu427 does indeed directly participate in H+ transfer, two corollaries follow. First, since E427Q- or E427D-mutated V-PPase is as sensitive to DCCD as wild type enzyme, inhibition by DCCD does not have a direct bearing on H+ translocation. By analogy with the results of protein chemical studies of F- and V-ATPases, the DCCD inhibitability of the V-PPase has been interpreted in terms of the participation of a DCCD-reactive residue in transmembrane H+-conduction (11). From the results presented here and the known reaction specificity of DCCD, such a conclusion is not warranted. Given that stable incorporation of dicyclohexylisourea and irreversible modification of a protein is contingent on the exclusion of water or other nucleophiles from the site of reaction (33), inhibition by DCCD simply indicates that catalytic activity is directly or indirectly dependent on carboxyl functions sequestered from bulk phase water; it does not automatically imply that the residue modified participates in H+ translocation. By the same token, lack of interaction of an acidic residue with DCCD, as is the case for Glu427, does not exempt it from involvement in H+ translocation.

Second, according to the revised topological model of the V-PPase, Glu427 has a cytosolic disposition (Fig. 2). If this residue is involved in H+ translocation, it seems likely that it forms part of an input channel responsible for the entry of H+ at the cytosolic face of the membrane. It is therefore conceivable that the E427Q mutation acts to neutralize the gamma -carboxyl group that would otherwise be present at this position, thereby blocking protonation of the other H+-carrying residues of the pump. Providing that the other H+-carrying residues in the transmembrane relay can switch into their output configuration at sufficient velocity, possibly as a result of a local increase in H+ activity due to a block in H+ transfer from this site in E427Q mutants, the pump will mediate futile cycling (become decoupled) and the coupling ratio will fall.

Similar decoupled mutants have been reported for bacteriorhodopsin (34), E. coli cytochrome-bo ubiquinol oxidase (35) and Rhodobacter sphaeroides cytochrome-c oxidase (36). In bacteriorhodopsin, four membrane-embedded Asp residues are essential for H+ translocation (37) and substitution of one of these has been shown to abolish H+ translocation while leaving the photocycle unaffected (34). In subunit I of E. coli cytochrome-bo ubiquinol oxidase, mutation of an Asp residue residing in a hydrophilic domain decouples H+ translocation from electron transfer (35). In R. sphaeroides cytochrome-c oxidase, mutation of an Asp residue at a position equivalent to that of cytochrome bo has a similar decoupling action (36).

The results reported here provide an indication of the identity of one potential starting point for PPi-driven H+ movement across the phospholipid bilayer by the V-PPase and dispel previous misconceptions of the mode of action of DCCD in this system, but many basic questions remain. It has yet to be determined if Glu305 and/or Asp504, because of their deeper insertion into the membrane, act at a point in the H+ relay downstream of Glu427 such that pump mutated at these positions, unlike pump mutated at position 427, is arrested in its output configuration, so stalling both H+ translocation and PPi hydrolysis. It is not known if H+-dissociable side chains other than those on acidic residues also participate in H+ translocation.


FOOTNOTES

*   This work was supported by Grant MCB93-05281 from the National Science Foundation (NSF) and Grant DE-FG02-91ER20055 from the Department of Energy (DOE) (both to P. A. R.) and DOE/NSF/USDA Triagency Plant Training Grant DE-FG02-94ER20162 awarded to the Plant Science Institute.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Dagger    To whom correspondence should be addressed. E-mail: parea{at}sas.upenn.edu.
1   The abbreviations used are: V-PPase, vacuolar H+-pyrophosphatase; DCCD, N,N'-dicyclohexylcarbodiimide; Mes, 4-morpholineethanesulfonic acid; NEM, N-ethylmaleimide; EDAC, 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide; V-, F-, and P-ATPases, vacuolar-, mitochondrial-, and plasma membrane-type ATPases, respectively.
2   The substrate-binding subunit of the V-PPase migrates at Mr 66,000 (64,500-72,000) on SDS gels, but its probable mass deduced from its amino acid sequence is 79-81 kDa. Thus, when referring to this polypeptide the terms "Mr 66,000 subunit" and "81-kDa subunit" are used interchangeably. Moreover, although it is now known that this subunit is probably the sole polypeptide species constituting the enzyme, it is referred to as the "substrate-binding subunit" because substrate hydrolysis and substrate-protectable covalent modification were the first functions assigned to it.
3   R.-G. Zhen and P. A. Rea, unpublished results.

ACKNOWLEDGEMENTS

We thank Dr. Elizabeth Jones (Carnegie Mellon University, Pittsburgh, PA) for the kind gift of S. cerevisiae strain BJ5459 and Dr. Yolanda Drozdowicz for critically reading this manuscript.


REFERENCES

  1. Nyren, P., Sakainore, Y., and Strid, A. (1993) Plant Cell Physiol. 34, 375-378 [Medline] [Order article via Infotrieve]
  2. Rea, P. A., and Poole, R. J. (1993) Annu. Rev. Plant Physiol. Plant Mol. Biol. 44, 157-180 [CrossRef]
  3. Cooperman, B. S., Baykov, A. A., and Lahti, R. (1992) Trends Biochem. Sci. 17, 262-266 [CrossRef][Medline] [Order article via Infotrieve]
  4. Rea, P. A., Kim, Y., Sarafian, V., Poole, R. J., Davies, J. M., and Sanders, D. (1992) Trends Biochem. Sci. 17, 348-353 [CrossRef][Medline] [Order article via Infotrieve]
  5. Baltscheffsky, M., and Baltscheffsky, H. (1993) in Molecular Mechanisms in Bioenergetics (Ernster, L., ed), pp. 331-348, Elsevier Science Publishers B.V., Amsterdam
  6. Zhen, R.-G., Kim, E. J., and Rea, P. A. (1997) Adv. Bot. Res. 25, 297-337
  7. Kim, E. J., Zhen, R.-G., and Rea, P. A. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 6128-6132 [Abstract]
  8. Kim, E. J., Zhen, R.-G., and Rea, P. A. (1995) J. Biol. Chem. 270, 2630-2635 [Abstract/Free Full Text]
  9. Sarafian, V., Kim, Y., Poole, R. J., and Rea, P. A. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1775-1779 [Abstract]
  10. Zhen, R.-G., Kim, E. J., and Rea, P. A. (1994) J. Biol. Chem. 269, 23342-23350 [Abstract/Free Full Text]
  11. Maeshima, M., and Yoshida, S. (1989) J. Biol. Chem. 264, 20068-20073 [Abstract/Free Full Text]
  12. Tanaka, Y., Chiba, K., Maeda, M., and Maeshima, M. (1993) Biochem. Biophys. Res. Commun. 190, 962-967 [CrossRef]
  13. Jones, E. W. (1991) Methods Enzymol. 194, 428-453 [Medline] [Order article via Infotrieve]
  14. Kunkel, T. A. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 488-492 [Abstract]
  15. Kunkel, T. A., Roberts, J. D., and Zabour, R. A. (1987) Methods Enzymol. 154, 367-382 [Medline] [Order article via Infotrieve]
  16. Baykov, A. A., Dubnova, E. B., Bakuleva, N. P., Evtushenko, O. A., Zhen, R.-G., and Rea, P. A. (1993) FEBS Lett. 327, 199-202 [CrossRef][Medline] [Order article via Infotrieve]
  17. Rea, P. A., and Turner, J. C. (1990) Methods Plant Biochem. 3, 385-405
  18. Peterson, G. L. (1977) Anal. Biochem. 83, 346-356 [Medline] [Order article via Infotrieve]
  19. Parry, R. V., Turner, J. C., and Rea, P. A. (1989) J. Biol. Chem. 264, 20025-20032 [Abstract/Free Full Text]
  20. Towbin, H., Staehelin, T., and Gordon, J. (1979) Proc. Natl. Acad. Sci. U. S. A. 76, 4350-4354 [Abstract]
  21. Harlowe, E., and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  22. Marquardt, D. W. (1963) J. Soc. Ind. Appl. Math. 11, 431-441
  23. Sipos, L., and von Heijne, G. (1993) Eur. J. Biochem. 213, 1333-1340 [Abstract]
  24. Jones, D. T., Taylor, W. R., and Thornton, J. M. (1994) Biochemistry 33, 3038-3049 [Medline] [Order article via Infotrieve]
  25. Kim, Y., Kim, E. J., and Rea, P. A. (1994) Plant Physiol. 106, 375-382 [Abstract/Free Full Text]
  26. Knight, H., Trewavas, A. J., and Knight, M. R. (1996) Plant Cell 8, 489-503 [Abstract/Free Full Text]
  27. von Heijne, G. (1992) J. Mol. Biol. 225, 487-494 [Medline] [Order article via Infotrieve]
  28. von Heijne, G. (1986) EMBO J. 5, 3021-3027
  29. Hartmann, E., Rapoport, T. A., and Lodish, H. F. (1989) Proc. Natl. Acad. Sci. U. S. A. 89, 5786-5790
  30. Leigh, R. A., Pope, A. J., Jennings, I. R., and Sanders, D. (1992) Plant Physiol. 100, 1698-1705
  31. Baykov, A. A., Bakuleva, N. P., and Rea, P. A. (1993) Eur. J. Biochem. 217, 755-762 [Abstract]
  32. Gordon-Weeks, R., Steele, S. H., and Leigh, R. A. (1996) Plant Physiol. 111, 195-202 [Abstract/Free Full Text]
  33. Hassinen, I. E., and Vuokila, P. T. (1993) Biochim. Biophys. Acta 1144, 107-124 [Medline] [Order article via Infotrieve]
  34. Heberle, J., Oesterhelt, D., and Dencher, N. A. (1993) EMBO J. 12, 3721-3727 [Abstract]
  35. Thomas, J. W., Puustinen, A., Alben, J. O., Gennis, R. B., and Wikstrom, M. (1993) Biochemistry 32, 10923-10928 [Medline] [Order article via Infotrieve]
  36. Fetter, J. R., Qian, J., Shapleigh, J., Thomas, J. W., Garciahorsman, A., Schmidt, E., Hosler, J., Babcock, G. T., Gennis, R. B., and Ferguson Miller, S. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 1604-1608 [Abstract]
  37. Mogi, T., Stern, L. J., Marti, T., Chao, B. H., and Khorana, H. G. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 4148-4152 [Abstract]
  38. Falk, G., and Walker, J. E. (1988) Biochem. J. 254, 109-122 [Medline] [Order article via Infotrieve]
  39. Huttly, A. K., Plant, A. L., Phillips, A. L., and Gray, J. C. (1990) Gene (Amst.) 90, 227-233 [Medline] [Order article via Infotrieve]
  40. Morikama, A., and Nakamura, K. (1987) Nucleic Acids Res. 15, 4692 [Medline] [Order article via Infotrieve]

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