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J. Biol. Chem., Vol. 277, Issue 25, 22209-22214, June 21, 2002

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H⁺-Pyrophosphatase of Rhodospirillum rubrum

HIGH YIELD EXPRESSION IN ESCHERICHIA COLI AND IDENTIFICATION OF THE CYS RESIDUES RESPONSIBLE FOR INACTIVATION BY MERSALYL^*

Georgiy A. Belogurov^§, Maria V. Turkina^§, Anni Penttinen, Saila Huopalahti, Alexander A. Baykov^§¶, and Reijo Lahti

From the  Department of Biochemistry and Food Chemistry, University of Turku, FIN-20014 Turku, Finland and the ^§ A. N. Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow 119899, Russia

Received for publication, March 27, 2002

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

H⁺-translocating pyrophosphatase (H⁺-PPase) of the photosynthetic bacterium Rhodospirillum rubrum was expressed in Escherichia coli C43(DE3) cells. Recombinant H⁺-PPase was observed in inner membrane vesicles, where it catalyzed both PP_i hydrolysis coupled with H⁺ transport into the vesicles and PP_i synthesis. The hydrolytic activity of H⁺-PPase in E. coli vesicles was eight times greater than that in R. rubrum chromatophores but exhibited similar sensitivity to the H⁺-PPase inhibitor, aminomethylenediphosphonate, and insensitivity to the soluble PPase inhibitor, fluoride. Using this expression system, we showed that substitution of Cys¹⁸⁵, Cys²²², or Cys⁵⁷³ with aliphatic residues had no effect on the activity of H⁺-PPase but decreased its sensitivity to the sulfhydryl modifying reagent, mersalyl. H⁺-PPase lacking all three Cys residues was completely resistant to the effects of mersalyl. Mg²⁺ and MgPP_i protected Cys¹⁸⁵ and Cys⁵⁷³ from modification by this agent but not Cys²²². Phylogenetic analyses of 23 nonredundant H⁺-PPase sequences led to classification into two subfamilies. One subfamily invariably contains Cys²²² and includes all known K⁺-independent H⁺-PPases, whereas the other incorporates a conserved Cys⁵⁷³ but lacks Cys²²² and includes all known K⁺-dependent H⁺-PPases. These data suggest a specific link between the incidence of Cys at positions 222 and 573 and the K⁺ dependence of H⁺-PPase.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The proton pumping pyrophosphatase (H⁺-PPase)¹ is an integral membrane protein that utilizes the energy released upon hydrolysis of PP_i to transport protons across the membrane against the electrochemical gradient (1-3). PP_i is a by-product of various nucleoside triphosphate-dependent reactions, and its hydrolysis makes these reactions practically irreversible. Hydrolysis of PP_i in the majority of living species is accomplished by soluble pyrophosphatase that dissipates released energy as heat. H⁺-PPase conserves part of this energy in the form of the proton electrochemical gradient.

H⁺-PPases represent a distinct class of ion translocases with no sequence similarity to ubiquitous ATP-energized pumps such as F-, V-, or P-type ATPases or ABC transporters (4). In prokaryotic species, H⁺-PPase resides in the cytoplasmic membrane and pumps protons away from cytoplasm, whereas in eukaryotic species the enzyme acidifies internal organelles such as vacuoles in plants (2, 3) and acidocalcisomes in protozoa (5). The proton-motive force (µ_H⁺) generated is used to transport a variety of solutes via secondary transporters and osmoregulation (6). In the photosynthetic bacterium Rhodospirillum rubrum, H⁺-PPase is capable of sustaining both PP_i hydrolysis coupled with uphill proton translocation and PP_i synthesis in conjunction with downhill proton translocation at significant rates under physiological conditions (7-11). The R. rubrum H⁺-PPase (R-PPase) is therefore often referred to as PP_i synthase by analogy with ATP synthase (12). Both PPase and proton translocation activities are associated with a single polypeptide of 66-90 kDa (13-15), which possibly forms a dimer (16, 17). About half of the PPase molecule is embedded in the membrane, as estimated from the 14-16 transmembrane spans predicted by computer modeling (18, 19).

All H⁺-PPases display an obligate requirement for Mg²⁺. H⁺-PPases from plant vacuoles, acidocalcisomes of protozoa, and fermentative bacteria additionally require millimolar concentrations of K⁺ for activity, whereas those from respiratory and phototrophic bacteria and archaea are relatively monovalent cation-insensitive (2, 3). Recently, K⁺-independent H⁺-PPases were identified in plant and protozoa; however, the subcellular localization of these proteins remains to be determined (20-22). A K⁺ transporting function was proposed for K⁺-dependent H⁺-PPases (23), but this issue is still a matter of controversy (14, 24). Both K⁺-dependent and K⁺-independent H⁺-PPases are inactivated by sulfhydryl modifying reagents (25-30).

Although the first H⁺-PPase discovered was of bacterial origin (7, 8), the proteins from plant and protozoa have been characterized in greater detail by genetic engineering techniques (2, 3). Four of these protein orthologs have been heterologously expressed in yeast Saccharomyces cerevisiae in forms capable of both PP_i hydrolysis and H⁺ translocation (15, 19, 20, 31). The interest in bacterial H⁺-PPases significantly increased in 1998, when the gene for R-PPase was cloned and sequenced (32). This was followed by the identification of several bacterial H⁺-PPase genes as a result of genome sequencing projects. A putative bacterial H⁺-PPase gene (from the hyperthermophile Thermotoga maritima) has been expressed in the yeast system, yielding a protein capable of PP_i hydrolysis but not H⁺ transporting activity (33). However, yeast appears to be an unsuitable host for the expression of other bacterial H⁺-PPases (including R-PPase) because of proteolytic degradation of the recombinant protein (33).

Here we report the high yield expression of fully functional R-PPase in Escherichia coli and use this expression system in conjunction with site-directed mutagenesis to identify the Cys residues responsible for R-PPase inactivation by mersalyl. Furthermore, phylogenetic analyses performed in this study reveal a unique relationship between the mersalyl-reactive Cys residues and K⁺ dependence in H⁺-PPases.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Plasmid Construction-- Full-length R-PPase gene (1) was amplified from an original cDNA clone (32) by PCR using Pfu Turbo DNA polymerase (Stratagene). The primers, AACGACATATGGCTGGCATCTATC (forward) and TTTTCTCGAGTTAGTGGGCCAGCACCGC (reverse), incorporated artificial NdeI and XhoI restriction sites (underlined), respectively. The PCR product was digested with these restriction enzymes and inserted into the multiple cloning site of pET22b(+) (Novagen). The resulting construct was further manipulated by introducing a KpnI restriction site via a silent mutation, 0.5 kb upstream of the C terminus of the R-PPase gene. Mutagenesis was performed by an overlapping PCR technique with a Stratagene QuikChange^TM mutagenesis kit or ordinary PCR in the case of C573A. The primers employed in our experiments are listed in Table I. R-PPase-encoding regions of the constructs were sequenced to confirm the presence of the required mutations and/or the absence of secondary substitutions.

R-PPase Expression-- E. coli C43(DE3) cells (34) were transformed with wild-type or variant R-PPase-pET22b(+) constructs, and selected for antibiotic resistance on LB plates containing 100 µg/ml ampicillin. The cells were grown in 2× YT medium (35) supplemented with 70 µg/ml ampicillin (2× YT-amp) at 37 °C with shaking at 250 rpm. A single colony was used to inoculate 5 ml of 2× YT-amp. The cells grown for 4 h were transferred to 4 °C and stored overnight. The following day, the cells were collected by centrifugation at 3800 × g for 15 min at 4 °C, resuspended in 1 ml of fresh 2× YT, and transferred to 30 ml prewarmed 2× YT-amp. After 1 h of incubation at 37 °C, the cells were induced with 1 mM isopropyl--D-thiogalactopyranoside for 6 h and then harvested by centrifugation at 3800 × g for 15 min at 4 °C. Next, the cells were resuspended in 1 ml of ice-cold buffer A (120 mM Tris-HCl, 40 µM EGTA, 2 mM Mg²⁺, 10% glycerol, pH 7.5) and pelleted by centrifugation at 7000 × g for 10 min at 4 °C. The washed cell pellets were frozen and stored at 70 °C until use.

Isolation of Inner Membrane Vesicles (IMV)-- The cell pellets were thawed on ice, resuspended in 1 ml of ice-cold buffer A, and disrupted by sonication using a 100 W ultrasonic disintegrator (MSE Ltd., London, UK) with a microtip at an operating frequency of 20 kHz and a 9-µm amplitude for 2 min in an ice-water bath. Unbroken cells and cell debris were removed by centrifugation at 20,000 × g for 2 min at 4 °C. The supernatant was diluted 20-fold with buffer A, and the membrane fraction was harvested by centrifugation at 150,000 × g for 1 h. The resulting pellet was resuspended in 1 ml of buffer A, homogenized by brief sonication with a microtip at 4 µm amplitude for 15 s, frozen in liquid nitrogen, and stored at 70 °C until use.

IMV used in proton translocation measurements were isolated in buffer B (20 mM Tris-HCl, 40 µM EGTA, 200 mM choline chloride, 5 mM MgCl₂, 250 mM trehalose, pH 7.5). The washed cell pellets were not frozen, cell disruption was performed in four 30-s pulses with 1-min breaks, and the IMV pellet was resuspended by agitation at 4 °C for 2-3 h, instead of homogenization by sonication.

Protein concentrations in IMV suspensions were measured by the Bradford assay (36). IMV quantities were calculated in terms of protein content.

Polyacrylamide Gel Electrophoresis and Western Analyses-- Electrophoresis was performed with 12% gels containing 0.1% SDS (37). The gels were stained using a Silver Stain Plus kit (Bio-Rad) (38). Prior to electrophoresis, IMV (2.5 mg protein/ml) were solubilized by mixing with an equal volume of cold 20 mM Tris-HCl buffer, pH 7.5, containing 1.2 M MgCl₂, 50% glycerol, 5 mM dithiothreitol, and 4% Triton X-100. The mixture was allowed to stand on ice for 15 min and diluted 5-fold with 125 mM Tris-HCl buffer, pH 6.8, containing 20% glycerol, 300 mM dithiothreitol, and 2.5% sodium dodecyl sulfate, and 5 µl (1.2 µg of protein)/lane was loaded onto the gel.

For Western blot analyses, only 0.3 µg of protein was loaded per lane. The electrophoresed samples were transferred to a nitrocellulose Hybond^TM ECL^TM membrane (Amersham Biosciences) in standard Towbin buffer (39) containing 20% (v/v) methanol for 1 h at 100 V in a Mini Trans-Blot apparatus (Bio-Rad). Transferred protein bands were stained with Ponceau S, and the R-PPase antiserum-reactive bands (32) were visualized using an ECL kit (Amersham Biosciences).

PP_i Hydrolysis Measurements-- PP_i hydrolysis was assayed by continuously recording P_i liberation with an automatic P_i analyzer (40). IMV suspensions (5-50 µl) were preincubated for 1 min with 25 ml of 0.12 M Tris-HCl buffer, pH 7.5, containing 5 µM gramicidin D and 40 µM EGTA. The reaction was initiated by the addition of 0.1 mM PP_i and 2 mM MgCl₂. The liberation of P_i was monitored for about 3 min. In experiments measuring the inhibiting effect of mersalyl and the protective effects of MgCl₂, these reagents were included in the preincubation medium.

PP_i Synthesis Measurements-- PP_i synthesis was assayed continuously by a coupled enzyme procedure (41, 42), using ATP-sulfurylase to convert PP_i into ATP and luciferase to monitor ATP formation. The assay mixture (at a total volume of 0.2 ml) contained 5.2 mM potassium phosphate (2 mM MgP_i complex), 7 mM MgCl₂ (5 mM free Mg²⁺), 5 µl luciferin/luciferase solution reconstituted with 5 ml of water (Sigma ATP assay mix; catalog number FL-ASC), 0.7 unit/ml ATP-sulfurylase (Sigma), 10 µM adenosine 5'-phosphosulfate, 1 mM dithiothreitol, 1 mg/ml bovine serum albumin, and 0.15 M HEPES/KOH buffer, pH 7.5. The reaction was initiated by adding 1 µl of IMV suspension, and the time course of luminescence was monitored with an LKB model 1250 luminometer.

Proton Translocation Measurements-- H⁺ translocation across the IMV membrane was assayed fluorometrically in 2 ml of buffer B, using 50 µg of IMV and 3 µM acridine orange as a pH indicator (43). The excitation and emission wavelengths were set at 495 and 540 nm, respectively. H⁺ translocation was initiated by the addition of 0.1 mM PP_i, 0.1 mM ATP, or 15 mM DL-lactate (sodium salts) and measured with a PerkinElmer Life Sciences MPF-2A fluorometer. All of the measurements were performed at 25 °C.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Heterologous Expression of R-PPase in E. coli-- The R-PPase gene was cloned into pET22b(+) vector under control of the T7/lac promoter. E. coli strain C43(DE3) (34) was transformed with either pET22b(+) only or the vector containing R-PPase gene. The resulting cell lines were cultivated in rich medium (2× YT) in the presence of 1 mM isopropyl--D-thiogalactopyranoside to induce recombinant protein production. IMV prepared from the C43(DE3) cells transformed with pET22b(+) containing the R-PPase gene (RPP-IMV) produced on a silver-stained SDS-polyacrylamide gel an intense 60-kDa band that reacted with polyclonal antiserum raised against R-PPase (32). This band was absent from pET-IMV prepared from C43(DE3) cells transformed with pET22b(+) only (Fig. 1). It should be noted that although the molecular mass of R-PPase calculated from the amino acid sequence is 71 kDa, it migrates as a 60-kDa protein in SDS-polyacrylamide gel electrophoresis because of extreme hydrophobicity (13).

View larger version (25K): [in this window] [in a new window]   Fig. 1.   Polyacrylamide gel electrophoresis of pET-IMV (lanes 1 and 3) and RPP-IMV (lanes 2 and 4) in the presence of sodium dodecyl sulfate. Lanes 1 and 2, silver staining; lanes 3 and 4, Western analyses using antibodies against R-PPase.

Functional Characteristics of R-PPase in IMV-- RPP-IMV displayed salt wash-resistant PPase activity with the following characteristics distinctive of H⁺-PPase: hypersensitivity to the H⁺-PPase inhibitor, AMDP (>95% inhibition by 20 µM AMDP), and low sensitivity to soluble PPase inhibitor, fluoride (5% inhibition by 0.25 mM fluoride) (44). The specific activity of IMV PPase was 1.7 µmol/min/mg protein, which is eight times greater than that in R. rubrum chromatophores (IMV prepared from R. rubrum cells) (13, 30). PPase activity of IMV was not significantly affected by the monovalent cations, K⁺ or Na⁺ (50 mM), but increased by 60% in the presence of 1 µM of the uncoupler carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (gramicidin was omitted from the assay medium in this experiment), consistent with the formation of a proton concentration gradient.

In contrast, the PPase activity of control pET-IMV was 20 times lower than that of RPP-IMV. This residual salt-washable and entirely fluoride-sensitive (>95% inhibition by 0.25 mM fluoride) activity was attributed to contaminating soluble E. coli pyrophosphatase.

Addition of PP_i to RPP-IMV resulted in significant intravesicular acidification, as monitored by acridine orange fluorescence quenching (Fig. 2). The sensitivity of H⁺ transport activity to AMDP and fluoride was comparable with that of PP_i hydrolysis activity. Neither the initial rate nor the extent of the PP_i-dependent intravesicular acidification was significantly affected by monovalent cations (K⁺ and Na⁺) or valinomycin plus K⁺ (data not shown). The pET-IMV exhibited no H⁺ transport activity (Fig. 2). However, both types of IMV maintained the [H⁺] gradient generated by the intrinsic E. coli membrane proteins, F₁F₀-ATPase and D-lactate dehydrogenase (observed upon the addition of ATP and lactate, respectively), indicating IMV integrity (data not shown).

View larger version (13K): [in this window] [in a new window]   Fig. 2.   PPi-driven H+ transport in IMV containing recombinant R-PPase. The process was initiated by the addition of PPi to RPP-IMV (curves 1-3) or pET-IMV (curve 4) and terminated with 10 mM NH4Cl. Curves 2 and 3 were obtained in the presence of 1 mM KF and 2 µM AMDP, respectively.

RPP-IMV exhibited appreciable PP_i synthesizing activity (Fig. 3). Its absence from pET-IMV and sensitivity to AMDP clearly indicated that PP_i synthesis is catalyzed by R-PPase. The observed rate of synthesis decreased with time, supposedly because of the action of intrinsic ATPase on ATP formed from PP_i by the action of ATP sulfurylase. This explanation is supported by the decay of the signal generated by added PP_i in Fig. 3. The activity calculated from the initial slope of curve 1 in Fig. 3 was 0.010 µmol/min/mg protein (representing 0.6% hydrolytic activity). This PP_i synthesizing activity was evident in nonenergized vesicles and thus characterizes the approach to equilibrium in the P_i/PP_i system.

View larger version (14K): [in this window] [in a new window]   Fig. 3.   PPi synthesis catalyzed by IMV containing recombinant R-PPase. The reaction was initiated by the addition of RPP-IMV (curves 1 and 2) or pET-IMV (curve 3). At the end of the incubation, 0.2 µM PPi was added to calibrate the assay. Curve 2 was obtained in the presence of 10 µM AMDP.

Mersalyl-reactive Cys Residues of R-PPase-- H⁺-PPases are highly sensitive to sulfhydryl reagents, such as mersalyl and N-ethylmaleimide (2). Inactivation of R-PPase by mersalyl occurs biphasically (30), suggesting the involvement of at least two Cys residues. From the seven Cys residues present in the R-PPase sequence (32), four (cysteines 87, 268, 335, and 395) are located within predicted membrane regions (18, 32) and therefore can hardly react with charged, membrane-impermeable mersalyl. However, the remaining three Cys residues (cysteines 185, 222, and 573) occur within regions assigned to cytoplasmic loops and are thus potentially reactive. To determine the cytoplasmically oriented residues responsible for inactivation by sulfhydryl modifying reagents, each Cys was replaced with an aliphatic residue (Ala or Val), and the effects of these substitutions on PP_i hydrolyzing and proton pumping activities and sensitivity to mersalyl were determined.

None of the Cys substitutions affected the PP_i hydrolyzing and H⁺ pumping activities to an extent significantly exceeding batch-to-batch scatter (25%), thus demonstrating dispensability of these residues for catalytic activity and the lack of gross structural changes in R-PPase upon substitution. However, all three substitutions slowed down R-PPase inactivation by the sulfhydryl modifying reagent, mersalyl, to some extent, with C185A exerting by far the largest effect (Fig. 4). Two phases of inactivation were observed in most cases, including rapid inactivation (by 20-30%) and slower inactivation to nearly zero activity. In these experiments, IMV were always preincubated with mersalyl before the addition of PP_i. If mersalyl was added after PP_i and Mg²⁺, the hydrolysis rate rapidly dropped by ~20%, with all of the R-PPase variants exhibiting the rapid inactivation phase in Fig. 4, and remained constant thereafter. These observations indicate that MgPP_i has no effect on the rapid phase but abolishes the slower phase. No rapid phase was observed with the C222V variant, suggesting that this phase specifically corresponds to Cys²²² modification. The effects of mersalyl on the H⁺ transport activity of the R-PPase variants were qualitatively similar to those presented in Fig. 4 (data not shown).

View larger version (14K): [in this window] [in a new window]   Fig. 4.   Time course of wild-type and variant R-PPase inactivation in RPP-IMV by 15 µM mersalyl in the presence of 2 mM MgCl2. The ordinate is scaled logarithmically; 100% refers to activity measured without mersalyl. The lines correspond to the initial slopes of the curves.

Similar experiments conducted with Mg²⁺ omitted from preincubation conditions generated qualitatively similar inactivation curves but with much faster kinetics. The pseudo-first-order rate constants calculated from the initial slopes of the slower phase of the inactivation curves were approximately 10 times larger in the absence of Mg²⁺ than in its presence (Table II). A R-PPase mutant protein in which all three Cys residues were substituted was completely resistant to mersalyl.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

E. coli versus Yeast as a Host Cell for H⁺-PPase Expression-- Initially, Kim et al. (15) expressed H⁺-PPase (from the plant Arabidopsis thaliana) in the yeast S. cerevisiae. Yeast was selected for heterologous expression, because it is a vacuolated host lacking endogenous vacuolar H⁺-PPase. Later, five more H⁺-PPases, including two plant vacuolar (one from Vigna radiata (19) and the second from A. thaliana (20)), one protozoal (from Trypanosoma cruzi (31)), one archaeal (from Pyrobaculum aerophilum (45)), and one bacterial (from T. maritima (33)) enzyme were expressed in yeast. The general applicability of the yeast system for expression of H⁺-PPase was thus established. However, problems were encountered in bacterial H⁺-PPase expression, including low yield of T. maritima H⁺-PPase and complete failure to express functional R-PPase (33). These problems were not unexpected, because compartmentalization of yeast cells presents a significant challenge for H⁺-PPase lacking eukaryotic sorting signals and may lead to mislocalization, improper post-translational modification, and, consequently, degradation or inactivation of H⁺-PPase. Moreover, differences in the interpretation of topogenic signals by eukaryotic and prokaryotic protein insertion machinery (46) may result in incorrect topology upon insertion of the bacterial protein into endoplasmic reticulum membrane. In E. coli, bacterial H⁺-PPase is targeted and inserted directly into the cytoplasmic membrane, which is its usual environment. In addition, E. coli lacks endogenous H⁺-PPase and is by far the most manipulatable and resilient host available.

In direct contrast to the yeast system where the specific activity of H⁺-PPase in yeast vacuoles was at least twice as low as that in plant vacuoles (15, 20), significant overexpression of H⁺-PPase was achieved with the E. coli system. The specific activity of R-PPase in E. coli IMV was an order of magnitude higher than that in R. rubrum chromatophores, signifying the potential applicability of IMV as a rich source for R-PPase purification. The high expression levels achieved in the present work was mainly due to the E. coli C43(DE3) strain employed, which is particularly suitable for overexpression of integral membrane proteins (34). We were additionally able to express the R. rubrum protein in the conventional E. coli BL21(DE3) strain (Novagen), although the expression level was four times lower than that in C43(DE3) (data not shown).

Expressed R-PPase Is Fully Functional-- Heterologously expressed R-PPase was competent in PP_i hydrolysis and synthesis and PP_i-energized H⁺ translocation. To our knowledge, this is the first study reporting H⁺ transport activity in a heterologously expressed bacterial H⁺-PPase. Although T. maritima H⁺-PPase was recently expressed in yeast (33), no H⁺ transport activity was demonstrated in this case. The high H⁺ transport activity observed with the E. coli IMV containing R-PPase is particularly notable. The initial rate of fluorescence decay in Fig. 2 (1200 F%/min/1 mg of IMV) was about seven times higher than that in similar assays utilizing yeast microsomes containing A. thaliana H⁺-PPase (18, 20). The difference in H⁺ transport activities is also confirmed by a greater steady-state quenching of acridine orange fluorescence, specifically, 1800 F%/mg for R-PPase (Fig. 2) versus 209 F%/mg for A. thaliana H⁺-PPase (18).

R-PPase may act as a PP_i synthase in vivo (7, 12, 47, 48) and thus appears functionally similar to ATP synthase. The two synthases act in parallel, with similar rates in chromatophore membranes driven by the light-induced µ_H⁺ (7, 10, 11) or artificial pH across the membrane (49). We showed that nonenergized IMV containing R-PPase exhibited appreciable PP_i synthesizing activity, as previously demonstrated with soluble PPase (50, 51). In our system, converting PP_i into ATP by ATP-sulfurylase provided a thermodynamic pull for PP_i synthesis. Attempts to detect this activity with nonenergized chromatophores were unsuccessful, because the ATPase present in the chromatophore membrane strongly interferes with the PP_i assay by consuming ATP formed by ATP-sulfurylase.² In the current system, this interference was markedly reduced because of the greater PPase/ATPase ratio and only affected the linearity of PP_i accumulation curves (Fig. 3). IMV containing R-PPase thus provide an efficient system to study nonenergized PP_i synthesis and may be of great importance in elucidating the mechanism by which H⁺-PPase couples scalar PP_i synthesis with vectorial H⁺ transfer.

Contribution of Different Cys Residues to Mersalyl Sensitivity-- The heterologous expression of fully functional R-PPase has paved the way for site-directed mutagenesis, a powerful technique for studies on enzyme structure and function. Here, we used this approach to identify the cytoplasmically oriented Cys residues responsible for R-PPase inactivation by the membrane-impermeable sulfhydryl reagent mersalyl.

The data presented in Fig. 4 clearly show that Cys²²² modification is responsible for the rapid inactivation phase, resulting in the loss of ~20% of activity. The slower inactivation phase evidently involves Cys¹⁸⁵ modification, because mutation of this residue had the most significant effect on the rate constant for this phase (Table II). However, the C185A variant exhibited both inactivation phases in the absence of Mg²⁺ (Table II), suggesting that Cys⁵⁷³ modification also contributes to the slower inactivation phase. Significantly, modification of Cys⁶³⁴ (corresponding to Cys⁵⁷³ of R-PPase) inactivates plant vacuolar H⁺-PPase (28, 29).

Interestingly, Cys²²² replacement not only prevented the rapid MgPP_i-insensitive inactivation phase but also decreased the rate constant for the slower inactivation phase attributable to Cys¹⁸⁵ and Cys⁵⁷³ modification by ~5-fold (Table II). A simple explanation is that adding a bulky mersalyl group to Cys²²² induces structural changes that are propagated to the environment of Cys¹⁸⁵ and Cys⁵⁷³, thus increasing their accessibility to the modification agent. Consistent with this interpretation, neither Cys¹⁸⁵ nor Cys⁵⁷³ is completely accessible in wild-type R-PPase, as indicated by their lower reactivities in comparison with Cys²²².

Mg²⁺ and MgPP_i are two other determinants of Cys accessibility in R-PPase. Mg²⁺ binding decreased k_i by ~10-fold in both wild-type and variant R-PPases (Table II), whereas MgPP_i binding completely arrested the slower inactivation phase. These observations suggest that Mg²⁺ and MgPP_i induce conformational changes in R-PPase. In plant vacuolar H⁺-PPase, only Cys⁶³⁴ (corresponding to Cys⁵⁷³ of R-PPase) was modified by sulfhydryl reagents. Although Mg²⁺ had a negligible effect in this case, the effect of MgPP_i was comparable with that in R-PPase (28, 29). This difference indicates a greater conformational flexibility of R-PPase.

Conservation of Cys Residues and Subtyping H⁺-PPase Family-- Fig. 5 reveals a clear relationship between the evolution pattern of H⁺-PPase and the occurrence of Cys at positions corresponding to the mersalyl-reactive Cys²²² and Cys⁵⁷³ of R-PPase. The whole set of 23 nonredundant H⁺-PPase sequences can be divided into two subfamilies, specifically, I (from A. thaliana to Carboxydothermus hydrogenoformans) and II (from Brucella melitensis to P. aerophilum). All subfamily II H⁺-PPases (except P. aerophilum H⁺-PPase) but none of the subfamily I proteins contain an equivalent of Cys²²². On the other hand, an equivalent of Cys⁵⁷³ of R-PPase that is only sporadically distributed within subfamily II is identified in all subfamily I H⁺-PPases. The absence of any Cys in P. aerophilum H⁺-PPase is not unexpected, taking into account the strong selection against cysteines in this organism adapted to tolerate oxygen at elevated temperatures. Cys¹⁸⁵ is unique to R-PPase. Moreover, at least in one strain of R. rubrum (R2), this residue is replaced by Gly².

View larger version (35K): [in this window] [in a new window]   Fig. 5.   Subtyping H+-PPases on the basis of phylogenetic analyses and Cys conservation patterns. The neighbor-joining phylogenetic tree and multiple sequence alignment were generated by Clustal X, version 1.81. Poorly aligned regions corresponding to positions 1-30 and 249-332 of R-PPase were excluded from the otherwise full-length sequence set used for tree calculation. The confidence of the branching order was verified by making 1000 bootstrap replicates. The tree was arbitrarily rooted with the C. hydrogenoformans sequence to mark out subfamilies. The columns to the right of the tree display residues corresponding to Cys185, Cys222, and Cys573 of R-PPase in the multiple sequence alignment, K+ requirement of functionally characterized H+-PPases (K+ dep), and protein sequence accession numbers in the GenBankTM or DNA contig numbers for preliminary sequences obtained from The Institute for Genomic Research (www.tigr.org). The question mark in the K+ dep column specifies the organisms possessing K+-dependent H+-PPases whose amino acid sequences have not been directly determined and therefore may be different from those used to construct the phylogenetic tree, because these organisms possibly contain multiple H+-PPases.

Surprisingly, subfamily I includes all heterologously expressed H⁺-PPases displaying K⁺ dependence, whereas all heterologously expressed K⁺-independent H⁺-PPases are categorized as subfamily II. Furthermore, the presence of K⁺-dependent H⁺-PPase in four organisms classified as family I was demonstrated (52-55), although the sequences shown in Fig. 5 may belong to an alternative H⁺-PPase of these organisms. Assuming that the K⁺ requirement was eliminated or acquired at a distinct point in the evolution of H⁺-PPases, one can assign almost all the sequences from subfamily I and II to K⁺-dependent and K⁺-independent H⁺-PPases, respectively (Fig. 5). Pérez-Castiñeira et al. (33) also classified H⁺-PPases into two phylogenetic groups based on their K⁺ dependence. However, they demarcated the groups differently because they did not take into account Cys conservation.

The sporadic occurrence of Cys⁵⁷³ in subfamily II versus complete lack of Cys²²² in subfamily I suggests that subfamily II descends from subfamily I enzymes. Thus, the incidence of Cys at position 222 and loss of K⁺ dependence may be parallel events in H⁺-PPase evolution. However, the loss of K⁺ sensitivity in family II is not a direct consequence of Cys²²² acquisition, because a reverse evolutionary substitution (C222A) did not confer K⁺ sensitivity to R-PPase. Strong conservation of Cys²²² in subfamily II and Cys⁵⁷³ in subfamily I implies important roles for these residues that are yet to be determined. The dispensability of these cysteine residues for catalysis and proton translocation established in the present work, in concurrence with earlier reports (29), rules out their direct participation in catalysis. A regulatory role involving Cys modification by formation of a disulfide with a protein factor or a low molecular mass thiol (for instance, glutathione) is thus the most likely theory.

	ACKNOWLEDGEMENTS

We thank Dr. J. E. Walker (Cambridge) for providing the E. coli C43(DE3) strain, Dr. M. Baltscheffsky and A. Schultz for providing R-PPase gene and antibody against the protein and helpful discussions, and A. B. Zyryanov for help with PP_i synthesis measurements.

	FOOTNOTES

^* This work was supported by Academy of Finland Grants 35736 and 47513 and Russian Foundation for Basic Research Grants 00-04-48310, 00-15-97907, and 01-04-06114.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.

^¶ To whom correspondence may be addressed. Tel.: 7-095-939-5541; Fax: 7-095-939-3181; E-mail: baykov@genebee.msu.su.

To whom correspondence may be addressed. Tel.: 358-2-333-6845; Fax: 358-2-333-6860; E-mail: reijo.lahti@utu.fi.

Published, JBC Papers in Press, April 15, 2002, DOI 10.1074/jbc.M202951200

² G. A. Belogurov and M. V. Turkina, unpublished observations.

	ABBREVIATIONS

The abbreviations used are: H⁺-PPase, proton translocating inorganic pyrophosphatase; AMDP, aminomethylenediphosphonate; IMV, inner membrane vesicles; R-PPase, R. rubrum H⁺-PPase; RPP-IMV, inner membrane vesicles containing R-PPase; pET-IMV, inner membrane vesicles lacking R-PPase; contig, group of overlapping clones.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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