TIBS 17 - SEPTEMBER 1992
REVIEWS MEMBRANE-BOUND H+-translocating adenosine triphosphatases (H'-ATPases) are the primary transducers by which living cells interconvert light, chemical and electrical energy 1. H*-ATPases establish and maintain electrochemical gradients across membranes for the H~coupled transport of other solutes, or in the special case of the energy-coupling membranes of mitochondria, chloroplasts and bacteria, transduce the H÷electrochemical gradients generated by membrane-linked, vectorial redox reactions to the synthesis of ATP2. Against this background, it is therefore surprising to find that the vacuolar membrane (tonoplast) of plant cells contains an inorganic pyrophosphate (diphosphate, PPL)-energized H+-pump (H÷-PPase; EC 3.6.1.1) as well as a more conventional H+-ATPase (EC 3.6.1.3) 3,4. Both enzymes catalyse electrogenic H+translocation from the cytosol to vacuole lumen, but the H+-PPase has the unusual characteristic of exclusively using PP~ as energy source 5. The importance of the H+-PPase and H+-ATPase for plant cell function derives from the shear bulk of the vacuole and the fact that most of its functions ultimately rely on energy-dependent H*-translocation across the vacuolar membraneE In terms of size, the vacuole is the dominant organelle of most plant cells, constituting 90-99% of the total intraceltular volume of a typical mature cell, and as such is the major storage compartment for nutrients, metabolites, storage proteins and hydrolases. Most, if not all, of these storage functions require sustained uphill transport of H+ from the cytosol to the interior of the vacuole. A multitude of H*-coupled uniporters, symporters and antiporters mediate the vacuolar accumulation of solutes (Fig. P. A. Rea, Y. Kim and V. Sarafian are at the Plant Science Institute, Department of Biology, Universityof Pennsylvania, Philadelphia, PA 19104, USA.R. J. Poole is at the Department of Biology, McGil~ University, 1205 Dr Penfield Avenue, Montreal, Quebec, Canada H3A 1B1. J. M. Davies and D. Sanders are at the Department of Biology, Universityof York, Heslington, York, UK Y01 5DD.
348
Vacuolar H+-translocating pyrophosphatases: a new category of ion translocase
The membrane surrounding the central vacuole of plant cells contains an H+-translocating ATPase (H+-ATPase) and an H+-translocating inorganic pyrophosphatase (H+-PPase). Both enzymes are abundant and ubiquitous in plants but the H+-PPase is unusual in its exclusive use of inorganic pyrophosphate (PPi) as an energy source. The lack of sequence identity between the vacuolar H+-PPase and any other characterized ion pump implies a different evolutionary origin for this translocase. The existence of the vacuolar H+-PPase, in conjunction with increasing recognition of PP~ as a key metabolite in plant systems, necessitates reconsideration of ATP as the primary energy source for membrane transport in plant cells.
1), while the localization of storage proteins and ]ysosomal-type hydrolases is dependent on the maintenance of an inside-acid transtonoplast pH difference (ApH). Any meaningful account of vacuolar function is therefore contingent on an understanding of the functional characteristics and origins of the primary energizers, the H*-PPase and H÷-ATPase. In this review we discuss recent advances in vacuolar H+-PPase research and show that its existence not only signifies a new category of ion translocase but also points to a broader role for PPi in the energy metabolism of plantsl
Species distribution of vacuolar H÷-PPase The vacuolar H+-PPase is widespread in plants. The enzyme has been demonstrated in vacuolar membrane fractions from all of the major vascular plant types (monocotyledons, dicotyledons, C3, C4 and CAM plants) 3, including their likely ancestors, the charophyte algae7 and the unicellular marine alga Acetabularia acetabulum 8. However, the situation in non-plant cells is not so clear. Although Lichko and Okorokov~ have reported PPi-driven H÷-trans!ocation in vacuolar membrane-enriched fractions from Saccharomyces carlsber-
genesis, attempts to identity an H+-PPase homologous to the plant enzyme either by immunological cross-reactivity with S. cerevisiae vacuolar membranes or by genomic analyses of S. cerevisiae, S. carlsbergenesis and Schizosaccharomyces pombe - have yielded negative results (R. Eisman, E. J. Kim and P. A. Rea, unpublished). Similarly, in chromaffin granule ghosts isolated from adrenal medulla, which are well documented for the presence of a H+-ATPase immunoto~ically cross-reactive with and homologous to the plant vacuolar H+-ATP-ase4, PP,-dependent H~-translo cation is not detectable (D. K. Apps and P. A. Rea, unpublished). Thus, pending data to the contrary, the vacuolar H+PPase is assumed to be restricted to plants and their antecedents.
Activity, abundance and polypeptide composition . The H÷-PPase is a major component of the vacuolar membranes of plant cells and can generate an H+-gradient of similar (if not greater) magnitude than the H+-ATPase on the same membrane 3,1°,n. The specific activity of the purified enzyme (12-20~molmg -~ min-l) ~°-12is consistent with a turnover © 1992,ElsevierSciencePublishers,(UK)
TIBS 17 -
SEPTEMBER1992
number of 34-82 s -1 and abundance estimates suggest that the H÷-PPase constitutes between 1% and 5-10% of total vacuolar membrane protein I°,H. Structural studies indicate that the vacuolar H+-PPase has a comparatively simple subunit composition and consists of a single 70 kDa polypeptide species. This component alone copurifies with PPase activity during detergent-solubilization and chromatography 1°-~3and is the only polypeptide of tonoplast vesicles susceptible to substrate (MgPPL)-protectable covalent modification by the sulfhydryl reagent N-ethylmaleimide (NEM)~1,a2.Accordingly, selective purification of the 70 kDa subunit and its insertion into artificial liposomes reconstitutes both MgPP~ hydrolysis and H+-translocationTM. These findings, together with the sequence data derived from cDNAs encoding this subunit, which show that the polypeptide specified satisfies the minimum structural requirements of an H+-pyrophosphatase (i.e. direct interaction with cytosolic substrate and continuity across the phospholipid bilayer) (Fig. 2), imply that subunits in addition to the 70 kDa polypeptide are not necessary for H+-PPase function.
PP~as anenergysource The ubiquity and abundance of the vacuolar H+-PPase gives rise to the question: is PP~ capable of playing the role of renewable cellular energy source? The free energy of hydrolysis of cytosolic ATP is sufficient to drive ATPase-mediated H÷-translocation, but is the same true of cytosolic PPi and the vacuolar PPase? The answer to this question is probably yes. First, it should be noted that all of the core biosynthetic pathways generate PPi and provide a steady supply of PP~ in biOsynthetically active tissues. Examples of PP,-generating reactions are the acylation of CoA in fatty acid synthesis, the aminoacylation of tRNA in polypeptide synthesis and the formation of phosphodiester bonds in polynucleotide and polysaccharide synthesis. Second, estimates of the free energy of hydrolysis of PP~ in the cytosol of plant cells yield values commensurate with transtonoplast H+ pumping. The textbook account of PP~, first proposed by Kornberg x~, is that it is a nucleotidyl transfer coproduct whose hydrolysis by soluble PPases serves to pull the primary (usually synthetic) reaction to completion:
2~
PPi
\ H+
\
zW= + 2 0 - 5 0 mV
3H +
~/ Na+
H÷
"~
ApH = 1.5--4.5 PHvac= 3 - 6
~ - -
Ca2+
~
~
sugar
.....
~ ....
.,L.
PHcy t = 7.5
Mal2-
NO3 H ÷
Ca 2+
ATP
ADP + Pi
Figure1 Electrogenic H÷-translocation by vacuolar H+oATPaseand H+-PPase. Both enzymes generate an inside-acid, inside-positive H+-electrochemicai potential difference (proton motive force, pmf) in which the chemical (ApH) and electrical potential difference (Ag) are related by the function pmf/F = A~/+ 59 ApH in mV at 250C. ApH is defined as pHcyt - PHvac and F is the Faraday constant. The resulting pmf may drive uniport or antiport. CI-, NOa- and malate (Mal2-) enter and Ca2+ exits in response to the inside-positive A~. Na+, Ca2+ and sugars enter in exchange for H+.
NTP + ROH -~ RONMP + PPi
AGO'= +ve
Pei ÷ H 2 0 ~ g e i
AGO'=-ve
(1) (2)
NTP +ROH + H20 -~ RONMP + 2Pi
AGO' = -ve
(3)
where AGo' is Gibbs free energy change at pH 7.0. In short, PPi hydrolysis should be accompanied by a significant release of free energy if biosynthesis is to proceed unimpeded. Although originally intended to explain the facilitatory action of soluble PPases on biosynthesis, it is now clear from measurements of cytosolic PP~ in plants that the Kornberg scheme is also consistent with (if not dependent on) the operation of the vacuolar H+-PPase. The most reliable estimates of cytosolic PP~ levels in plants fall in the range 0.2 to 0.3 mM16'17.Based on a cytoplasmic P~ concentration of 5 mMTM and an equilibrium Constant for PP~ hydrolysis of 2320 M~9, an overall free energy yield of 25 kJ mol-~ under physi01ogical
conditions would be predicted. This is more than sufficient to drive PPc dependent H+-translocation across the vacuolar membrane in v i v o 3. The potential d e p e n d e n c e of the Kornberg scheme on the vacuolar H+-PPase, on the other hand, is posed by the finding that in photosynthetic tissues soluble PPase activity is largely absent from the cytosol and almost exclusively located in the chloroplast stroma TM. Thus, two important conclusions may follow: (1) the vacuolar H*-PPase might be the sole enzyme responsible for the disposal of cytosolically produced PP~ in many plant tissues; and (2) it is not imperative that the energy liberated during PP~ hydrolysis be simply lost as heat, as is the case for soluble PPases the same driving force for biosyntheses would be provided by a H+-PPase which conserves a portion of this energy in the form of a transmembrane H* gradient.
349
TIBS 17 - SEPTEMBER 1 9 9 2
Vacuole
m m m
34
14
[, ivl:
IZl '~143 " [
r26
VI
485 ~ 563
v, [Viii i,× iv,,
Ii×,,
7"4\':17 ;"
216 "T 328 '~'369
×,,,
'4'
+++
Cytoplasm
Figure 2 Tentative topographic model of vacuolar H÷-PPase from computer-assisted hydropathy plots of the deduced amino acid sequence of cDNAs encoding the substrate-binding poiypeptide of the H+-PPase isolated from Arabidopsis. Transmembrane spans I-XIII, all of which appear t o be multimeric on the basis of their hydrophobic moment, were predicted from the HELIXMEM program of PC-GENE. The structure of the putative non-transmembranous regions was examined using the secondary structure predictions of Gamier eta/. 36 using both the GARNIER and GGBSM programs of PC-GENE. WW, c~-helix; , random coil; - - - , clusters of negative charge; + + + , clusters of positive charge; N, amino terminus; C, carboxyl terminus. The cytosolically oriented, non-transmembranous regions shown in bold contain the putative catalytic motifs depicted in Fig. 3.
Sequence and tentative structure A major advance in our understanding of the structure and origins of the vacuolar H÷-PPase has come from the recent isolation and sequencing of cDNAs encoding the enzyme from Arabidopsis thaliana 2° and Beta vulgaris (Y. Kim, E. J. Kim and P. A. Rea, unpublished). Partial cDNA clones were initially selected by probing expression libraries with antibody raised against the polypeptide purified from Vigna radiata vacuolar membranes TM and full-length clones were isolated by subsequent hybridization screens using the partial clones as probes. The identity of the clones was confirmed by alignment with direct internal sequence data acquired from the MgPPrbinding polypeptide of the enzyme purified from B. vulgaris ]5 and direct amino-terminal sequence data for the enzyme from E radiata I°. Both clones [designated AVP and BVP for Arabidopsis and Beta VaGuoiar (HL translocating) Pyrophosphatase~ respectively] encode a 770-amino acid polypeptide with a predicted mass of 81 kDa. [The discrepancy between apparent relative mass estimated from electrophoretic mobility (70 kDa) and computed
350
molecular weight (81 kDa) is a common feature of membrane proteins.] Computer-assisted hydropathy plots of the Arabidopsis and Beta vacuolar HL PPase amino acid sequences have established that AVP and BVP encode an extremely hydrophobii: integral membrane protein containing 13-16 amphipathic ('multimeric') transmembrane spans which may serve to generate and stabilize a relatively compact structure through the formation of inter-helical hydrogen bonds and salt bridges (Fig. 2). In addition, several of the putative hydrophilic domains are characterized by the presence of clusters of charged residues which may participate in cation (Mg2÷, Ca2÷, K~) or anion (pp4-, MgPPi2-) binding. Thus, the hydrophilic segment linking membrane spans I and II contains four contiguous glutamate residues (positions 64-67 in the Arabidopsis sequence), the segment linking spans IV and V contains four acidic residues in a stretch of eight amino acids (positions 222-229), the segment between sPans VIII and IX contains the sequence RXRXR (positions 525-529) and the penultimate hydrophilic domain, linking spans XII and XIII, contains
a preponderance of Lys residues. The overall orientation of t h e 81 kDa polypeptide is depicted as in Fig. 2 in accord with the ,positive-inside rule '21 wherein, for the majority of multiply spanning membrane proteins, positively charged amino acids are located predominantly towards the cytoplasmic surface of the membrane. Unique origins. The novelty of the vacuolar H÷-PPase has been confirmed by database searches of the sequence. All previously characterized phosphoanhydride-energized H÷pumps fall into three major categories: F, P and V, based on subunit organization, mechanism and inhibitor sensitivity22. They are subject to inhibition by azide, orthovanadate and bafilomycin, respectively, and the enzymes within each category show functionally significant sequence identities, indicating a common ancestry 4,22. By contrast, the vacuolar H÷-PPase is not inhibited by any of these typespecific inhibitors and computer searches of the nucleotide sequences of AVP and BVP and the deduced sequences of the protein products reveal no detectable homology between this pump and any other sequenced ion
TIBS 17 - SEPTEMBER 1 9 9 2
translocase 2°. The vacuolar H+-PPase is therefore likely to be a member of a Class of previously undescribed ion translocase. Close phylogenic links between the vacuolar H+-PPase and the soluble, nonenergy-conserving PPases are similarly unlikely: (1) all characterized soluble PPases have different subunit sizes from the vacuolar H+-PPase (20 kDa for the prokaryotic enzymes, 32-42 kDa for the eukaryotic enzymes)23; and (2) none of the known sequences for soluble PPases (from Arabidopsis, E. coli, Kluyveromyces and Saccharomyces) 23 align with the deduced sequence of the vacuolar H+-PPase. Of the two candidate homologs remaining - the energy-conserving H+PPases of mitochondria and those of phototrophic bacteria - the former are easily eliminated at both the biochemical and molecular levels. (1) Their structural characteristics are distinct from those of the vacuolar H+-PPase; mitochondrial H+-PPases are F-like in organization and consist of a peripheral 28-30 kDa catalytic subunit associated with the membrane through non-covalent interactions with a transmembranous sector 24. (2) Rat-liver mitochondrial H+-PPase does not cross-react with antibody raised against the vacuolar H+-PPase25. (3) Genomic clones of the 28-30 kDa subunit of the mitochondrial H+-PPase from Saccharomyces cerevisiae show 49% sequence identity to the soluble PPase from the same source 26, but no identity with the vacuolar H+-PPase2° (Y. Kim, E. J. Kim and P, A. Rea, unpublished). Evaluation of a more promising evolutionary connection between the vacuolar H+,PPase and the reversible H+translocating PPase (H+-PPi synthase) of phototrophic bacteria will require sequence data for the bacterial enzyme. The presence of a H+-PP~synthase on the energy-coupling membranes of phototrophic bacteria, notably the purple non-sulfur bacterium Rhodospirillum rubrum, has been known for some time 27, but it has only recently been shown that this translocase is an integral protein of approximately 56 kDa28. Two features of this polypeptide are significant: (1) it is immunologically cross-reactive with the MgPPcbinding subunit of the vacuolar H+-PPase (E. J. Kim, Y. Kim and P. A. Rea, unpublished) and, (2) unlike the 28-30 kDa peripheral, catalytic subunit of the mitochondrial H+-PPase, it is capable of mediating both MgPPi hydrolysis and H+-translocation 28. The likelihood of structural
AVP
Ath
119-1~.GF STDNKP C TYD
PPA-1
See
14 6-DzGzTDwKvIATD DZGzTDWKVIAID DZG~.TDwKvIvID D Q G E K D D K I IAVC
Kla
Spo A th •PPA-2
Sce
CONSENSUS
DDG~.LDWKVIvID
(D/E) xxxxDxKxxxxD
2 57-DVGADLVGKIE 4 8 -~..I P R W T N A K L I~Ira.I PRWTNAKLI~~. I P R W T Q A K L I~" ~..APT V F N C K V ~. ~..VPRWT T G K F ~.
(D/E) xxxxxxxKxE
Figure 3 Alignment of putative catalytic residues of vacuolar H÷-PPase (AVP), soluble PPases (PPA-1) and mitochondrial H÷-PPase (PPA-2). Active-site residues implicated in catalysis are indicated in bold 23. Residues of like charge that are common to all six sequences are shown in large letters. Ath, Arabidopsis thaliana; Sce, Saccharomyces cerevisiae; Kla, Kluyveromyces lactis; Spo, Schizosaccharomyces pombe; the PPA-1 and PPA-2 residue numbering is that of the mature soluble PPase protein from S. cerevisiae.
and functional homologies between the H+-PPases from Rhodospirillum, and possibly other phototrophic bacteria, and plant vacuolar membranes is therefore realistic and exciting. Genomic screens of Rhodospirillum using Arabidopsis H+-PPase cDNAs and their derivatives as probes are currently in progress in one of our laboratories (Y. Kim and P. A. Rea, unpublished). These studies will examine this proposal and explore the possibility that the vacuolar H+-PPase is as ancient as the progenores from which the non-sulfur, photosynthetic bacteria were derived. Putative catalytic site(s). Lack of extensive sequence homologies between the vacuolar H+-PPase and other characterized PPases does not automatically preclude functional analogies between the two classes of enzyme. A case in point is the soluble PPases themselves. Although gross sequence comparisons between soluble PPases from a wide range of organisms reveal only modest sequence identities of between 20 and 27%, residues implicated in catalysis show pronounced conservation 23. X-ray crystallographic and site-directed mutagenesis studies of the soluble PPase f r o m Saccharomyces have disclosed 17 residues that are thought to participate directly in catalysis, 11-16 of which (depending on alignment procedure) are conserved in all sequenced, soluble PPases. It is therefore of potential functional interest that eight of the activesite residues of soluble PPases fall into two configurations, EXTKXE and DX2EXDXKX4D (where X = any amino acid), beginning at positions 48 and 146,
respectively, in the sequence of the
Saccharomyces enzyme, while variants of both motifs (DXTKXEand DX4DXKX4D) are found at positions 257 and 119, respectively, in the deduced amino acid sequence of the vacuolar H+-PPase (Fig. 3). Thus, while the vacuolar H~-PPase and soluble PPases appear to be evolutionarily remote, they may share convergent motifs related to the need for both classes of enzyme to interact with the same substrates and cofactors (Mg2~, MgPPi and/or Mg2PP~). Both putative catalytic motifs have a cytosolic orientation (if the configuration of the vacuolar H+-PPase in Fig. 2 is correct) and the motif that starts at position 119 contains a Cys residue and is immediately flanked by another (Fig. 3). It is therefore tempting to speculate that the pronounced sensitivity of the vacuolar H+-PPase to inhibition and covalent modification by N-ethylmaleimide (NEM) and protection by substrate at the cytosolic face of the membrane u'12 is due to alkylation of one or both of these Cys residues.
Physiological function It is now established that the vacuolar H+-PPase and vacuolar H+-ATPase reside on the same membrane and pump H÷ into the same compartment (Fig. 1). The steady-state ApH and A~ generated by the PPase and ATPase of tonoplast vesicles are non-additive and subject to kinetic control by a common H÷ gradient 3, and intact vacuoles of ethyleneglycoltetraacetic acid (EGTA)-permeabilized cells of Chara7 and single, isolated patch clamped vacuoles of
351
TIBS 17 -
B. vulgaris 29 mediate both PPr and ATP-
dependent H+-translocation. Why are there two transport systems pumping the same ion into the same intracetlular compartment? This key question remains unanswered, but there are two possible explanations for the existence of the vacuolar H÷-PPase. Scavenging system, T h e v a c u o l a r H +PPase may simply scavenge the free energy of the PP~ formed in the pyrophosphorylytic reactions of polymer synthesis3°. While a soluble PPase would merely thermally dissipate the free energy of PP~ hydrolysis, a biologically useful output could be retrieved if some of this energy was conserved as a transmembrane H* gradient; this could then be used for subsequent energy-dependent solute transport. A remarkable characteristic of cellular PP~ levels in plants is their invariance. PP~ levels do not change detectably during light-dark transitions, during rapid changes in the rate of respiration or when tissues are subjected to anoxia or respiratory poisons ~6. Cellular ATP levels, on the other hand; change dramatically under these conditions. Coordinate stabilization of PP~ levels and operation of the vacuolar H*-PPase would provide a back-up system for the maintenance of vacuolar compartmentation despite (temporary) metabolic perturbation. Retention of a functional vacuolar H*PPase in plants may be explicable in terms of the greater range of environmental conditions experienced by plants versus mammalian cells. It has been suggested that H* pumps originally functioned as a pH-stat, disposing of the excess acidity generated by fermentative metabolism31. Conservation of this function and the sustained operation of the H*-PPase under conditions of low phosphate (ATP synthetic), which accompany anaerobiosis, would serve two purposes: elimination from the cytoplasm of the additional H+ produced fermentatively and maintenance of intracellular compartmentation. Mediating H÷ and K÷ translocation is a p o s s i b l e role for the vacuolar H~-PPase. Substrate hydrolysis and H÷ translocation by the vacuolar H*-PPase exhibit an almost absolute requirement for K÷ (Ref. 30) and patch-clamp studies on intact vacuoles show that ion translocation by the H+-PPase is specifically dependent on the presence of K* at the cytosolic, but not vacuolar, face of the membrane32. Since the reversibility of the vacuolar H÷-PPase is critically dependent on the transtonoplast K÷
352
gradient, it has been suggested that rather than merely acting as a supplementary H+ pump, the vacuolar H*PPase may serve to catalyse the transtonoplast translocation of both H* and K* (Ref. 32). Note that while H+coupled transport pathways are known for Na* and Ca2* at the vacuolar membrane, there is no information on the mechanism of K÷transport, despite the importance of this ion as a macronutrient and turgor regulant and the necessity for its energy-dependent transport into the vacuole against the prevailing inside~positive membrane potential.
SEPTEMBER 1992
through analysis of the physiological consequences of altered pump expression in transgenic plants that either express antisense transcripts or ectopically overexpress the H*-PPase. Acknowledgements
This work was supported by the National Science Foundation, the Department of Energy, the University Research Foundation, University of Pennsylvania, the Biological Sciences Research Group, University of Pennsylvania (awarded to P. A. R.) and the Agriculture and Food Research Council UK (awarded to D. S.). This article is Concluding remarks dedicated to the memory of Peter T h e ~abundance and ubiquity of the Mitchell, without whose revolutionary vacuolar H+-PPase in plant cells extends insights and single-minded devotion to conventional notions of the role of PP~ the problem of biological energy transin contemporary biological energy trans- duction none of this would have been duction. The novelty of the vacuolar H÷- possible. PPase and its apparently ancient origins, on the other hand, raise important References 1 Mitchell, P. (1961) Chemiosmotic Coupling in questions concerning the place of PP~ in Oxidative and Photosynthetic Phosphorylation, the evolution of cellular energy metabGlynn Research olism. The results of laboratory exper2 Harold, F. M. (1986) The Vital Force: A Study of Bioenergetics, Freeman iments designed to simulate primitive earth conditions suggest that the for- 3 Rea, P. A. and Sanders, D. (1987) Physiol. Plant 71, 131-141 mation of PP~ and other polyphosphates 4 Nelson, N. and Taiz, L. (1989) Trends Biochem. may have been pivotal for prebiotic Sci. 14, 113-116 5 Rea, P. A. and Poole, R. J. (1986) Plant Physiol. evolution: these molecules will catalyse 81, 126-129 the phosphorylation of adenosine, AMP 6 Raven, J. A. (1987) New Phytol. 106, and ADP and the synthesis of polypep357-422 tides 33. Moreover, since the synthesis of 7 Shimmen, T. and MacRobbie, E. A. C. (1987) Protoplasma 136, 205-207 PP~ from precipitated orthophosphate 8 Ikeda, M. et al. (1991) Biochim. Biophys. Acta is practicable at low temperatures 34and 1070, 77-82 these conditions approximate those for 9 Lichko, L. and Okorokov, L. (1991) Yeast 7, 805-812 template-directed oligomerization of nucleotides35, it is conceivable that the 10 Maeshima, M. and Yoshida, S. (1989) J. Biol. Chem. 264, 20068-20073 generation of PPi and the synthesis of 11 Rea, P. A, Britteni C. J. and Sarafian, V. Plant biopolymers were necessary concomiPhysiol. (in press) tants for the emergence of life. Any sys- 12 Britten, C. J., Turner, J. C. and Rea, P. A. (1989) FEBS Lett. 256, 200-206 tem which could harness the energy 13 Sarafian, V. and Poole, R. J. (1989) Plant contained in PP~ would consequently Physiol. 91, 34-38 have been at a replicative advantage. Is 14 Britten, C. J., Zhen, R., Kim, E. J. and Rea, P. A. J. Biol. Chem. (in press) the energy-conserving vacuolar H*15 Kornberg, A. (1963) in Horizons in Biochemistry PPase a product of life's prebiotic ori(Kasha, M. and Pullman, B., eds), gins before the emergence of the more pp. 251-264, Academic Press specific and recognizable structures of 16 Weiner, H., Stitt, M. and Heldt, W. (1987) Biochim. Biophys. Acta 893, 13-21 ATP (and other nucleotides) for nucleic 17 Takeshige, K. and Tazawa, M. (1989)J. Biol. acid synthesis, adenylation, pyrophosChem. 264, 3262-3266 phorylation and signal transduction? 18 Rebeille, F., Bligny, R. and Douce, R. (1984) Plant Physiol. 74, 355-359 From the standpoint of contemGuynn, R. W., Veloso, D., Lawson, J. W. R. and porary plant-cell function, the ready 19 Veech, R. L. (1974) Biochem. J. 140, 369-375 availability of cDNAs encoding the 20 Sarafian, V., Kim, Y., Poole, R. J. and Rea, P. A. (1992) Proc: Natl Acad. Sci. USA 89, vacuolar H*-PPase should allow the full 1775-1779 physiological significance of the en21 von Heijne, G. and Gavel, Y. (1988) Eur. J. zyme in vivo to be uncovered. The role Biochem. 174, 671-678 of the vacuolar H*-PPase in cellular 22 Pedersen, P. L. and Carafoli, E. (1987) Trends Biochem. Sci. 12, 146-150 energy conservation, turgor regulation, resistance to anaerobiosis and vacuolar 23 Cooperman, B. S., Lahti, R. and Baykov, A. A. Trends Biochem. ScL (in press) solute compartmentation in the intact 24 Mansurova, S. E. (1989) Biochim. Biophys. Acta plant is now amenable to investigation 977,237-247
TIBS 17 - SEPTEMBER 1992
.....
25 Maeshima, M. (1991) Eur. J. Biochem.
196, 11-17 26 Lundin, M., Baltscheffsky, H. and Ronn~i H. (1991) J. Biol. Chem. 266, 12168-121-72 27 Baltsheffsky, M. and Nyren, P. (1987) irl'/
29 Kurkdijan, A. and Hedrich, R. (1988) EMBO J. 7, 30 31
Phosphate Metabolism and Cellular RegUlation in Microorganisms (Torriani-Gorini,A. et al., eds), pp. 260-263, American So(~ietyfor
32
Microbiology
33
28 Nyren, P., Note, B. and Strid, A. (1991) Biochemistry 30, 2883-2887 i'. i'
THE EXISTENCE OF an enzymatic activity responsible for the termination of acetylcholine (ACh)-mediated neurbtransmission at cholinergic synapseg was postulated as early as 1914 by Dale (see Refs 1-3). Both of the AChdegrading enzymes, acetylcholinesterase (acetylcholine acetyl hydrolase, ACHE; EC 3.1.1.7) and butyrylcholinesterase (acylcholine acyl hydrolase, BCHE; EC 3.1.1.8), are evolutionarily conserved type B carboxylesterases that share extensive amino acid homology and display similar catalytic properties. There are, however, some important exceptions: ACHE specifically interacts with ACh and is subject to marked substrate inhibition, whereas BCHE is capable of degrading a wider range of choline esters and shows no evidence of substrate inhibition ~-3. Furthermore, each enzyme displays characteristic affinities to various selective inhibitors ~,2. Both enzymes appear as monomers, dimers and tetramers, as well as heavier forms bound to structural subunitsS; these are thought to participate in regulating the extent and duration of ACh interaetions with its cellular receptors. The ability of the subunits to interact with various inhibitors means that ACHE and BCHE are sensitive targets for natural and synthetic cholinergic toxins, including toxic glyco-alkaloids, highly poisonous organophosphorous (OP) and carbamate insecticides, snake venom peptides and diverse synthetic therapeutic agents ~-4. Cholinesterase (CHE) activities have been extensively studied by multidiSH. Soreq, A. Gnatt and Y. Loewenstein are at
the Departmentof Biological Chemistry, The Life Sciences Institute, The HebrewUniversity of Jerusalem, Israel 91904. L. F. Neville is at The David MayoneyInstitute of Neurological Sciences, University of Pennsylvania; PA 19104-6074, USA. :ii.... © 1992, Elsevier Science Publishers, (UK)
34
3661-3666 Rea, P. A. and Poole, R. J. (1985) Plant Physiol. 77, 46-52 Raven, J. A. and Smith, F. A. (1976) J. Theor. Biol. 57,301-302 Davies, J. M., Rea, P. A. and Sanders, D. (1991) FEBS Lett. 278, 66-68 Yamanaka, J., Inomata, K. and Yamagata, Y. (1988) Origins Life 18, 165-178 Hermes-Lima,M. (1990) J. Mol. Evol. 31, 353-358
35Acevedo, O. L. and Orgel, L. E. (1986) Nature 321, 790-792 36 Gamier, J., Osguthorpe, D. J. and Robson, B. (1978) J. Mol. Biol. 120, 97-120
See also the review by Barry S. Cooperman, 'Evolutionary conservation of soluble inorganic pyrophosphatases active site structure and mechanism' in the July issue of TIBS.
Excavations into the active-site gorge of cholinesterases
Acetyl- and butyrylcholinesterase (ACHE, BCHE) from evolutionarily distant species display a high degree of primary sequence homology and have biochemically similar catalytic properties, yet they differ in substrate specificity and affinity for various inhibitors. The biochemical information derived from analyses of ACHE and BCHE from human, Torpedo, mouse, and Drosophila, as well as that from the recombinant forms of their natural variants and site=directed mutants, can currently be re-examined in view of the recent X-ray crystallography data revealing the three-dimensional structure of Torpedo ACHE. The picture that emerges deepens the insight into the biochemical basis for choline ester catalysis and the complex mechanism of interaction between cholinesterases and their numerous ligands.
ciplinary approaches, which began with detailed biochemical studies of CHEs from several species. This resulted in intricate theories explaining the catalytic properties and inhibitor interactions that are characteristic of various CHEs1-5, More recently, the primary amino acid sequence of CHEs from evolutionarily diverse species was uncovered by gene cloning (for review see Refs 2, 5) and heterologous expression systems have been developed for recombinant CHEs from man 6-8, mouse ~, Torpedo(electric ray) 5,9 and Drosophila ~°, All of these approaches, in addition to affinity labeling H and monoclonal antibody studies ~2, suggested the involvement of several separate amino acid domains in the active center of CHEs, but could n o t identify these domains unequivocally. Recently, the three-dimensional analysis of Torpedo ACHE~3 has deepened, clarified and altered our understanding of CHEs considerably, providing structural infor-
mation on the positioning of catalytically important amino acid residues within these proteins ~3.In an attempt to integrate the information from these various disciplines, we superimposed highly homologous CHEs and their mutants into a single molecular model. The Catalytic- and ligand-binding properties of this composite CHE were analysed in relation to the threedimensional locations of the altered residues with a view to delineating CHE structure-function relationships.
Catalysis by cholinesterases Biochemical analyses demonstrated that catalysis by CHEs involves an active Ser residue (for review see Refs 1-5) and predicted that CHEs, like serine proteases (i.e. trypsin) hydrolyse choline esters through electron transfer within a catalytic triad. In the case of cholinesterases, a negatively charged residue draws a hydrogen atom from an adjacent His which, in turn, draws a
353
REVIEWS MEMBRANE-BOUND H+-translocating adenosine triphosphatases (H'-ATPases) are the primary transducers by which living cells interconvert light, chemical and electrical energy 1. H*-ATPases establish and maintain electrochemical gradients across membranes for the H~coupled transport of other solutes, or in the special case of the energy-coupling membranes of mitochondria, chloroplasts and bacteria, transduce the H÷electrochemical gradients generated by membrane-linked, vectorial redox reactions to the synthesis of ATP2. Against this background, it is therefore surprising to find that the vacuolar membrane (tonoplast) of plant cells contains an inorganic pyrophosphate (diphosphate, PPL)-energized H+-pump (H÷-PPase; EC 3.6.1.1) as well as a more conventional H+-ATPase (EC 3.6.1.3) 3,4. Both enzymes catalyse electrogenic H+translocation from the cytosol to vacuole lumen, but the H+-PPase has the unusual characteristic of exclusively using PP~ as energy source 5. The importance of the H+-PPase and H+-ATPase for plant cell function derives from the shear bulk of the vacuole and the fact that most of its functions ultimately rely on energy-dependent H*-translocation across the vacuolar membraneE In terms of size, the vacuole is the dominant organelle of most plant cells, constituting 90-99% of the total intraceltular volume of a typical mature cell, and as such is the major storage compartment for nutrients, metabolites, storage proteins and hydrolases. Most, if not all, of these storage functions require sustained uphill transport of H+ from the cytosol to the interior of the vacuole. A multitude of H*-coupled uniporters, symporters and antiporters mediate the vacuolar accumulation of solutes (Fig. P. A. Rea, Y. Kim and V. Sarafian are at the Plant Science Institute, Department of Biology, Universityof Pennsylvania, Philadelphia, PA 19104, USA.R. J. Poole is at the Department of Biology, McGil~ University, 1205 Dr Penfield Avenue, Montreal, Quebec, Canada H3A 1B1. J. M. Davies and D. Sanders are at the Department of Biology, Universityof York, Heslington, York, UK Y01 5DD.
348
Vacuolar H+-translocating pyrophosphatases: a new category of ion translocase
The membrane surrounding the central vacuole of plant cells contains an H+-translocating ATPase (H+-ATPase) and an H+-translocating inorganic pyrophosphatase (H+-PPase). Both enzymes are abundant and ubiquitous in plants but the H+-PPase is unusual in its exclusive use of inorganic pyrophosphate (PPi) as an energy source. The lack of sequence identity between the vacuolar H+-PPase and any other characterized ion pump implies a different evolutionary origin for this translocase. The existence of the vacuolar H+-PPase, in conjunction with increasing recognition of PP~ as a key metabolite in plant systems, necessitates reconsideration of ATP as the primary energy source for membrane transport in plant cells.
1), while the localization of storage proteins and ]ysosomal-type hydrolases is dependent on the maintenance of an inside-acid transtonoplast pH difference (ApH). Any meaningful account of vacuolar function is therefore contingent on an understanding of the functional characteristics and origins of the primary energizers, the H*-PPase and H÷-ATPase. In this review we discuss recent advances in vacuolar H+-PPase research and show that its existence not only signifies a new category of ion translocase but also points to a broader role for PPi in the energy metabolism of plantsl
Species distribution of vacuolar H÷-PPase The vacuolar H+-PPase is widespread in plants. The enzyme has been demonstrated in vacuolar membrane fractions from all of the major vascular plant types (monocotyledons, dicotyledons, C3, C4 and CAM plants) 3, including their likely ancestors, the charophyte algae7 and the unicellular marine alga Acetabularia acetabulum 8. However, the situation in non-plant cells is not so clear. Although Lichko and Okorokov~ have reported PPi-driven H÷-trans!ocation in vacuolar membrane-enriched fractions from Saccharomyces carlsber-
genesis, attempts to identity an H+-PPase homologous to the plant enzyme either by immunological cross-reactivity with S. cerevisiae vacuolar membranes or by genomic analyses of S. cerevisiae, S. carlsbergenesis and Schizosaccharomyces pombe - have yielded negative results (R. Eisman, E. J. Kim and P. A. Rea, unpublished). Similarly, in chromaffin granule ghosts isolated from adrenal medulla, which are well documented for the presence of a H+-ATPase immunoto~ically cross-reactive with and homologous to the plant vacuolar H+-ATP-ase4, PP,-dependent H~-translo cation is not detectable (D. K. Apps and P. A. Rea, unpublished). Thus, pending data to the contrary, the vacuolar H+PPase is assumed to be restricted to plants and their antecedents.
Activity, abundance and polypeptide composition . The H÷-PPase is a major component of the vacuolar membranes of plant cells and can generate an H+-gradient of similar (if not greater) magnitude than the H+-ATPase on the same membrane 3,1°,n. The specific activity of the purified enzyme (12-20~molmg -~ min-l) ~°-12is consistent with a turnover © 1992,ElsevierSciencePublishers,(UK)
TIBS 17 -
SEPTEMBER1992
number of 34-82 s -1 and abundance estimates suggest that the H÷-PPase constitutes between 1% and 5-10% of total vacuolar membrane protein I°,H. Structural studies indicate that the vacuolar H+-PPase has a comparatively simple subunit composition and consists of a single 70 kDa polypeptide species. This component alone copurifies with PPase activity during detergent-solubilization and chromatography 1°-~3and is the only polypeptide of tonoplast vesicles susceptible to substrate (MgPPL)-protectable covalent modification by the sulfhydryl reagent N-ethylmaleimide (NEM)~1,a2.Accordingly, selective purification of the 70 kDa subunit and its insertion into artificial liposomes reconstitutes both MgPP~ hydrolysis and H+-translocationTM. These findings, together with the sequence data derived from cDNAs encoding this subunit, which show that the polypeptide specified satisfies the minimum structural requirements of an H+-pyrophosphatase (i.e. direct interaction with cytosolic substrate and continuity across the phospholipid bilayer) (Fig. 2), imply that subunits in addition to the 70 kDa polypeptide are not necessary for H+-PPase function.
PP~as anenergysource The ubiquity and abundance of the vacuolar H+-PPase gives rise to the question: is PP~ capable of playing the role of renewable cellular energy source? The free energy of hydrolysis of cytosolic ATP is sufficient to drive ATPase-mediated H÷-translocation, but is the same true of cytosolic PPi and the vacuolar PPase? The answer to this question is probably yes. First, it should be noted that all of the core biosynthetic pathways generate PPi and provide a steady supply of PP~ in biOsynthetically active tissues. Examples of PP,-generating reactions are the acylation of CoA in fatty acid synthesis, the aminoacylation of tRNA in polypeptide synthesis and the formation of phosphodiester bonds in polynucleotide and polysaccharide synthesis. Second, estimates of the free energy of hydrolysis of PP~ in the cytosol of plant cells yield values commensurate with transtonoplast H+ pumping. The textbook account of PP~, first proposed by Kornberg x~, is that it is a nucleotidyl transfer coproduct whose hydrolysis by soluble PPases serves to pull the primary (usually synthetic) reaction to completion:
2~
PPi
\ H+
\
zW= + 2 0 - 5 0 mV
3H +
~/ Na+
H÷
"~
ApH = 1.5--4.5 PHvac= 3 - 6
~ - -
Ca2+
~
~
sugar
.....
~ ....
.,L.
PHcy t = 7.5
Mal2-
NO3 H ÷
Ca 2+
ATP
ADP + Pi
Figure1 Electrogenic H÷-translocation by vacuolar H+oATPaseand H+-PPase. Both enzymes generate an inside-acid, inside-positive H+-electrochemicai potential difference (proton motive force, pmf) in which the chemical (ApH) and electrical potential difference (Ag) are related by the function pmf/F = A~/+ 59 ApH in mV at 250C. ApH is defined as pHcyt - PHvac and F is the Faraday constant. The resulting pmf may drive uniport or antiport. CI-, NOa- and malate (Mal2-) enter and Ca2+ exits in response to the inside-positive A~. Na+, Ca2+ and sugars enter in exchange for H+.
NTP + ROH -~ RONMP + PPi
AGO'= +ve
Pei ÷ H 2 0 ~ g e i
AGO'=-ve
(1) (2)
NTP +ROH + H20 -~ RONMP + 2Pi
AGO' = -ve
(3)
where AGo' is Gibbs free energy change at pH 7.0. In short, PPi hydrolysis should be accompanied by a significant release of free energy if biosynthesis is to proceed unimpeded. Although originally intended to explain the facilitatory action of soluble PPases on biosynthesis, it is now clear from measurements of cytosolic PP~ in plants that the Kornberg scheme is also consistent with (if not dependent on) the operation of the vacuolar H+-PPase. The most reliable estimates of cytosolic PP~ levels in plants fall in the range 0.2 to 0.3 mM16'17.Based on a cytoplasmic P~ concentration of 5 mMTM and an equilibrium Constant for PP~ hydrolysis of 2320 M~9, an overall free energy yield of 25 kJ mol-~ under physi01ogical
conditions would be predicted. This is more than sufficient to drive PPc dependent H+-translocation across the vacuolar membrane in v i v o 3. The potential d e p e n d e n c e of the Kornberg scheme on the vacuolar H+-PPase, on the other hand, is posed by the finding that in photosynthetic tissues soluble PPase activity is largely absent from the cytosol and almost exclusively located in the chloroplast stroma TM. Thus, two important conclusions may follow: (1) the vacuolar H*-PPase might be the sole enzyme responsible for the disposal of cytosolically produced PP~ in many plant tissues; and (2) it is not imperative that the energy liberated during PP~ hydrolysis be simply lost as heat, as is the case for soluble PPases the same driving force for biosyntheses would be provided by a H+-PPase which conserves a portion of this energy in the form of a transmembrane H* gradient.
349
TIBS 17 - SEPTEMBER 1 9 9 2
Vacuole
m m m
34
14
[, ivl:
IZl '~143 " [
r26
VI
485 ~ 563
v, [Viii i,× iv,,
Ii×,,
7"4\':17 ;"
216 "T 328 '~'369
×,,,
'4'
+++
Cytoplasm
Figure 2 Tentative topographic model of vacuolar H÷-PPase from computer-assisted hydropathy plots of the deduced amino acid sequence of cDNAs encoding the substrate-binding poiypeptide of the H+-PPase isolated from Arabidopsis. Transmembrane spans I-XIII, all of which appear t o be multimeric on the basis of their hydrophobic moment, were predicted from the HELIXMEM program of PC-GENE. The structure of the putative non-transmembranous regions was examined using the secondary structure predictions of Gamier eta/. 36 using both the GARNIER and GGBSM programs of PC-GENE. WW, c~-helix; , random coil; - - - , clusters of negative charge; + + + , clusters of positive charge; N, amino terminus; C, carboxyl terminus. The cytosolically oriented, non-transmembranous regions shown in bold contain the putative catalytic motifs depicted in Fig. 3.
Sequence and tentative structure A major advance in our understanding of the structure and origins of the vacuolar H÷-PPase has come from the recent isolation and sequencing of cDNAs encoding the enzyme from Arabidopsis thaliana 2° and Beta vulgaris (Y. Kim, E. J. Kim and P. A. Rea, unpublished). Partial cDNA clones were initially selected by probing expression libraries with antibody raised against the polypeptide purified from Vigna radiata vacuolar membranes TM and full-length clones were isolated by subsequent hybridization screens using the partial clones as probes. The identity of the clones was confirmed by alignment with direct internal sequence data acquired from the MgPPrbinding polypeptide of the enzyme purified from B. vulgaris ]5 and direct amino-terminal sequence data for the enzyme from E radiata I°. Both clones [designated AVP and BVP for Arabidopsis and Beta VaGuoiar (HL translocating) Pyrophosphatase~ respectively] encode a 770-amino acid polypeptide with a predicted mass of 81 kDa. [The discrepancy between apparent relative mass estimated from electrophoretic mobility (70 kDa) and computed
350
molecular weight (81 kDa) is a common feature of membrane proteins.] Computer-assisted hydropathy plots of the Arabidopsis and Beta vacuolar HL PPase amino acid sequences have established that AVP and BVP encode an extremely hydrophobii: integral membrane protein containing 13-16 amphipathic ('multimeric') transmembrane spans which may serve to generate and stabilize a relatively compact structure through the formation of inter-helical hydrogen bonds and salt bridges (Fig. 2). In addition, several of the putative hydrophilic domains are characterized by the presence of clusters of charged residues which may participate in cation (Mg2÷, Ca2÷, K~) or anion (pp4-, MgPPi2-) binding. Thus, the hydrophilic segment linking membrane spans I and II contains four contiguous glutamate residues (positions 64-67 in the Arabidopsis sequence), the segment linking spans IV and V contains four acidic residues in a stretch of eight amino acids (positions 222-229), the segment between sPans VIII and IX contains the sequence RXRXR (positions 525-529) and the penultimate hydrophilic domain, linking spans XII and XIII, contains
a preponderance of Lys residues. The overall orientation of t h e 81 kDa polypeptide is depicted as in Fig. 2 in accord with the ,positive-inside rule '21 wherein, for the majority of multiply spanning membrane proteins, positively charged amino acids are located predominantly towards the cytoplasmic surface of the membrane. Unique origins. The novelty of the vacuolar H÷-PPase has been confirmed by database searches of the sequence. All previously characterized phosphoanhydride-energized H÷pumps fall into three major categories: F, P and V, based on subunit organization, mechanism and inhibitor sensitivity22. They are subject to inhibition by azide, orthovanadate and bafilomycin, respectively, and the enzymes within each category show functionally significant sequence identities, indicating a common ancestry 4,22. By contrast, the vacuolar H÷-PPase is not inhibited by any of these typespecific inhibitors and computer searches of the nucleotide sequences of AVP and BVP and the deduced sequences of the protein products reveal no detectable homology between this pump and any other sequenced ion
TIBS 17 - SEPTEMBER 1 9 9 2
translocase 2°. The vacuolar H+-PPase is therefore likely to be a member of a Class of previously undescribed ion translocase. Close phylogenic links between the vacuolar H+-PPase and the soluble, nonenergy-conserving PPases are similarly unlikely: (1) all characterized soluble PPases have different subunit sizes from the vacuolar H+-PPase (20 kDa for the prokaryotic enzymes, 32-42 kDa for the eukaryotic enzymes)23; and (2) none of the known sequences for soluble PPases (from Arabidopsis, E. coli, Kluyveromyces and Saccharomyces) 23 align with the deduced sequence of the vacuolar H+-PPase. Of the two candidate homologs remaining - the energy-conserving H+PPases of mitochondria and those of phototrophic bacteria - the former are easily eliminated at both the biochemical and molecular levels. (1) Their structural characteristics are distinct from those of the vacuolar H+-PPase; mitochondrial H+-PPases are F-like in organization and consist of a peripheral 28-30 kDa catalytic subunit associated with the membrane through non-covalent interactions with a transmembranous sector 24. (2) Rat-liver mitochondrial H+-PPase does not cross-react with antibody raised against the vacuolar H+-PPase25. (3) Genomic clones of the 28-30 kDa subunit of the mitochondrial H+-PPase from Saccharomyces cerevisiae show 49% sequence identity to the soluble PPase from the same source 26, but no identity with the vacuolar H+-PPase2° (Y. Kim, E. J. Kim and P, A. Rea, unpublished). Evaluation of a more promising evolutionary connection between the vacuolar H+,PPase and the reversible H+translocating PPase (H+-PPi synthase) of phototrophic bacteria will require sequence data for the bacterial enzyme. The presence of a H+-PP~synthase on the energy-coupling membranes of phototrophic bacteria, notably the purple non-sulfur bacterium Rhodospirillum rubrum, has been known for some time 27, but it has only recently been shown that this translocase is an integral protein of approximately 56 kDa28. Two features of this polypeptide are significant: (1) it is immunologically cross-reactive with the MgPPcbinding subunit of the vacuolar H+-PPase (E. J. Kim, Y. Kim and P. A. Rea, unpublished) and, (2) unlike the 28-30 kDa peripheral, catalytic subunit of the mitochondrial H+-PPase, it is capable of mediating both MgPPi hydrolysis and H+-translocation 28. The likelihood of structural
AVP
Ath
119-1~.GF STDNKP C TYD
PPA-1
See
14 6-DzGzTDwKvIATD DZGzTDWKVIAID DZG~.TDwKvIvID D Q G E K D D K I IAVC
Kla
Spo A th •PPA-2
Sce
CONSENSUS
DDG~.LDWKVIvID
(D/E) xxxxDxKxxxxD
2 57-DVGADLVGKIE 4 8 -~..I P R W T N A K L I~Ira.I PRWTNAKLI~~. I P R W T Q A K L I~" ~..APT V F N C K V ~. ~..VPRWT T G K F ~.
(D/E) xxxxxxxKxE
Figure 3 Alignment of putative catalytic residues of vacuolar H÷-PPase (AVP), soluble PPases (PPA-1) and mitochondrial H÷-PPase (PPA-2). Active-site residues implicated in catalysis are indicated in bold 23. Residues of like charge that are common to all six sequences are shown in large letters. Ath, Arabidopsis thaliana; Sce, Saccharomyces cerevisiae; Kla, Kluyveromyces lactis; Spo, Schizosaccharomyces pombe; the PPA-1 and PPA-2 residue numbering is that of the mature soluble PPase protein from S. cerevisiae.
and functional homologies between the H+-PPases from Rhodospirillum, and possibly other phototrophic bacteria, and plant vacuolar membranes is therefore realistic and exciting. Genomic screens of Rhodospirillum using Arabidopsis H+-PPase cDNAs and their derivatives as probes are currently in progress in one of our laboratories (Y. Kim and P. A. Rea, unpublished). These studies will examine this proposal and explore the possibility that the vacuolar H+-PPase is as ancient as the progenores from which the non-sulfur, photosynthetic bacteria were derived. Putative catalytic site(s). Lack of extensive sequence homologies between the vacuolar H+-PPase and other characterized PPases does not automatically preclude functional analogies between the two classes of enzyme. A case in point is the soluble PPases themselves. Although gross sequence comparisons between soluble PPases from a wide range of organisms reveal only modest sequence identities of between 20 and 27%, residues implicated in catalysis show pronounced conservation 23. X-ray crystallographic and site-directed mutagenesis studies of the soluble PPase f r o m Saccharomyces have disclosed 17 residues that are thought to participate directly in catalysis, 11-16 of which (depending on alignment procedure) are conserved in all sequenced, soluble PPases. It is therefore of potential functional interest that eight of the activesite residues of soluble PPases fall into two configurations, EXTKXE and DX2EXDXKX4D (where X = any amino acid), beginning at positions 48 and 146,
respectively, in the sequence of the
Saccharomyces enzyme, while variants of both motifs (DXTKXEand DX4DXKX4D) are found at positions 257 and 119, respectively, in the deduced amino acid sequence of the vacuolar H+-PPase (Fig. 3). Thus, while the vacuolar H~-PPase and soluble PPases appear to be evolutionarily remote, they may share convergent motifs related to the need for both classes of enzyme to interact with the same substrates and cofactors (Mg2~, MgPPi and/or Mg2PP~). Both putative catalytic motifs have a cytosolic orientation (if the configuration of the vacuolar H+-PPase in Fig. 2 is correct) and the motif that starts at position 119 contains a Cys residue and is immediately flanked by another (Fig. 3). It is therefore tempting to speculate that the pronounced sensitivity of the vacuolar H+-PPase to inhibition and covalent modification by N-ethylmaleimide (NEM) and protection by substrate at the cytosolic face of the membrane u'12 is due to alkylation of one or both of these Cys residues.
Physiological function It is now established that the vacuolar H+-PPase and vacuolar H+-ATPase reside on the same membrane and pump H÷ into the same compartment (Fig. 1). The steady-state ApH and A~ generated by the PPase and ATPase of tonoplast vesicles are non-additive and subject to kinetic control by a common H÷ gradient 3, and intact vacuoles of ethyleneglycoltetraacetic acid (EGTA)-permeabilized cells of Chara7 and single, isolated patch clamped vacuoles of
351
TIBS 17 -
B. vulgaris 29 mediate both PPr and ATP-
dependent H+-translocation. Why are there two transport systems pumping the same ion into the same intracetlular compartment? This key question remains unanswered, but there are two possible explanations for the existence of the vacuolar H÷-PPase. Scavenging system, T h e v a c u o l a r H +PPase may simply scavenge the free energy of the PP~ formed in the pyrophosphorylytic reactions of polymer synthesis3°. While a soluble PPase would merely thermally dissipate the free energy of PP~ hydrolysis, a biologically useful output could be retrieved if some of this energy was conserved as a transmembrane H* gradient; this could then be used for subsequent energy-dependent solute transport. A remarkable characteristic of cellular PP~ levels in plants is their invariance. PP~ levels do not change detectably during light-dark transitions, during rapid changes in the rate of respiration or when tissues are subjected to anoxia or respiratory poisons ~6. Cellular ATP levels, on the other hand; change dramatically under these conditions. Coordinate stabilization of PP~ levels and operation of the vacuolar H*-PPase would provide a back-up system for the maintenance of vacuolar compartmentation despite (temporary) metabolic perturbation. Retention of a functional vacuolar H*PPase in plants may be explicable in terms of the greater range of environmental conditions experienced by plants versus mammalian cells. It has been suggested that H* pumps originally functioned as a pH-stat, disposing of the excess acidity generated by fermentative metabolism31. Conservation of this function and the sustained operation of the H*-PPase under conditions of low phosphate (ATP synthetic), which accompany anaerobiosis, would serve two purposes: elimination from the cytoplasm of the additional H+ produced fermentatively and maintenance of intracellular compartmentation. Mediating H÷ and K÷ translocation is a p o s s i b l e role for the vacuolar H~-PPase. Substrate hydrolysis and H÷ translocation by the vacuolar H*-PPase exhibit an almost absolute requirement for K÷ (Ref. 30) and patch-clamp studies on intact vacuoles show that ion translocation by the H+-PPase is specifically dependent on the presence of K* at the cytosolic, but not vacuolar, face of the membrane32. Since the reversibility of the vacuolar H÷-PPase is critically dependent on the transtonoplast K÷
352
gradient, it has been suggested that rather than merely acting as a supplementary H+ pump, the vacuolar H*PPase may serve to catalyse the transtonoplast translocation of both H* and K* (Ref. 32). Note that while H+coupled transport pathways are known for Na* and Ca2* at the vacuolar membrane, there is no information on the mechanism of K÷transport, despite the importance of this ion as a macronutrient and turgor regulant and the necessity for its energy-dependent transport into the vacuole against the prevailing inside~positive membrane potential.
SEPTEMBER 1992
through analysis of the physiological consequences of altered pump expression in transgenic plants that either express antisense transcripts or ectopically overexpress the H*-PPase. Acknowledgements
This work was supported by the National Science Foundation, the Department of Energy, the University Research Foundation, University of Pennsylvania, the Biological Sciences Research Group, University of Pennsylvania (awarded to P. A. R.) and the Agriculture and Food Research Council UK (awarded to D. S.). This article is Concluding remarks dedicated to the memory of Peter T h e ~abundance and ubiquity of the Mitchell, without whose revolutionary vacuolar H+-PPase in plant cells extends insights and single-minded devotion to conventional notions of the role of PP~ the problem of biological energy transin contemporary biological energy trans- duction none of this would have been duction. The novelty of the vacuolar H÷- possible. PPase and its apparently ancient origins, on the other hand, raise important References 1 Mitchell, P. (1961) Chemiosmotic Coupling in questions concerning the place of PP~ in Oxidative and Photosynthetic Phosphorylation, the evolution of cellular energy metabGlynn Research olism. The results of laboratory exper2 Harold, F. M. (1986) The Vital Force: A Study of Bioenergetics, Freeman iments designed to simulate primitive earth conditions suggest that the for- 3 Rea, P. A. and Sanders, D. (1987) Physiol. Plant 71, 131-141 mation of PP~ and other polyphosphates 4 Nelson, N. and Taiz, L. (1989) Trends Biochem. may have been pivotal for prebiotic Sci. 14, 113-116 5 Rea, P. A. and Poole, R. J. (1986) Plant Physiol. evolution: these molecules will catalyse 81, 126-129 the phosphorylation of adenosine, AMP 6 Raven, J. A. (1987) New Phytol. 106, and ADP and the synthesis of polypep357-422 tides 33. Moreover, since the synthesis of 7 Shimmen, T. and MacRobbie, E. A. C. (1987) Protoplasma 136, 205-207 PP~ from precipitated orthophosphate 8 Ikeda, M. et al. (1991) Biochim. Biophys. Acta is practicable at low temperatures 34and 1070, 77-82 these conditions approximate those for 9 Lichko, L. and Okorokov, L. (1991) Yeast 7, 805-812 template-directed oligomerization of nucleotides35, it is conceivable that the 10 Maeshima, M. and Yoshida, S. (1989) J. Biol. Chem. 264, 20068-20073 generation of PPi and the synthesis of 11 Rea, P. A, Britteni C. J. and Sarafian, V. Plant biopolymers were necessary concomiPhysiol. (in press) tants for the emergence of life. Any sys- 12 Britten, C. J., Turner, J. C. and Rea, P. A. (1989) FEBS Lett. 256, 200-206 tem which could harness the energy 13 Sarafian, V. and Poole, R. J. (1989) Plant contained in PP~ would consequently Physiol. 91, 34-38 have been at a replicative advantage. Is 14 Britten, C. J., Zhen, R., Kim, E. J. and Rea, P. A. J. Biol. Chem. (in press) the energy-conserving vacuolar H*15 Kornberg, A. (1963) in Horizons in Biochemistry PPase a product of life's prebiotic ori(Kasha, M. and Pullman, B., eds), gins before the emergence of the more pp. 251-264, Academic Press specific and recognizable structures of 16 Weiner, H., Stitt, M. and Heldt, W. (1987) Biochim. Biophys. Acta 893, 13-21 ATP (and other nucleotides) for nucleic 17 Takeshige, K. and Tazawa, M. (1989)J. Biol. acid synthesis, adenylation, pyrophosChem. 264, 3262-3266 phorylation and signal transduction? 18 Rebeille, F., Bligny, R. and Douce, R. (1984) Plant Physiol. 74, 355-359 From the standpoint of contemGuynn, R. W., Veloso, D., Lawson, J. W. R. and porary plant-cell function, the ready 19 Veech, R. L. (1974) Biochem. J. 140, 369-375 availability of cDNAs encoding the 20 Sarafian, V., Kim, Y., Poole, R. J. and Rea, P. A. (1992) Proc: Natl Acad. Sci. USA 89, vacuolar H*-PPase should allow the full 1775-1779 physiological significance of the en21 von Heijne, G. and Gavel, Y. (1988) Eur. J. zyme in vivo to be uncovered. The role Biochem. 174, 671-678 of the vacuolar H*-PPase in cellular 22 Pedersen, P. L. and Carafoli, E. (1987) Trends Biochem. Sci. 12, 146-150 energy conservation, turgor regulation, resistance to anaerobiosis and vacuolar 23 Cooperman, B. S., Lahti, R. and Baykov, A. A. Trends Biochem. ScL (in press) solute compartmentation in the intact 24 Mansurova, S. E. (1989) Biochim. Biophys. Acta plant is now amenable to investigation 977,237-247
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.....
25 Maeshima, M. (1991) Eur. J. Biochem.
196, 11-17 26 Lundin, M., Baltscheffsky, H. and Ronn~i H. (1991) J. Biol. Chem. 266, 12168-121-72 27 Baltsheffsky, M. and Nyren, P. (1987) irl'/
29 Kurkdijan, A. and Hedrich, R. (1988) EMBO J. 7, 30 31
Phosphate Metabolism and Cellular RegUlation in Microorganisms (Torriani-Gorini,A. et al., eds), pp. 260-263, American So(~ietyfor
32
Microbiology
33
28 Nyren, P., Note, B. and Strid, A. (1991) Biochemistry 30, 2883-2887 i'. i'
THE EXISTENCE OF an enzymatic activity responsible for the termination of acetylcholine (ACh)-mediated neurbtransmission at cholinergic synapseg was postulated as early as 1914 by Dale (see Refs 1-3). Both of the AChdegrading enzymes, acetylcholinesterase (acetylcholine acetyl hydrolase, ACHE; EC 3.1.1.7) and butyrylcholinesterase (acylcholine acyl hydrolase, BCHE; EC 3.1.1.8), are evolutionarily conserved type B carboxylesterases that share extensive amino acid homology and display similar catalytic properties. There are, however, some important exceptions: ACHE specifically interacts with ACh and is subject to marked substrate inhibition, whereas BCHE is capable of degrading a wider range of choline esters and shows no evidence of substrate inhibition ~-3. Furthermore, each enzyme displays characteristic affinities to various selective inhibitors ~,2. Both enzymes appear as monomers, dimers and tetramers, as well as heavier forms bound to structural subunitsS; these are thought to participate in regulating the extent and duration of ACh interaetions with its cellular receptors. The ability of the subunits to interact with various inhibitors means that ACHE and BCHE are sensitive targets for natural and synthetic cholinergic toxins, including toxic glyco-alkaloids, highly poisonous organophosphorous (OP) and carbamate insecticides, snake venom peptides and diverse synthetic therapeutic agents ~-4. Cholinesterase (CHE) activities have been extensively studied by multidiSH. Soreq, A. Gnatt and Y. Loewenstein are at
the Departmentof Biological Chemistry, The Life Sciences Institute, The HebrewUniversity of Jerusalem, Israel 91904. L. F. Neville is at The David MayoneyInstitute of Neurological Sciences, University of Pennsylvania; PA 19104-6074, USA. :ii.... © 1992, Elsevier Science Publishers, (UK)
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3661-3666 Rea, P. A. and Poole, R. J. (1985) Plant Physiol. 77, 46-52 Raven, J. A. and Smith, F. A. (1976) J. Theor. Biol. 57,301-302 Davies, J. M., Rea, P. A. and Sanders, D. (1991) FEBS Lett. 278, 66-68 Yamanaka, J., Inomata, K. and Yamagata, Y. (1988) Origins Life 18, 165-178 Hermes-Lima,M. (1990) J. Mol. Evol. 31, 353-358
35Acevedo, O. L. and Orgel, L. E. (1986) Nature 321, 790-792 36 Gamier, J., Osguthorpe, D. J. and Robson, B. (1978) J. Mol. Biol. 120, 97-120
See also the review by Barry S. Cooperman, 'Evolutionary conservation of soluble inorganic pyrophosphatases active site structure and mechanism' in the July issue of TIBS.
Excavations into the active-site gorge of cholinesterases
Acetyl- and butyrylcholinesterase (ACHE, BCHE) from evolutionarily distant species display a high degree of primary sequence homology and have biochemically similar catalytic properties, yet they differ in substrate specificity and affinity for various inhibitors. The biochemical information derived from analyses of ACHE and BCHE from human, Torpedo, mouse, and Drosophila, as well as that from the recombinant forms of their natural variants and site=directed mutants, can currently be re-examined in view of the recent X-ray crystallography data revealing the three-dimensional structure of Torpedo ACHE. The picture that emerges deepens the insight into the biochemical basis for choline ester catalysis and the complex mechanism of interaction between cholinesterases and their numerous ligands.
ciplinary approaches, which began with detailed biochemical studies of CHEs from several species. This resulted in intricate theories explaining the catalytic properties and inhibitor interactions that are characteristic of various CHEs1-5, More recently, the primary amino acid sequence of CHEs from evolutionarily diverse species was uncovered by gene cloning (for review see Refs 2, 5) and heterologous expression systems have been developed for recombinant CHEs from man 6-8, mouse ~, Torpedo(electric ray) 5,9 and Drosophila ~°, All of these approaches, in addition to affinity labeling H and monoclonal antibody studies ~2, suggested the involvement of several separate amino acid domains in the active center of CHEs, but could n o t identify these domains unequivocally. Recently, the three-dimensional analysis of Torpedo ACHE~3 has deepened, clarified and altered our understanding of CHEs considerably, providing structural infor-
mation on the positioning of catalytically important amino acid residues within these proteins ~3.In an attempt to integrate the information from these various disciplines, we superimposed highly homologous CHEs and their mutants into a single molecular model. The Catalytic- and ligand-binding properties of this composite CHE were analysed in relation to the threedimensional locations of the altered residues with a view to delineating CHE structure-function relationships.
Catalysis by cholinesterases Biochemical analyses demonstrated that catalysis by CHEs involves an active Ser residue (for review see Refs 1-5) and predicted that CHEs, like serine proteases (i.e. trypsin) hydrolyse choline esters through electron transfer within a catalytic triad. In the case of cholinesterases, a negatively charged residue draws a hydrogen atom from an adjacent His which, in turn, draws a
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