calvin cycle

Chapter 2The Calvin Cycle and Its Regulation

William MartinInstitut fr Genetik, Technische Universitt Braunschweig,

Spielmannstr. 7, D-38023 Braunschweig, Germany

Renate ScheibePflanzenphysiologie, FB 5 Biologie/Chemie, Universitt Osnabrck,

D-49069 Osnabrck, Germany

Claus SchnarrenbergerInstitut fr Pflanzenphysiologie und Mikrobiologie, Freie Universitt Berlin,

Knigin-Luise-Str. 12-16a, D-14195 Berlin, Germany

SummaryI.II.

IntroductionThe Enzymes of the Calvin Cycle

A.B.C.D.E.F.G.H.

Ribulose-1,5-bisphosphate Carboxylase/oxygenasePhosphoglycerate KinaseGlyceraldehyde-3-phosphate DehydrogenaseTriosephosphate IsomeraseFructose-1,6-bisphosphate/Sedoheptulose-1,7-bisphosphate AldolaseFructose-1,6-bisphosphataseSedoheptulose-1,7-bisphosphataseTransketolase

I.J.K.

Ribulose-5-phosphate 3-epimeraseRibose-5-phosphate IsomerasePhosphoribulokinase

III.IV.

V.VI.

Calvin Cycle Gene Organization, Expression, and Regulation in EubacteriaCalvin Cycle Expression in Plants

A.B.C.D.E.

Quantification of ActivitiesExpression Studies of Enzyme Activities and TranscriptionGene Regulation Through High Sugar Sensing, and Redox StateRegulation in Specific SystemsCalvin Cycle Enzymes and Expression in Euglena gracilis

Enzyme Interactions and Multienzyme-like ComplexesBiochemical Regulation in Chloroplasts

The Ferredoxin/Thioredoxin SystemA.B.C.

Target EnzymesPhysiological Consequences

VII.VIII.

Studies of Calvin Cycle Enzymes with Antisense RNAConcluding Remarks

AcknowledgmentReferences

R. C. Leegood, T. D. Sharkey and S. von Caemmerer (eds), Photosynthesis: Physiology and Metabolism, pp. 951. 2000 Kluwer AcademicPublishers. Printed in The Netherlands.

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Summary

The Calvin cycle is the starting point ofcarbon metabolism in higher plants. It is a typically eubacterial pathway,as comparative biochemistry of all of its enzymes from prokaryotes and eukaryotes has revealed. The structuralbasis of Calvin cycle function is reviewed with an attempt at a balanced consideration of biochemical andmolecular findings. The structural diversity ofprokaryotic enzymes is emphasized, since the genes encoding thepathway in eukaryotes have all been inherited by plants from prokaryotes through endosymbiosis. Curiously,the enzymes that constitute the pathway in different organisms are often structurally unrelatedwhat isconserved in evolution is merely the set of substrate conversions, not the enzymes that catalyze them. Some ofthe structural and regulatory properties of the enzymes were present in the antecedents oforganelles, but otherswere newly acquired at the eukaryotic level. The expression of Calvin cycle genes is regulated by a widespectrum of factors, though the molecular details of the regulation have yet to be unraveled. Findings thatsuggest the existence of multienzyme-like Calvin cycle complexes are summarized. The molecular basis ofredox-modulated light regulation through the thioredoxin system and its importance for flexible control of thepathway under varying conditions is illustrated. Expression of Calvin cycle enzymes in response to external orinternal stimuli is briefly reviewed, as are newer findings from the expression of antisense constructs of Calvincycle enzymes in transgenic plants.

I. Introduction

The Calvin cycle is one of four known pathways offixation in nature, the three other pathways

being the reverse (or reductive) citric acid cycle(Evans et al., 1966; Beh et al., 1993; Schnheit andSchfer, 1995), the reductive acetyl-CoA (or Wood-Lungdahl) pathway (Fuchs and Stupperich, 1986;Ragsdale, 1991; Schnheit and Schfer, 1995), andthe recently discovered 3-hydroxypropionate pathway(Strauss and Fuchs, 1993;Ishii et al., 1996). However,the Calvin cycle is the only pathway of fixationknown to occur in plants (Fig. 1). It therefore figuresprominently in plant biochemistry, albeit undervarious acronyms, among them the reductive pentose

Abbreviations: 1,3BPGA - 1,3 bisphosphoglycerate; 3PGA 3-phosphoglycerate; Aldolase fructose- 1,6-bisphosphate aldolase;CBB Calvin-Benson-Bassham; cpDNA chloroplast DNA;CTE C-terminal extension; DHAP dihydroxyacetonephosphate; DTT dithiothreitol; E4P erythrose 4-phosphate;F1,6BP fructose 1,6-bisphosphate; F6P fructose 6-phosphate;FBPase fructose-1,6-bisphosphatase; FTR ferredoxin/thioredoxin reductase; GA3P glyceraldehyde 3-phosphate;GAPDH glyceraldehyde 3-phosphate dehydrogenase; MDH malate dehydrogenase; PGK phosphoglycerate kinase;inorganic phosphate; PRK phosphoribulose kinase; R5Pribose-5-phosphate; RPE ribulose-5-phosphate 3-epimerase;RPI ribose 5-phosphate isomerase; Ru1,5BP ribulose 1,5-bisphosphate; Ru5P ribulose-5-phosphate;Rubisco ribulose-1,5-bisphosphate carboxylase/oxygenase; SBPase sedohep-tulose-l,7-bisphosphatase; Su1,7BP sedoheptulose 1,7-bisphosphate; Su7P sedoheptulose 7-phosphate; Td thioredoxin; TKL transketolase; TPI triosephosphateisomerase; Xu5P xylulose 5-phosphate

phosphate pathway (RPPP), the photosyntheticcarbon reduction (PCR) cycle, the Calvin-Benson-Bassham (CBB) pathway, the Benson-Calvin cycle,the C3 cycle, and so on. The enzymes of the Calvincycle have been previously reviewed by Latzko andKelly (1979), Robinson and Walker (1981) andLeegood (1990). Regulation of the Calvin cycle hasbeen reviewed by Buchanan (1980), Macdonald andBuchanan (1990), Geiger and Servaites (1994) and,in cyanobacteria, by Tabita (1994). Historicaldevelopments surrounding the elucidation of thepathway have been briefly summarized elsewhere(Schnarrenberger and Martin, 1997).

In the Calvin cycle, ATP and NADPH from thelight reactions of the photosynthetic membrane areexpended to reduce to carbohydrate. From thestandpoint of ATP investment per mole of fixed,the Calvin cycle is the most costly of the fourfixation pathways known (Strauss and Fuchs, 1993).The basics of the pathway were clarified throughtracer studies in eukaryotic algae over 40 years ago(Calvin, 1956). The net reaction can be summarizedas

Mutants defective in fixation in the facul-tatively anaerobic, chemoautotrophic proteobacteriaRhodobacter sphaeroides (Gibson and Tabita, 1996)and Ralstonia eutropha (previously Alcaligeneseutrophus) (Kusian and Bowien, 1997) have been a

William Martin, Renate Scheibe and Claus Schnarrenberger

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powerful tool for understanding the molecular biologyand the genetic regulation of the pathway in theseorganisms. Molecular sequences are known for all ofthe enzymes of the pathway from spinach chloroplasts(Martin and Schnarrenberger, 1997) and from thegenome sequence of the cyanobacterium Synecho-cystis PCC6803 (Kaneko et al., 1996) and, with a fewexceptions, from Rhodobacter sphaeroides (Gibsonand Tabita, 1997) and Ralstonia eutropha (Bommeret al., 1996). Several lines of reasoning support theview that in order to understand the Calvin cycle ofhigher plant chloroplasts in a broader context, it isuseful to consider regulation and structural diversitywithin the pathway among eubacteria.

First, the pathway did not evolve de novo in plants,but rather was inherited from eubacteria via theendosymbiotic origins of organelles. As a conse-quence, manybut not allof the regulatoryproperties that are observed among the enzymes ofthe higher plant pathway arose at the prokaryoticlevel and were simply maintained within the plant

lineage, having been genetically transmitted fromthe cyanobacterial antecedents of plastids. Theenzymes of the pathway in chloroplasts are not allacquisitions from cyanobacteria, some are acquisi-tions from mitochondria (Martin and Schnar-renberger, 1997; Martin and Mller, 1998) that werererouted during evolution to a new target organelle(Martin and Herrmann, 1998).

Second, in most plastids, at least one enzyme ofthe pathway (one or both subunits of Rubisco) is stillencoded in chloroplast DNA (cpDNA), establishinga requirement for coordination of gene expressionbetween plastids and the nucleus in order to properlyexpress the pathway. Indeed, the plastids of somealgae even still possess the cbbR gene (Stoebe et al.,1998) which encodes a homologue of the trans-criptional regulator of Calvin cycle gene expressionin eubacteria.

Third, the quantitatively most important mechan-ism governing the activity of higher plant Calvincycle enzymeslight activation via the thioredoxin

Chapter 2 Calvin Cycle

12

systemis present and active in cyanobacteria. Thisregulatory mechanism was also inherited by plantsfrom the eubacterial antecedents of plastids, althoughthe molecularbasis for the covalent transitions in thetarget enzymes can differ between cyanobacteria andplants.

Fourth, although the mechanisms of Calvin cyclegene regulation in eubacteria are probably much lesscomplex than those in eukaryotes, by no means arethey irrelevant to our understanding of eukaryoticCalvin cycle gene regulation. On the contrary, due totheir simplicity and tractability, mechanisms ofCalvincycle gene regulation are much better understood ineubacteria than in eukaryotes. And with the recentdiscovery of a cyanobacterial homologue ofphytochrome (Hughes et al., 1997; Yeh et al., 1997),it appears that at least some of the basic machineryfor Calvin cycle gene regulation through light ineukaryotes were simply inherited from prokaryotesthrough endosymbiosis, although the actual signaltransduction pathways in prokaryotes and eukaryotesthat lead to gene regulation through light will, inmany cases, turn out to be quite different.

Finally, although the series ofsubstrate conversionsthat constitute the Calvin cycle are strictly conservedacross eubacteria and eukaryotes, the same degree ofconservation does not apply to the enzymes thatcatalyze those reactions. In fact, in this chapter wewill see that the pathway in proteobacteria,cyanobacteria and higher plant chloroplasts consistsof enzymes that catalyze identical reactions, but, insome cases, that are altogether unrelated at the levelof sequence, structure and reaction mechanism (seeMartin and Schnarrenberger, 1997).

Many of the subsequent chapters in this volumedeal, in one way or another, with various aspects ofthe Calvin cycle, including Rubisco itself (Chapters3, (Roy and Andrews) and 4 (von Caemmerer andQuick)), metabolite transport (Chapter 6, Flgge),C4 metabolism (Chapters 18 (Furbank et al.) and 19(Leegood)) and chloroplast-cytosol interactions(Chapters 7 (Aiken et al.) and 8 (Foyer et al.)). In thischapter, we will focus on structural, functional andregulatory aspects of the enzymes that constitute thepathway, emphasizing insights provided by molecularapproaches, but considering classical biochemicalaspects as well.

II. The Enzymes of the Calvin Cycle

A schematic comparison of Calvin cycle enzymes in

the Ralstonia eutropha (formerlyAlcaligenes eutrophus) (Bowien et al., 1993) andthose encoded in the genome of the cyanobacteriumSynechocystis PCC6803 (Kaneko et al., 1996a,1996b) reveals that pathways in these bacteriacomprise the same sets of substrate conversions, butin several cases with the help of enzymes that arenon-homologousor very nearly so (Fig. 2). Suchstructurally distinct but functionally homologousenzymes are traditionally designated as class I/classII enzymes, a term that will be used here. Differencesalso exist between the pathways in spinachchloroplasts and Synechocystis (e.g. use of class I vs.class II aldolase, respectively), but as depicted inFig. 2, these differences are less grave than across thetwo eubacteria compared. The following sectionsprovide a synopsis of structural and functionaldiversity for each Calvin cycle enzyme. Regulationof individual enzymes by covalent modificationthrough the ferredoxin/thioredoxin system (recentlyreviewed by Jacquot et al., 1997b) will be consideredlater in this chapter.

A. Ribulose-1,5-bisphosphate Carboxylase/oxygenase

Ribulose-1,5-bisphosphate carboxylase/oxygenase(EC 4.1.1.39, Rubisco) catalyzes the initialfixation step. The mechanism involves an activatingcarbamylation reaction between and thegroup of an active site lysine residue in the largesubunit (Lorimer and Miziorko, 1980). Car-bamylation is promoted by Rubisco activase (Portis,1992). For details of Rubisco kinetics, catalyticmechanism and regulation, see Chapters 3 (Roy andAndrews) and 4 (von Caemmerer and Quick). Thecrystal structure ofRubisco from spinach chloroplastsis known at great resolution, it is a stout cylindricaltetramer of dimers that are glued together byfour small subunits at each end (Shibata et al., 1996;Andersson, 1996). Two structurally distinct Rubiscoenzymes are known. Class I (or form I) Rubisco hasa native of about 560 kDa and consists of eightlarge subunits (LSU, ~55 kDa each) and eightsmall subunits (SSU, ~ 15 kDa each) comprisingthe holoenzyme. Assembly of the hetero-hexadecamer requires the aid of chaperonins in bothchloroplasts and eubacteria (Goloubinoff et al., 1989;Gatenby and Viitanen, 1994; Gutteridge and Gatenby,1995).


13

Class II Rubisco consists only of large subunits of~55 each holoenzyme) (Gibson and Tabita,

1996; Kusian and Bowien, 1997). The class I andclass II large subunits share about 30% amino acididentity, indicating that they share a common ancestor.Rubisco gene diversity is a complicated matter andhas been discussed in detail elsewhere (Watson andTabita, 1996; Martin and Schnarrenberger, 1997). Atleast two very ancient gene duplications (or lateraltransfers) have occurred in Rubisco evolution, onethat gave rise to the class I and class II enzymes anda second that gave rise to the two distinct families ofclass I Rubisco found in chlorophytic (green or G-type Rubisco) and rhodophytic plastids (red or R-type Rubisco), respectively (Martin and Schnar-renberger, 1997). Cyanobacteria and many andproteobacteria studied to date possess G-type Rubisco(Watson andTabita, 1996), whereas the proteobacteriaRalstonia, Rhodobacter and Xanthoflavus encode R-

type Rubisco in their cbb (Calvin-Benson-Bassham)operons. But Rhodobacter, like Rhodospirillum,Hydrogenovibrio and several other eubacteria, alsoencode and express the class II enzyme (Falcone andTabita, 1991; Stoner and Shively, 1993; Gibson andTabita, 1996). Notably, the curious Rubisco gene inthe Methanococcus genome encodes an active, O2-sensitive enzyme (Watson et al., 1999).

Higher plants, like cyanobacteria, possess G-typeclass I Rubisco, whereby in all species studied, thelarge subunit is encoded as a single copy in thecpDNA and the small subunit is encoded in thenucleus, usually as a gene family (see below). Theprimitive photosynthetic protist Cyanophoraparadoxa is an exception, however, in that both thelarge and the small subunits of its G-type Rubiscoare encoded in the cpDNA (Lambert et al., 1985).Rhodophytes and photosynthetic protists that haveobtained their plastids from rhodophytes via


14

secondary endosymbiosis (McFadden et al., 1996;Van de Peer et al., 1996) encode both subunits of R-type Rubisco in their cpDNA, and also encode ahomologue of cbbR, the transcriptional regulator ofCalvin cycle operons in proteobacteria. The onlyexamples in which eukaryotes have been shown topossess class II Rubisco have been described for thevery diverse group of photosynthetic protists ofsecondary symbiotic origin known as dinoflagellates(Morse et al., 1995; Rowan et al, 1996), where, quitesurprisingly, the gene for the class II Rubisco largesubunit is encoded in the nucleus. The diversity ofeukaryotic Rubisco genes is, in all likelihood, simplythe result of sampling from ancient eubacterial genediversity present in the common ancestor ofendosymbiotic organelles, very similar to allelesampling in population genetics, but on a geologicaltime scale (Martin and Schnarrenberger, 1997).

B. Phosphoglycerate Kinase

Phosphoglycerate kinase (EC 2.7.2.3, PGK) catalyzesthe reversible transfer of the of ATP tothe carboxyl group of 3-phosphoglycerate (3PGA),forming 1,3-bisphosphoglycerate (1,3 BPGA) for thesubsequent reduction step. In all prokaryotic andeukaryotic sources studied to date, the active enzymeis a monomer with an of ~44 kDa (Fothergill-Gilmore and Michels, 1993). The crystal structure ofthe enzyme from several sources is known. PGK isunusual in that substrate binding induces a dramaticconformational change: the two wings of thebutterfly structure are bent upon 3 PGA and ATPbinding by over 30 degrees, displacing distal regionsof the domains by some 27 (Bernstein et al., 1997).The chloroplast and cytosolic isoenzymes can beseparated with conventional techniques, roughly 90%of the PGK activity is localized in higher plantchloroplasts (Pacold and Anderson, 1975; Kpke-Secundo et al., 1990; McMorrow and Bradbeer,1990). In Chlamydomonas reinhardtii, a cytosolicisoenzyme seems to be lacking (Schnarrenberger etal., 1990; Kitayama and Togasaki, 1995). ChloroplastPGK from various sources shows biphasic kineticswith of~400 and of~500 atlow substrate concentrations, with a pH optimumaround 7.5 (Kpke-Secundo et al., 1990). The enzymehas not been found to be strongly regulated byallosteric effectors or by light (Leegood, 1990). The

enzyme has been cloned from several higher plants(Longstaff et al., 1989; Bertsch et al. 1993) and wasmapped in wheat (Chao et al., 1989). The higherplant nuclear genes for both the chloroplast and thecytosolic enzymes were obtained from cyanobacteriathrough endosymbiotic gene transfer (Brinkmannand Martin, 1996; Martin and Schnarrenberger,1997).

C. Glyceraldehyde-3-phosphateDehydrogenase

Glyceraldehyde-3-phosphate dehydrogenase (EC1.2.1.13, GAPDH) catalyzes thereversible reductive step of the Calvin cycle. Incatalysis, 1,3BPGA forms a highly reactive thioesterbond with the thiol moiety of the active site cysteineresidue under elimination of the acylphosphate. The carbonyl group of the covalentlybound intermediate is reduced to a hemithioacetal byhydride transfer from NADPH and glyceraldehyde3-phosphate (GA3P) is released from the enzymethrough cleavage of the hemithioacetal bond (Brndenand Eklund, 1980). ChloroplastGAPDH has activity with both NAD(H) andNADP(H). Although the of the enzyme istoo low to be relevant during (anabolic) fixation,NADPH being strongly preferred by the enzyme(Cerff, 197 8a), recent studies indicate that theactivity may play an important role during (catabolic)ATP-synthesis in the dark (Backhausen et al., 1998).The kinetic properties are complex and depend uponthe activation state of the enzyme (Cerff, 1978a;Wolosiuk and Buchanan, 1978). In the forwardreaction the fully active purified enzyme has a

of roughly 30 and a of 40(Baalmann et al., 1995). For the reaction in intactchloroplasts, a of1 has been estimated(Fridlyand et al., 1997).

Class I and class II GAPDH enzymes are knownthat share only 1520% sequence identity, both aretetramers of ~150 kDa, consisting of ~37 kDasubunits. Eukaryotes, eubacteria, and one halophilicarchaebacterium possess class I GAPDH (Pr etal., 1994; Brinkmann and Martin, 1996). The tertiarystructure of class I GAPDH is known for numeroussources (Biesecker et al., 1977; Michels et al., 1996).Class II GAPDH has been found only in archae-bacteria (Hensel et al., 1987; Fabry and Hensel,


15

1988; Zwickl et al., 1990). No crystal structures havebeen published for the class II enzyme.

Higher plantCalvin cycle GAPDH differs from allother known GAPDH enzymes in that it is anheterotetramer rather than a homotetramer (Cerffand Chambers, 1979; Ferri et al., 1990; Scagliarini etal., 1993).The tetrameric enzyme can reversiblyaggregate to a multimeric form of 600 to 800 kDa,being less active than the dissociated form (Cerff,1978a, 1978b; Pupillo and Faggiani, 1979; Wara-Aswapati et al., 1980; Trost et al., 1993). Thismultimeric form was probably the form ofthe enzymefirst purified from plants (Yonuschot et al., 1970).Cytosolic GAPDH (EC 1.2.1.12) and non-phos-phorylating GAPDH (EC 1.2.1.9), although tetra-mers, do not show this reversible oligomer formation(Pupillo and Faggiani, 1979). No strong allostericeffectors are known for higher plant GAPDH, but inthe cyanobacterium Synechocystis PCC6803 a lowMW fraction has been described that reduces theactivity of Calvin cycle GAPDH in the reverse(oxidative) direction (Koksharova et al., 1998).

A novel, plastid-specific GAPDH(GapCp) was recently described from Pinuschloroplasts that coexists with GAPDH of theCalvin cycle. It shows no detectable activity with

and has a of 62 pM and of344 (Meyer-Gauen et al., 1994; Meyer-Gauen etal., 1998). GapCp from Pinus is possibly similar tothe reported from isolated, non-photosynthetic plastids of developing cauliflowerbuds (Neuhaus et al., 1993). There is also biochemicalevidence for a similar plastid GAPDHin ripening sweet pepper fruits where, in cooperationwith an MDH, it appears to beimportant in the distribution of reducing equivalentsbetween plastidand cytosol (Backhausen et al., 1998).In some photosynthetic tissues, for example in pineseedlings, GapCp (an enzyme)appears to coexist with Calvin cycle GAPDH

(Meyer-Gauen et al., 1994;Schnarrenberger, unpublished). In some non-photosynthetic tissues, GapCp may functionallyreplace the enzyme. By analogy, in somephotosynthetic protists, an GAPDHenzyme has been recruited from anancestral enzyme (Liaud et al., 1997; Fagan et al.,1998). The nuclear gene for higher plant Calvincycle GAPDH was obtained by plants fromcyanobacteria, the cytosolic enzyme appears to havebeen obtained from the mitochondrial symbiont

Triosephosphate isomerase (EC 5.3.1.1, TPI)catalyzes the rapid and reversible ketose-aldoseisomerization of dihydroxyacetone phosphate(DHAP) and GA3P. The native enzyme in eubacteriaand eukaryotes is a homodimer of ~27 kDa subunits(Fothergill-Gilmore and Michels, 1993), in hyper-thermophilic archaebacteria TPI is a homotetramerof 25 kDa subunits (Kohlhoff et al., 1996). TheCalvin cycle enzyme of higher plant chloroplasts is ahomodimer of ~27 kDa subunits (Kurzok andFeierabend, 1984; Henze et al., 1994; Schmidt et al.,1995). For both the chloroplast and cytosolic enzymesseparated from leaves is ~2 mM andis ~700 (Kurzok and Feierabend, 1984). Thecrystal structure of the enzyme from many sources isknown (Velanker et al., 1997).

As for PGK, class I/class II forms ofTPI have notbeen described. Calvin cycle TPI of higher plantchloroplasts arose through a duplication of the pre-existing eukaryotic nuclear gene for cytosolic TPI,accompanied by the acquisition of a transit peptide(Henze et al., 1994; Schmidt et al., 1995). But sincethe prexisting nuclear gene was itself acquired viaendosymbiotic gene transfer from ancestors ofmitochondria (Keeling and Doolittle, 1997), theCalvin cycle of higher plant chloroplasts functionswith TPI enzyme of mitochondrial origin that wasrerouted to the plastid during evolution (Martin andSchnarrenberger, 1997).

genome (Martin et al., 1993; Henze et al., 1995). TheGapA and GapB subunits of the enzyme arosethrough gene duplication during chlorophyteevolution (Meyer-Gauen et al., 1994). The B subunitis implicated in regulatory properties of the enzyme(Scagliarini et al., 1998) and possesses a CTE ofroughly 30 amino acids relative to the A subunit thatis involved in thioredoxin-dependent regulation (seeSection VI).

D. Triosephosphate Isomerase

E. Fructose-1,6-bisphosphate/Sedoheptulose-1,7-bisphosphate Aldolase


16

Fructose-1,6-bisphosphate aldolase (EC 4.12.1.13,aldolase) catalyzes the reversible aldol condensationof dihydroxyacetone phosphate and either GA3P orerythrose-4-phosphate to yield fructose-1,6-bisphosphate or sedoheptulose-l,7-bisphosphate,respectively. Both activities are part of the Calvincycle. Two very distinct types of aldolase enzymesoccur in nature that differ in their catalytic mechanism(Rutner, 1964; Marsh and Lebherz, 1992). Class Ialdolase enzymes form a Schiff-base with thesubstrate during catalysis via condensation of theamino group of an active-center lysine residue withthe carbonyl group of the substrate. Class II aldolaseenzymes require divalent cations such as

as cofactors which stabilize the carbanionintermediate formed during the reaction. The dualspecificity for F1,6BP and Su1,7BP formation byaldolase applies to the chloroplast enzyme and to thecytosolic enzyme, both of the class I type in higherplants (Brooks and Criddle, 1966; Moorehead andPlaxton, 1990) and of the class II type in Cyanophoraparadoxa (Flechner et al., 1999). Class I aldolasesare homotetramers whereas class II aldolases arehomodimers. The subunit size of both classes ofaldolase enzymes is ~40 kDa, but class I and class IIaldolase monomers share no detectable sequencesimilarity. This, in addition to the different catalyticmechanisms and unrelated crystal structures forclass I (Blom and Sygusch, 1997) and class II (Cooperet al., 1996) aldolase, clearly indicates that these twoclasses of aldolase enzymes are the result ofevolutionary functional convergence. For separatedspinach chloroplast and cytosolic class I aldolase,

is 20 and 1 respectively, whereasis 6 and 4 respectively. The

corresponding values for chloroplast and cytosolicclass II aldolase from Cyanophora paradoxa are

1 mM and 660 respectively, whereasis 200 and 230 respectively

(Flechner et al., 1999).Class I and class II aldolases have a very complex

phylogenetic distribution across prokaryotes andeukaryotes (Henze et al., 1998). Most eubacteria,including all cyanobacteria studied to date, typicallypossess class II aldolase (Rutter, 1964; Antia, 1967),although class I aldolase is known in eubacteria(Witke and Gtz, 1993). Halophilic archaebacteriacan possess either class I or class II aldolase (Dharand Altekar, 1986). Interestingly, the Methanococcusgenome does not encode a recognizable homologueofeither class I or class II aldolase (Bult et al., 1996),

although methanogens are known to possess aldolaseactivity (Yu et al., 1994; Schnheit and Schfer,1995), raising the possibility that a class III aldolasewill eventually be found. A possible candidate forsuch a new class of aldolase has been described froma halophilic archaebacterium (Krishnan and Altekar,1991) that possesses a (mechanistically) class 1aldolase consisting of 27 kDa (rather than 40 kDa)subunits with novel properties. Among highereukaryotes, fungi typically possess class II aldolasewhereas metazoa and higher plants possess class Ialdolase (Schnarrenberger et al., 1990; Marsh andLebherz, 1992; Tsutsumi et al., 1994). Euglenagracilis is exceptional among eukaryotes in that itpossesses both class I aldolase (in the chloroplast)and class II aldolase (in the cytosol) (Pelzer-Reith etal., 1994b). In addition to class I and class II aldolase,ancient eubacterial gene duplications are known inclass II aldolase evolution that have given rise toaldolase-related enzymes specialized for substratesother than sugar phosphates (Plaumann et al., 1997).The Calvin cycle of both proteobacteria andcyanobacteria operates with class II aldolase, whilealdolase of higher plant chloroplasts is a class Ienzyme, and the paucity of sequences for class Ialdolase from prokaryotes makes it currentlyimpossible to tell whence the class I gene for thechloroplast enzyme arose (Plaumann et al., 1997).To add to this conundrum of diversity, the Calvincycle in cyanelles of Cyanophora paradoxa operateswith a class II aldolase (Gross et al., 1994). Thus,Calvin cycle aldolase of plastids has arisen at leasttwice in evolution, and the data for class I aldolase ofEuglenas chloroplasts suggest that a third inde-pendent origin ofCalvin cycle aldolase in plastids islikely (Plaumann et al., 1997).

F. Fructose-1,6-bisphosphatase

Fructose-1,6-bisphosphatase (EC 3.1.3.11, FBPase)catalyzes the cleavage of the phosphoester bond onC1 to yield fructose-6-bisphosphate (F6P). In mostproteobacteria and cyanobacteria, the FBPase andSBPase reactions of the Calvin cycle are catalyzedby a single enzyme (F/SBPase) with dual specificityfor both substrates (Gerbling et al., 1986; Gibson andTabita, 1988; Yoo and Bowien, 1995; Paoli et al.,1995). Xanthobacter flavus is an exception (seebelow). F/SBPase from cyanobacteria (Gerbling et


17

al., 1985) is a tetramer of ~40 kDa subunits, as isFBPase from spinach (Marcus and Harrsch, 1990).The crystal structure of spinach chloroplast FBPasehas been determined (Villeret et al., 1995).

FBPase catalyzes a highly exergonic reaction thatis virtually irreversible under physiological condi-tions, and it is one ofthe key targets for regulation ofthe Calvin cycle. Activity of the enzyme isundetectable in the dark (oxidized state), but increasesto maximum activities within a few minutes ofillumination due to thiol reduction via the thioredoxinsystem (Buchanan, 1980). FBPase is specificallyactivated by thioredoxin (hence the designation(Buchanan, 1980; Lopez-Jaramillo et al., 1997). Themechanism of chloroplast FBPase regulation wasrevealed by altered kinetics observed in the presenceof pH, and thiols (Zimmermann et al., 1976).Activation of chloroplast FBPase by reduced thiolsaffects a dramatic increase of substrate affinity of>20-fold (Charles and Halliwell, 1980),of the fully activated enzyme is 6 as compared to130 for the oxidized enzyme (Cadet and Meunier,1988b). The sensitivity of the chloroplast enzyme tolow concentrations ofmercuric ions has been studiedin several species (Ashton, 1998a).

Both chloroplast and cytosolic FBPase of higherplants are highly regulated (see Section VI), but byquite different mechanisms (Latzko et al., 1974;Zimmermann et al. 1976). The cytosolic enzyme is acontrol point for regulating flux through gluconeo-genesis. Like its homologues from the cytosol ofnon-photosynthetic eukaryotes, it is subject to strongallosteric inhibition by AMP and regulation throughF2,6BP (Stitt, 1990a, 1990b), whereas thioredoxinhas no effect. The chloroplast enzyme on the otherhand, is insensitive to both AMP and F2,6BP.Curiously, these distinct regulatory properties seemto have evolved specifically in the plant lineage. Thisis because higher chloroplast FBPase arose throughgene duplication of the prexisting nuclear gene forcytosolic FBPase, that itself appears to have beenacquired from mitochondria (Martin et al., 1996a;Schnarrenberger and Martin, 1997), indicating thatas in the case of TPIthe higher plant Calvin cyclefunctions with an FBPase enzyme of mitochondrialorigin. The archaebacteria Methanococcus mari-paludis and Haloarcula vallismortis possess highFBPase activity (Altekar and Rangaswamy, 1992; Yuet al., 1994) but the enzyme has not been purifiedfrom any archaeon and the Methanococcus genomedoes not encode a recognizable gene for FBPase

A highly specific sedoheptulose-1,7-bisphosphatase(EC 3.1.3.37, SPB) is not known from prokaryotes,although Xanthobacter flavus differentially expressestwo distinct F/SBPase isoenzymes that both acceptF1,6BP and Su1,7BP as substrates. The isoenzymeexpressed during autotrophic growth (CbbF) hasnearly equal activities with F1,6BP and Su1,7BP(3:1, respectively) as substrates, with a of3 the other isoenzyme possesses much loweractivity with Su1,7BP (van den Bergh et al., 1995).Similarly, Synechococcus PCC 7942 possesses twoimmunologically distinct tetrameric FBPase isoen-zymes, one ofwhich is specific for F1,6BP, the otherof which efficiently cleaves both F1,6BP and Su1 ,7BP(Tamoi et al., 1996). The cyanobacterial F/SBPaseisoenzymes thus have slightly more similar propertiesto those found in higher plants, where chloroplastFBPase and SBPase in plants are separate enzymesencoded by distinct but distantly related nucleargenes (Raines et al., 1988,1992; Martin et al., 1996a).In contrast to FBPase and bacterial F/SBPase, whichare tetramers, SBPase from higherplant chloroplastsis a dimer of ~35 kDa subunits (Nishizawa andBuchanan, 1981; Cadet et al., 1987). Also in contrastto many bacterial F/SBPase enzymes, higher plantchloroplast FBPase and SBPase show a highspecificity for their respective substrates, wherebychloroplast SBPase is highly, but not completelyspecific for Su1,7BP (Zimmermann et al., 1976;Breazeale et al., 1978; Cadet and Meunier, 1988b).Chloroplast SBPase, as FBPase, is redox-modulatedby thioredoxin (Breazeale et al., 1978; Nishizawaand Buchanan, 1981). The reduced (activated)enzyme has a of 50 and a of380 (Cadet and Meunier, 1988b). But since the

of reduced (activated) chloroplast FBPaseis 6 the F1,6BP activity of SPB is probably oflittle or no physiological relevance. Similar reasoningapplies to the SBPase activity of chloroplast FBPase(Ashton, 1998b). SBPase from wheat has beenexpressed in E. coli (Dunford et al., 1998). F2,6BP,a potent allosteric regulator ofcytosolic FBPase, hasno allosteric effect on chloroplast SBPase, but can

(Bult et al., 1996), raising the possibility thatstructurally unrelated class I and class II FBPaseenzymes may exist.

G. Sedoheptulose-1,7-bisphosphatase


18

act as a competitive inhibitor (Cadet and Meunier,1988b). The evolutionary relationship betweenchloroplast SBPase and eukaryotic FBPase andeubacterial F/SBPase enzymes is unclear (Martin etal., 1996a), but it appears that the specialization ofSBPase from a bifunctional F/SBPase ancestoroccurred at the prokaryotic level.

H. Transketolase

Transketolase (EC 2.2.1.1, TKL) catalyzes thereversible, thiamine diphosphate-dependent transferof a two carbon ketol group from either fructose-6-phosphate or sedoheptulose-7-phosphate (Su7P) toglyceraldehyde-3-phosphate to yield xylulose-5-phosphate (Xu5P) and either erythrose-4-phosphateor ribose-5-phosphate (R5P), respectively. TKL fromvarious sources is a homodimer of 74 kDa subunits(Feierabend and Gringel, 1983). The crystal structureof the yeast enzyme is known (Nikkola et al., 1994;Nilsson et al., 1997). The catalytic mechanisminvolves nucleophilic attack ofthe substrate carbonylgroup via the C2 carbanion of thiamine diphosphate(ThDP): the rate-limiting C2 deprotonation steprequires interaction of N1' in the ThDP pyrimidinering with (Kern et al., 1997).

Beyond the studies of Murphy and Walker (1982),who purified the enzyme 400-fold, and Feierabendand Gringel (1983), who found only a singlechloroplast species, little attention has been given tothe biochemistry of this Calvin cycle enzyme.Substrate affinities for the plant enzyme have beenreported as 100-130 for Xu5P, E4P and R5P(Murphy and Walker, 1982), for human erythrocytesthe values 20 30 and 2mM were found(Himmo et al., 1989). For the purifiedenzyme from spinach chloroplasts 77and 330 were found (Teige et al., 1998).TKL has been cloned from Craterostigma (Bernacciaet al., 1995), spinach (Flechner et al., 1996) andpotato (Teige et al., 1996). The enzyme from spinachchloroplasts has been expressed in highly activeform in E. coli (Flechner et al., 1996). Spinach leavesappear to possess only a single TKL enzyme, localizedexclusively in the chloroplast (Feierabend andGringel, 1983; Schnarrenberger et al., 1995). TKLshows structural similarity to several other enzymes

involved in ThDP-dependent C2 metabolism:pyruvate decarboxylase and the E1 subunit ofpyruvate dehydrogenase (Robinson and Chun, 1993).The nuclear gene for the Calvin cycle enzyme ofhigher plants was acquired from cyanobacteria(Martin and Schnarrenberger, 1997).

I. Ribulose-5-phosphate 3-epimerase

Ribulose-5-phosphate 3-epimerase (EC 5.1.3.1, RPE)catalyzes the reversible interconversion of ribulose-5-phosphate and xylulose-5-phosphate. RPE is ahomodimer of~23 kDa subunits in animals (Karmaliet al., 1983), Ralstonia (Kusian et al., 1992) andspinach (Nowitzki et al., 1995). The spinach enzymehas been purified to homogeneity and N-terminallysequenced (Teige et al., 1998). The purified spinachenzyme migrates as an octamer, the wasdetermined as 250 (Teige et al., 1998). RPEfrom the red alga Galdieria sulphuraria has aof ~830 (J. Girnus, W. Gross and C. Schnar-renberger, unpublished). Spinach leaves appear topossess only a single RPE enzyme, localized inchloroplasts (Schnarrenberger et al., 1995). RPE hasbeen cloned from sorghum and spinach (Nowitzki etal, 1995) and potato (Teige et al., 1995), the enzymefrom spinach chloroplasts has been expressed inactive form in E. coli (Nowitzki et al., 1995). Morerecently, the enzyme from spinach chloroplasts wascloned again, and was expressed in E. coli again(Chen et al., 1998). Neither the mechanism of catalysisnor the tertiary structure have been reported fromany source. Class I / class II RPE enzymes have notbeen described, but three very distantly rpe- relatedgenes exist in the E. coli genome, indicating thepresence of relatively ancient eubacterial genefamilies (Nowitzki et al., 1995). The nuclear gene forhigher plant Calvin cycle RPE was acquired fromcyanobacteria (Martin and Schnarrenberger, 1997).

J. Ribose-5-phosphate Isomerase

Ribose-5-phosphate isomerase (EC 5.3.1.6, RPI)catalyzes the reversible isomerization of ribose-5-phosphate and ribulose-5-phosphate. RPI has notbeen identified in the cbb operons of photosyntheticproteobacteria (Gibson and Tabita, 1996). No crystal


19

structures have been reported for this enzyme. Rutner(1970) purified RPI from spinach 2800-fold. Only asingle enzyme was found, a homodimer of 23 kDasubunits, as later shown for Arabidopsis (Babad-zhanova and Bakaeva, 1987), that had a of460 Chloroplast RPI from spinach (Martin etal., 1996b) has been cloned, it has sequence similarityto RpiA from E. coli (Hove-Jensen and Maigaard,1993). But E. coli also possesses a gene for a secondfunctional RPI enzyme, RpiB, that is a homodimerof 16 kDa subunits. It shows no sequence similarityto RpiA, but very high similarity to galactose-6-phosphate isomerases (Srensen and Hove-Jensen,1996). Thus for RPI, class I (e.g. spinach RPI orRpiA of E. coli) and class II (RpiB of E. coli)enzymes should be distinguished. No cytosolicisoenzyme of RPI was found in spinach leaves(Schnarrenberger et al., 1995). Calvin cycle (class I)RPI from spinach has identifiable homologuesencoded in the Synechocystis and Methanococcusgenomes, but due to paucity of reference sequences,the evolutionary origin of the plant nuclear gene instill unclear.

K. Phosphoribulokinase

Phosphoribulokinase (EC 2.7.1.19, PRK) transfersthe of ATP to the C1 hydroxyl group ofribulose-5-phosphate, regenerating the primaryacceptor. Class I and class II PRK enzymes areknown (Tabita, 1994; Brandes et al., 1996a; Martinand Schnarrenberger, 1997). Class I PRK is encodedin proteobacterial cbb operons. It is an octamer of~30 kDa subunits with allosteric inhibition throughAMP and allosteric activation through NADH(Runquist et al., 1995). Crystal structure data hasbeen reported for class I PRK (Roberts et al., 1995;DHT Harrison et al., 1998). Class II PRK is found incyanobacteria and higher plants. It is a dimer of ~44kDa subunits in chloroplasts (Porter et al., 1986;Clasper et al., 1994). The enzyme can associate totetramers in Synechocystis (Wadano et al., 1995).The 300-foldpurified enzyme from the chromophyticprotist Heterosigma carterae is a tetramer of 53 kDasubunits with a of 208 and of226 (Hariharan et al., 1998). Crystal structureshave not been reported for the class II enzyme. Thecatalytic properties of class I and class II PRK differmarkedly (Tabita, 1988). The nuclear gene for the

higher plant Calvin cycle enzyme was acquired fromcyanobacteria (Martin and Schnarrenberger, 1997).

PRK catalyzes a highly exergonic reaction and isstrongly regulated by the thioredoxin system(Buchanan, 1980). The oxidized (dark) enzymepossesses only about 2% of the activity of the fullyactive (reduced) form (Surek et al., 1985). Kineticvalues of of 60 and of 110were reported for the spinach enzyme expressed inthe yeast Pichia pastoris (Brandes et al., 1996a),similar to values determined for the purified nativeactivated wheat enzyme (Surek et al., 1985). Incontrast to GAPDH, FBPase, and SBPase, thio-redoxin activation of PRK does not involve loweringof values, but affects the (Porter et al., 1986).

III. Calvin Cycle Gene Organization,Expression, and Regulation in Eubacteria

Several excellent reviews on this topic haveappeared recently (Tabita, 1994; Gibson and Tabita,1996; Bommer et al., 1996; Gibson andTabita, 1997;Kusian and Bowien, 1997; Shively et al., 1998).Mutant strains of Rhodospirillum rubrum (Falconeand Tabita, 1993), Rhodobacter sphaeroides (Gibsonet al., 1991), and Ralstonia eutropha (formerlyAlcaligeneseutrophus) (Bowienetal.,1993)defectivefor autotrophic growth continue to uncover newgenes involved in Calvin cycle function andregulation. The structure and regulation of Calvincycle operons and gene clusters has been investigatedin several eubacteria. Among eubacteria, the mostcomplete picture of gene organization exists for the

Ralstonia eutropha and thecyanobacterium Synechocystis PCC6803. Thestructural organization ofCalvin cycle genes in theseorganisms could not possibly differ more.

Ralstonia eutropha possesses the largest cbboperon characterized to date (Bowien et al., 1993;Bommer, 1996). With two exceptions (ribose-5-phosphate isomerase and triosephosphate isomerase)it encodes the entire pathway, and is transcribed asone polycistronic mRNA under the regulation ofCbbR (Windhvel and Bowien, 1991), a member ofthe LysR family of transcriptional regulators (Tabita,1994). The opposite extreme is realized in Syne-chocystis, where no two genes for Calvin cycleenzymes occur as neighbors in the genome (Kanekoet al., 1996a, 1996b). The Synechocystis genes arenot separatedbyjustafewhundredorafewthousand


20

bases, they are strewn around the 3.6 Mb genomewith no recognizable pattern whatsoever. Even therbcL/rbcS operon is disrupted, the genes for the twosubunits being separated by an ORF of still unknownfunction, rbcX. Synechocystis possesses two geneshomologous to cbbR, but neither the function of theirproducts are known, nor whether Calvin cycle genesin Synechocystis form a regulon. Comparatively littleis known about regulation, coordinated or otherwise,of cyanobacterial Calvin cycle genes (Beuf et al.,1994; Li and Tabita, 1994; Gibson and Tabita, 1996;Xu and Tabita, 1996).

In Xanthobacter flavus, cbb genes are distributedacross at least two operons, the gap-pgk cluster is notcontiguous with the cbb operon, but it is part of a cbbregulon under CbbR control (Meijer et al., 1996). Asecond cbb operon is present on a large plasmid inRalstonia that is nearly identical to the chromosomaloperon (Bowien et al., 1993). The cbb operons studiedfrom Ralstonia Rhodobactersphaeroides, Rhodobacter capsulatus, Rhodo-spirillum rubrum, Xanthobacter flavus, and Nitro-bacter vulgaris (all show very littleconservation of gene order across species, other thanthe fact that cbbR is usually transcribed, as inRalstonia, on the opposite strand from a divergentpromoter (Gibson and Tabita, 1996). Given thedispersed nature of the Synechocystis genes, it isconceivable that either these operons were assembledin independent lineages from an ancestrally dispersedstate, or that fragmentation of an ancestral operonhas occurred in Synechocystis, accompanied byrearrangements in proteobacteria. Visible rearrange-ment of cbb genes in suggests thatconsiderable structural reorganization of cbb operonshas occurred in these genomes during evolution.There have been reports of Calvin cycle specificactivities in halophilic archaebacteria (Rawal et al.,1988; Altekar and Rajagopalan, 1990; Rajagopalanand Altekar, 1994) but the enzymes have not beencharacterized in detail.

A general picture of higher level control of signaltransduction and gene regulation for the Calvin cycleand its integration into the general metabolism ofphotosynthetic eubacteria is beginning to emerge.The presumably top level of hierarchy involves theRegA/RegB (PrrA/PrrB) two-component sensor-kinase system (Joshi and Tabita, 1996). This systemappears to integrate the control, expression andfeedback of regulons for photosystem biosynthesis(Sganga and Bauer, 1992; Eraso and Caplan, 1994;

Mosley et al., 1994; Allen et al, 1995), nitrogenmetabolism (nif-system), and fixation (Qianand Tabita, 1996; Joshi and Tabita, 1996). Theregulatory cascade is apparently influenced by theredox state of the cells, the level of oxygen, and thepresence of various carbon and nitrogen sources(Joshi and Tabita, 1996). Since precisely these factors(redox state, oxygen, carbon and nitrogen) are knownto have dramatic and, in some cases, interdependenteffects on plant metabolism and nuclear geneexpression (Turpin and Weger, 1990; Chen et al.,1993; Escoubas et al, 1995; Kozaki and Takeba,1996; Wingsle and Karpinski, 1996; Karpinski et al.,1997), these findings from bacterial systems mayhave a degree of model character for understandingrelated phenomena in eukaryotic systems, where themolecular basis of regulation is less thoroughlyunderstood. Although it seems unlikely at first sightthat these same prokaryotic molecular componentswill be found to be involved in plant signaltransduction, the principles of regulatory responseimplemented by eukaryotic signaling/regulationmachinery may ultimately prove to be very similar. Aremarkable study recently provided strong evidencethat some components for rapidly transducing redoxsignals in higher plants perceive the redox state ofthe plastoquinone pool directly in the thylakoidmembrane (Pfannschmidt et al., 1999). Whether ornot nuclear encoded bacterial two-componentsystems, which are still encoded in some chloroplastgenomes (Stoebe et al., 1998; Martin et al., 1998),might be involved in such processes, is an attractivequestion.

Regulation of the Calvin cycle enzyme expressionhas often been monitored using Rubisco as a markerfor the pathway. PRK, TKL RPI and RPE may fulfillthe same purpose, because they appear to be localizedin chloroplasts exclusively as well (Schnarrenbergeret al., 1995). The other enzymes may fulfill functionsin other pathways. For example PGK, GAPDH, TPI,aldolase and FBPase are involved in the gluconeo-genetic and partially in the glycolytic reactionsequence, and the oxidative pentose phosphatepathway relies on many enzymes of the Calvin cycle(Schnarrenberger et al., 1995). Expression studies of

IV. Calvin Cycle Expression in Plants

A. Quantification of Activities


21

Calvin cycle enzymes that possess cytosolichomologues require not only measurement of totalactivities in crude extracts but also quantification ofthe amount of activities attributable to chloroplastand cytosolic isoenzymes. This is particularlynecessary for the gluconeogenetic enzyme activitiesPGK, GAPDH, TIM, aldolase and FBPase isoen-zymes, which may be separated by ion-exchangechromatography but not by gel filtration. In specialcases, it is also possible to distinguish cytosol- andchloroplast-specific activities by virtue of theirdifferent substrate specificity for NADH and NADPH,as in the case of GAPDH, or by their differentialresponse to pH, sulfhydryl reagents, and as forFBPase. But compartmentation of plant carbohydratemetabolism is not an evolutionarily conservedproperty across species (Schnarrenberger et al., 1990),and within a given plant it varies across developmentalstages and tissues. Worse yet, across species,completely different enzymes are involved that mustbe assayed by different means, for example class Iand class II Calvin cycle aldolase (Gross et al., 1994;Plaumann et al., 1997; Flechner et al., 1999). Thus,Rubisco is a valuable marker for regulation of Calvincycle expression, but other enzymes may show verydifferent regulation patterns, and sweeping general-izations to the effect that, beyond the Calvin cycle,plantshave this, that or the other pathway of sugarphosphate metabolism in this or that compartmentare not possible.

The chloroplast activities of PGK, GAPDH,aldolase and FBPase in green leaves usually accountfor about 90% of the total activity (Heber et al., 1963;Latzko et al., 1974; Krger and Schnarrenberger,1983; Schnarrenberger and Krger, 1986; Lebherzet al, 1984; Kpke-Secundo et al., 1990; McMorrowand Bradbeer, 1990), chloroplast TPI accounts foronly about 50% of the total activity (Kurzok andFeierabend, 1984). For other isoenzyme activities ofstarch metabolism and the oxidative pentosephosphate pathway in green leaves, the cytosolicisoenzyme appears to account for most of theprevalent activity (Schnarrenberger, 1987). It appearsthat the activities of the regenerative part of theCalvin cycle (RPI, RPE, TKL) may not requireisoenzyme separation in most cases, since they areprobably located exclusively in the chloroplasts(Schnarrenberger et al., 1995), except in somespecialized tissues and species (e.g. TKL inCraterostigma: Bernacchia et al., 1995). For thesethree enzymes no chloroplast/cytosol isoenzymes

can be separated in spinach leaves (Schnarrenbergeret al., 1995). It is well known that various Calvincycle enzymes vary considerably in their maximalactivities among higher plants, various algae andeubacteria (Smillie, 1963; Heber et al., 1967; Latzkoand Gibbs, 1968; Latzko and Gibbs, 1969; Kelly andLatzko, 1979), as do activities in other pathways.

B. Expression Studies of Enzyme Activitiesand Transcription

The literature on expression of Calvin cycle enzymesis vast. One of the most widely studied aspects is theincrease of enzyme activity and mRNA levels inresponse to light. The influence of light on expressionof genes involved in photosynthesis has been reviewed(Chory et al., 1996; Kloppstech, 1997). Phytochromes(Schopfer, 1977; Pratt, 1995), blue light, and UV-receptors play important roles in this regulation, thatultimately reaches genes for many Calvin cycleenzymes. It is also well-known that the glycolytic,cytosolic isoenzymes of several Calvin cycle activitiesare, as a rule, not responsive to light and are inducedunder anaerobic conditions (Sachs, 1994; Kennedyet al., 1992).

Complete cDNAs have been characterized forseveral Calvin cycle enzymes from several sources,and all of the Calvin cycle enzymes from spinachchloroplasts have been cloned (Flechner et al., 1996;Martin et al., 1996a). Rubisco gene expression hasbeen studied in by far the greatest detail of all of theCalvin cycle enzymes. Transcription factors involvedspecifically in rbcS gene expression, e.g. GT-1 (Lamand Chua, 1990; Sarokin and Chua, 1992) and GT-2(Gilmartin et al., 1992) have been characterized. Ingeneral, expression of Calvin cycle genes in plants,particularly in etiolated seedlings, is stimulated bylight. This can occur through elevated transcription,or, as recent studies of the Cen gene in Chlamy-domonas mutants have shown, post-transcriptionallyat the level of mRNA stability (Hahn et al., 1996).Various cis elements have been described for Calvincycle genes from different sources, including theWF-1 element upstream ofthe genes for SBPase andFBPase ofwheat (Miles et al., 1993), the Gap andAEboxes upstream of the Arabidopsis GapA and GapBgenes (Conley et al., 1994; Kwon et al., 1994; Park etal., 1996), and an octameric motif in the first intronof the maize GapA1 gene (Donath et al., 1995;Khler et al., 1996). The FBPase promoter alsocontains a DNA binding site for the GT-1 factor


22

which mediates light activation ofexpression throughphytochrome in promoters of oat and rice (Lloyd etal., 1991b). In addition to the small subunit ofRubisco, for which numerous gene structures areknown (Wolter et al., 1988; DeRocher et al, 1993;Fritz et al., 1993), several higher plant Calvin cyclegene structures have been characterized. Theseinclude GapA from maize (Quigley et al, 1988),Arabidopsis (Shih et al., 1992), and several othersources (Kersanach et al., 1994), SBPase and FBPasefrom wheat (Raines et al., 1988, 1992), aldolasefrom rice (Tsutsumi et al., 1994) and Chlamydomonas(Pelzer-Reith et al., 1995), and PRK from wheat(Lloyd et al., 199la).

In studies of the transcript levels of all Calvincycle enzymes in various tissues during spinachdevelopment (Henze, 1997), it was observed thatmost mRNAs were present in all green leaf tissue inroughly the same relative quantities, with theexception of rbcS mRNA, that was present at roughly10-fold higher steady-state levels. In etiolatedcotyledons, all mRNA levels were reduced at least10- to 20-fold relative to green leaves. Uponillumination, mRNAs for rbcS, aldolase, SBPaseand PRK increased within 2 h after illumination,followed by the other mRNAs. After 24 h ofillumination, mRNA levels were indistinguishablefrom those in green leaf tissue. It is still too early totell whether genes of the Calvin cycle in higherplants are regulated as a unit, or whether their activityis simply modulated as part of a general greeningresponse of the gene regulatory machinery to lightand redox state. In the red alga Galdieria sulphuraria,that can grow heterotrophically or autotrophically(Gross et al., 1999), isoenzymes of many Calvincycle activities (aldolase, RPE, PGK, FBPase, andGAPDH) are specifically induced during thetransition from heterotrophic to autotrophic growth(J. Girnus, C. Schnarrenberger, W. Gross, unpub-lished)

There are far too many reports involving expressionstudies of Calvin cycle genes and enzyme activitiesto permit thorough review. In Table 1 we have tried toprovide access to some of that literature, includingmany studies that cannot be found by computer-searching (and omitting many studies that can).Rubisco is well known for its inducibility byphytochrome. The effects of phytochrome and otherlight receptors on the remaining Calvin cycle enzymeshave been studied in less detail and in many of theearly studies, the quantification of chloroplast vs.

cytosol enzyme activities was not considered. Table1 is by no means complete, but we hope that readersfind parts of it useful.

C. Gene Regulation Through High SugarSensing, and Redox State

A sugar-sensing system has been discussed in plantsthat may be able to significantly influence geneexpression (Sheen, 1990, 1994; Koch, 1996; vanOosten and Besford, 1996; Jang and Sheen, 1997;Chapter 10, Graham and Martin). Sugars like glucose,fructose and sucrose cause strong repression of genesfor photosynthetic functions, resulting in e.g.reduction of photosynthetic pigments and Calvincycle enzymes while other sugars are largelyineffective. Glucose feeding can reduce the steady-state mRNA levels of several Calvin cycle genes inwheat, including FBPase, SBPase, PGK and rbcS(Jones et al., 1996). Among the Calvin cycle enzymes,activity and protein of Rubisco decline steadily withinseveral days. This was demonstrated in cell suspensioncultures of Chenopodium rubrum and in intacttobacco and potato leaves which were cold-girdledfor 12 h to reduce assimilate export (Krapp et al.,1993), detached spinach leaves fed with glucosethrough the petioles (Krapp et al., 1991), mesophyllcells of tobacco (Criqui et al., 1992) and by comparinggreen and bleached leaves of transgenic tobaccoplants expressing a yeast-derived invertase in theapoplast (Stitt et al., 1990). In some ofthese systemsit was shown that glucose treatment resulted in therepression of these and other photosynthesis-relatedgenes. In cell suspension cultures of Chenopodiumrubrum, rbcS mRNA levels are reduced within severalhours and run-on experiments with isolated nucleiindicated that also the synthesis of rbcS is reduced,as is incorporation into the Rubiscoprotein, indicating an inhibition of de-novo synthesis(Krapp et al. 1993). In glucose-fed tobacco protoplastsand leaf discs, rbcS transcript levels are reducedwithin hours upon glucose addition, but rbcLtranscript levels are reduced at a much slower rate(Criqui et al., 1992). Thus, regulation takes placeprimarily at a transcriptional level. Information onglucose repression of other Calvin cycle enzymes islimited to FBPase and GAPDH, both of which, likeRubisco, show similarly declining activities in thepresence of glucose (Stitt et al., 1990; Krapp et al.1991, 1993).

Elevated (about 1000 ppm) can also influence


23

gene expression (reviewed by von Oosten andBesford, 1996), including those for some Calvincycle enzymes. For example, whole tomato plantsgrown at elevated relative to ambientgrown plants, for 10 days showed reduced Rubiscoactivity in the second half of this period, probablyaccounting for the long-term decline in photosyntheticefficiency under high (Yelle et al., 1989). rbcStranscript levels are greatly reduced in tomato plantswithin 4 days, while rbcL transcript levels declineless pronounced. This effect was enhanced in detachedleaves, indicating repression by elevated internalglucose levels (van Oosten et al., 1994). On thecontrary, low levels of resulted in an overex-pression of rbcS (Krapp et al., 1993). Besides Rubisco,also the activities ofPGK and GAPDH were reducedunder elevated condition, though only in fullydeveloped leaves (Besford, 1990). However, Krappet al. (1991) observed less inhibition at highthan under ambient at saturating irradiation andeven less under low irradiation. It should be notedthat any inhibition seen under these conditions mightalso be attributable to nitrogen limitation that canbecome apparent at increased growth rates (Kozakiand Takeba, 1996).

Van Oosten and Besford (1995) showed thattranscript levels of plastid-encoded rbcL and othergenes involved in photosynthesis (psbA, psaAB) werereduced in mature leaves by elevated Expressionof nuclear genes associated with the Calvin cyclesuch as Rubisco activase are also reduced by elevated

(van Oosten et al., 1994). Nuclear encoded rbcStranscript leaves were reduced in tomato plantsexposed to high as were plastid-encoded rbcLtranscript levels, though less markedly, and theseeffects could be simulated by sugar feeding (vanOosten and Besford, 1994). In terms of Rubiscocontent, tomato plants responded to high in amanner similar to plants grown with low nitrogensupply (van Oosten et al., 1995). In bird-cherry treesgrown under conditions where nutrients were notlimiting, Rubisco activity decreased in response tohigh (Wilkins et al., 1994). Clearly, there is aninterdependence between availability, nitrogen,redox state, sugar levels and light levels that influencegene expression. An Arabidopsis mutant defective ina gene that might be involved in integrating ortransducing sugar-related signals was recentlydescribed (van Oosten et al., 1997). Furthermore, theredox state of the thylakoid membrane itself has beenrecently shown to regulate the transcription of plastid

genes involved in maintaining redox balance(Pfannschmidt et al., 1999), a process that certainlyentails the Calvin cycle as the primary means forregenerating The question of which andhow many signaling pathways are involved inmaintaining redox balance in a manner that affectsthe Calvin cycle is still open.

D. Regulation in Specific Systems

A system involving preferential breakdown of 70Schloroplast ribosomes on Calvin cycle enzymes wasused in early studies, because it permitted the site ofenzyme synthesis to be determined long before thecoding capacity of chloroplast genomes had beendetermined (Feierabend and Schrader-Reichhardt,1967). Nuclear-encodedenzymes are still synthesizedon 80S ribosomes and imported to chloroplasts underpermissive low (22 C) temperatures or non-permissive high (32 C) temperatures. Among theenzymes of sugar phosphate metabolism assayed,GAPDH, PGK, TPI, TKL, FBPase, RPI, PRK andaldolase were recovered in chloroplasts at non-permissive conditions (Feierabend and Brassel, 1977;Feierabend, 1979, 1986; Feierabend and Gringel,1983; Kurzok and Feierabend, 1983,1986; Otto andFeierabend, 1989), however, Rubisco was absent(Feierabend, 1979), transcripts of Rubisco wererepressed as well (Winter and Feierabend, 1990).

Another well studied system of Calvin cycleexpression is green and white leaf tissue of thealbostrians mutant of barley. This mutant shows avariegated pattern of white and green striped leaveswith non-Mendelian inheritance (Hagemann andScholz, 1962). Rubisco, GAPDH, aldolase, andFBPase were strongly reduced in white leaf tissue(Brner et al., 1976; Bradbeer and Brner, 1978;Boldt et al., 1992). In contrast, the cytosoliccounterparts of the Calvin cycle enzymes, theenzymes of starch metabolism and the key enzymesof the oxidative pentose phosphate pathway werevirtually unchanged (Boldt et al., 1992). Transcriptsof Rubisco were totally repressed in white tissue butwere enhanced in green tissue through phytochrome(Hess et al., 1991). The transcripts of chloroplastPRK, GAPDH, PGK, aldolase, and FBPase wererepressed in white tissue, while those of cytosolicGAPDH and PGK were slightly enhanced (Hess etal., 1993, 1994; Boldt et al., 1994). The phenomenonis interpreted as the action of a plastid derived factoror signal which represses many (but not all) nuclear-


24 William Martin, Renate Scheibe and Claus Schnarrenberger

25Chapter 2 Calvin Cycle

26

encoded plastid proteins in the nucleus (Boldt et al.,1990; Hess et al., 1994; Hess et al., 1997). Thisfactor/signal also represses most genes of theglycolate pathway in plastids and peroxisomes,indicating functional unity of repression ofphotosynthetic functions (Boldt et al., 1997). Hahnet al. (1996) recently described a nuclear gene inChlamydomonas reinhardtii that posttranscriptionally

affects mRNA levels for several chloroplast proteins,including RPE.

Expression ofCalvin cycle genes has been studiedin the facultative CAM plant Mesembryanthemumcrystallinum (iceplant). GAPDH mRNA accumulatesin response to salt stress (Vernon and Bohnert, 1992;Vernon et al., 1993), whereby PRK expression isreduced by salt stress (Michalowski et al., 1992).


27

Transcript levels for cytosolic GAPDH increaseduring the transition from C3 to CAM metabolism(Ostrem et al., 1990). In C4 plants, only the bundlesheath cells contain a complete set of Calvin cycleenzymes (see also Chapter 18, Furbank et al.).

Enzymes of the Calvin cycle can also be found inplastids of non-green tissues. An extreme example isthe endosperm tissue of developing and germinatingcastor bean which never greens (Plaxton, 1996).Many of the plastidic isoenzymes of the Calvin cycleare present in this tissue and show an expressionpattern following the typical fat-to-sugar conversionwith maximum activities 5 days after germination(Nishimura and Beevers, 1981). The question iswhether these enzymes really function in fixationor whether they are due to a leaky expression.Alternatively, they could function in the oxidativepentose phosphate cycle which can provide NADPHfor nitrate reduction. This has been implied by workof Emes and Fowler (1978) on TKL and transaldolase.A cytosolic class I aldolase with specificity for bothF1,6BP and Su1,7BP was found in carrot storageroots but the plastidic homolog was missing(Moorhead and Plaxton, 1990). A general consider-ation ofmetabolism in chromoplasts was summarizedby Camara et al (1995).

E. Calvin Cycle Enzymes and Expression inEuglena gracilis

Euglena gracilis can grow autotrophically andheterotrophically on various substrates (Kitaoka etal., 1989; Brandt and Wilhelm, 1990). A first screenfor cytosolic and chloroplast enzyme activities ofsugar phosphate metabolism was presented by Smillie(1963). Enzyme levels related to photosynthesis anddegradative reactions like glycolysis seem to beantagonistically regulated under autotrophic andheterotrophic growth conditions, respectively. Mostenzyme activities ofthe Calvin cycle increase duringgreening though to various degrees and decreaseafter transfer to heterotrophic conditions (Latzkoand Gibbs, 1969; Kitaoka et al., 1989).

Enzyme activities in Euglena gracilis may also beregulated by light. This phenomenon was dissectedinto a blue- and a red-light reaction by the use ofmutants (Schmidt and Lyman, 1974): In wild-typecells blue light increased the activities of Rubisco,PRK, chloroplast GAPDH, and chloroplast aldolasetwice as effectively as red light. Mutant Y9ZNa1Lhad no chlorophyll and showed a blue-light but no

red-light effect. Mutant Y11P22DL had smallamounts ofchlorophyll and showed the same activityin red and blue light. Mutant W14ZNa1L had nochloroplasts and no Rubisco but the same activity ofGAPDH, PRK and aldolase as dark-grown wild-type cells, but this activity was not increased by light.In other mutants (W3BLU and W8BHL) with noplastid DNA small amounts of cytosolic class Ialdolase were recorded (Karlan and Russell, 1976).Chloroplast development in Euglena gracilis issensitive to glucose repression. When heterotrophiccells are transferred to autotrophic conditions,chloroplast development starts from proplastids. Thepresence of glucose inhibits greening and synthesisof Rubisco (Reinbothe et al., 1991a). Duringdedifferentiation from chloroplasts to proplastids,on the other hand, Rubisco synthesis ceasedimmediately upon transfer to the dark in the presenceof glucose (Reinbothe et al., 1991b).

Chloroplast and cytosolic GAPDH are both presentin Euglena gracilis cells, have been purified andshow no immunochemical cross-reaction (Grissonand Kahn, 1974; Theiss-Seuberling, 1984). Chloro-plast GAPDH is activated by dithiothreitol and/orthioredoxin (Theiss-Seuberling, 1981). Duringchloroplast development of Euglena gracilis in thelight NADP-GAPDH increases in activity (Hoven-kamp-Obbema and Stegwee, 1974). Both enzymesare encoded by nuclear genes and possess severalunusual sequence attributes (Henze et al., 1995).

The aldolase isoenzymes of Euglena gracilisbelong to the class I and class II type and arecompartmented in the chloroplasts and in the cytosol,respectively (Rutter, 1964; Mo et al., 1973; Pelzer-Reith et al., 1994b; Plaumann et al., 1997). Underautotrophic conditions, the chloroplast enzyme ismore active than the cytosolic enzyme (Mo et al.,1973; Karlan and Russel, 1976). This pattern isreversed during growth under heterotrophic condi-tions (Mo et al., 1973). While chloroplast andcytosolic aldolase of higher plants differ little in theirbiochemical parameters (Anderson and Pacold, 1972;Buckowiecki and Anderson, 1974; Krger andSchnarrenberger, 1983; Lebherz et al., 1984), thecytosolic class II aldolase of Euglena has a muchhigher value and a broader pH optimumthan class I aldolase (Pelzer-Reith et al., 1994b).Both aldolases of Euglena gracilis show endogenousrhythmicity in the light and in the dark (Pelzer-Reithet al., 1994a; Malik, 1997). The expression oftranscripts and the enzymes appears to be regulated


28

posttranscriptionally (Malik, 1997). Chloroplast andcytosolic isoenzymes of TPI were separated fromEuglena gracilis (Mo et al., 1973). The chloroplasttype A enzyme is high in autotrophic and low inheterotrophic cells while the cytosolic type Bisomerase predominates under heterotrophic growthconditions. An antagonistic regulation underautotrophic and heterotrophic growth conditions isalso implied for the chloroplast and cytosolic FBPasewhen measured at pH 8.5 and 6.9, respectively (Latzkoand Gibbs, 1969).

Structure and expression of nuclear genes forchloroplast proteins in Euglena are unusual in severalrespects. Many encode polyprotein precursors ofchloroplast proteins (Houln and Schantz, 1988,1993). An example of such a polyprotein is rbcS inEuglena. The nuclear gene is transcribed as an mRNAencoding eight nearly identical concatenate smallsubunits that are translated as a 140 kDa cytosolicpolyprotein, all eight subunits are imported intochloroplasts with the aid of a single transit peptide,and then proteolytically processed from thepolyprotein into individual subunits for Rubiscoassembly (Chan at al., 1990). This unusual polyproteinorganization appears to be restricted to some nuclear-encoded genes in protists of secondary symbioticorigin, i.e. protists that acquired their plastids byengulfing photosynthetic eukaryotes rather thanprokaryotes. [This notion was first suggested forEuglena (Gibbs, 1978) and subsequently demon-strated to be the case for several photosyntheticprotists (Maier, 1992; McFadden et al., 1994;Melkonian, 1996; Van de Peeret al., 1996; McFaddenet al, 1997)]. Such plastids are surrounded by threeor more membranes instead of two, and precursorimport is therefore more complex, involving ER-processing of a signal peptide prior to chloroplastuptake in the case of Euglena (Kishore et al., 1993;Sulli and Schwartzbach, 1996). It is likely that anumber of Euglenas nuclear genes for chloroplastproteins stem from the secondary symbiont and weretherefore transferred twice in evolution: once fromcyanobacteria to the chlorophyte nucleus, and oncemore from the chlorophyte nucleus to the nucleus ofthe Trypanosoma-like host (Henze et al., 1995;Plaumann et al., 1997). Euglenas nuclear genes forCalvin cycle GAPDH (Henze et al., 1995), aldolase(Plaumann et al., 1997), TKL and TPI (W. Martin,unpublished) are not encoded as polyproteins,indicating that this unusual organization is restrictedto certain transcripts. Euglenas nuclear genes for

rbcS and cytosolic GAPDH contain a novel class ofhighly structured introns that have not been describedfrom any other eukaryotes (Henze et al., 1995; Tessieret al., 1995). Also, spliced leader sequences arefound at the 5' end of many of Euglenas nucleartranscribed mRNAs (Tessier et al., 1991). Suchspliced leaders have been implicated in the RNA-processing ofpolycistronically transcribed eukaryoticoperons found in the euglenozoan lineage and inCaenorhabditis (Hirsch, 1994).

In other photosynthetic protists, little is known atthe molecular level about Calvin cycle enzymes andgene structure, but this can be expected to change inthe future since these organisms are turning up quitea number of surprising findings. For example thedinoflagellates Gonyaulax (Morse et al., 1995) andSymbiodinium (Rowan et al., 1996) use class IIRubisco in their Calvin cycle. Moreover, those class IIRubisco genes are nuclear encodedand inSymbiodinium as a polyprotein, as in the case ofEuglenas rbcS. Other photosynthetic protists seemto lack chloroplast- and cytosol-specific isoenzymesof sugar phosphate metabolism. Chlamydomonasreinhardtii is an extreme example, since this alga hasno cytosolic isoenzymes of sugar phosphatemetabolism for at least eight enzyme activities, amongthem aldolase (Schnarrenberger et al., 1994),suggesting that the general compartmentation ofcarbohydrate metabolism may be surprisinglyvariable across protists.

V. Enzyme Interactions and Multienzyme-like Complexes

In 1970, Rutner noted that ...there are now severalwell-documented cases of multi-enzyme complexes(e.g. fatty acid synthase, pyruvic dehydrogenase [...]),there is a tendency to implicate them in othersequential biochemical reactions (Rutner, 1970) anddelineated some straightforward mass-activitystoichiometric difficulties encountered when suchcomplexes are considered in the context ofthe Calvincycle. Since that time, there have been many reportsthat some enzymes of the Calvin cycle may formmultienzyme-like complexes, findings that have oftenbeen discussed in the context of metabolic channelingof intermediates. In enzymological studies prior to1980, these complexes were rarely observed. Themajority of reports deal with complexes isolatedfrom pea and spinach chloroplasts and from green


29

algae, similar associations between Calvin cycleenzymes have not been observed in any cyanobacteriaor photosynthetic proteobacteria. The reports differsubstantially with respect to the number and natureof protein-protein interactions observed. There arestill many open questions in this area, and there iscurrently no consensus concerning the nature,function or significance of such complexes. Variouscomplexes have been isolated by ultracentrifugationin sucrose gradients, by exclusion chromatographyin the presence of stabilizing compounds such asglycerol, and by ion-exchange chromatography. Themultienzyme-like complexes should be distinguishedfrom multimeric forms of individual enzymes whichthemselves can aggregate in purified form, forexample GAPDH (Baalmann et al., 1994), FBPase(Grotjohann, 1997) or RPE (Teige et al., 1998).

Several reports concern complexes consisting oftwo or three enzymes. A complex containing Rubisco,RPI (90 kDa) and PRK (54 kDa) was reported frompea leaves (Sainis and Harris, 1986) that catalyzedR5P-dependent fixation in the presence ofATP,and contained about 45% of RPI and PRK activitiesin the complexed form. In a similar complex fromspinach, 75% of PRK and 7% of RPI were found toassociate and copurify with Rubisco (Sainis et al.,1989). The ratio ofPRK to Rubisco was estimated tobe 1:1 to 1:3. In another report, a complex of PRKwith GAPDH was isolated from Scenedesmusobliquus (Nicholson et al., 1987). The stoichiometrywas estimated to be with a (too low)molecular mass of560 kDa. Also, PRK and GAPDHwere found to form a complex coexisting with thefree enzyme forms in spinach (Clasper et al., 1991).A novel, 12 kDa chloroplast protein (CP12) hasrecently been described from higher plants that shareshigh sequence similarity with the CTE of the GapBsubunit (Pohlmeyer et al., 1996). CP12 interacts withGAPDH in affinity chromatography and with PRKin the yeast two-hybrid system. Under oxidizingconditions, CP12 interacts with both proteins toform ~600 kDa complexes and has been suggested tobe involved in regulation (Wedel et al., 1997; Wedeland Soll, 1998).

A complex was isolated from Chlamydomonasreinhardtii consisting of with anaverage molecular mass of 460 kDa (Avilan et al.,1997; Lebreton et al., 1997), equal to the sum ofindividual masses of the free enzymes. Thedissociation of the complex can be achieved byreducing agents like DTT, NAD(P)H, reduced

ferredoxin or reduced thioredoxin, accompanied byan increase particularly in PRK activity. PRK isinactive in the oxidized form (see Section VI) andgained some activity during complex formation withGAPDH in a manner similar but not identical tochaperonin action (Lebreton et al., 1997). However,this increase corresponds to only a few percent oftheactivity of the reduced form present in the light. Thesubsequent dissociation of the complex by reducingagents causes a conformation change in PRK, another20-fold increase in PRK activity with a 4-folddecrease in and a 2-fold decrease inDuring complex dissociation, GAPDH showed arelative increase in favor of overactivity (Avilan et al., 1997). On the other hand, thecomplex could form spontaneously, upon addition of

or oxidized glutathione (Avilan et al., 1997;Lebreton et al., 1997). The association betweenGAPDH and PRK in Chlamydomonas involves twoenzymes that catalyze non-consecutive steps in thepathway and the complex is present under darkconditions where there should be no Calvin cycleactivity. Thus, the complex is unlikely to be involvedin channeling in the classical sense (Gontero et al.,1994; Ricard et al., 1994).

There have been reports of larger Calvin cyclemultienzyme-like complexes involving severaladditional enzymes. The first such larger complexcontained PRK, Rubisco, PGK, and GAPDH andwas reported by Mller (1972), who recognized thatthe fragile complex is dissociated by NADPH orATPand that the enzymes involved are activated duringdissociation. The complex had a molecular mass of400 kDa, less than the value of 700 to 800 kDaexpected. Sasajima and Yoneda (1974) found thatRPI, TKL and RPE copurify. More recent reportshave detected complexes with an in the range of500 to 1000 kDa (Gontero et al., 1988; Gontero et al.,1993; Rault et al., 1993; Sainis and Srinivasan, 1993;Sss et al., 1993, 1995). The enzymes involved inthese complexes and their stoichiometry differ inindividual laboratories, the function is generallyinterpreted as metabolic channeling.

Gontero et al. (1988) found a complex consistingof Rubisco, RPI, PRK, PGK and GAPDH. Thecomplex was fairly stable and homogeneous duringultracentrifugation. DTT increased the activity ofthe individual enzymes. The complex catalyzedfixation with R5P, ATP, NADPH and Themolecular mass ofthe complex was estimated as 520kDa and the ratio of the individual enzymes inferred


30

to be 2PRK:2GapA:2GapB:2RbcS:4RbcL, inaddition to some RPI and PGK (Rault et al., 1993).Because the molecular mass of the individualenzymes is anticipated to be much larger than that ofthe complex, it was suggested that Rubisco mightexist in an form (Rault et al., 1993), differingfrom that of the crystallized enzyme (Shibita et al.,1996). If the complex is subjected to SDS-PAGE,several protein bands are observed corresponding tothe bands of the individual enzymes.

A complex isolated from spinach containedRubisco, PRK, GAPDH, SBPase, ferredoxin-NADPreductase (FNR) and chaperonin 60 (Sss et al.,1993) and fixed from R5P. The complex wasstable at low salt conditions (250 mMKCl). Ammonium sulfate (1 M) or pH 4.5 completelydissociated the complex. All enzymes ofthe complexwere found almost exclusively attached to the outersurface of thylakoid membranes during goldimmunolabeling except Rubisco, which showed alsostromal localization (Sss et al., 1993a, 1993b; Adleret al., 1993). Similar association with thylakoids hadpreviously been reported for Rubisco (Grisson andKahn, 1974; McNeil and Walker, 1981), PRK (Fischerand Latzko, 1979) and GAPDH (Grisson and Kahn,1974). A newly described ~600 kDa complex fromtobacco contained Rubisco, PRK, RPI and carbonicanhydrase (Jebanathirajah and Coleman, 1998).

In other reports, complexes consisting of PGK-GAPDH (Malhotra et al., 1987; Macioszek andAnderson, 1987; Macioszek et al., 1990), GAPDH-TPI, aldolase-TPI, GAPDH-aldolase (Anderson etal., 1995), and PRI and PRK (Anderson, 1987;Skrukrud et al., 1991) have been found. CytosolicPGK and GAPDH were also found to form abienzyme complex (Weber and Berhard, 1982;Malhorta et al., 1987). Complexes of aldolase-GAPDH, PGK-GAPDH and aldolase-TPI fromchloroplasts were isolated and characterized in pea(Anderson et al., 1995). It has been suggested thatinteraction among GAPDH, TKL and aldolase aroundSBPase may lead to a direct transfer of GA3P amongthese enzymes (Marques et al., 1987). Finally, PRIand PRK were shown to have kinetics in a complexstate that differed from those anticipated for substratesused by non-complexed enzymes (Anderson, 1987).The theoretical kinetics ofCalvin cycle multienzymecomplexes have been modeled (Gontero et al., 1994;Ricard et al., 1994). In multienzyme complexes, thekinetics become increasingly complicated because

of the combined action of several enzymes.Note that the sum of molecular weights of the

native enzymes from spinach chloroplasts shown inFig. 1 is about 1500 kDa, Rubisco alone contributinga third of that. Since Rubisco is far more abundantthan any of the other Calvin cycle enzymes in plastids,it is clear that not all active Calvin cycle enzymes canexist in a complexed state (Rutner, 1970).

Summing up these findings on Calvin cyclemultienzyme complexes, it appears clear thatinteractions between various enzymes do exist, butthere is no consensus on which or how many enzymesinteract and whether the same enzymes interact indifferent species. The metabolic relevance of thesecomplexes is still unclear. Pressing problemsconcerning these associations have yet to be solved.

First, the isolated enzyme complexes are usuallydescribed to dissociate in the presence of reducingthiols (light), but the key regulatory factor of overallCalvin cycle activity is light-mediated (redox)activation through reduced thiols (see below). Thisdiscrepancy is difficult to reconcile with metabolicchanneling, since the kinetic data indicate higherenzyme activities for associated enzymes via transferof substrates in a consecutive reaction sequence, butflux through the pathway in the dark is basically nil(associated state) due to severe down-regulation ofFBPase, PRK and SBPase (and moderate down-regulation of GAPDH) in the dark (i.e. in absence ofreduced thioredoxins). But if the overall activities ofthe complexed enzymes are higher, as many suchstudies indicate, we are left with the question of thephysiological relevance of improved kinetics for(dark-) associated enzyme complexes, since the activeforms are dissociated in the light (see Section VI).

Second, if associations between enzymes are ascritical to Calvin cycle function as the interpretationsof many multienzyme studies would suggest, thenproblems in understanding the pathway ensue whenthe results from antisense inhibition of Calvin cycleenzymes are considered (see section VII). This isbecause antisense studies have shown that most (butnot all) Calvin cycle activities must be reduced onthe order of five- to ten-fold to affect a significantreduction in assimilation rate under normal growthconditions. If the brunt of assimilation occurs incomplexes, limiting any one component should beexpected to have a more drastic effect. Further workis needed to clarify the general significance of theseenzyme association phenomena.


31

VI. Biochemical Regulation in Chloroplasts

Light governs not only Calvin cycle gene expressionin higher plants, it is also the foremost determinantof enzyme activity, and hence flux through thepathway (Buchanan, 1980; Wolosiuk et al., 1993).Light regulation of the Calvin cycle is achieved bymodulation of enzyme activity of four enzymesthrough the ferredoxin/thioredoxin system: FBPase,SBPase, PRK and GAPDH (Buchanan, 1991). Thefirst three enzymes are obvious targets for regulation,since they catalyze reactions that are irreversibleunder physiological conditions, their regulatoryprinciple is the same: reduced activity as a result ofoxidation of regulatory cysteines by in the dark,full activity through reduction of regulatory cysteinesby reduced thioredoxin in the light. Activity iscontinuously adjusted in the light through continuedreoxidation and reduction (thioredoxin). Thesteady-state between these interconvertible enzymeforms is individually influenced by metabolites(Scheibe, 1990). The fourth reaction, catalyzed byGAPDH, is reversible. Its activity is dependent uponits state of aggregation, redox modulation being theprerequisite enabling this metabolite-inducedinterconversion under physiological conditions.

A. The Ferredoxin/Thioredoxin System

Ferredoxin (Fd) reduced by photosynthetic electronflow provides electrons for reduction via Fd/NADP reductase, for nitrite reduction via nitritereductase, for sulfite reduction via sulfite reductase,for the reductive generation of glutamate fromoxoglutarate via glutamate synthase (GOGAT), andfor the reduction of the thioredoxin (Td) viaferredoxin-thioredoxin reductase (FTR) (reviewedby Buchanan, 1980; Woodrow and Berry, 1988;Scheibe, 1990; Buchanan, 1991; Knaff and Hirasawa,1991; Wolosiuk et al., 1993; Jacquot et al., 1997b).FTR is composed of two different subunits, subunitA is rather variable between organisms, subunit B ismore highly conserved and contains an Fe-S clusterin addition to conserved cysteines involved in redoxtransfer, but it is not a flavoprotein (Tsugita et al.,1991; Falkenstein et al., 1994). Thioredoxins aresmall heat-stable proteins that occur in all organismsand in many compartments. In the chloroplast variousisoforms occur that differ in their primary structuresand specificities: Tdm, and (reviewed by

Eklund et al., 1991). Invitro,Tdm primarily activatesNADP-dependent malate dehydrogenase (NADP-MDH), and inactivates chloroplast glucose-6-phosphate dehydrogenase (G6PDH), whilepreferentially activates chloroplast FBPase, SBPase,PRK, GAPDH in addition to the chloroplast couplingfactor CF1 (reviewed by Buchanan, 1991). Whetherthis pattern of specificities also holds in stroma,where the protein concentration is very high, remainsto be established.

Traditionally, light/darkmodulationofchloroplastenzymes was considered as an all-or-nothing on/off-switch, but more recently it has become apparent thatit is also a means to fine-tune enzyme activities in thelight (Scheibe, 1990, 1991,1995). This is becausepresent at high concentrations in the chloroplast(Steiger et al., 1977) continuously reoxidizes thecysteines generatedbythioredoxin-mediatedelectronflow to the target enzymes. Light-modulated enzymesthus exist as two interconvertible enzyme forms thatare subject to covalent modification (reduction andreoxidation of cystine/cysteine residues), comparableto those enzymes that are subject to proteinphosphorylation/dephosphorylation (Scheibe, 1990).In both cases, energy is consumed to drive the cyclebetween the two forms, but in the light, energy in theform of reducing equivalents is abundant and posesno significant drain on the photosynthetic membrane.

Target enzymes as well as the chloroplastthioredoxins are characterized by the very negativemidpoint redox potentials of their regulatory cysteines(Faske et al., 1995). For NADP-MDH, FBPase andPRK these are around 380 mV, similar to that of thenonphysiological reagent dithiothreitol (DTT), andeven more negative than the value of 350 mV forTdm and (Gilbert, 1984). These redox potentialsare all more negative than those of NADP(H) (320mV), and of glutathione (-260 mV), indicating thatthese protein thiols cannot be reduced by cellularreductants other than reduced ferredoxin. In somecases, mixed disulfides can be formed with lowmolecular weight thiols such as glutathione(Ocheretina and Scheibe, 1994). That certainchloroplast proteins occur in an oxidized form is arather special attribute, since usually only extracellularproteins tend to exhibit this property (Fahey et al.,1977). For chloroplast enzymes it is this specificproperty which is the basis for a very flexibleregulatory system.


32

B. Target Enzymes

The light/dark-modulated chloroplast enzymes arecharacterized by their unusually negative redoxpotentials. As a result of this, they are only in thereduced state when electrons of very negative redoxpotential from ferredoxin are available in the light;otherwise they relax to their oxidized state. Theredox potentials themselves, however, are subject tochange by specific metabolites, mostly the substrateor the product of the respective enzyme reaction.These metabolites are also known to act as effectorsofthe reductive and/or the oxidative part ofthe redoxcycle. At equilibrium (with the redox buffer, in vitro)or at steady-state (in vivo) changes in the relativeeffector concentrations result in a more or lesspronounced shift of the ratio between oxidized andreduced enzyme form (in a concentration-dependentmanner) (Faske et al., 1995). Redox-modulatedchloroplast enzymes generally exhibit high simi-larities with their non-redox-modulated homologuesfrom other sources, but also tend to possess unique,cysteine-bearing sequence motifs that are responsiblefor their regulatory properties (Scheibe, 1990).

Chloroplast FBPase is the classical target for lightregulation via thioredoxin 1980). Thereis a strong dependence of FBPase activity upon theF1,6BP concentration, i.e. FBPase cannot easily beactivated by DTT (or in the light) in the absence ofF1,6BP. The resulting regulatory pattern is a strictfeedforward mechanism ofFBPase activation due toincreasing F1,6BP levels (Scheibe, 1991). Severalstudies have investigated the mechanism of activationusing FBPase overexpressed in E. coli (Jacquot et al.,1995; Hermoso et al., 1996; Jacquot et al., 1997a;Lopez-Jaramillo et al., 1997; Sahrawy et al., 1997).Chloroplast FBPase possesses a conspicuousinsertion of 1215 amino acids in the central regionof the primary structure with two conserved cysteineresidues separated by five amino acids andpreceded by a third conserved cysteine furtherN-terminal (Marcus et al., 1988; Raines et al., 1988).In vitro mutagenesis of and results inenzymatically active FBPase enzymes that can nolonger be regulated by thioredoxin, indicating thatthese may be specific targets of thioredoxin regulation(Jacquot et al., 1995). But in a more recent study,was also found to be responsible for redox dependence(Jacquot et al., 1997a). Replacing these three cysteineswith serine residues in rapeseed FBPase also resultedin enzymes that were active in a manner largely

independent of

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