Studieso n Lipoamide Dehydrogenase

CENTRALE LANDBOUWCATALOGUS

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Promotor : Dr.C .Veege r hoogleraar in de Biochemie

Co-promotor : Dr.A .d e Kok universitair hoofddocent JacqiiesA .E .Bene n

Studies on Lipoamide Dehydrogenase

Proef^chrift ter verkrijging van de graad van doctor in de landbouw- en milieuwetenschappen op geaagva n rector magnificus, Dr. H.C .va nde r Plas, in het ppenbaar te verdedigen op vrijdlag 8 mei 1992 des narniddagst e 13.30 uur in de Aula van daj Landbouwuniversiteit te Wageningen. Aan Anja Aan mijn moeder Ter nagedachtenis van mijn vader fj^optf^ ;v^

Hetverdien t aanbeveling deter m conservatief in relatie tot aminozuursubstituties met meeijomzichtighei d te benaderen.

Dit proefschrift, hoofdstuk 2-4.

De ejenvoudige manier om redoxpotentialen van flavoproteinen te bepalen zoals voorjesteld door Massey, werkt alleen voor eenvoudige flavoproteinen.

Massey, V. (1990) in Flavins and Flavoproteins 1990 (Curti,B. , Ronchi,S .an d Zanetti, G. eds) pp55-66 ,Walte rd e Gruyteran dCo ,Berli n - NewYork .

De uit de enzymologie bekende 'negatieve cooperativiteit' blijkt ook bij de Europese unififatie opgeld te doen.

Ricard,J .an dCornish-Bowden ,A . (1987) Eur.J. Biochem. 166, 255-272.

Devjoorspellend e waarde van de circulair dichroisme methode voor eiwitstructuren is vooijalsnog niet hoog.

Provencher, S.W . andGlockner , J. (1981) Biochemistry20, 33-37 . j Wishart, D.S. ,Sykes ,B . D.an d Richards, F. M. (1991) FEBSLet. 293,72-80 .

Intggenstellin g tot de bewering van Scrutton etal. isd e afwezigheid van reductie van het flavine in Cys47->Ser gemuteerd E. coliglutathio n reductase door NADPH niette verWachten.

Scrutton, N.S. , Berry ,A. , Deonarian,M . P.an dPerha m ,R . N.(1990 ) Proc. R. Soc. London242, 217-224.

De ^ewering door Borges ef al. dat in Pseudomonasfluorescens ee n tweede gen voor lipoimide dehydrogenase zou zijn aangetoond door Benen etal. i s tout.

Borges,A. , Hawkins,C . F., Packman,L C.an d Perham,R . N. (1990) Eur.J. Biochem. 194, 95-102. Benen,J .A . E.,Va n Berkel,W .J . H.,Va n Dongen , W. M.A . M.,Muller , F.an dD e Kok ,A . (1989) J. GenMicrobiol. 135, 1787-1797.

4oxidases een flavine-N5-sulfiet adduct vormen verdient aanpassing.

Muller,F . (1987) Free RadicalBiology Medicine 3, 215-230 .

De verwachte 5%dalin g van verkeersslachtoffers, ten gevolge van de maatregel overdag verplicht licht te voeren op alie gemotoriseerde vervoermiddelen, is niet gecorrigeerdvoo r de te verwachtentoenam e van het aantal slachtoffers onder motorrijders.

De excessieve veiligheidsmaatregelen waarmee asbest is omgeven zal, gezien de maatregelente n aanzien van AIDS preventie, ertoe leiden dat het afsluiten van een levensverzekering gepaard zai gaan met een asbest-antecedenten onderzoek.

Stellingen behorende bij het proefschrift

'Studies on lipoamidedehydrogenase '

Wageningen, 8 mei 1992. Jacques A. E.Benen . Voonvoord

Graagwi l ik op deze plaats iedereen bedanken die bij de totstandkoming van dit proefs^hrift heeft bijgedragen. Ce^s Veeger, mijn promotor, met wie pittige discussies werden gevoerd over ook 'zijn' efizym. Aart de Kok die alsco-promoto r optrad.Aar t bedankt voor je geduldig oor envoo r de vrijleid die je me hebt gelaten in onderzoek en vorming. WiHem van Berkel bij wie menig wetenschappelijk proeballonnetje werd opgelaten dat va|tk werd doorgeprikt endi e veel heeft bijgedragen aan het hele proefschrift. Adlie Westphal, die het gen coderend voor lipoamide dehydrogenase cloneerde, niet alleen een fijn collega en kamergenoot maar ook een gezellige kerel buiten werktijd. Miriam Gilissen, Fenneke Linker, Erna Hissink, Nicole Dieteren en Sandra Japenga die in het kader van een afstudeervak aan verschillende aspecten van lipoamide dehydrogenase werkten. Zdzislaw Zak who spent three months ontediou stitration s of mutated enzymes and who racordedth e CDspectra . Charles Williams Jr. who invited met o visit his lab inAn n Arbor, Michigan, USA,i n order to be able to do rapid scan analysis. DavidArscot t who made me familiar with the cofriplex apparatus. Charles and Davidthank s a lot. Without this oppertunity, fruitful{cooperatio n anddiscussion s Chapter 3woul d have had less impact. Ihop e the cooperation will be continued inth e future. Walter van Dongen die de correctie van hoofdstuk 5voo r zijn rekening nam. Andrea Mattevi, Bram Schierbeek en Wim Hoiva n de kristallografiegroep van de univer$iteit van Groningen die de structuur beschikbaar stelden voordat deze gepubliceerd was en die aanzienlijk meer mutaties voorstelden dan een promovendus kan maken en bestuderen. Allf medewerkers van de afdeling Biochemiedi e ervoor zorgden dat er altijd een prima|werksfeertj e was en waarbijj e je gal kon spuwen als het weer eens mis ging. Mijfi moeder,famili e en vrienden die altijd gei'nteresseerdware n in de voortgang. Ante,di e nooit klaagde als ikwee r te laat thuis was of in het weekend moest werken. Die me intijde n vantegensla g atijd uit de put hielp, kortom,di e een gewelaige steun voor mewas .Anj a bedankt I Chapters 2 and 5 have been published andchapter s 3 and 4 will be published separately.

Chapter 2: Benen,J. , Van Berkel,W. ,Zak ,Z. ,Visser ,T. ,Veeger ,C . andd e Kok,A . (1991) Eur.J. Biochem. 202,863-872 . Chapter 3: Benen,J. , van Berkel,W. ,Dieteren , N.,Arscott , D.,William sJr. , C, Veeger, C. andd e Kok,A . (1992) Eur.J. Biochem.,(submitted ) Chapter 4: Benen,J. , Van Berkel,W. ,Veeger , Can dd e Kok,A . (1992) Eur. J. Biochem.,(submitted ) Chapter 5: Benen, J.A . E.,Va n Berkel,W .J .H. ,Va n Dongen,W . M.A . M.,Muller ,F . andd e Kok,A . (1989) J. Gen.Microbiol. 135, 1787-1797.

The research described inthi s thesis was carried out atth e Department of Biochemistry, Agricultural University, Wageningen,Th e Netherlands, under the direction of Prof. Dr. C. Veeger. The investigations were financially supported by the Netherlands Organisation for Scientific Research (N.W . O.) underth e auspices of the Foundation for Chemical Research (S.O.N.). Contents

Page

Abbreviations 9

List ofjenzyme s 10

Chapter 1. Introduction 11

Chaptfr 2. Lipoamide dehydrogenase from Azotobacter vinelandii: 43 site directed mutagenesis of the His450-Glu455 diad. Spectral properties of wild-type and mutated enzymes.

Chapter 3. Lipoamide dehydrogenase from Azotobacter vinelandii: 53 site directed mutagenesis of the His450-Glu455 diad. Kinetics of wild-type and mutated enzymes.

Chapter 4. Lipoamide dehydrogenase from Azotobacter vinelandii: 79 Role of the C-terminus in catalysis and dimer stabilization.

Chapter 5. Molecular cloning and sequence determination of the 95 /pd-gene encoding lipoamide dehydrogenase from Pseudomonas fluorescens.

Chaptor 6. Lipoamide dehydrogenase from Pseudomonas fluorescens. 107 Acharacterizatio n of the enzyme.

General discussion 115

Samenvatting 123

Curriculum vitae 131

List of publications 131 Abbreviationi s AAD+ 3-aminopyridine adenine dinucleotide AcPyA0e+ 3-acetylpyridine adenine dinucleotide AcPyApeH 3-acetylpyridine adenine dinucleotide reduced BCOA0C branched-chain oxo acid dehydrogenase complex

Cl2lndi 2,6-dichlorophenolindophenol Cm midpoint concentration of Gdn/HCI required for unfolding CT 530 nmcharg e transfer species. AG standard Gibbs energy Eox oxidized (mutated) lipoamide dehydrogenase

EH2 2-electron reduced (mutated) lipoamide dehydrogenase with disulfide reduced (530 nmcharge-transfer - and tautomeric species). EH- EH2on e step deprotonated E2- EH2tw o steps deprotonated

EH4 general descriptive term for 4-electron reduced (mutated) lipoamide dehydrogenase EDTA ethylene diamine tetraacetate Em midpoint redox potential FAD(Hf) flavin adenine dinucleotide (reduced) FL fluorescent species. FIH- 2 electron reduced species with electrons on the flavin. Gdn/H0l guanidinium hydrochloride h Hill coefficient Hepps^EDTA Hepps/EDTA buffer, pH 8.0 k rate constant kb kilo basepairs kcat turnover rate,|j.mol e product per active site persecond .

Kd dissociation constant K| inhibition constant Km Michaelis Menten constant LipS2 D,L - 6,8 -thiocti c acidamide ; lipoamide

Lip(SH)2 reduced lipoamide /pd-gehe gene encoding lipoamide dehydrogenase NAD(R)+ nicotinamide dinucleotide (phosphate) oxidized NAD(Fl)H nicotinamide dinucleotide (phosphate) reduced Meq (sj>. Molar equivalent(s) Mr relative molecular mass

Nbs2 5,5'-dithiobis(2-nitrobenzoate) nm nanometer PDC pyruvate dehydrogenase complex pKa -log(proton dissociation constant) PPi/EDTA-buffer 50 mMsodiu m pyrophosphate, 0.5 mMEDTA , pH 8.0 PrtX unknown protein encoded by an open reading frame downstream ofth e /pcfgen e in Azotobacter vinelandiian d Pseudomonas fluorescens OGDC 2-oxo glutarate dehydrogenase complex ORF open reading frame SDS sodiumdodecylsulfate

6xSSC 0.9 MNaCI ,0.0 9 MNa 3citrate-2 H20, pH 7.0 SSE 20 %(w/v ) sucrose, 150 mMNaCI , 100 mMEDTA , pH 7.6 SucA gene encoding 2-oxo glutarate dehydrogenase SucB gene encoding succinyl transferase 4n melting temperature U unit of activity: 1umol e product per minute v apparent velocity, nmoles product per mgenzym e per minute V maximal velocity, u.moles product per mg enzyme per minute

List of enzymes

1. lipoamide dehydrogenase, NADH:lipoamide oxidoreductase (EC. 1.8.1.4) 2. DNA restriction endonucleases Alul,SamH1 , EcoR1, Hinc\\,Hind\\\, Kpn\, Notl, Psfl, Sau3A I,Sma\, Sph\, Ssti, (EC 3.1.24.4) 3. DNA polymerase I(Kleno w fragment)(EC 2.7.7.7) 4. DNA ligase (EC6.5.1.1 ) 5. T4-DNA polymerase (EC 2.7.7.7) 6. Glucose-6-phosphate dehydrogenase, D-Glucose-6-phosphate: NADP 1- oxidoreductase (EC 1.1.1.49) 7. Xanthine oxidase, xanthine: oxygen oxidoreductase (EC 1.1.3.22)

10 Chapter 1

Introduction

Occurrence and function of lipoamide dehydrogenase

Lipoamide dehydrogenase is the common name for NADH:lipoamide oxido- reductase (EC 1.8.1.4).Th e enzyme wasfirs t described by Straub [1] and has been knownjfo ryear s as Straub diaphorase as it readily transfers electrons from NADHt o artificial electron acceptors like 2,6-dichlorophenolindophenol and methylene blue [2]. Massey [3] and Searls and Sanadi [4] showed that Straub diaphorase is identical with lipoamide dehydrogenase. Lipoamide dehydrogenase is widely spread through nature as itwi s isolatedfro m many eukaryotic and prokaryotic species (Acomplet e list is given byWilliam s [5,6]) . Iti sa constituen t of several multi-enzyme complexes.Thes e complexes areth e pyruvate dehydrogenase complex [PDC] [7, 8] ,th e a-ketoglutarate dehydrogenase complex [OGDC] [3,4] ,th e branched chain oxo acid dehydrogenase complex [BCOADC] [9] andth e glycine cleavage system [10,11]. Lipoamide dehydrogenase functions also inth e galactose and maltose uptake system of E.coli [12,13.

Multienzyme complexes. In the multi-enzyme complexes in which lipoamide dehydrogenase catalyses the reoxidation of covalently bound dihydrolipoic acid and reduction of NAD+,th e lipoic acid isth ecarrie r of the acylfunctio n and is reduced after the acyl part has beentransferre dt o either CoA (inth e PDC,OGD C and BCOADC) or tetrahyijrofolat e (inth e glycine cleavage system). The lipoic acid (6,8-dithioctic acid) is in allcomplexe s coupled via amide linkage to a lysine of the acyl transferase component [14,15] or of a separate carrier protein inth e glycine oxidation system [16, 17].Thi s isshow n in Fig. 1. The flexible arm formed by lipoic acid and the lysine (the lipoylgroup ) is assumed to carry trie substrate from one active site ofth ecomple x to the other. In allth e complexes afterth e acyl part has beentransfere d to either CoA (inth e PDC,OGD C and BCOADC) or tetrahydrofolate (inth e glycine oxidation system) the lipoyl group is reduced and becorms reoxidized by lipoamide dehydrogenase. It has been shown that in the complexes only the 6R-lipoyl group is present [18]. For the 2-oxo acid dehydrogenase complexes the reaction sequence has been studied extensively [19] and isdepicte d in Fig.2 .Componen t E1 of the complexes isth e 2-oxo aiciddehydrogenas e that catalyses the decarboxylation of the 2-oxo acids and the sut'sequent alkylation of the lipoyl group that iscovalentl y attached to the

11 6S T

0=1 s—s ^i^ 6R

lysine 6R-lipoicaci d Lipoylgrou p

Fig. 1. Structureo flipoi caci dlinke dt oa lysin eresidue .Th easteris k (*) denotesa chira l carbonato mo flipoi caci dyieldin gth e 6R or6 Sconfiguratio n( Lo r Dconfiguratio n respectively).

E2component . The cofactor of component E1i sthiamin e pyrophosphate (TPP). Component E2i sth e acyltransferas e that catalysesth etransfe r ofth e acylgrou p from the alkylated lipoyl group to coenzyme A. The reduced lipoyl group left after release of the acyl group is reoxidized by component E3, lipoamide dehydrogenase. In order to be able to catalyse this oxidation, lipoamide dehydrogenase possesses a non-covalently linked FADan d a redox active active disulfide. The structure ofth e FAD isshow n in Fig.3 .

CoA-SH

R-C-COOH

CO,

NADH+H

NAD+

Fig. 2. Reaction sequence of the 2-oxo aciddehydrogenas e complexes.

12 pH<7 pH>pH>99 0 0 i I I II H2 CH2TICHOHI3—CHjO-pP—O-j-P —0—C

I

kjmitlovin riboftavin (vitomin82 ) hboflovin-5 -monophosphate (FMN)

llavin-adenine-dinucleotide (FAD)

Fig. J3.Structur e of FAD.

Tha reaction sequence forth e glycine oxidation system is presented in Fig.4 .Th e oxidation of glycine results inth e formation of methylene tetrahydrofolate, ammonia and CO2.Tjh e reaction requires four enzymes:a pyridoxal phosphate containing enzyme (P- or P1-protein) , an enzyme carrying the lipoyl group (H-o r P2-protein), an enzyme catalysing the tetrahydrofolate step of the reaction (T- or P4-protein) and lipoamide dehydrogenase (L- or P3 protein).Abbreviate d names are according to [11] and [10] respectively.

NHj.

CH2 ,THF

C02_ SN|. P CH2-COO H [pLpj _ A* HS' ' NH, N y ^w^CHa-THF +NH 3

P/P1 Y H/P2 [lip(SH)2l

\ FAD [PLF=NCH COOH] 2 COOH1 V •S [lipS2 •S L/P3 rFAD

l-SH NADH+H

NAD+

Fig. (4. Reaction sequence of the glycine oxidation system.

13 The role of lipoamide dehydrogenase inth e galactose-, maltose uptake system Isa s yet notclea r [13].

A family of enzymes

Lipoamide dehydrogenase as isolated from all sources investigated sofar is a homodimeric enzyme with one FAD and one redox-active disulfide bridge per monomer. These features are not exclusive to lipoamide dehydrogenase but are characteristic for agrou p of enzymes.A s afamily ,thes e enzymes are knowna sth e pyridine nucleotide disulfide oxidoreductases. Other, well studied members of this family are glutathione reductase [EC 1.6.4.2], trypanothione reductase [EC1.6.4.8], thioredoxin reductase [EC 1.6.4.5]an d mercuric ion reductase [EC 1.16.1.1] Less extensively studied are CoA-glutathione reductase [EC 1.6.4.6],cystin e reductase [EC 1.6.4.1], asparugasate reductase [EC 1.6.4.7], bis-y-glutamylcystine reductase and sulfhydryl oxidase. With the exceptions of mercuric ion reductase and sulfhydryl oxidase, all enzymes inthi s group catalyse electron transfer between NAD(P)(H) and a disulfide/dithiol. Lipoamide dehydrogenase and sulfhydryl oxidase are unique in this group since inth e physiological direction the flow of electrons is from reduced substrate via FADt o NAD(P)+,whil e this flow is reversed inth e other members. Nevertheless lipoamide dehydrogenase readily reduces lipoamide at the expense of NADH [3,4] . For an excellent overview onthi s family of enzymes the reader is referredt o a recent review by C.H.William s Jr.[6] .

Homology among pyridine nucleotide disulfide oxidoreductases

The genes encoding several of the pyridine nucleotide disulfide oxidoreductases have been cloned and sequenced,thu s establishing the primary structures of these enzymes. From different sources sequences of the /pdgene , encoding lipoamide dehydrogenase, were reported.Thes e comprise prokaryotic sequences, from Escherichiacoli [20] , Pseudomonas putida [21-23] and Azotobacter vinelandii[24] ,an d three eukaryotic sequences, human [25],porcin e [25] andyeas t [26,27] .I n Pseudomonasputida thre e /pdgene s were identified,tw o of thethre e /pdgene s encode lipoamide dehydrogenases that function inth e three different 2-oxo acid dehydrogenase complexes [9,28] .Th e third /pdgen e encodes a lipoamide dehydrogenase whose function inth e wild-type organism is not clear yet [23].Als o the genes encoding mercuric ion reductase from different origin were sequenced [29,30 ,

14 31], as were genes encoding glutathione reductase [32],trypanothion e reductase [33] andthioredoxi n reductase [34]. Not only DNA sequences yielded primary structures of the enzymes. Some parts of the amino acid sequences were obtained by peptide analysis long before the DNA-derived sequences became available as for example for lipoamWe dehydrogenase [35, 36],glutathion e reductase [37,] and thioredoxin reductase [38].Thes e partial sequences already showed that aconsiderabl e amount of amino £cid identity exists between the different enzymes. This strong homology was reinforcedb y allth e DNA derived primary structures. Table 1list s the sequence identity.

E.coli P. put. Pig Hum. E.coli Hum. Tn501 Tn21 LD LD LD LD GR GR MR MR

A. vin. LD 40 42 47 48 25 27 29 29 E.coli LD 42 44 40 26 28 29 27 P. put.L D 39 39 24 21 24 24 Pig LD 95 29 28 27 27 Hum. LD 27 28 27 27 E.coli G R 54 27 27 Hum. GR 26 26 Tn501 MR 86

Tabfe 1.Amin oaci dsequenc eidentit y matrixo fsevera ldisulfid eoxid oreductases . LD,fpoamid edehydrogenase ;GR ,glutathion ereductase ;MR ,mercuri c reductase; Hum., human;P. put, P. putida; A. vin., A. vinelandii.

Thijee dimensional x-ray structures. The homology between the enzymes is not only confined to the primairy structures but also extends to the three dimensional structures as revealed by the structures of glutathione reductase [39-43], lipoamide dehydrogenase [44-46],thioredoxi n reductase [47] and mercuric ion reductase [48]. Impressive isth e similarity of the three dimensional x-ray structures of lipoamide dehydrogenase from A. vinelandiian d human erythrocyte glutathione reductase. Although 'only' 27 %o f sequence identity exists between those two enzymes,th e overallstructure s are very similar [45]. Forbot h enzymes the structure ofth e monomer can bedivide d into four domains: the FAD binding domain,th e NAD+bindin g domain,th e central domain andth e interface domain [39-46].Th e interface domain contains the residues necessary for dimerformation . A schematic representation of the dimer is presented in Fig.5 .Th e FADappear s to be bound in an extended conformation in both enzymes [39, 45].Fo r

15 '2

Lip(SH)2

Fig. 5. Diagrammatic representation of the dimer structure of lipoamidedehydrogenas e from Azotobacter vinelandii. glutathione reductase crystal structures have been obtained with several ligands bound e.g. NADP+an dglutathion e [42,43] . Furthermore the structure ofth e 2-electron reduced enzyme was solved [42,43] . Until now for A. vinelandiilipoamid e dehydrogenase only the structure of the unliganded enzyme in its oxidized state was obtained, but from the extensive structural similarity with glutathione reductase and lipoamide dehydrogenase from P.putida [42 ,43 ,46 ] it iseviden t that binding of NAD+ occurs in an equivalent position to that of NADP+i n glutathione reductase. Fig. 6 shows a drawing of the active site.Thi s drawing is provided as an overview and is not claimedt o be an accurate detail.Th e numbering of the amino acids is according to the A. vinelandiilipoamid e dehydrogenase. A prime (') following a residue's number indicates that this residue isfro mth e second subunit. Thusth e drawing shows that each active site is built from elements of the FAD bindingdomain , the NAD binding andth e central domain of one subunit aswel l as from the interface domain of the other subunit. The isoalloxazine part of the FADdivide s the active site intotw o parts,a t the s/-side the dihydrolipoamide [lip(SH)2] binding site and atth e re­ side the NAD+bindin g site.Th e re-an d s/-faces of flavin are assigned following the rules of Hanson [49] with N-5 asth e central atom.Accordingl y the front and rear sideso f the flavin in Fig.3 ar e the si-an d re-faces, respectively. The histidine atth e sAside, His450', is the so-called 'active site base' and is oriented with N-3 toward the disulfide bridge, somewhat closer in the lipoamide dehydrogenase and glutathione reductase structures to the N-terminalCys48/5 8tha nt o Cys53/63 [41, 42,45] ,an d playsa ver y important role in catalysis as will become clear inth e following sections. Cys48 and Cys53 form the redox-active disulfide bridge.Th e sulfur of Cys48 faces out inth e lip(SH>2bindin g site and suggests that this sulfur reacts with the substrate.

16 His450 Pro451

°^HN N-Ala452 (\/G\uA55

NH2

Fig. 6.Schemati c representation of the active site of lipoamide dehydrogenase from Azo\obacter vinelandii.

Forglutathion e reductase, structural evidence forth e formation of this bond between glutathione and Cys58 (the equivalent of Cys48 of the lipoamide dehydrogenase structure) was obtained [42,43] .Therefor e Cys48 is referredt o asth e 'interchange thiol'. I pon reduction of the disulfide,th etw o protons associated with this process are presunfe dt o be shared between thetw o thiols andth e active site base.Thi s results,a s evidencedb y spectral studies, inth e formation of a red intermediate (thiswil l be discussedi nth e next sections). The redcolou r has been ascribedt o a charge-transfer complejx inwhic hth e FADi sth e acceptor anda thiolat e isth edono r [50].Th e structural data ndwclearl y show that the only possible candidate for a role as charge donor isth e thiolatejo f Cys53. The negative charge of the thiolate is proposedt o be stabilized by the positive charge on the protonated histidine [51] and by adipol e originating from an oc- helix by residues 325t o 340 [42,43 ,45] . Theglutamat e at position 455'form s ashor t strong hydrogen bondwit h N-1o f His450]. An identical feature was found inth e glutathione reductase structure andthi s prompted Pai and Schulz [43]t o suggest that this glutamate is responsible for raising the pKi ofth e active site base.Th e observation of ahig h pKa ofth e active site base originated from spectral and kinetic studies on pig heart and E. coli lipoamide dehydrogenase and glutathione reductase [51-57]. Tha structural data concerning the NAD+ binding to lipoamide dehydrogenase from P.putiqa show s that the binding toth e oxidized enzyme is in aconformatio n that cannot

17 leadt o effective electron transfer when bound in a similar way toth e reduced enzyme [46]. Inglutathion e reductase NADPHbind s in an extended conformation but before effective binding occurs,th e incoming NADPH must displace atyrosin e (Tyr197)tha t shields the FADfro m the solvent [43].Th e pyridinium ring adopts a stacked conformation withth e isoalloxazine with N-4 ofth e pyridinium close to N-5 of the isoalloxazine to allow good overlap of the Jt-orbitals (suggested already in 1960 [58]), andther e isa clos e approach of N-4 toth e Lys66/Glu201 ion pair [42,43 ]tha t forms a salt bridge beneath the FADi nth e vicinity of N-5. This ion pair isthough t to be catalytically involved [43,59] . In A. vinelandiilipoamid e dehydrogenase the Iys66/Glu201 ion-pair counterpart is formed byth e ion-pair Lys57/Glu194 [45], however the Tyr197counterpar t is avaline .

Spectral properties

The spectral properties of lipoamide dehydrogenase are closely linkedt o kinetic properties. Inorde rt o understand the spectral features, key elements of the kinetics, that will be discussed in depth in following sections,wil l be presented here. Although there aretw o active sites perdimer , here it is assumed for simplicity that the active sites act independently and equivalently. Therefore the phenomena described will refer to one active site unless explicitly stated otherwise. The presence oftw o redox-active groups inth e active site,th e FADan dth e disulfide, indicates that the enzyme can take upfou r reducing equivalents. It was shown that the reducing equivalents are transferred in pairs [60].Thus ,thre e reduction levels can bediscerned :th e non-reduced enzyme,o r as it will be referred to 'the enzyme in its oxidized state' [Eox],th e 2-electron reduced enzyme [EH2],an dth e 4-electron reduced enzyme [EH4].Wit h respect to effective catalysis,onl y Eox and EH2ar e relevant{60 , 61], during turnover the enzyme shuttles between Eox andEH2 . For all lipoamide dehydrogenases studied sofar catalysis inth e forward direction takes place according to a ping-pong mechanism [56, 58,62-6 4 ] although this has been questioned [65,66] .Th e reactions can be represented as follows:

Eox + lip(SH)2 s ** EH2+ lipS2 (1)

+ + EH2 + NAD v ^ Eox + NADH + H (2)

According to these reactions EH2 is a'free ' unliganded enzyme species and studies of this species might give more insight intoth e mechanism of the enzyme. EH2canno t

18 only bo obtained by reduction of Eox by lip(SH)2 but also by dithionite, borohydride or dithiotlireitol, avoiding any complications of the presence of lipS2-

Oxidized enzyme. In Fig. 7 representative spectra of Eox, EH2 and EH4o f pig heart ijpoamidedehydrogenas e are shown [52,58] .Whe n compared to free FAD,th e spectnjimo f Eox showstha t the major flavin banda t 455 nmi s5 nm red-shifted.Th etw o major absorption bands have much more vibrational fine structure as demonstrated

12 v° i-FAD 0 \ I-FADK E z 6 W f SH O M/ 1—FADH2 4 5 \1_USH EH4 z •" 2 UJ 0 1 1 r-* —-i\ 1 X^— 400 500 600 WAVELENGTH (nm)

Fig. 7. Spectrao f pig heart lipoamide dehydrogenase. The reductant was dithionite, pH 7.6. Froifi ref. [6]. byth ejshoulder s at 370,44 0 and 480 nm.Thes e features are indicative for aflavi n ina n apolar(environment [67].Th e extinction coefficient at 455 nm is 11.3 mM_1 cm"1[68] . Eox isjals o fluorescent. For glutathione reductase a similar absorbance spectrum was reported(X maxa t460nm ) howeverthi s enzyme is almost notfluorescen t due toth e presenfceo f a stackedtyrosine . From site directed mutagenesis studies on E.coli lipoamide dehydrogenase, the Eox spectrum of the wild-type enzyme almost identical to pig heart enzyme and also highly fluorescent, it appeared that similar spectral properties (shift of absorbance maximum to 462 nm and strong quenching of fluore: ence) as reported for glutathione reductase were obtained after changing Ile184, the counterpart of Tyr197o f glutathione reductase, intoTy r[69] .

2-eiectron reduced enzyme. In the late fifties and early sixties extensive studies on pigpear t lipoamide dehydrogenase were performed.Th e addition of one molar equivalent with respect to FAD of lip(SH)2 or dithionite resulted in atypica lspectrum , designated EH2i n Fig.7 . Studies with arsenite showedtha t the typical spectrum was

19 due to the presence of a reduced disulfide and mechanisms involving flavin-sulfur-bi- radicals were proposed [58, 60]. Itwa s Kosower [50] who suggestedtha t the typical spectrum arose from a charge-transfer interaction between athiolate and FAD in which thethiolat e isth e donor andth e oxidized FADth e acceptor. Therefore EH2wit h the spectrum showingth e typical absorbance at 530 nmi s nowcalle dth e 530 nmcharge - transfer species. The ability to isolate monomelic apoenzyme under nonreducing conditions showed that the disulfide is not between the subunits but within one subunit [63, 70]. Peptide sequence analysis of pig heart enzyme showedtha t the two cysteines forming the disulfide are only separated by five amino acids[36] . The EH2spectru m of pig heart enzyme thus showstha t the reducing equivalents mainly reside onth e disulfide and it was showntha t a large excess of lip(SH)2 did not reduce the enzyme beyond the EH2 level [60],demonstratin g that the EH2specie s is stable under anaerobic conditions. Further reduction than EH2wa s however easily accomplished by reduction with more than one molar equivalent of dithionite which resulted informatio n in EH4[4 ,52 ,58] . For E.coli lipoamid edehydrogenas e it was reported that a small excess of lip(SH)2 resulted in over-reduction to EH4an dth e addition of one molar equivalent of lip(SH)2 resulted in an EH2spectru m with a less intense 530 nm charge-transfer band [63].Thes e phenomena will be discussed later.

Thiol to FAD electron transfer. The characteristic EH2 spectra were also obtained for glutathione reductase and mercuric ion reductase after reduction with one molar equivalent of dithionite [5, 71, 72].Whe n both EH2o f pig heart lipoamide dehydrogenase and glutathione reductase were reacted with the thiolate specific reagent iodoacetamide it appeared that only one mole of iodoacetamide reacted per mole of FAD,whil e two moleso f thiol(ate)s are present per FAD[ 54,73-75] . Furthermore the spectrum of pig heart enzyme changed to a spectrum typical for FAD (enzyme) in its non-reduced state, however, 7 nm blue shifted,whil e the spectrum of glutathione reductase retained its typical EH2spectrum . For the monoalkylated glutathione reductase as well as the monoalkylated lipoamide dehydrogenase it was showntha t differential reactivity of thetw o nascent thiols exists andtha t in bothcase s the cysteine closer to the N-terminus was labeled [73].Th e resulting spectrum of monoalkylated glutathione reductase shows that the charge-transfer thiol is not alkylated [54]. For lipoamide dehydrogenase this was shown bytitratio n of the monoalkylated enzyme with NAD+[74 ,75 ] which resulted in a partial bleaching of the spectrum (see Fig.8 )wit h aconcomitan t rise ofth e absorbance at 380 nm.Onl y1. 2 moles of NAD+ reactedwit hth e 2 moles of active site per mole of dimer. Extrapolation ofth e spectrat o fullformatio n on both monomers resulted inth edotte d spectrum ofFig . 8. This spectrum bares close resemblance to a C4a-thio-substituted-dihydroflavin

20 12

0J6 — 448 nm s ^10 A\ "E 'E //A\ ,<> 04 o _ ^f ///3\\ \ = 8 3 05 £ 11//// \ \ i .—•-»'*390n m o v \vA\i o 1 1 1 1 1 ° 2 4 6 8 10 x 4 1 ° 1/NAD* (mM) Ul \ 1 \ 1 \ 2 - \

0 " ' 1 I^H II I I 400 500 600 WAVELENGTH (nm)

Fig.8 . Spectral changes induced by titrationo f monoalkylatedpi g heart enzyme (1) as mix((dwit h increasing concentrations of NAD+ (2-6 respectively). Equalconcentration s of NAt»+ were added to the reference cuvette.Th e dashed line isth e spectrum extrapolated to infinite NAD+b y usingth e 1/Ae448intercep t from adoubl e reciprocal plot (inset). The dottiidlin e is anestimat e of thespectru mo f the modifiedflavi nspecie s after correction from the contribution of residual unmodified flavin. From ref. [75].

Charge-transfer C4a-adduct reduced flavin

a .SlH Y .NH \ O P? -S" — S

f — SH HHis- — S His-

Schfeme 1. Formation and breakdown of thethiolat e to C4a-FAD adduct. From ref. [75]. deriva

21 hence its name 'interchangethiol' . Thethio lclose rt oth e FAD,th e C-terminalcystine , was named 'charge-transfer thiol'.

A base in the active site. For simplicity the above results are presented in a way asthoug h protons are not involved inth e process of reduction of lipoamide dehydrogenase. They are however involved, but it doesn't change the conclusions drawn. Williams [5] first suggestedth e involvement of a base inth e active site inorde rt o accept a proton of the substrate lip(SH)2 allowing facilitated nucleophilic attack of the disulfide byth e thiolate of the substrate. Evidence for this base was obtainedfro m studies on the pHdependenc e of the redox potentials andfro m pre-steady state kinetics (see sections on redox potentials and pre-steady state kinetics for details). Further evidence for this base was gained from studies on E.coli lipoamide dehydrogenase using a Afunctional arsenoxide [79, 80] which demonstrated that this base is a histidine. The latter finding was confirmed byth e three dimensional X-ray structure of glutathione reductase [39-43] and lipoamide dehydrogenase [44-46],an d by site directed mutagenesis experiments on both these enzymes where the histidine was replaced by other residues [81-83].

pH dependent spectral properties of Ehfe. The presence of the histidine and two cysteines inth e active sitethu s demonstrates that at leastthre e titratable groups can contribute to the spectral properties of EH2.A ver y careful analysis ofth e pH dependent properties of E.coli lipoamide dehydrogenase by Wilkinson and Williams [53, 84] yielded much insight intoth e distribution of tautomeric/electronic species atth e EH2 levelwhic h are depicted in Scheme 2.Th e central column shows the three forms that predominate at neutral pH.Th e central form isth e charge-transfer species which is non-fluorescent andfavore d by high pH.Th e upper form isth e fluorescent prototropic tautomer of the charge-transfer species favored by low pH.Th e lower species inth e central column hasth e electrons onth e flavin and is somewhat favored by low pH.Th e

assignment of the high pKat oth e histidine was based on kinetic studies [51].Fo r £ coli

enzyme the pKa values as attributedt o the charge-transfer thiol and histidine are 5.7 and 8.2 respectively [53],5. 9 and 7.5 respectively in 0.2 Mguanidiniu m hydrochloride [84]. Extrapolation to fully formed EH2a t each pHyielde d values for fluorescence and 530 nmcharge-transfe r absorbance which intur n were extrapolated to zero fluorescence and zero charge-transfer absorbance in order to obtain the maximal charge-transfer absorbance and the maximal fluorescence respectively. It appeared that the maximal charge-transfer reached an extinction coefficient of 3.3 mM-1 cm-1 and the maximal fluorescence was 68%o fth e Eox. From stopped flow experiments a similar high value for the extinction coefficient of the charge-transfer absorbance

22 -H+

pKal PKa2 ^FAD- -FAD

(I) Fl. -SH SH -SH HHIs S' HHIs

-FAD- ^-FAD-,

(») CT. SH HHIs SH His

FADH2

(Ill) REDUCED •S FLAVIN

Scheme 2. Modelfo rth e pH dependence ofth e spectroscopic properties of E. coli lipoamidedehydrogenas e atth e EH2 level.Th e protonation states are represented byth e columns labeledA , Ban dC .Th ethre espectrall ydistinc t speciesar e represented byrow s andjarelabele d I,I Ian d III.Th earrow so fform sM B andII Csignif yth echarge-transfe r interactions betweenth ethiolate san dth e FADan dar e drawnfro mth edono rt oth e acceptor. From ref.[53] . was obtained immediately after mixing of the enzyme with lip(SH)2 however it decreasedgraduall y intim e [53] as a result of tautomerisation andformatio n of the reduced FAD EH2specie s (FIH- species) and over-reduction to EH4.Th e maximal value ^f 3.3 mM"1 cm-1fo r the extinction coefficient was also obtained for EH2o f pig heart ^nzyme [55] showing that the lowercharge-transfe r absorbance of E.coli enzym e isdu e{t oth e presence of the other two species [53] andsuggestin g that in pig heart enzymjeth e charge-transfer species prevails. Acomplicatin g factor in the studies of the E.coli enzym e was a phenomenon known as disiroportionation [53,84 ]tha t accounted for a loss of charge-transfer absorbance of 20 %.Thi s means that 2EH2 react to Eox and EH4. Disproportionation appearedt o be enhanced inth e presence of lipS2o r reaction products after dithionite reduction [53]. Thereverse process ,formatio n of EH2fro m Eox and EH4wa s also observed andi s knowna s comproportionation [53]. In pig heart enzyme disproportionation was observedthoug h this contributedt o a much lesser extent [55].Th e difference in extent of disproportionation was attributed to differences in redox potentials between the enzymes. The pHdependenc e of the spectral properties of EH2o f pig heart lipoamide dehydrogenase and glutathione reductase essentially follows Scheme 2, with the

23 exception that the charge-transfer absorbance is much higher which is explained in terms of the equilibrium among the different electronic species being in favor of the charge-transfer species [55].Th e pKa values obtained for pig heart enzyme and glutathione reductase are respectively; 4.4 and 4.8 forth e charge-transfer thiol and 8.7 and9. 2 forth e histidine[55] . Ina n attempt to simplify the titration behaviour of Ehfesit e directed mutagenesis experiments were performed recently to replace either of the thiols orth e histidine by non-titratable residues [ 57,81-8 3 ,85] . Replacement of the interchange cysteine resulted in enzymes showing 530 nmcharge-transfe r absorbance in their oxidized state. The general conclusion istha t replacingth e interchange thiol affects both pKa'so f the wild-type enzyme;th e pKa ofth e charge-transfer thiol is lowered while the p«ao f the histidine is raised.Th e shift of the pKa's was explained by mutual influences of the charge-transfer thiolate and the imidazolium in analogy to papain [51]. Upon replacement of the histidine in E.coli lipoamid e dehydrogenase by glutamineth e pKa of the charge-transfer thiol increases to 7.4 [83].

Spectral changes in the presence of pyridine nucleotides. The spectral properties of lipoamide dehydrogenase in the presence of (reduced) pyridine nucleotides are quite different from those presented above. Fig.9 depict s pig heart enzyme reduced by 3 molar equivalents of NADH andth e effect of addition of NAD+t o this reduced species (from ref. [86]).Typica lfo r reduction by NADH of all lipoamide dehydrogenases studied inthi s respect isth e formation of a long wavelength absorbance, extending beyond 750 nm,whic h shows considerably different intensities at 750 nmdependin g onth e source of the enzyme. This longwavelengt h absorbance is attributedt o acharge-transfe r complex between reduced flavin and NAD+[58 ,60 , 87]. Incas e of pig heart enzyme the intensity at 750 nm is low while the absorbance at 530 nm is high (see Fig.9) .Thi s has ledt o the proposal that upon reduction with NADHi n pig heart enzyme the electrons mainly reside onth e disulfide. Further reduction beyond the level shown cannot be accomplished with NADH [86].Additio n of NAD+t o the NADH reduced enzyme resulted in a shift of the long wavelength absorbance with a maximum around 570-580 nm [86].Th e spectral change induced upon addition of NAD+appeare dt o bever y rapid (complete within 3 ms) andfro m this it was concluded that this species might be acatalyti c intermediate [86].Matthew s andcoworker s [88] titrated EH2,generate d by reduction with dithionite,wit h NAD+an d found a similar spectrum as shown in Fig.9 .Thei r conclusion wastha t the spectrum represented an enzyme species with the electrons onth e disulfide andwit h NAD+bound .Titratio n of pig heart lipoamide dehydrogenase inth e presence of NADase with NADH

24 400 500 600 Wavelength(nm )

Fig.9 . Visible absorbance spectrao fpi ghear t lipoamidea sobtaine d after reductionwit h3 molarequivalent so f NADH ( )followe db yadditio no f 100 molarequivalent so f NAD+ (- • -T- • -) atp H7.6 . ( ) Eox(3 5uM) . Fromref .[86] . preinciibated with NADase ledt o further reduction than shown in Fig. 9. Addition of NAD+restore dth e spectrum,though ,du et o the presence of NADase,th e spectrum gradually changedt oth e one of a more reduced FADwit h concomitant losso f the abforbance at 530 nman da rise at 750 nm [60]. Fromthis ,an dfro m kinetic studies (see section on kinetics) it was concludedtha t NAD+ preventsth e pig heart enzyme from biing over-reduced to the EH4state . Thi spectral changes upon reduction by NADH reported for E.coli lipoamide dehydrogenase are somewhat different than those reported for pig heart enzyme [63]. One mplar equivalent of NADH resulted in afurthe r reduction of the FADtha n reported for pig(hear t enzyme and a much lower 530 nm absorbance while the long wavelength absorbance (>600 nm )wa s higher [63].Als o the enzyme was easily further reduced towardth e EH4 level by excess of NADH indicating that there is adifferenc e in redox potential between pig heart and E.coli enzym e [61].Als o for E. colienzym e NAD+ appeals to prevent the enzyme from being over-reduced to EH4thoug h notfull y anda t much higher NAD+concentration s [61]. Fotfbot h E. colian d pig heart enzyme it was showntha t arsenite blocksth e disulfide of pre-jreducedenzym e [60,63] .Additio n of NADHt o these enzymes results in reduction of the FADwit h special characteristics:th e shoulders ofth e FADar e retained and a long wavelength absorbance develops with a maximum around 700-730 nm[60 , 63]. Treatment of the enzymes showing this spectrum with NADase resulted in a losso f this long wavelength absorbance. Therefore it was concludedtha t this absorbance

25 arises from acharge-transfe r complex between the reduced FADan d NAD+.Thi s charge-transfer complex could be different from the one previously mentioned inthi s section duet o the slightly shifted maximum. Spectral properties of the enzymes were also studied using other pyridine nucleo­ tidesthe n the physiological substrates NAD+an d NADH.Fo r both enzymes itwa s shown that NADPHwa s a reductant. The spectrawer e however similar to those obtained by reduction with dithionite or lip(SH)2, i.e. no long wavelength absorbance typical for acharge-transfe r interaction between reduced flavin and NADP+wa s detected [63,87 ] showing that nocorrec t binding of NADP(+)(H)occurred . Other NAD+ analogs, like 3-acetylpyridine adenine dinucleotide [AcPyAde+], 3-aminopyridine adenine dinucleotide [AAD+] and nicotinamide hypoxanthine dinucleotide [NHD+] and their reduced forms did however result in spectral characteristics upon reacting with either reduced or oxidized enzyme aswa s shown by Matthews andcoworker s [88]. A markedchang e of bothth e E.coli an dth e pig heart EH2spectru m occurred after the addition of AAD+:a shift ofth e 530 nmabsorbanc e to longer wavelength took place accompanied by a rise ofth e absorbance [84,88] .A st o the nature of the species exhibiting this spectrum no clues were obtained about the electron distribution between the disulfide andth e FAD[88] . Reduction of pig heart enzyme with AcPyAdeH inth e stopped flow instrument revealed a deadtim e spectrum with low long wavelength absorbance from 510 nm extending beyond 750 nmwit h noclea r maximum [88]. Furthermore it was showntha t inth e deadtim e noAcPyAde H was oxidized.Additionally , no isotope effect wasfoun d forth e formation ofth e deadtim e spectrum,whil e an isotope effect of 2.6 wasfoun dfo r the hydride transfer leading to formation of EH2.Thes e observations ledt o the conclusion that the deadtim e spectrum representedth e Michaelis complex of Eox and AcPyAdeH andtha t the low long wavelength absorbance was due to the charge- transfer complex between FADan dAcPyAde H [88]. Duet oth e very high rateo f reduction by NADH of any lipoamide dehydrogenase studied inthi s respect, no spectral evidence for an Eox-NADH complex was ever observed. Duringth e reoxidation of pig heart EH2b y AcPyAde+an d NHD+i nth e stopped flow instrument deadtim e spectra were obtained with a maximum around 580 nm[88] , similart oth e spectrum shown in Fig.9 after addition of NAD+t o the reduced enzyme. Also reduction by NHDHyielde d such a spectrum [88]. Itwa s concluded that these spectra represent a species with the disulfide reduced (dithiol),th e FAD oxidized and a pyridine nucleotide bound [88]. Fromth e change inthes e spectra when compared to EH2 it was inferred that the binding of a pyridine nucleotide influences the equilibrium distribution of electrons betweenth e FADan dth e reduceddisulfid e such ast o increase

26 the amount of reduced FAD [88].Thi s implication will beaddresse d inth e section on redox potentials. Recently, mutated E.coli lipoamide dehydrogenases were described in which Lys53wa s replaced with arginine or Ile184wit htyrosin e [59,69] .Th e lysine isth e counterpart of Lys66 of glutathione reductase whereth e lysine forms an ionpair with Glu201Thi s ionpair isver y close to N-5 of the FAD [45] and isthough t to be involvedi n modulatingth e redox potential ofth e FAD [43,59] .Ile18 4 isth e counterpart of Tyr197, the residue to be displaced when NADPH binds to glutathione reductase [43] (see section onthre e dimensional structures). Upon reduction of the Lys53->Arg and Ile184-t>Tyr mutated enzymes by one molar equivalent of NADHth e spectra revealed a much stronger reduction of the FADan d especially for enzyme Lys53->Arg an increased FADH--NAD+charge-transfe r when comparedt o the wild-type enzyme [59]. Thedifference s in FADH--NAD+charge-transfe r amongth e wild-type andth e mutated enzymus is most likely attributable to differences inaffinit y for NAD+[59] . Furthermore in the Ile184->Tyr mutated enzyme the spectrum of Eox closely resemblesth e spectrum of glutathione reductase, ashif t of the absorbance maximumt o 462 nm.Als o the fluorescence quantum yield of this enzyme decreased significantly and was similar to that asfoun dfo r glutathione reductase [59,69] .

Summary of spectral species. In Fig. 10 (p 28) all species of lipoamide dehydrogenase that were spectrally identified are presented together with the absorbance maxima of the charge-transfer band and visible appearance.

Redo* potentials

Inth e active site of lipoamide dehydrogenase two redox active groups are present, the disulfide andth e FAD.Th e formation of a stable EH2for m in pig heart enzyme with the reducing equivalents onth e dithiol as evidenced by the 530 nm charge-transfer absorbance indicates that the redox potentials of the disulfide andth e FAD are well separated. Inth e E.coli enzym e relatively easy over-reduction to the EH4stat e occurs, suggesting that the redox potentials of those two groups are lesswel l separated[53] . For bomenzyme s the redox potential of each couple wasdetermine d [52,53 ] For pig heart enzyme the reported potentials of the couple E2fo r the disulfide/dithiol

(Eox/EH2) ando f the couple E1fo r FAD/FADH- (EH2/EH4) are -0.280 V and -0.348V respectively at pH 7.0 [52].Th e pHdependenc e AE/ApHwa s shownt o be-0.0 6 V inth e pH ranje 5.5 to 7.6 for bothcouples . Fromthi s pHdependence ,showin g that two proton];a s well as two electrons are involved while simultaneously thiolate to FAD

27 -APADH -NHD+ NHD" FAD -FAD

NAD+ FADH FAD FADH" FADH J S -S J S" S" I His- HHis- +HHis-| HHis-J S -SH hSH -SH

FIH- EH2 EH4 EH4-NAD + leuco red leuco blue/grey 530 nm 750 nm

Fig.10 . Summaryo fspectrall yidentifie dcatalyti cintermediat especie so flipoamid e dehydrogenase. charge-transfer was observed, it was tentatively concluded that a base participates in the active site [52]. Forth e E.co//enzym eth e potentials at pH7. 0 are -0.264V fo r couple E2an d-0.31 7 Vfo rcoupl e E1[ref . [167] in [6]]wit hth e same pHdependenc e as found forth e pig heart enzyme [53]. Both potentials are more positive inth e E. coli enzyme than inth e pig heart enzyme andth e difference between them is less, making the E.coli enzym e more easily reducable to the EH4level .Th e redox potentials at pH

7.0 of the couples lipS2/lip(SH)2 and NAD+/NADH are -0.281 V and -0.320V respectively. Sinceth e pHdependenc e of the couple NAD+/NADH is-0.0 3 V/ pH unit, the reduction of EH2t o EH4b y NADHi s more easily accomplished at low pH[52] . During catalysis the electron flow isfro m dithiol via FADt o NAD+. Purely basedo n the redox potentials of the respective couples, especially inth e pig heart enzyme,th e electrons havet o overcome athermodynami c hill,th e FAD moiety. It istherefor e likely that the redox potential of the FADi nth e NAD+boun d enzyme is higher than inth e free enzyme andthu s NAD+serve s asa n effector to facilitate electrontransfe r from the dithiolt oth e FAD.Thi s effector role of NAD+wa s indicated intw o different ways.Th e first one was by spectral analysis of titration of pig heart EH2wit h NAD+an d NHD+.Thi s

28 resulted ina clea r shift of the charge-transfer absorbance maximum to 580 nman dwa s explained interm s of adecrease d excited state oxidation-reduction potential of the charge donor (thiolate) and acceptor (FAD) implying a decreased energy necessary for ground state electron transfer [52].Th e second indication was obtained by calculation of the rpdox potentials of the couples Ei and E2fo r pig heart enzyme based on steady state Wpetical- and dissociation constants which revealed that both couples were 30 mV mcfe positive inth e NAD+boun d enzyme[89] .

Kinetits of the forward and reverse reactions

Steady state kinetics. All lipoamide dehydrogenases studied thus far in this respect, except for lipoamide dehydrogenase from A. vinelandiian d pig heart [66,65] , function according to a ping pong bi bi mechanism inth e physiological direction [56, 60-64]. The kinetic parameters of the enzymes are presented inTabl e 2. For pig heart and ratilive r enzyme it was showntha t the reverse reaction proceeds viath e same mechanism [62,64] ,whic h wasquestione dto o [65],an d remainsth e same in pig heart enzymi when studied with lipoamide dehydrogenase complexed inth e OGDC [90]. For bothth e pig heart [65] andth e A. vinelandiienzym e [66] aternar y complex mechanism was proposed.Thi s was based onth e fact that the LB-plotswer e convergent ando n product inhibition studies [65].Thi s was supportedfo r pig heart enzyme byth e communication of two NAD+bindin g sites,on e high affinity site (Kd= 50-60 nM) and one lovyaffinit y site (Kd=200-250 nM) per monomer onth e oxidized enzyme [91]. However,Wilkinso n and Williams [61] argued that the apparent ordered mechanism could bjeexplaine d by assuming dead endcomplexe s to occur, a phenomenon that is commdnt o ping pong enzymes andwa s demonstrated forth e rat liver and E.coli enzymA [61,64]. The physiological reaction of E.coli lipoamide dehydrogenase proved difficult to study duet o severe product inhibition [63]. Use of an electron acceptor with higher midpoint potential than NAD+, in casuAcPyAde +, revealedtha t the mechanism of this enzyme isals o ping pong bi bi [63]. Later studies, usingth e stopped flow spectrophotometer, allowed the use of NAD+a s asubstrat e [56].Thes e studies reinforied the finding that the enzyme functions according to a ping pong bi bi mechanism. Furthermore these studies showedtha t the plot of NAD+versu s velocity did not oboy the simple Michaelis Menten equation, but required an equation taking cooperativity into account. The Hill coefficient of 1.1-1.4 indicated modest positive cooperativity atth e EH2level . From steady state kinetics nofurthe r indications of cooperativity for the other lipoamide dehydrogenases were reported except for rat liver enzymo which showed slight negative cooperativity atth e first substrate level at 25°C

29 Parameter pig heart rat liver E.coli

Forward reaction

Kmlip(SH) 2 (|iM) 300 490 -

Km NAD+(u.M ) 200 520 230 turnover number* 550 690 420

pKaV 6.3 7.0 6.7 Reverse reaction

Km NADH(uM ) - 62 -

Km lipS2 OiM) - 84 - turnover number* 530 248 -

pKaV 7.9 - - assay conditions 25°C,p H 7.6 37°C, pH 8.0 25°C,p H 7.5 Reference 88 64 56

Table2 .Kineti cparameter sfo rlipoamid edehydrogenases . # molessubstrat ex moleFAD -1x sec" 1.

[64]. For pig heart enzyme cooperativity was reported in monoalkylated enzyme titrated with NAD+[75 ] (see above) andfro mtitratio n experiments of Eox with NAD+[91] . Studies of pig heart enzyme inth e reverse reaction revealed asignifican t pH dependent lag in NADH oxidation by lipS2- The rate ofth e reaction accelerated as NAD+wa s produced,bu t was negligible if NADase was present inth e assay mixture. NAD+administratio n to the starting mixture abolishedth e lag-phase [60].Close r examination of this phenomenon showedtha t the acceleration,optimu m at pH 6.3, was not due to enhanced affinity of EH2fo r lipS2i nth e presence of NAD+bu t was merely due to the reoxidation of inactive EH4t o EH2b y NAD+an d so enhancing the steady state concentration of EH2[92] .Additionall y it was showntha t NAD+als o actsa sa product inhibitor though at higher concentrations (apparent Kactivation = 5 u.M,Kjnhibitio n = 280 uM)[92] . For E.coli enzym e the same autocatalytic time course asfoun dfo r pig heart enzyme was observed inth e reverse reaction [61]. Detailed analysis of this reaction showed that the same conclusions drawn from the pig heart enzyme also apply to the E.coli enzym e [61]. NAD+serve s both as anactivator , though with a pHdependen t apparent binding constant of NAD+wit hth e optimum at pH8. 0 (Ka = 0.4 mM),an d as an inhibitor (Kj = 1.0mM) . Itwa s showntha t inhibition was ofth e dead-endtyp e with EoxNAD+a s the dead-end complex. Furthermore,a dead-en d inhibition by NADH was shown at the EH2 level resulting eventually in formation of EH4.Th e activation by NAD+wa s again attributedt o reoxidation of EH4t o EH2[61] . InSchem e 3a model is presented ofth e catalytic cycle andth e dead end complexes.

30 E-NAD+ Kip V* p

Q E K A •X, KiS

E-lip(SH)2" "E-NADH" \ \ EH2-NPS2 EH2-NAD+

B K EH2 A .ar EH2-NADH

Schfm e3 .Mode ldepictin gth ecatalyti ccycl ean ddead-en dcomplexe si ncatalysi sb y lipoamidedehydrogenase .A ,B ,P andQ ar eNADH ,lipS2 ,NAD +an dlip(SH) 2respectively . Froriiref. [61].

Pre-i-steads y state kinetics. The pre-steady state kinetics have most extensively been studied with the pig heart enzyme. Massey,Gibso n andVeege r [62] wereth e first to determine the rates of reduction of the enzyme by lip(SH)2 and reoxidation of EH2b y NAD+. The reported rate constants at 25 °C, pH7.6 , 555 s-1 for lip(SH>2reductio n and> 800 s_1 for reoxidation in relation withth e steady stateturnove r number (575s _1)clearl y demonstratedtha t the reductive reaction is rate limiting in overall catalysis andtha t specie);a tth e EH2 level are relevant intermediates. Aver y detailed analysis of the pre-steady state kinetics of pig heart enzyme comprising reductive and oxidative reactions was reported by Matthews and co-workers and hasyielde d much insight inth e mechanism of the enzyme [51, 88].Th e reductiono f the enzyme by lip(SH)2 appeared to proceed at a rate of 830 s_1 between pH 5.5 and 8.0,thu s is independent of pH inthi s range.Th e rate ofth e reoxidation reaction of EH2 by lipSfehoweve r was maximal at pH 6.2 (880 s~1)an d appeared to begoverne d by a pKa ofb. 9 onth e EH2HPS2comple x [51].Thes e results,i nconjunctio n withth e pH dependence of the redox potential of E2,provide d conclusive evidence that a base is involved incatalysi s which has a low pKa (5.5) in Eox anda high pKa (7.9) in EH2[51] . The presence of the base as proposed byWilliam s [5], now knownt o be a histidine, substantiates the proposed mechanism of proton abstraction from the substrate byth e Hisfonnin g a thiolate anion and facilitating nucleophilic attack of the disulfide. According to this mechanism,presente d in Scheme 4, upon reaction of the activated

31 FAD FAD •? I 44-hlS- 1 —JN-tll»- T HN*-hls- \^U -S *v>* "O— *\J-*

-FAD SH / •FAD FAD -FAD SH ^=* SH -SH , ^3-

Scheme 4. Proposed reactionsequenc efo rth e reductive half reactionb y lip(SH)2- Iti sno t knownwhethe r the sulfura t C6o ra t C8 isinvolve di nmixe ddisulfid eformation .Adapte d from ref.[51] . substrate withth e disulfide a so called 'mixed disulfide' species is formed which by a second proton abstraction fromth e substrate byth e His leadst o attack ofth e mixed disulfide bond and subsequently to release of the oxidized product. This mechanism is the working hypothesis ever since. The reduction of the enzyme by NADHa t pH5. 9 andth e reoxidation of EH2b y NAD+a t pH8. 0 were shownt o proceed at rates fastertha n 3000 s-1 [88].Give ntha tth eturnove r number inth e physiological direction is83 5 s~1a tth e optimum,p H8. 0 [88],thi s demonstrates that the reductive reaction is rate limiting during catalysis as was already reported by Massey andco-worker s [62].Fro mth e pHdependenc e of Vmax ofth e physiological reaction,whic h isgoverne d by a pKa of 6.3,togethe r with the independence of pH of the rate of reduction by lip(SH)2, it was concluded that the decrease of Vmax at low pHcoul d beattribute dt o adecrease d rate of reoxidation of EH2b y NAD+[88] .Th e decreased rate of reoxidation was assignedt o the fact that the reoxidation is accompanied by proton release according to equation 2 (see page 18) which becomes thermodynamically unfavourable at low pH. Fromthi s it was inferred that both EH2,i n its protonated form,an d EH- bind NAD+,bu t proton release of EH2i s mandatory to complete the reoxidation reaction [88]. Inth e reverse reaction the rate of reoxidation of EH2b y lipS2wa s shownt o be rate limiting overth e entire pH interval studied and it was shown that the decrease ofth e rate of reoxidation of EH2b y lipS2depende d onth e protonation state of EH2,EH " being a non-productive species inthi s reaction [88].Whe n using the NADH analog AcPyAdeH

32 inth e Reverse reaction,th e reductive reaction appearedt o be rate limiting,th e rateo f reduction being decreased bytw o orders when comparedt o NADH. Inth e reaction lip(SH]2/ AcPyAde+ at pH8.0 ,th e reoxidation byAcPyAde +wa s shownt o be rate limiting. Here also the rate of the analog decreased two tothre e orders of magnitude,a t pH7.6jan d 6.3 respectively. Identical studies with the analogs NHD(H)(+) showedtha t the rate of reduction by NHDH haddecrease d one order andth e rate of reoxidation of EH2h^ ddecrease dtw o orders [88]. Th4 slower rates of the analogs in all reactions enabled the spectral identification of several possible intermediates inth e reaction mechanism of pig heart lipoamide dehydrogenase [88].Thes e species were presented in the section on spectral properties of lipoamide dehydrogenase. Attempts to identify the FAD-C4a adduct specie; were unsuccessful. For E.co//enzym e itwa s reportedtha t bothth e reductive reactions to EH2b y lip(SHfea s well as NADHar e very fast at pH 7.6 [63,69] ,an d are infac t complete within hedead-tim e ofth e stopped flow instrument. The further reduction to EH4 proceeds at slower rates, 100s -1 for NADH reduction and via a biphasic process for lip(SHfc reduction at rateso f 0.6 s~1an d0.3 5 s-1 [69]. All species involved incatalysis , identified and proposed,togethe r with species resulting from over-reduction to EH4ar e shown in Scheme 5 (p34) .

Sit* specifically altered enzymes. As already mentioned in the section on spectral properties, site directed mutagenesis experiments on E. coli lipoamide dehydrogenase were reported very recently. The residues that were subjected to mutagenesisar e Cys44,Cys49 , Lys53, Ile184an d His444 [85,59 ,69 ,82 , 83]. Bothcysteine s were replaced by serine [85]. Invie w of thever y important roleo f these residues incatalysi s it is not surprising that these mutated enzymes lack any activity;i nth e physiological reaction [85]. Nevertheless these mutated enzymes still possess some activity withth e artificial electron acceptor ferricyanide when NADH is the dolor. The turnover number is 10% an d5 0 %whe n comparedt o wild-type enzyme for enzyme Cys44->Ser and Cys49->Ser respectively [85]. Lys53form s a saltbridge with Glu188 (see section onthre e dimensional structures) and it was thought that the positive charge inth e pyridine nucleotide binding site was involvep in modulating the redox potential of the FAD.Therefor e Lys53 was replaced by Arg forminima l disturbance [59]. Forthi s mutated enzyme strong inhibition byth e electron donor was reported in both the lip(SH)2/NAD+ and NADH/lipS2 reaction,th e activity^bein g lesstha n 0.2 %o f wild-type enzyme in bothdirection s [59]. Itwa s not reportadwhethe r this inhibition is caused by over-reduction to EH4.

33 —* J> X X r- Q o < < z u. • i i i t

' lff-s J, + X X JO Q Q < < Z U. (0—

u. to—(/)

Scheme 5. Summary of identified and proposed intermediates inth e reaction mechanism of lipoamide dehydrogenase. The physiological reaction is counter clockwise from species 1 to 7. Species8 to 11sho wth eover-reductio n toth e EH4level .Th e arrows signify the charge-transfer interactions and aredraw nfro mth e donort oth e acceptor.

34 The rationale of the mutation Ne184->Tyr isdescribe d inth e section 'Spectral changas inth e presence of pyridine nucleotides' as areth e effects of this mutation on the spectral properties ofth e enzyme.Turnove r ofthi s enzyme inth e lip(SH)2/NAD+ reaction isdecrease d ten-fold with a similar pH dependence as found for the wild-type enzymo [59,69] .Th e NADH/lipS2 reaction isdecrease dtwo-fol d [59,69] . Forthi s enzymf too, inhibition by the electron donor was observed. Similarly as found for the wild-ty^e enzyme NAD+act s as an activator inth e reverse reaction ofth e mutated enzymithoug h the activating effect was maximal at 20u M NAD+instea d of 1m MNAD + as reportedfo r the wild-type enzyme [61, 69].Stoppe dflo w data indicate that inth e reductiono fth e mutated enzyme by NADH,th e first step,reductio n of the FAD,i sonl y slightly decreased but the subsequent hydride transfer to the disulfide is markedly inhibited [69]. Tha mutation His444->Gln was prepared to better understandth e role of the active site ba$e [82,83] .Onl y rapid reaction kinetics of the reduction by lip(SH)2 were reported forthi s mutated enzyme. Itwa s showntha t the apparent rate of reduction by5 molar equivalents of lip(SH)2 was decreasedt o 0.4 s_1 at pH 10.08 while the wild-type enzyme was reduced at a rate of 450-750 s_1 by 2 molar equivalents of lip(SH)2 at pH 9.85 [88],thu s clearly demonstrating the important function of this histidine. The rateo f reduction by lip(SH)2 increased as the pH was increased and this was explained in terms df a better reactivity ofth e substrate inth e mutated enzyme sinceth efirs t pKao f the suqstratewa s 9.3 [51],howeve r eventh e rates at high pHwer ever y slow. Itwa s arguedjthat the severe drop in reduction rate cannot reflect only the function of this histidin^e as a base, but might also reflect deteriorated binding of the lip(SH)2 anion in the apolar binding site andchange s inth e redox potential of the flavin,suggestin g a further fo\efo rth e active site base[83] .

Conformational stability

Initial isolation procedures for the pig heart enzyme contained a heating stept o 62°C [i] demonstrating that the enzyme inth e Eox state isver y heat stable.Thi s was reinforced at 70°C [93].Th e high stability of pig heart enzyme Eox was also demonstrated by ureadenaturation , in6. 5 Mure ath e enzyme did not denature [68]. However, reduction of the enzyme by NADH or lip(SH)2 was accompanied by a strongly decreased resistance towards urea denaturation which ledt o the conclusion that the active site disulfide was an intersubunit disulfide bridge holding the subunits together [68]. In addition it was suggestedtha t the instability could be a result ofth e reduction of the FAD. Later studies aimeda t the preparation of apoenzyme showed that it was possible to obtain monomeric apoenzyme under nonreducing conditions,

35 demonstrating that the active site disulfide bridge is an intrasubunit disulfide bridge [70], a finding that was confirmed later by peptide analysis of the fragment containing the active site disulfide [36]. Tryptic digestion studies with A. vinelandiienzym e have showntha t Eox is not digested in contrast to the N-ethylmaleimide labeled enzyme (the active site disulfide is labeled) indicating a conformational change upon reduction of the disulfide had occurred [94].Recentl y the stability ofth edifferen t redox states of A. vinelandiienzym e was addressed indetai l byva n Berkel andco-worker s [95].A T m of 80°C is reportedfo r Eox which decreases to 39°C and 42°C when reduced with dithionite or NADH respectively. Upon reduction with lip(SH)2 Tm is 53°C. Comparable results were obtained by studying the ureadenaturatio n in 7 Mure aa t 25°C. Upon reduction with NADH or dithionite the rate of denaturation comparedt o Eox increased 200 or 400 fold respectively. Administration of NAD+ha da protective effect onth e denaturation indicating that the formation of EH4i s an important factor in respect to denaturation as already suggested,thoug h rejected,b y Massey andco-worker s [68]. Reduction of the enzyme with lip(SH)2 in 7 Mure a increasedth e rate of denaturation only 2-3 fold. Spectral studies showedtha t in ureath e charge-transfer species was favored,thu s demonstrating that the EH2stat e is notconformationall y unstable. The relatively low thermostability of EH2wa s explained by assuming that att m of EH2th e enzyme becomes EH4du e to over-reduction byth e excess of lip(SH)2 present. Upon proteolytic digestion of Eox, EH2 prepared by lip(SH>2 reduction and EH4 prepared by NADH and dithionite reduction and apoenzyme, only the NADH anddithionit e reduced enzyme and apoenzyme appeared to be significantly digested demonstrating that a conformational change of the enzyme occurs upon reduction from EH2t o EH4whic h promotes dimer dissociation.

Outline of this thesis

At the onset of the investigations described inthi sthesi s progress was being made onth e elucidation of the crystal structure of the Azotobacter vinelandiilipoamid e dehydrogenase. Also the gene encoding this enzyme was cloned in our laboratory. By this, afir m basiswa s laidt o start site directed mutagenesis studies aimed at deepening the insight inth e reaction mechanism of lipoamide dehydrogenase. At the start of this work no site directed mutated mutated lipoamide dehydrogenases were reported though for the E.coli enzym e some mutated enzymes wereannounced . Thefirs t goalwa st o assess the function(s) ofth e active site base,histidine . Therefore the histidine was replaced by other residues and a glutamate, held responsible for affecting the pKa of the histidine was mutagenised. Itwa s expected that

36 the mutationswoul d alter the reaction rates of the different reaction intermediates and that th0se intermediates could be identified andstudied . Chapter 2describe s the steady state spectral properties and in Chapter 3th e steady state and rapid reaction kinetics andsom e rapid reaction spectral properties ofth e wild- type arjd mutated enzymes are presented. Sinie modeling studies suggested that Tyr16 was involved in substrate binding,thi s residua was replaced with other residues in order to stabilize the binding of the substrata, lipS2,an d hence to be ablet o grow crystals ofth e enzyme withth e substrate bound.Th e results obtained with these mutated enzymes are presented in Chapter 4. Prejiminary experiments with A. vinelandii, E.coli an d P. fluorescenslipoamid e dehydrogenase showed strong reactivity of the latter enzyme with antibodies raised against A. vinelandiienzym e while nocros s reactivity was observed with the E.coli enzymi. This indicates that high sequence homology is present between the A. vinelandiian dth e P. fluorescens enzymes andtha t the P. fluorescens enzyme constitutes a natural 'mutated' effective lipoamide dehydrogenase. Cloning and sequerjce analysis of this gene,describe d in Chapter 5 opensth e way to site directed mutagenesis. In Chapter 6 acharacterizatio n ofth e P. fluorescenswild-typ e enzyme is presented.

37 References

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41 Erratat o Chapter 2.

1) Pro455->Ala should read Pro451->Ala.

2) The multiplication factor 10"6 (p47 ) forth e quenching constants (KQ) shouldb e omitted.

42 Eur. J. Biochein. 202,863-87 2 (1991) © FEBS 1991

Chapter 2

LipoaimldeDehydrogenas e from Azotobacter vinelandii: site-directed mutagenesis of theHfc450-Glu45 5dia d Spectral propertieso f wild type and mutated enzymes

Jacques BENEN, Willem van BERKEL, Zdzislaw ZAK, Ton VISSER, Cees VEEGER and Arie de KOK Department of Biochemistry, Agricultural University, Wageningen, The Netherlands

(Receiced ApJil 22/July 30,1991) - EJB9 1052 2

I Three amino acid residues in the active site of lipoamide dehydrogenase from Azotobacter vinelandii were replaced by other residues. His450, the active-site base, was changed into Ser, Tyr and Phe. Pro451, in cis Conformation, was changed into Ala. Glu455 was replaced with Asp and Gin. Absorption, fluorescence and CD spectroscopy of the mutated enzymes in their oxidized state (E0„) showed nly minor changes with respect to the wild-type enzyme, whereas considerable changes were observed in the ;pectra of the two-electron-reduced (EH2) species of the enzymes upon reduction by the substrate ihydrolipoamide. Differences in extent of reduction of the flavin by NADH indicate that the redox potential of :he flavin is altered by the mutations. Enzyme Pro455->Ala showed the greatest deviation from wild type. The mzyme is very easily over-reduced to the four-electron reduced state (EH4) by dihydrolipoamide. This is probably ue to a change in the backbone conformation caused by the cis-trans conversion. From studies on the pH dependence of the thiolate charge-transfer absorption and the relative fluorescence f EH2 of the enzymes, it is concluded that mutation of His450 results in a relatively simple and easily interpreted istribution of electronic species at the EH2 level. For all three His450-mutated enzymes an apparent pK,, near 5.5 is calculated that is assigned to the interchange thiol. A second apparent pKi2 is calculated of 6.9, 7.5 and 7.1 for the His450-»Phe, -Ser and -Tyr enzymes, respectively, and signifies the deprotonation of the tautomeric equilibrium between the interchange and charge-transfer thiols. The difference in apparent pKl2 values between the His450-mutated enzymes is explained by changes in micropolarity. At the EH2 level the wild-type enzyme consists of multiple electronic forms as reported for the Escherichia coli enzyme [Wilkinson, K. D. and Williams C. H. Jr (1979) J. Biol. Chem. 254, 852-862]. Based on the results obtained with the His450-mutated enzymes, it is concluded that the lowest pK„i s associated with the interchange thiol. A model for the equilibrium species of the wild-type enzyme at the EH2 level is presented which takes three pX, values into account.

The results of the pH dependence of the electronic species at the EH2 level of Glu455-mutated enzymes essentially follow the model proposed for the wild-type enzyme. However mutation of Glu455 shifts the tautomeric equilibrium of EH2 in favor of the charge-transfer species. This effect is explained by impaired ion-pair formation between the protonated His and the interchange thiolate and a lowering of the pKt of His450.

Lipoamile dehydrogenase is the flavoprotein component it catalyses the oxidation by NAD + of a reduced lipoyl group of the pyruvate, oxoglutarate and branched-chain oxoacid which is covalently attached to the core compound of these dehydrogenase complexes [1,2]. In the physiological direction complexes. This lipoyl group can be replaced with free Correspotxience to Dr. A. de Kok, Department of Biochemistry, lipoamide. Lipoamide dehydrogenase belongs to the family of Agricultural ' Jniversity, Dreyenlaan 3, NL-6703 HA Wageningen, dimeric flavoenzymes that contain a disulfide participating in The Netherlaiid s catalysis [1], Other members of this family are glutathione Abbrevialions. Eox,oxidize d (mutated) lipoamide dehydrogenase; reductase [1], mercuric ion reductase [3], trypanothione re­ EH2,two-cleotro nreduce dlipoamid edehydrogenase ; EH4, four-elec­ ductase [4]an d thioredoxin reductase [5]. tron reduced Dipoamide dehydrogenase; LipS2, D,L-6,8-thioctic acid Extensive studies on lipoamide dehydrogenase have been amide, lipoanide; Lip(SH)2, reduced lipoamide; Nbs2, 5,5'- performed on the enzymes as isolated from pig heart, rat liver dithiobis(2-nirobenzoate); AcPyAde*, 3-acetylpyridine-adenine and Escherichia coli [6—14] . Potentially, the enzyme can take dinucleotide; PPj/EDTA, sodium pyrophosphate/EDTA bufTer, pH up four electrons/monomer, but during catalysis the enzyme 8.0; FADH~jreduced flavin; F1H~, EH2 specieswit hflavin reduced ; FL, fluorescent species; CT; charge transfer species. shuttles between the oxidized state (Eox) and the two-electron- Enzymes!Lipoamid e dehydrogenase, NADH:lipoamid e oxido- reduced state (EH2) [15]. The four-electron reduced state reductase (EC 1.8.1.4); DNA restriction endonuclease Notl, A/ml, (EH4) is inactive and originates from the dead-end complex Sstl, Smal (EC3.1.24.4) ;DN A polymerase I(Kleno w fragment) (EC EH2 •NAD H [12].Al l three lipoamide dehydrogenases were 2.7.7.7) shown to act according to a ping-pong mechanism in the

43 864 physiological reaction [6,8,14]. A reaction mechanism involv­ the polylinker from Bluescript KS was cut out by digestion ing a base in the active site that becomes protonated during with Sstl and Kpnl and ligated into Sril/AT/wiI-digested catalysis was proposed [16]. Initially a histidine was put for­ M13mpl8/19. As the insertion in the M13 vectors is in frame ward as the base. Later this idea gained strong support from with the lacZ gene, the blue/white feature of the Ml 3 vectors studies with a bifunctional arsenoxide [17]. isretaine d while the number of unique restriction sites suitable In the three-dimensional structure of E0„fro m Azotobacter for cloning is expanded. vinelandii, refined to 0.22 nm, a His residue is present in the Wild-type and mutated lipoamide dehydrogenase genes active site with the carboxylate function of a Glu in close wereexpresse d in transformed E. coliTG 2 grown in 5-1 batches contact [18]. In Eox the histidine is closer to the N-terminal of tryptone/yeast medium containing 75 ug/ml ampicillin at Cys48, the interchange thiol in the reductive reaction, than to 37° C with vigorous aeration. Cys53, the charge-transfer thiol. An identical feature was found in the related enzyme glutathione reductase in both the Construction of mutants oxidized and reduced state [19— 21]. It was also found that the active site is composed of both subunits. The residues that were to be replaced, His450 with Tyr, Both kinetic and spectral studies revealed that the active- Phe or Ser, Pro451 with Ala and Glu455 with Gin or Asp, are site His of lipoamide dehydrogenase and glutathione re­ all located in the C-terminal part of the enzyme. Therefore a ductase in EH2 showed a high pATa of about 8.5 whereas the 420-bp Notl —Kpnl fragment containing the coding region for charge-transfer thiol showed an unusually low pK„, about 4.5 amino acid residues 391—47 7 (C-terminus) and the putative [9. 13]. It has been argued that the pK„ shifts were due to transcription terminator was subcloned into M13mpbl9 and mutual influences [9, 13]. In order to assess the role of the used for mutagenesis according to the method described [24]. active-site histidine (His450) and the nearby glutamate Single-stranded uracil-containing DNA was isolated when the (Glu455), site-directed mutagenesis experiments were ratio of infectivity of the phage for E. coli RZ1032 to E. coli performed. His450 was replaced with Ser, Tyr or Phe and TG2 was higher than 106:1. Glu455 was replaced with Asp or Gin. Apart from these Single-stranded uracil-containing DNA was used for 'all- residues, we were also interested in the role of the backbone the-way-round' oligonucleotide directed in vitro mutagenesis. in this part of the active site. In the crystal structure of E„. To 1 ug single-stranded DNA (ssDNA), 10 pmol of phos- Pro451 is in cis-conformation thereby positioning the back­ phorylated oligonucleotide was annealed. The polymerase bone carbonyl of His450 towards the proton at N3 of the mixture contained 1 g annealed ssDNA, 250 uM of each of flavin (Mattevi, personal communication). This might have the dNTPs, 5 mM dithiothreitol, 10.0 mM Tris/Cl, 10.0 raM an influence on the redox properties of the flavin. Therefore MgCl2, pH 8.0, and 1.5 U Klenow DNA polymerase I in a Pro451 was replaced with Ala. final volume of 20 ul and was incubated for 2 h at 16 C. After In this paper we report on the pH-dependent spectral addition of 1 ul 10 mM ATP and 8 U T4-DNA ligase, the properties of the wild-type and mutated enzymes of Azoto­ incubation at 16°C was continued for 20 h. The reaction bacter vinelandii lipoamide dehydrogenase. mixture was used to transform E. coli TG2. Phages obtained from the transformation were grown and ss DNA was isolated and screened for the mutation according to the Sanger MATERIALS AND METHODS dideoxy-chain-termination method using deaza-dGTP [26]. Clones carrying the mutation were sequenced entirely to General screen for undesired mutations. Subsequently double- Restriction endonucleases were from Boehringer or stranded DNA (dsDNA) was isolated and from this the mu­ Bethesda Research Laboratories (BRL). E. coti DNA poly­ tated Notl —Kpnl fragment was isolated as a slightly longer merase I (Klenow fragment) was from BRL as was T4-DNA Notl —Smal fragment. In order to ligate the mutated fragment ligase. NAD+ (grade I), NADH (grade I) and all dNTPs and back into the gene, plasmid pAW104 was digested with Notl ddNTPs were from Boehringer. Lipoamide, 5,5'-dithiobis(2- and Smal, deleting the entire downstream region of the Notl + nitrobenzoate) (Nbs2), the NAD analog 3-acetyl-pyridine- site at position 2167. The mutated Notl —Smal fragment was adenine dinucleotide (AcPyAde+) and biological buffers were then ligated into the partial gene yielding an intact gene obtained from Sigma Inc. Dihydrolipoamide was synthesized carrying a point mutation. Proper 'in-frame' ligation of the according to Reed et al. [22 ] and the concentration was fragment was checked by subcloning the entire insert of the determined with Nbs2. Oligonucleotides for mutagenesis were new construct into M13mpl9 and sequencing the ligation synthesized with a BioSearch Inc. oligonucleotide synthesizer. point at the Notl site using the oligonucleotide used for mu­ [K-32P] dATP (3000 Ci mmol1; 11.1 TBq mmol-1) was pur­ tagenesis as a sequencing-primer. chased from New England Nuclear (NEN). Chromatography Sequencing of the mutated Notl —Kpnl cassette clearly resins were from Pharmacia. All chemicals used were of ana­ demonstrated that the desired mutations were made. lytical grade. E. coli TG2, a rec A' version of TGI [A(lac-pro), thi, sup Analytical methods E,\Res~ Mod- (k)] F' (traD36, proA* B+, foe/" Z,dM l 5)] was used throughout asa host for cloning and expression of cloned Large-scale isolation of the enzymes (up to 5-1 cultures) enzymes [23]. E. coli RZ1032 [SupE, dut', ung~] [24]wa s used was performed essentially as described [25] with one slight for generation of uracil-containing single-stranded DNA. modification. The Sepharose 6B enzyme eluate was concen­ Plasmid pAW104 containing the A. vinelandii wild-type trated using an ultrafiltration set (Amicon) equipped with lipoamide dehydrogenase gene has been described earlier [25] a YM30 membrane. The concentrate was then exhaustively and was kindly provided by Mr A. H. Westphal. dialyzed against 50m M potassium phosphate, 0.5 mM EDTA, Vectors M13mpbl8 and -19 were constructed by cloning pH 7.0, and stored as 1.0-ml aliquots (5— 7 mg/ml) under the polylinker from the plasmid vector Bluescript KS liquid nitrogen until use. The expression of the mutated (Stratagene Inc.) into vectors M13mpl8 and -19. For this lipoamide dehydrogenases was as high as for over-expressed

44 865 wild-type enzyme yielding 250 mg (mutated) lipoamide spectra were recorded until changes were complete. Additions dehydirogen ase . /5-1cultur e of transformed E. coli TG2. of dihydrolipoamide were stopped after reaching maximal FPLC analytical gel filtration was done as described [27]. charge transfer. Data, corrected for dilution, were analyzed For specl:al studies, enzyme solutions were freshly thawed with a Levenberg-Marquardt algorithm provided by the pro­ from liquid I itrogen and subsequently passed over BioGel P- gram 'Igor' from WaveMetrics Inc. run on an Apple 6DG10 chatg i ;e to the appropriate buffer, Macintosh. Routines were written to analyze the data for one, When n edording spectra under anaerobic conditions, en- two or three pKa values where the routines also varied the zyme solutions were transferred to anaerobic cuvettes that absorption coefficient to give best fit to the data. In a separate were closed 'vit h Subaseals. The cuvettes were subsequently set of experiments the pH-dependent fluorescence of both Eox evacuateds < times and gassed with argon. The argon was and EH2 (10 pM concentration) was studied. The excitation passed over a catalyst (BASF, R 3-11) to remove oxygen wavelength was 458 nm (4-nm bandpass) and the emission followed by I assage through an illuminated solution contain- wavelength 525 nm (4-nm bandpass). All titration experiments ing reduced methyl viologen, EDTA and proflavin (3,6- were performed at 25°C . diaminoacridine) to remove the last traces of oxygen. Solu- tions that w^re used to titrate/reduce the anaerobic enzymes Activity determinations were: deoxygi nated in an identical way. + NAD and NADH concentrations were determined in 50 Enzyme joncentrations were determined spectrophoto- mM sodium pyrophosphate, 0.5 mM EDTA, pH 8.0 (PPj/ metrically usngsa molar absorption coefficient e458 = 11300 1 -1 1 EDTA), using Ala is at 453 nm). 4 termined by converting AcPyAde* into AcPyAdeH (£ = sorption spectra were recorded at 25°C , on an 363 Visible al 9100 M"1 cm"1) using lipoamide dehydrogenase from -2a or computer-controlled Aminco DW-2000 Aminco D\ty Pseudomonas fluorescens [28]: to 950 ul PPi/EDTA, 20 ul eter. The scan speed was 5 nm s"'. Typically spectrophi dihydrolipoamide (final concentration 1.0 mM) and 20 ul of solutions were used. 45 uM enzynle several dilutions of the AcPyAde+ stock solution were added. Fluoresa nee emission spectra were recorded on an auto- The reaction was started with 10 pi of an 0.5 mg/ml enzyme mated Amin«o SPF-500 fluorimeter (Olis Inc., Jefferson, GA) solution and followed at 363 nm until changes were complete. at 25 C. Lipoamide dehydrogenase from Ps.fluorescens was used be­ Circular flichroic (CD) spectra were obtained with a Jobin cause this enzyme appeared to be insensitive to inhibition by Ivon mark V dichrograph connected to an IBM-PC. Quartz AcPyAdeH. cells with 1.1 cm path length were used in the wavelength + The forward reaction dihydrolipoamide/NAD was deter­ region 300 500 nm. In the ultraviolet range (190-300 nm), mined routinely in PP|/EDTA at 25°C . In a standard assay cells with 0.1 cm path length were used. The cell compartment + 1.0m M NAD and 1.0m M dihydrolipoamide were used and was thermostatted at 25° C and purged with nitrogen. The the formation of NADH was monitored at 340 nm. In some scan speed 1 nm s~' . The spectra reported are averages + cases AcPyAde* instead of NAD was used as electron ac­ of three indq jendent experiments on different samples of the ceptor and the formation of AcPyAdeH was followed at 363 same prepara tion All spectra were corrected for the appropri- nm. ate blank sol 1utions , recorded in the absence of enzyme. The values are e: x jressed in terms of molar ellipticity [6] and were calculated ua ng the equation: RESULTS 3300 General properties of mutated lipoamide dehydrogenases -XAE m Ix c The monomer/dimer ratio of the mutated enzymes in solu­ tion was estimated using FPLC. Upon elution of the mutated where Ae is the circular dichroism observed, / is the path enzymes on Superose-12, all profiles show one symmetrical length of the cell in cm and c is the molar concentration of peak with the same retention volume as found for dimeric the enzyme aiiallyzed; . Below 240 nm the ellipticity is expressed wild-type lipoamide dehydrogenase [29]. In this experiment it as mean resiilua l ellipticity [S\m, (A. vinelandii lipoamide de- was also seen that no FAD was liberated indicating that the hydrogenase is composed of 476 amino acid residues [25]). FAD binding of the mutated enzymes is not detectably altered In order to 0btai n spectra of two-electron-reduced enzymes, when compared to wild-type lipoamide dehydrogenase. enzyme solul ions were made anaerobic as described above Considerable differences in activity exist among the several and titrated vith a 5 mM dihydrolipoamide stock solution, (mutated) enzymes. The enzymes His450->Tyr, His450->Phe The reductiod of the enzymes was monitored by recording the and Pro455->Al a are almost completely inactive while enzyme absorption s[ectr a< ove r 320— 60 0 nm. When the characteristic His450-*Ser shows 0.5% activity of the wild-type enzyme. spectrum of :H2 was reached, CD spectra were recorded in The Glu455-mutated enzymes initially show kinetic traces the range 308 —60 0 nm. The enzyme concentration varied comparable to wild-type lipoamide dehydrogenase but, due from 1 uM (190- 240 nm) to 50 uM (300- 600 nm) in 50 mM to severe inhibition by the product NADH, kinetics can not potassium pi 1 osphate, 0.5 mM EDTA, pH 7.0. be studied. When using the NAD+ analog AcPyAde* as For pH < ependent titrations the following buffers were an electron acceptor, the inhibition is almost negligible. The used: 100 potassium acetate (pH 4.5—p H 5.3), 100 mM activity is 1.6%an d 1.3% of wild-type lipoamide dehydrogen­ )mlf + Mes (pH 5 6.5), 100 mM Hepes (pH 6.7-7.8) and 100 ase using AcPyAde forGlu455->Aspand Glu455->Gin mu­ mM Hepps (!>H!8. 0 —8.7) . The ionic strength was adjusted to tated enzymes, respectively. 150 mM witl 1.0 M K.C1. Enzyme solutions (45 jiM concen- tration) were nade anaerobic and subjected to titration using a Spectral properties syringe equipped with a dispenser enabling gas-tight Hal ultons For wild-type and mutated enzymes the ^2S0/^458 ratio the addition if defined aliquots of titrant. After each addition is 4.8—4.9. This demonstrates that the enzymes are saturated

45 866

A flavin and a thiolate that originates from reduction of the disulfide bond [30].Thi s complex will be referred to as the A 530-nm charge-transfer complex. The absorption at 530 nm is less intense than that of the EH2 form of the pig heart '5.0 enzyme but more intense than that of the E. collenzym e [7, •H1 A 14, 16]. ^L-IX At equilibrium, upon titration of the wild-type enzyme with one equivalent of NADH, the electrons are shared be­ tween the sulfur atoms, as evidenced by the 530-nm ab- sorbance, and the flavin molecule, as evidenced by the 750- nm absorbance, indicating FADH"/NAD+ charge-transfer complexformatio n [15].Not etha thal f the750-n mabsorbanc e is formed upon addition of the first equivalent of NADH. Furtherreductio nt oEH 4 resultsfro m theadditio no fa further equivalent of NADH (Fig. 1A) . With enzymes Glu455-»Asp and Glu455-»Gln the re­ duction to EH2 by dihydrolipoamide is also complete within mixing time.Th e spectra of EH2 (Fig. 1 B and 1C)sho w530 - nmabsorptio n remarkably higher than thewild-typ eenzyme . Upon titration ofenzym eGlu455-»As p with NADH, the absorption at 750 nm is considerably higher than that ob­ served for wild-type enzyme. The 530-nm absorbance ob­ served with the wild-type enzyme, after the initial addition of NADH, is not observed to the same extent in enzyme Glu455->Asp. With enzyme Glu455->Gln the absorbance at 750n mi scomparabl e towil d typebu t the 530-nm absorbance is very prominent (Fig. 1C). It is also noteworthy that the 400 500 600 700 400 500 600 700 extent of reduction of the FAD varies between wild type and enzymes Glu455->Asp and Glu455-»Gln after the addition WAVELENGTH (nm) of one or two equivalents of NADH (Fig. 1B and 1C). This difference suggests that the FAD in enzyme Glu455->Asp is Fig. 1. Visible absorption spectra ofA. vinelandii wild-type lipoamide moreeasil yreduce d inth epresenc eo fNA D+ than isth e FAD dehydrogenase andmutated enzymes. Spectra wererecorde d at 25° C in wild-type or Glu455->Gln enzyme. inPPi/EDT A(1 ) before ( )(E„ )an dafte r theadditio n of(2 )5. 0 Like the wild-type lipoamide dehydrogenase and the mol/moldihydrolipoamid e( )(EH 2) and before and after the addition of (3) 1.0 mol/mol NADH ( ) and (4) 2.0mol/mo l Glu455-mutated enzymes, the His450-»Ser enzymei s rapidly NADH (-• • •)- Spectra shown were recorded when changes were reduced by dihydrolipoamide and forms a stable EH2 (Fig. complete.(A )Wild-typ eenzyme ;(B )enzym eGlu455-*Asp ;(C ) en­ 1D). The spectrum is very similar to the Glu455-mutated zyme Glu455-*Gln; (D) enzyme His450-*Ser; (E) enzyme enzymes, showing an intense absorption band at 530 nm, His450-»Tyr; (F) enzyme His450-»Phe. For the mutated enzymes muchhighe rtha ntha to fwild-typ ea tthis pH .Initia l reduction His450-»Tyran dHis450-*Ph ea spectru mi spresente d(5 )tha tdepict s of the His450->Ser enzyme by NADH at pH 8.0 is charac­ the final spectrum that was isosbestic upon reduction with dihydrolipoamide (-•-•-) terized bya high 530-nm absorbance with only small changes in FAD absorbance and high absorbance at 340 nm due to free NADH. Eventually, after addition of excessNADH , the FAD becomes fully reduced and the spectrum shows a high with FAD and that the molar absorption coefficients for all absorbance at 750 nm indicative of the formation of an enzymes are almost identical. FADH"/NAD+ charge-transfer complex (results not shown). Fig. 1 shows the visible absorption spectra of wild-type The small extent of reduction of FAD of the enzyme and mutated enzymes. In the oxidized state flavin spectral His450->Ser suggests that the FAD is not as easily reduced properties of the enzymes are almost indistinguishable. The in thepresenc e of NAD+ as in thewild-typ eenzyme . enzymes His450->Tyr and His450->Phe show a more pro­ The His450-»Tyr and His450-*Phe enzymes are very nounced shoulder at 480 nm indicating a relatively more slowly reduced by dihydrolipoamide. The course of titration apolar environment of the flavin-isoalloxazine ring. This fea­ to EH2 is different with respect to the wild-type enzyme and ture isprominen t indifferenc e spectra (not shown).Th espec ­ mutated enzymes described above (Fig. 1E and 1F). While trum of enzyme Pro455-»Ala (not shown) however shows the spectra of thepreviousl y described enzymesar e isosbestic less pronounced vibrational fine structure. The absorption during reduction with dihydrolipoamide until maximal 530- maximum at 453 nm and closer resemblance of the spectrum nm charge transfer is reached, spectra of these enzymes are to that of free FAD are indicative of a more polar only initially isosbestic. Before maximal charge-transfer ab­ microenvironment of the isoalloxazine ring. sorbance isreache d the isosbestic points shift, indicating for­ At pH 8.0, thespectru m of wild-type Eox(Fig . 1 A)closel y mation of other reduced species. This effect is most pro­ resembles the spectra of pig heart and E. coltlipoamid e de­ nounced with enzyme His450-»Tyr: changes in spectra are hydrogenase [lj. Upon addition of a slight molar excess of isosbestic until a certain amount of charge-transfer ab­ dihydrolipoamide, EH2 isforme d within themixin gtime .Th e sorbance is reached (see Fig. 1E) which is retained for a few spectrum is characterized by a shift of the major flavin peak minutes. After this time interval, the charge-transfer ab­ toshorte rwavelengt h and atypica l absorption at 530n m that sorbanceincrease sfurthe r withsimultaneou sshif t ofisosbesti c isascribe d to acharge-transfe r complex between the oxidized points. With the enzyme His450->Phe this biphasic charge-

46 867

Table 1.Relative FAD fluorescence quantum yield of A. vinelandii wild-type lipoanide dehydrogenase andmutated enzymes The excitation wavelength was 450 nm and the emission wavelength 525 nm. All ex|icriment swer eperforme d at2 5° Ci n 50m M potassium phosphate, 0. I mM EDTA, pH 7.0. Enzyme concentrations were 2 jiM based on Ir AD absorption

Enzyme FAD quantum yield (relative to wild type) -t- Wild-type 1.00 Glu455->Asp 0.63 Glu455->Gln 1.14 0 100 200 300 400 500 His450->Phe 0.73 [Kl] (mM) His450->Ser 0.80 Fig.2 .Stern-Volmer plot of the FAD fluorescence quenching by iodide of His450-»Tyr 0.80 A.vinelandi i wild-typelipoamide dehydrogenase and mutated enzymes. Pro455->Ala 0.64 Thedat a arecorrecte d for salteffects . Theexcitatio n wavelength was 450 nm and the emission wavelength 525 nm. The enzyme concen­ trations,i n5 0m Mpotassiu m phosphate,0. 5m M EDTA,p H 7.0wer e adjusted to 2.0 uM (FAD absorption).The three His450enzyme s and enzyme Pro455->Ala gave nearly identical results and are presented transfer is not observed. Another typical aspect of ason ecurv e(O) . Wild typeenzym e (•); enzyme Glu455-*Gln (A); the spectrumjof EH2 of these two His450-mutated enzymes is enzyme Glu455^Asp (Q); FAD (A) the large shilt of the 458-nm peak of the flavin to 430-435 nm and a slift of the maximum from 530 nm to 540 nm, resulting in a clear resolution of the charge-transfer ab- sorbance. of the mutations is very local and does not disturb the overall Unlike :he slow reduction of His450->Phe and structure of the enzymes in their oxidized state. This con­ His450 Tyrjenzymes by dihydrolipoamide, the reduction by clusion is supported by the far-ultraviolet CD spectra (see NADH equilibrium approximately as fast as in wild- below). type lipoam^de dehydrogenase. The extent of reduction in Flavin accessibility of the mutated and wild-type enzymes both mutal ed enzymes clearly differs however. In was probed by iodide fluorescence quenching. Fig. 2show s the His450-Phe enzyme, FAD is almost fully reduced after the Stern-Volmer plots obtained after correction for salt effects. addition of equivalents NADH. In enzyme His450->Tyr, Obviously, mutation of Glu455 influences the accessibility of FAD is reduced to a lesser extent, slightly more than enzyme the flavin for iodide. Slight quenching is observed in the wild- His450->Ser. Thus apparently the ease of reduction of FAD type and mutated enzymes that still possess Glu455, yielding 6 1 of the mutat4d enzymes by NADH in the presence of NAD + quenching constants (KQ) of approximately 1xl0~ M" . is different ai l id i might be related to shifts of either or both the Similar results were reported for the pig heart enzyme [31]. redox potentjals of the disulfide and the FAD. Strong quenching at low KI concentrations was seen in the 6 1 Enzyme 'ro455->Ala behaves quite differently during re­ Glu-455 mutated enzymes (KQ = 15xl0" M" for 6 1 duction. Upoi in reduction with dihydrolipoamide no 'stable' Glu455^Gln and KQ = 11 x 10" M" for Glu455-Asp). This indicates that the carboxylate function of Glu455 is the EH2 species formed. The enzyme iscompletel y reduced after addition of iwo equivalents of dihydrolipoamide. Titration repelling force, preventing iodide from entering the active site. with dihydw lipoamide shows that initially the disulfide is The nonlinear behavior of the plots at higher KI concen­ reduced yiel«in g a transient 530-nm charge-transfer absorp- trations for the Glu455-mutated enzymes suggests minimally tion. After a few seconds charge transfer decreases with con- two possible conformations of the flavin binding region, one comitant fon lation ofreduce d flavin (results not shown). This accessible and one less accessible. The presence of monomers result indical s that in enzyme Pro455->Ala the electrons are was excluded by the fact that the F0/F ratio as a function of easily Iransfe rred from the sulfurs to the flavin and suggests enzyme concentration did not change at KI concentrations that the redqx potentials of the disulfide and the flavin are where strongest quenching was observed (not shown). At pre­ very close, n modelling studies on the crystal structure of sent it is not clear whether this different quenchability is an lipoamide dehydrogenase, it is deduced that the mutation intrinsic property of the Glu455 mutated enzymes or that it is Pro455->Ala probably leads to a changed position of the induced by the iodide present in the active site which might backbone re)ultin g in a breakage of the flavin N3-carbonyl cause cooperativity of the active sites. oxygen inter iction. Simultaneously the imidazole of His450 Possible charge effects of the quencher were probed using he plane Cys48-Cys53-His450-Glu455 (Mattevi CsCl as a quencher. Free FAD was not as strongly quenched shifts out of 6 personal corrjmunication) as found for KI; KQ (CsCl) = 0.5 x 10" M~' and KQ (KI) = 28x 10~6 M"1. Instead of quenching, a slight stimulation of fluorescence was observed with CsCl, of the same magnitude Fluorescence properties as found for NaCl and KC1. The shape of the fluorescence emission and excitation spectra of thi mutated enzymes are very similar to the shape Circular dichroic spectra of the spectn of wild-type lipoamide dehydrogenase (results not shown); only differences in relative quantum yield are In the ultraviolet region (190— 30 0 nm), only minor observed (se< Table 1). The shifts of the emission and exci- changes were recorded in the CD spectra of Eox. In the far- tation of enzyme Pro455->Ala are in accordance ultraviolet, the CD spectra are essentially identical, indicating with the : shifs of the two major flavin peaks of the visible that no significant changes in the overall secondary structure absorption sjiect:rum . From this we conclude that the effect between mutated and wild-type enzymes have occurred. In

47 868

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E /

"*£ if ^ -2 ij m -3 '/ '1 x -4. 1 © i -5 ' 260 280 300 400 500

WAVELENGTH Inml WAVEIENOTH Irani Fig. 3.Circtilar-dichroic spectra of A. vinelandii wild-typelipoamide dehydrogenase and some mutated enzymes. CDspectr ao fwild-typ eenzym e andenzyme s deviatingmos tfro m wild-type.Condition san d analysisar ea sdescribe di nMaterial san d Methods.( A— C ) CDspectr a ofE ox; wild-typeenzym e( );enzym e His450->Phe( );enzym eGlu455->As p( ).(A )Far-ultraviole t spectra (190— 25 0nm )o fth e enzymesar eessentiall yidentical ,therefor e onlywild-typ eenzym ei sshown . (B) Aromaticregio no fth espectru m (240— 30 0 nm); wild-type enzymean d enzymeHis450->Ph ear eidentical ,onl ywild-typ ean d Glu455->Asp enzymear eshown .(C )Visibl e region (300— 60 0nm) . (D) Visibleregio n(30 0— 60 0 nm)o fEH 2 ofwild-typ eenzym ean d twomutate denzyme sdeviatin gmos tfro m wild-typeenzym e(-——) , enzymes His450-.Phe( )an dGlu455-.Gl n( ) thenear-ultraviole t only theamplitud e of the positive Cotton showing that the thiolate-FAD charge-transfer complex has effect (,lm„ = 274nm )varie sslightl yi nal lenzymes .N odirec t rotational strength. The thiolate-FAD donor-acceptor inter­ relation between thearomati c amino acid residue engineered, action may lead to a shift of the (perturbed) first electronic e.g. in the His450-mutated enzymes, and the Cotton effect transition of the FAD as suggested by the strong negative could be established. Fig. 3A— C depicts the CD spectra of dichroism around 410 nm. The dichroism of the second ab­ the Eoxo fwild-typ e and twomutate d enzymes,Glu455-»As p sorption band, almost identical inal lenzymes ,decrease s with and His450->Phe, which show largest deviations from the respect to Eoxproportiona l toth e absorption spectrum witha spectrum of the wild-type enzyme. The CD spectra are very concomitant shift of themaximu m to 350nm . similar to the published spectra of the pig heart enzyme[32 , 33].I n thevisibl eregio n (300—60 0 nm),th eC D spectra ofal l mutated enzymesan d wild-typeenzym esho w a positive Cot­ pH dependence ofabsorption andfluorescence properties ton peak at 370nm ,an d two unresolved CD bandsa t 465n m of'EoxandEH2 and 490 nm, both either positive or negative, depending on In order to gain more insight into the influence of the the enzyme used. The positive bands at 370 nm, with nearly mutations on the catalytically important protonation states identicalellipticity ,correspon d toth esecon d absorption band of EH2, the pH dependence of the spectral properties of the of FAD. The two bands at 465 nm and 490 nm arise from enzymes was studied. For EOI of wild-type and mutated en­ vibronic transitions of the first absorption band. The major zymes,th emola rabsorptio ncoefficien t appearst ob eindepen ­ differences inth eC Dspectr a of E0,ar eobserve d at thelowes t dent of pH. However the charge-transfer absorbance at the energy absorption band of protein-bound FAD. The EH2 level ishighl ydependen t on pH and can be regarded as dichroisminthi srang evarie sfro m —3. 4x 103degM~1cm"1 a monitor of protonation states at this level of reduction. for enzymePro455->Al a to4. 8x 103degM~'cm_1 for wild- Dihydrolipoamide was used as reducing agent since with type.Th edifference s inmola rellipticit yindicat esligh tchange s dithionite no stoichiometric reduction could be achieved. inth e microenvironment of the flavin. Fig. 4 shows the 530-nm absorbance of wild-type and TheC Dspectr a of EH2 (Fig. 3D showsspectr a that devi­ mutated lipoamidedehydrogenase s asa function ofpH . Dur­ ated most from wild-type, Glu455-»Gln and His450->Ser), ing the course of the titrations the isosbestic points around indicate that differences occur between the enzymes at the 440n m shifted before maximal charge transfer was reached, lowest energycharge-transfe r transition of the FAD. Anega ­ indicatingth eformatio n ofothe rspecies .Th eshif t isinterpret ­ tive dichroism of different amplitude was observed in the ed as theformatio n of an EH2 species in which the electrons range around 400 nm and positive dichroism was observed aretransfere d toth e flavin [F1H~]and/o r disproportionation above44 0nm .Th e positivedichrois m extends far beyond the ofEH 2 into EH4 and EOIanalogou st o theE. coli enzyme [11]. upper limit of dichroic activity of the flavin as found in E„ The disproportionation might be enhanced due to the fact

48 869

Table 2. ApparentpK a valuesand absorption coefficients of the charge-transfer absorption of two-electron-reduced A. vinelandii wild-type lipoamidedehydrogenase andmutated enzymes Experimental^condition s and analysis are described under Materials and Methods. Data are presented in Fig.4

Enzyme p*.i PK.2 pK.3

M ' cm l M"'cm"' M-'crn"1 Wild-type 4.8 + 0.3 330 + 150 6.7 + 0.3 1060 + 170 8.7 + 0.5 1700 ± 800 Glu455->Aspi 5.9 + 0.1 2840 + 60 Glu455-.Glnj 5.6 + 0.1 3010 + 20 6.9 + 0.4 600+80 His450-Ser ' 5.5 ± 0.1 730 + 80 7.5 + 0.1 2730 + 80 His450-Tyr j 5.5 + 0.3 1270 + 80 7.1 + 0.1 2580 + 260 His450->Phe 5.3 + 0.2 1780 + 250 6.9 + 0.1 2150 + 230

-B The data of the His450-mutated enzymes are best fitted A/O5*- with two apparent pA, values (Fig. 4B). The pKt2 value shifts E 3 - to lower pH, proportional to the polarity of the side-chain engineered. In addition to this, the absorption coefficient of 2 S.2 the charge-transfer absorption is increased when compared to the maximal values reported for pig heart and E. coli enzyme -y. [11, 14]. In order to gain more insight into the groups that are 111 i i i associated with the apparent pKt values summarized in Table 2, the pH dependence of the fluorescence properties of the pH enzymes were studied. Since the presence of a thiolate in Fig. 4. pH dependence of thethiolate charge-transferabsorbance of A. the vicinity of the FAD acts as a fluorescence quencher, the vinelandiiwii type lipoamidedehydrogenase and mutated enzymes. fluorescence at the EH level may be regarded to as a monitor Conditions ai analysis are as described in Materials and Methods. 2 ids for the deprotonation of the charge-transfer thiol. The curves difcrwn through the data are calculated from the apparent For the E studied (wild-type, Glu455-»Gln, Glu455 p£a values and absorption coefficients at 530 nm obtained from the 0I best fit (Tabic 2). (A) Wild-type enzyme (•); enzyme Glu455-*Asp ->Asp, His450-»Phe and His450->Ser) the fluorescence is (D); enzyme Glu455->Gln (•). (B) Enzyme His450->Ser (A); en- dependent on pH, unlike the E. coli enzyme [11].Th e relative zymeHis45 0 •Tyr (A); enzyme His450->Phe (O) fluorescence quantum yield of the mutated enzymes with re­ spect to the wild-type enzyme as presented in Table 1i s grossly retained over the pH interval studied. that dihydr^lipoamide was used as reducing agent [11]. From Fig. 5A , Ban d C show the pH dependence of the fluores­ the degree reduction of the flavin it is estimated that these cence of E„, and EH2. The pK, values obtained from fitting species conti ibuted t to maximally 10-15 % of the final spec- are presented in Table 3. The pA", values of the fluorescence trum. The formation of these species is highest in the wild- of EOKo f the different enzymes demonstrate that at least one, type enzymi and almost negligible in enzyme His450->Ser. and in enzyme His450->Ser two, protonation steps influence No correctii>n s are made for the disproportionation. Thera­ the fluorescence of E0,. fore the caliulated py values are approximations and the Studies of the relative fluorescence of E. coli EH2 have correspondiii g absorption coefficients, especially for the wild- revealed that the extrapolated maximal fluorescence at zero type enzymi, are under-estimated. In Table 2, apparent pK, charge-transfer absorbance reaches 70% of the fluorescence values and a ^sorption coefficients of the charge-transfer com- of E„, [11] and thus the fluorescence quantum yield of fully plex of the qifferent enzymes are listed. protonated EH2 islowe r than that of EOI. Assuming similarity For the i rild--typl eenzym e the data are best fitted with three between the E. coli and A. vinelandii enzymes in this respect, pK, values (Fig- 4A). Assuming the maximal charge-transfer pKa values are calculated from varying the fluorescence quan­ 1 absorption ifflefficient of the wild-type is 2800-3200 M" tum yield of fully protonated EH2 between 70— 100% of Eox. cm"1 as re] :p irted f for E. coli and pig heart enzymes [11,14], the For enzyme His450-»Ser the maximal difference between the low, gradua absorption increase over the entire pH interval fluorescence quantum yield of EH2 and Eox is given by the indicates in A. vinelandii wild-type enzyme, as in E. coli value at lowest pH since only small changes occur at low pH. wild-type eizyme. EH2 is an equilibrium of several species The validity of the assumption is demonstrated by the large [ll]ofwhic| only one or two exhibit charge-transfer absorp- decrease in relative fluorescence of EH2 in the wild-type and tion. This c^inclusion is supported by the pH dependence of Glu455->Gln enzyme at low pH which cannot be correlated the fluorescence at the EH2 level (see below). to a proportional increase of charge-transfer absorption. Considerable changes when compared to wild-type en- Again, no corrections are made for any contributions from zyme occur in the pH profiles of both Glu455-mutated en- EOI resulting from disproportionation of EH2 and thus the zymes. The data of enzyme Glu455->Asp are well fitted as- relative fluorescence for the wild-type enzyme especially is suming one apparent pK, value (Fig. 4A). For enzyme over-estimated. Glu455-n the best fit is obtained with two apparent pK, The calculated apparent pK„ values for EH2 shown in Fig. values. The^s data indicate that the loss of the orienting effect 5 are presented in Table 3. The largest changes in relative of Glu455

49 870

Fig.5 .pH dependenceof the relative fluorescence of A. vinelandii wild-typelipoamide dehydrogenase and enzymes G/u455-*Gln andHis450-*Ser. Conditions and analysis are asdescribe d in Materials and Methods.Th ecurve sdraw n through the data arecalculate d from the apparentpK a values obtained from the best fit. (O) EOM;(•) , EH2- (A) Wild-type enzyme;(B )enzym e Glu455-»Gln; C; enzyme His450-»Ser

Table 3. ApparentpK a values of thefluorescence quantum yield of H+ H+ oxidized and two-electron reducedA. vinelandii wild-type lipoamide -JL _^L dehydrogenase andsome mutatedenzymes pKai pKa2 Experimental conditions and analysis are described under Materials I II and Methods. Data, except for enzyme His450-*Phe, are presented -FAD- -FAD- in Fig. 5 SH SH Enzyme Reduction level pKai pK,2 + SH H30 Wild-type E„ 6.5 + 0.2 Wild-type EH2 5.3 + 0.2 7.3 + 0.1 Glu455->Gln E„, 7.2 + 0.1 Glu455->Gln EH2 5.3 + 0.2 His450->Ser E„ 6.3 + 0.3 7.7 + 0.1 -FAD- -FAD- His450^Ser EH2 6.1 + 0.4 7.6 + 0.1 His450-*Phe EH2 6.0 + 0.2 7.0 + 0.1

protonated EH2 level is not precisely known, the data only ^ III IV have moderate value. Quite different in this respect are the Scheme I. Proposed modelfor the pH dependence ofthepredominant relative fluorescence data for enzymes His450->Ser and species of His450-mutaled A. vinelandii lipoamide dehydrogenases at His450-»Phe which very closely follow the charge-transfer theEH 2level. Species I is the fully protonated species, EH2 .Specie s absorption. IIan d IIIar eequilibriu m specieso f EH2 after thefirst deprotonation . Species VI is the E2~ species after deprotonation of both 'thiols' at high pH. The arrow signifies the charge-transfer interaction between thethiolat ean d theFAD .Fl. ,fluorescen t species.CT. , charge-transfer DISCUSSION species For a better understanding of the catalytic mechanism of lipoamide dehydrogenase from A. vinelandii, the active-site residue, His450, its hydrogen-bonding partner Glu455, and Pro451 were replaced with different amino acids. The low very easily over-reduced to EH4 [7]. The A. vinelandii wild- turnover rate of all mutated enzymes studied clearly shows type enzyme is less prone to over-reduction than the E. coli that all three amino acid residues are essential for effective enzyme, but is more easily over-reduced than the pig heart catalysis. The implications of the mutations on the kinetic enzyme. properties of the enzymes will be presented separately. Here From the crystal structures of A. vinelandii E„ and we have focussed on the spectral properties of the mutated oxidized and two-electron-reduced human glutathione re­ enzymes. ductase it is known that the glutamate and the histidine form With the exception of enzyme Pro455-»Ala, the absorp­ a short but strong hydrogen bond [19-21]. It has been tion, fluorescence and CD spectra of A. vinelandii wild-type suggested that this interaction results in a rise of the pK„ of and mutated E0I are highly similar showing that the mutations the histidine [19, 21]. The distance between the (protonated) have not introduced drastic structural changes. histidine and the interchange thiol is also short and allows The spectral similarities between the wild-type enzyme and ion-pair interaction. The data from the pH-dependent studies the mutated enzymes disappear upon reduction. For E. coli on theenzyme s willb e interpreted in the light of these structur­ and pig heart lipoamide dehydrogenase differences with re­ al features. It should be noted here that only the interchange spect to stabilization of EH2 have been reported [11, 14]. For thiol, Cys48, the charge-transfer thiol, Cys53, and His450 E. coli enzyme an equilibrium of at least three species was are considered to contribute to the observed pH-dependent proposed [11]. Only one of these species exhibits charge trans­ effects. fer. For pig heart enzyme the charge-transfer species prevails The results presented in this paper on the pH dependence [14,15]. The EH2 state of pig heart enzyme is very stable, i.e. of the charge-transfer absorption and the fluorescence of EH2 the enzyme cannot be over-reduced to EH4 by the substrate of the A. vinelandii wild-type enzyme show that, as in the E. dihydrolipoamide [6, 15]. In contrast, the E. coli enzyme is coli enzyme [11], at neutral pH, EH2 consists of multiple

50 871

H+ H+ S .H+ pKal pKa2 pKa3 n IV -FAD -FAD- p-FAD

-SH •SH -SH

SHHHi s •S' HHis -S" His

-FAD- ^FAD -FAD- FAD- r- SJ SH HHis SH His -S" HHis His

III ' VI ' VII

Scheme 2.Proposed model for thepH dependence of thepredominant species ofA. vinelandii Upoamide dehydrogenase at theEH 2 level. Species I isth e fully frotonated species, EH3. Specie s II and III are equilibrium species of EH2 after the first deprotonation. Species IV,V an dV I are theEH ~ lpecies after thesecon d deprotonation. Species VII isE 2~, presumably predominant at high pH.Th earro w signifies thecharge - transfer interaction between thethiolat e and the FAD.No t included areth especie s E0I and EH4 andth e EH2 species FIH". FL., fluorescent species, CT.,charge-transfe r species

electronic fojrms. The distribution of these forms is complex water, the interaction with an hydronium ion as proposed for and quantitiitio n is hampered by disproportionation. Before enzyme His450->Ser remains possible and will be enhanced discussing tlt e wild-type enzyme in more detail, the His450- in a more apolar environment. This results in less influence of mutated enzyme s will be addressed since these show less com­ the interchange thiolate on the charge-transfer thiol and plex behavicr . lowers the pKt of the latter. Moreover, upon deprotonation With all His450-mutated enzymes the data of the pH- of the charge-transfer thiol the negative charge is more com­ dependent < harge-transfer absorption were well fitted as­ pensated bya stronger interaction with the flavin as evidenced suming two apparent pK, values indicating the involvement by the unusual high charge-transfer absorption (e about 3900 of at least Iw o titratable groups. The data on the relative M ~' cm"' ) and a shift of the absorption maximum from 530 fluorescence of enzyme His450->Ser show that the large de­ run to 540 nm. crease in flu jrescence, expected from formation of a thiolate Based on the proposed Scheme 1fo r the His450-mutated near the flai in, occurs with a pK„ of 7.7, matching perfectly enzymes, Scheme 2depict s the protonation states of the wild- with thepA , obtained for thelarg e increase of charge-transfer type enzyme (adapted from [11]). Not included are the EH4 absorbance. Similar results were obtained for enzyme species and Eox resulting from disproportionation and the His450-»Phi where the pK, values for the largest increase in species F1H~ . charge-transfer absorbance and decrease in relative fluores­ At low pH, above pKti, the charge-transfer absorption is cence coincded. Scheme 1 depicts the major tautomeric low and the relative fluorescence high. Thus the major species species for tli eHis45 0 mutated enzymes. Thefac t that charge- at this protonation level is species II in Scheme 2. Based on transfer absiirptio n and high fluorescence is observed at low the results from the His450-mutated enzymes, it is concluded pH (apparel t pK, of 5.5 in all His450-mutated enzymes) dem­ that in the wild-type enzyme also deprotonation of the inter­ onstrates thi t tautomerization equilibrium between species II change thiol leads to tautomerization that results in charge- and III in Scheme 1 is almost completely shifted in favor transfer absorption. of species II. Therefore the deprotonation that leads to the At a pH above pA^,2, the tautomerization equilibrium tautomerizaiion , with an apparent pK„ of 5.5, is assigned to shifts in favor of the charge-transfer thiolate species V and the interchaiig e thiol. VI. Species IV is also present as indicated by the relative At high ]) H both thiols become deprotonated (species IV fluorescence. The third apparent pKt is assigned to the histi- in Scheme i). The presence of two negative charges in the dine in analogy to E. coli and pighear t enzyme [9,11,14] and active site stems conflicting. However, bearing in mind the glutathione reductase[14] . fact that theinterchang e thiol isreadil y accessible to water, the Scheme 2,presente d for thewild-typ e enzyme, also applies assumption is made that the negatively charged interchange to the Glu455-mutated enzymes. With these enzymes, full thiolate is stabilized by interaction with an hydronium ion. deprotonation is compressed to a JpH of 1—2 resulting in Apart from this, the charge-transfer thiolate is stabilized via co-titration of all three residues in EH2 with only a small interaction with the flavin. The positive end of a helix dipole contribution in charge-transfer absorption associated with an might conttibute to the stabilization of both thiolates apparent pK, of 6.9 for enzyme Glu455-»Gln. Replacement (Mattevi, personal communication). of Glu455 with Asp or Gin results in weakening or loss of the The decrease of pKal in enzymes His450->Tyr and hydrogen bond with the histidine which most probably will His450->Phi! supports the assumption that charge compen­ lead to an impaired orientation of the histidine relative to the sation occuis. As known from modelling studies with the interchange cysteine and a lowering of the p/C of the histidine crystal structure, the interchange thiolate is still accessible for in EH2. A distorted ion-pair interaction between the inter-

51 872 change thiolate and the protonated histidine and a lowered 10. Matthews, R. G., Ballou, D. P. & Williams, C. H. Jr (1979) J. p/C, of the histidine lead to less stabilization of the tautomeric Biol.Chem. 254, 4974-4981. species II in Scheme 2 and as a consequence species III is 11. Wilkinson, K. D.& Williams C. H.J r (1979)J. Biol.Chem. 254, favored. A lowered His450 p/f also results in formation of 852-862. a 12. Wilkinson, K. D.& William s C.H .J r (1981)J. Biol.Chem. 256, species VII at lower pH than in wild-type enzyme as the 2307-2314. protonated histidine ispar t of the majority of tautomerization 13. Sahlman, L. & Williams, C. H. Jr (1989) J. Biol. Chem. 264, species of Scheme 2. 8033-8038. Taken together, the results on the Glu455-mutated en­ 14. Sahlman, L. & Williams, C. H. Jr (1989) J. Biol. Chem. 264, zymes make it clear that the mutations lead to preference of 8039-8045. thecharge-transfe r species at the EH2 level of the enzymes and 15. Massey, V.& Veeger , C.(1961 ) Biochim. Biophys. Acta 48,33- demonstrate that Glu455 is an extremely important residue in 47. balancing the distribution of the equilibrium of electronic 16. Matthews,R .G . & Williams,C . H.J r (1976)J. Biol.Chem. 251, species at the EH2 level in the wild-type enzyme. 3956-3964. The results presented here show that mutation of His450 17. Adamson, S. R., Robinson, J. A. & Slevenson, K.. J. (1984) leads to a rather simple distribution of electronic species at Biochemistry 23, 1269-1274. 18. Mattevi, A., Schierbeek, A. J., Obmolova, G., Kalk, K. H. & the EH2 level that allows easy interpretation and has proved to be of great value for the interpretation of the complex Hoi, W. G. J. (1991) in Flavins andflavoproteins 1990 (Curti , B., Zanetti, G. & Ronchi, S., eds) pp 549-556. Walter de distribution in the wild-type enzyme and Glu455-mutated en­ Gruyter, Berlin, New York. zymes. Our results on the Glu455-mutated enzymes indicate 19. Pai, E. F. & Schulz, G. E. (1983) J. Biol. Chem. 258, 1752- that the effect of the histidine on the charge-transfer thiol is 1757. indirect and mediated through the interchange thiol. 20. Thieme, R., Pai, E. F., Schirmer, R. H. & Schulz, G. E. (1981) At present, studies are aimed at gaining more insight in J. Mol.Biol. 152, 763- 782. the relation of the distribution of the EH2 species in the 21. Karplus,P .A. , Pai,E .F .& Schulz,G . E.(1989 )Eur. J. Biochem. mutated enzymes and the impact of the distribution on the 178, 693-703. separate half reactions and overall catalysis of the (mutated) 22. Reed, L.J. , Koike, M., Lertich, M. E.& and Leach, F. R.(1958 ) enzyme(s). J.Biol. Chem.232, 143-149 . 23. Gibson, T. J. (1984) Ph.D Thesis, University of Cambridge, Wethan k Dr W. G. J. Hoi and Mr A. Mattevi for providing us England. with the coordinates of lipoamide dehydrogenase from A. vinelandii 24. Kunkel, T. A., Roberts, J. D. & Zakour. R. A. (1987) Methods prior to publication. We are indebted to Dr C. H. Williams Jr for Enzymol.154, 367-382 . helpful discussions. This work was supported by the Dutch-Foun ­ 25. Westphal, A. H. & De Kok, A. (1988) Eur.J. Biochem. 172, dation for Chemical Research (SON) with financial aid from the 299-305. Netherlands Organization for Scientific Research (NWO). 26. Sanger, F.,Nicklen , S.& Coulson,A . R. (1977)Proc. NatlAcad. Sci. USA 74, 5463-5467. REFERENCES 27. Van Berkel, W. J. H„ Van den Berg, W. A. M. A Muller, F. (1988) Eur. J. Biochem.178, 197-207. 1. Williams, C. H. Jr (1976) In Theenzymes (Boyer, P. D. ed.) vol. 28. Benen, J. A. E., Van Berkel, W. J. H., Van Dongen, W. M. A. 13,pp . 89— 173, Academic Press, New York. M., Muller, F. & De Kok, A. (1989) J. Gen.Microbiol. 135. 2. Sokatch,J . R., McCully, V.,Gebroski ,J . &Sokatch , D.J . (1981) 1787-1797. J. Bacteriol. 148, 639-646. 29. Berkel,W .J . H.van , Benen,J . A. E.& Snoek , M.C . (1991) Eur. 3. Fox, B.S .& Walsh , C.T . (1982)J. Biol.Chem. 257, 2498-2503 . 4. Shames,S .L. , Fairlamb,A .H. ,Cerami , A.& Walsh ,C .T .(1986 ) J. Biochem.797,769-779 . Biochemistry 25, 3519-3526. 30. Kosower, E. M.(1966 )i nFlavins andflavoproteins (Slater , E. C 5. Holmgren, A. (1980) Experienlia suppl. 36, 149-180. ed.) pp. 1 —14 , Elsevier, Amsterdam. 6. Massey, V., Gibson, Q. H. & Veeger, C. (1960) Biochem.J. 77, 31. Visser,A .J .W .G. , Grande, H.J. , Muller, F.& Veeger,C .(1974 ) 341-351. Eur. J. Biochem. 45,99-107 . 7. Williams,C . H. Jr (1965)J. Biol.Chem. 240, 4793-4800 . 32. Brady, A. H. & Beychok. S. (1969) J. Biol.Chem. 244, 4634- 8. Reed, J. K. (1973)J. Biol. Chem.248, 4834-4839 . 4637. 9. Matthews, R. G., Ballou, D. P.,Thorpe , C.& Williams , C. H.J r 33. Brady, A. H. & Beychok. S. (1971) J. Biol. Chem.246, 5498- (1977)7. Biol.Chem. 252, 3199-3207 . 5503.

52 Chapter 3

Lipoamide Dehydrogenase from Azotobacter vinelandii: site-directed mutagenesis of the His450-Glu455 diad. Kinetics of wild-type and mutatedenzymes .

Jacques Benen,Wille m van Berkel,Nicol e Dieteren, DavidArscott , Charles Williams Jr., Cees\ reeger and Arie de Kok.

Summary

Three amino acid residues in the active site of lipoamide dehydrogenase from Azotobacter vinelandiiwer e replaced with other residues. Histidine450,th e active site base, was replaced with serine, tyrosine and phenylalanine respectively. Proline451, from Xfray analysis found to be in c/s-conformation positioning the backbone carbonyl of His450clos e to N3o f the flavin,wa s changed into alanine. Glutamate455, from X-ray analysis expectedt o be involved in modulating the pKa ofth e base (histidine450), was replacedwit h aspartate andglutamine . The general conclusion istha t mutation of the His-Glu diad impairs intramolecular electron transfer between the disulfide/dithiol and the FApHVFAD. TJiewild-typ e enzyme functions according to a ping-pong mechanism in the physiological reaction in which the formation of NADH is rate limiting.Abov e pH 8.0 the enzymi is strongly inhibited byth e product NADH.Th e pHdependenc e of the steady state kinetics using the NAD+analo g 3-acetylpyridine adenine dinucleotide [AcPyAde+] revealsa pKa of 8.1i nth e pKmAcPyAde +plo t indicating that this pKa is relatedt oth e depro4nation of His450 [Benen et al. (1991) Eur.J. Biochem. 202,863-872 ] andt o the inhibition by NADH. Thq mutations considerably affect turnover. Enzymes Pro451->Ala, His450->Phe and His450->Tyr appear to be almost inactive in both directions. Enzyme His450->Ser is minimally active,V atth e pHoptimu m being 0.5% of wild-type activity inth e physiopgical reaction. Rapid reaction kinetics show that for the His450 mutated enzymesth e reductive half reaction using reduced 6,8-thioctic acid amide [lip(SH)2] is rate limiting and extremely slow when compared to the wild-type enzyme. For enzyme Pro45l)->Ala it is concludedtha t the loss of activity isdu et o over-reduction by lip(SH)2 and NADH.Th e Glu455 mutatedenzyme s arecatalyticall y competent but show strong inhibition byth e product NADH (Enzyme Glu455->Asp more than Glu455->Gln). The

53 inhibition can largely be overcome by using AcPyAde+ instead of NAD+i nth e physiological reaction. The rapid reaction kinetics obtained for enzymes Glu455->Asp and Glu455->Gln deviate fromth e wild-type enzyme. It isconclude dtha tthi sdifferenc e isdu et o cooperativity between the active sites inthi s dimeric enzyme. Rapid reaction kinetics of enzymes His450->Ser andGlu455->Gl n showth e existence of two intermediates atth e 2-electron reduced level:a species withth e NAD+bound ,th e flavin reduced andth e disulfide intact (oxidized) and a species with NAD+bound ,th e disulfide reducedan d the flavin oxidized. No spectral evidence is obtained for the participation of a proposed flavin C4a adduct intermediate [Thorpe,C . andWilliams ,C.H .Jr . (1981)Biochemistry 20, 1507-1513] in the reaction mechanism of enzymes His450->Ser and Glu455->Gln.

Introduction

Lipoamide dehydrogenase isth e flavoprotein component of the pyruvate-, oxoglutarate- and branched chain oxo acid dehydrogenase complexes [1,2]. Inth e physiological direction it catalyses the oxidation of a reduced lipoyl group,tha t is covalently attached toth e core protein of these complexes, by NAD+.Thi s lipoyl group can be replaced with free lipoamide. Lipoamide dehydrogenase belongs to the family of dimeric flavoenzymes that contain a redox active disulfide bridge participating in catalysis [1].Th e enzyme iscompose d oftw o identical subunits withth e two active sites built bycontact s betweenth e subunits. Other members of this family are glutathione reductase [1], mercuric ion reductase [3], trypanothione reductase [4] and thioredoxin reductase[5] . Extensive studies on lipoamide dehydrogenase have been performed on the enzymes as isolatedfro m pig heart, rat liver and Escherichiaco//[6-14] . Allthre e lipoamide dehydrogenases were shown to act according to a ping-pong mechanism in the physiological reaction [6,8,14] . Duringcatalysis ,th e enzyme shuttles between the oxidized [Eox] andth e 2-electron reduced [EH2] state [15]. The 4-electron reducedstat e [EH4]wa sshow nt o beinactiv e andt o result from over-reduction by NADH [12,15]. The over-reduction is manifested as product inhibition. Lipoamide dehydrogenase from E.coli has been shown to be very susceptible to product/,dead-end' inhibition [7, 12]. The inhibition is so severe that determination of steady state kinetics was only possible ina rapid reaction spectrophotometer [14].Th e nature ofth e dimeric structure indicates that communication between the active sites seems possible. For each enzyme a different degree of cooperativity was reported.Fo r pig heart enzyme negative cooperativity was reported (from NAD+bindin g studies) in mono-alkylated enzyme [16,

54 17] ancli n native enzyme [18]. For E. coli lipoamidedehydrogenas e [14] positive cooperativity at the EH2 level was reported while for rat liver lipoamide dehydrogenase a slighl negative cooperativity atth e first substrate levelwa s reported [8]. Forth e related enzymo mercuric ion reductase very strong cooperativity was reported recently that was explained in terms of alternating sites activity [19]. Detailed studies on pig heart enzyme comprising both steady state and rapid reaction kinetic^ ledt o the proposal of a reaction mechanism involving a base, histidine, inth e active $itetha t becomes protonated during catalysis [9,20 ,21] . Recently, conclusive evidence forth e role of the histidine was obtained by site directed mutagenesis studies on glutathione reductase [22,23] , E.coli [24 ] andA. vinelandiilipoamid e dehydrogenase [25]. Scheme 1depict s the proposed reaction intermediates during turnover in either direction andth e formation of the EH4species . Speciek (1) represents Eox. Species (2) has been proposed by Matthews andco ­ workers [9] and is known asth e mixeddisulfid e species.Afte r release of the oxidized substrcjte, species (3) isformed ,th e EH2specie s exhibiting the typical 530 nmcharge - transfer absorbance giving riset o a redcolor . It has been shown for E.coli [11 ] and A. vinalandiilipoamid e dehydrogenase [26] that this species consists of multiple electronic forms. The formation of the Michaelis complex of EH2an d NAD+,specie s (4), has beun proposed in analogy with pig heart EH2 reacting with the NAD+ analogs acetylp/ridine adenine dinucleotide [AcPyAde+] or nicotinamide hypoxanthine dinucleotide [10]. Itwa s shown that these intermediates had an altered charge-transfer absorbance with a maximum at 580 nm. Species (5),th e flavin C4a adduct intermediate characterized by an absorbance maximum at 384 nm,wa s proposed based on NAD+ binding]studie s with monoalkylated pig heart enzyme [17]. This species was also identified in wild-type and a mutated mercuric ion reductase [27, 28].Afte r completion of the reofddation of the disulfide the reducing equivalents reside onth e FAD as depicted by species (6). This species was first described by Massey andVeege r [15] studying reduction of pig heart enzyme with NADH inth e presence of arsenite. A green species was observed with an absorbance maximum at 680-700 nm.A similar green species was reported for pig heart enzyme reacting with reduced acetylpyridine adenine dinucleotide or nicotinamide hypoxanthine dinucleotide [10]. Finally FADH-i s reoxidized by NAD+ resulting in species (7). Spectral evidence for this species, showing low long wavelength absorbance extending beyond 700 nm and a maximum at 580 nm, has only been obtained by studies of pig heart enzyme reacting with reduced acetylpyridine adenine dinucleotide or nicotinamide hypoxanthine dinucleotide [10] and very recently with Lys53->Arg mutated E.coli enzyme [29] For E. coli lipoamide dehydrogenase over-reduction by a slight excess of reduced 6,8-thiqctic acid amide [lip(SH)2] was reported [7].Thi s was also shown for A.vinelandii

55 7 -NAD* NADH HHIs- H+ |-FADH" FAD J S I HIs- I His- -S

11 10 9 ' -NAD* , - -FADH"' 1, NAD -FADH UpSg - FADH"

1 UpSg -IH"""- NAD - -1HHHI^ *HHis- rsv. 1

Scheme 1. Proposed reaction intermediates for lipoamide dehydrogenase. The arrows indicate charge-transfer interaction and are drawnfro m donor to acceptor. Explanation intext . lipoamide dehydrogenase though at much higher concentrations of lip(SH)2 [26]. Species 8-10 are possible intermediates. Before reduction by lip(SH)2 can take place the disulfide must be in its oxidized state as shown by species (8).Analogou s to reduction of Eox the disulfide canthe n be further reduced viath e mixed disulfide intermediate, species (9). Species (10) isth e fully reduced EH4specie s with virtually no absorbance inth e visible region.Th e reduced flavin carries a negative charge on N(1) as evidenced by NMRstudie s [30].Specie s (11) represents the NAD+boun d EH4 speciestha t can beobtaine d by reduction of Eox with an excess of NADH via species (3) or by reacting species (10) with NAD+.Thi s species ischaracterize d by an absorbance maximum at 750 nm leading to a blue/grey color. In a previous paperth e rationale for the selection of amino acidst o be mutated was outlined andth e spectral properties of the wild-type and mutated enzymes were reported [26].Thi s paper deals with the kinetic properties and identification of severalo f the intermediate species as shown in Scheme 1 of A. vinelandii lipoamide dehydrogenase and mutated enzymes.

56 Materials and Methods

General. Construction of the mutated enzymes, isolation andtreatmen t of the enzymes were performed asdescribe d [26].NAD +(grad e I) NADH (grade I), NADP+, xanthirje, xanthine oxidase (from cow milk),glucose-6-phosphat e and glucose-6- phosptyatedehydrogenas e (from yeast, analytical grade) were from Boehringer. NAD+ analogjs, lipS2, dichlorophenolindophenol [Cl2lnd] and 5,5'-dithiobis (2-nitrobenzoate) [NbS2]land biological buffers were obtained from Sigma Inc.. All other chemicals used were of the highest purity available. NAD+, AcPyAde+, NADH, lip(SH)2 and enzyme concentrations were determined as described previously [26].Th ethio-NAD + concentration was determined as described for AcPyAde+.

Steady state kinetics. All activity measurements were performed on a Zeiss M4 QUIspectrophotomete r at 25°C.Th e forward reaction lip(SH)2/NAD+wa s determined routinely in5 0 mMsodiu m pyrophospate buffer, 0.5 mMEDT A pH8. 0 [PPi/EDTA-buffer]. Ina standar d assay 1.0 mMNAD +an d 1.0 mMlip(SH) 2wer e used andth e formation of NADHwa s monitored at 340 nm. Steady state kinetics ofth e forward reaction of the wild-ty|)e enzyme were studied as follows. To atemperatur e equilibrated cuvette containing 950 u.lPPi-buffer , 20 nl of each of the substrates was added followed byth e addition of 10|o, l of enzyme atth e proper dilution. Substrate concentration combinations were cnosen such ast o avoiderror s duet o inactivation of enzyme inth etim e course of the experiment. Therefore the experiment was started withth e lowest concentration of substrate A, where substrate Bwa svaried ,followe d byth e highest concentration of substrate A with Bvaried . Nextth e second lowest concentration of A was used followed by the(secon d highest concentration of A and so on.Triplicat e series were measured. pH(dependen t studies were performed in buffers containing 0.5 mM EDTA and adjustedt o 150m Mi nioni c strength with KCI. Buffers used:p H5.5-6.0-6.5,100 mM Mes;pt t 7.0-7.5-7.8,100 mMHepes ;p H8.0-8.6,10 0 mMHepps ;p H9.0-9. 3 100m M Ches.pat a were analyzed according to a ping-pong mechanism. Tho reverse reaction, NADH/lipS2,wa s determined as described [31].The NADH- dependent reduction of Cynd (diaphorase activity) was measured in PPi/EDTA-buffer (sceinii.,60 0 nm =2290 0 M"1 cm"1,p H8.0 ) by followingth e decrease of Cl2lnda t 600 nm. Ttie transhydrogenase reaction NADH/thio-NAD+was performed according to ref. [32].

Rabid reaction kinetics. Rapid reaction kinetics were carried out using a temperature-controlled single wavelength stopped-flow spectrophotometer, type SF-51, from High Tech Scientific inc. with 1.3m sdea dtime .Th e instrument was interfacedt o

57 an IBMcompute r for dataacquisitio n and analysis. Datawer e analyzed with a program from HighTec h Scientific inc.. Rapid scan experiments were performed with a stopped flow instrument having a2 c m light pathcel l and adea dtim e of 3 ms,an d using a Tracor Northern diode array spectrophotometer asth e detector (scantim e 5.42 ms).Th e detector was interfaced to aTraco r Northern computer for data acquisition andanalysis . All experiments were performed anaerobically at 21.0 °C in 100 mMGoo d buffers containing 0.5 mM EDTA adjustedt o 150 mMi n ionic strength with KCI. Enzyme concentrations after mixing were 26.7 nM. Generation of 2-electron reduced enzyme [EH2] was accomplished by reduction with a small excess of sodiumborohydride. Rate constants at infinite substrate concentration and Kdvalue s were determined from non-linear fitting of apparent rate constants obtained at,a t least,fiv e different substrate concentrations. Apparent rate constants represent the average of minimally four shots. Inorde r to achieve pseudo-first order conditions the lowest substrate concentrations usedwer e five times as high asth e enzyme concentration. Since only the L-enantiomer of lip(SH)2 reacts rapidly [6,9] ,substrat e concentrations were adjusted to those of the L-enantiomer. Lip(SH)2 and lipS2 were dissolved in buffer/ethanol to yield afina l ethanol content of 5% inth e mixing chamber. Simulation of rapid reaction kinetics was performed using the program 'KINSIM' [33] run on a VAX/VMS minicomputer.

Redox potential determinations. Visible absorption spectra were recorded on a temperature controlled Aminco DW2000 spectrophotometer at 25°C. Reductions were carried out inanaerobi c cuvettes in 50 mMpotassiu m phosphate buffer pH7.0 , 0.5m M EDTA. Redox potentials were determined inthre e different ways. Method one was essentially as described for pig heart enzyme [20]. Method two,th e xanthine/xanthine oxidase method,wa s recently described by Massey [34] and was applied without modification. The co-titrants used,3 0 |xMi nconcentration ,wer e safranine T, phenosafranine and benzylviologen. Methodthre e was a modification of methodtwo . Instead of using xanthine/xanthine oxidase as an electron generating system, NADPH was generated from NADP+ (75 \iM) using glucose-6-phosphate dehydrogenase (2-4 x 10-4 U) and glucose-6-phospate (400 p.M). NADPH proved useful as a reductant for lipoamide dehydrogenase since quantitative reduction could be achieved yielding spectra identical to reduction by lip(SH)2. Furthermore NADP+di d not give riset o the typical long wavelength absorbance characteristic for FADH7NAD+ charge-transfer interaction, indicating that no efficient binding occurs. For method one,th e titrant was added using a gastight Hamilton syringe equipped with a dispenser. Spectra were recorded untilchange s were complete. For methodtw o andthre e the reaction was started by the addition of enzyme (xanthine oxidase or glucose-6-phosphate

58 dehydrogenase) after anaerobiosis was established. In ordert o achieve equilibrium the amount of enzyme waschose n such ast o complete reduction in atim e span of 6-9 hrs. The anjiount of enzyme was halvedt o checkwhethe r equilibrium was established[34] .

Results

Steady state kinetics

WilfcJ-type lipoamide dehydrogenase. Under non-saturating conditions, the optimujn pH of the reaction lip(SH)2/NAD+i s 8.0,i n PPi/EDTA-buffer asfoun d for all other lipoamide dehydrogenases [6,7 , 8] Also the reaction Hp(SH)2/AcPyAde+ shows an optimum at pH8.0 . InFig . 1A and 1B th edoubl e reciprocal plots ofth e forward reactioh lip(SH)2/NAD+ are presented.Whe n the concentrations of lip(SH)2 and NAD+ are vasedi nfixe d ratios a straight line is obtained in a double reciprocal plot (not shown). These results clearly demonstrate that A. vinelandiilipoamid e dehydrogenase like all(othe r lipoamide dehydrogenases studied sofa r functions according to a ping- pong njiechanism.

10 15 2 4 6 1 1 1/[NAD+] (mM- ) 1/Illp(SH)2](mM- )

Figj 1. Steady state kinetics of thephysiological reaction ofAzotobacte r vinelandii wild-type Npoamidedehydrogenase at theoptimum pH 8.0. Thetemperatur e was 25°C,fo r other details sea Materials and Methods.A , LB-plot of the reaction lip(SH)2/NAD+wit h [NAD+]varie d and [ypjSH)2]fixed .Close dtriangle s represent the extrapolated velocities at infinite [lip(SH)2]. NAD+ coijcentrations (nM): 48, 72,92,147,194,511, 752 and 1044. Lip(SH)2 concentrations (nM): 104,200,295,397 and518 . B, asA wit h [lip(SH)2]varie d and [NAD+]fixed .Close d triangles represent the extrapolated velocities at infinite [NAD+].

Fram secondary double reciprocal plotsth e V and Kmvalue s for NAD+an d lip(SH)2 aredetermined :V =49 0 U/mg, kcat= 42 0 s-1, Kmlip(SH) 2= 39 0 uMan dK mNAD += 325nty . A Hill-plot constructed fromth e datafo r the wild-type enzyme reveals a coefficient of h= 1wit h NAD+a sa substrat e excluding cooperativity atth e EH2level . From Experiments in which the lip(SH)2 and AcPyAde+concentration s are varied in fixed ratios it can beconclude d that the mechanism ofth e reaction remains ping-pong. As isojea r from Table 1, the rate of reduction of AcPyAde+ byth e wild-type lipoamide

59 dehydrogenase is considerably lower thantha t of NAD+. From secondary double reciprocal plots the Kmvalue s for lip(SH)2 and AcPyAde* are determined. Kmlip(SH) 2 is lowered 4-fold whencompare d with NAD-*- as a substrate (see Table 1)whil e theK m

AcPyAde+i s2- 3fol d higher thanth e KmNAD+ . To explore the effect ofth e pH onth e activity, steady-state kinetics of wild-type enzyme was studied at different pHvalue s (see Fig.2 A andC ) using NAD+an d AcPyAde+ as electron acceptors. Unexpectedly, strong product inhibition is observed for wild-type enzyme using NAD+a t high pHvalues . This will bediscusse d below. As a

Enzyme lip(SH)2 / NAD* lip(SH)2 APAD* NADH Cl?lnd

kcat Km Hp(SH)2 Km NAD* kcat Km liP(SH)2 Km APAD- kcat Km NADH s-1 liM liM s-1 HM |lM s-' liM

Wild-type 420 390 325 100 140 1200 170 180 His4S0->Ser 1.6 1950 300 - - - 135 140 His4S0->Phe 0.0 - - - - - 170 150 His450->Tyr 0.0 - - - - - 170 200 Glu455->Asp nd - - 2.4 30 640 160 50 Glu455->Gln nd - - 1.9 240 1750 200 120 Pro451->Ala ? - - - - - ? -

Table 1.Steady state kineticparameters ofAzotoba&er vir\e\ar\d\i wild-typeand mutated enzymes.Th e parameters aredetermine d at pH8.0 ,th e pHoptimu mfo r each enzyme, except for enzyme Glu455->Aspwher eth e optimumwa s pH 7.0. ndsignifie stha t kinetics are not studied due to strong product inhibition.A questio n mark (?) for enzyme Pro451->Ala signifies that the parameters can not be determined duet o verytransien t activity (seetex t also).Th e concentration lip(SH)2 isfo rth e D,L-mixture.

result the Kman d V values determined at pHvalue s above pH 8.0 are not reliable and aretherefor e not included in Fig.2 A .Thi s inhibition is not observed with AcPyAde+a s acceptor.

Fromth e Log(V) plot in Fig.2 A ,a pK a value of 6.4 isobtaine dfo rth e rate limiting intermediate of wild-type enzyme. The invariance of pKm [lip(SH)2]an dth e change in the pKm [NAD+] around pH6. 4 suggeststha t protonation of the EH2.NAD+comple x is rate limiting.Thi s is supported by rapid reaction kinetics which show that at pH8. 0 and

6.8 the reoxidation by NAD+i s rate limiting (see below).Almos t identical pKa values as found forth e limiting protonation of the EH2.NAD+comple x of wild-type enzyme were reportedfo r pig heart [10]an d E.co//[14] enzyme,pK a =6. 2 and6. 7 respectively. This indicates that the enzymes obey the same mechanism of reoxidation. Fig. 2 Cshow stha t Vfo r AcPyAde+ reduction bywild-typ e enzyme does not reach a maximum inth e experimental pH interval.Therefor e no pKa can be assignedt o a rate limiting intermediate. Fromth e pKm [AcPyAde+] plot a pKao f 8.1 isdetermine d represen­ ting adeprotonatio n of EH2[35] .Thi s pKa value may reflect deprotonation of the histidine.

60 Fig.2 . pH dependence of thesteady state kinetics in thephysiological reaction of Azotobacter vinejandii wild-type lipoamide dehydrogenase and mutated enzymes His450->Ser and Gki4B5->Asp.Condition s are described in Materials and Methods.A ,wild-typ e enzyme using NADr as electron acceptor. B, enzyme His450->Ser using NAD+a selectro n acceptor. C, wild- typejenzym e using AcPyAde+ as electron acceptor. D, enzyme Glu455->Asp using AcPyAde* as electron acceptor.

61 Mutated enzymes. Table 1give s an overview of the specific activities in the various reactions of the wild-type and mutated enzymes. Considerable differences are observed between the wild-type andth e mutated enzymes inth e rate of either NAD+o r AcPyAde+dependen t oxidation of lip(SH)2.Th e diaphorase activity is however almost the same in wild-type and all mutated lipoamide dehydrogenases which indicates that the binding of NADHa t the re-side ofth e FADi s not appreciably affected byth e mutations onth e sAside.

His450 mutated enzymes. Changing the 'proton accepting' histidine450 [9] into tyrosine, phenylalanine or serine clearly demonstrates the important role of this residue in catalysis. With enzymes His450->Phe and His450->Tyr in either directions virtually no activity is observed.Wit h enzyme His450->Ser almost no activity was observed inth e reverse reaction, reduction of lipS2, monitored by the oxidation of NADH inth e presence of NAD+.Abou t 0.5%o fth e wild-type activity isfoun d inth e forward reaction, lip(SH)2/NAD+,a t the optimum pH8.0 . A kinetic analysis asdescribe d for wild-type enzyme demonstrates that enzyme His450->Ser also functions according to a ping- pong mechanism.Th e KmNAD +i sslightl y smaller inthi s mutated enzyme andth e

Km lip(SH)2 is 5-fold higher than found for the wild-type enzyme. Assuming His450 to abstract a proton fromth e substrate lip(SH)2 as proposed[9] , one might expect the activity of enzyme His450->Ser would increase at elevated pH values, where lip(SH)2 (pKai = 9.35, free in solution [9]) becomesdeprotonated . However V remains constant between pH 8.0 and pH9.0 . Inth e pH dependent studies no product inhibition by NADH isfoun d inth e experimental pH interval. From the Log(V) plot in Fig.2 B,a pKa value of 6.3 isobtaine dfo r the deprotonation ofth e rate limiting intermediate in enzyme His450->Ser. The 0.5% of activity remaining inth e forward reaction iscompatibl e with results recently reported for His-to-Alaan d His-to-GIn mutated enzymes of glutathione reductase [22,23 ] and His-to-GIn in E.coli lipoamide dehydrogenase [24]. This suggeststha t the function of the imidazole as a base,ca n becarrie d out by bound solvent, albeit ineffectively. The difference in effect of either tyrosine or phenylalanine at position 450 can not be assessed via kinetics. Spectral studies showedtha t these enzymes stillca n be reduced by lip(SH)2thoug h very slowly. It istherefor e concluded that the rather bulky aromatic residues impose structural constraints onth e enzyme that make it very difficult for lip(SH)2t o effectively interact with the disulfide to exchange reducing equivalents.

62 GluA55 mutated enzymes. With enzyme Glu455->Asp very strong product (NADH) inhibition is observed in the reaction lip(SH)2/NAD+. The results indicate that this enzyme is inhibited by NADH in a 'dead-end' manner. Addition of excess NAD+ after inhibition is complete does not restore activity. Revived NADH production isonl y observed after the addition of an extra aliquot of enzyme. This extra enzyme portion becomes inhibited approximately as fast asth e initial enzyme portion. No detailed studies!were carried out to investigate this phenomenon. However the results indicate that theinhibitio n is more dependent ontim e than onth e NADH concentration. Invie w of an increase of the dissociation constant forth e monomer-dimer equilibrium upon reduction to the EH4leve l [31] it istemptin gt o speculate that the irreversibility of the inhibition is caused by monomerization. Thejreverse reaction as well asth e transhydrogenase reaction NADH/thio-NAD+ are not detectable. When an electron acceptor with higher midpoint potential than NAD+o r thio-NAp1-i s used,in casuAcPyAde +i nth e forward reaction,an d Cl2lnd inth e diaphorase reaction,quantitativ e reduction of the acceptors is observed in the pH

interval studied (pH 5.5-9.0) (E'm NAD+/NADH = -320 mV [36],E' m thio-NAD+/thio-

NADH * -283 mV [36],E' mAcPyAde+/AcPyAde H = -258 mV[36 ] and E'm Cl2lnd/

CI2lndHJ2= +217 mV [37], pH7. 0 ). WithAcPyAde +"dead-end " inhibition is observed above bH7.5 . Enzyme Glu455->Gln differs in catalytic behavior from enzyme Glu455->Asp. Here a somewhat less strong inhibitory effect of NADH isfound .Wit h AcPyAde+a saccepto r no inhibitiajn is observed. Undjernon-saturatin g fixed substrate concentrations the optimum for the AcPyAde+ reduction is at pH 8.0 for enzyme Glu455->Gln and is loweredt o pH7.0-7. 5fo r enzyme Glu4554>Asp. Both Glu455 mutated enzymes function according to a ping-pong mechanism. Forbot hGlu45 5 mutated enzymesth e steady-state kinetics ofth e reaction Hp(SHM/AcPyAde+wer e studied as afunctio n of pH.Enzym e Glu455->Asp shows,a s mentioned above, 'dead-end' inhibition with AcPyAde+. This hampers the determination of the kjnetic constants above pH 7.5. Except for achang e inV ,th e kinetic parameterso f enzymaGlu455->As p do not change significantly in the experimental pH range (pH £ 75) , Fig. 2 D. The pH dependence of steady state kinetics of enzyme Glu455-|>Gln isquit e different from enzyme Glu455->Asp (results not shown).V change^ only slightly overth e experimental pH range (pH 5.5-9.0) showing an optimum

at pH8J0 .Th e Kmfo r lip(SH)2 increases at lower pH but KmAcPyAde + remains almost constant as in enzyme Glu455->Asp. No pKa values are determinable for this enzyme.

63 Rapid reaction kinetics

Inorde rt o get a more detailed insight intoth e separate half reactions,th e reductive reactions with lip(SH)2 and NADH andth e oxidative reactions with NAD+an d lipSa were studied in the stopped-flow spectrophotometer.

Wild-type lipoamide dehydrogenase. At pH 7.0 and 8.0 the reduction of Eox by lip(SH)2 and reoxidation of EH2b y NAD+an d lipS2 as monitored at 450 nman d 530 nmca n essentially be described by a single-exponential function. InTabl e 2th e extrapolated rateconstant s at infinite substrate concentration and corresponding Kd values are presented.

Enzyme Reduction Substrate pH ka Kti(a) kb K

Wild-type Eox lip(SH)2 8.0 2000 0.9 450, 530

Eox lip(SH)2 7.0 1000 0.7 530 Eox NADH 8.0 >3000 450, 530, 700

EH2 lipS2 8.0 400 0.5 450, 530

EH2 NAD* 8.0 1000 0.37 450, 530

EH2 NAD* 7.0 760 0.47 450, 530

His450->Ser Eox lip(SH)2 9.3 1.9 5.0 450, 530

Eox lip(SH)2 8.6 1.5 4.3 530

Eox lip(SH)2 7.8 1.3 4.7 530

Eox lip(SH)2 7.3 1.0 3.7 530

Eox lip(SH)2 6.6 0.8 3.5 530 Eox NADH 8.0 >3000 450, 530, 700

EH2 lipS2 8.0 <0.005 530

EH2 NAD* 8.0 4.0 0.33 530

EH2 NAD* 6.8 4.4 0.27 530

Glu455->Asp Eox lip(SH)2 8.0 ? ? 1000 4.7 450, 530

Eox lip(SH)2 7.0 ? ? 600 4.0 530

Eox lip(SH)2 5.9 230 5.4 530 Eox NADH 8.0 >3000

EH2 lipS2 8.0 ? ? 12 0.55 450, 530

Glu455->Gln Eox lip(SH)2 8.6 250 2.6 60 2.5 530

Eox lip(SH)2 7.9 ? ? 60 4.2 530

Eox lip(SH)2 7.3 ? 1 35 3.6 530

Eox lip(SH)2 5.9 44 4.8 530 Eox NADH 8.0 >3000 450, 530, 700

EH2 lipS2 8.0 <0.05 530

EH2 NAD* 8.0 58 0.17 14 0.30 530

Table 2. Rate constants and dissociation constants ofAzotobacte r vinelandii wild-type and mutated enzymes. For NADH reductionth e rate constant of reduction of the FAD is shown.A question mark (?) signifiestha tth e constants could notb edetermine d duet o the small absorbance changes associated at low substrate concentration (seetext) .K dvalue s for lip(SH)2 are expressedfo rth e L-enantiomer. Rateconstant sk a andk b are extrapolatedt o infinite [substrate].Th e faster rate constant isk awhil eth e slower rateconstan t iskb - Thewavelengt h at whichdat awer eobtaine di sindicate di nth e lastcolumn .

64 Ping-pong enzymes obey the following equations:

E + AF^ EA *-* E* +P k2 k4 0)

k5 k7 E'+ B*= ^ E'Bs-^E +Q (2) *6 k8 for whijchth e steady state relation

. k3 x k7 (3) •^"ka + k7

wasderive d[38] .Her e l<3 andk 7ar eth e first order rateconstant s ofth e reaction ofth e first arid second substrate respectively (ka ofth e appropriate substrate inTabl e 2).A kcat 1 wascalculate d as 670 S" a t pH 8.0 forth e reaction lip(SH)2/NAD+wit h data from Table

2 andequatio n (3).Thi s value agrees reasonably well with kcat calculated from steady _1 state wnetics:42 0 s .Compariso n of k3an dk 7demonstrate s that, at pH8.0 ,th e reductive reaction by lip(SH)2 is not rate limiting asfoun d forth e pig heart enzyme [6, 10]. Tho reductive reaction with NADH isextremel y fast andi s almost complete inth e deadtjm e of the stopped-flow instrument. Rapidsca n spectra (deadtim e spectrum 5.42 msaft^ r mixing) and acompilatio n of relative absorbance changes at 453,53 0 and 700

500 600

Wavelength(nm )

Fid.3 .Spectral changes of Azotobacte rvinelandi i wild-type lipoamidedehydrogenase upon reductionby NADH in the rapidreaction instruments. A.Reductio no fwild-typ eenzym eb y5 mola requivalent so fNAD Ha tp H7.8 ,recorde dwit h thejrapi dsca ninstrument .Eox , ; deadtim e(5.4 2m safte rmixing) , ;10.8 4m s aftlr mixing, ;54 2m safte rmixing , B. Compilationo frelativ eabsorbanc e changesdurin greductio no fwild-typ eenzym eb y1 0mola requivalent so f NADHi nth esingl e wavelength stopped-flow instrument.Absorbanc echange sa t45 3nm ,0 ; absorbanc e changesa t53 0nm ,x ;absorbanc echange sa t70 0nm , •.

65 nmcollecte d with the single wavelength instrument are shown in Fig.3 A and B.Th e absorbance at 700 nm is a monitor forth e charge-transfer complex between the reduced flavin [FADH-] and NAD+(specie s 6 in Scheme 1, see ref. 26 forth e spectrum of this species). Fig. 3A shows that approximately half of thetota l absorbance change at 453 nm takes place in 10 ms, but part Bshow stha t only 20% ofth etota l absorbance change takes place in 10ms .Th e fact thatth e 700 nmabsorbanc e isalread y upa t 0.5 ms indicates that aver y fast phase, involving a significant decrease in absorbance at 453 nm,wa s largely missed.Thes e initial changes are logically attributed to the formation of the FADH7NAD+charge-transfe r complex. The 530 nm absorbance increases overth e first 5 msec and indicatestha t the electrons (formally two electrons and a proton;a hydride equivalent) are passed over from the flavin to the disulfide (species 6t o 4 or 3 in Scheme 1)a n effect that is more pronounced inth e His450->Ser enzyme (see below).Change s occurring after 5 ms are most likely duet o further reduction byth e excess of NADH.

His450 mutated enzymes. The reductive reaction of Eox by lip(SH)2 is slow (His450->Ser) to extremely slow (His450->Phe and His450->Tyr) upon mutation of the active site base. No rate constants were determined for enzymes His450->Phe and-Y . Even atth e highest possible substrate concentration full reduction to EH2laste d25 0 seconds yielding an apparent first order rate constant of ±25x 10"3 s_1. For enzyme His450->Ser the reduction by lip(SH)2 is monophasic preceded by a lag-phase ( <30 0 ms). The reoxidation of EH2 by NAD+ is also monophasic, however the lag- phase is not observed inthes e traces.Th e rate constants and Kdvalue s as determined for enzyme His450->Ser are presented inTabl e 2.Th e rate constants and Kdvalue s for the substrates L-lip(SH)2 and L-lipS2shoul d be regarded as approximations since the determined Kdvalue s are approximately 3-4 times higher than the maximal experimental concentration of L-lip(SH)2 or L-lipS2 due to solubility limitations. Forth e forward overall reaction of enzyme His450->Ser from equation (3) a kcato f 1.0s- 1 iscalculate dtha t agrees reasonably well with kcat= 1- 6s- 1 as obtained from steady state kinetics. Contrary to the wild-type enzyme however, for enzyme His450->Ser the reductive reaction is rate limiting at both pH 6.8 and 8.0. The rate constant for the reductive reaction by lip(SH)2 increases only slightly with pH andth e traces remain mono-exponential upt o pH9.3 . The Kdfo r lip(SH)2 increases with pHsuggestin g that the substrate bindst o the mutated enzyme in its protonated state. The reoxidation reaction of His450->Ser EH2b y lipS2i s barely detectable at pH 8.0 indicating that the reactivity of the substrate isver y poor orth e affinity of the enzyme forthi s substrate isver y low.

66 Trw reoxidation of EH2b y NAD+proceed s faster inthi s enzyme than the reductiono f Eoxb y lip(SH)2 (Table 2) although the reaction is also extremely slow when compared to wild type enzyme. Previously we reported a pKa of 7.5 atth e EH2 levelfo rthi s 2 enzyrmfo rth e deprotonation EH7EH -[26] .Thi s pKa is not reflected inth e pH dependence of the reoxidation rate constants. A possible mechanism of electron transfer fromth e nascent thiols to the flavin involves the transient formation of aC4 a adduct between the charge-transfer thiolate andth o flavin [17, 27, 28],specie s 5 in Scheme 1.Thi s flavin C4a adduct species exhibits characteristic absorbance at 380 nm. Therefore the reoxidation of His450->Ser EH2wit h NAD+wa s monitored at this wavelength. No absorbance changes compatible withth e formation of a flavin C4aadduc t species were howeverdetected . Tha kinetics of the reduction of enzyme His450->Ser by NADH are compatible with a very fast reduction of the flavin (species 6 in Scheme 1)wit h subsequent slow transfer of the hydride equivalent to the disulfide (species 4 and3 in Scheme 1). Fig.4 A andB show rjapid scan spectra and a compilation of relative absorbance changes at 453,53 0 and 70|0 nm obtained with the single wavelength instrument. The profound stabilization of the l:ADH7NAD+charge-transfe r complex by afacto r of 1000 isdu et o impaired transfer of electrons to the disulfide. This intramolecular transfer of an hydride equivalent involves partial regain of absorbance at 453 nm, loss of FADH7NAD+ charge-transfer at 700 nman dgai n of the thiolate to FADcharge-transfe r absorbance at

A 0.4 V > ! 3 0.2

b 0 1 00 0 [V Relativ e absorbanc chang 3.001 0.01 0.1 1 10 100 Time(s ) 0.0 500 600 700

Wavelength(nm )

FigJ4. Spectral changes ofAzotobacte r vinelandii His450->Ser lipoamide dehydrogenase upon reduction by NADH in therapid reaction instruments. A. Reductiono f enzyme His450->Ser by 10 molar equivalents of NADH at pH7.8 , recorded withjthe rapidsca n instrument. Eox, ;dea dtim e (5.42m safte r mixing), •-•-•-; 10.84 msafte r mixing, ;54 2 msec,afte r mixing, B. Compilation of relative absorbance changes during reduction of enzyme His450->Ser by 10mola r equivalents of NADH inth e single wavelength stopped-flow instrument. Absorbance changes at 453 nm,0 ; absorbance changes at530 nm,x ;absorbanc e changes at 700 nm, •.

67 530 nm. Following reduction of the disulfide, reaction with a second mole of NADHi s very slow (species 3t o 11). Also inth e reductive reaction with NADH no38 0 nm absorbance is detected that would indicate the transient stabilization of a flavin C4a adduct species.

Glu455 mutated enzymes. For the Glu455 mutated enzymes the kinetic traces obtained upon reduction of Eox with lip(SH)2 or reoxidation of EH2wit h either lipS2o r NAD+a t pH> 6.0 are not mono-exponential as found forth e wild-type and His450->Ser enzymes (Table 2).Th e introduction of a second exponential demonstrates that the reaction proceeds in at least two phases. Inal lthre e reactions studiedth e biphasic nature becomes more apparent at high substrate concentrations asth e amplitude of the fast phase increases. Except for enzyme Glu455->Gln inth e reductive reaction at pH8. 6 and inth e reoxidation reaction with NAD+a t pH8. 0 no rateconstan t forth e fast phase can confidently be determined due to the small absorbance changes causing variability in apparent rate constants determined at low substrate concentration. For enzyme Glu455->Gln inth e reductive half reaction at pH8.6 ,th e contribution of the amplitude of the fast phase increases from 5% a t 250 nM lip(SH)2t o 30 %a t 1.7m Mlip(SH) 2 and extrapolates to 50 % at infinite lip(SH)2 concentration suggesting that only half of the enzyme reacts inth e fast phase.Thi s will bediscusse d below. Inth e oxidative reaction with NAD+a t pH 8.0,a simila r feature isobserved .Th e contribution of the amplitude of thefas t phase increases from 25% a t [NAD+]= Kd to 58% a t [NAD+]= 10time s Kd- In this reaction no absorbance changes at 380 nmar e found that indicate the formation of a C4a adduct species.Th e oxidative reaction of EH2o f enzyme Glu455->Gln with lipS2 at pH8. 0 isver y slow and almost beyonddetection . Fig. 5 shows rapid scan spectra obtained from reoxidation of EH2o f Glu455->Gln by 20 equivalents NAD+. EH2bind s NAD+rapidl y togiv e a spectrumtha t closely resembles the absorption spectra obtained fromtitratio n of pig heart EH2wit h NAD+[10 ] andtitratio n of pig heart enzyme with NADH inth e presence of high [NAD+] ref.[39] . This spectrum represents a reduced-disulfide-oxidized-flavin-NAD+ intermediate species (species 4 in Scheme 1). Intramolecular transfer of a hydride equivalent from thedithio lt oth e flavin is biphasic and relatively slow, and results in reduction of most of the flavin despite the presence of a large excess of NAD+.A tthi s stageth e reaction is thermodynamically blocked. The reductive reaction of Eox Glu455->Gln with 2equivalent s NADH is shown inFig . 6.Th edea dtim e spectrum showstha t the reduction of the FADi sver y fast (species 6i n Scheme 1).Th e subsequent intramolecular electron transfer to the disulfide is again relatively slow. Following reduction of the disulfide, reaction with a second moleo f NADH is slow.

68 A 0.5 B 8 °-4 ^-^__1 | 0.3 // -^2 i J 0.2 . 3 _ 0.1 ~~~^~^ 2 0.0 " i i i 0.01 0.1 1 10 Time (s)

1 I.I. 400 500 600

Wavelength(nm )

Fig 5. Spectral changes of2-electron reducedAzotobacte r vinelandii Glu455->Gln lipoamide dehydrogenase upon oxidation by NAD+in the rapid scan instrument. A. Oxidation of 2-electron reduced enzyme Glu455->Gln by 20 molar equivalents of NAD+a t pH]7.8 inth e rapid scan spectrophotometer. EH2, ;dea dtim e (5.42 ms after mixing), - • -; 306 msafte r mixing, ;30. 6s afte r mixing B.Tim edependenc eo f ab^orbance changes of panelA . Curve 1,457 nm;curv e 2,530 nm;curv e 3,670 nm.EH 2 wa|igenerate d by reductionwit h an excess of sodium borohydride withtim e forth e excess to sell destruct.

0.6 A

0.4

0.0 1 1

Wavelength(nm )

Rg. 6. Spectral changes ofAzotobacte r vinelandii Glu455->Gln lipoamide dehydrogenase upon reduction byNADH in the rapid scan instrument. A. Reductiono f enzyme Glu455->Gln by 2 molar equivalents of NADH atp H 7.8 inth e rapid scai spectrophotometer. Eox, ;dea dtim e (5.42 ms after mixing), ;61 2 ms aftar mixing, ;30. 6 safte r mixing B.Tim edependenc e of absorbance changeso f panel A. Curve 1,457 nm;curv e 2,530 nm;curv e 3,670 nm.

69 Enzyme Glu455->Asp differs from Glu455->Gln in spectral properties during reduction bytNADH (Table 2). At all NADHconcentration s used,enzym e Glu455+>Asp shows more extensive reduction of the flavin than enzyme Glu455->Gln andth e subsequent reactions are less clear (results not shown) The results obtained fromth e rapid reaction studies onth e Glu455 mutated enzymes clearly demonstrate that the intramolecular transfer of a hydride equivalent in either direction is slowed down though not as severely as in enzyme His450->Ser. The underlying mechanism might beth e same as for enzyme His450->Ser and will be discussed below.

Enzyme Pro451->Ala. The enzyme Pro451->Ala appears to be essentially inactive in both directions. Spectral studies however demonstrate that this enzyme can be reduced by both lip(SH)2 and NADH, but rapid over-reduction to the catalytically inactive EH4occurs . Stopped flow studies at 530 and 450 nmsho w the rapid formation of atransien t charge-transfer complex between thiolate and FAD upon reduction with lip(SH)2 (within 5 s) followed by a slower reduction of the flavin (results not shown). Enzyme Pro451->Ala also becomes fully reduced by NADH in 10 mswithou t any detectable changes at 530 nm.Th e fact that the reduction by NADH is much fastertha n the over-reduction by lip(SH)2 indicates that the absence of detectable activity inth e physiological reaction is mainly due to the over-reduction by NADH generated after one or afe w cycles of turnover.

Redox potentials. Spectral studies have suggested differences in redox potentials between wild-type and mutated enzymes [26].Th e different sensitivity of the enzymes toward inhibition by NADH also indicates that differences in redox potentials may exist. Quantitative estimation ofth e redox potentials as measured by three different methods however failed. The main reasonfo r this isth e uncertainty ast o the exact contribution of the various EH2species . During the reduction to EH2an d subsequently EH4n o isosbestic points are observed. Although no exact redox potentials are determinable, the results obtained allow an estimation of redox potentials of the enzymes in relation to each other. The arrangement of the enzymes in order of decreasing redox potentials (E'm EH2/EH4) is:Glu455->Asp , Glu455->Gln,wild-typ e and His450->Ser. This trend coincides with the sensitivity towards inhibition by NADH. Several lines of evidence indicate that binding of NAD+ induces an increase of the redox potential ofth e FAD [10, 40].Therefor e thecours e of reduction of wild-type enzyme and enzyme Glu455->Asp by NADPH in an NADPH generating system was studied inth e presence of 100|a. M AAD+[10,17 , 41],a non-reducible NAD+analo g (Kj

70 AAD+ for wild-type enzyme inth e reaction lip(SH)2/NAD+i s4 0 u.M).Th e formation of NADPI-fiis rate limiting in bothth e absence and presence of AAD+an d equal amounts of gluccjse-6-phosphate dehydrogenase were used.Th e results are very conclusive (Fig. 7,onl y wild-type enzyme shown): no NADPH absorbance is observed inth e presencje of AAD+ untilth e enzyme is almost completely reducedt o EH4.

0.4 - A B S 0.3 - a i\ •e 0.2 41£i$ \ il < 0.1 _ "-• v-"~-Sra^-L. 0.0 1 1 1 ,"1"""~i 400 500 600 700 400 500 600 700 Wavelength (nm) Wavelength (nm)

Fig. 7. Spectral changes upon reduction ofAzotobacte r vinelandii wild-type enzyme by NADPH in the absence/presence of AAD+. Conations for reductiono f wild-type enzyme by enzymatically formed NADPH are described inMaterial s and Methods.A . reduction ofwild-typ e enzyme by NADPH.Selecte d spectra are shoWn. B. reduction of wild-type enzyme by NADPH inth e presence of 100 u.MAAD+ . Selected spectra are shown.A correctio nwa s madefo rth e contribution of AAD+. The same amount of glucose-6-phosphate dehydrogenase was used in A and B.

Theise results for the first time clearly establish that binding of a pyridine nucleotide (AAD+) ndeed raises the EH2/EH4 redox potential. Additional support is obtained by the addition of AAD+t o EH2o f wild-type enzyme and enzyme Glu455->Asp, generated by the addiion of 5 molar equivalents of lip(SH)2, Fig.8 . Before addition of AAD+wa s startedti e EH2spectr a were at equilibrium.Additio n of 40 u,MAAD +result s ina shift towards longer wavelength and an increase of the charge-transfer absorbance for both

0.30 B 0.25 % 1 0.20 •*/ \ | 0.15 1 0.10 < 0.05 I-^. 0.00 1 1 1 1 400 500 600 700 400 500 600 700 Wavelength (nm) Wavelength (nm)

Fig. 8. Theeffect of addition ofAAD+ to EH2of Azotobacte r vinelandii lipoamide dehydrogenase. EH2wa s generated byth e addition of 5 molar equivalents of lip(SH)2 to the enzyme ina n anaerobic cuvette at pH 7.0. Spectra shownwer e recordedafte r changes were complete ,Eox ; , EH2; ,afte r additiono f 200 uMAAD+ .A , Wild-type lipoaJnide dehydrogenase; B, Glu455->Asp lipoamide dehydrogenase.

71 enzymes (spectra not shown) similart o the result reported forth e pig heart enzyme [10]. Subsequent additions of AAD+ (upt o 200 uM) have noeffec t onth e initially obtained spectrum of wild-type enzyme after addition of 40 uMAAD+ . However for enzyme Glu455->Asp a bleaching of the spectrum is observed,demonstratin g further reduction to EH4b yth e residual lip(SH)2. Forenzym e Glu455->Asp it isthu sclea rtha tAAD + effects an increase inth e redox potential ofth e FAD. Forwild-typ e enzyme the further reductiont o EH4i s not observedthoug h aprofoun d enhancement of the 530 nm absorbance occurs which may also be relatedt o achang e ofth e redox potential of the FAD aswa s suggested by Matthews andcoworker s [10] andThorp e andWilliam s [17].

Discussion

For the reductive reaction of pig heart lipoamide dehydrogenase with lip(SH>2a mechanism has been proposed in which a histidine functions as a proton acceptor/donor to yieldth e nucleophile necessary for attack of the disulfide [9,20] .A mixed disulfide species, species 2 in Scheme 1, was postulated as an intermediate and the breakdown of this species to yield EH2(specie s 3 in Scheme 1)wa s assumedt o be rate limiting forth e pig heart enzyme[9] . The rapid reaction kinetics for the wild-type enzyme shows a 2-fold increase in reductive rate by lip(SH)2 between pH7. 0 and8. 0 (Table 2). Inthis ,th e enzyme differs from pig heart enzyme where the rateconstant s were invariant between pH 5.5 and 8.0. At present we can not satisfactorily explain this discrepancy. For pig heart enzyme it hasbee n showntha t below pH 7.6 the reoxidation reaction becomes rate limiting in overall catalysis indicating that only the EH- species is capable of reacting with NAD+[10] . For A vinelandiienzym e the oxidation rate of EH2b yNAD + decreases from pH 8.0 to 6.8 indicatingtha t here also EH- isth e reactive species since this species prevails at higher pH[26] . The reaction of wild-type and all mutated enzymes with NADH,th e reverse reaction, allowsth e distinction of separate steps,a featur e that is most prominent in enzyme His450->Ser where the rate of reduction of the disulfide by NADH is slowed by more than 1000fold . The binding of NADH and subsequent reduction of the FAD is in all cases very fast yielding the FIH--NAD+specie s as an intermediate, species 6 in Scheme 1. The subsequent intramolecular transfer of a hydride equivalent leading to reduction of the disulfide and reoxidation ofth e FAD (species 4) isstrongl y affected byth e mutations. Also,th e forward reaction (species 4t o 6) isslowe d byth e mutations, more than 200 fold for enzyme His450->Ser.

72 Evidencefo r the formation ofth e proposed flavin C4aadduc t intermediate (species 5) as shown in Scheme 2 is not obtained under the experimental conditions applied.A plausible explanation for this istha t the rate of formation ofth e presumed C4aadduc t is slow in either direction when comparedt o its breakdown. Leichus and Blanchard [42] showedrecentl y that inth e forward reaction ofth e pig heart enzyme the formation of the proposedspecie s 5 is most likely the solvent isotopically sensitive transfer step under conditions of saturating [lip(SH)2] and [NAD+] variable. This is in agreement with our suggestion above,tha t inth e enzymes having either the base, His450, or itscharge - relay partner, Glu455,altered ,formatio n ofth e C4aadduc t isslo w relative to its breakdown.

Charge-transfer C4a-adduct reducedflavi n

N N Sy/ V \>°

\ oo / o O —S " —S / ° —s 1 + + _SH HHis- _s" HHis- 1 His- — S

Schpme 1. Formationan dbreakdow no fth ethiolat et oC4a-FA D adduct. Fromre f[17] .

ever, inth e reverse reaction of pig heart enzyme the formation of species 4i s ely solvent isotopically sensitive under conditions of saturating [lipS2] and variable [41].Thi s isi ncontras t withth e results obtained forth e mutated s where, again no C4a adduct is observed.Tw o possible reasons forthi s canb e :ed. First,th edeca y ofth e C4aadduc t might befaste r than itsformation .Second , bsence of a base,th e transfer of a hydride equivalent may become concerted, pening of the disulfide to formth e C4a adduct species, a negative charge s onth e interchange sulfur. It isquit e possible that a protonated histidine,th e necessary to stabilize this negative charge,an dtherefore ,th e second lity is probably favored. reoxidation reaction of EH2o f enzyme His450->Ser by NAD+i s not significantly d at pH 6.8 whencompare dt o pH8.0 .Thi s is unexpected in light of the ig. This mutated enzyme stabilizes thiolate to FADcharge-transfer . Therefore, n by lip(SH)2 requires the loss of 1proto n to reach afor m analogous to species

utth e base, EH\ ApK awa sdetecte d at 7.5 forth e His450->Ser enzyme and iteda s an EH-t o E2_deprotonatio n [26].Thi sdeprotonatio n shoulddisfavo r the n of NAD+.Th e almost complete absence of any reactivity atth e EH2leve lo f His450->Ser with lipS2a t pH 8.0 agrees withth e observation for pig heart

73 enzyme lacking reactivity of EH-wit h lipS2[9] ,an dou rconclusio n istha t the main electronic species of 2-electron reduced enzyme His450->Ser is E2_[26] . The reductive reaction of enzyme His450->Ser by lip(SH)2 isver y slow andfull y compatible with the mechanism proposed [1, 9].Th e observed short lag-phase is in agreement with a slow formation of the proposed mixeddisulfid e species (assuming that this species has no absorbance) asth e catalyst,th e histidine, is missing. Simulation ofth e reaction showstha ttrace s compatible with observedtrace s are only obtained when the breakdown of the mixeddisulfid e occurs 10-foldfaste r than itsformation .Fro m this we conclude that the rate limiting step inth e reduction of enzyme His450->Ser by lip(SH)2 isth e formation of the mixeddisulfid e intermediate. The Glu455 mutated enzymes arequit e different inthei r rapid reaction kinetics from wild-type enzyme. The data indicate that like the His450->Ser enzyme, intramolecular electron transfer between FAD/FADH- andth e disulfide/dithiol is severely impaired. Thus,an y alteration of the His-Gludia d affects not only deprotonation of the dithiol substrate, but also communication between the enzyme's two redox active couples. In addition,th e Glu455 mutated enzymes differ from His450->Ser enzyme in several details of their rapid reaction kinetics. Both enzymes Glu455->Asp and -Gin show clear biphasic kinetics during reduction of Eox with lip(SH)2 or reoxidation of EH2wit h either lipS2o r NAD+. An interesting feature of this biphasic behavior isth e fact that the amplitude of the fast reaction increases with substrate concentration and that it reaches about 50% o fth e total amplitude change.Thi s indicates that only one of the active sites of enzyme Glu455->Gln is reduced atth e rapid rate. Moreover, the KGln with lip(SH)2 at pH8.6 ) or lower than (enzyme Glu455->Gln EH2wit h NAD+a t pH8.0 ) those calculated forth e slower amplitude change. These results strongly indicate that cooperativity between the two active sites ofth e enzyme occurs. Itwa s shown previously that enzymes Glu455->Asp and -Gin show a remarkably different fluorescence quenching behavior when compared toth e other (mutated) enzymes [26]. Itwa s concludedtha t this was either duet oa n intrinsic property of these enzymes or due to cooperativity induced byth e quencher. Basedo n the present results it is now suggested that both the fluorescence quenching behavior andth e biphasic rapid reaction kinetics are due to cooperativity. Cooperativity was shown recently forth e related enzyme mercuric ion reductase [19] andwa s shown also for (mono-alkylated) pig heart enzyme reacting with NAD+ [16-18]. The results (Fig.7 ) obtained for the reduction of wild-type enzyme by NADPH inth e presence of AAD+ support cooperativity between the subunits. Since lipoamide dehydrogenase possesses only one NAD+bindin g site per active site, based onth e extensive structure homology between glutathione reductase and lipoamide dehydrogenase from P.putida [43] , binding of bothAAD +an d NADPH to one active site

74 simultaneously can be excluded.Th e rise ofth e redox potential ofth e flavin of the active site reducedt o EH2(sit e a) cantherefor e only be explained by binding of AAD+t o the other ajctive site (site b) or by simultaneous binding of either AAD+o r NADP(H)(+)t o site a andty. Whethe r site bi s Eox or EH2i s not clear. Alteration of Glu455,th e charge-relay residue inth e base diad, effects profound changes in catalysis. The lip(SH)2/NAD+ activity is unmeasurable due to over-reduction by the product NADH. The lip(SH)2/AcPyAde+activit y iscirc a 2%o f wild-type. Glu455 alters tne acid-base properties of His450 [26] and positions the imidazole ring presenting its N3towar dth e redox active disulfide. Both mutations can be considered conservative, Glu455->Gln and Glu455->Asp. Indeed,th e almost normal rateo f reduction of Glu455->Asp enzyme by lip(SH)2 demonstrates that the aspartate can mimicth ecatalyti c effect ofth e glutamate inth e enzyme substrate complex. Neitherth e aspartate north e glutamine residues are able however to accomplish the function of the base djad inth e intramolecular transfer of electrons between FAD/FADH- andth e disulfidB/dithiol. In contrast to the His450->Ser enzyme where the problem appears to be largely kinetic,th e effect with enzymes Glu455->Gln and Glu455->Asp seems to be both kinetic andthermodynamic . Thus,th e extent of reoxidation of 2-electron reduced enzym* Glu455->Gln by NAD+i sver y small (Fig.5) .Th e binding of NAD+ raisesth e redox potential of FAD relativet o that of the dithiol. Electrons pass,albei t slowly,t o the FAD bijt the resulting FADH- is not reoxidized.

Inhibition of A. vinelandiiwild-typ e enzyme by NADH above pH 8.0 is not observed in pig hteartenzym e while for E.co//enzym e this inhibition isobserve d at all pH values studied( For E.coli enzym e it was showntha t the inhibition iscause d by over-reduction of the ^nzymefro m EH2t o EH4b y NADH andtha t there isa direc t relation betweenth e inhibition andth e redox potential of the EH2/EH4coupl e [12]. Like the E.coli enzym e [7], inhibition of the A. vinelandiienzym e could becircumvente d in kinetic studies by using AcPyA

75 enzymes Glu455->Gln, His450->Ser and wild-type enzyme. These data taken together indicate that the over-reduction andthu s susceptibility to inhibition by NADH,i s directly related to differences in redox potentials. The relation between the relative redox potentials of the mutated enzymes andth e susceptibility to inhibition by NADH supports this hypothesis. Assuming that the change inth e redox potential with pHfo r the A. vinelandiienzym e relative to NADH is similar to that of pig heart enzyme [20],over-reductio n wouldb e favoured at lower pH.Th e inhibition however becomes apparent at a pH where the histidine likely deprotonates (pKa = 8.1, inferred fromth e pKm AcPyAde+ plot).Ou r results show that binding of a nucleotide raisesth e redox potential ofth e EH2/EH4 couple. Apparently this rise is enhanced when the histidine is deprotonated. This indicates that the protonated histidine, at the EH2 level,play s an important role in maintaining the redox potential of the active site carefully balanced to avoid over­ reduction by NADH.Th e results ofth e steady state kinetic studies onth e Glu455 mutated enzymes lend strong support to this hypothesis. Overth e entire pH range studied strong inhibition by NADH is observed for both enzymes while it was shown previously that the pKa of the histidine inthes e enzymes is considerably lowered (pKa< 6.5) when comparedt o wild-type [26].Thi s adds a second function to His450; not only does it serve asth e base for deprotonation of lip(SH)2,i t modulates the relative redox potentials of the disulfide/dithiol and FAD/FADH"couple s which may be reflected inth e markedly diminished rates of intramolecular electron transfer in the altered enzymes.

Acknowledgments.

Wethan k Dr.W.G.J .Ho ian d Mr.A . Mattevifo r providing uswit hth ecoordinate s of lipoamide dehydrogenase from Azotobacter vinelandiiprio r to publication. This work was supported by the "Dutch Foundation for Chemical Research' (SON) with financial aidfro m the 'Netherlands Organization for Scientific Research' (NWO), by the Health Services and Research Administration ofth e Department of Veterans Affairs andb y grant GM21444 (CHW) from the National Institute of General Medical Sciences.

References.

1. Williams,C.H .Jr . (1991) In Chemistryand Biochemistry of Flavoenzymes (Muller,F. , ed.)vo l III, inpress ,CR C Press Inc., BocaRaton .

76 2. £okatch,J.R. , McCully,V. ,Gebroski ,J .an dSokatch ,D.J . (1981) J. Bacterial. 148,639-646 . 3. fox, B.S. and Walsh,C.T . (1982) J. Biol.Chem. 257,2498-2503 . 4. £hames, S.L., Fairlamb,A.H. ,Cerami ,A . andWalsh ,C.T . (1986) Biochemistry )?5, 3519-3526. 5. Holmgren, A. (1980) Experientia suppl. 36, 149-180. 6. yiassey,V. , Gibson,Q.H .an dVeeger , C. (1960) Biochem. J. 77,341-351 . 7. iJMIIiams,C.H .Jr . (1965) J. Biol.Chem. 240,4793-4800 . 8. fteed, J.K. (1973) J. Biol.Chem. 248,4834-4839 . 9. l/latthews, R.G., Ballou,DP. ,Thorpe ,C .an dWilliams ,C.H .Jr . (1977) J. Biol. Chem.252, 3199-3207. 10. l/latthews, R.G., Ballou,DP . andWilliams ,C.H .Jr . (1979) J.Biol. Chem. 254, 4974-4981. 11. Wilkinson, K.D. and Williams C.H.Jr . (1979) J. Biol.Chem. 254,852-862 . 12. Wilkinson, K.D. and Williams C.H.Jr . (1981) J.Biol. Chem. 256, 2307-2314 . 13. ^ahlman, L. andWilliams , C.H.Jr . (1989a) J. Biol.Chem. 264, 8033-8038 . 14. ^ahlman, L. andWilliams , C.H.Jr . (1989b) J. Biol.Chem. 264, 8039-8045 . 15. ^assey, V. and Veeger, C. (1961) Biochim. Biophys. Acta48, 33-47 . 16. thorpe, C. andWilliams , C.H.Jr . (1976) J.Biol. Chem. 251, 7726-7728 . 17. Thorpe, C. andWilliams , C.H.Jr . (1981) Biochemistry20, 1507-1513. 18. ^/luiswinkelvan-Voetberg , Han d Veeger, C. (1973) Eur.J. Biochem.33, 285-29 1 19. filler, S.M., Massey,V. ,Williams ,C.H .Jr. , Ballou,DP . andWalsh ,C.T . (1991) tyiochemistry 30,2600-2612 . 20. Matthews, R. G. andWilliams , C. H.Jr . (1976) J. Biol. Chem. 251, 3956-3964. 21. Adamson, S. R., Robinson,J .A . and Stevenson, K. J. (1984) Biochemistry23, 1269-1274. 22. Berry,A. , Scrutton,N.S . and Perham, R.N. (1989) Biochemistry28, 1264-1269. 23. Deonarian, MP., Berry,A. ,Scrutton , N.S. and Perham,R.N . (1989)Biochemistry 28, 9602-9607. 24. Williams,C.H .Jr. , Allison,N. ,Russel ,G.C. , Prongay,A.J. , Arscott, D.L, patta, S., Sahlman, L. and Guest,J.R . (1989) Ann.N.Y. Acad.Sci. 573, 55-65. 25. Benen,J.A.E. , Berkelvan ,W.J.H .an d Kokde ,A . (1990) in Flavinsand Flavoproteins1990 (Curti, B., Ronchi,S .an dZanetti ,G .eds. ) pp557-564,Walte r qleGruyte r and Co. Berlin-New York 1991, 26.

77 28. Miller, S.M.,Massey ,V. , Ballou,D. ,Williams ,C.H .Jr. , Distefano,M.D. , Moore, M.J. andWalsh ,C.T . (1990) Biochemistry29, 2831-2841. 29. Meada-Yorita, K., Russel,G.C. ,Guest ,J.R. , Massey,V .an dWilliams ,C.H .Jr . (1991) Biochemistry30, i nth e press. 30. Vervoort, J. (1986) PhDThesis , University of Wageningen,Th e Netherlands. 31. Berkelvan ,W.J.H. ,Regeling ,A.G. ,Beintema ,J.J .an dd e Kok,A . (1991) Eur. J. Biochem.202, 1049-1055. 32. Broek vanden ,H.W.J . (1971) PhDThesis ,p . 8, University of Wageningen,Th e Netherlands. 33. Barshop, B.A., Wrenn, R.F. and Frieden,C . (1983) Anal.Biochem. 130, 134-145. 34. Massey, V. (1990) in Flavinsand Flavoproteins 1990 (Curti,B. , Ronchi,S .an d Zanetti, G.eds. ) pp.59-66 ,Walte r de Gruyter andCo . Berlin-New York 1991, 35. Dixon,M .an dWebb ,E.C .(1979 ) Enzymes, pp. 138-164, R. Clay Ltd.,Bungay , Suffolk, UK 36. Kaplan,N.O . (1960) in The Enzymes(Boyer , Lardy and Myrback eds.) vol.3 ,p . 151, Academic Press, NewYork . 37. Clarke,W.M . (1960) Oxidation-Reduction Potentials of OrganicSystems, The Williams andWilkin s Co., Baltimore, p.131 . 38. Palmer, G. and Massey, V. (1968) in Biologicaloxidations (Singer , T. P. ed.) part 2, p. 288, Interscience Publishers, New York. 39. Veeger, C. and Massey, V. (1963) Biochim.Biophys. Acta 67,679-681 . 40. Maeda-Yorita, K. andAki ,K . (1984) J. Biochem. 96,683-690 . 41. Fisher, T.L.,Vercellotti ,V . andAnderson , B.M. (1973) J. BiolChem. 248,4293 - 4299. 42. Leichus, B.N. and Blanchard,J.S . (1991) in Flavinsand Flavoproteins 1990 (Curti, B., Ronchi,S .an dZanetti ,G .eds. ) pp589-592 ,Walte rd e Gruyter andCo . Berlin-New York 43. Mattevi,A. ,Obmolova ,G. ,Sokatch ,J.R. , Betzel,C .an d Hoi,W.J.G .(1991 ) Submitted.

78 Chapter 4

Lipoamide Dehydrogenase from Azotobacter vinelandii. The role of the C-terminus in catalysis and dimer stabilization.

Jacques Benen,Wille mva n Berkel,Cee s Veeger andAri ed e Kok.

Summary

The|10 C-terminal residues are not visible inth e crystal structure of lipoamide dehydrogenase from Azotobacter vinelandii, but can be observed inth e crystal structures of the lipoamide dehydrogenases from Pseudomonasputida [Mattevi,A . et al., submitted] and Pseudomonas fluorescens. Inthes e structures,th e C-terminus folds backtoward s the active site and Is involved in interactions withth e other subunit. The(functio n of the C-terminus of lipoamide dehydrogenase from Azotobacter vinelandiiwa s studied by deletion of 5, 9 and 14 residues, respectively. Deletion of the last 5 residues does not influence the catalytic properties and conformational stability (thermojnactivation and unfolding by guanidinium hydrochloride). Removal of 9 residues results in an enzyme (enzyme A9) showing decreased conformational stability and higfi sensitivity toward inhibition by NADH.Thes e features are even more pronounced after deletion of 14 residues. In addition tyrosine16 , conserved in all lipoamide dehydrogenases sequenced thus far, andshown from the other structures to be likely involved in subunit interaction,wa s replaced by phenylalanine and serine. Mutation of Tyr16 also results in a strongly increased sensitivity toward inhibition by NADH.Th e conformational stability of both Tyr16 rotated enzymes is comparable to enzymeA9 . Thejresult s strongly indicate that a hydrogen bridge between tyrosine of one subunit (Tyr16 iji the A vinelandiisequence ) and histidine of the other subunit (His470 inth e A vinelanaiisequence) , exists inth e A vinelandiienzyme. I nth e A9 andA1 4enzyme s this interaction is abolished. It isconclude dtha t this interaction mediatesth e redox properties of the FADvi ath e conformation ofth e C-terminus containing residues450 - 470.

79 Introduction.

Lipoamide dehydrogenase is a member of the flavoprotein disulfide oxido reductases [1].Th e homodimeric enzyme catalyzes the NAD+dependen t reoxidation of dihydrolipoyl groups,whic h are in vivocovalentl y attachedt o the lipoate acyl transferase component of several multienzyme complexes [2]. Each active site contains an FAD and a redox active disulfide bridge and isconstitute d by amino acids from both subunits [3,4 ,5] . For lipoamide dehydrogenase from Azotobacter vinelandiith e crystal structure of the oxidized state has been determined at 0.22 nm resolution [4].Th e geometry of the active site is highly identicalt o the active site of humanglutathion e reductase [4]. Unfortunately, no crystal structure of reduced A. vinelandiilipoamid e dehydrogenase or mutated enzymes is available.

Tyr 18'

.Gin 26"

195 0„

Qu 442

Fig. 1. Schematicpicture of theinteractions madeby theC-terminal residues at the si-side of the flavin of lipoamide dehydrogenase from Pseudomonas putida. Residues of the opposite subunit are indicated by aprim e symbolfollowin g their sequence number. H-bonds (d< 0.34 nm) are shown by dashed lines,whil e the dotted lines indicate distances between residuesdirectl y or indirectly involved incatalysis . Corresponding numbering of relevant conserved residues of A. vinelandii enzyme in brackets:Tyr18 ' (16'),Gln26 ' (24'), Cys43' (48'), Cys48' (53'), Lys52' (57), Glu189' (194'), His437 (450),Glu44 2 (455),Glu44 6 (459), His457 (470).Thi s figure was kindly provided by Mr.A .Mattevi .

80 Lipiamide dehydrogenase from A. vinelandiicontain s a 15 amino acid C-terminal extension comparedt o glutathione reductase [6,7] .Th e last 10 residues are disordered and no:visibl e inth e electron density map [4]. It wassuggeste dtha t these residues constitute a mobile structure onth e outside of the molecule and might serve inth e interaction with the core component of the 2-oxo acidcomplexes ,th e acyitransferase . Deletiojri mutagenesis experiments however have shown that this hypothesis is not correct|[8]. Furthermore itwa s showntha t deletion of 9o r 14o f the C-terminal residues (enzymesdesignate d A9 andA14 ) strongly affects catalysis [8].A possible cluet o the role ofth e C-terminal residues is given byth e recently elucidated X-ray structures of lipoamfcledehydrogenas e from Pseudomonas putida [5] and P. fluorescens[Mattev i and Hql, personal communication]. Inthes e enzyme structures,th e C-terminus folds partly back intoth e protein structure comprising the active site.Th e relevant part of the structure is shown in Fig. 1. Thase new structures also showtha t atyrosin e (Tyr16 inth e A. vinelandiienzyme) , which is strictly conserved in all lipoamidedehydrogenase s sequenced thus far, could be involved in interaction withth e C-terminus ofth e other subunit. Here we report onth e characterization ofth e C-terminal deletion enzymes,A5 ,A 9 andA1 4 andth e mutated enzymes Tyr16->Phe,Tyr16->Ser . Based onth e results presented inthi s paper it is concludedtha t inA. vinelandiienzym e the C-terminus conformation issimila r totha t observed inth e Pseudomonasenzymes . Based on this feature)a coheren t explanation ofth eeffect s of the mutations isdiscussed .

Materials and Methods.

General. All biochemicals, bacterial strains and vectors used were described elsewhtere [9, 10].Th e mixed oligonucleotide allowing mutagenesis of Tyr16 into Phe and Sarwa s synthesized according to [9].

Construction of plasmids. The construction of plasmid pJABIOOO, carrying the wildtyp e Ipd-gene wasdescribe d in [8].A partial restriction map is shown in Fig.2 .Th e procedure for mutagenesis of Tyr16wa s identicalt o the one described in [9], except for the cassette. The cassette was obtained bydigestio n of pJABIOOOwit hth e restriction endonucleases Splr\ and Kpn1. The Spfrl/Kpn1 fragment, encoding the N-terminal part of the Brf-gene, containing Tyr16, wasclone d into Sp/71/Kpn1 digested M13mpb18 and subjectedt o mutagenesis. After screening forth e desired mutations by sequencing the cassette was excised by digestion with Hind\1 1an d Kpn1. Before religating the mutatepcassett e into pJABIOOOth etw o Kpn\ sites,on e immediately downstream of the

81 •I 5-a ii i i J L yi///yjf i r - - aceF Ipd il\sssss

i • 0 IO'OO 2000 basepair s

Fig. 2. Partial restrictionmap of theplasmid pJABWOO carrying theIpd-gen e from Azotobacter vinelandii, encoding lipoamide dehydrogenase, and part of the aceF-gene encoding acetyl transferase. The asterisk (*) indicatesth e positiono f Tyr16.

/pd-gene andth e other upstream,wer e destroyed.Th e upstream Kpn\ site was destroyed by digestion of pJAB1000 with Pst\ and Spfrl. The sticky ends were filled with T4-DNA polymerase and religated yielding plasmid pEH1. From this plasmidth e downstream Kpn1 site was removed by partially digesting pEH1 with Kpn1 and filling the sticky endwit h T4-DNA polymerase.Afte r checking for the desired change the H/nc/111/Kpnl fragment of the altered pEH1 (plasmid pEH12) was replaced withth e mutated cassette yielding an intact /pd-gene carrying either the Tyr16->Phe or Tyr16->Ser mutation. The construction of enzymes lacking the C-terminal residues (enzymes A5,A 9an d A14) was described previously [8].

Analytical methods. Purification of mutated enzymes was according to [9]. Spectral studies and stopped flow kinetics were performed in 100 mM Hepps,0. 5 mM EDTA, 150 mMioni c strength pH8. 0 (Hepps/EDTA buffer), as described [10]. Steady state kinetics were studied accordingt o [10] in 100 mMHepes ,0. 5 mMEDTA , 150m M ionic strength pH 7.0. Thermoinactivation was determined as described in [11] using lip(SH)2 and 3-acetylpyridine dinucleotide [AcPyAde+] as substrates in activity determinations for the Tyr16 mutated enzymes and NADH and dichlorophenolindophenol for enzymes A5,A 9 and A14. Unfolding experiments were performed according to [11]. Calculation of the fractional contribution of the different enzyme species,th e oxidized enzyme (Eox),th e 2-electron reduced enzyme (EH2) andth e 4-electron reduced enzyme (EH4) as shown in Fig.4 ,t oth e mixed spectra in Fig.5 i s performed asfollows . It isassume dtha t the EH2spectr a in Fig.3 as obtained after reduction with borohydride represent fully formed (100 %) EH2.Th e EH4spectru m was obtained by reducing the enzyme with 1M dithiothreitol .A species inwhic h the FAD is reduced andth e disulfide oxidized as a result of translocation ofth e electrons in EH2 (designated FIH")i s

82 spectrally indistinguishable from EH4.Sinc e Eox and FIH-/EH4d o not absorb at 540 nm, the ex!notio n at 540 nm ofth e mixed spectra in Fig.5 ca n directly be usedt o calculate the fraction of EH2. Since Eox and EH2ar e isosbestic ind e region of 445 nman y decrease in extinction of the mixed spectra atthi s wavelength whencompare dt oth e extinction of Eox and EH2i sa result fromth eformatio n of FIH-and/o r EH4.A tth e isosbestic point (see table 2)th e fractional contribution of FIH-+EH4 can be calculated in the following way:

EXmixed = FEOX X £XOX+ FEH2 XEA.EH 2 + (FRH"+FEH4) X EXEH4 (1) inwhia hexmixe di sth e extinction of the mixedspectru m atth e exact wavelength (X) of isosbeisticy of Eox and EH2, Fdenote s the fractional contribution ofth e species indicatedan dth e extinctions of Eox, EH2an d EH4a t wavelength X. are given by exx»x. EXEH2 find £XEH4 respectively and are derived from Fig.4 . Since e^oxan d £XEH2 are equal atth e isosbestic point and (FE0X + FEH2) = 0 - (FFIH~+FEH4))> equation (1)ca n be writtenas :

e^mixed =EXEH 2 x (1- (FFIH"+FEH4)) + (FFIH"+FEH4) X£x,EH 4 (2) resulting in equation (3):

eXEH2 miX ! FF|H-+FEH4= -^ ^ (3) 6XEH2"6XEH 4

Calculation of the individual fractions of FIH" and EH4i s possible by studying the electron distribution ina n equilibrating system atth e start and atth e endo f the equilibrium, provided that no extra input of electrons takes place during equilibration. In case ol the present study this calculation,wit h data presented in Table 2, isonl y done for reduction with 1 molar equivalent of lip(SH)2 (see below) where it can be assumed that no further reduction occurs. Reduction with 2 molar equivalents of lip(SH)2 shows a clear dscrease of Eox after equilibration indicating that further reduction hasoccurred . As un example the calculation of the fractional contribution of FIH" and EH4i s given for enzymeTyr16->Phe . As mentioned aboveth e FIH- species originates from electron transfer from the dithiolt oth e flavin. Iti sassume dtha t nofurthe r reduction by residual lip(SH)boccurs . The total fraction of electrons present for enzyme Tyr16->Phe immediately after reduction iseithe r0.6 5 or0.69 . (EH2+ FLH-o r EH4).Afte r equilibration the fraction of Eox has increased by 0.06 due to disproportionation at the expensteo f EH2.Consequentl y the fraction of EH4i s0.0 6 (2EH2->Eox +EH4) andth e fraction) of electrons herein is0.12 . Together withth e fraction electrons in EH2thi s adds

83 upt o 0.35. When it isassume dtha t the remainder isi nfac t FLH-thi sconstitute s a fraction of 0.30 (0.36-0.06 EH4).Th etota l ofth e fractions of electrons in EH2, EH4an d FLH-the n is0.6 5 which is exactly the same asth etota l immediately after reduction when it is assumedtha t here only is FLH'i s present. In an identical wayth e distribution of electrons for enzymes Tyr16->Ser andA 9ca n becalculated .

Results

Kinetics of the mutated enzymes. Previously it was reported that deletion of the 9 C-terminal residues (enzyme A9), residues results in strongly decreased activity inth e forward reaction [8].Thi s appearedt o because d by inhibition by NADH. Here activity could only be optimally detected with AcPyAde* as a substrate as observed with Glu455 mutated enzymes [10]. Enzyme A5 has kinetic properties essentially identical to wild type enzyme while enzyme A14i svirtuall y inactive (results not shown). TheTyr1 6 mutated enzymes also show severe inhibition by the product NADH inth e forward reaction while inth e reverse reaction, NADH/lipS2, no activity is observed in a standard assay. Using AcPyAde+ as an electron acceptor, theTyr1 6 mutated enzymes show an optimum at pH 7.0 instead of pH8. 0 asfoun dfo rth ewild-typ e enzyme [10].Tabl e1 shows the kinetic parameters of the Tyr16 mutated enzymes andth e wild-type enzyme at pH7.0 . The Lineweaver-Burk plots of the Tyr16 mutated enzymes are parallel (not shown),demonstratin g that the mechanism is still ping-pong. Mutation of Tyr16 into Phe or Ser only slightly affects the rate of turnover (Table 1). The reductive half reaction with

Enzyme V Km AcPyAde+ Km lip(SH)2

U/mg |iM \iM

wild-type 40 1000 70 Tyr16->Phe 20 800 80 Tyr16->Ser 14 1100 90

Table 1. Steady state kinetics of wild-typeand mutated enzymes Tyr16->Phe and Tyr1 6->Sero fAzotobacte r vinelandii lipoamide dehydrogenase. The kineticswer e determined with lip(SH)2 andAcPyAde + as substrates in 100m M Hepes,0. 5 mM EDTA, 150 mMioni c strength pH 7.0, at 25 °C. Thedat awer e analyzed accordingt o a ping-pong mechanism.

84 lip(SHfilc iissals o only slightly affected. Rapid reaction kinetics of the lip(SH)2 reduction at pH 8.0 result in rate constants of 2150s* 1 and 850 s-1 for enzyme Tyr16->Phe and Tyr16->Ser respectively withdissociatio n constants of 1.9m Man d 1.5m Mfo r L-lip(SH)2. Forwild-typ e enzyme this rate constant at pH8. 0 is200 0 S"1an d Kd= 0.9 mM[1W .Thes e results indicate that Tyr16 is not or only slightly involved in lip(SH)2 binding.

Spectral properties of the mutated enzymes. For the mutated enzymes A9, Tyr16-fPhe and -Serthe visible absorption spectra of Eox arever y similar to the wild- type eiizyme. Inal lthre e casesth e absorbance maximum is 1n mre d shiftedt o 458 nm. Enzyme A5 is spectrally indistinguishable from wild-type enzyme. The absorption spectrum of enzyme A14 shows less vibrational fine structure andth e absorbance maximum is4 nmblu e shiftedt o 453 nm (results not shown). FigJ3 show sth e spectra of enzymes Tyr16->Phe,Tyr16->Se r andA 9a s obtained after reduction with NADH. With enzymes Tyr16->Phe andA 9 reduction ofth e flavin is complete with only 1.5mola r equivalents (Meqs) of NADH added.Th e addition of1. 0

10 B 10-, \ v 8 " ' /~\ E T° 6 - '.>/'-«A 2 E t 4 / f \ I *C \ — — — 2 •* L - rr.r.--"-'—" 0 1 1 > 1 1 400 500 600 700 400 500 600 700 WAVELENGTH(nm ) WAVELENGTH (nm)

0 -\ c

8 -%i 6 4 ?

0 v 1— , 400 500 600 700 WAVELENGTH(nm )

FigJ3 . Visibleabsorption spectra of themutated Azotobacte r vinelandii lipoamide delwdrogenases Tyr16->Phe, Tyr16->Ser and ASupon NADH reduction. Allfire e enzymes were reducedwit h 1.0( )an d 1.5(2. 0 incase of Tyr16->Ser) ( ) molar equivalents of NADH.( ) Eox.Th e spectra shown arethos e recorded when changes were complete (after 15t o 20 min.). A, enzyme Tyr16->Phe; B, enzyme Tyri6->Ser; C enzyme A9.Th e experiments were conducted in Hepps buffer pH 8.0 at 251C.

85 Meqo f NADH results in more than 70%reductio n ofth e FAD.Thes e data demonstrate that the reducing equivalents mainly reside onth e flavin,favorin g the species FIH", instead of being mainly transferredt o thedisulfid e asfoun d for the wild-type [9],pi g heart [12,13] and mutatedenzym e His450->Ser [9]. Addition of upt o 1.0 mMNAD +t o enzyme reduced with 1.5Meq s of NADHdoe s not result in any reoxidation ofth e FAD. This failure of NAD+t o reoxidizeth e enzymet o EH2o r Eox isals o reflected inth e steady state kinetics. For enzyme Tyr16->Ser the spectra obtained upon NADH reduction are somewhat different from those of enzymes Tyr16->Phe andA9 .Wit hthi s enzyme slightly moretha n 2.0 Meqs of NADH are neededfo r complete reduction (Fig.3B) .Th e low 340 nm absorbance demonstrates that almost all NADH is oxidized inth e process of reduction, indicating formation of EH4.Again ,additio n of 1.0m Mo f NAD+t oth e fully reduced enzyme did not result in any reoxidation. Previously it was showntha t enzymes that are easily reduced by NADH and which

A B - A 10 r\ // \ \ 8 E \ • f\ •1 '*• \ ~ 6 \\// \\ "is 6 "U \\// M " / "" N I / * e 4 -'1 ** N \ / \ u - 2 ^l" \ 0 • " ~i-N ^-li; 400 50vV0 _ 60,0- 700 _ 400 500 600 700 WAVELENGTH (nm) WAVELENGTH (nm)

10 c ,• ^v^ 8 - /m\ 11 '\ N E A \ '/ \ 1 T° 6 _\ / \ \ 11 \l 2 W »1 I J *1 E 4 "J *" N \J *A w ' N v 2 _ V ^ \ % --.. •"-i-^- 0 1 d l--w.| 400 500 600 700 WAVELENGTH (nm)

Fig.4 . Visibleabsorption spectra of themutated Azotobacte r vinelandii lipoamide dehydrogenases Tyr16->Phe, Tyr16->SerandA9 after reduction with borohydride or dithiothreitol. The spectra of the borohydride reduction shown ( )wer e recorded immediately after additiono fth e reductant. Dithiothreitol reduction ( )wa s accomplished with 1.0M o f reductant. The spectra of the dithiothreitol reduction shown are those recorded when changes were complete. ( ) Eox.A , enzyme Tyr16->Phe; B, enzyme Tyr16->Ser; c enzyme A9.Th e experiments were conducted in Hepps buffer pH 8.0 at 25 °C.

86 show strong inhibition by NADH during turnover have a higher redox potential of the

FAD (EJmEH 2/EH4) [10,14,15]. In analogy tothi s it is suggestedtha t the redox potential ofth e HAD ofth e Tyr16 mutated enzymes andenzym e A9i s likely to be highertha ntha t of the wfild-typeenzyme . Fig.| 4show sth e spectra of enzymes Tyr16->Phe,Tyr16->Se r andA 9 as obtainedb y reduction with either excess of borohydride or dithiothreitol. Upon reduction with borohydrideth e three enzymes initially formth e typical EH2 spectrum withth e reducing equivalents resting onth e dithiol.Th e EH2specie s shown in Fig.4 , recorded immediately after addition ofth e reductant,ar ethu s only transiently stable.Th e least stable i$enzym e A9 andth e most stable isenzym e Tyr16->Phe. The spectra of enzymes Tyr16->|Phe,Tyr16->Se r andA 9ar e retained for approximately 15-20,5-1 0an d 2-3 minutedrespectivel y after which slow reduction^ the FADoccurs . InFig .5 spectrao f enzymesTyr16->Phe ,Tyr16->Se ran dA 9 respectively arepre ­ sentedu s obtained after reduction with lip(SH)2. Upon reduction of these enzymes with a small excess of lip(SH)2 over-reduction to EH4 and/or formation of FIH- and/ordis - proportionation isobserved .Typica l forthos e three enzymes isth e fact that the 530 nm charge-transfer absorbance during lip(SH>2 reduction never reaches values as high as foundwhe n using borohydride. Also the blue shift of the absorbance maximum is less,

400 500 600 700 400 500 600 700 WAVELENGTH (nm) WAVELENGTH (nm)

10 c 8 A /A -\ 6 '' *\\ //.''2\A\ 4 2 0 1 1 **-——r^T."B,-»i—... 400 500 600 700 WAVELENGTH (nm)

Rg.6 . Visibleabsorption spectra of themutated Azotobacte r vinelandii lipoamide dehydrogenases Tyr16->Phe, Tyr16->Ser and 49 upon Lip(SH)2 reduction. Thelspectra shown arethos e recordedimmediatel y afterth e additiono f 1.0( ,1 ) or 2.0 L ,3) molar equivalents of lip(SH)2an dafte rchange s are complete (after 2 houIs) afterth eadditio no f 1.0( ,2 )o r 2.0 ( • -, 4) molar equivalents. ( ) Eoxj A, enzyme Tyr16->Phe; B, enzyme Tyr16->Ser; C enzyme A9.Th e experiments weri conducted in Hepps buffer pH8. 0 at 25 CC.

87 suggesting that still Eox is present. Stoppedflo w studies show that for allthre e enzymes the 530 nnrvabsorbance rapidly increases mono-exponentially followed by a slow decrease at a rate compatible with the rate of over-reduction/ disproportionation observed inth e absorption spectra. Thisdemonstrate s that during recording of absorption spectra no initial high absorption at 530 nm ismissed . The spectra shown in Fig.5 ar ethos e obtained immediately after mixing with 1o r 2 Meqso f lip(SH>2an d after changes arecomplet e (after 2 hours).Whe n it is assumed that the EH2spectr a shown in Fig.3 represent fully formed EH2wit h maximalthiolat et o

FADcharge-transfe r it is possiblet o calculate the molefraction s of Eox, EH2 and FIH- +EH4i n each of the spectra shown in Fig.5 (see Materials and Methods). The calculatedfraction s are presented inTabl e 2. It isclea r that in each case the initial amount of EH2forme d decreases. After reduction with 1.0Me qo f lip(SH)2,th e decrease is mainly associated with an increase of FIH-+EH4 andonl y a small increase of Eox in case of enzyme Tyr16->Phe andA9 .Th e small increase of Eox relative to FIH-+EH4 indicates that no significant disproportionation occurs. This can be duet o either further reduction of Eox and EH2b y residual lip(SH)2 present, or intramolecular electron transfer fromth edithio lt oth e FAD.

Forenzym e Tyr16->Pheth e distribution of FIH"an d EH4 species, after the reduction with one equivalent of lip(SH)2 before and after equilibration, is outlined in the Materials and Methods section. Ina n identical wayth e fractionalcontributio n of FIH"an d EH4o f

Meqsl lip{SH)2 Tyr16->Phe Tyr16->Ser A9

Eox EH2 FIH-/ FIH-® EH4@ Eox EH2 FIH7 FIH-® EH4@ Eox EH2 FIH7 RH-@ EH4©

EH4* EH4* EH4*

1.0 0.35 0.61 0.04 0.04 0.00 0.44 0.52 0.04 0.04 0.00 0.26 0.68 0.06 0.06 0.00

1.0 final 0.41 0.23 0.36 0.30 0.06 0.44 0.24 0.32 0.32 0.00 0.40 0.14 0.46 0.32 0.14

2.0 0.17 0.70 0.13 - - 0.14 0.71 0.15 - - 0.12 0.73 0.15

2.0fina l 0.03 0.26 0.71 - - 0.11 0.13 0.76 - - 0.07 0.12 0.81

Table 2. Fractional contribution of oxidized,two- and four-electron reduced species to lip(SH)2 reduced Tyr16->Phe, Tyr16->Serand 69 mutatedAzotobacte r vinelandii lipoamide dehydrogenases. Spectraldat a are presented in Fig.4 .Analysi s isdescribe dunde r Materials and Methods. HMeqs :mola r equivalents added.# Fractionalcontributio n of FIH-+EH4.@ Fractional contribution of FIH" and EH4calculate da sdescribe d under Materials and Methods assuming that nofurthe r reductionoccurred . Final designates that changes inth e spectrum were complete after the amount of lip(SH)2 added. Eox and EH2o f enzymes Tyr16->Phe, Tyr16->Ser andA 9 are isosbestic at 445.4,445.4 and444. 7 nm respectively.

88 enzymes Tyr16->Ser andA 9 iscalculate d (Table 2). For allthre e mutated enzymes the calculatedtota l fraction of electrons after equilibration is exactly the same as inth e initial spectra. Fromthi s it isconclude dtha t intramolecular electron transfer fromth e dithiol to the FAD isdominan t inthes e 2-electron reduced mutated enzymes. For E. co//-enzyme the existence of the FLH- species wasdemonstrate d [14], although this species was only present in smallquantities .Th e occurrence oftranslocatio n suggests that the redox potentials of the disulfide andth e FAD are relatively close, however the slow rate at whichfi e electron transfer occurs suggeststha t this feature is relatedt o slow structural changes

Conformational stability. Recent conformational stability studies on wild-type lipoamlde dehydrogenase from A. vinelandiihav e shown that over-reduction of the enzymi to EH4 by NADH or dithionite promotes subunit dissociation thereby strongly mcrea ngth e sensitivity towards limited proteolysis,thermoinactivatio n and unfolding agents [11]. Exdept for enzyme A5,th etherma l stability ofth e mutated enzymes is decreased with renpectt o wild-type enzyme.Tabl e 3show stha t the melting temperatures (tm) of bothTyr1 6 mutated enzymes and enzyme A9 are clearly lower than the corresponding value afth e wildtyp e enzyme. For enzyme A14,thermostabilit y is strongly decreased yieldinga t m value which is only slightly higher as found for the monomeric wild type

Enzyme 4n Cm Gdn/HCI

°C M

wild-type 1) 80 2.45 A5 80 2.40 Tyr16->Phe 72 2.15 Tyr16->Ser 72 2.10 A9 65 2.05 A14 45 0.85 apoenzyme 1) 40 0.75

Tabl e3 . Conformational stability of mutated lipoamidedehydrogenase fromA .vinelandii . The meltingtemperature s (tm) and midpoint concentrations of unfolding (Cm Gdn/HCI) weni determined asdescribe di n ref. 11. (see also Fig.6) . 1)fijomref. 11.

89 350 400 450 500 550 1.0 1.5 2.0 Wavelength (nm) [Gdn/HCI] (M)

Fig. 6. Tryptophan fluorescence emission spectra ofTyr16->Ser lipoamide dehydrogenasefrom A .vinelandi iin Gdn/HCI. 2 uMenzym ewa s incubatedfo r3 0 rnini n10 0 mMK2HPO4 ,p H7. 0 andvariou s concentrations ofGdn/HC Ia t2 5 °C.Th e excitationwavelengt hwa s 295 nm.T oth e relativefluorescenc e ofth e nativeenzym e at52 4 nm,a valu e of 1.0i sassigned . (A) Fluorescence spectrumi nth eabsenc e ( )an di nth e presenceo f 1.6(1) ,2. 0 (2), 2.2 (3),2. 4 (4) and2. 8 MGdn/HC I(5) , respectively ( ). (B) Relativefluorescenc e at35 5 (0) and52 4 (•) nma sa functio no fth e unfoldingagent . apoenzyme [11].The high flavin fluorescence of A. vinelandiilipoamid e dehydrogenase is an intrinsic property ofth e dimeric enzyme [11]. Except for enzyme A14,thi s property isconserve d inth e mutated enzymes. For enzyme A14,flavi n fluorescence is strongly decreased andth e flavin ring is more accessible to solvent [16]. Lipoamide dehydrogenase from A. vinelandiicontain s a single tryptophan residue (Trp199) [17]. Both Trp199 andflavi n can be used as internal probest o study the unfolding of the (mutated) enzyme by guanidinium hydrochloride (Gdn/HCI) [11]. As an example,th e unfolding of enzyme Tyr16->Ser as afunctio n of Gdn/HCI concentration is recorded in Fig.6 . Fromth e midpoint concentrations of Gdn/HCI requiredfo r unfolding (Cm) it isclea r that the conformational stability of the mutated enzymesi s decreasedwhe n comparedt o wildtyp e enzyme (Table 3). Enzyme A14deviate s most showing a Cm value only slightly highertha n that reported for the monomeric wild type apoenzyme [11]. The dissociation constant for wildtyp e dimeric enzyme is below 1n M[11] . For enzymesA 9an dA14 , Kdvalue so f 5 nMan d2. 5u. M respectively were reported[8] . Therefore it is concludedtha t the decreased conformational stability of the mutated enzymes is a consequence of deteriorated subunit interaction.

Discussion

Recently the crystal structures of lipoamide dehydrogenase fromPseudomonas fluorescens[Mattevi , personal communication] and lipoamide dehydrogenase from the

90 N-terminal sequence

A. vinelandii SQKFDVIVIGAGPGGYVAAIKSAQLGLKTA (30) P. fluoresceins ******V*************RA******** (30) P.putida QQTIQTTLLI**A*********RAG***IP*E (32)

C-terminal sequence

A. vinelandii (447) VFAHPALSEALHEAALAVSGHAIHVANRKK- (476) P. fluorescens (447) **S**T************N*****I*****R (477) P. putida (434) IH***T*G**VQ****RAL***L*I (458)

Fig.7 . Sequencealignment ofthe N- and C- terminiof lipoamide dehydrogenase from Azdtobacter vinelandii [17], Pseudomonas fluorescens [18] and Pseudomonas putida W-val[19]. An asterisk(* )indicate sa residu eidentica lt o that in A. vinelandiilipoamid e dehydrogenase. branchJBd-chain oxo acid dehydrogenase complex from Pseudomonasputida with NAD+ioun d [5] were elucidatedt o 0.28 nman d 0.245 nm resolution respectively.I n bothstructure sth e C-terminal part folds back intoth e protein andi si ncontac t withth e N-terminal part ofth e other subunit (Fig. 1).Tyr18' ,th e equivalent ofTyr16 ' ofth e A. vinelandiienzyme , isfairl y well positionedt o form ahydroge n bond with ahistidin eo f the C-terminal parto fth e other subunit [4]. In view ofth e impressive sequence homology of the tpree lipoamide dehydrogenases, especially inth e N- and C-terminal region (see Fig. 7),jand the overlapping results obtained with the Tyr16 mutated enzymes and enzymi A9,i ti sconclude d that also inth e native A. vinelandiiwild-typ e enzyme the C-termfriusfold s back intoth e cavity,visibl e inth e structure. Recent further refinement of the A. vinelandiistructur e showstha t indeed density ispresen t that could account for the back folded C-terminus [Mattevi,persona l communication]. The properties of the mutated enzymes strongly indicate that the interaction between Tyr16' and His470 stabilizes the subunit interaction which iso futmos t importance for optimal catalysis, prevenfng the enzyme from inhibition/over-reduction byNADH .Th e question arises whether the observed differences insusceptibilit y toward inhibition/over-reduction between the E. coli, A. vinelandiian d pig heart wild-type enzymesi slinke dt o the dimer stability. Sinqeth e residues involved in the subunit interaction are relatively far away from the active ^ite,i ti spuzzlin g how the dimer interaction modulates the redox properties ofth e active 3ite. Iti sno wtemptin g to speculate that the observed diminished subunit interad|on leads to an increased redox potential of the FAD mediated viath e conformation of the C-terminus comprising residues 450-470.Th e isoalloxazine ofth e

91 FADi s located atth e interface ofth etw o subunits. Only onecontac t withth e isoalloxazirie,t o N-3, is provided byth e subunit whose C-terminus flanksth e active site: the backbone carbonyl oxygen of His450 [4] (His437 in Fig. 1). This carbonyl oxygen is positioned byth e c/s-proline451. Disturbance of this contact leadst o altered redox properties of theflavi n asshow n bystudie s of enzyme Pro451->Ala [9,10]. The strong over-reduction/inhibition of the Glu455 mutated enzymes [10] may occur in asimila r way since these enzymes also show decreased melting temperatures (67-70°C).

Acknowledgements

Wethan k Dr.W . Hoian d Mr.A . Mattevifo r preparing Fig. 1an dfo r making data available to us priort o publication.W eals o thank Mss. E. Hissink and M.d e Kortfo r technical assistance. This work was supported by the 'Dutch Foundation for Chemical Research' (SON) with financial aid from the 'Netherlands Organization for Scientific Research' (NWO).

References

1. Williams, C. H.Jr . (1991) in Chemistryand Biochemistry ofFlavoenzymes (Muller, F,. ed) vol3 ,i n press,CR C Press Inc.,Boc aRaton . 2. Reed,L J. (1974) Ace.Chem. Res. 7, 40-46. 3. Schierbeek,A .J. ,Swarte ,M . B.A. , Dijkstra,B .W. ,Vriend ,G. , Read,R .J. ,Hoi ,W .G . J., Drenth,J .an d Betzel,C . (1989) J. Mol. Biol. 206,365-379 . 4. Mattevi,A. , Schierbeek,A .J .an d Hoi,W . G.J . (1991) J.Mol. Biol. 220,975-994 . 5. Mattevi,A ,Obmolova ,G ,Sokatch ,J . R., Betzel.C.an d Hoi,W.J.G .(1991 )submitted . 6. Greer, S. and Perham, R. N. (1986) Biochemistry25, 2736-2742 . 7. Krauth-Siegel, R. L, Enders, B., Henderson,G . B.,Fairlamb ,A . H.an d Schirmer, R. H. (1987) Eur.J. Biochem. 164, 123-128. 8. Schulze, E. Benen,J .A . E.,Westphal ,A . H.an d De Kok,A . (1991) Eur.J. Biochem. 200, 29-34. 9. Benen,J. ,Va n Berkel,W. ,Zak ,Z , Visser,T ,Veeger , C.an d De Kok,A . (1991)Eur. J. Biochem.202, 863-872 . 10. Benen,J. ,Va n Berkel,W. , Dieteren,N. ,Arscott , D.,Williams ,C .Jr. ,Veeger , C.an d de Kok,A . (1992)submitte dt o Eur.J. Biochem. 11. Berkelvan ,W .J . H., Regelink,A .G. , Beintema,J .J .an d De Kok,A . (1991) Eur. J. Biochem202, 1049-1055.

92 12. Mussey,V . and Veeger,C . (1961) Biochim.Biophys. Acta 48,33-47 . 13. Vaeger,C and Massey, V. (1963) Biochim.Biophys. Acta 67,679-681 . 14. Wilkinson, K. D. andWilliam s C. H.Jr . (1979) J. Biol.Chem. 254,852-862 . 15. Wilkinson, K. D. and Williams C. H.Jr . (1981) J. Biol.Chem. 256, 2307-2314. 16. Benen,J .A . E., Van Berkel,W .J . H.an d De Kok,A . (1991) in Flavins and Fluvoproteins1990 (Curti, B., Ronchi, S. andZanetti ,G .eds. ) pp 557-564, Walter dejGruyte r and Co. Berlin-New York. 17. W^stphal,A . H.an d De Kok, A. (1988) Eur.J. Biochem. 172, 299-305. 18. B^nen,J .A . E.,Va n Berkel,W .J .H. ,Va n Dongen,W . M.A . M.,Miiller ,F. , andD e Kdk,A . (1989) J. Gen.Microbiol. 135, 1787-1797. 19. Bi^rns,G. , Brown,T. , Hatter, K. and Sokatch,J . R. (1989) Eur.J. Biochem. 179, 61- 6*

93 Journplof General Microbiology (1989),135 ,1787-1797 . Printed in Great Britain 1787

Chapter 5

Molecular Ooning and Sequence Determination of the Ipd Gene Encoding Lipoamide Dehydrogenase from Pseudomonas fluoresceins

By JACQUES A. E. BENEN,1 WILLEM J. H. VAN BERKEL,1 WALTER M. A. M. VAN DONGEN,1 FRANZ MULLER2 AND ARIE DE KOK1* 1 Department of Biochemistry, Agricultural University of Wageningen, Dreijenlaan 3, NL-6703HA Wageningen, The Netherlands 2 SandozAG, Agro Division, Toxicological Section, CH-4002 Basle, Switzerland

{Received 9December 1988;revised 17March 1989; accepted 5 April 1989)

The Ipd gene encoding lipoamide dehydrogenase (dihydrolipoamide dehydrogenase; EC 1.8.1.4) was isolated from alibrar yo fPseudomonasfluorescens D NA clone d in Escherichia coliTG 2 byus eo f serum raised against lipoamide dehydrogenase from Azotobacter vinelandii. Large amounts (up to 15% of total cellular protein) of the P. fluorescens lipoamide dehjdrogenas e were produced by the E. coli clone harbouring plasmid pCJB94 with the lipoimide dehydrogenase gene. The enzyme was purified to homogeneity by a three-step procedure. The gene was subcloned from plasmid pCJB94 and the complete nucleotide sequence of the subcloned fragment (3610bp ) was determined. The derived amino acid sequence of P.fluorescens lipoamide dehydrogenase showed 84%an d 42% homology when compared to the amino acid sequences of lipoamide dehydrogenase from A. vinelandii and E.a li,respectively . TheIpd gen eof P. fluorescens is clustere d inth egenom e with genesfo rth e othe• components of the 2-oxoglutarate dehydrogenase complex.

INTRODUCTION Lipoamide dehydrogenase (dihydrolipoamide dehydrogenase; EC 1.8.1.4) is the flavo- protii n component (E3)o f the multienzyme complexes that catalyse theoxidativ e decarboxyl- atioii of pyruvate (pyruvate dehydrogenase complex; PDC), 2-oxoglutarate (2-oxoglutarate dehjdrogenas e complex; OGDC) and branched-chain oxoacids (branched chain oxoacid dehjdrogenase complex; BCOADC) to acyl-CoA (Williams, 1976; Sokatch et al, 1981). Lipaamid edehydrogenas ecatalyse sth ereoxidatio n ofreduce d lipoylgroup stha t are covalently bouid to the dihydrolipoamide acyltransferase components (E2)o f the complexes. Lipoamide dehydrogenase belongs to the family of FAD-containing pyridine nucleotide oxidireductases . Other members of this family are glutathione reductase (Williams, 1976), menuri c reductase (Fox & Walsh, 1982), thioredoxin reductase (Holmgren, 1980) and trypiinothion e reductase (Shames et al., 1986). Common characteristics of these enzymes are (1)t lle yar eal ldimeri c and(2 ) they allcontai n oneredox-activ e disulphide bridge per subunit which participates in catalysis. Fiom several organisms genes encoding lipoamide dehydrogenase have been cloned and sequenced: from Escherichia coli (Stephens et al., 1983) and recently from man (small-cell carcinoma) and pig (adrenal medulla) (Otulakowski & Robinson, 1987), from bakers' yeast (Brovnin get al., 1988;Ros set al., 1988 )an d from Azotobactervinelandii (Westpha l &d eKok , 19891.

Abireviations: PDC,pyruvat e dehydrogenase complex; OGDC, 2-oxoglutarate dehydrogenase complex; BCCMJDC, branched-chainoxoaci ddehydrogenas ecomplex ;ORF , openreadin gframe .

0001-J286 © 1989 SGM 95 1788 J. A. E. BENEN AND OTHERS

InE. coliand A. vinelandiionl yon ecop yo f theIpd gen e ispresent . InE. colith egen e belongs to theace opero n togetherwit hth egene s encoding pyruvatedehydrogenas e (EC 1.2.4.1) (E1)an d dihydrolipoamide acetyltransferase (EC 2.3.1.12) (E2) of the PDC (Langley & Guest, 1977; Guest et al, 1981); in A. vinelandii, on the other hand, the Ipd gene was found to be located downstream of the gene encoding dihydrolipoamide succinyltransferase (EC 2.3.1.61) (E2) of the OGDC (Westphal & de K.ok, 1988; Hanemaaijer et al, 1988). In both organisms the Ipd genes can be transcribed independently from the genes for the other components in the respective clusters by use of separate promoters. In Saccharomyces cerevisiae (Dickinson et al., 1986) and in man, PDC and OGDC contain identical lipoamide dehydrogenases. In man this lipoamide dehydrogenase is also found in BCOADC (Otulakowski et al., 1988). In Pseudomonas putida and P. aeruginosa, on the other hand, two different lipoamide dehydrogenaseshav ebee nfoun d (Sokatchera/. , 1981;McCull yetal., 1986)an dthei r respective genes have been mapped in P. putida (Sykes et al, 1985). One lipoamide dehydrogenase (lpd-val), encoded by Ipdv, is part of the BCOADC; the second (lpd-glc), encoded by Ipdg, is parto f the OGDC, of the glycine oxidation system (Sokatch &Burns , 1984) and most likely of thePDC . TheIpdv gen e ispar to f acluste rcomprisin g thegene s encoding theBCOAD C and the Ipdg gene is part of the cluster encoding the OGDC. Sequence homology between carboxy-terminal peptides containing the active-site histidine residue of lpd-val from P. aeruginosa and P. putida and lipoamide dehydrogenase from E. coli has already been demonstrated by McCully et al. (1986) and extends to other lipoamide dehydrogenases. Therefore, adetaile d structure-function analysiso flipoamid e dehydrogenases, some of them cooperating in different complexes and others being specific for one complex, is very interesting. Ofgrea t value to this isth e determination of the three-dimensional structure of A. vinelandii lipoamide dehydrogenase, which is in progress (Schierbeek et al, 1989). Here we report on the molecular cloning and sequence determination of the gene encoding lipoamide dehydrogenase of the OGDC of P.fluorescens. Some general properties of the highly expressed, purified enzyme are described as well.

METHODS Bacterial strains, vectors and growth conditions. E. coli TG2 (Gibson, 1984), a recA version of TGI [Mfac-pro) thisupE [Res' Mod"(k) ]F'(traD36 proA+B* lacI*ZAM15)] was used throughout as a host for cloning andgrow n in YT medium. The P.fluorescens strain used in this study was that described by Howell elal. (1972)

and was grown in YT-medium or in minimal medium containing 200 mM-K2HP04,20-0 mM-NH^Cl, 800 mM- 1 NH4NOj, 85uM-MgSO. , 15-5mM-p-hydroxybenzoat e and 01 mg FeCl31" , pH 7-0. Plasmids pUC9 (Vieira & Messing, 1982)1pUC1 8an dpUC1 9(Yanisch-Perrone ral., 198S)an dpTZ18 R (Pharmacia)wer euse dfo rcloning ; M13mp9an d M13mpl8 wereuse d fornucleotid esequencin g(Norrande retal., 1983;Yanisch-Perro nera/. , 1985). Materials. Restriction endonucleases were obtained from Anglian Biotechnology, Boehringer or Bethesda Research Laboratories (BRL). T4 DNA ligase and E. coliDN A polymerase I(Kleno w fragment) were obtained from BRL. Universal sequencing primer, 7-deaza-dGTP and calf intestinal phosphatase were from Boehringer. Low- and high-gelling-temperature agarose were from Seakem. [a-32P]dATP (3000Cimmol"'; UITBq mmol"1) was purchased from New England Nuclear (NEN). Goat anti-rabbit IgG conjugated to alkaline phosphatase was from Promega Biotec. Sera against purified lipoamide dehydrogenase from A. vinelandii and E.coli wer ekindl yprovide db yM rA . H. Westphal(Westpha l& d eKok , 1988).Chromatograph y resinswer e from Pharmacia. All chemicals used were of analytical grade. Isolationofchromosomal and plasmid DNA. P.fluorescenswa sgrow nat 3 0° Ci n 1 litreo fminima l mediumt oa n ODMOo f 1-5-2-0.Cell swer eharveste d bycentrifugation (10min , 6000;)an dwashe donc e with 100m l20 %(w/v ) sucrose, 150mM-NaCl , 100mM-EDTA , pH 7-6(SSE) . The pellet was resuspended in 20m l SSE and lysozyme (10 mg) was added, followed by incubation at 37° C for 1 h. Next, 5 mg proteinase K was added, followed by another incubation at 37° C for 1h . The lysate was extracted five times with phenol, phenol/chlorofonn/isoamyl alcohol (25:24:1, by vol.)an d repeatedly extracted with chloroform/isoamyl alcohol (24:1, v/v) (Maniatis et al., 1982),unti l uponcentrifugatio n no pellicle was formed at the interface. Finally, the DNA was precipitated with ethanol,dissolve d in 10mic-Tris/HCl ,p H 8-0,0-1 mM-EDTA andstore dat 4 ° Cwit ha dro po fchlorofor m added. Large-scaleplasmi d DNA isolations were performed asdescribe d by Westphal &d e Kok (1988) and small-scale isolations as described by Bimboim & Doly (1979).

96 Nucleotide sequence of P. fluorescens Ipd gene 1789 I 1 W * - 5 - £ & £< o £ £ I 3?H I lt° I9Q ; sucA sucB ; /prf /jriA" -2-7 kb — i—70 kb pUC9 i

<*

SoulAI f =• AM | t= Well | £»R1 | 9 E=

Fig. 1. (a) Physical map of plasmid pCJB94 with the genes for 2-oxoglutarate dehydrogenase (sucA), dihydrolipoamide succinyltransferase (sucB),lipoamid e dehydrogenase (Ipd)an d an ORF encoding a polypeptide with yet unknown function (prtX). (b)Th e insert of plasmid pJB201 and the strategy for determination of the nucleotide sequence of subcloned fragments of pJB201 in M13 vectors. The arrowheads show the direction and extent to which Sou3AI, Alul, HineU and EcoRl fragments of pJB201 were sequenced. Numbers in panel (a) and at the top of panel (b) indicate the length of the fragments (in kb).

Coistruction and screening of a P. fluorescens gene library in E. coli. Construction of aP. fluorescens gen elibrar y in E.ca i and screening for clones that produce P.fluorescens lipoamid e dehydrogenase was performed essentially accoidin gt o the procedure used for the isolation of the A. vinelandiiIpd gen e asdescribe d by Westphal &d e Kok (1988) with the following slight modifications. Instead of 9-23 kb partial Sau3AI fragments, 7-15 kb SaulAl fragments of genomic P.fluorescens DN A were ligated into BomHI-digested pUC9 and instead of '35 I-labeIled protai nA , goatanti-rabbi t IgG conjugated toalkalin e phosphatase wasuse d fordetectio no f boundantibodies . As a source of primary antibodies, serum raised against A. vinelandii lipoamide dehydrogenase was used. Pllsmid pCJB94 was isolated from a clone that appeared positive upon initial screening and which produced high amounts of lipoamide dehydrogenase. The Ipdgen e was subcloned from this plasmid. Therefore, plasmid pCJB94 was partially digested with restriction enzyme Sa«3AI, the digested DNA was fractionated on a 1-2% (w/vllow-gelling-temperature agarose gel and fragments of 30-40 kb were isolated and'ligated into the BamHl site af pTZ18R. After transformation of E. coliTG 2 cells with the recombinant plasmids, the same screening procedure was applied as described above. Out of several positive clones E. coli TG2(pJB201) was chosen for sequence analysis of the cloned gene. Nucleotide sequencedetermination and analysis. The insert of plasmid pJB201 was isolated and digested with sevefil restriction enzymes. The resulting fragments were cloned in M13-derived vectors for sequence deterninatio n with the dideoxy chain-termination method of Sanger et al. (1977) according to the strategy displilye di n Fig. 1. Because of the high G + Ccontent , 7-deaza-dGTP was used instead of dGTP. Sequence data were compiled and analysed with a VAX computer using the programs of Staden (1982, 1984). Wistern blotting. This was done as described by Westphal &d e Kok (1988) with the same modification, with respa; t to detection of bound antibodies, as mentioned above. Sotahem blotting. Digestso f P.fluorescens DN A and plasmids pCJB94 and pJB201 were fractionated ina 0-6 % (w/v) agarose gel in 89mM-Tris/borate , 2mM-EDTA , pH 8-3, followed by transfer of the DNA to nitrocellulose accoi iingt oSouther n (1975). Isolated insert of pJB201 was nick-translated using [32P]dATP anduse d asa probe (Rig!y et al., 1977). Hybridization wascarrie d out at6 5° Cfo r 16h i n6 x SSC,5 x Denhardt'ssolutio n and0- 1% (w/v)SD S(Maniati set al., 1982).Th eblo twa swashe d at6 5 °Ci nbuffer scontainin g0- 1% (w/v )SD San d6 x SSC, 1x SSC and 0-1 x SSC, respectively, each wash lasting 45 min. Emyme purification.P. fluorescens lipoamid e dehydrogenase was isolated from £. coliTG2(pCJB94) , applying esseDiall y the same procedure as used for the isolation of lipoamide dehydrogenase from A. vinelandiiclone d in E.cot !TG 2 (Westphal &d e Kok, 1988). Briefly, cell-free extract wastreate d with protamine sulphate to remove nuclak acids and large complexes. After centrifugation, the supernatant was made 50% saturated with

97 1790 J. A. E. BENEN AND OTHERS ammonium sulphate, clarified by centrifugation and applied to a Sepharose 6B column Tor hydrophobic interaction chromatography. The enzyme, firmly bound at the top of the matrix, was eluted with 45%saturate d ammonium sulphate in SOmM-potassium phosphate, pH 70, OSmM-EDTA . FPLC analytical gel nitration was done as described by Van Berkel el al. (1988). Lipoamide dehydrogenase activity was assayed as described by Van den Broek (1971) using 1 M-potassium phosphate, pH 7-0, instead of 0-8M-sodiu m citrate, pH 6-5, as the buffer.

RESULTS AND DISCUSSION Isolation of E. coli clones thatproduce P.fluorescens lipoamide dehydrogenase Genomic P.fluorescens DN A was partially digested with restriction endonuclease SauiAI. Fragmentso f7-1 5k bwer eisolate d and ligated intoth eBamHl siteo fpUC9 .Th e recombinant plasmids were used to transform E. coliTG2 . Of the resulting clones, 1900wer e screened with antiserum raised against lipoamide dehydrogenase from A. vinelandii.On e clone, E. coli TG2(pCJB94),reacte d strongly with the antiserum:thi sclon e had a bright yellow appearance, suggesting that a large amount of a flavoprotein was produced. Cell-free extract of E.coli TG2(pCJB94)showe d a lipoamide dehydrogenase activity which was 30-fold higher than that ofcell-fre e extracto fE. co/i"TG2(pUC9) .SOS-polyacrylamidCge lelectrophoresi s (SDS-PAGE) ofthi scell-fre e extractreveale dlarg eamount so fa polypeptid ewit ha napparen t molecularmas s of 56kD a which was not detectable in E. coliTG2(pUC9 ) (results not shown). The P.fluorescens lipoamide dehydrogenase produced by E. coliTG2(pCJB94 ) was purified from cell-free extract of this clone following essentially the same procedure as described by Westphal& d eKo k(1988 )fo rpurificatio n oflipoamid edehydrogenas efro mA. vinelandiicloned in E.coli (Tabl e 1).Th e P.fluorescenslipoamid e dehydrogenase waselute d from the Sepharose 6Bcolum n at 45% ammonium sulphate saturation instead of 25% asfoun d for theA. vinelandii enzyme. This indicates that P.fluorescens lipoamid e dehydrogenase is less hydrophobic than A. vinelandii lipoamide dehydrogenase. SDS-PAGE of the purified protein showed a polypeptide witha napparen t molecularmas so f5 6kD atha twa svirtuall yfre e ofcontaminatin g polypeptides (Fig. 2a, lane 2). The mobility of this purified polypeptide on SDS-PAGE is different from E. colilipoamid e dehydrogenase, which has an apparent molecular mass of 50kD a (Fig. 2a, lane 1). The subunit composition of the purified enzyme wasestimate d byge lfiltration. Th e enzyme eluteda son esymmetrica l peak from aSuperos e 12column .Th e apparent molecularmas so fth e protein as estimated from the distribution coefficient was 110kDa , indicating that, like other lipoamide dehydrogenases, the enzyme is a dimer in its native state. The molecular mass as determined here isi n agreement withth e reported molecular masso f lipoamide dehydrogenase holoenzyme purified from P.fluorescens b y Scouten & McManus (1971). The amino acid sequence of the 22 amino-terminal residues of the purified protein was determined by automated Edman degradation. This sequence was different from the N-terminal sequence of E. coli lipoamide dehydrogenase, but was found to coincide with the sequence derived from the nucleotide sequence of the gene found in the cloned P. fluorescens fragment (vide infra),omittin g the initiating formylmethionine. This demonstrates that the purified lipoamide dehydrogenase was synthesized under the direction of the cloned P.fluorescens DN A fragment. Western blots of purified lipoamide dehydrogenases from E. coli, A. vinelandiian d P.fluorescens [obtaine d from E. coli TG2(pCJB94)] were incubated with antiserum raised againstlipoamid edehydrogenase s from E.coli or A. vinelandii(Fig .2b, c). ClonedP.fluorescens lipoamidedehydrogenas ereacte dwit hantiseru m againstA. vinelandiilipoamid edehydrogenas e (Fig.2 c )but not with serum against the E.coli enzym e (Fig.2b). I t can alsob econclude d from Fig. 2 that purified P. fluorescens lipoamide dehydrogenase as isolated from E. coli TG2(pCJB94) was not detectably contaminated with endogenous E. coli lipoamide dehydrogenase. A purification table of a large-scale isolation of the cloned P. fluorescens lipoamide dehydrogenase is presented in Table 1. From this table it can be estimated that the cloned enzyme accounts for approximately 15%o f the total cellular protein of E. coliTG2(pCJB94) .

98 Nucleotide sequence of P. fluoresce™ Ipd gene 1791

(fl) (A) (c) M 1 2 3 12341234

kDa 940- ' 670- < rats*

43-0-1

300-\

201- *

14-4-%

Fig.2 . Electrophoreticprofile so fpurifie d lipoamidedehydrogenase si na 12-5 %(w/v )polyacrylamid e gel with 01%SDS . (a) Staining with Coomassie brilliant blue R; (ft,c) Wester n blots of purified lipoamide dehydrogenases incubated with antisera against E. colilipoamid e dehydrogenase (ft) orA. vinelandiilipoamid edehydrogenas e(c) .Lane s 1,E. coli lipoamid edehydrogenas epurifie d fromisolate d pyruvatedehydrogenas ecomplex ;lane s2 ,P. fluorescein lipoamid edehydrogenas epurifie d fromE. coli TG2(pCJB94);lane s3 ,purifie d A. vinelandiilipoamid edehydrogenase ;lane s4 ,sampl ebuffer ; laneM : molecularmas smarkers .Th eban da tapproximatel y 60kD ai nal lfou rline si n(c )result sfro m reaction ofantiseru magains tA. vinelandii'lipoamidedehydrogenas ewit hsom ecomponen tpresen ti nth esampl e buffer. The bands at 100 kDa and 40kD a in E. colilipoamid e dehydrogenase represent residual pyruvate dehydrogenase anda degradatio n product of the latter.

Table 1. Purification of P. fluorescens lipoamide dehydrogenase from E. coli TG2(pCJB94)

Specific Total olume Protein activity* activity* Yield Step (ml) (mg) (U mg"') (U) (%) Cel^free extract 860 5525 19-2 107500 100 Prolimnini e sulphate supernatant 900 3240 32-7 106200 98 50% ammonium sulphate supernatant 1240 936 88-7 83040 77 Seplaroos: e 6B (pooled fractions) 215 512 1271 65575 61 * 1U = 1 umol NADH oxidized min"1.

Usihg standard assay conditions, the specific activity of the purified enzyme (127U mg"1; 1U|= 1 umol NADH oxidized min"1) is comparable to the specific activity of cloned A. vinelandii lipoamide dehydrogenase.

Nucleotide sequence andanalysis of the P.fluorescens Ipd gene lieIpd gen efro m P.fluorescens wa ssubclone d from the 7k b insert of pCJB94.Fro m oneo f resultant clones, £. coli TG2(pJB201)'(Fig. 1), plasmid was isolated and the 3-6k b insert I for sequence analysis. hesequencin g strategy isshow n in Fig. 1;al lsequenc e data werederive d from gel-readings 1a tleas ttw oindependentl y sequenced fragments. Thefinal 361 0b psequenc ewa scompile d

99 1792 J. A. E. BENEN AND OTHERS Succinyltransferase RBS DPARLLLDV* GATCCCGCTCGCCrrGCTCCTGCACGTCTGATTTACCfiTTrCTCTTCACCGCGGCCCCCTCGTGGCGCCGTCCGGTTTCACCCGAATAAGO 10 20 30 40 SO 60 70 80 90 ^_ SQKFDVVV IGAGPGGYVAAI RAAQLGL ATTGAGATATGAGCCAGAAATTCGACGTGGTAGTGATTGGTGCCGGCCCCGGCGGTTACGTTGCCGCCATCCGCGCTGCCCAGCTGGGCC 100 110 120 130 140 150 160 170 180

KTACIEKYIGKEGKVALGGTCLNVGCIPSK TGAAGACCGCCTGCATCCAGAAGTACATCGGCAAGGAAGGCAAGGTGGCGCTCGGCGGTACCTGCCTGAACGTTGGCTGCATCCCGTCCA 190 200 210 220 230 240 2S0 260 270

ALLDSSYKYIIEAKEAFKV1IGIEAKGVTIDV AGGCGCTGCTGGACAGCTCCTACAAGTACCATGAGGCCAAAGAAGCCTTCAAACTCCATGGTATCGAAGCCAAGGGCGTCACCATCGACG 280 290 300 310 320 330 340 350 360

PAMVARKAN1VKNLTGGIATLFKANCVTSF TTCCGnCGATCGTTGCGCGCAAGGCCAACATCGTCAAGAACCTGACCGGCGGCATCGCTACCCTGTTCAAGGCCAACGGCGTGACCTCCT 370 380 390 400 410 420 430 440 450

EGIIGKLLANKQVEVTGLDGKTQVLEAENVI TCGAAGGCCACGCCAAGCTGCTGGCCAACAAGCAGGTGGAAGTGACCGGTCTGGACGCCAAGACCCAGGTCCTCGAAGCCGAGAACGTGA 460 470 480 490 500 510 520 530 540

IASGSRPVEIPPAPLTDDI IVQSTGALEFQ TCATCGCTTCCGCCTCCCGCCCGGTGGAGATCCCGCCGGCCCCGCTGACCGACGACATCATCGTCGACTCCACCGGCGCCCTGGAATTCC 550 560 570 580 590 600 610 620 630

AVPKKLGVIGAGVIGI-ELGSVWARLGAEVT AGGCCGTGCCGAAGAAGCTCGGTGTGATCGGCGCCGGCGTCATCGCCCTGGAGCTGGGTTCGGTATGGGCCCGCCTGGGTCCCGAGGTGA 640 650 660 670 680 690 700 710 720

VLEALDKFLPAADEQIAKEALKyLTKQGLN CCGTGCTGGAAGCCCTGCACAAGTTCCTCCCGGCTGCCGACGAGCAGATCGCCAAGGAAGCCCTGAAGGTCCTGACCAAGCAAGGCCTGA 730 740 750 760 770 7E0 790 BOO 810

1RLGARVTASEVKKKQVTVTFTDANGEQKE ACATTCGCCTGGCCGCCCGCGTCACCGCCTCGGAAGTGAAGAAGAAGCAGGTCACCGTGACCTTCACCGATGCCAACGGCGAGCAGAAGG 820 830 840 850 860 870 680 890 900

TFDKL I VAVGRRPVTTDLLAADSGVTLDER AAACCTTCGACAAGCTGATCGTCGCnnTCGGCCCTCGCCCGGTGACCACCGACCTGCTGGCCGCCGACAGCGGCGTGACCCTGGACGAGC 910 920 930 940 950 960 970 980 990

GFIVVODIICKTSVI' GVFAIGDVVRGAMLAH GTGGCTTCATCTACGTCCACGACCACTGCAAGACCAGCGTTCCGGGCGTCTTCCCCATCGGTGACCTGGTTCGCGGCGCGATGCTCGCCC 1000 1010 1020 1030 1040 1050 1060 1070 1080

KASEEGVHVAER IACHKAQMNYDLI PSV I Y ACAAGGCCTCGGAAGAGGGCGTGATGGTCGCCGAGCGCATCGCCGGCCACAAGGCGCACATGAACTACGACCTGATCCCGTCGGTCATCT 1090 1100 1110 1120 1130 1140 1150 1160 1170

T H P E 1 AHVGKTEQTtKAEGVEVNVGTFl'FA ACACCCACCCGGAAATCGCCTGGGTCGGCAAGACCGAGCAGACCCTCAAGGCCGAGCGCGTCGAAGTCAACGTCGGCACCTTCCCGTTCG 1180 1190 1200 1210 1220 1230 1240 1250 1260

ASGRAMAANUTTGLVKV IADAKTDRVLGVH CCGCCAGCGGTCGTGCGATGGCTGCCAACGACACCACCGGCCTGGTCAAGGTCATCGCCGATGCGAAGACCGACCGCGTCCTGGGCGTCC 1270 1280 1290 1300 1310 1320 1330 1340 1350

V IGPSAAELVQQGA 1GHEFGTSAEDLGHHV ACGTGATCGGCCCGAGCGCCGCCGAr.CTGGTACAGCAAGGCGCGATCGGCATGGAATTCr>GCACCAGCGCCCAAGATCTGGGCATGATGG 1360 1370 1380 1390 1400 1410 1420 1430 1440

FSHPTI. SEAL H E A A I. AVNGHA11IIANRKKR TCTTCTCCCACCCGACGCTGTCCGAAGCCCTGCACGAAGCTGCCCTGGCGGTGAATGGCCATGCCATCCACATCCCCAACCGCAAGAAGC 1450 1460 1470 1480 1490 1500 1510 1520 1530

GCTrJWrTGATCCGACGCCCCWnTCACGCCAGGGGCGTCACAGCGACGCGGGACGCGTCGTCCGTCAGGCWGGACAGGTTCCTGGCCTT 1540 1550 1560 1570 1580 1590 1600 1610 1620

GTCGGGTCCGGGTTCTGTAGAATCCACGGTTTGCCGCCTGCGTCGGGATGACGCAGGCGGCAGACTTTCAGAAGAACCACGCCGGGTGAG 1630 1640 1650 1660 1670 1680 1690 1700 1710

CGCCGGCGACCGTTGCGAOrrGGCCGC(MCGTATGCCGCCGGACTGTTTTTTCGCGAGCCGGGTGGTTCGCGAAGCGACGCTCACAGGTC 1720 1730 1740 1750 1760 1770 1780 1790 1800 Fig.3 . Nucleotidesequenc eo fth eP. fluoresce™ Ipd gen ewit hth ederive damin oaci dsequenc ei none - character code. Indicated are a putativeribosome-binding site , RBS(overlined )an dthre e possible terminators,A B an d C(arrows) .Asterisk srepresen tsto pcodons .Th einitiatin gformylmethionin ei s omitted.

100 Nucleotide sequence of P.fluoresce™ Ipd gene 1793 fnimoverlappin gsequenc efiles, 7 8% o fth edat a(9 3% o fth ecodin gregio no fth eIpd gene )bein g derive dfro m gel-readingsofJx>t hstrands .Th einser to fpJB20 1(361 0bp )wa sfoun d tocompris e twD ope nreadin g frames (ORFs),on ebetwee n bp9 9an d 1532an d thesecon d between bp 1863 and 2990. The first ORF (bp99-1532; Fig. 3) could be identified as coding for lipoamide deiydrogenase . Identification was based on (1) the extensive homology of the derived amino ac dsequenc eo fth epolypeptid eencode d bythi sgen ewit hlipoamid edehydrogenas efro m both A. vinelandii(Westpha l& d eKok , 1988)an dE. coli (Stephen set al., 1983 )showin g84 %an d42 % homology, respectively (Fig.4) ,an d (2)th e identity of the derived amino-terminal amino acid s©juenc e with that of the protein purified from E. coli TG2(pCJB94). Thederive d aminoaci d sequenceo fP. fluorescens lipoamid edehydrogenas e (Fig. 3)contain s 477 amino acid residues excluding formylmethionine. The calculated molecular mass is 50033D a(5083 0D awhe nFA D isincluded) .Apparently, themolecula r massa sdetermine d by SI'S-PAGE (56kDa ) is overestimated by 10%. A similar observation was reported for lipoamide dehydrogenase from A. vinelandii (Westphal & de Kok, 1988). Upstream of the Ipdgen e noconsensu s E. coli-type promoter sequence could be identified in pJB201. Expression of P. fluorescens lipoamide dehydrogenase from plasmid pJB201 is dependen to nth e lacZpromote r ofth eclonin gvector .Thi swa sconfirme d byclonin g the insert in inverteddirectio n towardsth elacZ promote ri npUC19 .Th e startcodo nAU G isprecede db y a ]xrtentia l ribosome-binding site (bp87-94) . The nucleotide sequence downstream of the Ipd gene contains three regions of dyad sylnmetry .Fre eenerg ycalculation s(Tinoc oet al., 1973 )fo rth emR NA transcript so fth eregion s of dyad symmetry suggest that they may form very stable stem-and-loop structures: region A, A(r = -109 kJ mol"1; region B,A G = - 84k J mol"1; and region C,A G= -148 kJ mol"1. The indicated stem-and-loop structures might serve as transcription terminators. Homology between lipoamide dehydrogenases The primary structures of lipoamide dehydrogenases from E. coli(Stephen s et al., 1983), A. vinelandii (Westphal &d e Kok, 1988) and P.fluorescens (thi s study) are aligned in Fig. 4. Inlication s about functional domains in lipoamidedehydrogenase s werefirst obtaine d by Rice et al.(1984) . They compared the primary structures of E. colilipoamid e dehydrogenase and huma n glutathione reductase andfitted th e primary structure of lipoamide dehydrogenase into thi:three-dimensiona l structureo fglutathion ereductase .Schierbee ket al. (1989)determine d the thiee-dimensiona l structure of lipoamide dehydrogenase from A. vinelandii by X-ray cv stallography. The overall folding of the polypeptide chain of A. vinelandiilipoamid e deiydrogenas e appears to be very similar to that of glutathione reductase, with the same four- damai n structure. In the primary structure several features appear to be conserved in the ensymes:th eADP-bindin g folds of thecofactor s FAD and NAD(P)+ (positions 5-35 and 182- 211), respectively, relative to P.fluorescens lipoamid e dehydrogenase), the region around the adive-sit e disulphide bridge (positions 44-60) and the region around the active-site histidine (positions448-455) .Thes econserve d regionsar e indicated in Fig.4 ;the y areals oconserve d in P.fluorescens lipoamid e dehydrogenase. Besides the primary structures of the bacterial lipoamide dehydrogenases shown in Fig. 4, sequences of eukaryotic lipoamide dehydrogenases have been published: those of porcine adrenal medulla and human small-cell carcinoma (Otulakowski & Robinson, 1987) and of bakers' yeast (Browning etal., 1988;Ros set al., 1988).Als o in the primary structures of these euIcaryoti clipoamid edehydrogenase s theabove-mentione d features areconserve d (notshown) . |Localization of Ipdgenes inthe genome: P.fluorescens, E. coliand A. vinelandiicompared In pCJB94 theIpd gen ei sprecede d byth egene sfo r theothe r twocomponent so fth e OGDC ig. 1). Immediately upstream of the /pi gene, the gene for dihydrolipoamide succinyltrans- (sucB)wa s detected; this gene has now been subcloned and sequenced and is highly nologous to the dihydrolipoamide succinyltransferase of A. vinelandii (A. H. Westphal, onal communication) and E. coli (Spencer et al., 1984). Also the gene for the third uponent ofth eOGDC , 2-oxoglutaratedehydrogenas e (sucA), couldb e identified (resultsno t

101 1794 J. A.E .BENE NAN D OTHERS

I 10 20 30 40 II 50 60

priip SQKFDWVrG AGPGCYVAAI RAAQI.Cf.KTA CtF.KYIGKF.G KVALCGTCLN VGCIPSKALI. Avllp ""f Ecllti t...y.... "1 ,v% IV«R«NT -"T" " 70 80 90 )00 no 120 Pfllp DSSYKYIIF.AK EAFKVHGIPA KGVTIOVPAK VARKANIVKN LTCGIATLFK ANGVTSFEGH •S**I,**"S T Avlip .....F...„ Ecllp HVA-*VI*EA K'LAE*"VF GEPKTMOKI RTW*EKVINQ *"""L*GHA* GRKVKWNGL

130 110 150 160 170 180

priip CKLLANKQVK VTCI.OCKTQV LF.AENVIIA S GSRPVEIPPA PLTDOIIVDS TGALEFQAVP • .***,•• Avllp *.*..n.K.. ••AA^SS"" DT ..K...... •VDQ*VI*» " ••••0""N"" Ecllp ••FTCANTL• •E"EN"KT-' INFDNA""A ••**IQL*FI •HE'PR'W* •D*""LKE"

IN ,90 200 210 220 230 240

Pflip KKLCVICACV IGLFXGSVWA RLGAEVTVLE ALDKFLPAAD EQIAKEALKV LTKQGLNIRL Avllp

Ecllp ••••MGT*YH A**SQIO*V* MP'QVI"** KO*V-VFT"R 1S"-KF"LH«

2S0 2G O 269 279 289 299

Pfllp CARVTASEVK KKQVTVTFTD ANGE-QKETF DKLIVAVGRR PVTTDLLAAD SGVTLDERGF

Avllp N«,.,»K.V. *F**-KSQA*

Ecllp ETK"*V"A* EDGIY'TMEG KKAPAEPQRY •AVL*M**V "NGKN«D*GK A«*EV*D«"

309 319 329 339 349 359 Priip IYVODIICKTSVPGVFAIGO V VRGAMLAHKA SEEGVMVAER IAGHKAQNNY DL1PSVIYTH

Avllp aaaaaya/vaa aaa.y*aaaa .*«••«•••• aaaaayaaaa aaaaaaaaaa aaaaftaaaaa

Ecllp I"*KQIR"N •«III"*,"I •GQP"",G VEI,*IIVA"V "•K'llYFDP KV'-'IA'-E

369 379 389 309 409 419 Pfllp PE1AWVGKTE QTLKAF.GVEV NVCTFRFTTS GRAKAANDTT GLVKVIAORK TORVLGVHVI

Avllp ••».G...«. *A******AI •••V"P"AA" •••••••••A •F««««««A» •••••*•••• Ecllp -.v.*..!,..KF.A*F.K*IS Y ETA"PWAA' *"M"SDCAI> *MT*t."F"KE SH^M'GAIV

429 439 449 IV 459 469 Pfllp GPSAAELVQQ GAIGHEFGTS AEOLGHMVFS HPTLSEVvLIIE AALAVNGKA1 IHANRKKR

Avllp ••••-••••• a*a/vaaaaaa aaaaaaaaJ*A «.A»aa*a*« aaaaasaaaa av>aaaa

Ecllp "TNGC'LGE IGUI'M'CO "••1ALTIIIA "•"•ll'SVGL "EVFE'SIT OLP'P'AKKK

Fig. 4. Comparison of the amino acid sequence of the lipoamide dehydrogenases of P. fluorescens (Pflip), A. vinelandii(Avlip ; Westphal & de Kok, 1988) and E. coli (Eclip; Stephens et al., 1983). Identical residuesar e indicated byasterisks . Boxed residues indicate the ADP-binding site of FAD (I), theredox-activ edisulphid e bridge (II),th e ADP-binding site of NAD* (III)an d the region around the active-site histidine (IV).

shown).I norde rt oinvestigat e whetheranothe rcop yo fth eIpd gen ei spresen ti nP. fluorescens, genomic P.fluorescens DN A was hybridized withth eclone dIpd gen e (Fig. 5).Onl yon e DNA fragmentwa s foundt o hybridizewit hth e clonedIpd gene i nBamHh, Pstl- an d/ftndlll-digeste d P.fluorescens DN A (Fig. 5,lane s 1,2 an d 3,respectively) . In accordance withth epresenc eo f two EcoRl sites in the insert of pJB201 (Fig. 2), three fragments were found to hybridize in £coRI-digestedgenomi cDN A(Fig .5 ,lan e4) . Fromthi sexperimen tw eobtaine d no indication of the presence of a second Ipd gene in the P.fluorescens genom e with sufficient sequence homology to the cloned gene to be detected under the hybridization conditions applied. This suggests that, if another Ipd gene is present, as inothe r pseudomonads (Sokatch et al., 1981; McCully et al., 1986), only limited sequence homology exists between those Ipd genes. Theseresult sindicat etha ti nP. fluorescens, a si nA. vinelandii, butcontrar y toE. coli, the Ipd gene forms part of the OGDC cluster. Surprisingly, the homology in genetic organization betweenP. fluorescens an d A. vinelandiials oextend sdownstrea mo fth e Ipdgene .Downstrea mo f

102 Nucleotide sequence of P. fluoresce™ Ipd gene 1795

12 3 4 5 6 kb

231—

9-4-' 6-6-

4-3- 4

* 2-3—

2-0—

0-56- 1 Fig.5 . Autoradiographo fa Souther nblo tcontainin gP.ftuorescens chromosoma l DNA digests(lan e1 , BamHl;lan e 2, Pstl;lan e 3, Hindlll;lan e4 , EcoKl)an d EcoRldigest so fpCJB9 4(lan e S) andpJB20 1 (lane 6) hybridized with the "P-labelled insert of pJB201.

the Ipd gene large ORFs are found in both organisms (position 1863-2990 in the sequence of pJll201, prtX in Fig. 1) which might encode polypeptides that are 95% homologous when compared to each other. This indicates that the organization of the OGDC clusters of A. iinelandii and P.ftuorescens isver ysimilar . Formal proof that, asm A. vinelandii, noIpd gen e is preten t in the PDC cluster of P.ftuorescens, has to await the analysis of the genes of the PDC clutter.

Me are grateful to Dr R. Amons for the determination of the N-terminal sequence with the gas-phase seqienato ran dt oM r E. Keukensan d Ms M. C.Snoe kfo rtechnica lassistance .W ethan k MsM . F.va nEij k for edit] ng thetypescrip t and Mr M. M.Bouwman sfo rdrawin gth efigures. Thi sinvestigatio n wassupporte d byth e Netllerland sFoundatio nfo rChemica lResearc h(SON) ,wit hfinancial ai dfro mth eNetherland sOrganizatio n for Scientific Research (NWO).

REFERENCES BR^BOIM, H. C. & DOLY, J. (1979).A rapi d alkaline (19SS). Nucleotide sequence for yeast dihydrolip- e] traction procedure for screening recombinant amide dehydrogenase. Proceedings of the National piismi dDNA .Nucleic Acids Research 7,1513—1523 . Academyof Sciences of the UnitedStates of America BROVNTNO, K. S., UHUNGER, D. J. & REED, L. J. 85, 1831-1834.

103 1796 J. A. E. BENEN AND OTHERS

DICKINSON, J. R., ROY, D. J.&. DAWES, J. W. (1986). A tic sequences for related enzymes and identification mutation affecting lipoamide dehydrogenase, pyru­ of potential upstreamcontro l sites.Journal of General vate dehydrogenase and 2-oxoglutarate dehydrogen­ Microbiology 134, 1131-1139. ase activities inSaccharomyces cerevisiae. Molecular SANGER, F., NICKLEN, S. & COULSON, A. R. (1977). and General Genetics 204, 103-107. DNA sequencing withchai n terminating inhibitors. Fox, B.S . & WALSH, C.T . (1982). Mercuric reductase - Proceedingsof theNational Academy of Sciencesof the purification and characterization of transposon- United States ofAmerica 74, 5463-5467. encoded flavoprotein containing an oxidation- SCHIERBEEK, A. J., SWARTE, B. M. A., DUKSTRA, reduction-active disulfide. Journal of Biological B. W., VRIEND, G„ READ, R. J., HOL, W. J. O., Chemistry 257, 2498-2503. DRENTH, J. & BETZEL, C. (1989). Structure of GIBSON, T.J . (1984). Studies on the Epstein-Bar virus lipoamide dehydrogenase from Azotobactervinelan­ genome.Ph D thesis. University of Cambridge, UK. dii determined bya combinatio n ofmolecula r and GUEST, J. R., COLE, S. T. & JEYASEELAN, K. (1981). isomorphous replacement techniques. Journalof Organization and expression of the pyruvate Molecular Biology 206, 365-379. dehydrogenase complex genes of Escherichia coli SCOUTEN, W. H. & MCMANUS, I.R . (1971). Microbial K12. Journal ofGeneral Microbiology 127, 65-79. lipoamide dehydrogenase - purification and some HANEMAAIJER, R., JANSSEN, A., DE KOK, A. & VEEGER, characteristics ofth e enzyme derived from selected C. (1988). The dihydrolipoyltransacetylase compo­ microorganisms. Biochimica et biophysica acta 227, nent of the pyruvate dehydrogenase complex of 248-263. Azotobacter vinelandii.European Journal of Biochem­ SHAMES,S . L., FAIRLAMB, A. H., CERAMI, A. & WALSH, istry 174, 593-599. C T. (1986). Purification and characterizationo f HOLMGREN, A. (1980). Pyridine nucleotide-disulfide trypanothione reductase from Chrithidia fascicular, oxidoreductases. Experientia, Supplement 36, 149- a newly discovered member ofdisulfid e containing 180. flavoprotein reductase. Biochemistry25 , 3519-3526. HOWELL, L. G., SPECTOR, T. & MASSEY, V. (1972). SOKATCH, J. R. & BURNS, G. (1984). Oxidation of Purification andpropertie s of p-hydroxybenzoate glycin by Pseudomonas putida requires a specific hydroxylase from Pseudomonasftuorescens. Journal lipoamide dehydrogenase. Archivesof Biochemistry of Biological Chemistry 247, 4340-4350. and Biophysics 228, 660-666. LANGLEY, P. & GUEST, J. R. (1977). Biochemical SOKATCH, J. R., MCCULLY, V., GEBROSKI, J. & genetics of the o-ketoacid dehydrogenase complexes SOKATCH, D. J. (1981). Isolation of a specific of Escherichia coli K.12: isolation and biochemical dehydrogenase for a branched chain ketoacid properties of deletion mutants. Journalof General dehydrogenase from Pseudomonasputida. Journal of Microbiology 99, 263-276. Bacteriology 148, 639-646. MANIATIS, T., FRITSCH, E. F. & SAMBROOK, J. (1982). SOUTHERN, E. M. (1975). Detection of specific se­ Molecular Cloning: a Laboratory Manual. Cold quences among DNA fragments separated byge l Spring Harbor, NY: Cold Spring Harbor electrophoresis. Journal of Molecular Biology98 , Laboratory. 503-517. MCCULLY, V., BURNS, G. & SOKATCH, J. R. (1986). SPENCER, M. E., DARLISON, M. G., STEPHENS, P. E., Resolution ofbranche d chain oxoacid dehydrogen­ DUCKENFIELD, J. K. & GUEST, J. R. (1984). ase complex of Pseudomonas aeruginosa PAO. Bio­ Nucleotide sequence ofth e sucB gene encoding the chemical Journal233 , 737-742. dihydrolipoyl succinyl transferase of Escherichia coli NORRANDER, J., K.EMPE, T. & MESSING, J. (1983). K.12 and homology with the corresponding acetyl Construction ofimprove d M13 vectors using oligo- transferase. European Journal of Biochemistry 141, deoxynucleotide-directed mutagenesis. Gene 26, 361-374. 101-106. STADEN, R. (1982). Automation of the computer OTULAKOWSKI, G. & ROBINSON, B. H. (1987). Isolation handlingo fgelreadin g dataproduce d byth eshotgu n and sequence determination of cDNA clones for method of DNA sequencing. NucleicAcids Research porcine and human lipoamide dehydrogenases. 10, 4731-4751. Journal of BiologicalChemistry 262, 17313-17318. STADEN, R. (1984). A computerprogra mt oente r DNA OTULAKOWSKI, G., ROBINSON,B .H . &WILLARD ,H . F. gelreading data into a computer. Nucleic Acids (1988). Gene for lipoamide dehydrogenase maps to Research 12, 499-503. human chromosome 7.Somatic Cell and Molecular STEPHENS, P. E., LEWIS, H. M., DARLISON, M. G. & Genetics 14, 411-414. GUEST, 1. R. (1983). Nucleotide sequence of the RICE, D. W., SCHULZ, G. E. & GUEST, J. R. (1984). lipoamide dehydrogenase gene of Escherichia coli Structural relationships between glutathione reduc­ K12. EuropeanJournal of Biochemistry 135,519-527. tasean dlipoamid edehydrogenase .Journal of Molec­ SYKES, P. J., MENARD, J., MCCULLY, V. & SOKATCH, ular Biology 174, 483-496. J. R. (1985). Conjugative mapping of pyruvate, RlGBY, P. W.J. ,DlECKMANN , M., RHODES,C . &BERG , 2-ketoglutarate and branched chain ketoacid P. (1977). Labeling deoxyribonucleic acid to high dehydrogenase genes in Pseudomonas putida mu­ specific activity in vitro by nick translation with tants. Journal ofBacteriology 162, 203-208. DNA polymerase I. Journal of Molecular Biology TINOCO, I., JR,BORER , P. N., DENGLER, B., LBVINE, 113, 237-251. M. D., UHLENBECK, O. C, CROTKERS, D. M. & Ross, J., RED, G.A .& DAWES , J.W .(1988) . The GRALLA, J. (1973). Improved estimation ofsecon ­ nucleotide sequence of the LPD1 gene encoding dary structure in ribonucleic acids. NatureNew lipoamide dehydrogenase in Saccharomyces cerevi­ Biology 246, 40-41. siae: comparison between eukaryotic and prokaryo- VAN BERKEL, W. J. H., VAN DEN BERG, W. A. M. &

104 Nucleotide sequence of P. fluorescens Ipd gene 1797

MOlXBR, F. (1988). Large scale preparation and WESTPHAL, A.H .& D E KOK, A. (1988). Lipoamide reconstitution of apo-flavoproteins with special dehydrogenase from Azotobacter vinelandii;molecu - reference to butyryl-CoA dehydrogenase from lar cloning, organization andsequenc e analysiso f Megasphaeraelsdenii: hydrophobi c interaction chro- the gene. EuropeanJournal of Biochemistry 172, 299- matography. European Journal ofBiochemistry 178, 305. 197-207. WILLIAMS, C. H., JR (1976). Flavin containing VJkNDE N BROEK, H. W. J. (1971). Pyridine nucleotide dehydrogenases. In The Enzymes, vol. 13, pp. 89- Iranshydrogenase. PhD thesis. Agricultural Univer- 173. Edited by P. D. Boyer. New York: Academic sity of Wageningen, The Netherlands. Press. VIEIRA,J . & MESSING, J. (1982).Th e pUC plasmids,a n YANISCH-PERRON, C, VIEIRA, J. & MESSING, J. (1985). M13mp7 derived system forinsertio n mutagenesis Improved M13phag e cloning vectors and host and sequencing with synthetic universal primers. strains: nucleotide sequences of the M13mpl8 and Gene 19, 259-268. pUC19 vectors. Gene 33, 103-119.

105 Chapter 6

Lipaamidedehydrogenas e from Pseudomonas fluorescens. Som4 spectral properties and conformational stability

Jacquos Benen, Willem van Berkel andAri e de Kok.

Summary

Lipoamide dehydrogenase from Pseudomonasfluorescens i s characterized with respectt o some spectral properties andwit h respect to the conformational stability of the several redox states. Thl visible spectrum of the oxidized enzyme ischaracteristi c of aflavi n in an apolar environment. Reduction of the enzyme by NADH shows that the reducing equivalents mainly:resid e onth e disulfide. Excess NADHdoe s notfull y reduceth e enzyme to the four-electron reduced level.Th e enzyme is not inhibited by the product NADH. Th^ oxidized enzyme isthermostabl e showing a melting temperature of 86 °C.Th e thermit stability drastically decreases upon reduction.Th e oxidized and lip(SH)2 reduced enzyme are insensitive to unfolding by urea. Unfolding of NADH-reduced enzymteb y urea is inhibited inth e presence of NAD+. It isconclude dtha t over-reduction resultsi n a decreased stability of the dimer.

Intreduction

Th* gene encoding lipoamide dehydrogenase from Pseudomonasfluorescens ha s recently been cloned andexpresse d in Escherichiacoli [1]. The enzyme shows 84 % amino acid sequence identity with lipoamide dehydrogenase from Azotobacter vinelandii. The crystal structures of three lipoamide dehydrogenases have been determined at present,fro m AYmelandiiat 0.22 nmresolutio n [2] andfro m P.putida [3 ] andfro m P. fluorescens [Mattevi, personal communication] at 0.25 and0.2 8 nm resolution respectively. The three structures appear to be almost identical. Inth e Pseudomonas structures the last ten amino acid residues, which were initially not visiblelinth e electron density map of A. vinelandiienzyme , appear to fold back into the proteinjstructur e that constitutes the active site.Apar t from the,fo r A. vinelandiienzyme ,

107 nine established intersubunit hydrogen bonds,thi s backfolded C-terminus makes additional contacts with the other subunit [3]. Site-specific mutagenesis experiments withth e A. vinelandiienzym e have showntha t the C-terminus adopts a similar conformation inthi s enzyme (Chapter 4) [4]. Incontras t to the highdegre e of amino acid and structural identity the A. vinelandiian d P. fluorescens enzyme differ strongly with respect to FAD induced dimerization of the apoenzyme [5] indicating differences inth e enzyme's structures which are not revealed atth e current level of resolution. Lipoamide dehydrogenases, as isolated from different sources, strongly differ with respect to inhibition byth e product NADH [6-8]. This inhibition is mainly due to over­ reductiont o thefour-electro n reduced state (EH4),a sfirs t demonstrated withth eE.coli enzyme [9]. For lipoamide dehydrogenase from A. vinelandiii t was shown that over­ reduction strongly promotes subunit dissociation andtha t dimerization isth e driving force for theconformationa l stability of the protein [10]. Interestingly, both product inhibition andconformationa l stability are affected by mutations located in the vicinity of the interface[4] . Preliminary experiments in our laboratory indicated that lipoamide dehydrogenase from P. fluorescensi s insensitive to product inhibition despite the conservation of amino acids atth e C-terminus. Inthi s Chapter some catalytic properties and conformational stability of lipoamide dehydrogenase from P. fluorescensar e reported.

Materials and Methods

Biochemicals and concentration determinations have been described in Chapters 2 and 3 [8, 10]. Lipoamide dehydrogenase from P. fluorescens, asclone d in E.coll, was isolated according to Chapter 5 [1].Thermoinactivatio n and unfolding experiments were performed essentially as described elsewhere [11]. Absorption spectra under anaerobic conditions were recorded at 25 °C on an Aminco DW-2000 spectrophotometer, essentially as reported earlier. Rapid reaction and steady-state kinetics were studied as reported previously [8] with one slight modification:fo r steady state kinetics stock solutions of dihydrolipoamide (lip(SH)2 were prepared in pure methanol.

Results

Spectral properties. Fig. 1 shows the visible absorption spectra of lipoamide dehydrogenase from P. fluorescensbefor e and after anaerobic reduction with one and two molar equivalents of NADH.A t pH8.0 ,th e flavin absorption spectrum of the

108 —1 n 0.5 1 /'••. -. 1' 0.4 _ \ " o § 0.3 _/ AW

§ 0.2 - / i < 0.1 --, / tv~~^ i 1 —i —i 400 500 600 700 Wavelength (nm)

Fig 1. Visibleabsorption spectra of Pseudomonas fluorescens lipoamide dehydrogenase. Spectrawer e recordeda t 25°C in 100m M Hepps, 0.5 mM EDTA, pH8.0,15 0 mMioni c strength, before ( :Eox ) and afterth e additiono f 1.0( )an d2. 0 molar equvalertt so f NADH( )an dafte rth e additiono f 4.0 mMNAD +(-•-•-• ) to 2 molar equivalentso f NADH reduced enzyme. Spectra shownwer e recorded when changes weie complete. oxidizejd enzyme is very similar to those reported for lipoamide dehydrogenases from other sjources. Addition of one equivalent of NADH yields a spectrum resembling that reportejdfo r the pig heart enzyme [12]. The relatively high absorbance at 530 nm and the lozj absorbance around 700 nmdemonstrat e that the reducing equivalents mainly resideio nth e disulfide,characteristi c forth e 2-electron reduced enzyme (EH2). Full reduction ofth e enzyme to the four-electron reduced state [EH4] can not be accomplished by NADH as also reported forth e pig heart enzyme [12]. This suggests that th« redox potentials of the EH2/EH4couple s of both enzymes are very alike and more negative than those reported for lipoamide dehydrogenase from A. vinelandiian d E.co/ / 10,13]. This is also indicated by kinetic studies:i nth e forward reaction,a t pH 8.0, no significant inhibition by NADH isobserved . Upon addition of a large excess of NAD+t o stoichiometric NADH-reduced enzyme, a speaes is formed exhibiting maximal charge-transfer absorbance at 650-700 nm(Fig . 1).A smilar species was reported for pig heart enzyme [12]. From work onA. vinelandii mutated lipoamide dehydrogenase [8] it is concluded that this is a species with NAD+ bound,th e flavin oxidized and with the disulfide reduced.

Conformational stability. Fig. 2 shows the thermostability of lipoamide dehydrogenase from P. fluorescensi nth e absence and presence of reducing agents. The oxidized enzyme is highly thermostable yielding a midpoint temperature of thermal

109 0.1-

0.2

CD a 0.3 _*: 0.4

TEMPERATURE (C )

Fig2 . Thermostability of Pseudomonas.fluorescen s lipoamide dehydrogenase in the absence and presence of reducing agents. The apparent first-order rateconstan t for enzyme inactivation is plotted as afunctio n of the incubationtemperature . 2.0 uM enzyme was incubated atdifferen t temperatures. For other details see Materials and Methods. Oxidized enzymes (o),i nth e presence of 1 mM lip(SH)2 (•), inth e presence of 1m Mdithionite (x).

denaturation (tm) of 86 °C. Thisvalu e is 6°C highertha ntha t reported forth e A. vinelandiienzym e [11]. Upon reduction with excess dithionite or NADH,thermostabilit y strongly decreases, yielding apparent fm values (44an d 47 °C, respectively) inth e same range as reported forth e monomericapoenzym e [5]. Reduction with excess lip(SH)2give s riset o aver y sharp break inth e meltingcurv e at 61 °C (Fig.2) .Thi s sharp break is also observed in similar studies with A. vinelandiienzym e [11] and ascribedt o over-reduction to the EH4stat e at elevated temperatures. The oxidized forms of lipoamide dehydrogenase from pig heart and A. vinelandiiar e highly resistant towards unfolding by urea [11, 14]. Unfolding of these enzymes by urea is induced upon reduction by NADH. Inactivation ofth e P. fluorescens enzyme in 7 Murea , pH7.0 ,follow s first-order kinetics. As can be concluded from Table 1, in urea, both oxidized and lip(SH)2- reduced enzyme are very slowly inactivated,wherea s rapid inactivation occurs when the enzyme is reduced with dithionite ort o some lesser extent, with excess NADH.Th e

110 2 k 6

UREA [M] UREA [M]

Fid. 3. Inactivation ofPseudomona sfluorescen s andAzotobacte r vinelandii lipoamide dehydrogenase by urea either in the absence orpresence of reducing agents 2.0 uM enzyme was incubated in 100m M K2HPO4,p H 7.0 andvariou s concentrations of uraaa t 25°C . The residual lipoamide activity after 30 minincubatio ni splotte d against the concentration urea. A, P. fluorescens lipoamide dehydrogenase. B, A. vinelandii lipoamide dehydrogenase. Oxidized enzymes (o),i nth e presence of 1 mM lip(SH)2 (x), in tha presence of 1 mMNAD H (•)

3 1 Reductant k'inact x 10 min-

none 4.1

lip(SH)2 5.9 NADH/NAD+ 7.6 NADH 150 dithionite 230

Tattle 1.Inactivation of Pseudomonasfluorescen s lipoamide dehydrogenase in urea. Th

111 I 1 i A ' I tin? / V A_ 0.2 / V B_ -z. /fy-sK\ < /I C\ \ 7 2 \\ \ - CQ —/ \ \ — OH ~^2 \ 1 _ 0.1- 1/° 10 u1-' - CQ < - i V i i v.. 400 480 560 640 720 400 480 560 640 720 WAVELENGTH(nm )

Fig. 4. Absorption spectra of anaerobically NADH-reduced Pseudomonas fluorescens lipoamide dehydrogenase in thepresence of urea. 20 u.Menzym e in 100 mMK2HPO4 , pH7.0 , inth e presence of 3 Mure awas anaerobically reducedwit h 1m M NADHa t 25 °C. Spectrawer e recorded attim e intervals as indicated below.Afte r 30 minincubation , airwas admittedan d spectrawer e recorded again.(A ) Eox (1); 1mi nafte r NADH reduction (2);2 0 minafte r NADH reduction (3);afte r air-inlet (4).(B ) Eox inth e presence of 1m MNAD +(1) ;2 0 minafte r NADH reduction (2);afte r air-inlet (3). rates of inactivation are somewhat lowertha nthos e reportedfo r A. vinelandiienzym e [11] Fig. 3A shows the conformational stability of oxidized and reduced enzyme species at various ureaconcentrations . The oxidized and lip(SH)2 reduced enzyme are relatively urea-insensitive (cf.Tabl e 1) while the activity of NADH reduced enzyme becomes affected above 4 Murea . The apparent midpoint concentration of urea inactivation (Cm urea) forthi s species is 5.5 Man d much higher than the value observed forth e A vinelandiienzym e (Cm urea= 1.5M , Fig.3B) . In urea,th e shape of the absorption spectrum of the oxidized enzyme remains virtually unchanged with agradua l shift of the absorption maximum from 457 nm (no urea) to 460 nm (7 Murea) .Comparabl e effects were reportedfo r pig heart lipoamide dehydrogenase [14]. This indicates that inth e oxidized state urea induces asmal l change in the polarity of the microenvironment of the isoalloxazine ring of the protein bound FAD. As already mentioned above, inth e absence of urea,ful l reduction to EH4b y NADH is not observed. Fig.4 shows the spectral properties of lipoamide dehydrogenase from P. fluorescens after reduction by excess NADHan d inth e presence of NAD+an d3. 0 M urea. In3 Murea , reduction by excess NADH leadst o reduction of the flavint oth e same extent as foundfo r Eox, howeverth e 700 nmabsorbanc e is lost (Fig.4A) .Thi s loss of 700 nm absorbance is most likely due to an urea induced change inth e polarity ofth e active site, rathertha nt o losso f NAD+bindin g (see below). After reoxidation by

112 air inlet,a typica l EH2spectru m is obtained indicating that under the conditions used, the dinieric structure of the enzyme is retained.Gelfiltratio n ofthi s particular sample over Btogel P-6DG reveals the presence of about 15% fre e FAD. For A.vinelandii enzyme (Cm urea= 1.5 M),unde r identical conditions,ful l reduction to EH4occur s and moretha n 90%o f FAD is released after reoxidation (not shown). In3 Mure a and excess of NAD+,over-reductio n of the P. fluorescensenzym e by NADHp s almost completely blocked (Fig.4B) .Th e absorption spectrum, showing an intense charge transfer banda t 530 nm remains virtually unchanged,eve n after reoxidition. The relatively high extinction at 530 nm might again be duet o an urea- inducepeffec t onth e polarity of the active site. Underth e same conditions,th e A. vinelandiii s only slightly protected from over-reduction by NADH;gelfiltratio n of the reoxidired A. vinelandiisampl e reveals the presence of about 80%fre e FAD (not shownj. The above results clearly confirm earlier conclusions that over-reduction of lipoamjde dehydrogenase to EH4 promotes subunit dissociation [11].

Discussio n

The results described inthi s Chapter show that, despite their very high degree of sequence homology and structural identity, P. fluorescensan dA. vinelandiilipoamid e dehydrogenase differ with respect redox properties. The most striking differences are observed upon reduction with excess NADH.Wit h lipoamide dehydrogenase from P.-fluotescens, binding of NAD+shift s the distribution of two-electron reduced species in favor of the disulfide reduced enzyme, whereas inth e A. vinelandiienzym e the electrons mainly reside onth e flavin.Ther e isthu s aclea r relation between the redox potential of the couple EH2/EH4 andth e sensitivity toward urea denaturation:th e lower the couple,th e lower the sensitivity toward ureadenaturation . The higher thermostability of the P. fluorescens enzyme when comparedt o the A.vinelandii enzyml, as reflected inal l redox states, is ascribedt o a higher dimer stability of the P. fluorescens enzyme. Sitefspecific mutagenesis experiments onth e A. vinelandiienzym e [4] indicate that small structuralchange s inth e vicinity of the subunit interface may leadt o drastic effects on enzyme catalysis. The catalytic properties andconformationa l stability ofth e mutated enzymes strongly suggest that the interaction of the C-terminus withth e other subunit influencesth e redox properties ofth e active site. Recent results onth e crystal structure of lipoatnide dehydrogenase from A. vinelandiiindicat e that inthi s enzyme the C- terminup also folds back intoth e protein. Refinement of the crystal structure of lipoamide dehydrogenase from P. fluorescens istherefor e of utmost importance. Together with

113 site-specific mutagenesis experiments, this will yield valuable information on the structural features involved inth e regulation of the redox properties of thistyp e of flavoenzymes.

Acknowledgements.

This work was supported in part byth e Netherlands Foundation of Chemical Research (S.O.N.) with financial aid from the Netherlands Organization for Scientific Research(N.W.O.) .

References

1. Benen,J .A . E.,Va n Berkel,W .J .H. ,Va n Dongen,W . M.A . M.,Muller , F.,an dD e Kok,A . (1989) J. Gen.Microbiol. 135, 1787-1797. 2. Mattevi,A. , Schierbeek,A .J .an d Hoi,W . G.J .(1991 ) J. Mol. Biol. 220,975-994 . 3. Mattevi,A ,Obmolova ,G ,Sokatch ,J . R., Betzel.C.an d Hoi,W.J.G .(1991 ) submitted. 4. Benen,J. , Van Berkel,W. ,Veeger , C. and De Kok,A . (1992) submitted. 5. Van Berkel,W .J . H.,Benen ,J .A . E.an dSnoek , M.C . (1991) Eur.J . Biochem. 197, 769-779. 6. Massey,V . Gibson,Q . H.an dVeeger , C. (1960) Biochem. J. 77, 341-351. 7. Williams,C . H.Jr . (1965) J. Biol.Chem. 240,4793-4800 . 8. Benen,J. , Van Berkel,W. , Dieteren,N. ,Arscott , D.,Williams ,C .Jr. ,Veeger , C.an d de Kok,A . (1992)submitted . 9. Wilkinson, K. D.an dWilliam s C. H.Jr . (1981) J. Biol.Chem. 256, 2307-2314 . 10. Benen,J. ,Va n Berkel,W. ,Zak ,Z. ,Visser ,T ,Veeger , C.an d De Kok,A . (1991)Eur. J. Biochem.202, 863-872 . 11. Berkelvan ,W .J .H. , Regelink,A .G. , Beintema,J .J .an d De Kok,A . (1991) Eur. J. Biochem 202, 1049-1055. 12. Veeger, Can d Massey,V . (1963) Biochim.Biophys. Acta 67,679-681 . 13. Wilkinson,K . D. andWilliam s C. H.Jr . (1979) J. Biol.Chem. 254,852-862 . 14. Massey,V. , Hofmann,T . and Palmer, G. (1962) J. Biol.Chem. 237, 3820-3828

114 I General discussion

Thewor k described inthi s thesis mainly focussed on mechanistic studies on lipoarride dehydrogenase from Azotobacter vinelandiia s approached by site directed mutagenesis. This enzyme was chosen as asubjec t of investigation to enhance our knowledge about the pyruvate dehydrogenase complex of which this enzyme is a constituent. For many yearsthi s complex, particularly the acetyl transferase core component, has been a major subject of investigation inou r laboratory. Atth e start of the investigations described inthi s thesis the genes encoding the components ofth e complex from Azotobacter vinelandiiwer e cloned in our laboratory by Westphal and Haanamaayer and initial progress had been made on the elucidation of the crystal structure by Schierbeek from the crystallography group at the Groningen University, headed by Prof. Hoi. Refinement studies, ultimately resulting in a resolution of 0.22 nm, were performed by Mattevi. Biochemical knowledge of this particular enzyme was howevjer scarce. Infi e sixties and seventies considerable progress was made on the elucidation of the reaction mechanisms of the pig heart and E.coli lipoamid e dehydrogenases by careful kinetic and spectral studies and by means of chemical modification. The enzymes were found to contain aflavi n and a redox active disulfide bridge. The two redox active groups allow the distinction of three redox states:th e oxidized enzyme (Eox),th e 2-electron reduced enzyme (EH2) andth e 4electro n reduced enzyme (EH4). Only Box and EH2ar e kinetically important, EH4constitute s an inactive species. The kinetical patterns of these enzymes inth e forward direction, reoxidation of dihydrolipoamide (lip(SH)2> with NAD+a s electron acceptor, were in general compatible with a ping pong mechanism.A major finding of these studies was the discovery of a base inth e active site, later identified as a histidine. The postulated function of the histidine isth e activation of the substrate dihydrolipoamide (lip(SH)2) by proton(abstractio n resulting in facilitated nucleophilic attack onth e disulfide bridge. This base also appeared to be involved inth e distribution of electronic species atthe , catalytlcally important, EH2 level of these enzymes. Inprde r to understandth e role of the histidine incatalysis ,w e replaced this histidine (His450) by several other residues: phenylalanine, serine and tyrosine. Apart from tljese, for catalysis seemingly 'drastic' mutations, additional mutated enzymes were pjrepared in which supposedly either the position of the histidine was altered (enzyme Pro451->Ala) or its chemical properties (enzymes Glu455->Asp and Glu455- >Gln),a s judged from the crystal structure of the highly homologous glutathione reductase. Inglutathion e reductase Pro468 (the equivalent of Pro451) is inth ecis- conformation anddirect s His467 (the equivalent of His450) toward the active site

115 disulfide bridge and Glu472 (the equivalent of Glu455) forms a short strong hydrogen bondwit h His450 andthu s might modulate its pKa.Thes e features were later shownt o be identical inth e structure of A vinelandiilipoamid e dehydrogenase. A detailed analysis ofth e spectral and kinetic properties of the wild-type and mutated enzymes isdescribe d in Chapters 3 and 4 respectively. The mutations all affect turnover, which will bediscusse d below. They also considerably affect the visible absorbance and fluorescence properties especially at the EH2level . Remarkable isth e strong increase of absorbance attributedt o the thiolate to FADcharge-transfe r complex at high pH (530 nmabsorption ) in all mutated enzymes when compared to wild-type enzyme (enzyme Pro451->Ala only transiently formsth e EH2species) . An increase in charge-transfer absorbance can beth e result of several factors or a combination thereof: 1) a smaller distance between the charge-donor (thiolate) and charge-acceptor (FAD), 2) a lower dielectrical constant, 3) an increase of the difference in redox potential between the charge-donor and charge-acceptor and 4) an increase inth e concentration of donor or acceptor. Incas e of factors 1-3, the charge-transfer absorbance not only increases, but alsoth e maximum shifts to longer wavelength. Incas e of factor 4,th e maximum remains unchanged. For the Glu455 mutated enzymes and enzyme His450->Ser no significant shift of the thiolate to FADcharge-transfe r absorbance maximum is observed,wherea s in enzymes His450->Phe and -Tyr aclea r red-shift occurred. It istherefor e concluded that the increased charge-transfer absorbance inth e mutated enzymes is primarily due to an increased concentration of thiolate as a result of a higher degree of deprotonation of the charge-transfer thiol relative toth e wildtyp e enzyme (FAD is constant) due to pKa changes of the participating groups, with an additional micropolarity effect inth e enzymes His450->Phe and -Tyr. Based onthi s conclusion the pH profiles of thethiolat e to FADcharge-transfe r absorbance as presented in chapter 2, Fig.4 wer e interpreted. The model for the distribution of electronic species developed for E.coli an dpi g heart EH2,take s onlytw o deprotonation steps into account, the histidine andth e charge-transfer thiol. The interchange thiol remains protonated at high pH.Thi s model predictstha t upon removal of the active site baseth e pHdependenc e of the thiolate to

FADcharge-transfe r absorbance would mainly obey one pKa, of the charge-transfer thiol. Our results withth e His450 mutated enzymes however clearly showtha t the pH dependence of thethiolat e to FADcharge-transfe r isgoverne d by two pKa values.

116 Injvie wo f the pig heart modelth e low pKa, equal inal lthre e His450 mutated enzymes, must be assignedt o the charge-transfer thiol. The high pKa must then be assignedt o at least one group that 1)influence s the charge-transfer absorbance (for moretha n 55% in enzyme Hi4450->Phe to more than 75%i n enzyme His450->Ser),

2) whose pKa is affected by polarity of the side-chain engineered and

3) isocountsfo rth e largest dropi nfluorescenc e despite the facttha t atth e pKao f thingrou p at least 95 %o fth e charge-transfer thiol isdeprotonated . There is however nodirec t structural evidence for a residue that could fulfill allthre e requiroments. Therefore we have developed a new model forth e A.vinelandii lipoarrid e dehydrogenase in which the interchange thiol as a third deprotonatable group participates.

InIhi s modelth e low pKa found forth e His450 mutated enzymes isassigne dt o the interchange thiol with the charge-transfer absorbance as a result of tautomerization withth echarge-transfe r thiol. The high pKa, associated withth e 3 marked effects menticne dabove , is logically assigned to the charge-transfer thiol (in effect, both prototiopic tautomers). Inth e wild-type enzyme,wher e three pKa values werefound , the low pKa is assigned toth e interchange thiol,th e intermediate pKa toth e charge- transfarthio l andth e high pKat oth e protonated histidine. Forth e Glu455 mutated enzymes it isconclude dthat , at low pH, co-titration of allthre e residues occurs due to a loweradpK a of His450 and losso fth e orienting effect of Glu455 onth e histidine toward the interchange thiolate. Despite the fact that the microscopic pKa model presented explains the observed macroscopic pKa values it should be kept in mindtha t the assignments made are still disputable and other residues might contribute as well. Therefore the model should be regarded as a working hypothesis.

Th(istead y state kinetics of the forward reaction reaction ofth e wild-type enzyme is compatible with a ping-pong bi bi mechanism. Previous studies in our laboratory had showntha t this enzyme functions according to aternar y complex mechanism.A t present we cannot satisfactorily explain this discrepancy. The Glu455- andTyr16 - (Chapler 4) mutated enzymes and enzyme His450->Ser also function according to a ping pong bi bi mechanism indicating that this mechanism is indeed correct. It should be notodtha tth e steady state kinetics ofth e Glu455-an dTyr1 6 mutatedenzyme s was studied with another electron acceptor, in casu,3-acetylpyridin e dinucleotide (APAD+). This was done because these enzymes show strong inhibition by the product NADH. The steady-state kinetics of the His450 mutated enzymes clearly showsth e important function ofthi s histidine as acatalyst . Enzymes His450->Phe and His450)->Tyr appear to be almost inactive in both directions. Enzyme His450->Ser is

117 still,thoug h minimally, active:V atth e pHoptimu m being 0.5% of wild-type activity in the physiological reaction. Rapid reaction kinetics showstha t for these enzymes the reductive half reaction using lip(SH)2 is rate limiting and extremely slow when comparedt oth e wildtyp e enzyme.Thi s important function ofth e histidine was also reported inth e course of our investigations from site directed mutagenesis studieso f E.coli lipoamidedehydrogenas e and E.coli glutathion e reductase. Enzyme Pro451->Ala is essentially inactive in both directions and it isconclude d that the loss of activity isdu et o over-reduction to EH4b y lip(SH)2 andNADH . Iti s shown forth e Glu455 mutated enzymestha t the activity inth e forward reaction isconsiderabl y lowertha n for the wild-type enzyme. Rapid reaction studies with lip(SH)2 showed that the rate ofth e reductive reaction of enzyme Glu455->Asp was halved and for enzyme Glu455->Gln this reaction was decreased approximately 10 fold. This indicates that inth e substrate bound enzyme Glu455->Asp the histidine still can act as abase . This function isclearl y hampered in substrate bound enzyme Glu455->Gln. The reduction of the flavin of the wild-type enzyme and His450- and Glu455 mutated enzymes by NADH proceeds very fast. Inth e wild-type enzyme this reduction isfollowe d by afas t reduction of the disulfide with concomitant reoxidation of the FAD. Inth e mutated enzymes this intramolecular transfer of a hydride equivalent is severely affected. Alsoth etransfe r of a hydride equivalent fromth e dithiolt oth e FAD,effecte d by NAD+, is severely diminished.Thes e properties of the mutated enzymes have enabled the spectral identification of species 4 and 6 of the reaction intermediates as presented in Scheme 1i n chapter 3. Species 5 inthi s scheme isth e so-called flavin C4a adduct species with acovalen t bond between the charge-transfer thiolate and C4a ofth e flavin.W e have obtained no spectral evidence for the participation of the proposed flavin C4aadduc t species during studies of the transfer of a hydride equivalent inth e mutatedenzyme s from species 4t o 6o r vice versa. This questions the existence of the C4a adduct species as acatalyti c relevant intermediate. The effect ofth e mutations oncatalysi s inchapte r 3 are interpreted inth e light of the currently 'established' involvement of this C4aadduc t species. Forth e reverse direction oftransfe r of a hydride equivalent (from species 6t o 4) it is concluded that this intermediate is probably not involved and direct hydridetransfe r occurs from the reducedflavi n to the disulfide. Similarly, asa result of microscopic reversibility, in the direction 4t o 6 itca n beconclude d that hydridetransfe r canaccoun t for the lacko f identification of the C4aadduc t species. One might arguetha t the mutations alterth e mechanism of transfer of a hydride equivalent from participation of the C4aadduc t species to direct hydride transfer or a concerted reaction. Before doing this one must consider the following.Th e flavin C4a

118 adductspecies has until now never been reported in any (mutated) lipoamide dehydrogenase except for pig heart enzyme where the interchange thiol was alkylated. Only idwil d type mercuric reductase this species has been observed upon reduction with N^DPH at low pH.Othe r reports onth e C4aadduc t were with mercuric reductase of whidh the interchange thiol is replaced by alanine andwit h thioredoxin reductase of which #ie flavin was replaced by 1-deaza-FAD.Th e majority of reports onth e C4a adductjspecies isthu s with enzymes that arefa r from 'healthy'. Interestingly too inthi s respect isth e fact that no C4a adduct formation was reported in monoalkylated glutathione reductase and no reports ofthi s species were made on E.co/ ;glutathion e reductase or E. coli lipoamide dehydrogenase in which the interchange thiol was mutated. Therefore it is proposed here that in wild-type lipoamide dehydrogenase the C4aadduc t is not a relevant catalytic intermediate andth e electron transfer from disulfide/dithiol to FAD/FADH-occur s via hydride transfer.

For(the Glu455 mutated enzymes strong inhibition by NADHwa s observed indicatingtha t the redox potential of the EH2/EH4coupl e was higher than in the wild- type enzyme. Therefore the quantitation of the redox potentials was investigated. These ^tudies were however frustrated as discussed inchapte r 3. Nevertheless it could bjeconclude d that the redox potential ofth e EH2/EH4coupl e of these enzymes is higher^ha ni n the wild-type enzyme. Spectral studies with these enzymes showedtha t EH2generate d by reduction by lip(SH)2 is rather stable anddoe s not result in easy reductiono fth e FAD (incontras t toth eTyr1 6 mutated enzymes andenzym e A9,cf chaptei 4 and below). Addition of AAD+ (a non-reducable NAD+analog ) to EH2o f enzyrm Glu455->Asp resulted in reduction of the FAD,strongl y indicating that binding of a nucleotide increasesth e redox potential of the FAD relative to that of the disulfide bridge. Similar results were obtained for this enzyme andth e wild-type enzyme in a study in which reduction was performed with enzymatically generated NADPH inth e presence and absence of AAD+. For bothth e wildtyp e enzyme andth e Glu455 mutated enzyme the binding of a nucleotide thus increases the redox potential of the FAD.wh y is itthe n that in enzyme Glu455->Asp strong inhibition is observed and not inwild-^yp e enzyme?. A possible explanation istha t the redox potential of the FAD- nucleotktecomple x is higher in enzyme Glu455->Asp than it is inth e wild-type enzyma.Thi s is supported bytw o lines of evidence. First,th e addition of AAD+t o EH2 of bothjenzyme s results in reduction of FADonl y in enzyme Glu455->Asp andsecond , transhyjlrogenation from NADH to thio-NAD+ was only observed with wild-type enzymei. Apparently, the redox potential of APAD+i s high enough to accept electrons fromtl" 4 reduced FAD in enzyme Glu455->Asp. Initially it wasthough t thatth e His450, which i$influence d by Glu455 is directly involved in modulation of the redox potential

119 ofth e FAD.However ,th e results obtainedwit hth eTyr1 6 mutatedenzyme s andC - terminal deletion mutants described in Chapter 4, show that the increased redox potential of the EH2/EH4coupl e ofthes e mutated enzymes isdirectl y relatedt o a diminished subunit interaction.Evidenc e isobtaine dtha t this may beth e case for the Glu455 mutated enzymes too in addition to a possible role for His450. The 10C-termina l residues are not visible inth e crystal structure of lipoamide dehydrogenase from A. vinelandii, but can be observed inth e crystal structures of lipoamide dehydrogenase from Pseudomonas putidaan d Pseudomonasfluorescens. Inthes e structures,th e C-terminus folds backtoward sth e active site and is involved in interactions with the other subunit. The function of the C-terminus of lipoamide dehydrogenase from Azotobacter vinelandiiwa s studied by deletion of 5, 9an d 14 residues, respectively. Tyrosine 16, conserved in all lipoamide dehydrogenases sequenced thus far, and shown from the other structures to be likely involved in subunit interaction,wa s replaced by phenylalanine and serine. Deletion of the last 5 residues does not influence the catalytic properties and conformational stability (thermoinactivation and unfolding by guanidinium hydrochloride). Removal of 9 residues results in an enzyme (enzyme A9) showing decreased conformational stability and high sensitivity toward inhibition by NADH. These features are even more pronounced after deletion of 14 residues. Mutation of Tyr16 also results in a strongly increased sensitivity toward inhibition by NADH and results in slow translocation of reducing equivalents from the dithiol to the FAD demonstrating that EH2i s not 'stable'. Similar conclusions can be drawn for enzyme A9.Th e conformational stability of bothTyr1 6 mutated enzymes is comparable to enzymeA9 . The results strongly indicate that a hydrogen bridge between tyrosine of one subunit (Tyr16 inth e A. vinelandiisequence ) and histidine of the other subunit (His470 inth e A. vinelandiisequence) , also exists inth e A. vinelandiienzyme . Inth e A9an d A14 enzymes this interaction is missing. It isconclude d that this interaction mediates the redox properties of the FADvi ath e conformation of the C-terminus comprising residues 450-470.

Chapter 5 describes the molecular cloning and sequence determination of lipoamide dehydrogenase from P. fluorescens. The derived amino acid sequence shows aver y high degree of sequence identity in comparison to the A.vinelandii sequence especially inth e region of residues which constitute the active site. Despite the high sequence homology and high degree of structural identity differences between the A. vinelandiian d P. fluorescenslipoamid e dehydrogenases

120 are manifest as presented inChapte r 6.Th e spectral differences are in accord withth e conclusion that the redox potential of the couple EH2/EH4 inth e nucleotide bound enzymo is lower inth e P. fluorescensenzyme . Duet oth e fact that NADHdoe s notfull y reduceth e enzyme to the EH4leve l (the redox potential of the FADi s lower inth e P. fluorescensenzyme ) the conformational stability of NADH reduced enzyme is much higheritha nfoun d for A vinelandiienzyme . Thetemperatur e stability of the P. fluoresipens enzyme is highertha n the A vinelandiienzym e andtherefor e strongly suggests a general relation between the dimer stability andth e redox properties.A relatiort which may also extend to other lipoamide dehydrogenases.

121 Samenvatting

Levende organismen, van bacterien tot complexe meercelligen, vertonen twee kenm^rkende eigenschappen: ze planten zich voort en ze vertonen alda n niet waarneembare activiteit. Teneinde deze eigenschappen tot uiting te brengen bedient een organisme zich van allerlei organische verbindingen die varieren in complexiteit. Tot dq meest complexe verbindingen horen het DNA end e eiwitten. Het DNA bevat erfelijke informatie die in pakketjes, genaamd genen, lineair gerangschikt zijn in het DNA. Voor het merendeel van de genen geldt dat de informatie die erin besloten ligt, codeert voor een eiwit. Een eiwit bestaat uit een keten van relatief eenvoudige bouwstenen, de aminozuren, waarvan in het algemeen 20 verschillende gebruikt worden om een eiwit sament e stellen. Met deze 20 aminozuren is in een lange keten, in priricipe, een oneindig aantal variaties te maken. Welke aminozuren op welke plaatsdoor de cellulaire eiwit-fabriekjes genaamd ribosomen in de groeiende eiwitketen ingebouwd moeten worden, ligt besloten in het gen coderend voor dat eiwit. be aminozuurketen, het eiwit, is op een voor ieder eiwit karakteristieke wijze tot een kuwen samengevouwen. Eiwitten kunnen grofweg verdeeld worden intwe e categorieen. Tot de eerste categerie behoren de enzymen en tot de tweede categorie worden hier alle andere eiwitten gerekend. Deze tweede categorie omvat onder andere signaaleiwitten en structiirele eiwitten. De enzymen zorgen ervoor dat readies die onder normale omstandigheden niet of nauwelijks plaatsvinden, snel en efficient verlopen.Z e wordan hierbij niet zelf omgezet. Met andere woorden: enzymen zijn katalysatoren. Voor elke natuurlijke organische verbinding is erzowel een synthese route als een afbraak route via enzymen. Een voorbeeld. Suiker in droge vorm is stabiel aan de lucht en dus reageert de suiker niet met de aanwezige zuurstof. Insterie l water is suiker eveneens stabiel. Suiker is, zoals bekend, een prima energie leverancier voor in levende organismen. De energie die opgeslagen zit in de opgeloste suiker wordt in de levende eel vrij gemaukt door de suiker met behulp van enzymen in verschillende stappen af te breken. In zuurstof afhankelijke organismen reageren deze afbraakproducten vervobens met zuurstof tot de uiteindelijke bouwstenen van de suiker, water en kooldipxide. Deze eindproducten kunnen dan weer door planten met behulp van enzymen weer worden omgezet in suiker onder invloed van zonlicht, dat voor de noodaakelijke energie zorgt. Enzymen zijn zeer specifiek voor wat betreft hun substra(a)t(en), zoals een sleutel past dp een slot. De reactie vindt plaats in het zogenaamde 'actieve centrum'. Naast enzymendi e reacties met een substraat katalyseren,zij n er ook enzymen die twee of

123 meerdere substraten met elkaar laten reageren. De volgorde van binding van de substraten aan het enzym is bepalend voor het mechanisme volgens welk het enzym werkt. Zo wordt er bijvoorbeeld onderscheid gemaakt tussen een geordend en een groeps-overdracht mechanisme (ook wel ping-pong mechanisme genoemd). In het geordend mechanisme binden eerst detwe e substraten,steed s in vaste volgorde, waarna de reactie plaatsvindt en beide producten vervolgens loslaten. Ingeva l van een groeps-overdracht mechanisme bindt altijd eerst een substraat waarbij een groep van het substraat overgedragen wordt op het enzym en het veranderde substraat, nu product genaamd, loslaat. Vervolgens bindt hettweed e substraat. Hierop wordt dan de tijdelijk op het enzym geparkeerde groep overgedragen, gevolgd door loslaten van het tweede product. De enzym gekatalyseerde reacties zijn in het algemeen omkeerbaar, dus uitgaande van de producten is het vaak mogelijk de oorspronkelijke substraten terug te vormen. Dit hangt af van de relatieve concentraties van de substraten en producten en van de 'kinetische karakteristieken' van het enzym Inbeginse l is hetz o dat als beide substraten in,t . o.v . het enzym,hog e concentraties aanwezig zijn en er geen producten aanwezig zijn bij de start van de reactie, na een korte aanloop fase (meestal minder dan een seconde) gedurende langere tijd een lineair verband is in detij d tussen de gebruikte substraat concentraties en de waargenomen snelheid van productvorming. Dit wordt dan de snelheid bij een gegeven substraatconcentratie genoemd. Naarmate een substraatconcentratie hoger wordt loopt de snelheid op. Dit blijft echter niet eeuwig doorgaan. Op een gegeven moment is het zoda t eentoenam e ind e substraatconcentratie niet meer resulteert in een toename van de snelheid. Dit heet de maximale snelheid (Vmax) en is kenmerkendvoo r ieder enzym. Door de afhankelijkheid van de snelheid van het enzym van beide substraten nauwkeurig te bestuderen is het mogelijk inzicht te krijgen of er bijvoorbeeld sprake is van een geordend mechanisme of van een groeps-overdracht mechanisme. Omdat er onder condities gewerkt wordt waarbij de snelheid in principe onveranderlijk is in de tijd bij een gegeven combinatie van substraten, is noodzakelijkerwijs de snelheid van binding van de substraten en snelheid van loslaten van de producten gelijk. Met andere woorden de concentraties van vrij enzym, substraat gebonden enzym en product gebonden enzym zijn constant in detij d en dus 'statisch'. Deze toestand wordt in Engelse bewoordingen 'steady state' genoemd. Bestudering van deze steady-state is vergelijkbaar met het beschrijven van de karakteristieken van een auto waarbij gedurende detest s steeds met een andere constante snelheid gereden werd. Deze karakteristieken zullen zeker interessant zijn maar echt leukword t het pas als ook de accelleratie van de auto bekeken wordt. Dit geldt evenzeer voor enzymen. De accelleratie fase van de auto is vergelijkbaar met

124 de aanhopfas eto t de steady-state kinetiek van het enzym. Dezefas e wordt de'pre - steady utate' fasegenoemd . Omdatdez e pre-steady state fase slechts zo kortduurt ,i s het noadzakelijk hiervoor geavanceerde apparatuur te gebruiken. De informatie die uit de nauwkeurige analyse van de pre-steady state fase gehaald wordt, geeft inzicht in de srelhei d van binding van ieder van de substraten en producten en maakt het soms n^ogelijko mtussenstappe n ind e omzettingva n substraat naar product of omgek^erd te identificeren als iedere tussenstap andere meetbare eigenschappen heeft. Na(di tintermezz o terug naar de suiker afbraak. Suikerafbraak door organismen onder ^uurstofloze condities leidt tot de vorming van eenvoudige zuren (bv. zuurkool en spieipijn) of alcohol. Hierbij wordt echter maar een kleingedeelt e van de in de suiker aanwezige energie benut. Inaanwezighei d van zuurstof is het mogelijk tot 600% meer energie uit de suiker te halen door 'verbranding' tot water en kooldioxide. Dit proces is veel complexer dan dat onder zuurstofloze condities en omvat dan ook een geweldig grotere hoeveelheid en varieteit aan enzymen. Eenva n de sleutel enzymen indi t proces is het pyruvaat dehydrogenase. Dit enzym is onderdeel van een enzymcomplex bestaande uit drie verschillende enzymen. Het substraat van dit complex, pyruvaat, isd e directe voorloper van de producten zoals verkregen onder zuurstofloze suikerafbraak. Het complex neemt dus het substraat voor deze zuurstofloze routes weg. Dedri e enzymen die deel uitmaken van dit complex zijn het pyruvaat dehydrogenase, het transacetylase en het lipoamide dehydrdgenase. Het pyruvaat dehydrogenase splitst kooldioxide af van pyruvaat en draagt het resterende deei van het substraat, een acetylgroep, over op een lipoylgroep van het transacetylase. Het transacetylase koppelt vervolgens de acetylgioep aan Coenzym A, dat afgeleid is van het vitamine panthotheenzuur (uit de groepva n de B-vitamines). Het acetyl-CoenzymA isd e verbinding die in een complex van realties wordt omgezetto t water en kooldioxide. Ook is het mogelijk met deze verbinding andere voor het organisme noodzakelijke verbindingen te maken. Nadat het transacetylase het acetyl-Coenzym A heeft afgegeven, resteert een lipoylgroep die niet in dejuist e toestand is om opnieuw een acetylgroep te ontvangen. De toestami waarin de lipoylgroep verkeert heet de 'gereduceerde' vorm. De juiste toestami om opnieuw een acetylgroep te ontvangen heet de 'geoxideerde' vorm. Het omzettan van de lipoylgroep van de gereduceerde in de geoxideerde vorm vindt plaats coor het lipoamide dehydrogenase. In feite doet het lipoamide dehydrogenase niets arders dan twee electronen (e-) van de lipoylgroep afpakken en deze overdragen op NAD+ (afgeleid van vitamine B1). Teneinde de electro-neutraliteit te handha/en zijn hierbij twee protonen betrokken (H+). Op de lipoylgroep worden door lipoamide dehydrogenase dus twee -SH groepen (dithiol) omgezet in een-S-S -

125 groep (disulfide) en ontstaat vrij NADH + H+. Dittyp e van readies wordt redox readies genoemd. De readie van lipoamide dehydrogenase verloopt volgens een groeps- overdracht mechanisme en kan dus alstwe e deelreadies worden voorgesteld:

Eox + lip(SH)2 ^=^ EH2+ lipS2 (1)

+ + EH2 + NAD s ^ Eox + NADH + H (2)

In het algemeen bevatten enzymendi e redox readies katalyseren een extragroe p zijnde een metaalatoom of prosthetische groep, of beide. Lipoamide dehydrogenase bevat als extra groep een flavine molecuul. Dit flavine is afgeleid van vitamine B2e n heeft in zijn geoxideerde toestand een felgel e kleur. Uit de literatuur is bekend dat lipoamide dehydrogenase naast het flavine nog een tweede redoxadieve groep bevat: een disulfide-brug. Deze brug is analoog met de disulfide-brug van de lipoylgroep en kan dus ook geopend worden. Het verschil met de lipoylgroep zit in het feit dat detwe e SHgroepe n in het enzymafkomsti g zijn van detwe e aminozuren, cysteines, die een dusdanige positie ten opzicht e van elkaar hebben dat de brug vorming mogelijk is. De twee redoxadieve groepen, het flavine en de disulfide-brug, zorgen ervoor dat het enzym in principe drie redoxtoestanden kent: de geoxideerde vorm (Eox),d e 2-electronen gereduceerde vorm (EH2) en de 4-eledronen gereduceerde vorm (EH4). Aangetoond is dat voor katalyse alleen de vormen Eox en EH2 relevant zijn. Devormin g van EH4blee k niet produdief. Detwe e eledronen die in de katalyse betrokken zijn gaan over in paren. Eerst wordt de disulfide-brug gereduceerd en vervolgens worden de eledronen gezamenlijk overgedragen op het flavine. Voor bestudering van het gezuiverde enzym wordt als substraat gebruik gemaakt van gereduceerd lipoamide (lipSH)2> in plaats van de lipoylgroep van het transacetylase. Het feit dat de twee eledronen overgaan in paren resulteert in een volledig gereduceerd flavine. Dit gereduceerde flavine is kleurloos. Dus zou men slechts een kleurverandering in het enzym verwachten, van geel naar kleurloos. Reductie van het enzym zoals ge'isoleerd uit varkenshart tot het EH2 niveau gaf echter duidelijk de vorming van een rood-gekleurd enzym te zien. Aanvankelijk werd dit gei'nterpreteerd als een 1-eledron gereduceerd flavine (dat ook rood is van kleur). Later bleek dat deze rode kleur het gevolg is van een zogenaamd ladings-overdracht complex (charge-transfer complex) tussen het flavine en een gedeprotoneerde SH groep (een thiolaat ion:S -). Ditcharge-transfe r complex bestaat alleen als er licht opvalt , m.a .w . het is een aangeslagen toestand. In het donker ise r weliswaar een soort basale interadie tussen het thiolaat en het flavine, doch er vindt geen ladingsoverdracht

126 plaats. Ind e EH2-toestand zitten de electronen dus hoofdzakelijk op de gereduceerde disulfide. Nanst de vorming van een thiolaat-flavine charge-transfer complex bleek ook de vorming van een gereduceerd flavine-NAD+comple x opt e treden. Dit uit zich door een gi^en tot grijs/blauwe kleur van het enzym. De verschillende kleur veranderingen (spect^ale vormen) van het enzym ind e verschillende toestanden heeft het mogelijk gema^kt ind e pre-steady state fase een aantal reactie-tussenstappen te identificeren. Naijwkeurige analyse van kinetische studies end e spectrale vormen had ertoe geleidjda t er een basewer dvoorgestel d in het actieve centrum van het enzym.D e functiejva n die base is het abstraheren van een proton van het substraat lip(SH>2me t als getolg dat het gevormde thiolate gemakkelijker op de disulfide-brug van het enzymka n aanvallen. Deze base bleek een histidine te zijn. Dit is een aminozuur met een zii^eten die een proton kan opnemen of afstaan bij een ongeveer neutrale pH.D e pHwaard e waarbij 50 %va n de verbinding een proton heeft afgestaan noemt men per definitie de pKa. DepK a waarde van de histidine is niet onveranderlijk maar is afhanMelijk van de omgeving waarin de histidine zich bevindt. Zo zal de aanwezigheid van ean negatieve lading ervoor zorgen dat de pKa omhooggaat , dus zal de histidine mindejmakkelij k een proton afstaan.Omda t deze histidine zich in het actieve centrum bevindt, dicht bij de disulfide-brug kan er een proton worden opgenomen dat eigenlijk bij 66n| van de door-reductie-onstane thiolen hoort. Dat dit gebeurt, blijkt onder andere door dfecharge-transfe r absorptie die de rode kleur tot gevolg heeft. Veranderingen in het en^ym die de pKa van deze histidine veranderenzulle n dus ook een effect hebbefi op de rode kleur. Daarnaast zullen deze veranderingen effect hebben op de katalytjsche activiteit van het enzym daar de histidine hierbij betrokken is. Onlangs is de 3-dimensionale structuur van het lipoamide dehydrogenase uit Azoto^acter vinelandiiopgehelderd . Uit deze structuur blijkt dat het enzym uit twee identieke subeenheden bestaat. Per enzym molecuulzij n twee actieve centra aanwefeig. leder actief centrum blijkt opgebouwd uit aminozuren die deel uitmaken van bgide subeenheden. Zo behoort de redoxactive disulfide-brug bij de ene subeepheide n de histidine bij de andere subeenheid. Het flavine zit als het ware tussenbeide subeenheden in, naast de disulfide-brug en de histidine. Vlak bij de histidine blijkt nog een glutamaat te zitten. Dit aminozuur is negatief geladen en kan dus van invloed zijn op de eigenschappen van de histidine. Omdat het actieve centrum tussen beide subeenheden gelegen is, mag verwacht worden dat de stabiliteit van de subeenheid-interactie van belang is voor de katalyse.

Mel de hierboven geschetste structurele en de biochemische kennis gewapend werd besloten nat e gaan wat er gebeurt met het enzym als de histidine vervangen

127 zou worden door een aminozuur met andere eigenschappen. Daarnaast werd er ook gekeken naar het effect van vervanging van de glutamaat naast de histidine. De achterliggende gedachte hierbij is om meer inzicht te krijgen in het reactiemecha- nisme van lipoamide dehydrogenase. De vervanging van de aminozuren is heel eenvoudig te bewerkstelligen door de DNA-code voor het enzym iets te veranderen. De resultaten en ideeen,di e bestudering van deze veranderde enzymen heeft opgeleverd,worde n beschreven in hoofdstuk 2 en 3 van dit proefschrift. In hoofdstuk 2 worden de spectrale eigenschappen van de veranderde enzymen beschreven. De nadruk is daarbij gelegd op de bestudering van de spectrale eigenschappen van de katalytisch belangrijke EH2 toestand (zie reactie vergelijkingen boven) met name de rode kleur die een maat is voor de graad van protonering van dethiolaa t die het dichtst bij het flavine zit (de charge-transfer thiolaat). De pH-afhankelijkheid van de rode kleur van de veranderde enzymen bleek niet te beschrijven met een model,gebaseer d optwe e pKa-waarden:d e histidine en de charge-transfer thiol, zoals voorgesteld voor andere lipoamide dehydrogenases.

Daaromwer d een nieuw model voorgesteld dat uitgaat van drie pKa-waarden. Nieuw aan dit model is niet alleen de toevoeging van een extra pKa-waarde maar bovendien laat dit model deprotonering van de andere thiol (de 'interchange-thiol') toe engaa t er zelfs van uitda t deze thiol als eerste deprotoneert. Dit laatste is gebaseerd op resultaten verkregen met enzymen waarin de histidine is vervangen door serine, tyrosine of phenylalanine. De enzymen waarin de glutamaat vervangen werd door glutamine of aspartaat, gaven een beeldte zien dat sterk afweek van het normale enzym. De resultaten zijn evenwel nog steeds verklaarbaar met het model,doc h met

het verschilda t de drie pKa-waarden zo dicht bij elkaar liggen dat ze infeit e als $en pKa gemeten worden. Deze resultaten geven duidelijk aan dat de pKa van de histidine door de glutamaat bepaald wordt. In hoofdstuk 3word t het effect van de veranderingen in de enzymen opd e katalyse beschreven. Alle veranderde enzymen geven verlaagde activiteit te zien. Dit is het sterkst bij de enzymen waarbij de histidine vervangen is. Hierbij iszowe ld e eerste als de tweede deelreactie sterk vertraagd. De oorzaak van de trage eerste deelreactie ligt in het feit dat de histidine niet meer aanwezig is enzodoend e geen proton meer van het substraat kan afpakken. Deomgekeerd e reactie blijkt ook heel sterk vertraagd. Devoornaamst e oorzaak hiervan isdezelfd e als die in de voorwaartse richting:d e overgang van de electronen en een proton van de gereduceerde disulfide-brug naar het flavine of omgekeerd is zeer sterk vertraagd. De enzymen waarin de glutamaat werd vervangen werden zeer sterk geremddoo r het product NADH. Dit iszee r waarschijnlijk toe te schrijven aan de vorming vand e inactieve EH4vorm . Met een aangepast substraat, i.p . v. NAD+,blee k wel activiteit te

128 metenjmaa r deze was sterk verminderd invergelijkin g met het normale enzym. Analyse van de deelreacties leerde dat niet zo zeer de eerste deelreactie hiervoor veranliivoordelijk is maar ook weer de overgang van de electronen en een proton van de gefeduceerde disulfide naar het flavine. Dejtrage overgang van de electronen en een proton hebben het mogelijkgemaak t twee rJBactie-intermediairen te identificeren. Een reactie-intermediair dat in de literatyur was voorgesteld en dat in principe geidentificeerd had kunnen moeten worden, werd niet waargenomen. Dit duidt erop dat het in de literatuur voorgestelde intermediair mogelijk niet voorkomt. In hoofdstuk 4 is een onderzoek naar het effect van de subeenheid interactie opd e katalyse beschreven. Hierbij is met name gekeken naar de invloed van de verwijdering van enkele aminozuren aan de staart (C-terminus) van het enzym. Deze aminoruren waren in de structuur niet te zien en leken ogenschijnlijk overbodig. Ook werd er een aminozuur veranderd van de ene subeenheid dat mogelijk een interactie met d* staart van de andere subeenheid heeft. De resultaten geven zeer duidelijk aan dat dejstaar t van de subeenheid en de wederzijdse subeenheid-interactie een sterk effect hebben op de katalyse; in het bijzonder bepalend zijn voor de mate waarin het enzyn^gerem dword t door het product NADH. In (loofdstuk 5 wordt de opheldering van de DNA code van het gen voor lipoamide dehydrogenase van Pseudomonasfluorescens beschreven . De aminozuren van dit enzyrrj blijken voor 84% identiek aan dat van Azotobacter vinelandii. Op grand hiervafi zou men geneigd zijn dat deze enzymen veel overeenkomst zouden moeten vertonJ9n. In hoofdstuk 6 zijn experimenten beschreven waaruit blijkt dat dit niet zo hoeft te zijn. Er is een aanzienlijk verschil tussen deze enzymen wat betreft de neiging tot de vormirjg van de inactieve EH4vorm . Het enzym uit P. fluorescens blijkt veel sterker bestartdtege n de door-reductie naar deze EH4vorm . Er zijn aanwijzingen dat dit mogelijk verband houdt met de stabiliteit van het enzym:d e subeenheid interactie.

129 Curriculum vitae

Jacques Alex Erwin Benen werdgebore n op 6 november 1959te Nuth. In 1978 behaal^Je hlj het diploma Atheneum Baa n het St. Janscollege te Hoensbroek. In datzelf^iejaa r werd begonnen met de studie blologie aan de Katholieke Universiteit te Nijmegen. Het kandldaatsexamen werd in 1981 behaald. De doctoraalfase werd in septerrjber 1985 afgesloten en omvatte de hoofdvakken moleculaire biologie (Prof. Dr. J. G.G JSchoenmakers ) en microbiologie (Prof. Dr. G. D.Vogels ) en het bijvak dierfysiplogie (Prof. Dr. S. E. Wendelaar-Bonga). In 1981 werd het diploma stralingsdeskundige op het C-niveau behaald. Van 1 augustus 1986to t 31 maart 1991 was hij als promovendus verbonden aan de afdelinj Biochemie van de Landbouwuniversiteit te Wageningen. Tot oktober 1987 is hij werkzaam geweest met de klonering van het gen coderend voor para- hydroxybenzoaat hydroxylase van Pseudomonasfluorescens. Vana f november 1987 tot maart 1991 werden de werkzaamheden verricht die hebben geresulteerd in dit proefsdhrift. Resultaten uit hoofdstuk 2,3 e n 4werde n gepresenteerd op het Tenth International Symposium on Flavins and Flavoproteins' gehouden van 15to t 20jul i 1990te |Como , Italie. Injanuar i 1991 werd gedurende drie weken een werkbezoek gebracjit aan de afdeling Biological Chemistry van de University of Michigan,An n Arbor, Michigan, USA bij Prof. Dr. C. H.William s Jr. j

List ojf publications

1) ^enen,J.A.E. ,va n Berkel,W.J.H. , van Dongen,W.M.A.M. ,Muller , F.an dd e Kok, A. (1989). "Molecular cloning and sequence determination of the Ipd-gene •ncoding lipoamide dehydrogenase from Pseudomonasfluorescens." J. Gen. Microbiol. 135,1787-1797 . 2) Westphal,A.H. , Eschrich,K. , van Dongen,W.M.A.M. ,Benen ,J.A.E. ,d e Kok,A . and van Berkel,W.J.H . (1990). "Site-directed mutagenesis of para-hydroxy lienzoate hydroxylase from Pseudomonasfluorescens." i n Flavinsand Flavoproteins 1990 (Curti, B., Ronchi,S .an dZanetti ,G .eds. ) W.d e Gruyter and Co. Berlin-New York, pp.231-234 . 3) Benen,J.A.E. ,va n Berkel,W.J. H andd e Kok,A . (1990). "Site-directed ijnutagenesis studies on lipoamide dehydrogenase from Azotobacter vinelandii." iji Flavinsand Flavoproteins 1990 (Curti, B., Ronchi,S . andZanetti ,G .eds. ) W. $e Gruyter and Co. Berlin-New York, pp.557-564 .

131 4) Benen,J.A.E. ,Dieteren ,N.A.H.M. ,va n Berkel,W.J.H .an dd e Kok,A . (1990). "Lipoamide dehydrogenase from Azotobacter vinelandii- kinetic studies onwil d type and mutated enzymes." in Flavinsand Flavoproteins 1990 (Curti, B., Ronchi, S. andZanetti ,G .eds. ) W.d e Gruyter andCo . Berlin-New York, pp. 565- 568. 5) Schulze, E., Benen,J.A.E. ,Westphal ,A.H. ,va n Berkel,W.J.H .an dd e Kok,A . (1990). "Binding studies of the dihydrolipoamide dehydrogenase component (E3) inth e pyruvate dehydrogenase complex from Azotobacter vinelandii." in Flavinsand Flavoproteins 1990 (Curti, B., Ronchi,S .an dZanetti ,G . eds.)W .d e Gruyter and Co. Berlin-New York, pp.569-572 . 6) van Berkel,W.J.H. ,Benen ,J.A.E .an d Snoek, M.C. (1991)."O nth e FAD induced dimerization of apo-lipoamide dehydrogenase from Azotobacter vinelandiian d Pseudomonas fluorescens - Kinetics of reconstitution." Eur.J. Biochem. 197, 769-779. 7) Schulze, E., Benen,J.A.E. ,Westphal ,A.H .an dd e Kok,A . (1991)."Interactio no f lipoamide dehydrogenase with the dihydrolipoyl transacetylase component of the pyruvate dehydrogenase complex from Azotobacter vinelandii." Eur.J. Biochem.200, 29-34 . 8) Snoep,J.L , Westphal,A.H. , Benen,J.A.E. ,Teixeir ad e Mattos,M.J. ,Neyssel , O.M. andd e Kok,A . (1991). "Isolation andcharacterisatio n ofth e pyruvate dehydrogenase complex of anaerobically grown Enterococcus feacalisNCT C 775.". Eur.J. Biochem., inth e press 9) Benen,J. , van Berkel,W. ,Zak ,Z. ,Visser ,T. ,Veeger , C. andd e Kok,A . (1991). "Lipoamide dehydrogenase from Azotobacter vinelandii:site-directe d mutagenesis of the His450-Glu455 diad. Spectral properties of wild-type and mutated enzymes." Eur.J. Biochem. 202,863-872 . 10) Benen,J. , van Berkel,W. , Dieteren,N. ,Arscott , D.,William s Jr, C, Veeger,C . and de Kok, A. (1992) "Lipoamide dehydrogenase from Azotobacter vinelandii: site-directed mutagenesis of the His450-Glu455 diad. Kinetics of wild-type and mutated enzymes." Eur.J. Biochem., submitted 11) Benen,J. ,va n Berkel,W. ,Veeger , C. andd e Kok,A . (1992) "Lipoamide dehydrogenase from Azotobacter vinelandii: Role ofth e C-terminus in catalysis anddime r stabilization." Eur.J. Biochem.,submitte d 12) Bastiaens, P. I.H. ,va n Hoek,A. , Benen, J.A . E., Brochon,J .C .an dVisser ,A .J . W. G. (1992) "Conformational dynamics and intersubunit energy transfer in wild type and mutant lipoamide dehydrogenase from Azotobacter vinelandii. A multidimensional time-resolved polarized fluorescence study." Biophys.J. , submitted.

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