Biochimica et Biophysica Acta - Tahoma Clinic · Review The multifaceted pyridoxal...

Biochimica et Biophysica Acta 1814 (2011) 1497–1510

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Review

The multifaceted pyridoxal 5′-phosphate-dependent O-acetylserine sulfhydrylase☆

Andrea Mozzarelli a,b,⁎, Stefano Bettati a,b, Barbara Campanini a,b, Enea Salsi a, Samanta Raboni a,Ratna Singh a, Francesca Spyrakis a,b, Vidya Prasanna Kumar c, Paul F. Cook c,⁎⁎a Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italyb Institute of Biostructures and Biosystems, Rome, Italyc Department of Chemistry and Biochemistry, University of Oklahoma, Stephenson Life Sciences Research Center, Norman, OK, USA

Abbreviations: AA, α-aminoacrylate external Schphosphosulphate; ATPS, ATP sulfurylase; BCA, β-chsynthase; EA, external Schiff base; IA, internal Schiff baO-acetylserine; OASS, OAS sulfhydrylase; OPS, O-phosdrylase; PAPS, 3′-phosphoadenosine 5′-phosphosulfateRSAP, reductive sulfate assimilation pathway; SAT, serthio-2-nitrobenzoate; GdnHCl, guanidinium hydrochlor☆ This article is part of a Special Issue entitled: Pyrid⁎ Correspondence to: A. Mozzarelli, Department o

Biology, University of Parma, Via GP. Usberti 23/A, 4312⁎⁎ Correspondence to: P.F. Cook, Department of

University of Oklahoma, Stephenson Life Sciences ResParkway, Norman, OK 73019-5251, USA.

1570-9639/$ – see front matter © 2011 Elsevier B.V. Aldoi:10.1016/j.bbapap.2011.04.011

a b s t r a c t
a r t i c l e i n f o
Article history:Received 9 March 2011Received in revised form 17 April 2011Accepted 20 April 2011Available online 28 April 2011

Keywords:PLP catalysisEnzyme mechanismSpectroscopyVitamin B6

Cysteine is the final product of the reductive sulfate assimilation pathway in bacteria and plants and servesas the precursor for all sulfur-containing biological compounds, such as methionine, S-adenosyl methionine,iron–sulfur clusters and glutathione. Moreover, in several microorganisms cysteine plays a role as a reducingagent, eventually counteracting host oxidative defense strategies. Cysteine is synthesized by the PLP-dependent O-acetylserine sulfhydrylase, a dimeric enzyme belonging to the fold type II, catalyzing a beta-replacement reaction. In this review, the spectroscopic properties, catalytic mechanism, three-dimensionalstructure, conformational changes accompanying catalysis, determinants of enzyme stability, role of selectedamino acids in catalysis, and the regulation of enzyme activity by ligands and interaction with serineacetyltransferase, the preceding enzyme in the biosynthetic pathway, are described. Given the key biologicalrole played by O-acetylserine sulfhydrylase in bacteria, inhibitors with potential antibiotic activity have beendeveloped. This article is part of a Special Issue entitled: Pyridoxal Phospate Enzymology.

iff base; APS, adenosine 5′-loro-L-alanine; CS, cysteinese; NAS, N-acetylserine; OAS,phoserine; OPSS, OPS sulfhy-; PLP, pyridoxal 5′-phosphate;ine acetyltransferase; TNB, 5-ideoxal Phospate Enzymology.f Biochemistry and Molecular4 Parma, Italy.Chemistry and Biochemistry,earch Center, 101 Stephenson

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© 2011 Elsevier B.V. All rights reserved.

1. Role of OASS in cysteine metabolism

1.1. Cysteine in the context of sulfur metabolism

Inorganic sulfur is the most abundant element on Earth aftercarbon, oxygen and nitrogen, but, differently from the aforemen-tioned elements, it is contained in a limited number of biologicalcompounds [1]. This situation is attributed to the high reactivity ofsulfur and its limited use in the more oxidized states [2]. In fact, onlylower oxidation states of sulfur participate in anabolic reactions [2]. Inthe presence of oxygen, it is usually found as sulfate and from thisform it must be reduced to sulfide before assimilation into organicmolecules like cysteine.

The central role played by cysteine in organic sulfur metabolism isdemonstrated by the fact that mobilization of sulfur from cysteine isthe first step of many biosynthetic pathways that supply the cell withbiomolecules such as Fe–S clusters, modified tRNAs (thiouridine),thiamine, biotin, glutathione andmycothiol [2–4]. Whereas mammalsrely on sulfated amino acids for sulfur supply, mostly the essentialmethionine, most bacteria and plants do perform the assimilation ofsulfur into cysteine through the reductive sulfate assimilationpathway (RSAP).

Beside its function as a protein building block and as a component ofimportant biomolecules, cysteine plays an essential role in the life cycleof pathogens such as Trichomonas vaginalis,Mycobacterium tuberculosis,and Salmonella typhimurium. In fact, cysteine participates directly or as aprecursor of reducingagents in themaintenanceof the redox state of thecell. This function is of special interest to microorganisms that spendpart of their life cycle in highly oxidizing environments, such as humanmacrophages. The ability of counteracting host oxidative defenses iscritical for pathogen survival during long latent phases.

1.2. Cysteine biosynthesis in prokaryotes

Cysteine biosynthesis in prokaryotes generally adheres to thecommonly accepted scheme shown in Fig. 1 (see [1,5] for compre-hensive reviews), although many exceptions exist that are of limitedinterest to this review. For this reason, here we will focus on the well-characterized sulfur assimilation in enteric bacteria and we willstress some of the main differences found in pathogens such as

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1498 A. Mozzarelli et al. / Biochimica et Biophysica Acta 1814 (2011) 1497–1510

M. tuberculosis for which sulfur metabolism has been suggested as anemerging drug target (see below).

Genes involved in sulfur assimilation in S. typhimurium andEscherichia coli are organized in a regulon, i.e. a group of operonswhose expression is collectively regulated. A comprehensive review onthe chromosomal organization and on the regulation of the cysteineregulon was presented by Kredich for S. typhimurium and E. coli [5,6].Five regions containing genes coding for proteins of RSAP wereidentified in S. typhimurium: cysG, cysE, cysB, cysAMK and cysCDHIJ(see also Fig. 1) [5]. InE. coli the organizationof genes is similar,with fewdifferences (see [1] for an exhaustive overview of E. coli sulfurmetabolism and enquire the web site EcoCyc [7] for information onthe metabolic pathway, chromosomal organization and the groupinginto transcriptional units). Whereas all the aforementioned genes codefor proteins involved in sulfate transport, activation, reduction andincorporation into cysteine, CysB is a LysR type regulator that activatesthe transcription of most of the cysteine regulon genes (see paragraphon 1.3. Regulation of cysteine biosynthesis in prokaryotes).

Sulfur assimilation in enteric bacteria, like E. coli or S. typhimurium,can be split into two main steps: an energetically highly demandingreduction process that converts sulfate (+6) to sulfide (−2), followedby the incorporation of sulfide into cysteine [1]. RSAP begins with theactive transport of sulfate into the cell and its subsequent reduction tosulfide with consumption of 1 mol of GTP, 2 mol of ATP and 1 mol ofNADPH for eachmole of cysteine synthesized. The need to couple sulfatereduction with ATP/GTP hydrolysis is due to the fact that sulfate is apoorly reactive molecule and needs to be activated prior to reduction[1,8]. It isworthmentioning that the pathway for sulfate reduction leadsto the synthesis of an intermediate, 3′-phosphoadenosine 5′-phospho-sulfate (PAPS), that is a universal sulfuryl group donor ([8,9] andreferences therein). RSAP can be considered a “superpathway”, built bythe convergence of two branches, the sulfate reduction pathway and theO-acetylserine (OAS) biosynthetic pathway that originates fromglycolysis. The two branches converge on the substitution reactionthat involves OAS and sulfide, catalyzed by the pyridoxal 5′-phosphate(PLP)-dependent enzyme OAS sulfhydrylase (OASS).

The reaction catalyzed by OASS is a β-substitution, i.e. a β-elimination reaction of acetate, followed by the addition of sulfide to aPLP covalently attached intermediate (see below) [10,11]. Thisreaction, with some exceptions in organisms such as T. vaginalis,involves the acetylation of the β-hydroxyl group of serine catalyzedby serine acetyltransferase (SAT). The reaction catalyzed by SAT is atthe branch point of sulfur, carbon and nitrogen assimilation [12] (see[13] for a general scheme). It is well assessed that OASS and SAT forma complex, known as cysteine synthase (CS) [13–15], whose functionin plants as a sensor of sulfur levels inside the cell has been proposed[16]. However, differences in metabolism and compartmentalizationbetween plants and bacteria do not allow the direct extension of thishypothesis to bacteria and, thus, CS function in enteric bacteria is stilla matter of debate. One important aspect of cysteine biosynthesis inprokaryotes lies in the existence of two isozymes of OASS, namelyOASS-A and OASS-B, the products of cysK and cysM, respectively.OASS-A is the best characterized of the two proteins [10,11,17–31]and is highly expressed at basal levels [32]. Conversely, OASS-Bfunction is still controversial and its activity accounts for only 20% oftotal cysteine biosynthesis at most [33]. The two enzymes share a 43%sequence identity [34], have an almost superimposable threedimensional structure [34,35], and similar activities, with OASS-Bbeing more promiscuous and able to accept bigger substrates [21,34].Although early reports stated that OASS-B was mainly expressedunder anaerobic conditions [21], reports on the differential expressionof the two isozymes are scarce and do not deal with the distinct rolespossibly played by the two enzymes in infection and long-termsurvival. Redundancy in enzymatic function points to the central roleplayed by cysteine biosynthesis in bacteria, especially for thosepathogens, like S. typhimurium and M. tuberculosis, that have to face

highly oxidizing environments during the persistent phase of their lifecycle [8,36]. One important difference between the two isozymes isthe ability to form CS: only OASS-A is able to interact with SAT [37,38].This is a central point in the understanding of the differential functionof the two enzymes and needs further investigation. Also, thebiological significance of the ability of OASS-B to use thiosulfateas a sulfur source is unclear [33] because the product of this reaction,S-sulfocysteine, needs to be converted into cysteine. In E. coli theexistence of such an enzymatic activity has not been demonstrated[1]. The scenario regarding the product of cysM is clearer inM. tuberculosis [39,40] and T. vaginalis [41], where this isoform hasbeen thoroughly investigated and proposed as an antibacterial drugtarget. In M. tuberculosis three genes annotated as OASS weredescribed: cysK1 shows high homology to OASS-A, cysM shows highhomology to OASS-B, and the third one, still poorly characterized, ishomologous to both OASS-A and OASS-B and is often referred to ascysK2 [42]. Whereas CysK1 is a typical OASS and interacts with SAT[42], it was shown that CysM is a O-phosphoserine (OPS) sulfhy-drylase [39,40]. This finding follows the results of early works onT. vaginalis [41] and Aeropyrum pernix [43] where the existence of anOPS sulfhydrylase (OPSS) activity was detected for the first time.Following these discoveries, it was suggested that the gene productCysK codes for OASS, whereas the gene product CysM codes for OPSSeven in enteric bacteria like E. coli [41]. The picture is actually morecomplicated because CysM from M. tuberculosis uses a sulfurdonor other than sulfide. In fact, Actinomycetales, which includesM. tuberculosis, is at present the only group of bacteria that possess aprotein, indicated as CysO, that becomes thiocarboxylated at the C-terminus [40,44,45]. This protein directly interacts with CysM andinserts its thiocarboxylated C-terminal arm into CysM active site. Thenegatively charged sulfur atom attacks the α-aminoacrylate interme-diate (see below) to give free CysM and a CysO-cysteine thatrearranges to S-CysO-cysteine and is eventually hydrolyzed to CysOand cysteine [40]. It was proposed that this pathway has evolved toprovide cysteine in harsh environments, such as in macrophages [46]and hyperthermophilic niches because it relies on more stable formsof substrates for cysteine biosynthesis, i.e. OPS in place of OAS andthiocarboxylated CysO in place of free sulfide [47]. At present, theexistence and role of such an alternative pathway in enteric bacteriaare uncertain. In fact, although the hallmark for OPS vs. OAS specificityof CysM has been identified as a conserved arginine residue [41,48],also present in E. coli, Haemophilus influenzae and S. typhiumuriumOASS-B, no OPSS activity has been shown for S. typhimurium andH. influenzae proteins (PFC, unpublished observation).

Regardless of the differences among microorganisms in RSAP, thecentral role played by cysteine in the metabolism of prokaryotes pointsto the exploitation of this pathway as a new source of targets forantibacterial drugs [46,49–55]. In fact, as nicely summarized in [8],targeting cysteine biosynthesis would affect the ability of bacteria tocounteract host defenses during the persistent phase of infection.Proteins and enzymes involved in sulfur assimilation are dispensable forlife of the pathogen outside the host and, in some cases, also duringinfection, but become essential for long-term survival. Because OASScatalyzes theessential,final stepof cysteinebiosynthesis, it is at theheartof the regulatory mechanisms of RSAP (see below). This enzyme is notpresent in mammals, in contrast to other proteins, like ATP sulfurylase[50], and thus targeting OASS is proposed as an effective strategytowards the development of new antibiotics [54]. This is demonstratedby the fact that CysM in M. tuberculosis is essential for survival inmacrophages [56,57] and CysK/CysMknockout strains of S. typhimuriumare impaired in their ability to develop antibiotic resistance [53].

1.3. Regulation of cysteine biosynthesis in prokaryotes

The central role played by cysteine biosynthesis in bacterialmetabolism is witnessed by its multilevel, strict regulation:

Fig. 1. Reductive sulfate assimilation pathway in bacteria. Enzymes involved in RSAP are shown in bold with the name of their coding genes, following the scheme proposed for E. coli [1].Induction of transcriptional activity by NAS occurs by the action of the regulator CysB. Abbreviations used are as follows: ATPS, ATP sulfurylase; APSK, 5-phosphosulfate kinase; PAPS R,3-phosphoadenosine 5-phosphosulfate reductase; NADPH-SR, NADPH-dependent sulfite reductase.

1499A. Mozzarelli et al. / Biochimica et Biophysica Acta 1814 (2011) 1497–1510

transcriptional activation, feedback inhibition and regulatory protein–protein interactions. Interestingly, most of cysteine biosyntheticregulation affects either OASS or SAT activity/expression and onceagain substantiates the importance of CS in RSAP.

As it is common for bacterial biosynthetic pathways, the mainmechanism for the control of enzymatic activity is at the level oftranscription. Genes or gene clusters belonging to the cysteineregulon, with the exception of cysE, are under the positive control ofthe LysR-type regulator CysB [5,32,58] that also represses its ownsynthesis [5]. LysR-type regulators require inducers and anti-inducers tofulfill their function. In the case of the cysteine regulon, N-acetylserine(NAS) acts as an inducer of the four transcriptional units. OAS, the finalproduct of the “glycolysis branch” of RSAP, is an unstable moleculethat generates NAS by O- to N-acetyl migration at basic pH [24,59]. Theaccumulation of NAS inside the cell signals a deficiency in the sulfursupply, ultimately sulfide, which is needed in the final step of cysteinebiosynthesis. Sulfate and thiosulfate are believed to act as anti-inducers of the cysteine regulon [1].

The activity of SAT is not under the positive, transcriptional controlof CysB but is mainly regulated by the feedback inhibition exerted bycysteine [14,60–63]. Cysteine accumulation leads to the inhibition ofSAT activity, which causes a depletion of OAS. The final effect of thisfeedback mechanism is the arrest of the highly energy demandingsynthesis of sulfide by sulfate reduction. The inhibitory effect ofcysteine on OASS activity might be non-physiologic because of thehigh measured inhibition constant [5], whereas for sulfide the Ki isabout 25 μM.

Perhaps the most interesting mechanism of regulation of cysteineproduction is the formation of protein complexes. In particular, OASShas been demonstrated, in different organisms, to be able to interactspecifically with various proteins of the RSAP. For example, in Bacillussubtilis, OASS forms an active complex with the transcriptional

repressor CymR, stabilizing its interaction with DNA [64]. This resultsin the repression of sulfate reduction that is associated with a highintracellular concentration of cysteine. Interestingly, in B. subtilisOASS does not interact with SAT and the formation of the CymR-OASScomplex is the basis for the ability of this organism to respond tochanges in cysteine levels inside the cell. In E. coli a more complexpicture for the regulation of RSAP by protein–protein interactions isemerging, and likely extends to other enterobacterial RSAPs [9,13–15,38,62,65–68]. In fact, OASS is the central player in a high ordermulti-protein complex composed, based on our current knowledge, ofOASS itself, SAT and ATP sulfurylase (ATPS). In the proposedregulatory mechanism, OASS interacts with ATPS and activates theenzyme for the forward, thermodynamically unfavorable reaction,i.e. adenosine 5′-phosphosulphate (APS) synthesis. On the other hand,OASS is also involved in the formation of CS, a tight complex (Kdiss

about 1 nM in E. coli and H. influenzae) where the active site of OASS isthe anchor point for the insertion of SAT C-terminal sequence, andthus for complex formation [68]. The complex is stabilized by sulfideand dissociated by direct competition between SAT and OAS, with adecreased OASS activity in the complex [14,67,69–71]. Thus, amultienzyme complex, possibly comprising even more proteins ofRSAP, is at the basis for the fine tuning of cysteine biosynthesis inprokaryotes, as already proposed in plants [16,72].

2. Spectroscopic properties

2.1. Absorption spectra

The absorption spectrum of OASS-A and -B is typical of a PLP-containing enzyme and is characterized by a band at 412 and 414 nm,respectively (Fig. 2A), that is pH-independent in the range of proteinstability [19]. This band is attributed to the protonated ketoenamine


species of the internal aldimine (IA) and indicates that the iminenitrogen of the cofactor is protonated and hydrogen bonded to the 3′-oxygen atom of PLP. The ratio of the intensity at 280 and either412 nm for the A-isozyme or 414 nm for the B-isozyme is 3.4–3.6 [19]and 4.0–4.2 [34], respectively. This difference is well accounted by thepresence of two tryptophan residues in the A-isozyme, W50 andW161, and three tryptophan residues,W28,W159, andW212, in the Bisoform.

In the presence of the substrate OAS, the band shifts to 470 nmdue to the formation of the ketoenamine of the α-aminoacrylateintermediate (AA) [11,19,73]. Concomitantly, a band at 330 nmappears, indicative that the reaction mixture also contains theenolimine tautomer of AA. The Kdiss of OAS at pH 6.9 is about 3 μMand increases an order of magnitude per pH unit [19]. When OASS isexposed to the product analog, L-serine, the 412 nm band shifts to418 nm for the A-isozyme with an intensity decrease and to 427 nmfor the B-isozyme with no intensity change [34,74]. In the case of theA-isozyme a new band appears at 325 nm, likely the enoliminetautomer of the external aldimine (EA). In the presence of the productL-cysteine, similar spectral changes occur in both isozymes. Theenzyme affinities for L-serine and L-cysteine are pH-dependent, andincrease with increasing pH. For OASS-B pKa values of 7.7 and 6.9were determined for the group controlling the strength of L-serineand L-cysteine binding, respectively. The pH-independent values ofKdiss, above pH 8.0, for L-serine and L-cysteine are about 30 and 3 mM,respectively [34]. For OASS-A the dependence of affinity on pHexhibits a bell-shape curve with slopes of 2 and −1 yielding anaverage pKa of 7.4 for the low pH range and a pKa of 9.6 at high pH. ThepH independent value of L-cysteine dissociation constant is 120 μM[23]. A similar analysis, carried out for L-serine, gave a limiting valueof +2with pKa values of 7.6 and 8.4. The pH-independent value of theL-serine dissociation constant is 4.2 mM [23] (see also below).

2.2. Steady-state and time-resolved fluorescence

The fluorescence spectrum of OASS with excitation at 298 nm ischaracterized by a band at about 340 nm due to the emission oftryptophan residues (Fig. 2B) [22,75,76]. Another emission band ispresent at 500 nm, less intense for OASS-B than for OASS-A [34,74].This band originates from an energy transfer between the tryptophanemission and the IA enolimine absorption band. The tryptophanresidues and the PLP ring in OASS-A and -B are within distances thatallow this process to take place (Fig. 3). An excited state protontransfer converts the enolimine to ketoenamine that emits at 500 nm.This mechanism is supported by the following evidence [76]: i) theapo-enzyme does not exhibit an emission at 500 nm, ii) the sameemission at 500 nm is observed with excitation at 412 nm, and iii) thedirect excitation at 330 nm leads to emission bands at 400 and500 nm. The distinct contribution to the emission of tryptophanresidues of OASS-B (Fig. 3B) was investigated by characterizingemission properties of single and double tryptophan mutants [74].Residue W28 was found to be mainly responsible for directtryptophan fluorescence emission, whereas W212, located in a highlyflexible region near the active site, is mainly responsible for theemission at 500 nm. W159 contributes slightly to direct emission andenergy transfer to PLP. Comparison of the emission properties ofOASS-A and -B indicates that the OASS-B active site is likely to bemorepolar and flexible than OASS-A, in agreement with a broader substratespecificity and higher catalytic efficiency.

In the presence of either L-serine, L-cysteine or acetate, at high pH,the fluorescence emission band is centered at 490 nm and issignificantly increased with respect to the emission of IA [22,74,75].This behavior is explained on the basis of a ligand-induced closure ofthe active site that causes an increased apolarity and decreased PLPmobility. The formation of AA leads to an emission that is significantlylower than that observed for EA. Upon excitation at 330 nm, the

emission exhibits two bands centered at 420 and 540 nm, whereasupon excitation at 420 nm the emission bands are centered at 500 and540 nm, and upon excitation at 470 nm, the main absorbance peak ofAA, the emission spectrum shows a band at 540 nm. Not surprisinglygiven the distances, the tryptophan emission band is not significantlyaffected by occupancy of the active site.

To probe the active site conformation along the catalytic pathway,time-resolved fluorescence measurements of OASS-A upon excitationat 420 nm were carried and analyzed [28,75]. For IA, the emissiondecays were characterized by two lifetimes of about 0.6 and 3.8 nswith almost equal fractional amplitude [75]. The emission decays ofthe EA in the presence of either L-serine or L-cysteine were best fittedusing two components with lifetimes of about 1.1 and 3.8 ns, with thefractional intensity of the slow component being 0.92 with L-cysteineand 0.75with L-serine, respectively [75]. The fast component, emittingat 530 nm, is attributed to a dipolar species formed in the excitedstate by proton dissociation, and the slow component, emitting at490 nm, is attributed to a ketoenamine tautomer of the EA. The slowcomponent for the EA fluorescence decay is characterized by the samelifetime value as that of IA with an increased fractional intensity,indicating that the distribution between the ketoenamine and thedipolar species is shifted toward the ketoenamine tautomer in EA,compared to IA. Differences in the equilibrium distribution of theketoenamine and enolimine tautomers well account for differences inthe emission properties of EAs of L-cysteine and L-serine.

The fluorescence decay of AA, upon excitation at 330 nm, is bestfitted using three components with lifetime values similar to thosefound for the IA, with the slow component predominating [75]. Theweak intensity and the short lifetime of the emission with excitationat 465 nm indicate that the AA enolimine tautomer interactssignificantly with neighboring groups of the protein matrix and mayhave a higher mobility than EA.

Subtle conformational differences between OASS-A and -B weredetected by comparing the absorption spectra and steady-state andtime-resolved emission of the IA in the absence and presence of theproduct acetate and of the EA with L-serine [34,74]. The two isozymesshow a different equilibrium distribution of the enolimine andketoenamine tautomers, likely as a result of a more polar active sitefor OASS-B. The tautomeric distribution is affected by the occupancyof the enzyme active site. In the presence of acetate, which binds to theα-carboxylate subsite, the population of the ketoenamine tautomerincreases due to a conformational change. This finding is in agreementwith structural data and suggests a higher degree of conformationalflexibility for OASS-B with respect to OASS-A.

The tryptophan phosphorescence properties of wild type OASS-Aand single tryptophan mutants were characterized over the temper-ature range 170–273 K [77]. The comparison of apo- and holo-OASSphosphorescence indicates that the coenzyme causes about 70%reduction in emission intensity and lifetime. The phosphorescencelifetime in fluid medium reveals conformational heterogeneity inOASS-A that is affected by cofactor, substrates, and pH. In particular,coenzyme binding decreases protein flexibility around W161,whereas OAS and L-serine exhibit an opposite effect.

3. Structural characterization

OASS is a homodimer formed by two identical subunits (Fig. 4),with a buried surface area at the interface of 1350 Å2 (PDB ID: 1oas,[26]). The subunits thatmake up the dimer are formed by a large and asmall domain. The large domain includes an internal β-sheet formedby five parallel (β2, β7–10) and one antiparallel (β1) β-strands,surrounded on both sides by three (α5–7) and two (α8, α9) α-helices, respectively. Similarly, the smaller domain contains aninternal β-sheet (β3–6) contoured by four α-helices (α1–4) [34].The active site is located in the protein core at the interface betweenthe two domains, close to the PLP bound to K41. As observed for OASS-

Wavelength (nm)350 400 450 500 550O

AS

S-A

rel

ativ

e flu

ores

cenc

e in

tens

ity

0.0

0.1

0.2

0.3

OA

SS

-B relative fluorescence intensity0.0

0.2

0.4

0.6

0.8

1.0

B

Wavelength (nm)300 400 500 600

Abs

orba

nce

0.0

0.1

0.2

0.3

0.4

0.5

0.6A

Fig. 2. Absorbance spectra (A) and fluorescence emission spectra for excitation at298 nm (B) of OASS-A (solid line) and OASS-B (dashed line).

PLP

W212

W28

W159

21 Å

24 Å

12 Å

A

B

PLP

W161 W50

20 Å24 Å

Fig. 3. Location of the tryptophan residues in S. typhimurium OASS-A (A; PDB ID: 1oas[26]) and OASS-B (B; PDB ID: 2jc3 [34]). Tryptophan residues and the PLP cofactor arehighlighted in black-capped sticks.


A from S. typhimurium [26], the cofactor is strongly anchored throughits dianionic phosphate group to the proteinmatrix by eight hydrogenbonds, six from G176, T177, G178 and T180 in the PLP loop and twofrom water molecules. Moreover, the N1 nitrogen of the coenzymepyridine ring is within hydrogen bonding distance to the hydroxyloxygen of S272, while the 3′-phenolic oxygen of the cofactor interactswith the side-chain amide nitrogen of N71.

The structure of OASS-B from S. typhimurium in the openconformation has been recently solved (PDB ID: 2jc3, [34]). Asshown in Fig. 4, OASS-B exhibits a fold very similar to the A isoform,presenting a large and a small domain containing PLP at the domaininterface. Moreover, as already observed for OASS-A, the proteincontains a sub-domain, including residues 86–130, and is able toundergo a similar conformational rearrangement upon the ligandbinding. Nevertheless, comparison of the catalytic intermediates andthe spectrofluorometric properties of the isozymes exhibited thepresence of a diverse equilibrium distribution between the enolimineand the ketoenamine tautomers. This is possibly related to a higherflexibility of the B isozyme, as also suggested by the highercrystallographic B-factors observed for OASS-B with respect toOASS-A [34]. The PLP phosphate group is anchored to the proteinmatrix via an extended hydrogen bond network mainly formed withthe phosphate-binding loop including residues 174–178. Comparisonof the active sites of OASS-A (Fig. 5A) and OASS-B (Fig. 5B) reveals thepresence of a well conserved region belonging to the small domain,and substitution of numerous residues on the larger domain side ofthe site. Thus, while T68, S69, G70, N71 and T72 exhibit similarpositions, the residues on the large domain have different locations,giving a different conformation of the loop containing these residues.In particular, the presence of R211 and W212 in the B isozyme makesthe binding pocket more polar and apparently less accessible

compared to the A isozyme. The different orientation of M119 alsocontributes to the closing of the active site, which is only accessible viaa narrow channel.

The IA structure of OASS-A exhibits an open conformation of thebinding site, easily accessible to incoming ligands. In contrast, thestructure of the mutant OASS-A where K41 has been replaced by analanine residue and L-methionine is bound to PLP forming an EA,exhibits a closed conformation (Fig. 6) (PDB ID: 1d6s, [83]). Thesuperposition of the two structures shows a significant conformationalchangemainly affecting the substrate-binding loop (Fig. 7). It has beensuggested that the conformational change is mainly triggered by theinteraction of the substrate carboxylic moiety with the substrate-binding loop (residues 67–71), highly conserved from bacteria toplants [84]. The closure of the binding pocket stabilizes the EA,contributing to the proper positioning of substrate-binding groups andexcluding water molecules from the active site. These effects lead to astabilization of the AA, favorably orienting the protein residues as wellas the substrate for the elimination reaction [83]. In particular, thebinding loop closes to properly localize S69, which moves about 7 Å,and forms two new hydrogen bonds with the carboxylate group of thesubstrate and with the phenolic oxygen of the PLP cofactor. Thecofactor also tilts by about 13°. This local and apparently restrictedrearrangement induces a significantly larger conformational changeaffecting a sub-domain (residues 87–131) formed by β-strand 4, α-helix 3, β-strand 5 andα-helix 4, whichmoves as a rigid body towards

Large domain

Small domain

Fig. 4. Superposition of the crystallographic structures of OASS-A (light gray; PDB ID:1oas [26]) and OASS-B from S. typhimurium (dark gray; PDB ID: 2jc3 [34]). The PLPcofactor is shown in black-capped sticks.

A

I229

M119

PLP

K41

T72

S69

T68

Q227

G228

Q142G230A231

G232

F233

S272N71

B

M119

PLP

K41P207

T72

T68

S69

W212R211

R210

I209

G208

S255

Q140

N71

Fig. 5. Comparison of the binding pocket of OASS-A (A, PDB ID: 1oas [26]) and OASS-B(B, PDB ID: 2jc3 [34]). The PLP cofactor and the more significant residues lining the activesite and likely interacting with the incoming ligands and substrates are represented inblack-capped sticks.

S69

N71

PLP

S272

G228

I229

G230

A231

G232

F233

M119

Q142

T68

T72

Q227

MET

A41

Fig. 6. Close-up view of methionine external aldimine of S. typhimuriumOASS-AmutantK41A (PDB ID: 1d6s [83]).


the active site by a rotation of 7°. Thus, residues far apart in the openconformation are brought into close proximity as the sub-domainmoves and new hydrophobic and electrostatic interactions are formedbetween the N-terminal and the C-terminal domains. Overall, thismovement reduces the severe twist of the central β-sheet, anddecreases the size of the binding pocket entrance, which can now beoccupied only by small molecules, such as the second substrate sulfideor the first product acetate. Moreover, the C-terminal residues 316–322, disordered in the open state, assume a more fixed position as thebinding pocket closes, packing against the sub-domain formed byresidues 87–131 in the corresponding sub-domain of the othermonomer in the dimeric structure [83]. Nevertheless, and in contrastto what has been observed for other PLP-dependent enzymes, e.g., theβ-subunit of tryptophan synthase, these conformational adjustmentsthat occur as catalysis takes place are not affected by the crystalpacking since catalysis proceeds without cracking the crystals [26],although not all OASS crystals are fully catalytically competent [27].The hypothesis that the catalytic process is assisted by a conforma-tional adjustment associated with the transition from an open to aclosed state is further supported by the observation that AA [75–77,85]and the enzyme that is allosterically inhibited [86] by a chloride anion(PDB ID: 1fcj, [87]) exhibit slightly different conformations. By singlecrystal polarized absorption microspectrophotometry [88,89], it wasdemonstrated that the catalytic competence of OASS towards OASdepends on the crystallization conditions, indicating that lattice forcesprevent catalytically relevant conformational transitions [27]. On thecontrary, the reaction of the crystalline enzyme with either L-cysteineor L-serine led to the formation of the EA with dissociation constantsand pH dependence close to those observed in solution [27].

4. Thermodynamic stability and unfolding mechanism

The observation of a discrete number of common catalyticmechanisms, sequences and structural motives allows the classificationof PLP-dependent enzymes into functional groups or structural foldtypes [78,79]. Members of this large super-family often belong to thesame group in spite of very low or marginally significant sequenceidentity/homology. For instance, OASS-A and the β2 dimer of trypto-phan synthase from S. typhimurium belong to theβ-family and fold typeII, in spite of a sequence identity of only about 20%. This raised interest in

trying to compare the folding mechanism of PLP-dependent enzymesbelonging to the same or different groups, in order to identify specificstructure and function determinants encoded in the primary sequence.An additional point of interest is the role of the coenzyme in structuralstabilization and in the control of the folding/unfolding mechanism. Todate, this kind of investigation has been pursued with some depth for alimited number of PLP-dependent proteins, compared to the hugeamount of structural and functional information gathered for manymembers of the PLP family.

Fig. 7. Superposition of S. typhimurium OASS-A internal aldimine in the openconformation (light gray; PDB ID: 1oas [26]) and the methionine external aldimine inthe closed conformation (dark gray; PDB ID: 1d6s [83]). Closure of the sub-domainlocated within the small domain is highlighted by the arrow. The PLP cofactor is shownin black-capped sticks.


A detailed investigation of the unfolding mechanism of holo- andapo-OASS-A from S. typhimurium was carried out with a variety ofspectroscopic techniques: absorption, circular dichroism, steady stateand time-resolved fluorescence, 31P nuclear magnetic resonance, andphoton correlation [80,81]. The effect ofmutation of the two tryptophanresidues of OASS-A on structure, stability and enzymatic function wasalso explored, being both residues partially exposed on the surfaceof the protein (Fig. 3A) [82]. Homologous studies of the B isozyme ofS. typhimurium OASS are currently ongoing in our laboratories.

4.1. Unfolding mechanism and structural role of PLP in OASS-A

Equilibration of holo-OASS solutions with increasing concentra-tions of the denaturant guanidinium hydrochloride (GdnHCl) causesi) a red-shift and an increased intensity of tryptophan directfluorescence emission upon excitation at 298 nm, and ii) the loss ofthe emission at 500 nm attributed to energy transfer to the coenzyme[81]. These observations are consistent with an increased exposure tosolvent of tryptophans and loss of the energy transfer to PLP. AtGdnHCl concentrations higher than about 2.0 M, the emission spectraof holo-OASS become very similar to the apo-form of the enzyme. Thissuggests that the coenzyme is released at denaturant concentrationshigher than 2 M, in agreement with the blue shift of the IA absorptionpeak from 412 nm to 392 nm, a wavelength typical of free PLP, andwith the results of 31P NMR [81], time-resolved fluorescence, andfluorescence anisotropy of both tryptophan and PLP [80].

Fitting of the denaturant-dependence of tryptophan fluorescenceemission to a two-state unfolding process allowed an estimation of thethermodynamicparameters for equilibriumunfolding [81]. The transitionmidpoints and the unfolding free energies are 1.57±0.15 MGdnHCl and2.72±0.40 kcal/mol, and0.47±0.02 MGdnHCl and1.47±0.11 kcal/molfor holo- and apo-OASS, respectively. The equilibrium unfolding curvescalculated from circular dichroism spectroscopy (from the denaturantdependence of the ellipticity at 222 nm) yield similar parameters for apo-OASS, but indicate a somewhat lower stability for theholo-enzyme. Threerelevant conclusions can be drawn from these results: i) the thermody-namic stability of OASS is relatively low, about 2.7 kcal/mol, a result thatis only apparently surprising taking into account thatmarginal stability isoften a necessary counterpart of dynamic flexibility required forenzyme function and regulation; ii) the coenzyme elicits a significantstabilizing effect onOASS structure; and iii) the different unfolding curvescalculated from fluorescence and circular dichroism data, that are

sensitive to the tryptophan microenvironment and to the global contentof secondary structure elements, respectively, suggest that the twodomains of the OASS protomer are differently affected by the stabilizinginteractions of the coenzyme. The latter observation was furtherinvestigated by time-resolved fluorescence studies [80].

The tryptophan residues of S. typhimurium OASS-A are in helicalregions of the N-terminal (W50) and C-terminal domains (W161)(Fig. 3A). Both tryptophans are located about 20 Å away from the PLP,a distance consistent with a significant Förster resonance energytransfer to the coenzyme. Due to the different orientation of thetransition dipoles, the energy transfer predominantly occurs betweenW50 and the PLP [76]. Different from steady-state fluorescence, time-resolved fluorescence data collected by phase and modulationtechnique allowed resolution of the individual contribution oftryptophan residues to the complex emission decay, and investigationof the differential stabilizing effect elicited by bound PLP on regionsbelonging to the N- or C-terminal domains [80]. Results demonstratethat, although PLP is covalently bound to K41 in the N-terminaldomain, it stabilizes the C-terminal domain to a higher extent in theholo-enzyme. This effect is likely mediated by the network ofhydrogen bonds between the phosphate group of PLP, the N-terminusof helix 7 and some residues (176–178, 180) of the C-terminal domain[26]. Consistently, no significant difference in the unfolding behaviorof the two domains, that are endowedwith a high degree of structuralhomology [26] (Fig. 7), is observed in the apo-enzyme. The differencein stability of the two domains of the holo-OASS protomer is likely tohave functional significance, since a higher structural plasticity isrequired for the large movement of the subdomain containingresidues 87–131 (in the N-terminal domain) that occurs upon formationof the EA [83] (Fig. 7).

Dynamic light scattering experiments carried out to address whendissociation of the dimer to monomers occurs along the unfoldingpathway [81], indicated that this is a late event in OASS denaturation,and must be preceded by the cleavage of the covalent bond betweenPLP and K41, and by extensive disruption of the tertiary andsecondary structure. This is likely a consequence of the strength ofinter-monomer interactions in OASS, and explains the observed two-state unfolding behavior of such a complexmolecule. No intermediatespecies having spectroscopic properties different from the nativedimer or the unfolded monomer are populated at equilibrium.

Interestingly, even in the absence of denaturant, circular dichroismspectroscopy highlighted differences in the secondary structurecontent of holo- and apo-OASS [81], indicating that in addition to aglobally stabilizing effect, the presence of bound PLP is necessary toachieve the native fold of the enzyme. The role of the coenzyme inassisting proper folding of the native structure is not consistentlyobserved in all PLP-dependent enzymes investigated, and appears notto be related to the fold type ([81] and references therein).

4.2. Role of tryptophans in OASS-A structure, stability and function

Tryptophan residues are often highly conserved in proteinsequence and play essential structural and functional roles, includingstructural stabilization, inter-protein interactions, and ligand binding[82]. We have investigated the effect of W50Y and W161Y mutationsin OASS-A, with the aim of singling out the role of each tryptophan onstructure, function and thermodynamic stability, and to obtainseparate probes of structure and dynamics of the N- and C-terminaldomain [82]. W161 is highly conserved in eukaryotes and bacteria(63% of 62 analyzed sequences), while W50 is not.

Neither mutation significantly alters the OASS-A secondarystructure, as indicated by circular dichroism spectroscopy in the far-UV region, or formation of the IA, as shown by absorbance spectra.However, a slightly higher population of the enolimine tautomer wasobserved in the W161Y mutant, suggesting some modest alterationof the active site environment. The perturbation of the coenzyme


environment is confirmed by acrylamide fluorescence quenchingdata, indicating a reduced accessibility to the active site, intermediatebetween the open and closed conformation of the wild type protein(see paragraph 3 on Structural characterization). Data suggest thatmutation of W161 might induce a partial closure of the active sitecleft, in agreement with the increased thermodynamic stability of theN-terminal domain, likely due to stronger inter-domain interactionswith respect to the wild type IA (W161 makes no direct interactionswith the N-terminal domain). This observation, together with thehigh conservation of W161 and its location in a flat, surface region ofthe protein, points to a possible involvement of this residue in theinteraction surface of OASS with SAT. Indeed, the proposed mecha-nism underlying the regulatory effect of SAT on OASS activity involvesthe closure of the active site of OASS upon binding of the SAT C-terminalpeptide [13]. Notably, the increase in the Km for OAS observed inthe W161Y mutant is comparable to that induced by the interactionwith SAT.

Fluorescence emission spectra indicate that the W50Y mutationcauses an increase in the energy transfer efficiency fromW161 to PLPthat is normally minor in the wild type enzyme. This could be aconsequence of a mutation-induced rearrangement of the position ofhelix 1, bearing bothW50 and the Schiff base K41, causing a change inthe distance or relative orientation of PLP and W161. The observedchanges in the catalytic activity, mainly affecting the second halfreaction, are actually consistent with changes in the orientation of theAA species. Surprisingly, the W50Y mutation dramatically affects theunfolding pathway of OASS by strongly destabilizing the subunitinterface, as shown by a coupling of protein denaturation and subunitdissociation that is not observed in the wild type or the W161Ymutant enzymes, a remarkable result for mutation of a single, surface-exposed, non-conserved residue.

5. Kinetic mechanism

The kinetic mechanism of OASS-A and -B from S. typhimuriumwasstudied using a combination of initial rate studies, stopped-flowkinetic studies and isotope effects. Initial velocity data are consistentwith a ping pong kinetic mechanism with competitive substrateinhibition by OAS and SH− [11,90]. OAS binds to the IA and isconverted to the AA with the release of acetate. This is followed bybinding of bisulfide, which is converted to cysteine with regenerationof the IA. Bisulfide exhibits strong competitive substrate inhibition(KI SH=50 μM; KSH=6 μM) [90]. The inhibition was originally thoughtto result from binding of bisulfide to free enzyme, competing withOAS [11,90]. However, a structure with chloride, a bisulfide analog,bound to an allosteric site suggested allosteric substrate inhibition bybisulfide [86,87].

The kinetic mechanism is supported by product and dead-endinhibition studies. Cysteine was noncompetitive vs. OAS and SH−

indicating it binds to the IA and AA forms of the enzyme. Of interest,product inhibition by acetate gives S-parabolic noncompetitiveinhibition vs. OAS with TNB as the nucleophilic substrate. Theparabolic slope effect results from acetate binding to the AA andreversing the reaction to generate the IA to which acetate also binds[90]. Dead-end inhibition by thiocyanate is competitive vs. SH−, anduncompetitive vs. OAS. The turnover number for OASS-A is 130 s−1.

In a ping pong mechanism, the two half reactions are independentof one another and thus the second order rate constant for one half ofthe reaction should be independent of the other reactant used. This isnot observed for OASS-A. V/KOASEt is 1×105 M−1 s−1 with SH− as thenucleophilic substrate, but only 37 M−1 s−1 with 5-thio-2-nitrobenzoate(TNB) as the substrate [90]. In addition, with β-chloro-L-alanine (BCA) asthe amino acid substrate, V/KBCAEt is 2 M−1 s−1 with SH− as thesubstrate, but 0.016 M−1 s−1 with TNB as the substrate. On the otherhand, the V/KSHEt and V/KTNBEt are independent of the amino acidsubstrate. Data suggest a significant OAS:lyase activity, with conversion of

the AA to pyruvate and ammonia, with TNB being slower and much lessefficient compared to SH− [19].

The overall kinetic mechanism can be depicted as shown inScheme 1.

5.1. Location of slow steps

The first half of the OASS reaction, conversion of OAS and the IA tothe AA and acetate, is the slowest for the overall reaction. Pre-steadystate experiments carried out by rapidly mixing OAS and enzymeresulted in a decrease in the absorbance of the IA at 412 nm and aconcomitant appearance of the AA at 470 nm; no other intermediateswere observed [91]. The maximum rate of formation of the AA, kmax,is 300 s−1, similar to V/Et obtained from steady state studies. Thedependence of the first order rate constant for the AA formation onOAS concentration gives a KEA similar to the Km for OAS. Data areconsistent with the elimination of acetate as the major contributor torate limitation for the overall reaction. In agreement, the primarydeuterium isotope effect on V/KOAS is 1.7 at neutral pH, equal to theisotope effect on V. The effect on V/KOAS increases to 2.8 at pH 5.5, andthis is equal to the isotope effect on kmax, which reflects the formationof the AA from the OAS-EA complex. The isotope effect data allow anestimation of the stickiness factor for OAS, cf=1.5, indicating that theoff-rate constant for OAS from EA is 1.5 times faster than conversion ofthe EA to the AA. The cf term can be divided into cf in, kmax/kMichaelis,which is ~0, and cf ex, kmaxkEA/kMichaeliskoff OAS, which is 1.5. The rateconstants are defined as follows: kmax is as defined above, kEA is therate constant for formation of the EA from the Michaelis complex,kMichaelis is the rate constant for formation of the Michaelis complexfrom the EA, and koff OAS is the rate constant for dissociation of OASfrom the Michaelis complex.

The V/KbisulfideEt is 2×107 M−1 s−1, close to the diffusion limit [90].Rapidly mixing the AA with bisulfide results in formation of E and L-cysteine in the mixing time of the instrument, even at low μMconcentrations of bisulfide, suggesting all rate processes within thesecond half reaction have rate constants ≥1000 s−1 [91]. A carefulstudy of the kinetics of the second half reaction using a number ofnucleophilic substrate analogs showed that the slow step is release ofcysteine from the cysteine EA [92]. In fact, carrying out the second halfreaction in D2O at low pD gives a stable EA.

Rapidmixingexperimentswith theproduct of the reaction, L-cysteine,gave a rapid transient formation of the AA with a kmax of 110 s−1; thiswas followed by a more rapid formation (kESB~1000 s−1) of an ESB. TheEA is formed as a result of attack of a cysteine thiolate on C3 of the AA togive the lanthionine EA [91].

5.2. Acid–base chemical mechanism

The pH dependence of kinetic parameters provided information onthe optimum protonation state of ionizable functional groups onreactant and enzyme, important for binding and/or catalysis. Themostaccurate information was obtained using BCA and TNB as substrates,since OAS is unstable at pH values above 7, undergoing an O to Nmigration to give N-acetyl-L-serine [73]. The V/K for BCA decreases atlow and high pH giving pKa values of 6.7 and 7.4. The pKa of 7.4 reflectstheα-amine of BCA, while that of 6.7 reflects an enzyme group. Theα-amine must be neutral for nucleophilic attack on C4′ of the IA, andthus the two groups have reverse protonation states, i.e., the groupwith a pKa of 6.7must be protonated, while that with a pKa of 7.4mustbe unprotonated for optimum activity. The pH dependence of V/KTNB

representing the second half reaction reflects two pKa values of 6.5and 8.2. The pKa of 8.2 is also observed in the pH dependence of theOAS:acetate lyase activity, and reflects the side chain of K41, whichforms the IA with PLP, and must be protonated in the second halfreaction to donate a proton to Cα as the product EA is formed. Thegroupwith a pKa of 6.5 is likely the same as that observed in the V/KBCA

Scheme 1. Proposed kinetic mechanism of OASS-A. The binding of SH− to an allosteric site results in inactive enzyme, and SH− is competitive with OAS. The process represented byk13 is the OAS:lyase activity. The mechanism is otherwise a typical ping pong reaction (AA represents the α-aminoacrylate species).


pH-rate profile. A reasonable mechanism incorporating the datadiscussed above is provided in Scheme 2 below.

The pH dependence of the Kd for the L-serine [85] and L-methionine [83] EAs is in agreement with the above data. The affinityfor both reactants decreases as the pH decreases exhibiting therequirement for two groups that must be unprotonated for optimumbinding. In the case of serine, estimated pKa values are 7.6 and 8.4,while they are 6.8 and 8.3 for methionine; pH independent Kd valuesfor the two amino acids are 4 mM and 78 μM, respectively. Datasuggested the α-amine of the amino acid and an enzyme group mustbe unprotonated for optimum binding. The pH dependence of the Kd

for L-cysteine EA exhibited the requirement for two groups unproto-nated (the average value was 7.4) and one protonated (9.6) foroptimum binding [85]. By analogy to data obtained for serine andmethionine, the α-amine of cysteine and an enzyme group must beunprotonated, while the side chain thiol must be protonated foroptimumbinding; reverse protonation states are observed for the sidechain thiol and the α-amine pKa.

Data can be interpreted in terms of theminimalmechanism shownin Scheme 2. The pKa of 6.5–7.5, observed in all profiles, is for anenzyme group that must be unprotonated for optimum activity/binding. A pKa of 7.2 was measured from the pH dependence of theenhancement of the long wavelength fluorescence that occurs uponbinding of acetate to the IA such that it occupies the amino acid α-carboxylate subsite [22]. The authors suggested the conformationalchange that was induced by acetate binding was pH dependent, andresulted in closing or partially closing the active site. This is likely thesame pH dependence observed in V/K pH-rate profiles and pKi profilesdiscussed above. Thus, in Scheme 2, the active form of the enzyme inthe first half reaction is that with the enzyme group unprotonated. Inaddition, the α-amine of the amino acid must be neutral for optimumbinding and activity (attack on C4′) as seen in the V/KOAS pH-rateprofile and the pKi profiles for serine, methionine and cysteine. In thesecond half reaction, the unprotonated form of the enzyme group isagain required to generate the active conformation, and K41, whichparticipated in the IA, must be protonated. Second order rateconstants for nucleophilic substrates with a pKaN7 are pH dependent,while those with a pKab7 are pH independent, consistent with theionized form as the active reactant [92]. In other words, the reactantmust be a nucleophile to act as a substrate. Thiols, including H2S,which have a pKa value above 7, must be unprotonated to act as anucleophile.

A quinonoid intermediate (Q) is observed transiently in thetryptophan synthase reaction [93]. One might anticipate this interme-diate in the OASS-A reaction, but attempts to observe this species havebeenunsuccessful. However, acetate is a good leaving group and requiresno proton transfer as acetate is formed. The structure of the nativeenzyme exhibits N1 of PLP within hydrogen bonding distance to S272,such that the serine likely donates a hydrogen bond to N1 (Fig. 5A). As aresult, a quinonoid could exist only transiently as the α-proton isabstracted in an E1 mechanism, as serine donates a proton, perhaps, in alow barrier hydrogen bond.

In the direction of the AA formation fromOAS, abstraction of theα-proton is the slowest step along the reaction pathway. As a result, onemight argue that, if Q was present, it would not be seen because itwould not build up in the direction of the AA formation. Tai and Cook[94] measured rate of formation of the OAS EA beginning with the AAand acetate in H2O and D2O; a solvent deuterium kinetic isotope effectof 2.5 was measured. The slow step in this reaction direction would beprotonation of Cα after Q formation, and it should be observed ifformed, especially in the presence of D2O, which slows protonation ofCα by at least 2.5 fold. However, not even transient formation of Qwasobserved in H2O or D2O, and, thus, the reaction appears to be aconcerted α,β-elimination.

5.3. Site-directed mutagenesis

Site-directed mutagenesis was used to probe the mechanismproposed in Scheme 2. In the tryptophan synthase reaction, convert-ing S377, which is within hydrogen bond distance to N1 of PLP, to D orE resulted in buildup of Q. The intermediate was thus presumablystabilized as the carboxylate forms a stronger hydrogen bondwith theprotonated N1 of the cofactor [95]. Changing S272 in OASS-A to D or Edoes not result in buildup of Q [96]. The proximity of the N1 ofpyridine to the positive dipole of helix 10 in OASS-A likely preventsdevelopment of a positive charge on N1. In the S272D, E mutantenzymes, V/KOAS is identical to that measured for wild type but kmax

and the KEA for OAS are both decreased to maintain the same secondorder rate constant. Data suggest nonproductive binding of OAS, i.e.,positioning of the EA once formed is such that only a small percentageis competent for the elimination reaction. A significant decrease in thevisible circular dichroism molar ellipticity of the EA, and a decrease inthe long wavelength fluorescence of the mutant enzymes areconsistent with a change in the position of the pyridine ring of theEA. In addition, S377 in the β-subunit of tryptophan synthase is nothomologous to S272 of OASS-A [95]. It is not in the vicinity of a helixdipole, and would allow a positive charge on N1 of the pyridine ring.

Converting K41 to A has a number of consequences [25]. Thestructure of the K41Amutant enzyme showsmethionine in EA linkagewith the active site PLP (Fig. 6). The thioether side chain, mimickingthe acetoxy leaving group of OAS, extends from the re face of PLPtoward the active site entrance, while the Cα―H proton is on the siface of PLP [83]. This can be compared to other enzymes that catalyzea β-elimination reaction, which have the Cα―H proton and theleaving group on the si face [94]. All of these catalyze the eliminationof a relatively poor leaving group, and require a proton transfercarried out by the Schiff base lysine. In addition, the apo-K41Amutantenzyme exhibits a slow formation of the EA of OAS, but proceeds nofurther along the reaction pathway. As a result, transimination ismuchmore rapid than formation of the imine from the pyridoxal formof PLP and the free amine, and PLP-dependent enzymes have evolvedto use this mechanism. Buildup of the EAwith no production of the AAis consistent with Cα―H proton abstraction being catalyzed by K41.

Scheme 2. Minimal mechanism for OASS-A. A pH dependent conformational change precedes reactant binding to generate the active enzyme. IA, EA, and AA are as defined in the text.


Formation of the EA results in closing the active site as shown inthe structure of the K41A mutant enzyme (Fig. 7) [83]. The wild typeenzyme Schiff base is rapidly reduced in the mixing time by NaBH4

[25], while the K41A mutant enzyme was resistant to reduction of itsSchiff base for days. The trigger to close the active site appears to beinteraction of the α-carboxylate of the substrate with the substratebinding loop, comprised of residues 69–71, and Q142. Recently, site-directed mutagenesis was used to change T68, S69, N71, and Q142 toA, creating four mutant enzymes [84]. All of the mutations affectclosing of active site but to different extents. No AA is observed for theT68A and Q142A mutant enzymes, but both give a much greater rateof pyruvate production compared to wild type enzyme consistentwith a decreased stability of the AA. T68 and Q142 donate a hydrogenbond to the carboxylate oxygen of the substrate EA likely properlyorientating the EA for conversion to the AA. On the other hand, the AAis observed in equilibrium with the EA for the S69A and N71A mutantenzymes; the AA is only about 10% the level found for the wild typeenzyme. Both residues donate a hydrogen bond to the EA carboxylate,and N71 is also within hydrogen bonding distance to O3′ of thecofactor. It appears that the N71 and S69 contribute to stabilizingthe EA and AA, while T68 and Q142 contribute most to stabilization ofthe AA.

5.4. Stereochemistry and transition state structure

The concerted α,β-elimination of the elements of acetic acid takesplace by abstraction of the 2S proton of the OAS EA, which is directedaway from the si face of PLP, by K41, the lysine that forms the EA[25,83,94,96]. Structural data also support the thermodynamicallyfavored anti E2 reaction, with the acetoxy group directed away fromthe re face of the cofactor. The addition of the elements of H2S acrossthe C2–C3 double bond of the AA proceeds with retention ofconfiguration as shown by Floss [97]. Addition of SH− must be tothe same face as elimination of the acetoxy group, and protonation ofCα must be at the si face of the cofactor by K41.

Primary and β-secondary kinetic deuterium and solvent kineticdeuterium isotope effects have been measured for OASS-A [98]. The

primary deuterium kinetic isotope effect measured as a function ofpH, gives a value of 2.8 once the values have been extrapolated to lowpH to eliminate substrate stickiness (see 5.1, Location of slow stepsabove). A similar value is measured on kmax in the pre-steady state anda value of 3.3±0.9 is calculated from the measured deuterium andtritium isotope effects. Thus, the value of 2.8 is the intrinsic isotopeeffect on hydride transfer, or is at least very close to it. In agreement isthe value of 2.5 measured for the solvent deuterium isotope effectfrom the direction of EA formation from the AA and acetate [94]. Thevalue of about 3 could be consistent with either an early or latetransition state for Cα―H proton abstraction, with a value of about 7–10 expected for a symmetric transition state [99]. The β-secondarykinetic deuterium isotope effect is 1.1, quite small compared to themeasured β-secondary deuterium equilibrium isotope effect of 1.8.The secondary isotope effect reflects the amount of double bondcharacter at C3 in the transition state. Thus, the transition state forconcerted α,β-elimination exhibits very little Cβ―O bond cleavage,but significant Cα―H cleavage, i.e., data are consistent with anasynchronous transition state with Cβ―O bond cleavage laggingbehind C―H bond cleavage (Scheme 3). The low degree of Cβ―Obond cleavage is likely a result of some stabilization of the developingCα carbanion by the pyridine ring.

5.5. Dynamics

Information on the dynamics of the OASS-A reaction can beobtained from a combination of spectral and structural data. One ofthe most useful techniques in this regard is 31P NMR (see [100]).The 31P chemical shift (δ) is sensitive to the tightness of binding of the5′-phosphate of PLP, and this is directly related to the conformation ofthe bound cofactor and the enzyme. For example, the IA of OASS-A hasa δ of 5.2 ppm [19], consistent with a tightly bound 5′-phosphate. Anupfield chemical shift (a lower value) of 5.2 indicates a looser boundphosphate, while a downfield shift (a higher value) indicates a tighterbound phosphate compared to the IA. Visible circular dichroism andlong wavelength fluorescence provide information on the

image of Scheme�2

Wavelength (nm)

450 500 550 600 650

Rel

ativ

e flu

ores

cenc

e in

tens

ity

0.0

0.4

0.8

1.2

1.6

2.0

[MNENI] (mM)0.0 0.2 0.4 0.6 0.8 1.0 1.2F

luor

esce

nce

inte

nsity

at 5

00 n

m

0.9

1.1

1.3

1.5

1.7

1.9

Fig. 8. Fluorescence emission spectra for excitation at 412 nm of OASS-A in the presenceof increasing concentrations of MNENI. Inset: dependence of emission intensity at500 nm on MNENI concentration. The line through data points represents the fitting toa binding isotherm.

A


conformation of the bound cofactor relative to the protein matrix andenzyme tryptophan residues, respectively.

The EA of lanthionine can be formed upon addition of cysteine tothe IA, since initially formed AA is captured by a second cysteine thiolto give the EA. Lanthionine binds relatively tightly (pH independentKd is 130 μM), likely because of its longer hydrophobic thioether sidechain; this is likely the same reason methionine is found as an EA inthe K41Amutant enzyme (pH independent Kd is 78 μM). Both of thesegive a slight downfield shift (δ=5.3 ppm) compared to the IAconsistent with a tighter binding as a result of closing the site on theEA. The EA formed with serine, however, gives 2 resonances, δ=3.5and 5.3 ppm; a weighted average value of 4.4 ppm is observed with aline width of 50.5 Hz, suggesting two species in exchange. One ofthese at 3.5 ppm is shifted upfield from the IA and suggests weakerbinding of the 5′-phosphate, while the other is equivalent to that ofthe lanthionine EA. UV–visible spectra show a 1:1 equilibriummixture of ketoeneamine and enolimine tautomers of the serine EA.Taken together with the information on the IA and lanthionine EA,data suggest a mixture of open (enolimine) and closed (ketoenamine)forms of the enzyme in equilibrium. The tighter binding of the 5′-phosphate in the IA likely results from the positioning of the cofactorin Schiff base linkage with K41, so that even though the enzyme existsin an open conformation the strain generated by the Schiff baselinkage is felt by the 5′-phosphate. The lanthionine EA causes theactive site to close on the long hydrophobic side chain, while anequilibrium mixture is observed for the shorter more flexible serineside chain. In agreement, a single resonance is observed for the AAwith a chemical shift that is significantly upfield (δ=3.95) comparedto the IA, which exhibits a narrow line width (18.5 Hz) consistentwith the 5′-phosphate tumbling with the protein [101]. As expected,the site is more mobile for the AA and the serine EA because of theirsmaller side chain.

In agreement with the above are data obtained for the K120Qmutant enzyme [102]. As the active site of OASS-A closes, K120 formsa new hydrogen bond with the carbonyl of A231 on the other side ofthe active site. As a result, K120 stabilizes the closed form of theenzyme. The K120Q mutant enzyme gives 31P NMR chemical shiftsthat are identical to those of wild type enzyme, but with narrower linewidths for the serine and lanthionine EAs. Data are consistent with anincrease in the rate of opening and closing the OASS-A active site.Overall, data suggest alternating open and closed conformations asthe reaction pathway is traversed: open (IA), closed (OAS EA), open(AA), closed (cysteine EA), open (IA).

Scheme 3. Proposed transition state structure for the concerted anti-α,β-elimination ofacetic acid from the OAS ESB. The Cβ―O bond to the leaving acetoxy group is directedtoward the viewer, while the Cα―H bond is directed away from the si face of thecofactor.

5.6. OASS-B

The mechanism and structure of OASS-B are very similar to thoseof OASS-A. There are a few differences that are worthy of note andthese will be discussed briefly below.

The kinetic mechanism proposed for the B-isozyme is qualitativelyvery similar to that obtained for the A-isozyme. However, there aretwo important differences: i) there is no dependence of the secondorder rate constants on the identity of the other substrate, and ii) there

B

PLP

I267

N266

E265N264

N72

T73

S70

T69

Q143

Fig. 9. A) Three-dimensional structure and B) close-up view of the active site of OASS-Afrom H. influenzae complexed with the MNENI peptide (PDB ID: 3iqi [54]).

image of Scheme�3

image of Fig.�8


is no allosteric inhibition by bisulfide [34,90]. Thus, the B-isozymebehaves as a prototypical Bi Bi ping pong enzyme.

The active site of the B-isozyme is more hydrophilic than that ofthe A-isozyme, largely as a result of the presence of C280 and D281 inOASS-B, which replace the neutral S300 and P299 in OASS-A (Figs. 4and 5) [34]. D281 is positioned above the re face of the cofactor withinhydrogen bonding distance to Y286, while C280 is 3.4 Å from N1 ofPLP. The more hydrophilic site has consequences. The IA and EA existalmost solely as the ketoenamine tautomer. The conformation of theB-isozyme also appears to be much more flexible than that of the A-isozyme, which allows a relaxed nucleophilic substrate specificity.

The pH dependence of kinetic parameters is very similar for bothisozymes, but the much faster B-isozyme cannot be as easily studiedin the pre-steady state [34]. Saturation by OAS of the observed firstorder rate constant for formation of the AA could not be achieved. ThepH dependence of the second order rate constant, kmax/KEA, shows therequirement for a group with pKa≥6.5 that must be unprotonated foroptimum activity and/or binding, as also found for the A-isozyme. Inaddition, the pH dependence of the KEA for lanthionine and serineexhibits a single pKa of 7.0–7.5 for a group that must be unprotonatedfor optimum binding, as opposed to results for the A-isozymediscussed above under acid–base chemical mechanism, which arequite complex. Failure to observe the pKa values for the α-amine ofcysteine or serine suggests there must be compensatory ionization ofa group on the enzyme, possibly C280. Finally, the rate of the OAS:acetate lyase activity of the B-isozyme is 40-fold higher than that ofthe A-isozyme, likely because of the increased conformationalflexibility of the enzyme. The pH dependence of the lyase activitygives a pKa of 9 for K41, a pH unit higher than that of the A-isozyme,likely a result of increased negative charge in the site (D281 andC280).

Spectral data indicate subtle differences in the overall conforma-tion of the B- compared to the A-isozyme. The biggest difference isseen in the 31′P NMR data [103]. The IA exhibits a δ of 6.2 ppm, 1 ppmfurther downfield than that observed for OASS-A, the highest δobserved for any PLP-dependent enzyme to date. Data suggest a verytightly bound 5′-phosphate that generates torsional strain about theC5–C5′ bond and a change in the O4–C5′–C5–C4 torsion angle. Thelanthionine and serine EAs are slightly upfield and downfield,respectively, of the IA with values of 6.0 and 6.3 ppm, respectively.Although it is difficult to imagine the reaction pathway dynamics ofthe B-isozyme are significantly different than those of the A-isozyme,given the identical reaction catalyzed, more work will be required onthis aspect of the mechanism.

6. Interaction of OASS-A with SAT

As stated above, OASS-A, but not OASS-B, is inhibited by itsinteraction with SAT, the preceding enzyme in the cysteine biosyn-thesis pathway. The interaction of OASS-A and SAT has beenthoroughly investigated. Yeast two-hybrid system [104] and inhibi-tion studies [66] have demonstrated that the main determinant of theformation of an OASS-SAT complex is the SAT C-terminal decapeptide.SAT lacking the 20 C-terminal amino acids does not form the complex[66]. A key residue for establishing the interaction is the last aminoacid, isoleucine, as it was found that a peptide lacking isoleucine doesnot bind to OASS-A. The Kdiss between OASS and SAT is nanomolar,whereas the Kdiss between OASS-A and either the C-terminaldecapeptide or pentapeptide of H. influenzae is 580 nM [15] and50 μM [54], respectively. These measurements were carried outexploiting the change in the fluorescence intensity of the PLP bandcentered at 500 nm, triggered by the transition from an open to aclosed conformation upon ligand binding. Stopped-flow fluorescencespectroscopy was used to characterize the fast events involved in therecognition between SAT and OASS, determining the time courses as afunction of temperature and pH, and in the presence of the

physiological regulators cysteine and bisulfide [13]. Results indicatethat OASS–SAT assembly occurs via a two-step mechanism involvingrapid formation of an encounter complex, followed by a slowconformational change. The conformational change likely resultsfrom the closure of the active site of OASS upon binding of the SAT C-terminal peptide. Bisulfide stabilizes the complex by decreasing thereverse rate of the isomerization step, while cysteine only slightlydecreases the rate of complex formation.

Complexes ofOASS-A fromH. influenzae [68], M. tuberculosis [42] andA. thaliana [105]with the corresponding C-terminal peptide of SAT in anopen conformation, have been solved, detecting the last four aminoacids in the two former structures and the last eight in the latter. Thissuggests that these residues provide the predominant contribution tothe free energy of binding, with the particular relevance of the terminalisoleucine, as above mentioned. This residue makes several hydrogenbondswithN72, T73, T79 andQ143, and other hydrophobic interactions[54]. On the basis of the structural and energetic analysis of theH. influenze OASS–SAT complex, four hundred MNXXI peptides weregenerated in silico and docked into the OASS-A binding site. The affinityof 14 selected peptides was experimentally determined (Fig. 8) and thestructures of OASS complexed with the three higher affinity pentapep-tides, MNYDI (PDB ID: 3iqh), MNENI (PDB ID: 3iqi) and MNWNI (PDBID: 3iqg)were solved [54]. The structure of OASS-A complexedwith theMNENI pentapeptide (Fig. 9) shows only the four last residues [54]. Thepeptide fully occupies thebindingpocket forcing theprotein toassumeaconformationwith the terminal isoleucine (I267) deeply buried into theprotein core and stabilized by several interactions (Fig. 9B). One oxygenof the carboxylate moiety contacts N72 and T73, while the secondinteracts also with T69 and Q143. The carboxamide group of N266 ishydrogen bonded to S70, the carboxylate side-chain of D265 directedtoward the solvent while the amino moiety of N264 interacts with thebackbone carbonyl of N264 (Fig. 9B). A pharmacophoric analysis of theactive site indicates that a tight binding is achieved placing twohydrogen bond acceptor groups, better if negatively charged, inpositions corresponding to I267 and N266, and a hydrophobic groupnext to N264 [54]. This investigation opens theway to the developmentof peptidomimetic compounds, potentially useful as antibiotics.

Acknowledgements

This work was supported by the Grayce B. Kerr endowment to theUniversity of Oklahoma to support the research of P.F.C. and by fundsfrom the Italian Ministry of University and Research to A.M.

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