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Seminars in Cell & Developmental Biology 22 (2011) 285–292

Contents lists available at ScienceDirect

Seminars in Cell & Developmental Biology

journa l homepage: www.e lsev ier .com/ locate /semcdb

eview

acterial glyoxalase enzymes

thaiwan Suttisansanee, John F. Honek ∗

epartment of Chemistry, University of Waterloo, 200 University Avenue, Waterloo, Ontario, Canada N2L 3G1

r t i c l e i n f o

rticle history:vailable online 15 February 2011

a b s t r a c t

The glyoxalase system is composed of two metalloenzymes, Glyoxalase I and Glyoxalase II. This systemis important in the detoxification of methylglyoxal, among other roles. Detailed studies have determinedthat a number of bacterial Glyoxalase I enzymes are maximally activated by Ni2+ and Co2+ ions, but are

eywords:lyoxalaseacteriascherichia colilostridiumickel

inactive in the presence of Zn2+. This is in contrast to the Glyoxalase I enzyme from humans, which iscatalytically active with Zn2+ as well as a number of other metal ions. The structure–activity relationshipsbetween these two classes of Glyoxalase I are serving as important clues to how the molecular structuresof these proteins control metal activation profiles as well as to clarify the mechanistic chemistry of thesecatalysts. In addition, the possibility of targeting inhibitors against the bacterial versus human enzyme

incetalloenzyme

has the potential to lead to new approaches to combat bacterial infections.© 2011 Elsevier Ltd. All rights reserved.

ontents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2851.1. Glyoxalase I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2851.2. Glyoxalase II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2891.3. Glyoxalase III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291

. Introduction

The glyoxalase system is composed of two enzymes con-isting of Glyoxalase I (GlxI) and Glyoxalase II (GlxII) thatonvert metabolically produced �-ketoaldehydes into their cor-esponding 2-hydroxycarboxylic acids (d-lactate in the casef methylglyoxal (pyruvaldehyde)), using an intracellular thiols a cofactor/cosubstrate [1]. The first enzyme, GlxI (S-d-actoylglutathione methylglyoxal lyase (isomerizing), EC 4.4.1.5),onverts a hemithioacetal, the product of the non-enzymatic reac-ion between cytotoxic methylglyoxal (MG) and a thiol suchs glutathione (GSH), to S-d-lactoylglutathione, which itself hasmportant cellular properties such as in the control of bacterial

bacteria (such as Streptococcus agalactiae and Enterococcus faecium)is glutathione, where its biosynthesis has been investigated [3–6].Actinomyces such as Mycobacteria, however, produce an unusualmycothiol (MSH), an acetylated cysteine attached to an inositol-linked glucosamine [6,7]. The Glyoxalase I using this type of thiol isbelieved to employ a parallel pathway to that of the system usingGSH and this cofactor is currently under investigation in our labo-ratory [8]. Other organisms such as the protozoa Trypanosoma andLeishmania have been found, in some cases, to utilize their majorintracellular thiol, trypanothione, as cofactor for their GlxI (Fair-lamb, in this issue). This article discusses some of the properties ofthe bacterial glyoxalase enzymes.

otassium efflux pumps [2]. This compound is then hydrolyzedy GlxII (S-2-hydroxyacylglutathione hydrolase, EC 3.1.2.6) to pro-uce non-toxic d-lactate and regeneration of the correspondinghiol. The major intracellular thiol found in most Gram-negativeacteria, cyanobacteria, purple bacteria and some Gram-positive

∗ Corresponding author. Tel.: +1 519 888 4567; fax: +1 519 746 0435.E-mail address: jhonek@uwaterloo.ca (J.F. Honek).

084-9521/$ – see front matter © 2011 Elsevier Ltd. All rights reserved.oi:10.1016/j.semcdb.2011.02.004

1.1. Glyoxalase I

Being a metalloenzyme, it has been of interest to deter-mine the metal specificities of various bacterial Glyoxalase I.

Very early on, it was determined that the metalloenzyme GlxIexhibits a Zn2+-dependence [9]. It was also reported that the GlxIfrom Pseudomonas putida is a Zn2+-activated enzyme [10]. It wastherefore naturally assumed that all GlxI from bacteria wouldexhibit an identical metal activation characteristic. However, it was

286 U. Suttisansanee, J.F. Honek / Seminars in Cell & Developmental Biology 22 (2011) 285–292

Fig. 1. Multiple sequence alignment of bacterial Glyoxalase I (organism name follows by National Center for Biotechnology Information (NCBI) accession number) includingE. coli (NP 310387), Y. pestis (ZP 01887743), N. meningitidis (CAA74673), P. aeruginosa GloA1 (AAG06912), P. aeruginosa GloA2 (AAG04099), P. aeruginosa GloA3 (AAG08496),P ino aca whileZ t andT rate a

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ss(Zasc(Za

. putida (AAN69360) and C. acetobutylicum (AAK80149). The GlxI with shorter amnd C. acetobutylicum are non-Zn2+-activated enzymes (being Ni2+/Co2+-activation),n2+-activated enzymes. The metal binding residues are marked with grey highlighhe alignment was created using CLC Free Workbench (version 3.0.1) with the accu

iscovered that GlxI isolated from the Gram-negative bacteriumscherichia coli is inactive with Zn2+, but is activated in the pres-nce of Ni2+ and Co2+ ions [11]. Similar metal activation profilesere also found in GlxI from other Gram-negative bacteria such

s Yersinia pestis and Neisseria meningitides [12,13] as well as theram-positive bacterium Clostridium acetobutylicum (unpublishedata) and from the trypanosome protozoa, Leishmania major [14].hese results divide GlxI into two classes according to metal activa-ion, Zn2+-activated and non-Zn2+-activated (Ni2+/Co2+-dependentnzymes). Based on current reports, it is likely that bacterial GlxInzymes are predominantly Ni2+/Co2+-dependent with the pos-ible exception of pseudomonads. As mentioned, early studiesn the field determined that the GlxI from P. putida is a Zn2+-ctivated enzyme [10,15]. However, Pseudomonas aeruginosa haseen reported to possess three genes including gloA1, gloA2 andloA3 that code for active GlxI enzymes (GloA1, GloA2 and GloA3,espectively) [16]. Metal activation analyses determined that GloA1nd GloA2 are Ni2+/Co2+-activated enzymes, while GloA3 is Zn2+-ctivated [16]. While non-Zn2+-activated GlxI possess high metalpecificity (high activity being observed in the presence of Ni2+ ando2+ and only trace activities found with other divalent metals suchs Mn2+ and Cd2+) [11], the Zn2+-dependent enzymes can be acti-ated by a variety of divalent metals (for example, GloA3 can bectivated by Ni2+, Mn2+, Co2+, Cd2+, Zn2+ and Mg2+) [16].

It appears that the amino acid length of the protein and itsequence can be used to make very good predictions as to the metalpecificity of a new GlxI. GlxI with shorter amino acid sequences∼130 amino acids in length) tend to be Ni2+/Co2+-activated (non-n2+-activated), while longer proteins (∼180 amino acids in length)re likely Zn2+-activated enzymes (Fig. 1). Analyses of multiple

equence alignments and X-ray crystallographic structures indi-ate that the Zn2+-activated GlxI possess a long N-terminal armregion A in Fig. 1) that wraps around another subunit [17]. Thisn2+-activated form also employs three extra loops (regions B, Cnd D in Fig. 1) that are absent in the Ni2+/Co2+-activated enzymes.

id sequences from E. coli, Y. pestis, N. meningitides, P. aeruginosa (GloA1 and GloA2)GlxI with longer amino acid sequences from P. aeruginosa (GloA3) and P. putida arebold letters indicate loop regions (A–D) that only exist in the Zn2+-activated GlxI.

lignment algorithm (http://www.clcbio.com).

The X-ray crystallographic structure of human GlxI (PDB: 1QIN), aZn2+-activated enzyme, suggests that one out of these three regionsis a three-turned �-helix (corresponds to region B, residues 81–95[17]) that blocks one side of the catalytic pocket. This loop mightplay a significant role in determining metal preference for variousGlxI, as it locates next to the active site. Two small loops (regions Cand D) locate at the outer layer of the catalytic pocket. These loopsmight be expected to contribute much less, if any, to the metalspecificity and enzymatic activity. Therefore, it would be interest-ing to investigate how the amino acid sequence relates to metalactivation of GlxI and how these extra loops might affect the metalactivation profile. Investigations in this area are currently beingundertaken.

GlxI is a member of the ����� superfamily of proteins, whichincludes GlxI, methylmalonyl-CoA epimerase (MMCE), extradioldioxygenase (DIOX), fosfomycin resistance protein, bleomycinresistance protein and mitomycin C resistance protein [18,19]. Thecatalytic/binding cleft is formed by arrangement of the �����motif either within one subunit, as in the monomeric Plasmodiumfalciparum GlxI (a subunit fusion as determined by the secondarystructural prediction and multiple sequence alignment [20]), orat the interface of a homodimer as in the case of Homo sapi-ens GlxI (PDB: 1QIN) [17]. A monomeric GlxI would thereforeform the active site from a single polypeptide (two active sitesper monomer), while a homodimeric protein would form twointermolecular domain–domain interfaces that require active siteresidues being contributed from both subunits (two active sites perdimer) (Fig. 2A).

The structure of the bacterial GlxI investigated so far is ahomodimeric structure, consisting of two subunits with two active

sites. The overall structure of the homodimeric E. coli GlxI isthat of the ����� superfamily structure [21]. Each monomerconsists of two domains, the N-terminal domain (residues 3–60)with a �1�1�2�3�4 arrangement and the C-terminal domain(residues 72–126) containing a �5�2�6�7�8 motif. Both domains

U. Suttisansanee, J.F. Honek / Seminars in Cell & Developmental Biology 22 (2011) 285–292 287

F ellow,a obutyli single

atl1tmubHpdf

ftaTptmtalbcaa

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ig. 2. (A) The ribbon structure of the overall structure of E. coli GlxI (green and yctive sites are formed at the dimeric interface. (B) The overall structure of C. acetntramolecular domain–domain interfaces where the active site is formed within a

re connected by an extended loop (residues 61–71). Even thoughhese two domains employ high structural similarity, they possessow sequence identity and similarity (2% sequence identity and3% sequence similarity), suggesting possible structural evolutionhrough gene fusion and mutagenesis steps from a common one-

odule ancestor. A series of 8 �-sheets (4 from each subunit) curvesp to form a catalytic pocket at the dimeric interface. The metalinding sites are formed by four metal binding residues (His5, Glu56,is74 and Glu122) that lie at the bottom of the catalytic pocket. Thisocket is created by Leu41, Phe43 and Val54 from the N-terminalomain of one subunit and Ala76, Val103, Ile110, Phe112 and Ile124

rom the C-terminal domain of another subunit.The X-ray crystallographic structure of the active Ni2+-bound

orm of the E. coli GlxI (PDB: 1F9Z) possesses an octahedral geome-ry with four metal binding residues (His5, Glu56, His74 and Glu122)nd two water molecules around the metal center (Fig. 3A) [21].he corresponding inactive form with Zn2+-bound (PDB: 1FA5)ossesses a five-coordinated trigonal bipyramidal geometry withhe same four metal binding residues but with only one water

olecule (Fig. 3B) [21]. Regardless of the metal-activation classhe GlxI belongs to (Zn2+-activated, or Zn2+-inactive but Ni2+/Co2+-ctivated), all active forms of GlxI appear to require an octahedraliganded arrangement around the catalytic metal. The differenceetween the active and inactive forms of GlxI may be explained byonsideration of their active site geometry [21,22]. This structuralrrangement must be critical to the mechanism of the enzyme buts yet the details of the catalytic steps remain elusive.

Interestingly, the isolation of a GlxI from a Gram-positiveacterium, C. acetobutylicum, provides an example of a bacterialnzyme with a different dimeric arrangement, wherein the activeites are formed at the two intramolecular domain–domain inter-aces (two active sites per dimeric enzyme) (Fig. 2B). Yet the overallrotein fold arrangement (����� motif) of each domain on the C.cetobutylicum GlxI is similar to that of the E. coli enzyme. The C.cetobutylicum GlxI possesses a shorter amino acid sequence, thus,his enzyme lacks the N-terminal arm and the extra three loopshat exist only in the Zn2+-activated enzymes. However, instead oforming two active sites at the dimeric interface, C. acetobutylicumlxI creates an active site within each subunit (PDB: 2QH0). Thenzyme from this Gram-positive bacterium is capable of bindingither Zn2+ or Ni2+ atoms in its X-ray crystallographic structurePDB: 2QH0 and 3HDP, respectively). However, the Ni2+-bound

nzyme exhibits an octahedral coordinated active site with fouretal binding residues (His5, Glu52, His75 and Glu124) and twoater molecules, while the Zn2+-bound form possesses a trigonal

ipyramidal geometry with the same metal binding residues butnly one water molecule liganded to the divalent metal. Investiga-

PDB: 1F9Z) employing two intermolecular domain–domain interfaces, where theicum GlxI (PDB: 2QH0) showing a different type of dimeric arrangement, the twosubunit.

tion of enzyme activity has shown that the C. acetobutylicum GlxIfalls into the non-Zn2+-activated class (being Ni2+/Co2+-activation,unpublished data), supporting the hypothesis that the shorter GlxIenzymes are Ni2+ and Co2+-activated enzymes even though theoverall protein arrangement of the homodimeric enzymes can bedifferent.

Analysis of the X-ray crystallographic structure of an inhibitor(S-(N-hydroxy-N-p-iodophenylcarbamoyl)glutathione) boundhuman GlxI (PDB: 1QIN) suggested that the hemithioacetal sub-strate likely binds in the active site in a “Y” conformation, wherethe thiol of the cysteine moiety is the base, and the glutamate andglycine moieties are the forks [23]. The free carboxylate groupsof �-glutamate and the C-terminal glycine in GSH point outwardin the crystal structure and interact with enzyme residues atthe entrance of the active site. Thus, the entrance to the activesite of GlxI requires a large space in order to support the sizeof its substrate (the distance from the �-carbon of glutamateto the carboxyl-carbon of the glycyl residue in GSH measures8.93 A). Even though there are two possible entrances for thehemithioacetal substrate in E. coli GlxI, the loop connecting �3and �4 (residues 46–54) partially blocks the active site, thuscreating a wall at one site of the catalytic pocket that prevents thesubstrate from contacting the aqueous solvent and might directthe incoming substrate to enter from the other wider entrance.E. coli GlxI contains four possible hemithioacetal binding residuesincluding Arg9, Asn60, Arg98 and Lys104 at this entrance (Fig. 4A).Arg9 and Asn60 from one subunit may stabilize the carboxylate andamino groups of the �-glutamate of the GSH moiety, while Arg98

and Lys104 from another subunit may interact with the carboxylgroup of its glycyl residue. The two oxygens from the three-carbonsubstrate might replace the two water molecules at the metalcenter, thus maintaining the octahedral coordination environmentaround the metal center.

The superimposed structures of Ni2+-bound E. coli GlxI (PDB:1F9Z) and the E. coli enzyme with a bound transition statehydroxamate analogue ([24] and unpublished data) contributeknowledge as to the likely conformational changes that occur inprotein binding of the GSH moiety. These two structures possesshigh structural similarity with the root mean square deviation(r.m.s.d.) of 0.69 A for 103 C� pairs with high flexibility at theloops (residues 34–39 and 99–110) and the C-terminus of theenzyme. The loop that locates between �2 and �3 (residues

34–39) and the loop that locates between �6 and �7 (residues99–110) move toward the active site, covering the catalytic pocketfrom above (Fig. 4B). The movement of the loop residues 99–110causes a significant change in the position of the substrate bind-ing residues, Arg98 and Lys104 (C� moves 1.386 A and 4.896 A,

288 U. Suttisansanee, J.F. Honek / Seminars in Cell & Developmental Biology 22 (2011) 285–292

F etry( ge and

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ig. 3. The metal coordination of E. coli GlxI can form either (A) an octahedral geomPDB: 1FA5). The water molecules are shown in blue, while the Ni2+ atom is in oran

espectively, away from its unbound structure) such that theiride chain amines are likely to form hydrogen bonding interac-ions with the carboxyl group of the glycyl residue in the GSH

oiety. Another two substrate binding residues, Arg9 and Asn60,re only slightly altered in these structures. The last nine aminocids at the C-terminus (residues 126–135) are absent from the-ray crystallographic structure but are observed upon binding of

he hydroxamate analogue (unpublished data). Although no directnteraction of this C-terminus and the hydroxamate analogue isetected, it is possible that this loop flips back to cover one sidef the active site once the substrate enters to protect it from thexternal environment. However, self-interactions within the C-erminus (such as Glu126/O–Ala129/N and Asp128/O–Arg131/N)s well as the interactions of the C-terminus and the loop residues9-110 (such as Thr107/N–Leu133/O, Thr107/O�1–Gly134/Ond Thr108/O�1–Gly132/O) are observed. These interactionsreate a network of interactions that likely maintains theydrophobic environment within the catalytic site and possi-

ly causes the structure of the C-terminus to become moreefined.

Previous metal titration versus enzyme activation studies, andsothermal titration calorimetry (ITC) and nuclear magnetic reso-ance (NMR) experiments have been utilized to show that the GlxI

ig. 4. (A) A ribbon structure of E. coli GlxI (green and yellow) showing interactions of tys104 with the bound transition state analogue of hydroxamate, TSI (unpublished structDB: 1F9Z) and the enzyme with bound hydroxamate analogue (green and yellow, unpubhe active site upon binding of the inhibitor. The Ni2+ atom in E. coli GlxI with bound hydroound inhibitor is in grey.

with Ni2+ atom (PDB: 1F9Z) or (B) a trigonal bypiramidal geometry with Zn2+ atomthe Zn2+ atom is in magenta.

from E. coli and C. acetobutylicum possess 1 mol of metal per mol ofdimeric enzyme for both Ni2+ and Co2+-activation. These analysessuggest that the metal ion binds tightly in the catalytic pockets ofthese two enzymes and that only one active site may be functionalat any one time ([11,25,26] and unpublished data). This situationmay be explained in terms of asymmetry in the subunits which leadto asymmetric active sites. The information obtained from analy-sis of the X-ray crystallographic structures and 15N–1H HSQC NMRexperiments on E. coli GlxI suggested a slight asymmetry betweenthe two active sites, in which one active site possesses a higherbinding affinity for metal and substrate than the other site [21,26].It was previously found that the apo-enzyme contains two asym-metric active sites with one site having a higher binding affinity formetal [21,25,26]. Addition of metal to the first active site occursrapidly and induces a subtle but measurable protein conforma-tional change [26]. The binding of metal to the second unoccupiedactive site requires additional time, and both active sites of metal-reconstituted enzyme regain increased symmetry (r.m.s.d. of the

2+

apo E. coli GlxI and its Ni bound form (PDB: 1FA8 and 1F9Z, respec-tively) is 0.201 A for 128 C� pairs) [21,26]. However, the flexibility(based on analysis of the B-factor in the protein structures) of theloops (residues 99–110 and 34–39) which act as lids to cover theactive sites is different, indicating that one active site might be more

he four possible hemithioacetal binding residues including Arg9, Asn60, Arg98 andure). (B) The superimposed structure of Ni2+-bound E. coli GlxI (magenta and cyan,lished data) showing the enzymatic movement of the loop residues 99–110 towardxamate analogue is shown in orange, while the Ni2+ atom in E. coli enzyme without

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U. Suttisansanee, J.F. Honek / Seminars in C

uitable to binding substrate and which might possess a higher sub-trate binding affinity [21]. Differences between the two active sitesn a single subunit GlxI (P. falciparum; Deponte, in this issue) as wells protozoan two subunit GlxI (Fairlamb, in this issue) have alsoeen observed [14,20].

The distinct difference in the loop region in both active sitesan be observed in the superimposed structures of Ni2+-bound. coli GlxI without bound inhibitor (PDB: 1F9Z) and its structureith bound hydroxamate analogue (unpublished data). This anal-

sis indicates that one active site employs a larger movement inhe loop residues 99–110 and a smaller movement in the loopesidues 34–39 compared to those of the other active site. Theifference in degree of movement may affect the network con-ections within the enzyme itself as well as the interactions withhe GSH moiety. The different flexibility of the substrate stabilizingesidues (Arg98 and Lys104) in each active site is possibly crucialor interactions with the glycine residue of GSH moiety and theoss of these enzyme–substrate interactions would be expected toause a decrease in activity as had been observed previously forhe human GlxI (the mutation of M157A in human GlxI, the residuehat corresponds to Lys104 in E. coli GlxI, causes significant lossn activity [27]). As well, the substrate might choose to enter thective site which has a greater loop flexibility. However packingf the enzyme in the crystal lattice in these structures will likelyontribute to some extent to the differing flexibility of the loopsn these two subunits, but its magnitude is unknown. The NMRtructural investigation however does not suffer from this possi-le contributing factor and does indicate the presence of active sitesymmetry [26].

In evolution of the protein structural scaffolds for GlxI, it haseen previously proposed that the variety of domain swapping forlyoxalase I enzymes are the result of gene duplication and fusionvents of a common one-module ancestoral gene [10,19,28]. A two-otif pseudosymmetric metallomonomer from P. putida with one

ctive site was believed to have evolved from this single domainrecursor by formation of a link between the two domains [10].his process was suggested based on the discovery of a monomericorm of GlxI from P. putida that forms upon treatment of then2 dimeric protein with EDTA (apo-monomer) or glutathione (Znonomer) [10]. It has been suggested that the Zn monomer is

n intermediate between the Zn2 dimer and apo-monomer andccommodates a non-covalently bound glutathione in the activeite, in which its linker is rearranged in order to improve GSH bind-ng [10]. This Zn monomeric enzyme exhibited some enzymaticctivity (kcat ∼ 115 ± 40 s−1) but this was significantly lower thants fully dimeric two-domain structure (kcat ∼ 500 ± 100 s−1). Uponlutathione removal, the Zn monomer regains its dimeric structure10].

Due to remarkable structural and noticeable sequence similarityf the two domains, it is possible that the dimeric GlxI (four domainsf two polypeptides) with two active sites was a second genera-ion of the gene duplication-fusion event. The dimeric interactionf bacterial GlxI, so far, is now been shown to form two possiblerrangements. The dimeric GlxI from C. acetobutylicum is formedith a back-to-back interaction of two monomers with one active

ite formed within one subunit. Each subunit of C. acetobutylicumlxI likely employs a protein fold that might be expected for thease of the P. putida Zn-monomer. On the other hand, the dimertructure observed for the GlxI from E. coli, if it did evolve from thisonomeric two-domain GlxI (i.e. monomeric P. putida GlxI), would

equire the evolution of a shift from a single subunit active site by

utation, to a structure that forms a new dimeric interaction, cre-

ting an active site at the interface of the dimer. These processesre far more complicated than the formation of dimeric structurexhibited by the C. acetobutylicum GlxI. Therefore, it is easy to imag-ne (although difficult to prove!) that this enzyme might be an

evelopmental Biology 22 (2011) 285–292 289

intermediate between the monomeric two-domain and the dimerictwo-domain GlxI enzyme structures.

1.2. Glyoxalase II

Although a number of bacterial GlxI have now been character-ized, few bacterial GlxII have been investigated. GlxII from E. coliand Salmonella typhimurium have been examined and character-ized, while two other putative GlxII enzymes from Campylobacterjejuni and N. meningitides are mentioned through amino acidsequence searching [13,29–32]. GlxII exists in both the cytosol andthe mitochondria of eukaryotes. In animals, multiple forms of GlxIIare found in mitochondria (both in the intermembrane space and inthe matrix), while there appears to be only one form in the cytosoland only the enzyme from the intermembrane space appears toresemble the cytosolic GlxII form [33–36]. Plants, on the otherhand, appear to possess a mitochondrial GlxII form, but multiplecytosolic forms [37–39], while yeast employ a cytosolic as well as amitochondrial form [40,41]. The discovery of GlxII in mitochondriasuggests the possible transport of S-d-lactoylglutathione from thecytosol to the mitochondria in eukaryotes [36]. Since mitochon-dria contain d-�-hydroxyacid dehydrogenase, an enzyme that isresponsible for the formation of pyruvate from d-lactate, it is pos-sible that pyruvate can be formed directly in the mitochondria [36].Bacterial GlxII, however, appear to exist as one single gene copy[30,31].

GlxII is a metalloenzyme with binuclear active sites permonomer that can be activated by various metals depending onthe particular source organism. This property has been compared tothe related metallo-�-lactamase group of enzymes where mononu-clear metal ion-containing active sites are observed [42]. Analysisof the properties of the GlxII from S. typhimurium indicated thatthe enzyme can be activated in the presence of Fe2+, Fe3+, Zn2+ andMn2+ (0.21 mol zinc, 0.64 mol iron and 0.3 mol manganese), thoughoptimal activity is observed with Mn2+ [31]. These variations inmetal content do not have significant effects on the enzymaticactivity [31] but may contribute to a cellular mechanism to con-trol the enzyme’s activity [43]. On the other hand, as isolated, E. coliGlxII binds 1.7 mol of zinc per mol monomeric enzyme, while othermetals were not detected [30]. The activity of the metal chelator-treated enzyme (apo-enzyme), however, could be restored withthe addition of Mn2+ or Co2+ [30]. Interestingly, this apo-enzymecannot be fully reactivated by the addition of Zn2+, suggesting thatZn2+ inhibition might also occur, possibly due to interactions ofthe metal with cysteine residues in the protein [30]. Unlike GlxI,GlxII does not possess two distinct metal activation classes suchthat no enzymatic activity was found in attempts to reconstitutethe apo-enzyme with Ni2+ [30].

The Zn2+-bound E. coli GlxII (1.7 mol of zinc per mol monomericenzyme) with S-d-lactoylglutathione possesses a kcat of 53 s−1

and Km of 184 ± 22 �M, and a similar enzymatic activity wasobserved with the Mn2+-reconstituted enzyme [30]. The kineticsof S. typhimurium GlxII with bound Mn2+ on the same substrateexhibits a kcat of 395 ± 11 s−1 and Km of 241 ± 18 �M [31]. However,the metal content of this latter enzyme is only approximately 1 molMn2+ per mol monomeric enzyme, suggesting that the enzyme withonly one metal atom is catalytically active and the second metalmight not be necessary for catalysis [31]. These results are fur-ther supported by the discovery of the properties of the GlxII fromSalmonella enteric serovar Typhimurium (YcbL), where this GlxIIpossesses only one metal binding site with optimal activity being

observed in the presence of iron [32] and the human enzyme alsobeing reported as active in its mono-metallated Zn2+ form [44].

GlxII is a member of the metallo-�-lactamase superfamily [45].The proteins in this superfamily share a common ��/�� foldand contain the conserved motif THxHxDH which serves in metal

290 U. Suttisansanee, J.F. Honek / Seminars in Cell & D

Fig. 5. (A) The overall structure of Salmonella typhimurium GlxII (PDB: 2QED) show-ing the ��/�� fold of the N-terminal domain (green) and the �-helix containingC-terminal domain (yellow). (B) The metal binding sites of the two iron atoms(magenta) showing the metal binding residues (His53, His55, His110, Asp57, His58,His165 and Asp127) and water molecules (blue) that form a square pyrimidal geome-try around the metal centers (magenta). (C) The predicted substrate binding siteconsisting of five substrate binding residues (Phe138, Arg136, Tyr167, Arg245 andLs2

baaaioncodT

typhimurium GlxII with a water molecule in YcbL, while all metal

ys248) that stabilize glutathione, in which its presence is from the superimposedtructure of human GlxII (PDB: 1QH5) and Salmonella typhimurium GlxII (PDB:QED).

inding. So close is the structural relatedness that beta-lactamasectivity can even be detected in Glyoxalase II that has beenltered by directed evolution experiments [46]. This superfamilylso contains a quorum-quenching N-acyl-l-homoserine lactonase,mportant in controlling the concentrations of a particular classf bacterial sensor molecules. This enzyme’s structural related-ess to GlxII has been noted [47,48]. The overall structure of GlxII

an be divided into two parts according to its predominant sec-ndary structure and fold, the �-sheet containing the N-terminalomain and the �-helix containing the C-terminal domain (Fig. 5A).he N-terminal domain possesses a similar protein fold to the

evelopmental Biology 22 (2011) 285–292

metallo-�-lactamase superfamily, which consists of a four-layered� sandwich with two mixed � sheets flanked by �-helices. The C-terminal domain mainly consists of �-helices and is essential forsubstrate binding [49]. The enzyme contains two metal bindingsites, which are located in the N-terminal domain on the edge ofthe �-sandwich, while the substrate binding site lies at the inter-face of the two domains. The X-ray crystallographic structure of theS. typhimurium GlxII (PDB: 2QED) contains two iron atoms in themetal binding sites. One iron atom is coordinated by His53, His55,His110 and a water molecule, while the other iron atom is boundto Asp57, His58, His165 and another water molecule (Fig. 5B). Thesetwo metals are bridged by a third water molecule and Asp127, form-ing an octahedral coordination around each metal center (Fig. 5B).Interestingly, an investigation on human GlxII with bound S-(N-hydroxy-N-bromophenylcarbamoyl)glutathione (HBPC-GSH, PDB:1QH5), a slow substrate of GlxII, reported that the active site geom-etry is maintained upon binding of the substrate analogue, wherethe carbonyl oxygen replaces a water molecule and is bound to theZn1 atom [49]. This metal coordination might be significant for themechanism of the reaction as has been proposed for the humanGlxII [49]. However, the metal coordination environment for theZn2 atom results in a square pyramidal geometry due to the loss ofone water molecule. The change in the active site geometry of thesecond metal Zn2 might create an unsuitable environment for thesubstrate to bind, thus supporting the previous observation on opti-mal activity being observed with only one occupied metal bindingsite on the enzyme.

The superimposed structures of human GlxII with bound GSH(PDB: 1QH5) and S. typhimurium GlxII (PDB: 2QED) help predictthe substrate binding residues in the latter enzyme (Phe138, Arg136,Tyr167, Arg245 and Lys248, Fig. 5C). The carboxylate of the glycinemoiety of GSH might interact with Arg245 and Lys248 from the C-terminal domain. The amide nitrogens of the glycine and cysteinemoieties of GSH are located within hydrogen bonding distance ofthe hydroxyl group of Try167. This Tyr residue can also possiblyform an interaction with the sulfur atom from the cysteine moietyof GSH. The amine side chain of Lys143 in human GlxII (correspond-ing to Arg136 in S. typhimurium GlxII) can interact with the carbonyloxygen of the �-glutamate moiety of GSH while the side chain ofArg136 in S. typhimurium GlxII swings away such that it locates out-side the hydrogen bonding distance of the thiol. Instead, the amidenitrogen of this residue can stabilize the carbonyl oxygen of the cys-teine in GSH. The carbonyl oxygen of the �-glutamate moiety of GSHis also stabilized by interaction with Tyr145 from the human GlxII(corresponding to Phe138 in S. typhimurium GlxII). This interaction,however, is eliminated in the studied GlxII from other organismssince Tyr145 is substituted by Phe (except in the mitochondrial GlxIIfrom Arabidopsis thaliana, which possesses Ser), suggesting that thisresidue is not required for substrate recognition [49]. The molec-ular recognition of the substrate is believed to occur through theGSH moiety and the interaction of the thioester group with metalions.

Interestingly, the unusual GlxII from S. enteric serovarTyphimurium, YcbL (PDB: 2XF4), contains only one metal bind-ing site and lacks the C-terminal domain [32]. Even though all themetal binding residues are conserved (His56, His58, Asp60, His61,Asp151, His132 and His192), only Asp60, His61, Asp151 and His192

as well as a water molecule form a trigonal bipyramidal geom-etry around the metal center. Analysis of the superimposition ofthe molecular structures of YcbL (PDB: 2XF4) and S. typhimuriumGlxII (PDB: 2QED) shows the replacement of the second metal in S.

binding residues are well overlapped. As well, since there is onlyone metal ion in the catalytic site, the bridging water molecule is notpresent. In addition, because of the lack of the C-terminal domainin YcbL, only two out of five substrate binding residues remain

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U. Suttisansanee, J.F. Honek / Seminars in C

Arg160 and Phe163). The superimposed structures of human GlxIIith bound GSH (PDB: 1QH5) and YcbL (PDB: 2XF4) suggest that

he guanidino side chain of Arg160 might be able to interact withhe carbonyl oxygen of the �-glutamate moiety of GSH. As YcbLacks the essential substrate stabilizing residues in the C-terminalomain and its metal coordination might not be suitable for cat-lytic reaction, the significantly lower enzymatic activity of thislxII to other bacterial enzymes might be explained [32].

.3. Glyoxalase III

A report has appeared that provides evidence for a singlenzyme in E. coli, termed Glyoxlase III, which directly convertsethylglyoxal to d-lactate without the need for an intracellular

hiol cosubstrate [50]. Little information is available on this enzymeowever one report indicates the possible role of RNA polymeraseigma factor (rpoS) in the bacterial regulation of this enzyme activ-ty [51].

Summary: New discoveries in the structure and function ofhe bacterial glyoxalase enzymes ere improving our understand-ng of the contributions that protein structure make to metalctivation profiles for these enzymes. This fundamental knowl-dge contributes to our basic information on the function of thesenzymes and their molecular products in the biochemistry of bac-eria [52–54].

cknowledgments

The authors gratefully acknowledge research support fromSERC (Canada) to J.F.H., and the University of Waterloo and theovernment of Thailand for graduate support and scholarships to.S. The authors wish to gratefully acknowledge our collaborators,r. S.K. Burley, Dr. G. Davidson, Dr. M.M. He, Dr. S. Lagishetty, Dr..J. Maroney, Dr. B.W. Matthews, Dr. K.N. Rao, Dr. J.M. Sauder, Dr.

. Swaminathan.

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