- Email: [email protected]
Vital roles of an interhelical insertion in catalase–peroxidase bifunctionality
BBRC Biochemical and Biophysical Research Communications 318 (2004) 970–976 www.elsevier.com/locate/ybbrc
Vital roles of an interhelical insertion in catalase–peroxidase bifunctionalityq,qq Yongjiang Li and Douglas C. Goodwin* Department of Chemistry and Program in Cell and Molecular Biosciences, Auburn University, Auburn, AL 36849-5312, USA Received 25 March 2004 Available online 7 May 2004
Abstract The loop connecting the F and G helices of catalase–peroxidases contains a 35 amino acid structure (the FG insertion) that is absent from monofunctional peroxidases. These two groups of enzymes share highly similar active sites, yet the monofunctional peroxidases lack appreciable catalase activity. Thus, the FG insertion may serve a role in catalase–peroxidase bifunctionality, despite its peripheral location relative to the active site. We produced a variant of Escherichia coli catalase–peroxidase (KatG) lacking its FG insertion (KatGDFG ). Absorption spectra indicated the heme environment of KatGDFG was highly similar to wild-type KatG, but the variant retained only 0.2% catalase activity. In contrast, the deletion reduced peroxidase activity by only 50%. Kinetic parameters for the peroxidase and residual catalase activities of KatGDFG as well as pH dependence studies suggested that the FG insertion supports hydrogen-bonded networks critical for reactions involving H2 O2 . The structure also appears to regulate access of electron donors to the active site. Ó 2004 Elsevier Inc. All rights reserved. Keywords: KatG; Deletion mutagenesis; Heme; Catalase–peroxidase; Cytochrome c peroxidase; ABTS; Synechocystis KatG
Catalase–peroxidases are heme enzymes that use a single active site to catalyze both catalase and peroxidase reactions [1,2]. These enzymes have generated much interest because catalase–peroxidase from Mycobacterium tuberculosis is essential for the activation of the front-line antitubercular agent, isoniazid [3]. Indeed, it has been estimated that over 70% of isoniazid-resistant strains of M. tuberculosis carry mutations that alter the function of catalase–peroxidase [4]. Furthermore, several highly virulent pathogens carry unique periplasmic catalase–peroxidases that non-pathogenic relatives of these organisms lack, suggesting that these catalase– peroxidases may be virulence factors [5–8]. Clearly, the
q Funds to support this research were provided by the Petroleum Research Fund of the American Chemical Society (PRF 38802-G4) and Auburn University. qq Abbreviations: KatG, E. coli catalase–peroxidase; KatGDFG, 0 KatG lacking FG insertion; ABTS, 2,2 -azino-bis(3-ethylbenz-thiazoline-6-sulfonic acid); CT, charge transfer transition. * Corresponding author. Fax: 1-334-844-6959. E-mail address: [email protected] (D.C. Goodwin).
0006-291X/$ - see front matter Ó 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2004.04.130
evaluation of catalase–peroxidase structure and function has important ramifications. The peroxidase and catalase catalytic cycles both begin with the oxidation of the ferric heme by two electrons by a hydroperoxide (typically H2 O2 ) to form the ferryl-oxo porphyrin/protein radical intermediate known as compound I. The catalase cycle is completed by the reduction of compound I to the ferric enzyme by H2 O2 , resulting in the production of molecular oxygen. Conversely, in the peroxidatic cycle, compound I is typically reduced in two sequential one-electron steps by an exogenous reducing substrate. Two equivalents of substrate radical are generated and the enzyme is reduced to the ferric state via the ferryl-oxo intermediate known as compound II (Fig. 1). In spite of their efficient catalase activity, catalase– peroxidases bear no sequential or structural resemblance to monofunctional catalases. Conversely, the catalase– peroxidases are highly similar to the monofunctional peroxidases. Indeed, two recent crystal structures of catalase–peroxidases demonstrate that most of the major amino acid side chains in the active site are virtually
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Fig. 1. Catalytic cycle of catalase–peroxidases. A typical exogenous peroxidase reducing substrate as (AH) and its corresponding radical (A ) are also shown.
superimposable on those of cytochrome c peroxidase [9,10]. Nevertheless, cytochrome c peroxidase and other monofunctional peroxidases lack appreciable catalase activity. Given the high similarity of the monofunctional peroxidase and catalase–peroxidase active sites, it is reasonable to suggest that protein structures external to the catalase–peroxidase active site have an important role in fine-tuning the active site for its bifunctional abilities. In support of this hypothesis, comparisons of the protein structures and amino acid sequences of these two groups of enzymes reveal three structural features that are unique to the catalase–peroxidases. These are two interhelical insertions (35 amino acids each) and a C-terminal domain (300 amino acids) [11,12]. All three of these structures are peripheral to the catalase–peroxidase active site [9,10]. One of these interhelical insertions appears within a loop that connects the F and G helices of the catalase– peroxidase N-terminal domain (Fig. 2). Thus, we refer
971
to this structure as the FG insertion. Because this structure is uniformly absent from monofunctional peroxidases, it may have an important role in catalase– peroxidase bifunctionality. However, until the present time, the function of neither the FG insertion nor any residue composing it has been investigated. We employed a novel deletion mutagenesis procedure to generate a clone for recombinant Escherichia coli catalase–peroxidase (KatG) lacking its FG insertion (KatGDFG ). This variant was expressed, isolated, and characterized. Relative to wild-type KatG (wtKatG), KatGDFG retained appreciable peroxidase activity but very little catalase activity, demonstrating that this structure is essential for the unique bifunctional capabilities of catalase–peroxidases. Additionally, the kinetic parameters for the remaining peroxidase activity as well as the spectral properties of KatGDFG have provided valuable information on specific contributions of this structure to catalase–peroxidase function.
Materials and methods Reagents. All materials for cloning, mutagenesis, protein expression, and protein purification were purchased as previously delineated [13]. Hydrogen peroxide (30%) and hemin were purchased from Sigma (St. Louis, MO). All buffers and media were prepared using water purified through a Millipore Q-PakII system (18.2 MX/cm resistivity). Cloning and expression of wtKatG and KatGDFG . To generate the expression constructs for KatGDFG , we developed a procedure adapted from Seamless cloning (Stratagene). The plasmid for expression of recombinant His-tagged E. coli KatG (pKatG1) was used as a template. Primers were designed to: (1) amplify pKatG1 excluding the codons corresponding to the targeted deletion (Pro 277–Thr 311), (2) include a restriction site for Eam1104 I, and (3) yield compatible overhanging sequences upon digestion with Eam1104 I. Because Eam1104 I is a type IIS endonuclease, it cleaves target DNA downstream of its recognition sequence. Thus, the resulting overhanging sequences can be designed for religation such that only the codons corresponding to the targeted deletion are removed. No introduction
Fig. 2. Position of the FG insertion relative to key amino acids within the KatG active site. (A) Maximizes visibility of key active site side chains and their positions relative to the a carbons of the FG insertion (ribbon). The positions of N-terminus (P277) and C-terminus (T311) of the FG insertion are indicated. Two important H-bonds are shown as dark lines. (B) Represents a 120° leftward rotation of the structure. The N-terminus of the FG insertion (P277) is indicated. All numbering is according to E. coli KatG. Coordinates are from the B. pseudomallei KatG structure (1MWV) [9].
972
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
of new restriction sites or extra codons is involved, nor is the removal of codons outside the target sequence required. We used primers ECCP D201 (TAC ATC TCT TCT AGA ACC GGC ACC GTG GGT TTT ACC CAG) and ECCP D202 (GAT GCC TCT TCC TCT GGT CTG GAA GTA GTC TGG). The Eam1104 I recognition sequence is underlined, and the overhanging sequence generated upon hydrolysis by Eam1104 I is in bold. The PCR product was digested with Eam1104 I and ligated using T4 DNA ligase (in the presence of Eam1104 I). The ligated plasmid was used to transform E. coli (XL-1 Blue). Colonies were selected on the basis of ampicillin resistance, and candidate plasmids were verified by DNA sequence analysis to ensure the accuracy of the mutagenesis procedure. As we have done with pKatG1, the E. coli expression host (BL-21 [DE3] pLysS) was transformed with pKatGDFG . Expression of wtKatG and KatGDFG was accomplished as described for another catalase–peroxidase [13]. Purification of wtKatG and KatGDFG . All cell pellets were resuspended in Bugbuster reagent supplemented with 0.1 mM phenylmethylsulfonyl fluoride. Following homogenization, benzonase (250 U) was added, and the mixture was incubated at 23 °C with gentle stirring for 1 h. The cell lysate was then centrifuged at 16,000g. For wtKatG and KatGDFG , the supernatant was loaded onto a Ni– NTA column by recirculating the solution through the column bed overnight (1 mL/min). The column was then washed with buffer A (50 mM phosphate buffer, pH 8.0; 200 mM NaCl) supplemented with 2 mM imidazole. A second wash was then performed with buffer A supplemented with 20 mM imidazole. Finally, protein was eluted off the column with buffer A supplemented with 200 mM imidazole. Excess imidazole was then removed by dialysis or gel filtration chromatography. Enzyme reconstitution. We have observed that wtKatG and KatGDFG can be reconstituted with hemin following purification. Titration of each of the apoproteins with hemin results in a linear increase in activity up to 1 equivalent. Addition of hemin beyond this point does not result in any additional activity. To minimize contribution of free or adventitiously bound hemin to spectral or kinetic measurements, each protein was reconstituted with 0.75 equivalents hemin. Hemin concentration was determined by the method of Falk [14]. Enzyme activities and UV–visible absorption spectra. Catalase and peroxidase activities as well as UV–visible absorption spectra were obtained and evaluated as described previously for another catalase– peroxidase [13].
Results Mutagenesis, expression, and isolation of KatGDFG We developed a procedure to specifically delete the FG insertion from E. coli catalase–peroxidase (KatG)
(see Materials and methods). Based on sequence alignments of catalase–peroxidases in comparison to those for other class I peroxidases (i.e., cytochrome c peroxidase and ascorbate peroxidases) [11,12], we designed primers to seamlessly eliminate the codons corresponding to Pro 277–Thr 311 (i.e., the FG insertion) (Fig. 2). The resulting variant, KatGDFG , was expressed in a T7-based system and isolated in a soluble form. Electrophoretic migration of the KatGDFG monomer evaluated by SDS–PAGE was consistent with elimination of the targeted 35 amino acid segment (data not shown). UV–visible spectra of wtKatG and KatGDFG Following reconstitution with hemin (see Materials and methods), we recorded UV–visible absorption spectra for the ferric, ferrous, and ferri-cyano forms of KatGDFG (Fig. 3). For ferric KatGDFG , the Soret maximum (408 nm) and charge transfer maxima (501 and 637 nm) are consistent with a mixture of states dominated by pentacoordinate and hexacoordinate high-spin heme iron. A very weak shoulder at 535 nm suggests a minor contribution from hexacoordinate low-spin heme. The pentacoordinate high-spin species dominates in the ferrous state, as indicated by a Soret maximum of 438 nm in addition to a strong b band at 561 nm and a relatively weak a band at 590 nm. All spectra for KatGDFG were highly similar to those for wtKatG (Table 1), suggesting that removal of the FG insertion did little to disrupt the immediate environment around the heme group. The most notable spectral difference between wtKatG and KatGDFG was the relatively low molar absorptivity of the ferric state. One possible explanation would be that a portion of the hemin from reconstitution remained unbound by the enzyme. However, we have recently been able to express and isolate KatGDFG in its holoenzyme form using an expression system developed in our laboratory [15]. We observe spectral and catalytic properties highly similar to those of the reconstituted enzyme (data not shown).
Fig. 3. Absorption spectra recorded for KatGDFG . Spectra were obtained for ferric (bold solid line), ferri-cyano (dotted line), and ferrous KatGDFG (dashed line). (A) shows these spectra from 350 to 500 nm, and (B) shows them from 475 to 725 nm. Ferri-cyano KatGDFG was generated by the addition of 2 mM NaCN to ferric KatGDFG . Ferrous KatGDFG was generated by the addition of a few grains of solid dithionite to ferric KatGDFG . All spectra were recorded at 23 °C in 100 mM phosphate, pH 7.0.
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
973
Table 1 Absorption characteristics of wtKatG and the KatGDFG varianta Absorption maxima [nm] (absorptivity [mM1 cm1 ])
Enzyme
Soret
b
a
CT2b
CT1c
Ferric wtKatG KatGDFG
408 (120.7) 408 (77.4)
—d
—
—
—
502 (17.5) 501 (10.9)
629 (9.7) 637 (4.0)
Ferri-cyano wtKatG KatGDFG
423 (96.1) 423 (86.9)
542 (15.6) 541 (10.8)
—
—
—
—
—
—
Ferrous wtKatG KatGDFG
439 (79.8) 438 (88.2)
561 (15.1) 560 (13.4)
581 (10.7) 590 (8.5)
—
—
—
—
a
All spectra were collected in 100 mM phosphate buffer, pH 7.0, 23 °C. b CT2 short wavelength charge transfer transition (usually between 490 and 510 nm). c CT1 long wavelength charge transfer transition (usually between 600 and 650 nm). d Transitions that were absent or too weak to make unequivocal assignments.
Table 2 Steady-state kinetic parameters for catalase and peroxidase activities of wtKatG and KatGDFG Activitya (substrate)
Enzyme wtKatG
KatGDFG
b
Catalase (H2 O2 ) KM (mM) kcat (s1 ) kcat =KM (M1 s1 ) Peroxidase (H2 O2 )c KM (mM) kcat (s1 ) kH2 O2 (M1 s1 ) Peroxidase (ABTS) KM (mM) kcat (s1 ) kABTS (M1 s1 )
4.0 0.4 (1.2 0.4) 104 (3.1 0.5) 106
4.5 0.7 26 4 (5.7 0.2) 103
0.83 0.07 58 3 (7.1 0.9) 104
1.5 0.1 26 1 (1.8 0.1) 104
0.23 0.05 52 8 (2.2 0.2) 105
0.012 0.001 26 2 (2.2 0.1) 106
a
All assays were performed at 23 °C. All catalase assays were carried out in 100 mM phosphate buffer, pH 7.0. c All peroxidase assays were carried out in 50 mM acetate buffer, pH 5.0. b
Catalase and peroxidase activities of wtKatG and KatGDFG Consistent with previous reports [13], our preparation of wtKatG showed substantial catalase activity (Table 2). Removal of the FG insertion produced a precipitous reduction in catalase activity, decreasing the apparent second-order rate constant by a factor of 550 relative to wtKatG. This effect was due primarily to a decrease in kcat from 1.2 104 s1 for wtKatG to 26 s1 for KatGDFG . The apparent KM for H2 O2 as measured from catalase activity was unchanged. In addition to the loss of activity, shifts in pH profile were observed. The pH optimum shifted from 6.5 to 7.0, and although a similar dependence of relative rate on pH was observed at high pH values, the catalase activity of KatGDFG
Fig. 4. Effect of pH on catalase activities of wtKatG and KatGDFG . Catalase activities of wtKatG (s) and KatGDFG (j) were evaluated in the presence of 20 mM H2 O2 and 100 mM phosphate at 23 °C.
decreased more sharply below 7.0 than observed for wtKatG (Fig. 4). KatGDFG retained appreciable peroxidase activity (Fig. 5). With respect to H2 O2 or ABTS, kcat values (26 s1 in each case) were roughly half of that observed for wtKatG (Table 2). A moderate increase (twofold) in the apparent KM for H2 O2 was observed. Conversely, the apparent KM for the reducing substrate, ABTS, decreased by a factor of 10 upon removal of the FG insertion. The kinetic parameters obtained with respect to H2 O2 have been used to describe reactions leading to the formation of compound I [16]. Elimination of the FG insertion led to a fourfold decrease in apparent second-order rate constant (kH2 O2 ) governing these steps. On the other hand, kinetic parameters obtained with respect to the reducing substrate (in this case ABTS) describe the reactions that result in conversion of compound I back to ferric enzyme. The apparent second-order rate constant (kABTS ) governing these steps increased by a factor of 10 (Table 2). In addition to a dramatically reduced KM for ABTS, we also observed ABTS-dependent inhibition of KatGDFG peroxidase activity at concentrations above
974
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Fig. 5. Comparison of the peroxidase activities of wtKatG and KatGDFG . The effects of H2 O2 (A) and ABTS (B) concentrations on the rate of ABTS oxidation by wtKatG (d) and KatGDFG () were evaluated. (A) ABTS concentrations were selected to obtain optimum peroxidase activity: 1 mM for wtKatG and 0.1 mM for KatGDFG . (B) All reactions contained 10 mM H2 O2 . All reactions were carried out at 23 °C in 50 mM acetate buffer, pH 5.0. The [E]T values were calculated based on heme content.
0.1 mM (Fig. 5B). At 1 mM ABTS, a 33% decrease in activity from the recorded maximum was observed. No inhibition was apparent for wtKatG over this same concentration range. Likewise, the peroxidase activity of neither wtKatG nor KatGDFG was sensitive to peroxidedependent inhibition (Fig. 5A).
Discussion Catalase–peroxidases have the capacity to catalyze two reactions using a single active site. Remarkably, this bifunctional active site is highly similar to that of the less versatile monofunctional peroxidases. Catalase–peroxidases all have an insertion (35 amino acids) that extends the loop connecting the F and G helices relative to that of monofunctional peroxidases. This FG insertion is peripheral to the active site. Nevertheless, our results demonstrate that this structure is essential for catalase–peroxidase bifunctionality. Furthermore, spectra recorded for KatGDFG showed only subtle changes in heme environment, indicating that observed effects on catalase activity were not due to dramatic changes in the coordination sphere of the heme iron. Roles of the FG insertion in reactions with H2 O2 The catalase cycle involves two principal reactions with H2 O2 , compound I formation and compound I reduction. Recent site-directed mutagenesis studies have identified three mechanisms by which the catalase activity of catalase–peroxidase is supported by the active site. Each participant in a novel covalent link (Trp 105, Tyr 226, and Met 252 [E. coli KatG numbering]) has been shown in at least one catalase–peroxidase to be essential for catalase but not peroxidase activity [17–19]. Our data to date suggest KatGDFG still forms this covalent structure. A hydrogen-bonded network involving Asp 135 and Asn 136 also appears to be important. The analogous residues in Synechocystis KatG (Asp 152 and Asn 153) are essential for full catalase activity. Interestingly, replacement of each of these residues
diminishes catalase activity, apparently by differing mechanisms. Disruption of a hydrogen bond between the distal histidine and Asn 153 primarily affects compound I formation [20], whereas replacement of Asp 152 disrupts compound I reduction [21]. The rate-determining step for catalase activity is thought to be compound I formation [22]. The change in pH profile for KatGDFG is consistent with either a change in the rate-determining step or a dramatic shift in the pKa s associated with it. The pH profile is reminiscent of that observed for D152N Synechocystis KatG, where H2 O2 -dependent compound I reduction is the principal reaction affected by the substitution [21]. As observed for D152N Synechocystis KatG, KatGDFG also shows minimal effects on peroxidase activity as compared to the substantial reduction in catalase activity. Indeed, our kABTS value for KatGDFG suggests that, if anything, peroxidatic reduction of compound I is enhanced by elimination of the FG insertion. Indirect evidence suggests that compound I formation has also been diminished to some extent. Our kH2 O2 values derived from peroxidase activity decreased fourfold relative to wtKatG. This is in spite of the near absent catalase activity. Normally, consumption of H2 O2 by compound I is expected to diminish observed peroxidase activity because it is “non-productive” consumption of H2 O2 (i.e., it does not result in oxidation of ABTS or other reducing substrates). Indeed, catalasenegative catalase–peroxidase variants often show an increase in peroxidase activity [17,18,21]. To the present time, we have been unable to directly observe H2 O2 dependent KatGDFG compound I formation by stoppedflow. However, rates of CN binding (often monitored as a reaction analogous to the initial interaction of H2 O2 with the heme iron [20,21]) were diminished relative to wtKatG. An increase (34-fold) in the dissociation constant for CN was also observed.1 In these properties, KatGDFG resembles the N153A and N153D Synechocystis KatG variants [20].
1
Li, Y., and Goodwin, D.C., unpublished observations.
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Given the consecutive appearance of Asp 135 and Asn 136 in catalase–peroxidase sequences, it is reasonable to suggest that H-bonding interactions involving each of them may be disrupted by structural modification such as that described here. Indeed, the ascending and descending limbs of the FG insertion run roughly parallel to the Hbonding network outlined by His 106 (the distal histidine), Asn 135, and Leu 130 (carbonyl oxygen) (Fig. 2). Roles of the FG insertion in reactions with reducing substrates The dramatic increase in kABTS observed for KatGDFG is also noteworthy. One explanation is that removal of the FG insertion increases access to the heme edge for electron donors, facilitating electron transfer. Catalase– peroxidases have only a narrow aperture through which substrates (especially peroxides) enter the active site. The N-terminal (P277-S279) and C-terminal (G306-T311) ends of the FG insertion form a significant portion of the barrier separating the bulk solvent from the active site. It is reasonable to expect that the absence of this barrier would enhance electron transfer in the reductive steps of the peroxidase cycle. In principle, one would expect that enhanced electron transfer would be evident from kABTS and kcat obtained with respect to ABTS. However, should some step other than ABTS oxidation be rate-determining, compound I formation for example, only an increase in kABTS would be predicted concomitant with the decrease in apparent KM . Interestingly, the interaction of ABTS with KatGDFG results in inhibition of peroxidase activity at moderate concentrations. The enhanced access of the reducing substrate to the active site may, at these concentrations, interfere with access of other substrates to the enzyme active site. In summary, we have used a novel procedure for deletion mutagenesis to eliminate a catalase–peroxidaseunique structural feature, the FG insertion. The resulting variant, KatGDFG , has very little catalase activity but retains peroxidase activity comparable to that of the wild-type enzyme. The selective loss of catalase activity is observed despite the peripheral position of the FG insertion relative to the active site. This indicates that the FG insertion has a role in fine-tuning the catalase– peroxidase active site for its unique bifunctionality. The spectral and kinetic properties of KatGDFG suggest that it supports hydrogen-bonded networks that are integral to catalase activity and regulates access of peroxidatic electron donors to the active site.
Acknowledgments The authors thank Dr. Holly Ellis for helpful discussion. Funds were provided by the American Chemical Society’s Petroleum Research Fund and Auburn University.
975
References [1] P.C. Loewen, Bacterial catalases, in: J.G. Scandalios (Ed.), Oxidative Stress and The Molecular Biology of Antioxidant Defenses, Cold Spring Harbor Laboratory Press, Plainview, NY, 1997, pp. 273–308. [2] A. Claiborne, I. Fridovich, Purification of the o-dianisidine peroxidase from Escherichia coli B. Physicochemical characterization and analysis of its dual catalatic and peroxidatic activities, J. Biol. Chem. 254 (1979) 4245–4252. [3] Y. Zhang, B. Heym, B. Allen, D. Young, S. Cole, The catalaseperoxidase gene and isoniazid resistance of Mycobacterium tuberculosis, Nature 358 (1992) 591–593. [4] A. Rattan, A. Kalia, N. Ahmad, Multidrug-resistant Mycobacterium tuberculosis: molecular perspectives, Emerg. Infect. Dis. 4 (1998) 195–209. [5] E. Garcia, Y.A. Nedialkov, J. Elliott, V.L. Motin, R.R. Brubaker, Molecular characterization of KatY (antigen 5), a thermoregulated chromosomally encoded catalase-peroxidase of Yersinia pestis, J. Bacteriol. 181 (1999) 3114–3122. [6] W. Brunder, H. Schmidt, H. Karch, KatP, a novel catalaseperoxidase encoded by the large plasmid of enterohaemorrhagic Escherichia coli O157:H7, Microbiology 142 (1996) 3305–3315. [7] P. Bandyopadhyay, H.M. Steinman, Catalase-peroxidases of Legionella pneumophila: cloning of the katA gene and studies of KatA function, J. Bacteriol. 182 (2000) 6679–6686. [8] J. Amemura-Maekawa, S. Mishima-Abe, F. Kura, T. Takahashi, H. Watanabe, Identification of a novel periplasmic catalaseperoxidase KatA of Legionella pneumophila, FEMS Microbiol. Lett. 176 (1999) 339–344. [9] X. Carpena, S. Loprasert, S. Mongkolsuk, J. Switala, P.C. Loewen, I. Fita, Catalase-peroxidase KatG of Burkholderia pseudomallei at 1.7A resolution, J. Mol. Biol. 327 (2003) 475–489. [10] Y. Yamada, T. Fujiwara, T. Sato, N. Igarashi, N. Tanaka, The 2.0 A crystal structure of catalase-peroxidase from Haloarcula marismortui, Nat. Struct. Biol. 9 (2002) 691–695. [11] M. Zamocky, G. Regelsberger, C. Jakopitsch, C. Obinger, The molecular peculiarities of catalase-peroxidases, FEBS Lett. 492 (2001) 177–182. [12] K.G. Welinder, Bacterial catalase-peroxidases are gene duplicated members of the plant peroxidase superfamily, Biochim. Biophys. Acta 1080 (1991) 215–220. [13] C.L. Varnado, K.M. Hertwig, R. Thomas, J.K. Roberts, D.C. Goodwin, Properties of a novel periplasmic catalase-peroxidase from Escherichia coli O157:H7, Arch. Biochem. Biophys. 421 (2004) 166–174. [14] J.K. Falk, in: K.M. Smith (Ed.), Porphyrins and Metalloporphyrins, Elsevier Publishing, New York, 1964, pp. 804–807. [15] C.L. Varnado, D.C. Goodwin, System for the expression of recombinant hemoproteins in Escherichia coli, Protein Expr. Purif. 35 (2004) 76–83. [16] H.B. Dunford, J.S. Stillman, On the function and mechanism of action of peroxidases, Coord. Chem. Rev. 19 (1976) 187–251. [17] C. Jakopitsch, M. Auer, A. Ivancich, F. Ruker, P.G. Furtmuller, C. Obinger, Total conversion of bifunctional catalase-peroxidase (KatG) to monofunctional peroxidase by exchange of a conserved distal side tyrosine, J. Biol. Chem. 278 (2003) 20185–20191. [18] A. Hillar, B. Peters, R. Pauls, A. Loboda, H. Zhang, A.G. Mauk, P.C. Loewen, Modulation of the activities of catalase-peroxidase HPI of Escherichia coli by site-directed mutagenesis, Biochemistry 39 (2000) 5868–5875. [19] L.J. Donald, O.V. Krokhin, H.W. Duckworth, B. Wiseman, T. Deemagarn, R. Singh, J. Switala, X. Carpena, I. Fita, P.C. Loewen, Characterization of the catalase-peroxidase KatG from Burkholderia pseudomallei by mass spectrometry, J. Biol. Chem. 278 (2003) 35687–35692.
976
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
[20] C. Jakopitsch, M. Auer, G. Regelsberger, W. Jantschko, P.G. Furtmuller, F. Ruker, C. Obinger, The catalytic role of the distal site asparagine-histidine couple in catalase-peroxidases, Eur. J. Biochem. 270 (2003) 1006–1013. [21] C. Jakopitsch, M. Auer, G. Regelsberger, W. Jantschko, P.G. Furtmuller, F. Ruker, C. Obinger, Distal site aspartate is essential in
the catalase activity of catalase-peroxidases, Biochemistry 42 (2003) 5292–5300. [22] G. Regelsberger, C. Jakopitsch, M. Engleder, F. Ruker, G.A. Peschek, C. Obinger, Spectral and kinetic studies of the oxidation of monosubstituted phenols and anilines by recombinant Synechocystis catalaseperoxidase compound I, Biochemistry 38 (1999) 10480–10488.
Vital roles of an interhelical insertion in catalase–peroxidase bifunctionalityq,qq Yongjiang Li and Douglas C. Goodwin* Department of Chemistry and Program in Cell and Molecular Biosciences, Auburn University, Auburn, AL 36849-5312, USA Received 25 March 2004 Available online 7 May 2004
Abstract The loop connecting the F and G helices of catalase–peroxidases contains a 35 amino acid structure (the FG insertion) that is absent from monofunctional peroxidases. These two groups of enzymes share highly similar active sites, yet the monofunctional peroxidases lack appreciable catalase activity. Thus, the FG insertion may serve a role in catalase–peroxidase bifunctionality, despite its peripheral location relative to the active site. We produced a variant of Escherichia coli catalase–peroxidase (KatG) lacking its FG insertion (KatGDFG ). Absorption spectra indicated the heme environment of KatGDFG was highly similar to wild-type KatG, but the variant retained only 0.2% catalase activity. In contrast, the deletion reduced peroxidase activity by only 50%. Kinetic parameters for the peroxidase and residual catalase activities of KatGDFG as well as pH dependence studies suggested that the FG insertion supports hydrogen-bonded networks critical for reactions involving H2 O2 . The structure also appears to regulate access of electron donors to the active site. Ó 2004 Elsevier Inc. All rights reserved. Keywords: KatG; Deletion mutagenesis; Heme; Catalase–peroxidase; Cytochrome c peroxidase; ABTS; Synechocystis KatG
Catalase–peroxidases are heme enzymes that use a single active site to catalyze both catalase and peroxidase reactions [1,2]. These enzymes have generated much interest because catalase–peroxidase from Mycobacterium tuberculosis is essential for the activation of the front-line antitubercular agent, isoniazid [3]. Indeed, it has been estimated that over 70% of isoniazid-resistant strains of M. tuberculosis carry mutations that alter the function of catalase–peroxidase [4]. Furthermore, several highly virulent pathogens carry unique periplasmic catalase–peroxidases that non-pathogenic relatives of these organisms lack, suggesting that these catalase– peroxidases may be virulence factors [5–8]. Clearly, the
q Funds to support this research were provided by the Petroleum Research Fund of the American Chemical Society (PRF 38802-G4) and Auburn University. qq Abbreviations: KatG, E. coli catalase–peroxidase; KatGDFG, 0 KatG lacking FG insertion; ABTS, 2,2 -azino-bis(3-ethylbenz-thiazoline-6-sulfonic acid); CT, charge transfer transition. * Corresponding author. Fax: 1-334-844-6959. E-mail address: [email protected] (D.C. Goodwin).
0006-291X/$ - see front matter Ó 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2004.04.130
evaluation of catalase–peroxidase structure and function has important ramifications. The peroxidase and catalase catalytic cycles both begin with the oxidation of the ferric heme by two electrons by a hydroperoxide (typically H2 O2 ) to form the ferryl-oxo porphyrin/protein radical intermediate known as compound I. The catalase cycle is completed by the reduction of compound I to the ferric enzyme by H2 O2 , resulting in the production of molecular oxygen. Conversely, in the peroxidatic cycle, compound I is typically reduced in two sequential one-electron steps by an exogenous reducing substrate. Two equivalents of substrate radical are generated and the enzyme is reduced to the ferric state via the ferryl-oxo intermediate known as compound II (Fig. 1). In spite of their efficient catalase activity, catalase– peroxidases bear no sequential or structural resemblance to monofunctional catalases. Conversely, the catalase– peroxidases are highly similar to the monofunctional peroxidases. Indeed, two recent crystal structures of catalase–peroxidases demonstrate that most of the major amino acid side chains in the active site are virtually
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Fig. 1. Catalytic cycle of catalase–peroxidases. A typical exogenous peroxidase reducing substrate as (AH) and its corresponding radical (A ) are also shown.
superimposable on those of cytochrome c peroxidase [9,10]. Nevertheless, cytochrome c peroxidase and other monofunctional peroxidases lack appreciable catalase activity. Given the high similarity of the monofunctional peroxidase and catalase–peroxidase active sites, it is reasonable to suggest that protein structures external to the catalase–peroxidase active site have an important role in fine-tuning the active site for its bifunctional abilities. In support of this hypothesis, comparisons of the protein structures and amino acid sequences of these two groups of enzymes reveal three structural features that are unique to the catalase–peroxidases. These are two interhelical insertions (35 amino acids each) and a C-terminal domain (300 amino acids) [11,12]. All three of these structures are peripheral to the catalase–peroxidase active site [9,10]. One of these interhelical insertions appears within a loop that connects the F and G helices of the catalase– peroxidase N-terminal domain (Fig. 2). Thus, we refer
971
to this structure as the FG insertion. Because this structure is uniformly absent from monofunctional peroxidases, it may have an important role in catalase– peroxidase bifunctionality. However, until the present time, the function of neither the FG insertion nor any residue composing it has been investigated. We employed a novel deletion mutagenesis procedure to generate a clone for recombinant Escherichia coli catalase–peroxidase (KatG) lacking its FG insertion (KatGDFG ). This variant was expressed, isolated, and characterized. Relative to wild-type KatG (wtKatG), KatGDFG retained appreciable peroxidase activity but very little catalase activity, demonstrating that this structure is essential for the unique bifunctional capabilities of catalase–peroxidases. Additionally, the kinetic parameters for the remaining peroxidase activity as well as the spectral properties of KatGDFG have provided valuable information on specific contributions of this structure to catalase–peroxidase function.
Materials and methods Reagents. All materials for cloning, mutagenesis, protein expression, and protein purification were purchased as previously delineated [13]. Hydrogen peroxide (30%) and hemin were purchased from Sigma (St. Louis, MO). All buffers and media were prepared using water purified through a Millipore Q-PakII system (18.2 MX/cm resistivity). Cloning and expression of wtKatG and KatGDFG . To generate the expression constructs for KatGDFG , we developed a procedure adapted from Seamless cloning (Stratagene). The plasmid for expression of recombinant His-tagged E. coli KatG (pKatG1) was used as a template. Primers were designed to: (1) amplify pKatG1 excluding the codons corresponding to the targeted deletion (Pro 277–Thr 311), (2) include a restriction site for Eam1104 I, and (3) yield compatible overhanging sequences upon digestion with Eam1104 I. Because Eam1104 I is a type IIS endonuclease, it cleaves target DNA downstream of its recognition sequence. Thus, the resulting overhanging sequences can be designed for religation such that only the codons corresponding to the targeted deletion are removed. No introduction
Fig. 2. Position of the FG insertion relative to key amino acids within the KatG active site. (A) Maximizes visibility of key active site side chains and their positions relative to the a carbons of the FG insertion (ribbon). The positions of N-terminus (P277) and C-terminus (T311) of the FG insertion are indicated. Two important H-bonds are shown as dark lines. (B) Represents a 120° leftward rotation of the structure. The N-terminus of the FG insertion (P277) is indicated. All numbering is according to E. coli KatG. Coordinates are from the B. pseudomallei KatG structure (1MWV) [9].
972
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
of new restriction sites or extra codons is involved, nor is the removal of codons outside the target sequence required. We used primers ECCP D201 (TAC ATC TCT TCT AGA ACC GGC ACC GTG GGT TTT ACC CAG) and ECCP D202 (GAT GCC TCT TCC TCT GGT CTG GAA GTA GTC TGG). The Eam1104 I recognition sequence is underlined, and the overhanging sequence generated upon hydrolysis by Eam1104 I is in bold. The PCR product was digested with Eam1104 I and ligated using T4 DNA ligase (in the presence of Eam1104 I). The ligated plasmid was used to transform E. coli (XL-1 Blue). Colonies were selected on the basis of ampicillin resistance, and candidate plasmids were verified by DNA sequence analysis to ensure the accuracy of the mutagenesis procedure. As we have done with pKatG1, the E. coli expression host (BL-21 [DE3] pLysS) was transformed with pKatGDFG . Expression of wtKatG and KatGDFG was accomplished as described for another catalase–peroxidase [13]. Purification of wtKatG and KatGDFG . All cell pellets were resuspended in Bugbuster reagent supplemented with 0.1 mM phenylmethylsulfonyl fluoride. Following homogenization, benzonase (250 U) was added, and the mixture was incubated at 23 °C with gentle stirring for 1 h. The cell lysate was then centrifuged at 16,000g. For wtKatG and KatGDFG , the supernatant was loaded onto a Ni– NTA column by recirculating the solution through the column bed overnight (1 mL/min). The column was then washed with buffer A (50 mM phosphate buffer, pH 8.0; 200 mM NaCl) supplemented with 2 mM imidazole. A second wash was then performed with buffer A supplemented with 20 mM imidazole. Finally, protein was eluted off the column with buffer A supplemented with 200 mM imidazole. Excess imidazole was then removed by dialysis or gel filtration chromatography. Enzyme reconstitution. We have observed that wtKatG and KatGDFG can be reconstituted with hemin following purification. Titration of each of the apoproteins with hemin results in a linear increase in activity up to 1 equivalent. Addition of hemin beyond this point does not result in any additional activity. To minimize contribution of free or adventitiously bound hemin to spectral or kinetic measurements, each protein was reconstituted with 0.75 equivalents hemin. Hemin concentration was determined by the method of Falk [14]. Enzyme activities and UV–visible absorption spectra. Catalase and peroxidase activities as well as UV–visible absorption spectra were obtained and evaluated as described previously for another catalase– peroxidase [13].
Results Mutagenesis, expression, and isolation of KatGDFG We developed a procedure to specifically delete the FG insertion from E. coli catalase–peroxidase (KatG)
(see Materials and methods). Based on sequence alignments of catalase–peroxidases in comparison to those for other class I peroxidases (i.e., cytochrome c peroxidase and ascorbate peroxidases) [11,12], we designed primers to seamlessly eliminate the codons corresponding to Pro 277–Thr 311 (i.e., the FG insertion) (Fig. 2). The resulting variant, KatGDFG , was expressed in a T7-based system and isolated in a soluble form. Electrophoretic migration of the KatGDFG monomer evaluated by SDS–PAGE was consistent with elimination of the targeted 35 amino acid segment (data not shown). UV–visible spectra of wtKatG and KatGDFG Following reconstitution with hemin (see Materials and methods), we recorded UV–visible absorption spectra for the ferric, ferrous, and ferri-cyano forms of KatGDFG (Fig. 3). For ferric KatGDFG , the Soret maximum (408 nm) and charge transfer maxima (501 and 637 nm) are consistent with a mixture of states dominated by pentacoordinate and hexacoordinate high-spin heme iron. A very weak shoulder at 535 nm suggests a minor contribution from hexacoordinate low-spin heme. The pentacoordinate high-spin species dominates in the ferrous state, as indicated by a Soret maximum of 438 nm in addition to a strong b band at 561 nm and a relatively weak a band at 590 nm. All spectra for KatGDFG were highly similar to those for wtKatG (Table 1), suggesting that removal of the FG insertion did little to disrupt the immediate environment around the heme group. The most notable spectral difference between wtKatG and KatGDFG was the relatively low molar absorptivity of the ferric state. One possible explanation would be that a portion of the hemin from reconstitution remained unbound by the enzyme. However, we have recently been able to express and isolate KatGDFG in its holoenzyme form using an expression system developed in our laboratory [15]. We observe spectral and catalytic properties highly similar to those of the reconstituted enzyme (data not shown).
Fig. 3. Absorption spectra recorded for KatGDFG . Spectra were obtained for ferric (bold solid line), ferri-cyano (dotted line), and ferrous KatGDFG (dashed line). (A) shows these spectra from 350 to 500 nm, and (B) shows them from 475 to 725 nm. Ferri-cyano KatGDFG was generated by the addition of 2 mM NaCN to ferric KatGDFG . Ferrous KatGDFG was generated by the addition of a few grains of solid dithionite to ferric KatGDFG . All spectra were recorded at 23 °C in 100 mM phosphate, pH 7.0.
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
973
Table 1 Absorption characteristics of wtKatG and the KatGDFG varianta Absorption maxima [nm] (absorptivity [mM1 cm1 ])
Enzyme
Soret
b
a
CT2b
CT1c
Ferric wtKatG KatGDFG
408 (120.7) 408 (77.4)
—d
—
—
—
502 (17.5) 501 (10.9)
629 (9.7) 637 (4.0)
Ferri-cyano wtKatG KatGDFG
423 (96.1) 423 (86.9)
542 (15.6) 541 (10.8)
—
—
—
—
—
—
Ferrous wtKatG KatGDFG
439 (79.8) 438 (88.2)
561 (15.1) 560 (13.4)
581 (10.7) 590 (8.5)
—
—
—
—
a
All spectra were collected in 100 mM phosphate buffer, pH 7.0, 23 °C. b CT2 short wavelength charge transfer transition (usually between 490 and 510 nm). c CT1 long wavelength charge transfer transition (usually between 600 and 650 nm). d Transitions that were absent or too weak to make unequivocal assignments.
Table 2 Steady-state kinetic parameters for catalase and peroxidase activities of wtKatG and KatGDFG Activitya (substrate)
Enzyme wtKatG
KatGDFG
b
Catalase (H2 O2 ) KM (mM) kcat (s1 ) kcat =KM (M1 s1 ) Peroxidase (H2 O2 )c KM (mM) kcat (s1 ) kH2 O2 (M1 s1 ) Peroxidase (ABTS) KM (mM) kcat (s1 ) kABTS (M1 s1 )
4.0 0.4 (1.2 0.4) 104 (3.1 0.5) 106
4.5 0.7 26 4 (5.7 0.2) 103
0.83 0.07 58 3 (7.1 0.9) 104
1.5 0.1 26 1 (1.8 0.1) 104
0.23 0.05 52 8 (2.2 0.2) 105
0.012 0.001 26 2 (2.2 0.1) 106
a
All assays were performed at 23 °C. All catalase assays were carried out in 100 mM phosphate buffer, pH 7.0. c All peroxidase assays were carried out in 50 mM acetate buffer, pH 5.0. b
Catalase and peroxidase activities of wtKatG and KatGDFG Consistent with previous reports [13], our preparation of wtKatG showed substantial catalase activity (Table 2). Removal of the FG insertion produced a precipitous reduction in catalase activity, decreasing the apparent second-order rate constant by a factor of 550 relative to wtKatG. This effect was due primarily to a decrease in kcat from 1.2 104 s1 for wtKatG to 26 s1 for KatGDFG . The apparent KM for H2 O2 as measured from catalase activity was unchanged. In addition to the loss of activity, shifts in pH profile were observed. The pH optimum shifted from 6.5 to 7.0, and although a similar dependence of relative rate on pH was observed at high pH values, the catalase activity of KatGDFG
Fig. 4. Effect of pH on catalase activities of wtKatG and KatGDFG . Catalase activities of wtKatG (s) and KatGDFG (j) were evaluated in the presence of 20 mM H2 O2 and 100 mM phosphate at 23 °C.
decreased more sharply below 7.0 than observed for wtKatG (Fig. 4). KatGDFG retained appreciable peroxidase activity (Fig. 5). With respect to H2 O2 or ABTS, kcat values (26 s1 in each case) were roughly half of that observed for wtKatG (Table 2). A moderate increase (twofold) in the apparent KM for H2 O2 was observed. Conversely, the apparent KM for the reducing substrate, ABTS, decreased by a factor of 10 upon removal of the FG insertion. The kinetic parameters obtained with respect to H2 O2 have been used to describe reactions leading to the formation of compound I [16]. Elimination of the FG insertion led to a fourfold decrease in apparent second-order rate constant (kH2 O2 ) governing these steps. On the other hand, kinetic parameters obtained with respect to the reducing substrate (in this case ABTS) describe the reactions that result in conversion of compound I back to ferric enzyme. The apparent second-order rate constant (kABTS ) governing these steps increased by a factor of 10 (Table 2). In addition to a dramatically reduced KM for ABTS, we also observed ABTS-dependent inhibition of KatGDFG peroxidase activity at concentrations above
974
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Fig. 5. Comparison of the peroxidase activities of wtKatG and KatGDFG . The effects of H2 O2 (A) and ABTS (B) concentrations on the rate of ABTS oxidation by wtKatG (d) and KatGDFG () were evaluated. (A) ABTS concentrations were selected to obtain optimum peroxidase activity: 1 mM for wtKatG and 0.1 mM for KatGDFG . (B) All reactions contained 10 mM H2 O2 . All reactions were carried out at 23 °C in 50 mM acetate buffer, pH 5.0. The [E]T values were calculated based on heme content.
0.1 mM (Fig. 5B). At 1 mM ABTS, a 33% decrease in activity from the recorded maximum was observed. No inhibition was apparent for wtKatG over this same concentration range. Likewise, the peroxidase activity of neither wtKatG nor KatGDFG was sensitive to peroxidedependent inhibition (Fig. 5A).
Discussion Catalase–peroxidases have the capacity to catalyze two reactions using a single active site. Remarkably, this bifunctional active site is highly similar to that of the less versatile monofunctional peroxidases. Catalase–peroxidases all have an insertion (35 amino acids) that extends the loop connecting the F and G helices relative to that of monofunctional peroxidases. This FG insertion is peripheral to the active site. Nevertheless, our results demonstrate that this structure is essential for catalase–peroxidase bifunctionality. Furthermore, spectra recorded for KatGDFG showed only subtle changes in heme environment, indicating that observed effects on catalase activity were not due to dramatic changes in the coordination sphere of the heme iron. Roles of the FG insertion in reactions with H2 O2 The catalase cycle involves two principal reactions with H2 O2 , compound I formation and compound I reduction. Recent site-directed mutagenesis studies have identified three mechanisms by which the catalase activity of catalase–peroxidase is supported by the active site. Each participant in a novel covalent link (Trp 105, Tyr 226, and Met 252 [E. coli KatG numbering]) has been shown in at least one catalase–peroxidase to be essential for catalase but not peroxidase activity [17–19]. Our data to date suggest KatGDFG still forms this covalent structure. A hydrogen-bonded network involving Asp 135 and Asn 136 also appears to be important. The analogous residues in Synechocystis KatG (Asp 152 and Asn 153) are essential for full catalase activity. Interestingly, replacement of each of these residues
diminishes catalase activity, apparently by differing mechanisms. Disruption of a hydrogen bond between the distal histidine and Asn 153 primarily affects compound I formation [20], whereas replacement of Asp 152 disrupts compound I reduction [21]. The rate-determining step for catalase activity is thought to be compound I formation [22]. The change in pH profile for KatGDFG is consistent with either a change in the rate-determining step or a dramatic shift in the pKa s associated with it. The pH profile is reminiscent of that observed for D152N Synechocystis KatG, where H2 O2 -dependent compound I reduction is the principal reaction affected by the substitution [21]. As observed for D152N Synechocystis KatG, KatGDFG also shows minimal effects on peroxidase activity as compared to the substantial reduction in catalase activity. Indeed, our kABTS value for KatGDFG suggests that, if anything, peroxidatic reduction of compound I is enhanced by elimination of the FG insertion. Indirect evidence suggests that compound I formation has also been diminished to some extent. Our kH2 O2 values derived from peroxidase activity decreased fourfold relative to wtKatG. This is in spite of the near absent catalase activity. Normally, consumption of H2 O2 by compound I is expected to diminish observed peroxidase activity because it is “non-productive” consumption of H2 O2 (i.e., it does not result in oxidation of ABTS or other reducing substrates). Indeed, catalasenegative catalase–peroxidase variants often show an increase in peroxidase activity [17,18,21]. To the present time, we have been unable to directly observe H2 O2 dependent KatGDFG compound I formation by stoppedflow. However, rates of CN binding (often monitored as a reaction analogous to the initial interaction of H2 O2 with the heme iron [20,21]) were diminished relative to wtKatG. An increase (34-fold) in the dissociation constant for CN was also observed.1 In these properties, KatGDFG resembles the N153A and N153D Synechocystis KatG variants [20].
1
Li, Y., and Goodwin, D.C., unpublished observations.
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
Given the consecutive appearance of Asp 135 and Asn 136 in catalase–peroxidase sequences, it is reasonable to suggest that H-bonding interactions involving each of them may be disrupted by structural modification such as that described here. Indeed, the ascending and descending limbs of the FG insertion run roughly parallel to the Hbonding network outlined by His 106 (the distal histidine), Asn 135, and Leu 130 (carbonyl oxygen) (Fig. 2). Roles of the FG insertion in reactions with reducing substrates The dramatic increase in kABTS observed for KatGDFG is also noteworthy. One explanation is that removal of the FG insertion increases access to the heme edge for electron donors, facilitating electron transfer. Catalase– peroxidases have only a narrow aperture through which substrates (especially peroxides) enter the active site. The N-terminal (P277-S279) and C-terminal (G306-T311) ends of the FG insertion form a significant portion of the barrier separating the bulk solvent from the active site. It is reasonable to expect that the absence of this barrier would enhance electron transfer in the reductive steps of the peroxidase cycle. In principle, one would expect that enhanced electron transfer would be evident from kABTS and kcat obtained with respect to ABTS. However, should some step other than ABTS oxidation be rate-determining, compound I formation for example, only an increase in kABTS would be predicted concomitant with the decrease in apparent KM . Interestingly, the interaction of ABTS with KatGDFG results in inhibition of peroxidase activity at moderate concentrations. The enhanced access of the reducing substrate to the active site may, at these concentrations, interfere with access of other substrates to the enzyme active site. In summary, we have used a novel procedure for deletion mutagenesis to eliminate a catalase–peroxidaseunique structural feature, the FG insertion. The resulting variant, KatGDFG , has very little catalase activity but retains peroxidase activity comparable to that of the wild-type enzyme. The selective loss of catalase activity is observed despite the peripheral position of the FG insertion relative to the active site. This indicates that the FG insertion has a role in fine-tuning the catalase– peroxidase active site for its unique bifunctionality. The spectral and kinetic properties of KatGDFG suggest that it supports hydrogen-bonded networks that are integral to catalase activity and regulates access of peroxidatic electron donors to the active site.
Acknowledgments The authors thank Dr. Holly Ellis for helpful discussion. Funds were provided by the American Chemical Society’s Petroleum Research Fund and Auburn University.
975
References [1] P.C. Loewen, Bacterial catalases, in: J.G. Scandalios (Ed.), Oxidative Stress and The Molecular Biology of Antioxidant Defenses, Cold Spring Harbor Laboratory Press, Plainview, NY, 1997, pp. 273–308. [2] A. Claiborne, I. Fridovich, Purification of the o-dianisidine peroxidase from Escherichia coli B. Physicochemical characterization and analysis of its dual catalatic and peroxidatic activities, J. Biol. Chem. 254 (1979) 4245–4252. [3] Y. Zhang, B. Heym, B. Allen, D. Young, S. Cole, The catalaseperoxidase gene and isoniazid resistance of Mycobacterium tuberculosis, Nature 358 (1992) 591–593. [4] A. Rattan, A. Kalia, N. Ahmad, Multidrug-resistant Mycobacterium tuberculosis: molecular perspectives, Emerg. Infect. Dis. 4 (1998) 195–209. [5] E. Garcia, Y.A. Nedialkov, J. Elliott, V.L. Motin, R.R. Brubaker, Molecular characterization of KatY (antigen 5), a thermoregulated chromosomally encoded catalase-peroxidase of Yersinia pestis, J. Bacteriol. 181 (1999) 3114–3122. [6] W. Brunder, H. Schmidt, H. Karch, KatP, a novel catalaseperoxidase encoded by the large plasmid of enterohaemorrhagic Escherichia coli O157:H7, Microbiology 142 (1996) 3305–3315. [7] P. Bandyopadhyay, H.M. Steinman, Catalase-peroxidases of Legionella pneumophila: cloning of the katA gene and studies of KatA function, J. Bacteriol. 182 (2000) 6679–6686. [8] J. Amemura-Maekawa, S. Mishima-Abe, F. Kura, T. Takahashi, H. Watanabe, Identification of a novel periplasmic catalaseperoxidase KatA of Legionella pneumophila, FEMS Microbiol. Lett. 176 (1999) 339–344. [9] X. Carpena, S. Loprasert, S. Mongkolsuk, J. Switala, P.C. Loewen, I. Fita, Catalase-peroxidase KatG of Burkholderia pseudomallei at 1.7A resolution, J. Mol. Biol. 327 (2003) 475–489. [10] Y. Yamada, T. Fujiwara, T. Sato, N. Igarashi, N. Tanaka, The 2.0 A crystal structure of catalase-peroxidase from Haloarcula marismortui, Nat. Struct. Biol. 9 (2002) 691–695. [11] M. Zamocky, G. Regelsberger, C. Jakopitsch, C. Obinger, The molecular peculiarities of catalase-peroxidases, FEBS Lett. 492 (2001) 177–182. [12] K.G. Welinder, Bacterial catalase-peroxidases are gene duplicated members of the plant peroxidase superfamily, Biochim. Biophys. Acta 1080 (1991) 215–220. [13] C.L. Varnado, K.M. Hertwig, R. Thomas, J.K. Roberts, D.C. Goodwin, Properties of a novel periplasmic catalase-peroxidase from Escherichia coli O157:H7, Arch. Biochem. Biophys. 421 (2004) 166–174. [14] J.K. Falk, in: K.M. Smith (Ed.), Porphyrins and Metalloporphyrins, Elsevier Publishing, New York, 1964, pp. 804–807. [15] C.L. Varnado, D.C. Goodwin, System for the expression of recombinant hemoproteins in Escherichia coli, Protein Expr. Purif. 35 (2004) 76–83. [16] H.B. Dunford, J.S. Stillman, On the function and mechanism of action of peroxidases, Coord. Chem. Rev. 19 (1976) 187–251. [17] C. Jakopitsch, M. Auer, A. Ivancich, F. Ruker, P.G. Furtmuller, C. Obinger, Total conversion of bifunctional catalase-peroxidase (KatG) to monofunctional peroxidase by exchange of a conserved distal side tyrosine, J. Biol. Chem. 278 (2003) 20185–20191. [18] A. Hillar, B. Peters, R. Pauls, A. Loboda, H. Zhang, A.G. Mauk, P.C. Loewen, Modulation of the activities of catalase-peroxidase HPI of Escherichia coli by site-directed mutagenesis, Biochemistry 39 (2000) 5868–5875. [19] L.J. Donald, O.V. Krokhin, H.W. Duckworth, B. Wiseman, T. Deemagarn, R. Singh, J. Switala, X. Carpena, I. Fita, P.C. Loewen, Characterization of the catalase-peroxidase KatG from Burkholderia pseudomallei by mass spectrometry, J. Biol. Chem. 278 (2003) 35687–35692.
976
Y. Li, D.C. Goodwin / Biochemical and Biophysical Research Communications 318 (2004) 970–976
[20] C. Jakopitsch, M. Auer, G. Regelsberger, W. Jantschko, P.G. Furtmuller, F. Ruker, C. Obinger, The catalytic role of the distal site asparagine-histidine couple in catalase-peroxidases, Eur. J. Biochem. 270 (2003) 1006–1013. [21] C. Jakopitsch, M. Auer, G. Regelsberger, W. Jantschko, P.G. Furtmuller, F. Ruker, C. Obinger, Distal site aspartate is essential in
the catalase activity of catalase-peroxidases, Biochemistry 42 (2003) 5292–5300. [22] G. Regelsberger, C. Jakopitsch, M. Engleder, F. Ruker, G.A. Peschek, C. Obinger, Spectral and kinetic studies of the oxidation of monosubstituted phenols and anilines by recombinant Synechocystis catalaseperoxidase compound I, Biochemistry 38 (1999) 10480–10488.