Peptidoglycan Modification by the Catalytic Domain of Streptococcus

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Peptidoglycan modification by the catalytic domain of Streptococcus pneumoniae OatA follows a ping-pong bi-bi mechanism of action David Sychantha, and Anthony John Clarke Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00301 • Publication Date (Web): 29 Mar 2018 Downloaded from http://pubs.acs.org on March 29, 2018

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Biochemistry

Peptidoglycan modification by the catalytic domain of Streptococcus pneumoniae OatA follows a pingpong bi-bi mechanism of action David Sychantha† and Anthony J. Clarke* Department of Molecular & Cellular Biology, University of Guelph, Guelph, ON N1G 2W1, Canada

* E-mail: [email protected]. Phone: +1-519-824-4120 ext. 54124

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ABSTRACT

Streptococcus pneumoniae amongst other Gram-positive pathogens produces O-acetylated peptidoglycan using the enzyme OatA. This process occurs through the transfer of an acetyl group from a donor to the hydroxyl group of an acceptor sugar. While it has been established that this process involves the extracellular, catalytic domain of OatA (SpOatAC), mechanistic insight is still unavailable. This study examined the enzymatic characteristics of SpOatAC-catalyzed reactions through analysis of both pre-steady and steady-state kinetics. Our findings clearly show that SpOatAC follows a ping-pong bi-bi mechanism of action involving a covalent acetyl-enzyme intermediate. The modified residue was verified to be the catalytic nucleophile, Ser438. The pH dependence of the enzyme kinetics revealed that a single ionizable group is involved consistent with the participation of a His residue. Single turn-over kinetics of esterase activity demonstrated that k2 ≫ k3, revealing that the rate-limiting step for the hydrolytic reaction was the breakdown of the acetyl-enzyme with a half-life of >1 min. The previous assignment of Asn491 as an oxyanion hole residue was also confirmed as its replacement with Ala resulted in a 50-fold decrease in catalytic efficiency relative wild-type SpOatAC. However, this loss of catalytic efficiency was mostly due to a large increase in KM, suggesting that Asn491 contributes more to substrate binding.


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Biochemistry

INTRODUCTION There has been an emerging interest in bacterial cell surface modification as a target for developing novel agents to overcome antibacterial resistance. This is because such alterations to the bacterial cell envelope are often needed for pathogenicity and/or improved fitness within a host organism.1,2 For example, maintaining the integrity of the peptidoglycan (PG) sacculus is essential for bacterial cell viability. However, the β(1→4) linkage between its repeating units of N-acetylmuramoyl (MurNAc) and N-acetylglucosaminyl (GlcNAc) residues is the target of lysozyme, a major enzyme of the eukaryotic innate immune system.3 To counter this, many bacterial pathogens persist within a host by O-acetylating the C6-OH of MurNAc which provides steric protection against lysozyme hydrolysis.4 Prior to the identification, characterization and, particularly, enhancement of inhibitors of cell wall modifying enzymes, detailed knowledge of their catalytic mechanisms is essential. Currently, the only polysaccharide O-acetyltransferase which has had its mechanism verified through detailed studies is PatB from Neisseria gonorrhoeae (NgPatB).5 Whereas the structure of NgPatB is still unknown, homology modelling predicted that it possesses an SGNH-hydrolase fold (atypical α/β-hydrolase fold) with a Ser-His-Asp catalytic triad.6 Indeed, these features appear to be common amongst the known structures of other polysaccharide Oacetyltransferases.7–10 Structure-guided mutagenesis studies involving these latter enzymes have also suggested that they may commonly follow a ping-pong bi-bi mechanism of action. However, definitive proof of this for these enzymes is still lacking. The O-acetylation of PG occurs as a post-synthetic modification following the transglycosylation of Lipid II precursors into the existing sacculus (reviewed in 4). In Grampositive bacteria, this modification is catalyzed by O-acetyltransferase A (OatA).11 This 69.2 ACS Paragon Plus Environment

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kDa integral membrane protein is predicted to be composed of two domains, an N-terminal membrane spanning domain and an extracellular C-terminal domain.12 The N-terminal domain is proposed to translocate acetyl groups form a cytoplasmic source across the cytoplasmic membrane for their transfer to PG catalyzed by the C-terminal O-acetyltransferase domain.4 Despite being recognized as a virulence factor in a number of Gram-positive pathogens, including

Staphylococcus

aureus,11,13,14

Enterococcus

faecalis15,16

and

Streptococcus

pneumoniae,17 little progress has been made in exploiting OatA for antibiotic development. This is likely due to the complexities of biochemically characterizing a membrane-bound enzyme that acts on an insoluble substrate such as PG. Recently however, we circumvented these issues by producing soluble recombinant forms of the C-terminal catalytic domains of the OatA orthologues from both S. aureus (SaOatAC) and S. pneumoniae (SpOatAC),10 and developing assays for their kinetic analyses using pseudosubstrates.10,18 Using these assays, we found that SaOatAC and SpOatAC catalyze both esterase and Oacetyltransferase reactions.10 Also, we solved the crystal structure of SpOatAC with high resolution. Although sharing minimal sequence identity to the well-characterized PatB5,6,18,19 (18.7 % identity and 39.1 % similarity between SpOatAC and NgPatB), SpOatAC also adopts an SGNH-hydrolase fold and it has a catalytic triad composed of Ser438, His571, and Asp568. Sitespecific replacements demonstrated these residues to be essential for catalysis and a mechanismbased inhibitor was used to further support the identification of Ser438 as the catalytic nucleophile. However, still lacking is a detailed kinetic analysis of OatA, and thus confirmation that it follows a ping-pong bi-bi reaction mechanism, as well as identification of its rate limiting step. Herein, we present both a pre-steady and steady-state kinetic analysis of SpOatAC using a pseudosubstrate donor and a variety of oligosaccharides as acceptors. By trapping the acetylACS Paragon Plus Environment

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Biochemistry

enzyme intermediate, we provide confirmation that Ser438 serves as the catalytic nucleophile. Together, these data help to explain why the esterase activity of OatA observed in vitro is limited, and that in vivo the enzyme would function solely as an O-acetyltransferase. MATERIALS AND METHODS Materials. Luria-Bertani medium was purchased from Difco (Detroit, MI), while acetone and acrylamide were obtained from Fisher Scientific (Nepean, ON). Sodium dodecyl sulfate (SDS) and 4-methylumbelliferyl acetate (4MU-Ac) were purchased from Sigma-Aldrich (Oakville, ON), and chitooligosaccharides were obtained from Megazyme (Bray, Ireland). Complete His-Tag purification resin for immobilized metal affinity chromatography (IMAC) was purchased from Roche (Mississauga, ON), and Source Q is a product of GE healthcare (Piscataway, NJ). Unless otherwise stated, all other chemicals were obtained from Bio Basic Inc. (Markham, ON). Engineering and Production of SpOatAC and the N491A Variant. SpOatAC from S. pneumoniae R6 covers residues 435-603 and it was engineered with an N-terminal His6-tag encoded in the pBAD-His A expression vector. This construct produced the plasmid pDSAC-81.8 The AsN491Ala variant was produced previously by site-directed mutagenesis to generate pDSAC-81-N491A.8 The wild-type and derivative proteins were produced and purified as previously described.8 As before, SpOatAC was overproduced in Escherichia coli BL21 in LB medium for 3 h using 0.2 % (w/v) arabinose as inducer. The protein was isolated from cell lysate using IMAC and dialyzed into 25 mM sodium phosphate buffer, pH 8. The sample was subsequently applied to a Source Q anion-exchange column and eluted with a 0–50 % gradient of 1 M NaCl in phosphate buffer over 50 mins at 1 mL/min. Fractions were analyzed for purity by SDS-polyacrylamide gel electrophoresis, pooled, and quantified using the calculated extinction coefficient (32,430 M-1·cm-1) at λ = 280 nm. ACS Paragon Plus Environment

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Steady State Kinetic Analyses. Analyses of enzyme-catalyzed hydrolysis of 4MU-Ac were made using a fluorometric assay (λex = 390 nm; λem = 420 nm) as described previously.10 To measure enzyme-catalyzed esterase activity, SpOatAC (1 µM) in 25 mM sodium phosphate buffer, pH 6.5 was incubated with varying amounts of 4MU-Ac dissolved in DMSO. Final volumes were 75 µL and the final concentration of DMSO was 5 % (v/v), which did not affect enzyme activity. Enzyme-catalyzed O-acetylation of acceptor ligands was performed using the same conditions, except the concentration of 4MU-Ac was fixed at 100 µM in reaction mixtures, while varying amounts of chitooligosaccharide (chitotetraose, GlcNAc4; chitopentaose, GlcNAc5; or chitohexaose, GlcNAc6.) were included. Transferase rates were corrected for the combined background rates of enzyme-catalyzed and spontaneous 4MU-Ac hydrolysis. The reactions were monitored in black 96-well microtiter plates using a BioTek Synergy H1 plate reader (BioTek Instruments, Inc., Winooski, VT) over 2 min at 25 ºC. Each reaction was performed in duplicate. To obtain the KM and kcat values, the data were fitted to the MichaelisMenten equations 1 and 2 for esterase and transferase activities, respectively, using non-linear regression with GraphPad prism 4,

kcat Et S K M  S

(1)

kcat Et  A   B K  A   K MA  B   AB

(2)

vo 

vo 

B M

where, A and B represent donor and acceptor substrates, respectively. Lineweaver-Burk analyses to determine the mechanism of action of SpOatAC were performed using the conditions described above. SpOatAC (1 µM) was mixed with reaction buffer containing 4, 2, 1, and 0.5 mM GlcNAc4. The reactions were initiated with the addition of 4MU-Ac at fixed concentrations of 80, 64, 32, and 16 µM. The effect of pH on the kinetics of


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Biochemistry

SpOatAC-catalyzed 4MU-Ac hydrolysis and acetyltransfer was determined using 50 mM citratephosphate buffer at pH 4.7–8.5. Experiments were performed at 25 ºC in duplicate. To obtain pKa values, Dixon-Webb plots20 were generated by fitting the data to eq. 3 using GraphPad Prism 4,

1

H

(3)

where (kcat)max is the maximum rate when the enzyme is in its optimal ionized form. Pre-Steady State Kinetic Analyses. Single turnover assays were performed using an Applied Photophysics stopped-flow fluorescence spectrophotometer (Leatherhead, UK) equipped with excitation and emission monochromators, and a constant-temperature circulating water bath set to 25 ºC. Experiments were performed by rapidly mixing the contents of two syringes, one loaded with 10 µM SpOatAC in reaction buffer and the second with varying concentrations of 4MU-Ac (500, 250, 125, 62.5, 32.15, and 16.3 µM) in reaction buffer. The burst amplitude was measured by fluorometry (λex = 390 nm; λem = 420 nm). The pseudo-first order reaction was fitted to eq. 4 to obtain kobs,

(4)

1

where P(t) is product formed over time (t) and A is the amplitude of the pre-steady state phase of reaction. To obtain k2 values, the kobs values were plotted as a function of 4MU-Ac concentration and fitted to eq. 5,

kobs 

where

K Macetyl 

k2  k1 k1

and

k S K  S 2 acetyl M

K M  K Macetyl

(5)

k3 k2  k3

To determine the concentration of acetyl-enzyme, the amplitude of the burst phase was obtained ACS Paragon Plus Environment

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by finding the y-intercept of the extrapolated steady-state phase. As implied by Scheme 1,

Scheme 1. Reaction pathway for hydrolysis of 4MU-Ac.

the k3 value was determined using eq. 6

k3 

vo EAc

(6)

where EAc is acetyl-enzyme, and the half-life of the acetyl-enzyme was calculated using eq. 7. t1/2 =

(7)

Trapping the Acetyl-SpOatAC Intermediate. Direct observation of the covalent acetylSpOatAC intermediate was achieved by real-time analysis of a reaction by liquid chromatography-mass spectrometry (LC-MS) using an Agilent 1200 HPLC interfaced with an Aglient UHD 6520 Q-TOF MS (Agilent Technologies Inc., Santa Clara, CA) housed in the Mass Spectrometry Facility of the Advanced Analysis Centre, University of Guelph. A reaction mixture was prepared in a MS-grade sample vial on ice. Using the conditions described above for 4MU-Ac hydrolysis, the reaction was initiated with the addition of 100 µM 4MU-Ac. A 2 µL sample of the mixture was immediately injected into a C18 column (Zorbax 300SB-C18, 1 × 50 mm; 5µm) equilibrated with 10 % acetonitrile in 0.1 % formic acid; the injection sequence took approximately 60 s to complete. Residual substrate and buffer were separated from the protein with the application of a linear gradient to 85 % acetonitrile in 0.1 % formic acid over 7 mins at 0.2 mL/min. The MS electrospray capillary voltage was maintained at 4.0 kV and the drying gas temperature at 350° C with a flow rate of 13 L/min. Nebulizer pressure was 40 psi and the fragmentor was set to 150. Nitrogen was used as both nebulizing and drying gas. The mass-to-charge ratio was scanned across the m/z range of 300-3200 m/z in 4GHz extended ACS Paragon Plus Environment

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Biochemistry

dynamic range positive-ion mode. The instrument was externally calibrated with the ESI TuneMix (Agilent). A control reaction without 4MU-Ac added was also performed to serve as the initial time point. Data analysis was performed using the MassHunter Qualitative Analysis Version B.06.00 software (Agilent). Deconvolution of the m/z spectrum was achieved using the Maximum Entropy algorithm within the BioConfirm software (Agilent). Identification of the site of O-acetylation was achieved through analysis of a tryptic digest of an acetone-quenched SpOatAC-catalyzed hydrolysis reaction. 4MU-Ac (100 µM final concentration) was added to 50 µL of SpOatAC (10 µM) in reaction buffer. After 1 min incubation at 25 C, the reaction was quenched by dilution (20-fold) in cold acetone and chilled to -80 ºC. The precipitated protein was resuspended in 20 µL 50 mM ammonium bicarbonate buffer, pH 7.5 and incubated with 20 µg/mL trypsin (10 µL) for 4 h at 37 C. The digest was subsequently analyzed by LC-MS/MS using the MS system described above. Peptides were separated on a C18 column (Aglilent AdvanceBio Peptide Map, 100 mm × 2.1 mm; 2.7 µm) equilibrated with 2 % acetonitrile in 0.1 % formic acid. Following sample injection, a linear gradient to 45 % acetonitrile in 0.1 % formic acid was applied over 40 min at 0.2 mL per min, followed by an increase to 55 % acetonitrile over 10 min; the first 2 mins of eluant were sent to waste and not the spectrometer. The mass spectrometer electrospray capillary voltage and nebulizer pressure were maintained as above. The fragmentor was set to 150 and nitrogen was used as both nebulizing and drying gas, and collision-induced gas. The mass-to-charge ratio was scanned across the m/z range of 300-2000 m/z in 4GHz (extended dynamic range positive-ion auto MS/MS mode). Three precursor ions per cycle were selected for fragmentation. The sample injection volume was 100 µl. Data analysis was performed using Mmass (www.mmass.org/).


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RESULTS Determination of Steady-State Kinetic Parameters for SpOatAC. Initial kinetic studies were performed to optimize the enzyme assay for SpOatAC using the pseudo acetyl-donor 4MUAc. This substrate was chosen to increase sensitivity and reduce background hydrolysis of a former assay, which was performed with the more labile donor, p-nitrophenyl acetate (pNPAc).10 To maintain solubility of this substrate, reactions required 5 % (v/v) DMSO (final concentration) as a co-solvent, which did not affect enzyme activity. Since SpOatAC can catalyze both esterase and transacetylation reactions, we first determined the kinetic parameters for 4MUAc hydrolysis in the absence of acceptor (Scheme 1). The reactions were consistent with Michaelis-Menten behavior and the kcat (0.011 s-1) and KM (13 μM) values for 4MU-Ac (Table 1) were ~1.4 and ~5-fold lower than those previously reported for pNP-Ac,10 respectively. The lower KM value for 4MU-Ac hydrolysis suggested that either binding of 4MU-Ac is tighter or the rate of acetyl-enzyme formation is more rapid. We then determined the kinetic parameters for transacetylations using chitooligosaccharides as acceptors with a fixed concentration of 4MU-Ac (Scheme 2). Compared to the previously reported kinetic parameters, which used pNP-Ac as donor substrate and ethanol (5% v/v) as a cosolvent,10 we observed ~3–5-fold lower KM values for the chitooligosaccharides (Table 1). To ensure that these differences were not a consequence of the specific choice of donor, we determined the kinetic parameters for GlcNAc5 using a fixed concentration of pNP-Ac but with DMSO as a co-solvent. The results confirmed that the observed KM value was comparable to that involving 4MU-Ac, indicating that the type of co-solvent used affects chitooligosaccharide binding. Consequently, the lower KM values resulted in overall higher kcat/KM values. Despite this, a positive correlation between kcat/KM and acceptor length was observed, which is consistent with the previously determined parameters10 (Table 1).


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Biochemistry

Table 1. Steady-state kinetic parameters for SpOatACa Parameters Acetyl Donor Acceptor b Donor Parameters 4MU-Ac Water 4MU-Ac GlcNAc4 4MU-Ac GlcNAc5 Acceptor Parametersc 4MU-Ac GlcNAc4 4MU-Ac GlcNAc5 4MU-Ac GlcNAc6 pNP-Ac GlcNAc6

kcat (s-1)

KM (mM)

kcat /KM (M-1s-1)

0.011±0.00032 ND ND

0.013±0.0011 ND ND

853±70 742±9 758±12

0.087±0.003 1.7±0.10 51±4 0.092±0.0005 0.87±0.10 106±15 0.126±0.004 0.53±0.06 238±029 0.186±0.009 0.95±0.10 195±22 a Reactions performed with 1 M enzyme in 25 mM sodium phosphate buffer, pH 6.5 containing 5 % (v/v) DMSO. b Donor concentration varied with acceptor concentration fixed at 10  KM (GlcNAc4, 18 mM; GlcNAc5, 8 mM). c Acceptor concentration varied with donor concentration fixed at 10  KM (4MU-Ac, 100 M; pNP-Ac, 500 M). ND, not detected.

Order of the SpOatAC-Catalyzed Reaction. We determined a series of kinetic parameters for GlcNAc4 as acceptor at fixed concentrations in which the 4MU-Ac concentrations were varied, and then vice-versa where the donor concentration was fixed while varying acceptor concentrations. The data were analyzed with Lineweaver-Burk plots, revealing a family of parallel lines consistent with a ping-pong bi-bi mechanism of action (Figure 1). According to this mechanism, the transfer of acetyl from 4MU-Ac to the enzyme will occur in the absence of the second substrate (Scheme 2).

Scheme 2. Reaction pathway for acetyl transfer from 4MU-Ac to a carbohydrate (CHO) acceptor.

Therefore, the rate of enzyme acetylation (k2) is expected to be independent of acceptor concentration. We then determined kinetic parameters for 4MU-Ac using single, fixed and saturating (×10 KM) concentrations of GlcNAc4 and GlcNAc5 as acceptors. Under these


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Figure 1. Determination of reaction order of SpOatAC by steady-state kinetic analysis. Lineweaver-Burk plots of initial rates of reaction of 1 μM SpOatAC in 25 mM sodium phosphate buffer, pH 6.5 at 37 C with (A) 4MU-Ac as acetyl donor fixed at concentrations of (□) 16 M, (○) 32 M, (■) 64 M, and (●) 80 M and chitotetraose as acceptor at the concentrations shown; and (B) 4MU-Ac as donor acetyl at the concentrations shown with fixed concentrations of chitotetraose as acceptor ligand fixed at (□) 0.5 mM, (○) 1 mM, (■) 2 mM, and (●) 4 mM. The data represent the average of three experiments; error bars hidden by symbols. Figure 2. Steady-state and pre-steady-state kinetic analyses of SpOatAC. A) Steady-state kinetics of SpOatAC-catalyzed turnover of 4MU-Ac in the absence (▼) and presence of 18 mM GlcNAc4 (○) and 8 mM GlcNAc5 (●). B) Representative presteady state time-course for a single turnover of 4MU-Ac hydrolysis in the (●) absence, and presence of (■) 1 μM and (▲) 2 μM SpOatAC. The dashed lines represent the interpolated slopes of the steadystate phase. The experimental data were fit to eq. 4 to obtain values of kobs. C) Determination of rate constants for 4MU-Ac hydrolysis. The slope of the line was used to determine k2/KMacetyl, and the data were fit to eq. 5 to estimate the value of k2.


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Biochemistry

conditions, release of 4MU did not follow Michaelis-Menten behavior, as the reactions could not be saturated (Figure 2A). However, we were able to obtain second-order rate constants (kcat/KM) (Table 1). These values were approximately equal to that of the kcat/KM for 4MU-Ac hydrolysis. Overall, these observations implied that 4MU-Ac is a poor substrate for SpOatAC and suggest that the rate of enzyme acetylation is proportional to 4MU-Ac concentration. Therefore, the overall reaction rate (kcat) is dependent on enzyme deacetylation, and hence k2 ≫ k3. Pre-steady state kinetic analysis of SpOatAC-catalyzed 4MU-Ac hydrolysis. The steadystate kinetic analyses implied that SpOatAC catalysis involves a rapid acetylation step followed by a slower rate-limiting deacetylation step, especially for the hydrolytic reaction (Scheme 1). This reaction order should allow for the observation of a rapid burst of 4MU release that represents a single-turnover event. Indeed, we did observe this with stopped-flow fluorometry, which showed a burst of activity in an enzyme concentration dependent manner, lasting approximately 30 s (Figure 2B). The burst amplitude was determined by obtaining the yintercept of the extrapolated steady-state slope, which was equal to 95 ± 6 % of the known amount of SpOatAC. The pre-steady state was also observed to be dependent on 4MU-Ac concentration. However, the reaction could not be saturated with the substrate concentrations tested (15–500 μM) due to the low solubility of 4MU-Ac. Despite this, we obtained the secondorder rate (k2/KM) constant of 699 ± 5 M-1·s-1 through linear-regression analysis (Figure 2C). This constant was similar to the kcat/KM value of 4MU-Ac turn-over in the absence and presence of acceptor, which confirms that k2 is independent of acceptor concentration. The k2 and KM values were estimated to be 3.16 ± 1.0 s-1 and 3.9 ± 1.4 mM, respectively. We calculated the deacetylation (k3) step from the amount of acetylated enzyme and the steady-state rate for each substrate concentration using eq 6. This provided a k3 value of 0.0093 ± 0.0002 s-1, which is a close approximation of the overall kcat for 4MU-Ac hydrolysis (Table 1). ACS Paragon Plus Environment

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Applying eq. 7, we calculated the half-life (t1/2) of the acetyl enzyme to be 77 ± 0.31 s. With a k2/k3 ratio of approximately 350, these data confirm that a k2 >> k3, and that deacetylation is the rate-limiting step. Trapping the Covalent Acetyl-SpOatAC Intermediate. We confirmed the acetyl-enzyme accumulation observed in single-turnover experiments by directly monitoring the reaction using LC-ESI-MS. Initially, we determined the molecular mass of the resting enzyme in the absence of added substrate to be m/z 21,547. We then repeated the LC-ESI-MS analysis, but with 4MU-Ac (100 µM) introduced to the sample immediately prior to LC injection. The time-lapse between the addition of substrate and separation of the protein within the column was approximately 60 s. The consequent mass spectrum revealed a single peak with m/z 21,589 (Figure 3A). This indicated that free-enzyme had been completely converted into a new species with a difference of m/z 42 relative to the resting enzyme. This mass shift is consistent with the accumulation of an acetyl-enzyme intermediate. To identify the acetylated residue, we quenched a larger scale reaction with cold acetone, digested the recovered protein with trypsin and then separated the resulting peptides by LCMS/MS. Comparison of peptides produced from trypsin digests of the quenched SpOatACcatalyzed reaction and a control digest of apo enzyme identified two ions (m/z 1431.20 [M+2H]2+ and m/z 1452.20 [M+2H]2+) with a neutral mass difference of m/z 42. These masses corresponded well with the native and acetylated forms of the peptide 414WGSELEADANSLGIADGTMLSIGDSVALR442 (Ser438 in bold). MS/MS analyses of

these parent ions verified the sequence of the peptide, and mapped the position of the acetyl group to Ser438, the putative catalytic Ser residue (Figure 3B, Table 2).


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Biochemistry

Table 2. Identification of O-acetylated peptide in SpOatC by Q-TOF LC-MS/MSa

Figure 3. Direct observation of the acetyl-enzyme intermediate by MS. A) SpOatAC was incubated in the absence and presence of 0.1 mM 4MU-Ac and immediately injected onto a C-18 reverse-phase HPLC column (total incubation time, 60 s) with MS detection. The resulting mass spectra of SpOatAC incubated in the absence (black) and presence (red) of 4MU-Ac are off-set and show a molecular mass difference of 42, consistent with the formation of a covalent acetyl-enzyme intermediate. B) LC-ESIMS/MS fragmentation of a tryptic peptide of SpOatAC from an acetone-quenched reaction. The peptide with m/z 1452.2 [M+2H]2+ corresponds to the amino acid sequence shown and the fragmentation profile is consistent with Oacetylation occurring on Ser438. See Table 2 for a list of peptide masses identified.

Ion # b1 b2 b3 b4 b5 b6 b7 b8 b9 b10 b11 b12 b13 b14 b15 b16 b17 b18 b19 b20 b21 b22 b23 b24 b25 b26 b27

[M+H] 186.08 244.11 331.14 460.18 573.27 702.31 773.35 888.37 959.41 1073.45 1160.49 1273.57 1330.59 1443.68 1514.71 1629.74 1686.76 1787.81 1918.85 2031.93 2145.02 2202.04 2317.07 2446.11 2545.18 2616.21 2729.30

Residue W G S E L E A D A N S L G I A D G T M L I G D S* V A L R

[M+H] 2717.33 2660.31 2573.28 2444.23 2331.15 2202.11 2131.07 2016.04 1945.01 1830.96 1743.93 1630.85 1573.83 1460.74 1389.70 1274.68 1217.66 1116.61 985.57 872.48 759.40 702.38 587.35 458.31 359.24 288.20 175.12

Ion # y27 y26 y25 y24 y23 y22 y21 y20 y19 y18 y17 y16 y15 y14 y13 y12 y11 y10 y9 y8 y7 y6 y5 y4 y3 y2 y1

a

b and y ions correspond to the fragments identified in Figure 3 with a tolerance of ± 0.03.

pH dependence of SpOatAC kinetic parameters. Previous structural analysis of SpOatAC indicated that Ser438 belongs to a classical catalytic triad involving His571 and Asp568. Through site-specific replacement studies, we demonstrated that these residues were necessary for catalysis and that the enzymatic of activity SpOatAC was affected by pH.10 To better ACS Paragon Plus Environment

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understand the influence of pH on SpOatAC catalysis, we determined its steady state MichaelisMenten parameters over a the range of 4.7–8.5. We were limited to these pH values because of the low fluorescence response and instability of 4MU-Ac in acidic and basic conditions, respectively. The pH dependence of kcat (enzyme-substrate complex) and kcat/KM (free-enzyme) for 4MU-Ac hydrolysis adhered to a sigmoidal curve implying the involvement of a single ionizable group (Figure 4A and 4B). The estimated pKa values for this group in the enzymesubstrate complex and free enzyme were at 6.9 ± 0.1 and 7.3 ± 0.2, respectively. Similarly, the pH dependence of the kinetic parameters for acetyltransfer to GlcNAc4 was also sigmoidal (Figure 4C and 4D). However, the estimated pKa values of 5.5 ± 0.2 and 6.0 ± 0.1 for the enzyme-substrate complex and the free enzyme were lower by 1.4 and 1.3 pH units, respectively. These data are consistent with the identification of His571 as the ionizing residue involved in catalysis, and that its protonation state affects hydrolysis and transacetylation to different extents.

Figure 4. pH dependence of SpOatAC-catalyzed esterase and O-acetytransferase activities. Steady-state kinetic parameters for were determined for 1 μM SpOatAC in 50 mM citrate-phosphate buffer at the pH values indicated. A) kcat and B) kcat/KM values for 4MU-Ac hydrolysis, and C) kcat and D) kcat/KM values for GlcNAc5 O-acetylation are plotted as a function of pH. Values of pKa were determined by fitting the data to eq. 3. Error bars denoted s.d. (n=3). ACS Paragon Plus Environment

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Biochemistry

Contribution of Asn491 to catalysis. Former crystallographic analysis of SpOatAC showed that a covalent complex involving Ser438 with methane sulfonylfluoride mimics the tetrahedral transition state. Further, this adduct was stabilized by Asn491, to which it was hydrogen bonded. When this Asn residue was replaced with Ala, SpOatAC was inactive as a transferase. However, surprisingly it had 45 % residual hydrolysis activity.10 This was unexpected since Asn491 is one of only two potential residues that could stabilize the transition-states of each half of the pingpong bi-bi reaction pathway. Hence, with new knowledge of the SpOatAC-catalyzed reaction, we conducted a kinetic analysis of the esterase activity of the Asn491Ala variant using 4MU-Ac as substrate. The kcat value of 0.0037±0.0002 s-1 was 34 % of that for the wild-type, a decrease that is comparable to what was previously observed with specific activity measurements.10 However, the KM value of 230 ± 31 M for 4MU-Ac was 20-fold higher, with the consequence that the kcat/KM of 16 ± 2 M-1·s-1 was 53-fold lower.

DISCUSSION OatA-dependent O-acetylation of PG is a protective cell surface modification that occurs in most Gram-positive bacteria.4,21 As this process is typical in drug-resistant pathogens, OatA is considered to be important for virulence and it thus represents a potential target for the development of new antibacterial agents. We began to explore this potential with the biochemical and structural studies on the catalytic O-acetyltransferase domain of OatA. This led to our proposal that it functions using a ping-pong bi-bi mechanism of action.10 Direct experimental evidence for this mechanism is provided in this study by both pre-steady and steady-state kinetic analyses and with the trapping of the acetyl-enzyme intermediate. A ping-pong bi-bi mechanism is consistent with the structure of SpOatAC, which possesses an SGNH-hydrolase fold involving a catalytic triad comprised of Ser438-His571ACS Paragon Plus Environment

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Asp569. The crystal structure of the enzyme inactivated by the mechanism-based inhibitor methane sulfonylfluoride10 supported Ser438 of this triad as the catalytic Ser residue.12 The trapping and direct observation of O-acetyl-Ser438 in this study confirmed both the identification of this catalytic nucleophile and that the reaction mechanism proceeds through a covalent intermediate. The participation of a His571 residue in the mechanism is supported by the pHdependence of the reaction, which revealed an essential ionizable residue with pKa values consistent with a His residue. These data correspond well with previous kinetic and mechanistic analyses of PG O-acetyltransferase B (PatB) of Gram-negative bacteria, which was also shown to use a ping-pong bi-bi mechanism.5 Whereas the structure of PatB has not yet been determined, and despite the overall low sequence identity, homology models show that its structure could be similar to SpOatAC.5,6 OatA is predicted to be an integral membrane protein comprised of an N-terminal membrane-spanning domain (OatAN), and the C-terminal extracellular domain OatAC. OatAN has 11 hypothetical membrane helices and amino acid sequence similarity to the Acyltransferase 3 (Acyl_transf_3) superfamily of enzymes (Pfam PF01757; InterPro IPR002656). As such, it has been proposed to translocate acetyl groups from a cytoplasmic source, presumably acetyl-CoA, across the cytoplasmic membrane for presentation to OatAC for their subsequent transfer to muramoyl groups in the peptidoglycan sacculus.10 It remains to be established if OatAN indeed catalyzes this translocation, and whether or not an intermediate acetyl carrier molecule is involved. Regardless, given the expected localization of OatAC, acetylation of its catalytic Ser438 would occur at the outer leaflet of the cytoplasmic membrane. As the structure of SpOatAC resembles those of esterases from the SGNH-hydrolase family, previous work attempted to explain why this domain catalyzes acetyl transfer instead of hydrolysis. Indeed, such hydrolysis would be wasteful with the loss of the acetate product to the


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Biochemistry

external milieu, a reaction that the enzyme would need to minimize if not preclude in vivo. The methyl sulfonate adduct to the catalytic Ser of Serine esterases is generally thought to mimic the putative tetrahedral transition state for attack by water.22 The crystal structure of SpOatAC in complex methyl sulfonate showed that the adduct is stabilized by only two hydrogen bond donors (the backbone NH of Ser438 and the Nδ2 of Asn491) and its orientation appeared to be set by the carbonyl oxygen of Val460.10 This orientation of the adduct is different to those in structures of most SGNH-hydrolases in complex with similar inhibitors (phosphonylfluorides), which use three H-bond donors (Asn, Gly, and Ser) for stabilization. Based on this, the consequent orientation of the SpOatAC acetyl-enzyme was proposed to disfavor hydrolysis, thereby minimizing any wasteful esterase activity.10 This proposal is supported by our current pre-steady state kinetic analysis of the esterase activity catalyzed by SpOatAC where k3 was found to be the rate limiting step for hydrolysis. The t1/2 of 77 s for the hydrolytic deacetylation step is consistent with that observed for the esterase activity of hamster arylamine acetytransferase 2 (t1/2 = 88 s)23 while the kinetic parameters are in striking contrast to those of authentic SGNH carbohydrate esterases. For example, the k2 and k3 values for the carbohydrate esterase CtCE2 from Clostridium thermocellum24 are 7.2 × 103 and 5.25 × 105fold greater than those for SpOatAC , while the k2/k3 ratio is only ~4.6 compared to ~350 for SpOatAC. Given the very low k3 value for the hydrolytic reaction of SpOatAC in vitro, coupled with the required localization of the enzyme between the outer leaflet of the cytoplasmic membrane and the PG sacculus, it is highly unlikely that any release of free acetate would occur. Instead, the acetylcharged enzyme would appear be primed and wait for its interaction with muramoyl residues along a peptidoglycan strand for subsequent transfer of the acetyl to free C6 hydroxyl groups. In this regard, the high turgor pressure of the cytoplasm would force the cytoplasmic membrane, and hence any of its associated enzymes, up against newly incorporated PG repeat units on the ACS Paragon Plus Environment

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Page 20 of 26

PG sacculus, the substrate for SpOatAC.10 Furthermore, this juxtaposition and tight association may help to limit water availability. Asn491was predicted to comprise the oxyanion hole of SpOatAC based on sequence alignments

with

other

SGNH

hydrolases

and

preliminary

characterization

of

the

(Asn491Ala)SpOatAC variant supported this role of the residue.10 However, the current kinetic analysis suggests that its role to stabilize transition states is minimal as its replacement does not significantly affect kcat, at least of for the hydrolytic reaction when using pseudosubstrates with good leaving groups. Although the overall rate of hydrolysis is low, attack on the acetyl-enzyme by water and stabilization of the subsequent transition state appears to only require a single Hbond, which is provided by Ser438; Asn491 is the only other appropriate residue within Hbonding distance10. While contributing a little to this stabilization, Asn491 appears to have a larger role in substrate binding as reflected by the significant increase in KM value upon its replacement. Unfortunately, it is not possible to directly assess any role of Asn491 in stabilizing the transition state for the transfer of acetyl to acceptor carbohydrate because (Asn491Ala)SpOatAC is devoid of measurable transferase activity.10 It remains to be determined what role, if any, Asn469 plays in binding acceptor substrate or stabilizing the second transition state as, despite considerable effort, we have not achieved any success in obtaining a crystal structure of OatAC in complex with an appropriate ligand. In general, it has been implied that many polysaccharide O-acetyltransferases with a Ser-His-Asp triad employ a ping-pong bi-bi mechanism of action. However, in most cases, with the exception of PatB,5 the catalytic mechanism has not been thoroughly studied. This work provides the first determination of both the pre-steady and steady-state kinetic parameters of OatAC, providing a foundation for the future characterization of other members of this medically relevant enzyme class.


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Biochemistry

AUTHOR INFORMATION Corresponding Author E-mail: [email protected]. Phone: +1-519-824-4120 ORCID Anthony Clarke: 0000-0003-4076-0488 Present Address †

Michael G. DeGroote Centre For Learning - Room 2301, McMaster University, 1200 Main St.

West, Hamilton, ON, L8N 3Z5, Canada Author Contributions DS conceived, designed and performed experiments, and AJC supervised the study; DS and AJC wrote the manuscript. Funding Sources These studies were supported by operating grants to AJC from GlycoNet (a Canadian National Centre of Excellence) and the Canadian Institutes for Health Research. Notes The authors declare no competing financial interests. Acknowledgments We thank both Dr. Dyanne Brewer and Dr. Armen Charchoglyan of the Mass Spectrometry Facility (Advanced Analysis Centre, University of Guelph) for expert and technical assistance. We also thank Joseph Ciufo for his technical assistance with some of the enzyme assays.

ABBREVIATIONS PG, peptidoglycan; OatA, O-acetyltransferase A; SaOatAC and SpOatAC, C-terminal Oacetyltransferase domains of OatA from S. aureus and S. pneunmoniae, respectively; MurNAc,


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N-acetylmuramic

acid;

GlcNAc,

N-acetylglucosamine;

Page 22 of 26

Oat,

O-acetyltransferase;

Pat,

peptidoglycan O-acetlytransferase;4MU-Ac, 4-methylumbelliferone; pNP-Ac; p-nitrophenyl acetate; (GlcNAc)4 and (GlcNAc)5, chitotriose and chitopentaose, respectively; MSF, methane sulfonylfluoride;

IMAC,

immobilized

metal

affinity

chromatography;

LC,

liquid

chromatography; ESI, electrospray ionization; MS, mass spectrometry; MS/MS, tandem mass spectrometry; CE2, carbohydrate esterase family 2; SDS-PAGE, sodium dodecyl sulfatepolyacrylamide gel electrophoresis; DMSO, dimethylsulfoxide.

REFERENCES (1) Davis, K. M., and Weiser, J. N. (2011) Modifications to the Peptidoglycan Backbone Help Bacteria to Establish Infection. Infect. Immun. 79, 562–570. (2) Whitfield, G. B., Marmont, L. S., and Howell, P. L. (2015) Enzymatic Modifications of Exopolysaccharides Enhance Bacterial Persistence. Front. Microbiol. 6, 471. (3) Callewaert, L., and Michiels, C. W. Lysozymes in the Animal Kingdom.(2010) J. Biosci. 35, 127–160. (4) Moynihan, P. J., Sychantha, D., and Clarke, A. J. (2014) Chemical Biology of Peptidoglycan Acetylation and Deacetylation. Bioorg. Chem. 54, 44–50. (5) Moynihan, P. J., and Clarke, A. J. (2014) Mechanism of Action of Peptidoglycan OAcetyltransferase B Involves a Ser-His-Asp Catalytic Triad. Biochemistry 53, 6243–6251. (6) Moynihan, P. J., and Clarke, A. J. (2010) O-Acetylation of Peptidoglycan in GramNegative Bacteria: Identification and Characterization of Peptidoglycan O-Acetyltransferase in Neisseria gonorrhoeae. J. Biol. Chem. 285, 13264–13273.


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(7) Sychantha, D., Little, D. J., Chapman, R. N., Boons, G.J., Robinson, H., Howell, P. L., and Clarke, A. J. (2018) PatB1 is an O-Acetyltransferase that Decorates Secondary Cell Wall Polysaccharides. Nat. Chem. Biol. 14, 79–85. (8) Baker, P., Ricer, T., Moynihan, P. J., Kitova, E. N., Walvoort, M. T. C., Little, D. J., Whitney, J. C., Dawson, K., Weadge, J. T., Robinson, H., Ohman, D. E., Codée, J. D. C., Klassen, J. S., Clarke, A. J., and Howell, P. L. (2014) P. aeruginosa SGNH Hydrolase-Like Proteins AlgJ and AlgX Have Similar Topology but Separate and Distinct Roles in Alginate Acetylation. PLoS Pathog. 10, e1004334. (9) Riley, L. M., Weadge, J. T., Baker, P., Robinson, H., Codee, J. D. C., Tipton, P. A., Ohman, D. E., and Howell, P. L. (2013) Structural and Functional Characterization of Pseudomonas aeruginosa AlgX: Role of AlgX in Alginate Acetylation. J. Biol Chem. 288, 22299–22314. (10) Sychantha, D., Jones, C. S., Little, D. J., Moynihan, P. J., Robinson, H., Galley, N. F., Roper, D. I., Dowson, C. G., Howell, P. L., and Clarke, A. J. (2017) In Vitro Characterization of the Antivirulence Target of Gram-Positive Pathogens, Peptidoglycan O-Acetyltransferase a (OatA). PLoS Pathog. 13, e1006667. (11) Bera, A., Herbert, S., Jakob, A., Vollmer, W., and Götz, F. Why Are Pathogenic Staphylococci So Lysozyme Resistant? (2005) The Peptidoglycan O-Acetyltransferase OatA is the Major Determinant for Lysozyme Resistance of Staphylococcus aureus. Mol. Microbiol. 55, 778787. (12) Bernard, E., Rolain, T., Courtin, P., Guillot, A., Langella, P., Hols, P., and ChapotChartier, M.P. Characterization of O-Acetylation of N-Acetylglucosamine: A Novel Structural Variation of Bacterial Peptidoglycan. (2011) J. Biol. Chem 286, 2395023958.


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(13) Bera, A., Biswas, R., Herbert, S., Gotz, F. The Presence of Peptidoglycan OAcetyltransferase in Various Staphylococcal Species Correlates with Lysozyme Resistance and Pathogenicity. (2006) Infect. Immun. 74, 45984604. (14) Herbert, S., Bera, A., Nerz, C., Kraus, D., Peschel, A., Goerke, C., Meehl, M., Cheung, A., and Götz, F. Molecular Basis of Resistance to Muramidase and Cationic Antimicrobial Peptide Activity of Lysozyme in Staphylococci. (2007) PLoS Pathog. 3, e102. (15) Hébert, L., Courtin, P., Torelli, R., Sanguinetti, M., Chapot-Chartier, M.P., Auffray, Y., and Benachour, A. (2007) Enterococcus faecalis Constitutes an Unusual Bacterial Model in Lysozyme Resistance. Infect. Immun. 75, 5390-5398. (16) Le Jeune, A., Torelli, R., Sanguinetti, M., Giard, J.C., Hartke, A., Auffray, Y., and Benachour, A. (2010) The Extracytoplasmic Function Sigma Factor SigV Plays a Key Role in the Original Model of Lysozyme Resistance and Virulence of Enterococcus faecalis. PLoS One 5, e9658. (17) Crisóstomo, M. L., Vollmer, W., Kharat, A. S., Inhülsen, S., Gehre, F., Buckenmaier, S., and Tomasz, A. (2006) Attenuation of Penicillin Resistance in a Peptidoglycan O-Acetyl Transferase Mutant of Streptococcus pneumoniae. Mol. Microbiol. 61, 1497–1509. (18) Moynihan, P. J., and Clarke, A. J. (2013) Assay for Peptidoglycan O-Acetyltransferase: A Potential New Antibacterial Target. Anal. Biochem. 439, 73–79. (19) Moynihan, P. J, Clarke, A. J. (2014) Mechanism of Action of Peptidoglycan OAcetyltransferase B Involves a Ser-His-Asp Catalytic Triad. Biochemistry 53, 6243–6251. (20) Dixon, M., and Webb, E. C. The Enzymes, 2nd ed.; Longmans Green: London, 1964; p 108.


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(21) Moynihan, P. J., and Clarke, A. J. (2011) O-Acetylated Peptidoglycan: Controlling the Activity of Bacterial Autolysins and Lytic Enzymes of Innate Immune Systems. Int. J. Biochem. Cell Biol. 43, 1655–1659. (22) Myers, D. K., and Kemp, A. (1954) Inhibition of Esterases by the Fluorides of Organic Acids. Nature 173, 33–34. (23) Wang, H., Vath, G. M., Gleason, K. J., Hanna, P. E., and Wagner, C. R. (2004) Probing the Mechanism of Hamster Arylamine N-Acetyltransferase 2 Acetylation by Active Site Modification, Site-Directed Mutagenesis, and Pre-Steady State and Steady State Kinetic Studies. Biochemistry 43, 8234–8246. (24) Montanier, C., Money, V. A., Pires, V. M. R., Flint, J. E., Pinheiro, B. A., Goyal, A., Prates, J. A. M., Izumi, A., Stålbrand, H., Morland, C., Cartmell, A., Kolenova, K., Topakas, E., Dodson, E. J., Bolam, D. N., Davies, G. J., Fontes, C. M. G. A., and Gilbert, H. J. (2009) The Active Site of a Carbohydrate Esterase Displays Divergent Catalytic and Noncatalytic Binding Functions. PLoS Biol. 7, e71.


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For Table of Contents Use Only Peptidoglycan modification by the catalytic domain of Streptococcus pneumoniae OatA follows a ping-pong bi-bi mechanism of action David Sychantha and Anthony J. Clarke


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Peptidoglycan Modification by the Catalytic Domain of Streptococcus

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