A Random-Sequential Kinetic Mechanism for Polysaccharide

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A random-sequential kinetic mechanism for polysaccharide monooxygenases John Hangasky, and Michael A Marletta Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00129 • Publication Date (Web): 23 Apr 2018 Downloaded from http://pubs.acs.org on April 23, 2018

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Biochemistry

A random-sequential kinetic mechanism for polysaccharide monooxygenases

John A. Hangaskya and Michael A. Marlettaa,b,c,*

aCalifornia

Institute for Quantitative Biosciences (QB3), University of California,

Berkeley, California, USA 94720 bDepartment cDepartment

of Chemistry, University of California, Berkeley, California, USA 94720 of Molecular and Cell Biology, University of California, Berkeley,

Berkeley, California, USA 94720 *Corresponding author: Michael A. Marletta ([email protected])

Keywords: monooxygenase, oxygen activation, enzyme mechanism

ACS Paragon Plus Environment

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Abbreviations AA, auxiliary activity CDH, cellobiose dehydrogenase CO, carbon monoxide Glc6, cellohexaose HAA, hydrogen atom abstraction HPAEC, high performance anion exchange chromatography HRP, horseradish peroxidase LPMO, lytic polysaccharide monooxygenase MES, 2-(N-morpholino)ethanesulfonic acid MOPS, 3-morpholinopropane-1-sulfonic acid N. crassa, Neurospora crassa PASC, phosphoric acid swollen cellulose PMO, polysaccharide monooxygenase


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Biochemistry

Abstract Polysaccharide monooxygenases are mononuclear copper enzymes that catalyze the hydroxylation of polysaccharides leading to the scission of the glycosidic bond. The mechanism in which PMOs utilize molecular oxygen to oxidize the polysaccharide substrate still remains largely unknown. Here, steady-state kinetics assays were used to probe the mechanism of oxygen-dependent cellohexaose oxidation catalyzed by MtPMO9E. Kinetic analysis indicated that both kcat/KM(O2) and kcat/KM(Glc6) were dependent on the concentration of the second substrate. Inhibition studies using carbon monoxide were also carried out. In addition, KD values for Glc6 were determined for the Cu(I) and Cu(II) forms of the enzyme. Taken together, PMOs follow a random-sequential kinetic mechanism to form a ternary ES-O2 complex. The optimal pH for MtPMO9E turnover was determined to be between pH 6.00 and pH 7.00. Furthermore, the kinetic parameters kcat, kcat/KM(O2) and kcat/KM(Glc6) demonstrate a decrease in PMO activity at low pH, and provide equivalent kinetic pKas of 5.10. This points to the protonation of a general base required for turnover. These results provide a basis for the initial chemical steps in the mechanism of PMOs.


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Introduction Polysaccharide monooxygenases (PMOs), also known to as lytic PMOs (LPMOs), are a large family of extracellular, copper-dependent enzymes that degrade a wide range of polysaccharides via hydroxylation of the carbohydrate backbone.1–4 These enzymes serve vital physiological roles for their host organisms5,6 and also hold promise in biomass conversion technologies.7,8 The Carbohydrate Active Enzyme database (CAZy; www.cazy.org) classifies PMOs into six auxiliary activity (AA) families based on primary sequence similarity: AA9, AA10, AA11, AA13, AA14 and AA15.9 The predominate nomenclature used for naming PMOs begins with an abbreviation of the organism of origin [e.g. Mt (Myceliophthora thermopilia)], followed by (L)PMO with the CaZy family (e.g. (L)PMO9) and a letter to identify the specific PMO (e.g A, B, C, etc). Although the primary sequences may vary, PMOs share structural similarities, including a surface exposed Cu active site with a flat substrate-binding surface and a rigid β-sheet core connected by flexible loops.10 The Cu(II) center is coordinated by the N-terminal histidine, the N-terminal amine and a second histidine ligand in a histidine brace motif. The active site is positioned on the flat substrate-binding surface of the protein exposing the oxidative center of the PMO to surface of the polysaccharide substrate.11

While general kinetic mechanisms for copper monooxygenases have been extensively studied,12 there have been limited mechanistic studies with PMOs. Postulated mechanisms for PMOs have relied on crystallographic and computational data, as well as limited in vitro kinetic studies.13–17 Kinetic studies have been complicated by the fact that initially all known PMOs used insoluble polymeric polysaccharide substrates. The reduced Cu(I) active site, generated from electron transfer from cellobiose dehydrogenase (CDH)18,19 or small molecule reductants such as ascorbate,20,21 is then followed by reactivity with O2 to form a Cu-superoxo intermediate.16 The timing and delivery of a second electron and two protons dictate the Cu/O2 intermediate responsible for hydrogen atom abstraction (HAA) and regioselective oxidation of the polysaccharide (Scheme 1).22 Although the


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Biochemistry

identity of this intermediate is unknown, it is generally proposed to be a Cusuperoxo or Cu-oxyl species.23,24

Scheme 1 Minimal scheme for polysaccharide oxidation. R = polysaccharide substrate, E = PMO

One of the challenges impeding mechanistic studies of PMOs is the uncoupling of oxygen reduction and substrate oxidation. PMOs can reduce O2 to generate H2O2, with or without a polysaccharide substrate present. H2O2 can serve as a co-substrate for PMOs leading to a radical based mechanism,25,26 therefore, it can be difficult to distinguish oxygenase activity from peroxygenase activity (Scheme 1). However, high cellohexaose (Glc6) concentrations suppress H2O2 formation27,28 and lead to a fully coupled reduction of O2 and Glc6 oxidation.29

MtPMO9E is an ideal enzyme for kinetic studies because it is active on soluble substrates (e.g. Glc6), which removes the problems associated with insoluble substrates. Moreover, the suppression of uncoupled turnover by Glc6 and the addition of H2O2 scavengers, such as horseradish peroxidase (HRP), to reaction mixtures, enables the direct study of O2-dependent oxidations.29 Reported here are a comprehensive series of steady-state assays with MtPMO9E with the overall conclusion that PMOs operate by a random-sequential kinetic mechanism. Coupling ratios confirmed that saturating concentrations of Glc6 lead to stoichiometric conversion of O2 and oxidized Glc6. Additionally, the pH dependence of MtPMO9E reactivity showed pH independent and optimal turnover between pH 6.00 and pH 7.00. Reversible, pH dependent inactivation (pKa ~5.10) of an active site base was also observed, but the origin of this phenomenon could not be definitively assigned


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to a specific residue. These results provide insight into the initial steps of the chemical mechanism PMOs use to oxidize polysaccharides.

Materials and Methods

Materials All materials were purchased from commercial vendors and used without further purification. Glc6 was purchased as a lyophilized powder with >95% purity (Megazyme, Ireland) and resuspended in 50 mM MES, 50 mM MOPS, pH 5.20–7.00 with the ionic strength (I = 100 mM) adjusted using NaCl. A 96-well plate formatted BCA assay was used to determine the reducing-end concentration of the Glc6 stock as previously described.30 MtPMO9E was expressed in N. crassa and purified as previously described.31

Steady-state kinetic assays varying Glc6. Steady-state kinetic measurements varying Glc6 were obtained at a fixed O2 concentration (50, 125 206, 300, 500 or 800 µM). Reaction solutions contained MtPMO9E (0.5–1.0 µM) with Glc6 (0–500 µM), HRP (0 or 1.3 µM), Amplex Red (0 or 200 µM), and a saturating concentration of ascorbic acid (2 mM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 5.20–7.00 at 40 °C. Buffer containing Glc6 was equilibrated at 40 °C for two minutes prior to adding ascorbic acid (2 mM). The reaction was then initiated by the addition of MtPMO9E. Aliquots (20 µL) of the reaction mixture were taken over a specified time course and quenched with 0.2 M NaOH (20 µL).

Initial rates were determined by quantifying the consumption of Glc6 over time using high performance anion exchange chromatography (HPAEC). Standard curves for Glc6 were generated from peak integration from Glc6 standards of known concentration. Initial rates were determined from the consumption of no more than 15% of the limiting reactant in the assay; this was imperative as MtPMO9E could


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Biochemistry

potentially oxidize the accumulating product cello-oligosaccharides. No lag or burst phases were observed and initial rates plotted as a function Glc6 concentration were fit to the Michaelis Menten equation yielding the apparent kinetic parameters kcat and kcat/KM(Glc6) as well as the Michaelis constant KM(Glc6).

The initial rates determined at a given fixed Glc6 concentration were also re-plotted as a function of O2 concentration and then fit to the Michaelis Menten equation. The resulting kinetic parameters, kcat and kcat/KM(O2) and the Michaelis constant KM(O2), were used to provide insight to the O2 activation kinetics involved in the coupled reaction. At Glc6 concentrations below 25 µM, O2 saturation could not be achieved and the data was fit to Equation 1, where kcat/KM is the catalytic efficiency and KM is the Michaelis constant.

[]

=

∗[ ] [ ]

Equation 1

Steady-state kinetic assays varying O2 Oxygen consumption assays using a Clark-type oxygen electrode (Oxytherm, Hansatech) were utilized to monitor assays with varying O2 concentrations. Buffer (50 mM MES, 50 mM MOPS, I = 100 mM, pH 5.20–7.00) containing a fixed concentration of Glc6 (0, 10, 25, 50, 100, 200, 375 or 500 µM) was stirred in a thermostatted chamber at 40 °C until the temperature and O2 concentration were equilibrated. A saturating concentration of ascorbic acid (2 mM) was added to determine the background rate of O2 consumption prior to the addition of MtPMO9E (1–2 µM).

Although the consumption of O2 could be continuously monitored for several minutes with no observed lag or burst phases, initial rates were determined from the linear phase of time courses accounting for 5–10% turnover of the limiting reactant. The rate of O2 consumption from ascorbic acid was subtracted to yield the


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final initial rate. Initial rates were plotted as a function of the O2 concentration and fit to the Michaelis-Menten equation to obtain the apparent kinetic parameters kcat and kcat/KM(O2).

CO inhibition Assays varying either O2 or Glc6 in the presence of CO were performed in sealed reaction vials in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.50. A saturated stock solution of CO (950 µM) was prepared by continuous sparging with CO gas for 30 minutes. Mixtures of the reaction components were combined to give the desired O2/CO concentration, using N2 to maintain a partial pressure of one atmosphere in the closed reaction vial. Assays varying O2 (50–615 µM) contained MtPMO9E (1 µM), ascorbic acid (2 mM) and Glc6 (500 µM). Assays with varying Glc6 (0–500 µM) used the same assay conditions, with the exception of a fixed [O2] (216 µM) and the addition of HRP (1.3 µM) and Amplex Red (200 µM) to the reaction mixture. Assays were initiated by the addition of ascorbic acid. To determine initial rates, reaction aliquots were quenched in NaOH and analyzed by HPAEC as described above.

Using GraphPad Prism software, global analysis of the initial rate data was used to determine the inhibition constant, Ki. The initial rates collected as a function of [O2] were fit to equations 2 and 3, where kcat is the maximum velocity, is the observed KM(O2), and Ki is the inhibition constant. , () , and ! were shared among all data points.

= []

" ∗[ ] $% # () [ ]

() = () ∗ &1 +

Equation 2 [)] #*

+

Equation 3

Initial rate data collected as a function of [Glc6] were fit to equation 4 and 5, where !-. is the observed turnover number at each given [CO], is the maximum


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Biochemistry

velocity without CO, (/012) is the Michaelis constant and ! is the inhibition constant. , (/012) , and ! were shared among all data points. []

=

*34 "

∗[562]

# (789) [562]

!-. =

"

[:] *

Equation 4 Equation 5

pH Dependence Steady-state assays were performed as described above between pH 5.20 and 7.00. The log of the kinetic parameters kcat, kcat/KM(O2), and kcat/KM(Glc6) were plotted as a function pH to determine the pH dependence of each parameter. The pH profiles were fit to Equation 6, where EH is the maximal kinetic parameter of the protonated enzyme form, E is the kinetic parameter of the conjugate base, x is the pH at which the kinetic parameter was determined and pKa is the kinetic pKa.

log( >?@A>B CDEDFA@E) =

G∗HIJ ∗HIK HIJ HIK

Equation 6

H2O2 Suppression Endpoint measurements were used to determine the extent of H2O2 formation in the presence of Glc6. Reactions containing MtPMO9E (1 µM) and Glc6 (0–1 mM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.50 were initiated by the addition of ascorbic acid (50 µM) and incubated for 10 minutes at 40 °C before recording the absorbance of resorufin (λmax = 570) using a SpectraMax340 (Molecular Devices). To correct for the background photo-oxidation of Amplex Red, the absorbance reading from corresponding control reactions without added PMO were subtracted from the reported values (n = 3).


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Substrate KD Determination Quenching of the intrinsic tryptophan fluorescence of MtPMO9E resulting from Glc6 binding was used to determine Cu(II) MtPMO9E–Glc6 and Cu(I) MtPMO9E–Glc6 binding constants. Samples contained MtPMO9E (1 µM) and Glc6 (0–2.5 mM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.50 prepared in a fluorescence cuvette. The Glc6 binding constant to the Cu(II) enzyme form was performed aerobically, and that for the Cu(I) enzyme form, anaerobically. Anaerobic samples were prepared in an glovebag following thorough degassing via argon sparging. The Cu(I) enzyme form was generated by a 20-minute incubation with ascorbic acid (2 mM), which was subsequently removed using a Micro Bio-Spin 6 column. Fluorescence measurements

were

recorded

using

a

Horiba

Scientific

FluoroMax-4

spectrofluorometer. Samples were excited at 290 nm and emission spectra were recorded from 300–600 nm. The emission intensity at 354 nm was plotted as a function of Glc6 concentration and fit to equation 7, where I is the measured fluorescence intensity, Io is the initial fluorescence intensity measurement, If is the final fluorescence intensity measurement, E is the enzyme concentration, n is the number of binding sites, S is the Glc6 concentration and KD is the binding affinity. LMLN

LO ML

=

[][P]#Q MR([][P]#Q ) MS[][P][]

Equation 7

RESULTS Kinetics of O2 Reduction Since PMOs are known to uncouple O2 reduction from polysaccharide oxidation,27 two assays formats were employed to determine the kinetic parameters of O2 reduction. First, O2 consumption assays were used to account for all PMO turnovers, regardless if O2 reduction was coupled to Glc6 oxidation. Second, Glc6 consumption assays were used to study only the coupled reaction (i.e. concomitant O2 reduction and Glc6 oxidation). Using both of these assay formats, steady-state kinetic assays with varying O2 were performed at several fixed concentrations of Glc6.


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Biochemistry

The initial rates determined using the O2 consumption assay were fit to the Michaelis-Menten equation (Figure S1) and kinetic parameters are summarized in Table 1. The results show that kcat decreased with increasing Glc6 concentrations, plateauing at saturating (>200 µM) Glc6 concentrations (Table 1). In the absence of Glc6, kcat (26 ± 2 min–1) is the maximal rate of the uncoupled reaction. At saturating Glc6 concentrations kcat (17 ± 1 min–1) is the catalytic rate of the coupled reaction resulting in Glc6 hydroxylation. As kcat reports on all kinetic steps after substrate binding, the difference in kcat reflects a change in an irreversible chemical step indicating the reaction mechanism of the PMO has changed. Based on coupling ratios (see below), this is a result of coupling O2 reduction to substrate hydroxylation. The kcat/KM(O2) (0.074 ± 0.016 µM–1 min–1) was insensitive to the Glc6 concentration (Figure 1A), due to the saturating ascorbate concentration maintaining the reduced Cu(I) form of the enzyme. The Michaelis constant for O2 revealed an inverse correlation between the apparent KM(O2) and the Glc6 concentration, suggesting synergistic binding of O2 and Glc6.

Steady-state kinetic assays with varied O2 using the HPAEC-based Glc6 consumption assay were carried out next to probe the O2 reaction kinetics of the coupled reaction (Table 1 and Figure S2). kcat increases with increasing Glc6 concentrations, which is consistent with a larger fraction of the total enzyme forming an ES complex (S = Glc6) that leads to product formation. At saturating concentrations of O2 and Glc6, kcat (17 ± 1 min–1) is agreement with values determined from the O2 consumption assay (see above). O2 saturation of the enzyme could not be achieved at low Glc6 concentrations, preventing the accurate determination of kcat. However, the kcat/KM(O2) and KM(O2) were determined using Equation 1 (Table 1). Unlike the O2 consumption assays, the Glc6 consumption assays indicated the second order rate constant for the coupled reaction, kcat/KM(O2), was dependent on Glc6 concentration. The regression analysis of kcat/KM(O2) as a function of Glc6 concentration resulted in a secondary plot, which intersects the origin and plateaus at high (>200 µM) Glc6 concentrations (Figure 1A).


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Figure 1. Secondary Plots of kcat/KM. (A) kcat/KM(O2) as a function of Glc6. The kcat/KM(O2) was determined using O2 consumption assays () and Glc6 consumption assays (). Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), a fixed [Glc6] (0–385 µM) and a varied [O2] (0–800 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.5 at 40 °C (n = 3). (B) kcat/KM(Glc6) as a function of [O2]. The kcat/KM(Glc6) was determined in the presence () and absence () of HRP (1.3 µM) and Amplex Red (200 µM). Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (5–500 µM) and a fixed [O2] (50–800 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM pH 6.5 at 40 °C (n = 3).

Table 1. Apparent kinetic parameters measured for MtPMO9E at fixed [Glc6]. O2 Consumption Assaysa [Glc6]

kcat

kcat/KM(O2)

KM(O2)

Glc6 Consumption Assaysb kcat

kcat/KM(O2)


KM(O2)

Reaction

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Biochemistry

(µM)

(min–1)

(μM–1 min–1)

(μM)

(min–1)

(μM–1 min–1)

(μM)

Couplingc

0

26 ± 2

0.071 ± 0.012

365 ± 45

ND

ND

ND

ND

10

22 ± 2

0.073 ± 0.012

300 ± 40

ND

0.001 ± 0.001

710 ± 185

>70

25

20 ± 2

0.078 ± 0.011

258 ± 44

ND

0.004 ± 0.002

630 ± 100

20 ± 10

50

18 ± 1

0.077 ± 0.018

235 ± 36

6±1

0.012 ± 0.007

524 ± 81

6.4 ± 4.0

100

17 ± 1

0.074 ± 0.013

230 ± 35

10 ± 1

0.033 ± 0.011

300 ± 83

2.2 ± 0.9

200

17 ± 2

0.074 ± 0.016

235 ± 33

17 ± 2

0.065 ± 0.017

257 ± 60

1.2 ± 0.3

375

17 ± 1

0.074 ± 0.011

233 ± 37

17 ± 2

0.070 ± 0.017

240 ± 50

1.1 ± 0.3

500

17 ± 1

0.074 ± 0.012

235 ± 41

17 ± 2

0.076 ± 0.019

228 ± 46

1.0 ± 0.3

a.

Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), a fixed [Glc6] (0–500 µM) and a varied [O2] (0–800 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.5 at 40 °C (n = 3).

b.

Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), a fixed [Glc6] (0–500 µM), HRP (1.3 µM), Amplex Red (200 µM) and a varied [O2] (0–800 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.5 at 40 °C (n = 3). ND = Not determined.

c.

This ratio reflects the number of O2 equivalents consumed to that incorporated into Glc6. Value calculated from kcat/KM(O2) (O2 consumption) / kcat/KM(O2) (Glc6 consumption).

Reaction Coupling Comparison of the kcat/KM(O2) values determined using the two different assay methods showed that O2 reduction can uncouple from polysaccharide oxidation (Figure 1A). The ratio of the kcat/KM(O2) values determined by the O2 consumption and Glc6 consumption assays, provide a quantitative measure of the reaction coupling. The coupling ratio (C) was calculated from comparing the kcat/KM(O2) (O2 consumption)

to that of the kcat/KM(O2) (Glc6 consumption ) (Table 1). The extent of coupling

was strongly dependent on the concentration of Glc6. Low Glc6 concentrations led to the highest ratios (C >70) indicating the reduction of O2 did not often lead to incorporation an O-atom into the polysaccharide substrate. However, at saturating Glc6 concentrations, every reduction of O2 led to oxidation of the cellooligosaccharide indicated by a coupling ratio of unity (C = 1.0 ± 0.3). Similarly, spectrophotometric assays using HRP and Amplex Red indicated that H2O2 was the product formed in the uncoupled reaction, as reported previously.27 MtPMO9E readily produced H2O2 in the absence of Glc6, but H2O2 formation was suppressed


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by increasing the Glc6 concentration (Figure S3). The concentration of Glc6 reduces the extent of the uncoupled reaction yielding similar results to those reported in Table 1.

Kinetics with Glc6 Steady-state kinetic assays with Glc6 as the varied substrate were performed with several fixed concentrations of O2 to characterize the steps involved in Glc6 binding and oxidation (Figure S4). Assays were carried out in the presence of HRP (1.3 µM) and Amplex Red (200 µM) to scavenge H2O2 generated from uncoupling ensuring that only O2-dependent turnover was measured. The initial rates were fit to the Michaelis-Menten equation resulting in the apparent kinetic parameters summarized in Table 2. Under ambient O2 concentration, kcat (10 ± 1 min–1) and kcat/KM(Glc6) (0.11 ± 0.02 µM–1 min–1), as well as the KM(Glc6) (98 ± 14 µM) for MtPMO9E are similar to values previously reported for PMOs.13,32–34 Both kcat and kcat/KM(Glc6) increase with O2 concentrations such that at high O2 concentrations (800 µM) both parameters approach values that likely reflect the intrinsic kinetic parameters. It was also observed that increased O2 concentrations led to decreased KM(Glc6) values, again suggesting synergistic binding of the two substrates.

The analysis of kcat/KM(Glc6) as a function of O2 concentration resulted in a secondary plot that passed through the origin and plateaued at O2 concentrations greater than 300 µM (Figure 1B, Table 2), indicating that kcat/KM(Glc6) reflects kinetic processes involved in O2 binding. When HRP was excluded from the reaction conditions, the plot of kcat/KM(Glc6) as a function of O2 concentration does not exhibit an obvious plateau in kcat/KM(Glc6) at high O2 concentrations (Figure 1B and Figure S5). Under these conditions, this parameter reflects activity originating from H2O2 generated in situ from the free PMO in solution.

Table 2. Apparent kinetic parameters measured for MtPMO9E at fixed [O2].


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Biochemistry

– H2O2 Scavengera

+ H2O2 Scavenger Presentb

[O2]

kcat

kcat/KM(Glc6)

KM(Glc6)

kcat

kcat/KM(Glc6)

KM(Glc6)

(µM)

(min–1)

(µM–1 min–1)

(µM)

(min–1)

(µM –1 min–1)

(µM)

50

2.0 ± 0.2

0.038 ± 0.008

52 ± 7

1.8 ± 0.1

0.017 ± 0.005

104 ± 24

125

4.6 ± 0.4

0.102 ± 0.020

45 ± 6

5.2 ± 0.6

0.054 ± 0.012

96 ± 15

206

8.9 ± 0.5

0.300 ± 0.043

30 ± 4

10 ± 1

0.105 ± 0.018

98 ± 14

300

11 ± 1

0.440 ± 0.081

25 ± 4

15 ± 1

0.165 ± 0.024

91 ± 12

500

14 ± 1

0.636 ± 0.152

22 ± 5

18 ± 1

0.214 ± 0.050

84 ± 19

800

17 ± 2

0.739 ± 0.155

23 ± 4

17 ± 1

0.223 ± 0.040

76 ± 13

a.

Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (5–385 µM) and a fixed [O2] (50– 800 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.5 at 40 °C (n = 3).

b.

Assays were performed using MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (5–500 µM), a fixed [O2] (50–800 µM), HRP (1.3 µM) and Amplex Red (200 µM) in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.5 at 40 °C (n = 3).

CO inhibition CO was utilized as an inhibitor to help resolve the steady-state mechanism for PMOs. Analysis of the resulting kinetic parameters determined the mechanism of inhibition with respect to each substrate. Steady-state kinetic assays with varying O2 were performed using three fixed concentrations of CO (Figure 2A). The kinetic data showed the KM(O2) increases with increasing CO concentration and that kcat was unaffected, consistent with competitive inhibition of CO with O2 (Table 3). The global fit of these data resulted in an inhibition constant for CO (Ki = 155 ± 25 µM). Next, steady-state kinetic assays with varying Glc6 were performed using the same three concentrations of CO (Figure 2B). The kinetic data showed kcat decreased with increasing CO concentrations and the KM(Glc6) remained constant, consistent with noncompetitive inhibition of CO with Glc6 (Table 3). Determination of the Ki (256 ± 21 µM) by global analysis showed an increase in the Ki when Glc6 was the varied substrate.


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 2. CO inhibition of MtPMO9E. (A) Steady-state kinetic assays with varying O2. Assays contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (500 µM), O2 (50–800 µM) and CO (0, 100, 200 or 400 µM). (B) Steady-state kinetic assays with varying Glc6. Assays contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (0–385 µM), O2 (206 µM), CO (0, 100, 200 or 400 µM), HRP (1.3 µM) and Amplex Red (200 µM). All assays were performed in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.50 at 40 °C (n = 2). See Table 3 for the fits of the data and the corresponding kinetic parameters.

Table 3. Kinetic Parameters in the presence of CO.a


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Biochemistry

[CO]

kcatb

KM(O2)b

K id

kcatc

KM(Glc6)c

K ie

(µM)

(min–1)

(µM)

(µM)

(min–1)

(min–1)

(µM)

0

18 ± 2

230 ± 47

12 ± 1

104 ± 18

100

18 ± 2

300 ± 56

9±1

108 ± 32

155 ± 25

256 ± 21

200

18 ± 4

518 ± 150

7±1

109 ± 27

400

19 ± 9

842 ± 311

5±1

107 ± 33

a.

All assays were performed in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 6.50 at 40 °C (n = 2).

b.

Assays contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (500 µM), O2 (50–800 µM)and CO (0, 100, 200 or 400 µM).

c.

Assays contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (0–385 µM), O2 (206 µM), CO (0, 100, 200 or 400 µM), HRP (1.3 µM) and Amplex Red (200 µM).

d.

Determined from the global fit of initial rate data using a competitive inhibition model. See equations 2 and 3. See Table S1 for kinetics parameters of global fit.

e.

Determined from the global fit of initial rate data using a noncompetitive inhibition model. See equations 4 and 5. See Table S2 for kinetics parameters of global fit.

Binding affinity for Glc6 The quenching of the intrinsic tryptophan fluorescence of MtPMO9E upon Glc6 binding was used to determine the Glc6 binding affinity for MtPMO9E. MtPMO9E– Glc6 binding constants were determined for the Cu(II) and Cu(I) enzyme forms (Supplemental Figures S6 and S7). Glc6 binding to the oxidized enzyme form was very weak (KD = 994 ± 194 µM), similar to a previously reported binding constant determined using ITC.14 A significantly tighter binding affinity (KD = 175 ± 24 µM) was determined for the reduced enzyme in the absence of O2. Combined, these data indicate MtPMO9E is capable of binding Glc6 without O2 coordinated to the Cu active site and that the reduced enzyme has a significantly tighter affinity for Glc6.

pH dependent kinetics Steady-state assays were performed between pH 5.20 and pH 7.00 to probe for mechanistically-relevant ionizable groups on MtPMO9E (kcat), the MtPMO9E/O2 complex (kcat/KM(O2)) and the MtPMO9E/Glc6 complex (kcat/KM(Glc6)).35 Low activity


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

of MtPMO9E below pH 5.20 prevented analysis at lower pH values. MtPMO9E exhibited pH-dependent activity under conditions of low substrate (kcat/KM(O2), kcat/KM(Glc6)), as well as at saturating concentrations of Glc6 and O2 (kcat) (Table 4). The pH profiles for kcat, kcat/KM(O2) and kcat/KM(Glc6) appeared hyperbolic in shape with maximum activity between pH 6.00 and pH 7.00 (Figure 3); this data was fit to a model for a single essential base (Equation 6).

The pH dependence of kcat/KM(O2) (pKa = 5.08 ± 0.11) and kcat/KM(Glc6) (pKa = 5.11 ± 0.09) suggests there is an general base involved in forming the ES-O2 complex (S=Glc6). As this could be a result of diminished ability for the PMO to bind substrate, assays were repeated at pH 5.20 with increased substrate concentrations. There was no significant change in the resulting kinetic parameters (Table 4, second pH 5.20 entry), indicating this base was not involved in the binding of O2 or Glc6. In agreement with this finding, the pH dependence on kcat (pKa = 5.11 ± 0.08 min–1) indicated that this base is essential for chemistry. Additional assays showed that the protonation was reversible. The observed rate of O2 consumption by an aliquot of MtPMO9E incubated overnight in pH 5.20 buffer and then assayed at pH 6.50 with a saturating concentration of Glc6 (500 µM) (kobs = 5.1 ± 0.3 µM min–1) was within error of the control (kobs = 5.5 ± 0.2 µM min–1).


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Biochemistry

Figure 3. pH Dependent Turnover of MtPMO9E. (A) Regression plot of log(kcat) as a function of pH. The O2 consumption assays contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (500 µM), and O2 (50–800 µM). (B) Regression plot of log(kcat/KM(O2)) () and log(kcat/KM(Glc6)) () as a function of pH. O2 consumption assays were used to determine kcat/KM(O2) under the reaction conditions described for panel (A). Glc6 consumption assays were used to determine the kcat/KM(Glc6) and contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (0–385 µM), ambient O2 (206 µM), HRP (1.3 µM) and Amplex Red (200 µM). All assays were performed in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 5.20 – 7.00 and were performed at 40 °C (n = 3).


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Table 4. Kinetic parameters measured for the coupled turnover of MtPMO9E.a kcatb

kcat/KM(O2)b

kcat/KM(Glc6)c

(min–1)

(µM–1 min–1)

(µM–1 min–1)

5.20

4.4 ± 0.2

0.014 ± 0.003

0.037 ± 0.009

5.20

4.1 ±

0.3d

5.50

8.5 ± 0.5

0.034 ± 0.008

0.059 ± 0.017

6.00

16 ± 1

0.070 ± 0.012

0.095 ± 0.018

6.50

17 ± 2

0.074 ± 0.011

0.110 ± 0.018

7.00

17 ± 1

0.072 ± 0.012

0.110 ± 0.013

pH

0.015 ±

0.003d

0.043 ± 0.009e

a.

Assays were performed in 50 mM MES, 50 mM MOPS, I = 100 mM, pH 5.20–7.00 at 40 °C

b.

Initial rate data was collected monitoring O2 consumption. Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (500 µM), and O2 (50–800 µM) (n = 3).

c.

Initial rate data was collected, monitoring Glc6 consumption, in the presence of HRP. Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (0–385), ambient O2 (206 µM), Amplex Red (200 µM) and HRP (1.3 µM) (n = 3).

d.

Initial rate data was collected monitoring O2 consumption. Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (1 mM) and O2 (50–800 µM) (n = 2).

e.

Initial rate data was collected, monitoring Glc6 consumption, in the presence of HRP. Reactions contained MtPMO9E (1 µM), ascorbate (2 mM), Glc6 (0–385), O2 (800 µM), Amplex Red (200 µM) and HRP (1.3 µM) (n = 2).

Discussion

Multi-substrate enzymatic reactions can be mechanistically complicated and multiple probes are often needed to determine the steady-state kinetic mechanism. The elucidation of the O2-dependent mechanism for PMO reactivity has been hindered by the insoluble nature of polysaccharide substrates. The ability of MtPMO9E to oxidize Glc6 overcomes many of the issues associated with the use of insoluble substrates and has afforded insight into the mechanism of PMO reactivity with O2. Even so, careful experimental design of the reaction conditions is necessary to observe O2-dependent activity and distinguish it from H2O2-dependent activity resulting from uncoupled turnover.29


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Biochemistry

The steady-state binding order of substrates can be determined from assays in which one substrate is varied while the other is held at a fixed concentration in combination with inhibition studies.36 For MtPMO9E, both second order rate constants, kcat/KM(Glc6) and kcat/KM(O2) were dependent on the second substrate (Figure 1), indicating that each kinetic parameter encompassed the binding of both substrates. The inhibition studies show that CO is competitive with O2 and noncompetitive with Glc6 (Figure 2). These results are consistent with a randomsequential mechanism, as well as a steady-state ordered mechanism in which O2 binds prior to Glc6. Additional kinetic studies were not possible since product inhibition was not observed, there are currently no known competitive inhibitors of Glc6 and the irreversible O–O bond cleavage step prevented isotopic exchange experiments. Although not a kinetic probe, titrations with Glc6 indicated that reduced MtPMO9E was capable of binding Glc6 in absence of O2. The overall interpretation of the data presented herein is that PMOs follow a steady-state random-sequential mechanism to form a ternary ES-O2 complex (S = Glc6) (Scheme 2). The binding of O2 and Glc6 generates a Cu-superoxo complex which can be stabilized by second coordination sphere residues,37–39 but the subsequent chemical steps of O-atom incorporation into the polysaccharide substrate remain unresolved.

Scheme 2. Proposed mechanism for PMO activity.


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

One of the advantages PMOs hold in catalyzing polysaccharide degradation is the ability to interact with the lattice surface of insoluble polysaccharides because of a flat substrate-binding surface.40 The reduced active site promotes polysaccharide binding (Supplemental Figures S7 and S8),14,41 implying that reduction of the PMO occurrs prior to the binding of the polysaccharide substrate. This is consistent with the proposed random-sequential mechanism, as the PMO can either bind O2 or Glc6 upon reduction of the copper active site (Scheme 2). The random binding of substrates also implies that the enzyme can uncouple if the second electron transfer to the E-O2– complex occurs before polysaccharide binding. Thus, precise temporal control of electron transfer is key to minimize the uncoupled reaction. In the absence of the polysaccharide, a reduced active site is not only prone to generate H2O2, but also to react with H2O2 with deleterious effects to PMO function.25,29,42 Although H2O2 reactivity proceeds much faster than that of O2,42 recent data suggests that there is limited H2O2 reactivity in the native PMO environment.29

The pH dependence of MtPMO9E turnover has also been observed in other PMOs. 43,44

The optimal pH range for these PMOs closely mirror the optimal pH range for

cellobiose dehydrogenase reduction.21 It is possible this pH dependence serves as a regulatory function to prevent uncoupled turnover. MtPMO9E exhibits similar pKa’s on kcat , kcat/KM(Glc6) and kcat/KM(O2) signifying the protonation of an base that is essential for chemistry and not involved in substrate binding (Figure 3). Similarly, a decrease in the apparent melting temperature of NcLPMO9C was observed below pH 5.0, indicative of a destabilizing protonation.41 Moreover, the binding of Glc6 and O2 did not perturb the pKa (Figure 3A). Hydrogen bonding networks can shift the pKa of amino acid side chains, such that the kinetic pKa of MtPMO9E could be attributed to the protonation of either an imidazole and carboxylate side chain, making it difficult to definitively assign the site of protonation. Interestingly, crystal structures of Ls(AA9)A, a PMO with 42% sequence identity to MtPMO9E, exhibit a pH-dependent rotation of a Cu-coordinating histidine.45 Upon substrate binding, the histidine rotates back into the Cu-equatorial plane,45 which is presumably the active


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Biochemistry

form of the enzyme. Nonetheless, the activity of MtPMO9E is still impaired at low pH, even with saturating concentrations of Glc6, suggesting other factors may govern O2 reactivity.

Conclusion

Taken together, the results indicate PMOs use a random-sequential kinetic mechanism to bind the substrates Glc6 and O2 (Scheme 2), providing insight into the chemical mechanism of fungal PMOs. Following reduction of the active site, the PMO can bind either substrate. The binding of the second substrate forms the a ternary Michaelis complex generally accepted as a Cu(II)-superoxo. The chemical steps after the formation of this complex are still unknown. Resolving these irreversible chemical steps will be crucial to the proposal of a complete chemical mechanism for PMOs. Additional studies, including kinetic isotope effects and advanced spectroscopic techniques, will be necessary to elucidate the reactive species responsible for HAA.

Acknowledgements

The authors would like to thank Stefan Kapczynski for assistance in the preparation of MtPMO9E and Professor Judith Klinman for the use of the Hansatech Oxygraph instrument and useful conversions. We would also like to thank Elise Span and Dr. Christopher Lemon for critical reading of the manuscript.

Notes

The authors declare no competing financial interest.

Supporting Information


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Supporting Information, including initial velocity vs. substrate plots, H2O2 suppression data and pH dependence controls, is available free of charge on the ACS Publications website.

Funding acknowledgement

This work was supported by the National Science Foundation (NSF grant #1565770).


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Biochemistry

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A random-sequential kinetic mechanism for polysaccharide monooxygenases

John A. Hangasky and Michael A. Marletta


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A Random-Sequential Kinetic Mechanism for Polysaccharide

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