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Kinetic Characterization of Prenyl-flavin Synthase from Saccharomyces cerevisiae Nattapol Arunrattanamook, and E. Neil G. Marsh Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.7b01131 • Publication Date (Web): 12 Dec 2017 Downloaded from http://pubs.acs.org on December 13, 2017

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Biochemistry Arunrattanamook and Marsh 2017

Kinetic Characterization of Prenyl-flavin Synthase from Saccharomyces cerevisiae

Nattapol Arunrattanamook ‡† and E. Neil G. Marsh*†§

†

Department of Chemistry, ‡Department of Chemical Engineering and §Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109, United States

* Corresponding Author: Prof. Neil Marsh e-mail: [email protected] Tel:

+ 734 763 6096

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Page 2 of 17 Arunrattanamook and Marsh 2017

ABSTRACT We have characterized the kinetics and substrate requirements of prenyl-flavin synthase from yeast.

This enzyme catalyzes the addition of an isopentenyl unit to reduced flavin

mononucleotide (FMN) to form an additional 6-membered ring that bridges N-5 and C-6 of the flavin nucleus, thereby converting the flavin from a redox cofactor to one that supports the decarboxylation of aryl carboxylic acids. In contrast to bacterial enzymes, the yeast enzyme was found to use dimethylallyl-pyrophosphate, rather than dimethylallyl-phosphate, as the prenyldonor in the reaction. We developed a coupled assay for prenyl-flavin synthase activity in which turnover was linked to the activation of the prenyl-flavin-dependent enzyme, ferulic acid decarboxylase. The kinetics of the reaction are extremely slow: kcat = 12.2 ± 0.2 h-1 and KM for dimethylallyl-pyrophosphate = 9.8 ± 0.7 µM; the KM for reduced FMN was too low to be accurately measured. The kinetics of reduced FMN consumption were studied under pre-steady state conditions. The reaction of FMN was well described by first-order kinetics with kapp = 17.4 ± 1.1 h-1. These results indicate that a chemical step, most likely formation of the carbon-carbon bond between C6 of the flavin and the isopentenyl moiety is substantially rate-determining in the reaction.

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Various strains of yeast decarboxylate substituted trans-phenylacrylic acid derivatives, including ferulic acid (4-hydroxy-3-methoxy-phenylacrylic acid) a breakdown product of lignin, to the corresponding styrene derivatives (Figure 1A). 1-3 This serves as a mechanism to detoxify these carboxylic acids that are inhibitory to the growth of the yeast.4, 5 The enzyme responsible for this reaction, ferulic acid decarboxylasea (FDC), has recently been shown to require a novel, modified flavin mononucleotide (FMN) derivative for activity termed prenyl-FMN (pr-FMN, Figure 1A). Pr-FMN contains an isopentyl moiety appended between the C6 and N5 positions of the isoalloxazine flavin nucleus to form an additional 6-membered ring .6, 7 This modification converts the flavin from a redox cofactor to one capable of stabilizing the negatively charged intermediates formed during decarboxylation.8, 9 For FDC, decarboxylation is proposed to occur through the formation of a covalent adduct between the substrate and pr-FMN by an unusual 1,3 cycloaddition reaction;7,

10-13

however, this mechanism may not be true for other pr-FMN-

dependent enzymes.14 Pr-FMN is synthesized by a specialized prenyl transferase that uses dimethylallyl phosphate to install the additional 6-membered ring on the reduced form of FMN (Figure 1B).6, 15

In yeast, this enzyme has been known as PAD1, an abbreviation for phenylacrylic acid

decarboxylase, as a result of early genetic studies that linked this gene to decarboxylase activity.1 However, it is now clear that this enzyme is not a decarboxylase and therefore we refer to it here as prenyl-FMN synthase (PFS) and propose this name be adopted in future. Homologs of FDC and PFS are found in many microorganisms, indicating that pr-FMNdependent decarboxylases are likely widespread in Nature.

In many bacteria, they are

represented by UbiD and UbiX proteins respectively, which form part of the ubiquinone biosynthesis pathway.16-19 UbiX has been characterized as a PFS and has been the subject of

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detailed crystallographic studies.15,

20

UbiD, a homolog of FDC, is believed to catalyze the

decarboxylation of ubiquinone biosynthetic intermediate 4-hydroxy-3-octaprenyl-benzoate21 although this activity has yet to be experimentally demonstrated. Recently, a second pr-FMNdependent enzyme has been characterized from Enterobacter cloacae and Klebsiella pneumoniae that catalyzes the reversible decarboxylation of 3,4-dihydroxybenzoic acid.14 A mechanism for pr-FMN synthesis has been proposed based on crystallographic studies of the PFS represented by UbiX from Pseudomonas aeruginosa.15 The enzyme catalyzes the reaction of reduced FMN with dimethylallylphosphate (DMAP). The first step is proposed to be nucleophilic attack of N5 on DMAP to form N5-alkylated FMN with the loss of the phosphate group.

Formation of the C6 bond occurs next, presumably by a mechanism involving

protonation of the dimethylallyl double bond. Finally, oxidation of the flavin nucleus is required to generate the active pr-FMN cofactor (Figure 1B). It is unclear whether, in vivo, this step occurs spontaneously or with the assistance of the cognate pr-FMN-dependent decarboxylase. A

B

Figure 1: A The decarboxylation reaction catalyzed by prenyl-FMN-dependent ferulic acid decarboxylase and the structure of prenyl-FMN. B The reaction catalyzed by the UbiX class of prenylflavin synthases between FMNH2 and DMAP. The reaction is thought to occur through reaction at N5 of the flavin first. The reduced pr-FMNH2 undergoes subsequent oxidation to the active pr-FMN cofactor.

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Here we describe studies on the eukaryotic PFS from Saccharomyces cerevisiae, scPFS (referred to in earlier literature as PAD1). We have developed a coupled assay for scPFS that has allowed us to determine the steady state kinetic parameters for enzyme. Moreover, whereas the previously studied bacterial enzymes use DMAP as a substrate, which is not commonly used by prenyl transferase enzymes, we find that scPFS prefers dimethylallylpyrophosphate (DMAPP) as a substrate, which is a common precursor in isoprenoid biosynthesis.

MATERIALS AND METHODS Materials. FMN, and dithiothreitol were purchased from MP biomedicals; dimethylallyl pyrophosphate was purchased from Isoprenoids LC, Tampa FL; potassium ferricyanide was purchased from Acros Organics. All other chemicals were purchased from Sigma-Aldrich. Holo-FDC, apo-FDC and scPFS (tPAD) were recombinantly expressed in E. coli and purified as described previously.6 Assay for pr-FMN formation by scPFS. Typically assays were performed in 100 mM HEPES buffer, pH 8.5, containing 1.0 mM MgCl2 at room temperature under anaerobic conditions. The buffer was purged with nitrogen gas and then equilibrated in a Coy anaerobic chamber. Other components of the assay, FMN, DMAP, DMAPP, scPFS, DTT, sodium dithionite and potassium ferricyanide were made up as stock solutions and transferred into the anaerobic chamber. Assays were typically performed at room temperature in 1.5 mL Eppendorf tubes with a total assay volume of 1 mL. It was found necessary to pre-reduce scPFS with 5 mM DTT prior to assay to obtain consistent activity measurements. Assays contained 400 - 600 nM scPFS and 4 µM apoFDC. The substrate concentrations varied between 0.75 to 50 µM for FMN and between 1 to 500 µM for DMAP or DMAPP. Reactions were initiated by addition of sodium dithionite (1 5


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mM final concentration), which rapidly reduced FMN to FMNH2. Reactions were quenched at various times by addition of potassium ferricyanide (5 mM final concentration) which rapidly oxidized the remaining FMNH2 and any pr-FMN formed. The amount of pr-FMN formed was determined from the activity of the re-constituted FDC as described below. Assay of reconstituted FDC. To determine how much pr-FMN had been generated by scPFS, the activity of FDC with cinnamic acid as substrate was measured as follows. 125 µL of the quenched scPFS reaction mixture was diluted to 500 µL with 100 mM HEPES buffer, pH 8.5, containing 1.0 mM MgCl2. The mixture was equilibrated at room temperature for 45 min to ensure that all pr-FMN had bound to FDC. Then, cinnamic acid was added to the assay to a final concentration of 2.5 mM.

After 4 min, the assay mixture was quenched with HCl, final

concentration 0.27 M, and the styrene produced extracted by vortexing with 400 µL ethyl acetate containing 94.7 µM undecane as an internal standard. The organic and aqueous phases were separated by centrifugation at 10,000 rpm for 10 min in a tabletop microcentrifuge. The amount of styrene produced was determined by GC-MS as described previously.6 HPLC analysis of FMN. Assays containing 100 µM DMAPP, 2 µM scPFS and 1.5 µM FMN were set up in a similar manner to those described above, but included 9 µM riboflavin as internal standard. After quenching with potassium ferricyanide, the amount of FMN in the samples was determined by reverse phase HPLC using a Shimadzu HPLC system equipped with a UV-visible diode array detector with monitoring at 450 nm. Chromatography was performed using EC 250/4.6 Nucleodur C18 “Gravity” reverse phase column. The column was equilibrated with 5 mM ammonium acetate (A) and analytes eluted with a gradient of increasing acetonitrile (B) at a flow rate of 0.5 mL/min. The solvent composition was held at 100% (A) for 4 min, and then increased linearly to 15% (B) over 16 min, followed by isocratic elution at 15% (B) for 25

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min, then returned to 100% (A) over 5 min and held for 5 min for column equilibration. Under these conditions FMN eluted at 24.5 min and riboflavin eluted at 35.7 min. FMN concentrations were calculated from peak integration and normalized with respect to the riboflavin internal standard.

RESULTS AND DISCUSSION Our initial efforts to develop a quantitative assay for scPFS activity attempted to exploit the fact that phosphate (Figure 1B) is produced during the reaction. Given that a variety of methods have been developed for orthophosphate and pyrophosphate detection, this appeared to be a promising strategy to assay PFS activity. Therefore, with DMAP as the substrate, we attempted to detect the orthophosphate formed during the reaction using commercial kits for phosphate detection. Neither assays based on orthophosphate binding to malachite green dye (Cayman Chemicals), nor assays based on the purine nucleoside phosphorylase-catalyzed reaction of orthophosphate with 2-amino-6-mercapto-7-methylpurine ribonucleoside (EnzChek® phosphate Assay Kit) proved successful in measuring scPFS activity. Either unacceptably high background rates were encountered (malachite green assay) or other components of the assay interfered with the colorimetric measurement (enzymatic assay).

These problems were

exacerbated by the very slow rate at with scPFS catalyzes the formation of pr-FMN. Later attempts to measure enzymatically the production of pyrophosphate, using DMAPP as substrate and an assay based on the successive reactions of maltose phosphorylase; glucose oxidase and horseradish peroxidase (PiPer™ Pyrophosphate Assay kit) were also unsuccessful. In this case, the pyrophosphate assay proved incompatible with other components of the scPFS assay and yielded false positive signals.

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Development of a coupled assay for scPFS. Given the difficulties encountered with measuring phosphate production, we investigated the possibility of coupling the production of pr-FMN to the activation of FDC. This would allow the formation of one molecule of pr-FMN to be coupled to the production of many molecules of styrene, thereby affording a considerable amplification of the signal. For the purposes of the assay fully apo-FDC was necessary to prevent background activity.

This was purified from an UbiX-deficient E. coli strain as

described previously.6 For the response of this assay to be linear, it is necessary to assume that all the pr-FMN synthesized by scPFS is taken up by apo-FDC so that the activity of FDC measured in the second step reflects the amount of pr-FMN synthesized. This requires both that the concentration of apo-FDC in the assay is much higher than that of scPFS and that the affinity of FDC for pr-FMN be relatively high. Although the Kd for FDC binding pr-FMN is not known, the fact that holoFDC can be purified intact from recombinant E. coli cells that over-express scPFS suggests that pr-FMN is bound quite tightly. In contrast, scPFS does not appear to bind pr-FMN tightly because when this enzyme was purified from recombinant E. coli cells and characterized, only residual FMN was found bound to the protein.6 Preliminary experiments determined that scPFS activity could be reliably measured using FDC as a coupling enzyme and established the optimal conditions for the assay. Typically assays were performed in 100 mM HEPES buffer pH 8.5 containing 1.0 mM MgCl2 in final volume of 1 mL in a Coy anaerobic chamber at 20o C. Assays contained 400 nM scPFS that was pre-incubated with 5 mM DTT for 10 min. This step was found to be necessary to obtain consistent activity measurements; presumably this reduces disulfide bonds between cysteine residues in the protein that may have become oxidized during the purification of the enzyme.

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After reduction of scPFS, 50 µM FMN and 4 µM apo-FDC, (final concentrations) were added to the assay, followed by addition of either DMAP or DMAPP to 500 µM final concentration. The reaction was initiated by addition of sodium dithionite to 1 mM final concentration, which rapidly reduced the FMN. After time periods varying from 30 s to 40 min, the reaction was stopped by addition of potassium ferricyanide to 5 mM final concentration. This rapidly oxidized unreacted FMNH2, thereby stopping the reaction, and also efficiently oxidized the reduced pr-FMNH2 formed in the reaction to the catalytically active form (re-oxidation of pr-FMNH2 in air gave less consistent results). The assays were removed from the anaerobic chamber and the oxidized mixture was allowed to stand for 45 min to ensure complete binding of pr-FMN to apo-FDC. 125 µL of the assay mixture was diluted to 500 µL in assay buffer and the activity of the reconstituted FDC was measured by addition of trans-cinnamic acid, 2.5 mM final concentration. The KM for trans-cinnamic acid is 0.18 mM,6 so the reaction is effectively run under Vmax conditions. After 4 min, the assay mixtures were quenched by addition of HCl, 0.27 M final concentration, and the styrene produced was extracted with ethyl acetate and quantified by GC-MS as described previously.6 The amount of pr-FMN produced was calculated from the amount of styrene produced in the assay per unit time, assuming that FDC catalyzes the conversion of 276 mol of styrene per min for each mol active enzyme.6 Identity of the prenyl donor in the reaction. Guided by reports that the bacterial PFS was specific for DMAP,7 our initial experiments on the eukaryotic enzyme used DMAP as a substrate. However, the very low levels of activity obtained with DMAP caused us to reevaluate whether DMAPP, which is a much more common prenyl-donor, might be a substrate for scPFS. This was found to be the case. The rate of pr-FMN formation was compared with either

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100 µM DMAPP or 500 µM DMAP present as the prenyl-donor, conditions that represent saturating concentrations of these substrates. Under these conditions, scPFS exhibited ~20-fold higher activity with DMAPP than DMAP (Figure 2); therefore, further experiments were performed using DMAPP as the substrate.

Figure 2: Comparison of the rates of FMN prenylation catalyzed by scPFS with either DMAP (500 µM) (blue squares) or DMAPP (100 µM) (red circles) as the prenyl donor. For details see the text.

Steady state kinetic analysis of scPFS. Having demonstrated that the FDC-coupled assay gave linear and reproducible results, we investigated the kinetics of pr-FMN formation by scPFS. These assays employed 400 nM scPFS that had been pre-reduced with 5 mM DTT for 10 min and a 10-fold molar excess of apo-FDC. To determine the KM for DMAPP, the concentration of FMN was fixed at 50 µM (assumed saturating) and the concentration of DMAPP varied between 1 and 500 µM (Figure 3A). The data were fitted to the Michaelis-Menten equation to give KMapp = 9.8 ± 0.7 µM for DMAPP and kcatapp = 12.2 ± 0.2 h-1. To investigate the KM for FMN, the concentration of DMAPP was fixed at 500 µM (assumed saturating) and the concentration of

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FMN varied between 0.75 and 50 µM (Figure 3B). From these measurements, the apparent KM for FMN was estimated as 0.61 ± 0.17 µM. However, we note that at the lowest concentrations of FMN used in these assays the assumptions of the Michaelis-Menten equation are not valid and therefore this value should be regarded as an upper bound.

B

A

Figure 3: Steady state kinetic analysis of scPFS measured using the FDC-coupled assay. A: DMAPP as the variable substrate; B: reduced FMN as the variable substrate. The data are shown fitted into Michaelis-Menten equation.

Single turnover kinetics of FMN consumption. Taking advantage of the very slow turnover rate exhibited by scPFS, we sought to examine the kinetics in more detail under single turnover conditions. To accomplish these measurements, assays were set up with higher concentrations of scPFS, typically 600 nM, and sub-stoichiometric concentrations of FMN, typically 300 nM. Given affinity of scPFS for FMN, estimated from the steady-state measurements, most of the FMN in the assay should be bound to the enzyme at these concentrations. The other components of the assay were present at the same concentrations as for the steady-state measurements. The reaction was initiated by the addition of DMAPP (500 µM final concentration). At various times aliquots were withdrawn and the quenched by addition of K3Fe(CN)6. The amount of pr-FMN

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formed was then determined from the activity of FDC, as described above.

Under these

conditions the production of pr-FMN was well described by a first-order kinetic model (Figure 4) with an apparent first order rate constant of 17.5 ± 1.1 h-1.

Figure 4: Pre-steady state kinetic analysis of scPFS under single turnover conditions. In separate experiments the reaction was followed either monitoring the production of pr-FMN using the FDC-coupled assay (red circles), or by monitoring the decrease in FMN (blue squares). For details see the text.

The pre-steady state formation of pr-FMN by scPFS could also be followed more directly by the disappearance of FMN. We initially attempted to detect the formation of pr-FMN by reverse phase HPLC; however, the small amount of product formed and the absence of a strongly absorbing long-wavelength chromophore, combined with the instability of the cofactor and interference from contaminating analytes prevented us from reliably being able to quantify prFMN by this method. In contrast, consumption of FMN could be followed relatively easily by quantifying the FMN remaining in the assay by HPLC. FMN was detected by chromatography on a C-18 reverse phase HPLC column equilibrated in 5 mM ammonium acetate and developed

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with an increasing gradient of acetonitrile, with peaks monitored at 450 nm, as described in the Methods section. The pre-steady state kinetics of FMN consumption mirrored that formation of pr-FMN followed by the discontinuous coupled assay described above. The data were well fitted by a single exponential with an observed rate constant, kobs = 20.3 ± 3.8 h-1, a result that is, within error, the same as the rate constant for pr-FMN formation (Figure 4). The pre-steady state measurements indicate that a chemical step is significantly rate determining in the mechanism of scPFS, as the rate constants are only slightly faster than kcat measured under steady state conditions. Most likely, formation of the carbon-carbon bond at C6 of the flavin ring is the slow step. This hypothesis is supported by the fact that for the bacterial PFS, UbiX, the intermediate prior to formation of the carbon-carbon bond accumulates during the reaction; this intermediate, in which the N5 position of the flavin is prenylated, was observed crystallographically.

Furthermore, although the mechanism remains unclear, carbon-carbon

bond formation at C6 must involve transient loss of the aromatic ring system, which would be energetically unfavorable. In conclusion, we have developed a coupled assay that has allowed us to undertake the first kinetic analyses of the newly discovered class of flavin prenyl transferases. Our findings demonstrate that, in contrast to the bacterial enzymes that are specific for DMAP, the yeast PFS utilizes DMAPP far more efficiently than DMAP. Whereas DMAPP is the starter unit for isoprenoid biosynthesis in both prokaryotic and eukaryotic organisms, DMAP is not a common metabolite and its biosynthetic origins – whether by hydrolysis of DMAPP or phosphorylation of prenol – currently remain unclear. Somewhat surprisingly, it appears that scPFS catalyzes the formation of reduced pr-FMN extremely slowly, with kcat only ~ 12 h-1. This may reflect the energetic barrier to forming a carbon-carbon bond between the isoprene moiety and the aromatic

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flavin ring. We also note that the slow kinetics may reflect a lack of evolutionary pressure to become more catalytically proficient, because, as a cofactor, pr-FMN is expected to be required in only very small quantities in the cell. AUTHOR INFORMATION Corresponding author: *email: [email protected], Phone: 734 763 6096 Funding This work was supported by National Science Foundation grant CHE 1152055, to E.N.G.M.; N.A. acknowledges the support of a Royal Thai Government Scholarship. Notes The authors declare no conflict of interest. ABBREVIATIONS FDC, ferulic acid decarboxylase; pr-FMN, prenylated flavin mononucleotide; scPFS, Saccharomyces cerevisiae prenyl-flavin synthase; PAD1, phenylacrylic acid decarboxylase; DMAP, dimethylallyl phosphate; DMAPP, dimethylallyl pyrophosphate

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REFERENCES [1] Clausen, M., Lamb, C. J., Megnet, R., and Doerner, P. W. (1994) Pad1 Encodes Phenylacrylic Acid Decarboxylase Which Confers Resistance to Cinnamic Acid in Saccharomyces cerevisiae, Gene 142, 107-112. [2] Goodey, A. R., and Tubb, R. S. (1982) Genetic and Biochemical-Analysis of the Ability of Saccharomyces cerevisiae to Decarboxylate Cinnamic-Acids, J. Gen. Microbiol. 128, 2615-2620. [3] Huang, Z. X., Dostal, L., and Rosazza, J. P. N. (1994) Purification and Characterization of a Ferulic Acid Decarboxylase from Pseudomonas fluorescens, J. Bacteriol. 176, 5912-5918. [4] Baranowski, J. D., Davidson, P. M., Nagel, C. W., and Branen, A. L. (1980) Inhibition of Saccharomyces cerevisiae by Naturally-Occurring Hydroxycinnamates, J. Food Sci. 45, 592-594. [5] Stratford, M., Plumridge, A., and Archer, D. B. (2007) Decarboxylation of sorbic acid by spoilage yeasts is associated with the PAD1 gene, Appl. Environ. Microbiol. 73, 6534-6542. [6] Lin, F., Ferguson, K. L., Boyer, D. R., Lin, X. N., and Marsh, E. N. G. (2015) Isofunctional Enzymes PAD1 and UbiX Catalyze Formation of a Novel Cofactor Required by Ferulic Acid Decarboxylase and 4-Hydroxy-3-polyprenylbenzoic Acid Decarboxylase, ACS Chem. Biol.y 10, 1137-1144. [7] Payne, K. A. P., White, M. D., Fisher, K., Khara, B., Bailey, S. S., Parker, D., Rattray, N. J. W., Trivedi, D. K., Goodacre, R., Beveridge, R., Barran, P., Rigby, S. E. J., Scrutton, N. S., Hay, S., and Leys, D. (2015) New cofactor supports alpha,beta-unsaturated acid decarboxylation via 1,3dipolar cycloaddition, Nature 522, 497-501. [8] Leys, D., and Scrutton, N. S. (2016) Sweating the assets of flavin cofactors: new insight of chemical versatility from knowledge of structure and mechanism, Curr. Opin. Struct. Biol. 41, 19-26. [9] Piano, V., Palfey, B. A., and Mattevi, A. (2017) Flavins as Covalent Catalysts: New Mechanisms Emerge, Trends Biochem. Sci. 42, 457-469. [10] Ferguson, K. L., Arunrattanamook, N., and Marsh, E. N. G. (2016) Mechanism of the Novel Prenylated Flavin-Containing Enzyme Ferulic Acid Decarboxylase Probed by Isotope Effects and Linear Free-Energy Relationships, Biochemistry 55, 2857-2863. [11] Ferguson, K. L., Eschweiler, J. D., Ruotolo, B. T., and Marsh, E. N. G. (2017) Evidence for a 1,3Dipolar Cyclo-addition Mechanism in the Decarboxylation of Phenylacrylic Acids Catalyzed by Ferulic Acid Decarboxylase, J. Am. Chem. Soc. 139, 10972-10975. [12] Tian, G., and Liu, Y. J. (2017) Mechanistic insights into the catalytic reaction of ferulic acid decarboxylase from Aspergillus niger: a QM/MM study, Phys. Chem. Chem. Phys. 19, 77337742. [13] Lan, C. L., and Chen, S. L. (2016) The Decarboxylation of alpha,beta-Unsaturated Acid Catalyzed by Prenylated FMN-Dependent Ferulic Acid Decarboxylase and the Enzyme Inhibition, J. Org. Chem. 81, 9289-9295.

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Kinetic Characterization of Prenyl-flavin Synthase from

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