STRUCTURAL AND MECHANISTIC CHARACTERIZATION OF ENZYMES IN

PERSULFIDE OXIDATION AND MONOLIGNOL

BIOSYNTHESIS PATHWAYS

By

STEVEN ANDREW SATTLER

A dissertation submitted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY School of Molecular Biosciences

MAY 2018

© Copyright by STEVEN ANDREW SATTLER, 2018 All Rights Reserved

© Copyright by STEVEN ANDREW SATTLER, 2018 All Rights Reserved To the Faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of STEVEN

ANDREW SATTLER find it satisfactory and recommend that it be accepted.

ChulHee Kang, Ph.D., Chair

Susan Wang, Ph.D.

Luying Xun, Ph.D.

Margaret Black, Ph.D.

ii ACKNOWLEDGMENT

There are many people who I would like to thank for helping place me in a position to earn this degree. Foremost, I would like to thank my academic advisors, friends, and family for their guidance, support, and patience while I spent several years as an undergraduate and doctoral student. Most of my achievements are attributed to your respective influences on my education or disposition, and for that I will always be grateful. Even if you are not mentioned by name in this acknowledgment, know that your influence is appreciated.

I would first like to thank my doctoral advisor, Dr. ChulHee Kang, for providing me with an opportunity to earn this degree. His guidance and willingness to listen to my ideas were crucial to my development as a scientist and prepared me for a professional career in theoretical work. I would also like to thank Dr. Luis Matos, who provided me with my first research opportunity as an undergraduate. I partially owe any success that I have had as a graduate student to his guidance.

The final academician I would like to recognize by name is Susan Wick Johnson. As a teacher and friend, she was the first person to guide me in understanding the rewards of scholastic success.

To members of my graduate research group, both past and present, I would like to thank

Abigail Green, Robert Hayes, Kevin Lewis, Timothy Moural, Alexander Walker, Se-Young Jun, and Shin-Hwa Wu. The alliances we have developed through the years are both personal and professional in nature, and I would like to thank all of you for your friendship and collaboration as we were all trying to make difficult transitions in our lives. I will never forget the impact that that our relationships had and continue to have on my development as a scientist and person.

To my family and close friends, the ways and magnitudes of how you have affected my life for the better are immeasurable. I would like to thank my parents, Scott and Stacie Sattler and

Roni and Bill Jarrell, my grandparents Richard and Sandy Stone and Edith and Larry Sattler, and

iii my siblings Chris Sattler, Brian Sattler, Brook Sattler, J.R. Jarrell, Carlee Jarrell, Austin Sattler, and Corey Jarrell. I would also like to thank April Frers, Rahul Dhal, Rob Jones, Josh Swider,

Justin Johnson, Jason Price, Deborah Bohnen, Enrique Alvarado, and Brett Vanderwurff. I have always greatly appreciated your friendship, love and support, and have learned valuable lessons from each of you. Finally, I would like to thank the administrative staff of the School of Molecular

Biosciences, particularly Kelly McGovern and Tami Breske. I never missed a deadline because of them.

iv STRUCTURAL AND MECHANISTIC CHARACTERIZATION OF ENZYMES IN

PERSULFIDE OXIDATION AND MONOLIGNOL

BIOSYNTHESIS PATHWAYS

Abstract

by Steven Andrew Sattler, Ph.D. Washington State University May 2018

Chair: ChulHee Kang

We conducted investigations of the structural and biochemical properties of two distinct types of enzyme from different pathways: cinnamoyl-CoA reductases (CCRs) and the persulfide dioxygenases (PDOs). CCR is an enzyme of the monolignol biosynthesis pathway in plants that uses NADPH to reduce any of three major hydroxycinnamoyl-CoA thioesters, thus catalyzing the formation of hydroxycinnamaldehydes. These aldehydes are further reduced by cinnamyl alcohol dehydrogenase (CAD) to form the alcohol substrates required for lignin biosynthesis. One CCR,

SbCCR1, was crystallized in the presence of NADPH and the structure was determined at 2.9 Å resolution. It was determined—through site-directed mutagenesis, ITC, molecular docking, and kinetics assays—that not only is feruloyl-CoA the preferred substrate for SbCCR1, but residues

Thr154 and Tyr310 are essential to binding functional groups about the aromatic ring of the substrate. Production of a T154Y mutant of SbCCR1 resulted in significant reduction in substrate preference for feruloyl-CoA over p-coumaroyl-CoA, providing support for the hypothesis that substitutions can be made at this position leading to favorable reductions in or softening of lignin

v content in S. bicolor. Furthermore, confirmed the existence of an additional CCR in S. bicolor through multiple-sequence alignment and kinetics analyses.

PDOs catalyze the oxidation of a sulfane sulfur on glutathione persulfide (GSSH) or higher glutathione polysulfanes (GSSnH, where n > 1) during the process of detoxifying dihydrogen sulfide (H2S). In this study, the structures of PDOs from M. xanthus (MxPDO1) and P. putida

(PpPDO2) were determined in the presence of a catalytic iron, as well as in the presence of both iron and product molecule glutathione for the PDO from P. putida. Structure alignments with other

PDOs, in addition to the 1.46 Å PpPDO2/GSH complex structure, revealed key substrate-binding residues and provided a basis for activity differences between PDOs and a closely related group of enzymes, the glyoxalases II. Static light scattering further showed that, despite high identity to the glyoxalases II, PDOs are likely dimeric. In summary, we described interactions between PDOs and GSH, as well as provided information as to how two highly related groups of enzymes could differ in form and function.

vi TABLE OF CONTENTS

Page

ACKNOWLEDGMENT...... iii

ABSTRACT ...... v

LIST OF TABLES ...... ix

LIST OF FIGURES ...... x

CHAPTERS

CHAPTER ONE: INTRODUCTION ...... 1

1.1 General Overview ...... 1

1.2 Function of CCR ...... 1

1.3 Mutations to Monolignol Biosynthetic Enzymes and the “Lignin Problem” ...... 1

1.4 Function of PDOs ...... 2

1.5 PDOs and Ethylmalonic Encephalopathy (EE) ...... 3

1.6 Structural and Evolutionary Relationships between PDOs and the Glyoxalases II ...... 3

1.7 Chapter Summaries ...... 5

1.8 References ...... 10

CHAPTER TWO: CHARACTERIZATIONS OF TWO PERSULFIDE DIOXYGENASES OF THEMETALLO-β-LACTAMASE SUPERFAMILY ...... 14

2.1 Contributions...... 14

2.2 Abstract ...... 14

2.3 Introduction ...... 15

2.4 Materials and Methods ...... 17

2.5 Results ...... 21

vii

2.6 Discussion ...... 24

2.7 Acknowledgments...... 28

2.8 References ...... 43

CHAPTER THREE: STRUCTURAL AND BIOCHEMICAL CHARACTERIZATION OF CINNAMOYL-COA REDUCTASES ...... 47

3.1 Contributions...... 47

3.2 Abstract ...... 47

3.3 Introduction ...... 48

3.4 Materials and Methods ...... 51

3.5 Results ...... 56

3.6 Discussion ...... 62

3.7 Conclusion ...... 67

3.8 Acknowledgments...... 68

3.9 References ...... 85

CHAPTER FOUR: CONCLUSIONS...... 92

viii LIST OF TABLES

Table 2.1: Crystallographic data for the PpPDO2 and MxPDO1b structures ...... 29

Table 2.2: Thermodynamic parameters for binding substrate analogs by PpPDO2 ...... 30

Table 3.1: X-ray diffraction data and refinement statistics for SbCCR1 (PDB identifier 5TQM) ...... 69

Table 3.2: Thermodynamic properties of interaction between SbCCR1 and various ligands ...... 70

Table 3.3: Thermodynamic properties of interaction between wild-type SbCCR1 or mutants T154A and Y310F and feruloyl-CoA ...... 71

Table 3.4: Kinetic values for wild-type SbCCR1 in the presence of three hydroxycinnamoyl- CoA substrates ...... 72

Table 3.5: Kinetic values for wild-type and mutant forms of SbCCR1, SbCCR2, Sobic.002G146000, and Sobic.010G066000 in the presence of feruloyl-CoA or p-coumaroyl- CoA ...... 73

ix LIST OF FIGURES

Page

Figure 1.1: A cladographic illustration of the relationships between the three types of persulfide dioxygenase, the glyoxalases, and two metallo-β-lactamases ...... 7

Figure 1.2: Modeling of the CnPDO1 active site within experimentally derived electron density ...... 9

Figure 2.1: Ribbon diagrams representing the oligomeric and superimposed structures of PDOs ...... 32

Figure 2.2: GSH complex and ligand-free forms of PpPDO2 ...... 33

Figure 2.3: The oligomeric state of PpPDO2 in solution...... 35

Figure 2.4: Measurements of PpPDO2 binding for the product and substrate analogs via ITC ....37

Figure 2.5: Multiple sequence alignment of four PDOs and three glyoxalases II ...... 38

Figure 2.6: Metal-binding and secondary coordination sphere architectures of PDOs and glyoxalases II ...... 40

Figure 2.7: Active site representations of PpPDO2 and human glyoxalaseII in complex with GSH...... 42

Figure 3.1: Ribbon diagram of the global structure of SbCCR1 in complex with NADP+ and manually docked feruloyl-CoA...... 74

Figure 3.2: (A) The observed NADP+ in the binding pocket of SbCCR1; (B) Coniferaldehyde docked into the putative phenylpropanoid-binding region of SbCCR1 ...... 75

Figure 3.3: (A) ITC curves for wild-type SbCCR1 upon titration with various ligands; (B) ITC comparison between wild-type SbCCR1 and mutants T154A and Y310F upon titration with feruloyl-CoA ...... 76

Figure 3.4: Multiple sequence alignment of the amino acid sequences for SbCCR1 and several related enzymes ...... 77

Figure 3.5: Superimposed tertiary structures of SbCCR1 and other CCRs ...... 79

Figure 3.6: (A) Michaelis-Menten curves for wild-type SbCCR1 in the presence of three hydroxycinnamoyl-CoA substrates; (B) Michaelis-Menten curves for wildtype SbCCR1 and two mutant forms, T154A and Y310F, in the presence of feruloyl-CoA ...... 80

x

Figure 3.7: (A) Reaction velocities in the presence of feruloyl-CoA or p-coumaroyl-CoA for several forms of CCR; (B) Thr-154 and Tyr-310 in SbCCR1 are shown binding functional groups of coniferaldehyde; (C) Fold-recognition model of the active site of Sobic.004G065600 (SbCCR3) ...... 82

Figure 3.8: Proposed catalytic reaction mechanism of SbCCR1 ...... 84

xi Dedication

To Chris and Brian.

xii

CHAPTER ONE

INTRODUCTION

1.1 General Overview

Structural and biochemical characterization of proteins is the central focus of the research performed in this laboratory. Throughout the course of this research and joined with such techniques as fast protein liquid chromatography (FPLC), site-directed mutagenesis, isothermal titration calorimetry (ITC), enzyme kinetics assays and static light scattering, X-ray diffraction has been used to determine the molecular structures of multiple proteins and any relevant ligand complexes. Characterizations of a cinnamoyl-CoA reductase (CCR) from Sorghum bicolor and two persulfide dioxygenases (PDOs) from Pseudomonas putida and Myxococcus xanthus are presented here from the results obtained by employing these techniques.

1.2 Function of CCR

CCR catalyzes the penultimate reaction in the monolignol biosynthesis pathway of

S. bicolor (1). In short, it produces the phenylpropanals that are reduced by cinnamyl alcohol dehydrogenase (CAD) to yield phenylpropanols guaiacyl (G), syringyl (S), and p-hydroxyphenyl

(H) alcohols (2). The resulting phenylpropanols then undergo oxidative radical coupling to produce the mature lignin heteropolymers (3). This dissertation provides information for the first published structure of monomeric enzyme CCR from S. bicolor (4), which uses the reducing power of NADPH to reduce phenypropanoyl-CoA thioesters (5).

1.3 Mutations to Monolignol Biosynthetic Enzymes and the “Lignin Problem”

Lignin is a complex heteropolymer composed of radically coupled G, S, and H monolignols

(Fig. 2) (6,7). Seemingly constructed in random fashion with undetermined selectivity for specific monolignols at any time during its polymerization, lignin provides hydrophobicity for water

1

transport in the xylem, structural rigidity to the plant, and protects against injurious agents (4,8).

Though absolutely required for plant viability, the presence of lignin surrounding hemicellulosic material presents a barrier to extraction of polysaccharides from the plant by bioethanol production facilities. Being energetically and financially expensive, as well as perhaps ecologically hazardous do to the chemicals used in thermochemical pre-treatment, the ability to modulate plant lignin biosynthesis is perhaps the most desirable mechanism of control for improving cost efficiency of biofuel production (9).

That there are natural mutations to maize and sorghum monolignol biosynthetic enzymes resulting in viable plants with reduced or softer lignin sets precedence for generation of additional mutants with similar phenotypes (10). The natural mutants, presenting with characteristic brown mid-ribs in the leaves due to high phenylpropanoic acid deposition, are intuitively referred to as

“BMR” mutants (10). BMR mutants known at present include bmr2, bmr6, and bmr12, corresponding respectively to mutations in 4-coumarate ligase (4CL), CAD, and caffeate O- methyltransferase (COMT) (10-12). The crystal structures determined for phenylalanine ammonia-lyase (PAL), caffeoyl-CoA O-methyltransferase (CCoAOMT), hydroxycinnamoyltransferase (HCT), COMT, CAD, peroxidase (PRX), and CCR in the presence or absence of substrates and products have provided information required for rational mutant design according to residue function. An example of such a study is provided in Chapter Three

(2,4,11,13-16).

1.4 Function of PDOs

PDOs are metalloenzymes critically important for the oxidation of H2S (17,18). These proteins oxidize the terminal sulfane sulfer of GSSnH, leading to the production of cations sulfite and sulfate for excretion (19). Our work in characterizing PDOs from P. putida and M. xanthus

2

provided not only the first datasets for bacterial PDOs, but the first dataset for a PDO in complex with reaction product glutathione (20). These data yielded information about residues functional in active-site binding of GSH, giving insight on the molecular basis for disease ethylmalonic encephalopathy.

1.5 PDOs and Ethylmalonic Encephalopathy (EE)

Ethylmalonic encephalopathy (EE) is a severe, early-onset mitochondrial disorder that is invariably fatal (21). Signs and symptoms include vascular lesions leading to petechiae purpura, orthostatic acrocyanosis, brain failure, failure to thrive, and a host of others (21,22). Those suffering from the disease, which is often leads to death by the age of ten years, also present with underdeveloped muscle tone and an unusually wide gait (23,24). The first case report was made in

1991, but it was not until 2004 that the EE locus was mapped to chromosome 19; its gene then being aptly named ethe1 (22,24). It was discovered shortly after that that ethe1 encodes a persulfide dioxygenase, and mutations to the gene would ultimately disrupt normal catabolism of H2S

(18,23). Since the earlier days of the gene’s identification, the structure of human persulfide dioxygenase – the protein being referred to as hETHE1 in the original report – and the GSH complex with PpPDO2 reported shortly after led to understanding of the functional significance of residues implicated in the disease state (17,20).

1.6 Structural and Evolutionary Relationships between PDOs and the Glyoxalases II

The glyoxalase system of enzymes, present in both eukaryotic and prokaryotic systems, is a GSH-dependent system that catalyzes the oxidation of 2-oxoaldehydes in a two-step process

(25,26). First, glyoxalase I (GloA) performs an isomerization of the GSH/2-oxoaldehyde conjugates that form readily without enzymatic catalysis (26). Glyoxalase II (GloB) uses a

3

dinuclear metallic center, with both catalytic metals tending to be zinc, along with a co-substrate water molecule to produce lactate as a final oxidative step (27,28).

Along with the recent discovery of PDO apo-form and complex structures through X-ray data acquisition and processing, several groups have noted the close relationships between PDOs and GloB enzymes (Fig. 1) (17,20,29). The two types of enzyme contain metallo-β-lactamase folds and generally align well with respect to Cα positioning in most regions outside of their active sites

(20). Curiously, although both enzymes contain similar seven-residue, histidine-rich sequences with involvement in metal- or substrate-binding, the degree of involvement of these residues is dependent on enzyme type and thus far has not been rationalized with structural data that is currently available (20). Therefore, the structural basis for the two enzymes’ differences in metal- binding capabilities is not understood.

To date, all structures produced from X-ray data for PDOs indicate that the proteins are exclusively dimeric (17,20). That crystal packing for these enzymes represents the true oligomeric state is supported by several lines of evidence, including static light scattering experiments with

PpPDO2 by our own group (17,20). In contrast to PDOs, the GloBs are shown to be exclusively monomeric through X-ray diffraction and biochemical experiments (20,30). That these two enzymes differ structurally at the quaternary level, given their differences in metal-binding preferences or capabilities and significant structural similarities, two reciprocal hypotheses to explain this phenomenon may be proposed: metal-binding capability dictates oligomerization state, or oligomerization state dictates metal-binding capability.

Recent unpublished experiments with an ETHE1 from Cupriavidus necator (herein referred to as CnPDO1 for consistency with earlier reports) may provide insight into nature of the metal-binding/oligomerization relationship between the two enzymes. The crystal structure for

4

CnPDO1 has been determined at 1.4 Å resolution and its tertiary structure aligns closely with that of a previously published crystal structure for MxPDO1 (r.m.s.d = 1.30 Å). Such close alignment does not surprise given their sequence identity (54%), however CnPDO1 was monomeric according to its crystal packing and bound a single iron atom with an unprecedented four-histidine, one-aspartate pentad of residues (Fig. 2). Such an arrangement would preclude binding of GSH or molecular oxygen – both of which are required for PDO activity – and thus eliminate the possibility that CnPDO1 was crystallized in its natural form.

Upon examination of the enigmatic CnPDO1 structure and comparisons with several

GloBs, it was noticed that CnPDO1 used a subset of the seven-residue, histidine-rich motif shared between the PDOs and GloBs. Further examination of the electron density difference maps following a series of refinements revealed a large molecule bound in what would be the dimerization interface for the PDO1s. Additional refinements using known solvent molecules failed to clear density in the difference map in that region, possibly suggesting that multiple types of solvent molecule or multiple configurations of the same molecule were bound in the region prior to crystallization. Purification or storage buffer components were not large enough for adequate placement within the region. Additional crystallization trials are needed to rule out the possibility that there is a solvent molecule disrupting dimerization.

1.7 Chapter Summaries

The first chapter of this dissertation includes introductory information supplementary to the introductions provided in the second and third chapters. While the first chapter was produced uniquely for this dissertation, the latter two chapters are each composed entirely of published, first-author manuscripts. Consequently, the reference lists are provided at the end of their respective chapters in order to preserve original publication reference formats. The citation style

5

used in Chapter One will be that of the Journal of Biological Chemistry to provide stylistic consistency with the manuscript in the second chapter.

In Chapter Two, published structural and biochemical information is provided for two bacterial PDOs and one complex between a PDO and GSH. From the data gathered, we draw comparisons between a human PDO (referred to as hPDO in the report, but is commonly referred to as hETHE1) and the bacterial PDOs as it applies to understanding the molecular basis for ethylmalonic encephalopathy. We also present information about the proteins’ oligomeric states in aqueous solutions and align their primary structures with multiple PDOs for other domains of life.

Chapter Three includes the published manuscript describing structural and biochemical characteristics of monolignol biosynthesis enzyme CCR. As in the second chapter, data and descriptions are presented that provide insight on the oligomeric structure of CCRs, primary structure alignments, the complex between SbCCR1 and its co-product NADP+. Additionally, we present the results of kinetics assays and provide evidence for the existence of another CCR in

S. bicolor. Molecular docking results from this study further contributed to understanding of how

CCRs interact with phenylpropanoid substrates and products via identification of residues that interact directly with hydroxyl or methoxy functional groups around the aromatic ring of phenylpropanoids. From those results, site-directed mutagenesis was performed to produce CCR mutants with altered kinetic activity. The final chapter concludes this dissertation and provides a summary of our findings in addition to future directions in each project.

6

7

Figure 1.1: A cladographic illustration of the relationships between the three types of persulfide dioxygenase, the glyoxalases, and two metallo-β-lactamases. The PDOs are divided into three sub-types: β-lactmase-like-hydrolase (Blh), SdoA (PDO2), and ETHE1

(PDO1). Close structural relationships exist between PDOs and GloB2 enzymes, particularly with respect to the histidine-rich sequences conserved between them that are involved in binding catalytic metals.

8

Figure 1.2: Modeling of the CnPDO1 active site within experimentally derived electron density. The central gray star represents the catalytic Fe(II). Pink stars indicate water molecules. The blue mesh surrounding the stick model represents electron density. As is evident from this original model, four histidines and a single aspartate surround the metal, a peculiar arrangement for PDOs.

9

1.8 References

1. Pan, H., Zhou, R., Louie, G. V., Muhlemann, J. K., Bomati, E. K., Bowman, M. E., Dudareva, N., Dixon,

R. A., Noel, J. P., and Wang, X. (2014) Structural studies of cinnamoyl-CoA reductase and cinnamyl- alcohol dehydrogenase, key enzymes of monolignol biosynthesis. The Plant cell 26, 3709-3727

2. Jun, S. Y., Walker, A. M., Kim, H., Ralph, J., Vermerris, W., Sattler, S. E., and Kang, C. (2017) The

Enzyme Activity and Substrate Specificity of Two Major Cinnamyl Alcohol Dehydrogenases in Sorghum

(Sorghum bicolor), SbCAD2 and SbCAD4. Plant physiology 174, 2128-2145

3. Ralph, J., Guillaumie, S., Grabber, J. H., Lapierre, C., and Barriere, Y. (2004) Genetic and molecular basis of grass cell-wall biosynthesis and degradability. III. Towards a forage grass ideotype. C R Biol 327,

467-479

4. Sattler, S. A., Walker, A. M., Vermerris, W., Sattler, S. E., and Kang, C. (2017) Structural and

Biochemical Characterization of Cinnamoyl-CoA Reductases. Plant physiology 173, 1031-1044

5. Sarni, F., Grand, C., and Boudet, A. M. (1984) Purification and properties of cinnamoyl-CoA reductase and cinnamyl alcohol dehydrogenase from poplar stems (Populus X euramericana). European journal of biochemistry / FEBS 139, 259-265

6. Van Vliet, W. F. (1954) The enzymic oxidation of lignin. Biochimica et biophysica acta 15, 211-216

7. Palmer, N. A., Sattler, S. E., Saathoff, A. J., Funnell, D., Pedersen, J. F., and Sarath, G. (2008) Genetic background impacts soluble and cell wall-bound aromatics in brown midrib mutants of sorghum. Planta

229, 115-127

8. Chabannes, M., Barakate, A., Lapierre, C., Marita, J. M., Ralph, J., Pean, M., Danoun, S., Halpin, C.,

Grima-Pettenati, J., and Boudet, A. M. (2001) Strong decrease in lignin content without significant alteration of plant development is induced by simultaneous down-regulation of cinnamoyl CoA reductase

(CCR) and cinnamyl alcohol dehydrogenase (CAD) in tobacco plants. The Plant journal : for cell and molecular biology 28, 257-270

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9. Cotana, F., Cavalaglio, G., Gelosia, M., Coccia, V., Petrozzi, A., and Nicolini, A. (2014) Effect of double- step steam explosion pretreatment in bioethanol production from softwood. Appl Biochem Biotechnol 174,

156-167

10. Sattler, S. E., Funnell-Harris, D. L., and Pedersen, J. F. (2010) Efficacy of singular and stacked brown midrib 6 and 12 in the modification of lignocellulose and grain chemistry. J Agric Food Chem 58, 3611-

3616

11. Green, A. R., Lewis, K. M., Barr, J. T., Jones, J. P., Lu, F., Ralph, J., Vermerris, W., Sattler, S. E., and

Kang, C. (2014) Determination of the Structure and Catalytic Mechanism of Sorghum bicolor Caffeic Acid

O-Methyltransferase and the Structural Impact of Three brown midrib12 Mutations. Plant physiology 165,

1440-1456

12. Sattler, S. E., Saballos, A., Xin, Z., Funnell-Harris, D. L., Vermerris, W., and Pedersen, J. F. (2014)

Characterization of novel Sorghum brown midrib mutants from an EMS-mutagenized population. G3

(Bethesda) 4, 2115-2124

13. Jun, S. Y., Sattler, S. A., Cortez, G. S., Vermerris, W., Sattler, S. E., and Kang, C. (2018) Biochemical and Structural Analysis of Substrate Specificity of a Phenylalanine Ammonia-Lyase. Plant physiology 176,

1452-1468

14. Walker, A. M., Hayes, R. P., Youn, B., Vermerris, W., Sattler, S. E., and Kang, C. (2013) Elucidation of the Structure and Reaction Mechanism of Sorghum Hydroxycinnamoyltransferase and Its Structural

Relationship to Other Coenzyme A-Dependent Transferases and Synthases. Plant physiology 162, 640-651

15. Moural, T. W., Lewis, K. M., Barnaba, C., Zhu, F., Palmer, N. A., Sarath, G., Scully, E. D., Jones, J.

P., Sattler, S. E., and Kang, C. (2017) Characterization of Class III Peroxidases from Switchgrass. Plant physiology 173, 417-433

16. Walker, A. M., Sattler, S. A., Regner, M. R., Jones, J. P., Ralph, J., Vermerris, W., Sattler, S. E., and

Kang, C. (2016) Determination of the structure and catalytic mechanism of Sorghum bicolor caffeoyl-CoA

O-methyltransferase. Plant physiology

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17. Pettinati, I., Brem, J., McDonough, M. A., and Schofield, C. J. (2015) Crystal structure of human persulfide dioxygenase: structural basis of ethylmalonic encephalopathy. Hum Mol Genet 24, 2458-2469

18. Tiranti, V., Mineri, R., Viscomi, C., Tiveron, C., Rimoldi, M., and Zeviani, M. (2008) Characterization of the Ethe1 protein by cellular and animals models: Towards an understanding of its role in ethylmalonic encephalopathy. Neurology 70, A484-A484

19. Kabil, O., and Banerjee, R. (2012) Characterization of patient mutations in human persulfide dioxygenase (ETHE1) involved in H2S catabolism. The Journal of biological chemistry 287, 44561-44567

20. Sattler, S. A., Wang, X., Lewis, K. M., DeHan, P. J., Park, C. M., Xin, Y., Liu, H., Xian, M., Xun, L., and Kang, C. (2015) Characterizations of Two Bacterial Persulfide Dioxygenases of the Metallo-beta- lactamase Superfamily. The Journal of biological chemistry 290, 18914-18923

21. Di Meo, I., Lamperti, C., and Tiranti, V. (1993) Ethylmalonic Encephalopathy. in GeneReviews((R))

(Adam, M. P., Ardinger, H. H., Pagon, R. A., Wallace, S. E., Bean, L. J. H., Stephens, K., and Amemiya,

A. eds.), Seattle (WA). pp

22. Burlina, A., Zacchello, F., Dionisi-Vici, C., Bertini, E., Sabetta, G., Bennet, M. J., Hale, D. E., Schmidt-

Sommerfeld, E., and Rinaldo, P. (1991) New clinical phenotype of branched-chain acyl-CoA oxidation defect. Lancet 338, 1522-1523

23. Tiranti, V., Viscomi, C., Hildebrandt, T., Di Meo, I., Mineri, R., Tiveron, C., Levitt, M. D., Prelle, A.,

Fagiolari, G., Rimoldi, M., and Zeviani, M. (2009) Loss of ETHE1, a mitochondrial dioxygenase, causes fatal sulfide toxicity in ethylmalonic encephalopathy. Nature medicine 15, 200-205

24. Tiranti, V., D'Adamo, P., Briem, E., Ferrari, G., Mineri, R., Lamantea, E., Mandel, H., Balestri, P.,

Garcia-Silva, M. T., Vollmer, B., Rinaldo, P., Hahn, S. H., Leonard, J., Rahman, S., Dionisi-Vici, C.,

Garavaglia, B., Gasparini, P., and Zeviani, M. (2004) Ethylmalonic encephalopathy is caused by mutations in ETHE1, a gene encoding a mitochondrial matrix protein. American journal of human genetics 74, 239-

252

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25. Murata, K., Inoue, Y., Rhee, H., and Kimura, A. (1989) 2-Oxoaldehyde metabolism in microorganisms.

Can J Microbiol 35, 423-431

26. Thornalley, P. J. (2003) Glyoxalase I--structure, function and a critical role in the enzymatic defence against glycation. Biochem Soc Trans 31, 1343-1348

27. Vander Jagt, D. L. (1993) Glyoxalase II: molecular characteristics, kinetics and mechanism. Biochem

Soc Trans 21, 522-527

28. Campos-Bermudez, V. A., Leite, N. R., Krog, R., Costa-Filho, A. J., Soncini, F. C., Oliva, G., and Vila,

A. J. (2007) Biochemical and structural characterization of Salmonella typhimurium glyoxalase II: new insights into metal ion selectivity. Biochemistry 46, 11069-11079

29. Liu, H., Xin, Y., and Xun, L. (2014) Distribution, diversity, and activities of sulfur dioxygenases in heterotrophic bacteria. Applied and environmental microbiology 80, 1799-1806

30. Cameron, A. D., Ridderstrom, M., Olin, B., and Mannervik, B. (1999) Crystal structure of human glyoxalase II and its complex with a glutathione thiolester substrate analogue. Struct Fold Des 7, 1067-

1078

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CHAPTER TWO

CHARACTERIZATIONS OF TWO BACTERIAL PERSULFIDE DIOXYGENASES OF THE

METALLO-Β-LACTAMASE SUPERFAMILY

Sattler SA, Wang X, Lewis KM, DeHan PJ, Park CM, Xin Y, Liu H, Xian M, Xun L, Kang C. (2015). Characterizations of Two Bacterial Persulfide Dioxygenases of the Metallo-β-lactamase Superfamily. Journal of Biological Chemistry, 290(31), 18914-23.

All text, tables, and figures in this chapter are directly from the published manuscript.

2.1 Contributions

C.K., L.X., and M.X. conceived and coordinated the study. C.K., L.X., S.A.S., K.M.L.,

M.X., and C.P. wrote the paper. X.W., Y.X., and H.L. generated clones of PDO genes. S.A.S.,

X.W., and P.J.D. expressed, purified, and crystallized the proteins. C.P. and M.X. performed the synthesis of GSSO3 and GSSCH3, which were used for ITC and protein crystal soaking. C.P. performed NMR to assess purity of the synthetic compounds. S.A.S. and K.M.L. obtained x-ray diffraction data for PpPDO2 and MxPDO1b. S.A.S. and P.J.D. performed ITC and multiangle light scattering. S.A.S. solved the molecular structures of PpPDO2 and MxPDO1b. All authors contributed to experimental design, reviewed the results, and approved the final version of the manuscript.

2.2 Abstract

Persulfide dioxygenases (PDOs), also known as sulfur dioxygenases (SDOs), oxidize glutathione persulfide (GSSH) to sulfite and GSH. PDOs belong to the metallo-β-lactamase superfamily and play critical roles in animals, plants, and microorganisms, including sulfide detoxification. The structures of two PDOs from human and Arabidopsis thaliana have been reported; however, little is known about the substrate binding and catalytic mechanism. The crystal structures of two bacterial PDOs from Pseudomonas putida and Myxococcus xanthus were

14

determined at 1.5- and 2.5-Å resolution, respectively. The structures of both PDOs were homodimers, and their metal centers and β-lactamase folds were superimposable with those of related enzymes, especially the glyoxalases II. The PDOs share similar Fe(II) coordination and a secondary coordination sphere-based hydrogen bond network that is absent in glyoxalases II, in which the corresponding residues are involved instead in coordinating a second metal ion. The crystal structure of the complex between the Pseudomonas PDO and GSH also reveals the similarity of substrate binding between it and glyoxalases II. Further analysis implicates an identical mode of substrate binding by known PDOs. Thus, the data not only reveal the differences in metal binding and coordination between the dioxygenases and the hydrolytic enzymes in the metallo-β-lactamase superfamily, but also provide detailed information on substrate binding by

PDOs.

2.3 Introduction

Accumulating evidence indicates that hydrogen sulfide (H2S) plays significant roles as a signaling molecule in animals (1, 2). The cellular H2S concentration is maintained by equilibrium between its formation and oxidation. It is produced from sulfur-containing amino acids by cystathionine β-synthase, cystathionine γ-lyase, and mercaptopyruvate sulfur transferase (3). In eukaryotes, H2S is oxidized in the mitochondrion. First, sulfide:quinone oxidoreductase oxidizes it to sulfane sulfur that is likely present as a cysteinyl persulfide intermediate within the active site

(4). Sulfide:quinone oxidoreductase passes the sulfur to a sulfane sulfur acceptor. Although sulfite is an effective acceptor, GSH is more likely to be the physiological acceptor to produce glutathione persulfide (GSSH) (5). Then, persulfide dioxygenase (PDO), which is also known as sulfur dioxygenase (EC 1.13.11.18), oxidizes GSSH to sulfite and GSH (6). Furthermore, a rhodanese or sulfurtransferase is involved in the metabolism (5, 7, 8). Mutations in the human PDO gene ethe1

15

are the cause of the recessive hereditary disease ethylmalonic encephalopathy, which can lead to unusually high excretion of short-chain carboxylic acids in the urine, brain defects, and early death

(9). PDO activity is also important in plants; its gene inactivation in Arabidopsis thaliana has been reported to produce defects in seed development and embryonic arrest by the early heart stage

(10). Thus, H2S oxidation plays a major physiological role in both plants and animals. The activity of PDO was initially discovered in bacterial cell extracts, and GSSH was identified as a substrate

(11). The human PDO (referred to as hPDO in this report, and is known as hETHE1) is the first characterized PDO (10, 12). Sequence analysis reveals that it belongs to the metallo-β-lactamase superfamily, which consists of mainly hydrolytic enzymes (13). Proteins in the family share the same structural fold of two stacked β-sheets surrounded by α-helices. The metal-binding residues are located at one edge of the β-sheets, and substrate binding is from the loops and α-helices surrounding the metal center. These proteins have six or seven conserved amino acid residues for binding one or two metal ions at the metal center. Because the other members of the superfamily are mainly hydrolytic enzymes, their reaction mechanisms cannot apply to explain the dioxygenase activity of PDOs. Using the hPDO sequence, we have identified a wide distribution of PDO genes in sequenced bacterial genomes and have recently characterized 10 bacterial PDOs (14). On the basis of sequence analysis, we have proposed three subclasses of PDO: (i) ETHE1, which is present in animals, plants, and bacteria, (ii) persulfide dioxygenase A (PdoA), also known as sulfur dioxygenase A (SdoA), which is common in Proteobacteria, and (iii) Blh, which is an acronym for

“β-lactamase-like hydrolase” (14). To be consistent with scientific names, we rename the bacterial

ETHE1 type as PDO type I, and the PdoA type as PDO type II throughout this report. Our new structural data also support this classification. Apo-form crystal structures of hPDO and A. thaliana

PDO (referred to as AtPDO in this report, and is also known as AtETHE1) have been determined

16

(15, 16). In this report, we present structural and biochemical analyses for a PDO from

Pseudomonas putida (PpPDO2, also known as PpSdoA) and for a PDO from Myxococcus xanthus

(MxPDO1b, also known as MxETHE1b because M. xanthus possesses three type I PDOs). The crystal structure of PpPDO2 with GSH in its binding pocket permits identification of the amino acid residues involved in substrate binding. In addition, structural comparison of PpPDO2 with

MxPDO1b reveals that there are differences in the GS-moiety binding sites between them.

Furthermore, the change of metal binding in the PDOs in comparison with other members of the

MBL superfamily is discussed for the evolution of a dioxygenase from a hydrolase.

2.4 Materials and Methods

Chemicals and Enzymes

Chemicals were obtained from Sigma or Fisher Scientific. Crystallization screens were obtained from Hampton Research and Qiagen.

Cloning and Enzyme Purification

Genes encoding PpPDO2 from P. putida (ABQ76243) and MxPDO1b from M. xanthus

(WP_011554322, ex. YP_632494) were cloned into pET-30 Ek/LIC with Escherichia coli

BL21(DE3) as the host (14). For expression and purification of C terminally His-tagged PpPDO2 or MxPDO1b, cultures were grown at 37 °C in LB broth containing 30 μg/mL of kanamycin. The cultures were allowed to reach an A600 of 0.6 prior to inducing protein expression, which was done by adding 0.5 mM isopropyl -D-thiogalactopyranoside to the media and incubating for 22 h at

20 °C. Cells were harvested by centrifugation at 5000 x g, frozen, and then suspended in 50 mM

Tris buffer, pH 8.0, supplemented with 300 mM NaCl and 20 mM imidazole. Cells were lysed by sonication and lysates were cleared by centrifugation at 15,000 x g. The supernatant was stirred into nickel-nitrilotriacetic acid-agarose resin (Qiagen), the column was washed with 2 volumes of

17

the lysis buffer, and recombinant enzyme was eluted with buffer containing 50 mM Tris, pH 8.0,

300 mM NaCl, and 250 mM imidazole. The eluted sample was concentrated and exchanged into

20 mM Tris, pH 7.5, applied to a 6-mL Resource Q column (GE Healthcare), and the enzyme was eluted with a 50 mM stepwise NaCl gradient in the same buffer. PpPDO2 or MxPDO1b-containing fractions, which eluted at 200 and 100 mM NaCl, respectively, were pooled, buffer-exchanged, and concentrated into the appropriate buffer for crystallization or biochemical experiments. To ensure higher occupancy of ferrous iron in the active sites, all enzyme preparations were incubated with ferrous ammonium sulfate and ascorbic acid on ice for 2 h prior to use, and at concentrations equal to those of the enzymes. Purity was monitored for all protein preparations by SDS-PAGE and protein concentrations were determined with the method of Bradford, using BSA as a standard.

Protein Crystallization and Structure Determination

Crystals of PpPDO2 and MxPDO1b were grown using the hanging-drop, vapor-diffusion method. For PpPDO2 crystallization, purified protein at 30 mg/mL in 20 mM Tris, pH 8.0, was mixed with an equal volume of reservoir solution and equilibrated against the same solution at

4 °C. The reservoir solution was 100 mM HEPES, pH 7.5, 200 mM ammonium acetate, and 25%

(w/v) PEG 3350. Crystals of this enzyme typically appeared within 3 days. To obtain structural data for PpPDO2 in complex with GSH, crystals were soaked in 5 mM GSH for 1 h prior to harvest.

Adequate cryoprotection was achieved by passing crystals through a small drop of storage buffer/mother liquor mixture (50% of each component by volume) that was brought to 18% glycerol. For MxPDO1b crystallization, purified protein at 11 mg/mL in 20 mM MOPS, pH 7.1, was mixed with an equal volume of reservoir solution and equilibrated against the same solution at 4 °C. The reservoir solution was 200 mM calcium chloride, 100 mM HEPES, pH 7.5, 15% (w/v)

PEG 400, and 15% glycerol (w/v). Because the mother liquor was a sufficient cryoprotectant, no

18

additional glycerol was required to prevent freezing. The space groups of PpPDO2 and MxPDO1b were P43 and P61, respectively; the asymmetric units of the unit cells contained both molecules of the respective homodimers. Diffraction data were collected up to 1.5-Å resolution for PpPDO2 in complex with GSH and 2.5 Å for the apo-forms of both MxPDO1b and PpPDO2 at the Berkeley

Advanced Light Source (ALS, beamline 8.2.1). The diffraction data were processed with the

HKL2000 package (17). The statistics for the diffraction data are listed in Table 1. Initial phasing of apo-form PpPDO2 diffraction data were conducted by molecular replacement with the PDB coordinates of model 4EFZ using PHENIX Phaser (18). MxPDO1b initial phasing was conducted by molecular replacement as well, using the atomic coordinates of the unpublished structure for a

PDO1 protein from Cupriavidus necator. Iterative model building and refinement was conducted using the programs COOT (19) and PHENIX. The coordinate and diffraction data have been deposited in the Protein Data Bank.

Multiangle Light Scattering and Isothermal Titration Calorimetry (ITC)

To determine a predominant oligomeric state, multiangle laser light scattering was performed for PpPDO2 as previously described (20). Briefly, 200 μg of PpPDO2 were loaded onto a Yarra 3-μm SEC-200 column (Phenomenex) and eluted isocratically at 0.5 mL min-1. The elution buffer for PpPDO2 was 20 mM MOPS, pH 6.8, supplemented with 100 mM NaCl. The eluate was successively passed through a UV detector (Gilson), an Optilab DSP interferometric refractometer

(Wyatt Technology), and a Dawn EOS laser light scattering detector (Wyatt Technology). Data analysis was performed using the Zimm fitting method with software (ASTRA) provided by the instrument manufacturer.

Isothermal calorimetry titrations were executed for PpPDO2 and several substrate and product analogs in an ITC200 instrument (Malvern Instruments). The protein was prepared by extensive

19

buffer exchange into the titration buffer, which consisted of 20 mM MOPS, pH 7.1. The concentration of protein in the calorimetric titration cell was diluted to 200 µM. All titrations were performed at 25 °C with a stirring speed of 750 rpm and 27 injections (1.4 µl each). All ligands were brought to a concentration of 2 mM in the titration buffer and injected into the protein solution, and the heats of binding were recorded. Ligands were also titrated against buffer alone to account for the heats of dilution. Ligand concentrations were adjusted to obtain significant heats of binding, and the time intervals between injections were adjusted to ensure proper baseline equilibration. All samples were degassed prior to titration.

Preparation of GSSCH3

To a solution of GSH (11 mg, 0.036 mmol) in 2 mL of sodium phosphate buffer (50 mM, pH 7.4), 10 eq of methyl methanethiosulfonate in 1 M N,N-dimethylformamide were added. The resulting solution was vigorously stirred for 1.5 h at room temperature. The solution was frozen on a dry ice/acetone bath and then lyophilized. The residue was washed and filtered with cold diethyl ether (3 X 5 mL) to remove excess methyl methanethiosulfonate. The white solid was collected by filtration to afford a product in 90% yield and characterized by 1H NMR, which was

1 compared with a previous report (21). H NMR (300 MHz, D2O) δ 4.80 (m, CHCH2S, 1H), 3.97

(s, CH2CO2, 2H), 3.82 (t, J = 6.3 Hz, CHNH2, 1H), 3.25 (dd, J = 14.6, 4.9 Hz, CH2S, 1H), 3.02

(m, CH2S, 1H), 2.53 (q, J = 6.9 Hz, CH2CH2CO, 2H), 2.43 (s, SSCH3, 3H), 2.17 (q, J = 7.6 Hz,

CH2CH2CO, 2H).

- Preparation of GSSO3

The compound was synthesized following a reported procedure (22). Freshly prepared 2 mL of Na2SO3 solution (200 mM: sodium phosphate buffer, pH 7.1) was directly added to freshly prepared 2 mL of GSNO solution (80 mM: sodium phosphate buffer, pH 7.1), and the resulting

20

- 1 mixture was stirred for 1 h at room temperature. The formation of GSSO3 was monitored by H

- 1 NMR and the spectral characteristics of GSSO3 were compared with previously reported H NMR

1 data (22). H NMR (300 MHz, D2O, pD 7.4, 2 M KCl, 80 mM sodium phosphate, 100 M EDTA)

δ 5.01 (dd, J = 8.9, 4.6 Hz, CHCH2S, 1H), 3.97–3.89 (overlapped-m, CHNH2, CH2CO2, 3H), 3.76

(dd, J = 14.6, 4.7 Hz, CH2S, 1H), 3.57 (dd, J = 14.7, 8.9 Hz, CH2S, 1H), 2.72–2.59 (m, CH2CH2CO,

2H), 2.32 (m, CH2CH2CO, 2H).

2.5 Results

Global Structure

Purified recombinant PpPDO2 andMxPDO1b were crystallized in the P43 and P61 space groups and solved at 1.5- and 2.5-Å resolution, respectively. The structures of both PpPDO2 and

MxPDO1b molecules consisted of two similarly sized, six-stranded central β-sheets surrounded by α-helices (Fig. 1, A and B), a structural motif strongly conserved among the metallo-β-lactamase superfamily. Although there were several insertions and deletions between

PpPDO2 and MxPDO1b, which were primarily located in loops and N/C termini, the Cα positions of the two molecules were superimposable with a r.m.s. deviation value of 1.18 Å (Fig. 1C). The

C-terminal eight residues of MxPDO1b were not visible in the final 2Fo - Fc electron density map, indicating their disordered nature.

Throughout this report, the residue numbers for PpPDO2 were used, with those of

MxPDO1b in parentheses unless stated otherwise. In both PpPDO2 and MxPDO1b structures, a single ferrous iron was coordinated by three residues occupying one face of the coordination sphere, including His74 (His57), His149 (His112), and Asp170 (Asp129) (Fig. 2A). This is a typical coordination pattern among mononuclear non-heme iron (II) oxygenases and is known as the

2-His-1-carboxylate facial triad (13, 23). The locations and orientations of participating residues

21

observed in both PpPDO2 and MxPDO1b were identical and all three residues were from the same edge of the central β-sheets. The nitrogen atoms in Fe(II)-coordinating imidazole rings of His74

(His57) and His149 (His112) were within a hydrogen bond distance from the neighboring residues,

Arg181 (Arg138) and Thr73 (Thr56), respectively (Fig. 2A). Thus both His residues could not only fix the orientation but also tune the redox potential of the Fe(II). Three water molecules (W1, W2, and W3) were noticed occupying the opposite face of the Fe(II) coordination sphere (Fig. 2A). W3 was hydrogen bonded to the side chains of His76 (His59), Asp78 (Asp61), and His212 (His170), closely mimicking the 2-His-1-carboxylate facial triad. W2 was also hydrogen-bonded to the backbone of

Ala77 (Ala60). W1, which was displaced by GSH upon its binding, was connected only to the bulk solvent.

Oligomeric Structure of PpPDO2 and MxPDO1b

The asymmetric units of both PpPDO2 and MxPDO1b were composed of two tightly associated monomers in a non-crystallographic, 2-fold manner (Fig. 1, A and B). This dimeric status of PpPDO2 is maintained in solution and was verified by multiangle laser light scattering experiments (Fig. 3). The dimer interface had a symmetrically oriented inter-subunit β-sheet between two C-terminal peptides (Val255–Leu262), resulting in a large area of hydrophobic interaction. The observed dimer interface contributed to stabilizing one face of the substrate binding pocket, indicating the dimer as a functional unit (Fig. 2A).

GSH Complex

The Fo - Fc map of GSH-soaked PpPDO2 crystals showed the corresponding electron density for the bound molecules located on the other side of the facial triad open to the bulk solvent (Fig.

2B). The r.m.s. deviation value for Cα atoms between apo-form and GSH complex PpPDO2 was

0.3 Å, indicating little change upon GSH binding (Fig. 2C). One of the metal-coordinating water

22

molecules (W1) was replaced by GSH and the distance between the sulfur atom of GSH and Fe(II) was 2.5 Å (Fig. 2, A and B). A deep binding pocket was established by Asp78, His149, Asp170,

Arg181, Tyr214, Arg250, Arg253, Val261, and Leu262. Noticeably, the apo-form structure of PpPDO2 had a few water molecules at equivalent positions of the GSH ligand (Fig. 2A). The backbone of

Arg181 and the side chains of Arg181, Tyr214, Arg250, and Arg253 were in direct interaction with GSH

(Fig. 2B). Specifically, a glycinyl carboxyl oxygen of GSH was electrostatically interacting with the guanidinium groups of the Arg250 and Arg253 side chains, and the two backbone carbonyl oxygen atoms of GSH were within hydrogen bond distance from the backbone nitrogen and guanidinium group of the Arg181 side chain. In addition, the cysteinyl amide hydrogen of GSH established a hydrogen bond with the phenolic hydroxyl group of Tyr214. The glutamyl carboxyl group of GSH displayed both direct and indirect interactions through a water molecule-mediated hydrogen bond with neighboring residues’ backbone atoms. Contrary to the result of our efforts

- with GSH, our numerous attempts to diffuse GSSO3 into the apo-form PpPDO2 crystals were not successful, resulting in non-diffracting crystals or diffraction data of very low resolution.

ITC Data

We used ITC to confirm the differential binding affinities among the PpPDO2 product and

- two of its analogs. A small amount of heat was released when GSH or GSSO3 was titrated into

PpPDO2-containing solutions (ΔH = -0.3 and 2.2 kcal mol-1, respectively) (Table 2, Fig. 4).

Further analysis of the ITC data revealed favorable entropic contributions for both GSH and

- -1 -1 GSSO3 binding (ΔS = 25.5 and 15.1 cal mol K ), probably indicating that several solvent molecules were displaced from the pocket upon binding of either compound. Supporting this, there were several water molecules in the substrate-binding pocket of the apo-form crystal structure of

23

- PpPDO2. The calculated Kd values for GSH and GSSO3 (Table 2) were 1.6 and 12 µM, respectively. PpPDO2 did not appear to have affinity for GSSCH3 (Fig. 4).

2.6 Discussion

To establish the proper classification of bacterial PDOs and to identify any unique signatures, comparisons with available structures in the Protein Data Bank (PDB) were executed using Dali (24) and BLAST (25) searches. The superimposed three-dimensional structures of

PpPDO2, MxPDO1b, and the related enzymes displayed that most of the regions with high sequence similarity were located around the residues of the facial triad. Significantly, the

Fe(II)-coordinating residues and critical residues in the second coordination sphere were completely conserved among those closely related PDOs, including AtPDO and hPDO. Those residues and their physical arrangements may be conserved to maintain the orientation of

Fe(II)-coordinating residues and water molecules, as well as for facilitating rearrangements of electrons and protons of substrates.

As shown by our Dali and BLAST searches, both PpPDO2 and MxPDO1b share a high level of sequence identity with PDOs and glyoxalases II (Fig. 5). In addition, their β-lactamase folds are superimposable with those of PDOs and glyoxalases II from various species. Their metal-binding centers also closely resemble that of the first metal-binding site of the glyoxalases

II (Fig. 6), which use di-metallic reaction centers to hydrolyze S-lactoylglutathione.

The secondary coordination sphere with hydrogen bond networks observed in both

PpPDO2 and MxPDO1b are essentially the same among the PDOs (Fig. 6A, B). This similarity, however, does not extend to the glyoxalases II, because the three-dimensional structures of those enzymes do not show any secondary coordination sphere with hydrogen bond networks. The corresponding residues in the glyoxalases II are instead involved in coordinating a second metal

24

ion (Fig. 6, C and D). Given the fact that most members of the metallo-β-lactamase superfamily are hydrolytic enzymes with binuclear metal centers (26), PDOs likely evolved from a hydrolytic enzyme that has two coordinated metal ions. Evolution has led to loss of affinity for a second metal and gain of the coordination for water molecules together with a hydrogen bond network in their second coordination sphere, which is likely critical to Fe(II) positioning and catalysis, as we proposed in the case of 2,6-dichloro-p-hydroquinone 1,2-dioxygenase (27). Consequently, the metal-binding center is highly conserved among PDOs.

Both primary and tertiary structures of MxPDO1b closely resemble hPDO and AtPDO in regard to the GS-moiety binding pocket and to Fe(II) coordination, thus supporting our classification scheme for type I and type II PDOs (14). On the contrary, the structure of PpPDO2 is more similar to those of the glyoxalases II than to hPDO and AtPDO in terms of its three specific arginine residues (Arg181, Arg250, and Arg253) electrostatically interacting with GSH (Figs. 2B and

7). The structure of PpPDO2 in complex with GSH showed that the two interior carbonyl oxygens of GSH were anchored by Arg181, and the glycinyl carboxyl group of GSH electrostatically interacted with two other arginine residues (Arg250 and Arg253) (Fig. 2B).

Significantly, both location and orientation of the bound GSH in PpPDO2 were similar to those in the crystal structure of human glyoxalase II (PDB code 1QH5 (28)) (Fig. 7). Furthermore, the positions of the three specified arginine residues in PpPDO2 were conserved and superimposable when the three-dimensional structures of glyoxalases II and PpPDO2 were aligned. However, due to large insertions or deletions just before those basic residues among PDOs and glyoxalases II, all alignment programs used failed to capture that structural/functional conservation, which was only possible to grasp with the aid of tertiary structure alignment. In type

I PDOs, including hPDO, AtPDO, and MxPDO1b, there is a single basic residue (Arg214 of hPDO

25

or Arg189 of MxPDO1b) located in the set of conserved residues NPR(L/V), suggesting the possibility of convergent evolution to accommodate the binding of GSSH. As shown in Fig. 5, many sequence alignment programs aligned Arg253 of PpPDO2 with Arg221 of MxPDO1b and the

Arg246 of hPDO, which turned out meaningless. The latter two residues are located in a different

α-helix that is irrelevant to substrate binding, and their guanidinium groups instead point to the bulk solvent. Significantly, the His-rich region of the glyoxalases II includes a one-turn α-helix, and the 1st, 3rd, 5th, and 6th residues in its HHHXDH motif are involved in metal coordination

(Figs. 5 and 6, C and D). On the contrary, in the corresponding region in PDOs, only the first histidine of HXHXDH is involved in Fe(II) coordination (Figs. 5 and 7, A and B). The remaining residues are involved in the hydrogen bond network, with the Fe(II)-coordinating water molecules and neighboring residues establishing the core of the secondary coordination sphere. Therefore,

HHHXDH versus HXHXDH could serve as a signature sequence for distinguishing between glyoxalases II and PDOs.

Nonsense mutations of Gln12 and Gln63 and missense mutations of Tyr38, Leu55, Thr136,

Thr152, Cys161, Arg163, Thr164, Asp165, Leu185, and Asp196 of hPDO have been implicated in ethylmalonic encephalopathy (6, 9, 30). These residues are conserved among PDOs (Fig. 5). The structure of PpPDO2 in complex with GSH suggests that the backbone amide and side chain of

Arg163 in hPDO (Arg181 in PpPDO2) jointly anchor the flanking carbonyl oxygens of GSH

(Fig. 2B). The residues near Arg163 of hPDO showed similarity with those of the glyoxalases II

(Fig. 5). Tyr38 of hPDO (Fig. 1C), which is conserved among PDOs, establishes a side chain-mediated hydrogen bond to the backbone carbonyl oxygen of the residue located at the

β-strand of the opposite side in the central β-sheet. Consequently, the observed Y38C mutation can impact the overall folding of hPDO and proper configuration of the Fe(II) site. Thr136, which

26

is also conserved, is located next to the Fe(II)-coordinating histidine residue, forming a β-bulge structure through its side chain hydrogen bond. Thus, the T136A/T136G mutations will cause the loss of proper geometry for Fe(II) coordination. Leu185, which is conserved in PDOs and glyoxalases II, interacts with neighboring hydrophobic residues. Alteration to a charged residue such as arginine (L185R) will greatly reduce the stability of enzymes or result in misfolding. It may also be possible that conservation of hydrophobic residues in this region is necessary to shield iron-bound oxygen from bulk solvent, thus preventing formation of reactive oxygen species in non-productive side reactions.

Active Site of PpPDO2 and Plausible Reaction Mechanism

Both apo-form and GSH binary complex structures of PpPDO2 are suggestive of a likely mechanism. The resting state of PpPDO2 in its Fe(II) form is coordinated by two histidines, an aspartate, and three water molecules. The incoming substrate, GSSH, replaces the solvent-exposed water molecule (W1 in Fig. 2A) coordinated to Fe(II) and liberates several water molecules from the GSSH-binding pocket (Fig. 2, A and B). Upon docking into the binding pocket, the carboxyl groups of GSSH are anchored by hydrogen bonds and salt bridges to the backbones and side chains of the residues that constitute the pocket. The pKa of the sulfhydryl group in GSSH has been estimated to be 7.2, which is lower than that of GSH (31). Thus, it is likely deprotonated or being deprotonated as it approaches Fe(II), which is facilitated by the Lewis acid nature of the metal, the entropic effect of the departing water molecules, and the lower dielectric constant upon departure of those waters. In addition, the Fe(II)-coordinating water molecule, W3 (Fig. 2A), could act as a proton relay to facilitate the deprotonation of GSSH. Association of substrate into the active site should change the electronic properties of Fe(II) and the hydrogen bond network, which triggers replacement of a water molecule in the primary coordination sphere by O2. It is tempting to

27

184 speculate that a coordinating water (W2), which is proximal to Phe , is replaced by O2 (Fig. 2A).

Phe184 could act as a gate, because two alternate conformations of its side chain were detected in the electron density maps of both apo-form PpPDO2 and the enzyme in complex with GSH. The two conformers lend plausibility to a mechanism for transient exposure of the hydrophobic environment to the bulk solvent area.

Conclusion

Our results for PpPDO2 and MxPDO1b indicate a close relationship among PDOs, especially around the Fe(II) binding site. Although the shape and location of their binding sites for the GS moiety are superimposable between the type I and type II of PDOs, the key amino acid residues for substrate binding originated from different parts of the proteins, supporting the idea for grouping of them into individual subclasses. Our structural characterization also helps to gain a comprehensive picture of their binding and catalytic mechanisms that is likely conserved among

PDOs.

2.7 Acknowledgement

Original research was supported by the NSF (MCB 1021148, DBI 0959778) and the M.J.

Murdock Charitable Trust. X. Wang was supported by China Scholarship Council (CSC, No.

201208370133) during a visit to Washington State University.

28

Crystallographic data for the structures MxPDO1b PpPDO2 for and data Crystallographic

.

1

2.

Table

29

Table 2.2. Thermodynamic parameters for binding substrate analogs by PpPDO2

30

31

Figure 2.1: Ribbon diagrams representing the oligomeric and superimposed structures of

PDOs. A, PpPDO2 and B, MxPDO1b are represented in their dimeric states. The orange spheres in both enzymes represent Fe(II). C, the superimposed structures of MxPDO1b (tan),

PpPDO2 (light blue), hPDO (blue), and AtPDO (purple). The corresponding N and C termini are marked N and C, respectively. Residues corresponding to ethylmalonic encephalopathy-causing mutations are highlighted in red. The residues are numbered according to the hPDO sequence, with equivalent residues from PpPDO2 in parenthesis. This figure was generated using Chimera (UCSF) (32).

32

33

Figure 2.2: GSH complex and ligand-free forms of PpPDO2. A, the stereo image of the Fe(II) coordinated by three water molecules in the apo-form of PpPDO2. The Fe(II)-coordinated water molecule W1 represents the water that is displaced upon binding of GSH, whereas water

W2 is in the proposed site of O2 binding. The third water, W3, is engaged in hydrogen bonding with neighboring residues and may function as a proton relay. Notably, the lipophilic region of the Lys44 side chain in chain B interfaces with neighboring hydrophobic residues of chain A, likely contributing to binding pocket stability. B, the difference Fourier map clearly shows

PpPDO2 in complex with GSH. Surrounding residues important for binding and catalysis are displayed and their residue numbers are indicated. The water molecules are shown as red spheres. C, the ribbon diagrams for Cα atoms of apo-form PpPDO2 (tan) and the same enzyme in complex with GSH (blue) are superimposable with a r.m.s. deviation of 0.3 Å. This figure was generated using Chimera (UCSF) (32).

34

Figure 2.3: The oligomeric state of PpPDO2 in solution. The elution profile for PpPDO2 was monitored with multiangle laser light scattering and is shown as absorbance (left y axis) and molecular weight (right y axis) versus elution volume (mL). The solid line represents changes in absorption at 280 nm. The thick black cluster in the middle of the peak indicates the calculated molecular mass of 67.4 kDa from the light scattering, illustrating the dimeric nature of PpPDO2.

35

36

Figure 2.4: Measurements of PpPDO2 binding for the product and substrate analogs via

− ITC. The trends of heat released during serial injections of GSH (▴), GSSO3 (●), and

GSSCH3 (■) into a solution of PpPDO2 are presented. The data reveal low micromolar affinity

− for GSH and GSSO3 , but no apparent binding of GSSCH3.

37

38

Figure 2.5: Multiple sequence alignment of four PDOs and three glyoxalases II. Included in the alignment are PDOs from P. putida (PpPDO2), M. xanthus (MxPDO1b), human (hPDO), and Arabidopsis (AtPDO), as well as glyoxalase II enzymes from human (hGloB), A. thaliana

(AtGloB), and Salmonella typhimurium (StGloB). The α-helices and β-strands are highlighted in red and yellow, respectively. Functionally significant residues are bolded, and residues involved in binding metals are underlined. Single conserved residues are marked with an asterisk, whereas conserved motifs are enclosed by a box. Multiple sequence alignment was performed with CLUSTALW2 using a BLOSUM weighting matrix. Secondary structure for each enzyme was calculated from PDB coordinates using DSSP (version 2.0)(29).

39

40

Figure 2.6: Metal-binding and secondary coordination sphere architectures of PDOs and glyoxalases II. The two structures shown at the top panel correspond to PpPDO2 (A) and

MxPDO1b (B). At the bottom panel are glyoxalase II enzymes from human (C) and S. typhimurium (D). The extensive hydrogen bond network that appears to be characteristic of PDOs is substituted with a conserved, one-turn α-helix (resides 56–60 of human and 55–59 of S. typhimurium) in the glyoxalases II that is involved in coordinating the second metal.

41

Figure 2.7: Active site representations of PpPDO2 and human glyoxalase II in complex with

GSH. A, the binding pocket of human glyoxalase II (Protein Data Bank code 1QH5) with a bound GSH molecule is shown. The two gray spheres represent the catalytic Zn(II) ions

(28). B, for comparison, the GSH-binding modes and participating residues of PpPDO2

(green) and human glyoxalase II (gray) were superimposed. Residue labels that correspond to residues of the glyoxalase binding pocket are indicated in parentheses. Bond lengths are written near their respective bond representations. This figure was generated using PyMOL

Molecular Graphics System, version 1.7.0.3 (Schrödinger, LLC).

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2.8 References

1. Coletta, C., Papapetropoulos, A., Erdelyi, K., Olah, G., Módis, K., Panopoulos, P.,

Asimakopoulou, A., Gerö, D., Sharina, I., Martin, E., and Szabo, C. (2012) Hydrogen sulfide and nitric oxide are mutually dependent in the regulation of angiogenesis and endothelium-dependent vasorelaxation. Proc. Natl. Acad. Sci. U.S.A. 109, 9161–9166

2. Kolluru, G. K., Shen, X., and Kevil, C. G. (2013) A tale of two gases: NO and H2S, foes or friends for life? Redox Biol. 1, 313–318

3. Singh, S., and Banerjee, R. (2011) PLP-dependent H2S biogenesis. Biochim. Biophys. Acta

1814, 1518–1527

4. Jackson, M. R., Melideo, S. L., and Jorns, M. S. (2012) Human sulfide: quinone oxidoreductase catalyzes the first step in hydrogen sulfide metabolism and produces a sulfane sulfur metabolite.

Biochemistry 51, 6804–6815

5. Libiad, M., Yadav, P. K., Vitvitsky, V., Martinov, M., and Banerjee, R. (2014) Organization of the human mitochondrial hydrogen sulfide oxidation pathway. J. Biol. Chem. 289, 30901–30910

6. Kabil, O., and Banerjee, R. (2012) Characterization of patient mutations in human persulfide dioxygenase (ETHE1) involved in H2S catabolism. J. Biol. Chem. 287, 44561–44567

7. Melideo, S. L., Jackson, M. R., and Jorns, M. S. (2014) Biosynthesis of a central intermediate in hydrogen sulfide metabolism by a novel human sulfurtransferase and its yeast ortholog.

Biochemistry 53, 4739–4753

8. Hildebrandt, T. M., and Grieshaber, M. K. (2008) Three enzymatic activities catalyze the oxidation of sulfide to thiosulfate in mammalian and invertebrate mitochondria. FEBS J. 275,

3352–3361

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9. Tiranti, V., Briem, E., Lamantea, E., Mineri, R., Papaleo, E., De Gioia, L., Forlani, F., Rinaldo,

P., Dickson, P., Abu-Libdeh, B., Cindro-Heberle, L., Owaidha, M., Jack, R. M., Christensen, E.,

Burlina, A., and Zeviani, M. (2006) ETHE1 mutations are specific to ethylmalonic encephalopathy. J. Med. Genet. 43, 340–346

10. Holdorf, M. M., Owen, H. A., Lieber, S. R., Yuan, L., Adams, N., DabneySmith, C., and

Makaroff, C. A. (2012) Arabidopsis ETHE1 encodes a sulfur dioxygenase that is essential for embryo and endosperm development. Plant Physiol. 160, 226–236

11. Rohwerder, T., and Sand, W. (2003) The sulfane sulfur of persulfides is the actual substrate of the sulfur-oxidizing enzymes from Acidithiobacillus and Acidiphilium spp. Microbiology 149,

1699–1710

12. Tiranti, V., Viscomi, C., Hildebrandt, T., Di Meo, I., Mineri, R., Tiveron, C., Levitt, M. D.,

Prelle, A., Fagiolari, G., Rimoldi, M., and Zeviani, M. (2009) Loss of ETHE1, a mitochondrial dioxygenase, causes fatal sulfide toxicity in ethylmalonic encephalopathy. Nat. Med. 15, 200–205

13. Bugg, T. (2001) Oxygenases: mechanisms and structural motifs for O2 activation. Curr. Opin.

Chem. Biol. 5, 550–555

14. Liu, H., Xin, Y., and Xun, L. (2014) Distribution, diversity, and activities of sulfur dioxygenases in heterotrophic bacteria. Appl. Environ. Microbiol. 80, 1799–1806

15. Pettinati, I., Brem, J., McDonough, M. A., and Schofield, C. J. (2015) Crystal structure of human persulfide dioxygenase: structural basis of ethylmalonic encephalopathy. Hum. Mol.

Genet. 24, 2458–2469

16. McCoy, J. G., Bingman, C. A., Bitto, E., Holdorf, M. M., Makaroff, C. A., and Phillips, G. N.,

Jr. (2006) Structure of an ethe1-like protein from Arabidopsis thaliana. Acta Crystallogr. D Biol.

Crystallogr. 62, 964–970

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17. Otwinowski, Z., and Minor, W. (1997) Processing of x-ray diffraction data collected in oscillation mode. Methods Enzymol. 276, 307–326

18. Adams, P. D., Grosse-Kunstleve, R. W., Hung, L.-W., Ioerger, T. R., McCoy, A. J., Moriarty,

N. W., Read, R. J., Sacchettini, J. C., Sauter, N. K., and Terwilliger, T. C. (2002) PHENIX: building new software for automated crystallographic structure determination. Acta Crystallogr.

D Biol. Crystallogr. 58, 1948–1954

19. Emsley, P., Lohkamp, B., Scott, W. G., and Cowtan, K. (2010) Features and development of

Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501

20. Webb, B. N., Ballinger, J. W., Kim, E., Belchik, S. M., Lam, K. S., Youn, B., Nissen, M. S.,

Xun, L., and Kang, C. (2010) Characterization of chlorophenol 4-monooxygenase (TftD) and

NADH:FAD oxidoreductase (TftC) of Burkholderia cepacia AC1100. J. Biol. Chem. 285, 2014–

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21. Arisawa, M., Suwa, A., and Yamaguchi, M. (2006) RhCl3-catalyzed disulfide exchange reaction using water solvent in homogeneous and heterogeneous systems. J. Organomet. Chem.

691, 1159–1168

22. Choi, L.-S., and Bayley, H. (2012) S-Nitrosothiol chemistry at the singlemolecule level.

Angew. Chem. Int. Ed. Engl. 51, 7972–7976

23. Hegg, E. L., and Que, L., Jr. (1997) The 2-His-1-carboxylate facial triad: an emerging structural motif in mononuclear non-heme iron(II) enzymes. Eur. J. Biochem. 250, 625–629

24. Holm, L., and Sander, C. (1993) Protein structure comparison by alignment of distance matrices. J. Mol. Biol. 233, 123–138

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25. Altschul, S. F., Madden, T. L., Schäffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman,

D. J. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25, 3389–3402

26. Bebrone, C. (2007) Metallo--lactamases (classification, activity, genetic organization, structure, zinc coordination) and their superfamily. Biochem. Pharmacol. 74, 1686–1701

27. Hayes, R. P., Green, A. R., Nissen, M. S., Lewis, K. M., Xun, L., and Kang, C. (2013)

Structural characterization of 2,6-dichloro-p-hydroquinone 1,2- dioxygenase (PcpA), a new type of aromatic ring-cleavage enzyme. Mol. Microbiol. 88, 523–536

28. Cameron, A. D., Ridderström, M., Olin, B., and Mannervik, B. (1999) Crystal structure of human glyoxalase II and its complex with a glutathione thiolester substrate analogue. Structure 7,

1067–1078

29. Kabsch, W., and Sander, C. (1983) Dictionary of protein secondary structure: pattern recognition of hydrogen-bonded and geometrical features. Biopolymers 22, 2577–2637

30. Mineri, R., Rimoldi, M., Burlina, A. B., Koskull, S., Perletti, C., Heese, B., von Döbeln, U.,

Mereghetti, P., Di Meo, I., Invernizzi, F., Zeviani, M., Uziel, G., and Tiranti, V. (2008)

Identification of new mutations in the ETHE1 gene in a cohort of 14 patients presenting with ethylmalonic encephalopathy. J. Med. Genet. 45, 473–478

31. Stockdreher, Y., Venceslau, S. S., Josten, M., Sahl, H. G., Pereira, I. A., and Dahl, C. (2012)

Cytoplasmic sulfurtransferases in the purple sulfur bacterium Allochromatium vinosum: evidence for sulfur transfer from DsrEFH to DsrC. PLoS One 7, e40785

32. Pettersen, E. F., Goddard, T. D., Huang, C. C., Couch, G. S., Greenblatt, D. M., Meng, E. C., and Ferrin, T. E. (2004) UCSF Chimera: a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612

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CHAPTER THREE

STRUCTURAL AND BIOCHEMICAL CHARACTERIZATION OF CINNAMOYL-COA

REDUCTASES

Sattler SA, Walker AM, Vermerris W, Sattler SE, and Kang C. (2017). Structural and Biochemical Characterization of Cinnamoyl-CoA Reductases. Plant Physiology, 173(2), 1031- 1044.

All text, tables, and figures in this chapter are directly from the published manuscript.

3.1 Contributions

S.A.S., W.V., S.E.S., and C.K. conceived this project and designed experiments; S.A.S. and A.M.W. performed experiments; S.A.S., A.M. W., and C.K. analyzed data; S.A.S., A.M.W.,

W.V., S.E.S., and C.K. wrote the article.

3.2 Abstract

Cinnamoyl-coenzyme A reductase (CCR) catalyzes the reduction of hydroxycinnamoyl- coenzyme A (CoA) esters using NADPH to produce hydroxycinnamyl aldehyde precursors in lignin synthesis. The catalytic mechanism and substrate specificity of cinnamoyl-CoA reductases from sorghum (Sorghum bicolor), a strategic plant for bioenergy production, were deduced from crystal structures, site-directed mutagenesis, and kinetic and thermodynamic analyses. Although

SbCCR1 displayed higher affinity for caffeoyl-CoA or p-coumaroyl-CoA than for feruloyl-CoA, the enzyme showed significantly higher activity for the latter substrate. Through molecular docking and comparisons between the crystal structures of the Vitis vinifera dihydroflavonol reductase and SbCCR1, residues threonine-154 and tyrosine-310 were pinpointed as being involved in binding CoA-conjugated phenylpropanoids. Threonine-154 of SbCCR1 and other

CCRs likely confers strong substrate specificity for feruloyl-CoA over other cinnamoyl-CoA thioesters, and the T154Y mutation in SbCCR1 led to broader substrate specificity and faster

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turnover. Through data mining using our structural and biochemical information, four additional putative CCR genes were discovered from sorghum genomic data. One of these, SbCCR2, displayed greater activity toward p-coumaroyl-CoA than did SbCCR1, which could imply a role in the synthesis of defense-related lignin. Taken together, these findings provide knowledge about critical residues and substrate preference among CCRs and provide, to our knowledge, the first three-dimensional structure information for a CCR from a monocot species.

3.3 Introduction

Cinnamoyl-CoA reductase (CCR; EC 1.2.1.44) catalyzes the first committed reaction toward the biosynthesis of monolignols in plants. It hydrogenates hydroxycinnamoyl-CoA thioesters, producing the corresponding hydroxycinnamaldehydes and CoASH in an NADPH- dependent reaction. CCR activity was first reported by Wengenmayer et al. (1976) in soybean

(Glycine max) suspension cultures; the cloning of a CCR cDNA, from Eucalyptus gunnii, was first reported by Lacombe et al. (1997). The position of CCR in the phenylpropanoid pathway gives it a pivotal role in directing metabolic flux toward flavonoids and stilbenes or monolignols, hydroxycinnamic acids, and, depending on the species, hydroxycinnamoyl esters. The most common monolignols are p-coumaryl, coniferyl, and sinapyl alcohols. They are directed to the secondary cell walls, where they undergo oxidative coupling to form lignin polymers. The structures in lignin derived from the aforementioned monolignols are referred to as p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) residues (Ralph et al., 2004). Lignin fills the voids left between the complex network formed by cellulose microfibrils and hemicellulosic polysaccharides in secondary cell walls. The addition of lignin provides hydrophobicity to xylem cells and improves the mechanical strength of cells, providing structural support. Lignin also can

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be formed as a mechanical barrier in response to fungal infections or attacks from insects

(Vanholme et al., 2010).

The cloning of E. gunnii CCR enabled the cloning of several of its orthologs. This resulted in the identification of two CCR genes in maize (Zea mays; Pichon et al., 1998) and Arabidopsis

(Arabidopsis thaliana; Lauvergeat et al., 2001). Expression studies revealed that, in both species,

CCR1 is involved in lignification of stem tissues, whereas CCR2 is involved in lignification in response to attack by pathogens. In barrel clover (Medicago truncatula), however, the different

CCR paralogs appear to have distinct substrate specificity, with CCR1 displaying a preference for feruloyl-CoA and CCR2 for p-coumaroyl-CoA and caffeoyl-CoA (Zhou et al., 2010). The role of

CCR in lignification has been investigated with the use of mutants and transgenics. The vascular tissue of transgenic tobacco (Nicotiana tabacum), in which CCR was down-regulated, had thinner walls, brown vascular tissue, reduced lignin concentration, an increase in the S-G ratio, and displayed reduced growth and tissue abnormalities (Piquemal et al., 1998). The secondary cell walls in these plants contained tyramine ferulate, which was hypothesized to be a sink for the accumulated feruloyl-CoA esters (Ralph et al., 1998). The Arabidopsis irregular xylem4 mutant, identified based on its collapsed xylem phenotype, was shown to have a defective CCR1 gene

(Jones et al., 2001). This phenotype could be reproduced by transgenic down-regulation of CCR1

(Goujon et al., 2003). Excess feruloyl-CoA resulting from reduced CCR1 expression was converted to feruloyl malate (Derikvand et al., 2008). Two independent Arabidopsis ccr1 mutants contained less lignin than the wild-type control. The lignin of these mutants had a higher proportion of H residues and a lower proportion of S residues (Van Acker et al., 2013). In contrast, in maize, analysis of the transposon insertion mutant Zmccr1−, which had 31% residual CCR1 expression, showed slightly reduced lignin concentration in stem tissue, a slight increase in the S-G ratio, and

49

a reduction in H residues. In addition, the organization of the sclerenchyma fibers was affected.

The expression of several cell wall-related genes was decreased in this mutant, while several genes involved in flavonoid biosynthesis showed higher expression levels (Tamasloukht et al., 2011).

Despite species-specific differences, taken together, these data show a pivotal role of CCR in phenylpropanoid metabolism and in ensuring the integrity of secondary cell walls.

While the role of lignin is important for the survival of a plant, lignin also reduces the efficiency of the industrial use of plant biomass. The presence of lignin reduces the digestibility of forage crops (Jung and Buxton, 1994; Fontaine et al., 2003) and impedes the enzymatic saccharification of biomass to fermentable sugars that can be fermented to renewable fuels and chemicals (Chen and Dixon, 2007; Van Acker et al., 2013). As a consequence, a thermochemical pretreatment is necessary to reduce the recalcitrance of the plant cell wall to cellulolytic enzymes

(Hu and Ragauskas, 2012; Leu et al., 2013). The phenolic compounds derived from lignin during this process also are harmful to the microbes used during the fermentation (Ximenes et al., 2011).

The C4 grass sorghum (Sorghum bicolor) is receiving considerable attention as a lignocellulosic feedstock for the production of renewable fuels and chemicals, in part because of its ability to grow under harsh conditions, which include low-fertility soils, high temperatures, and extended periods of drought (Farré and Faci, 2006; Wang et al., 2014). Manipulation of lignin concentration and lignin subunit composition in sorghum through the incorporation of certain brown midrib (bmr) mutations had been shown to result in improved rumen digestibility (Porter et al., 1978) and greater efficiency of the enzymatic saccharification of sorghum biomass (Saballos et al., 2008; Dien et al., 2009) Given the important role of lignin for the survival of the plant, there is an inherent risk of reduced yields or lower biomass quality associated with the reduction in lignin concentration and/or change in lignin composition. These effects can be balanced with the

50

use of specific mutant alleles, as has been shown for the Bmr12 gene, which encodes caffeic acid

O-methyltransferase (Bout and Vermerris, 2003; Sattler et al., 2012). In this instance, the bmr12-

34 allele reduced lignin concentration to a level that was intermediate between wild-type sorghum and the bmr12-ref allele, which is a null allele, while still offering the same benefit of enhanced efficiency of enzymatic saccharification (Sattler et al., 2012).

Manipulation of the monolignol biosynthetic genes, such as cinnamyl alcohol dehydrogenase (Bmr6; Saballos et al., 2009; Sattler et al., 2009) and 4-coumarate-CoA ligase

(Bmr2; Saballos et al., 2012), also improved biomass conversion in sorghum (Saballos et al., 2008;

Scully et al., 2016). In order to maximize the toolkit for the manipulation of cell wall composition and the redirection of metabolic flux to secondary metabolites with health-promoting properties, it is important to have a detailed understanding of substrate specificity and catalytic mechanisms of the enzymes involved in the biosynthesis of monolignols. We have recently reported detailed structural and catalytic analyses of hydroxycinnamoyltransferase (Walker et al., 2013), caffeic O- methyltransferase (Green et al., 2014), and cinnamoyl-CoA O-methyltransferase (Walker et al.,

2016). Although previous studies on CCR have provided significant insight on the structure- function and kinetics of this enzyme (Pan et al., 2014), the structural basis for substrate selection and preference of CCR remains largely unknown. In this study, we provide detailed biochemical and structural findings that support our proposal for differentiating substrate selectivity among

CCRs. Knowledge from this study is expected to facilitate the engineering of sorghum as an improved livestock forage or bioenergy feedstock.

3.4 Materials and Methods

Chemicals and Enzymes

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Chemicals were obtained from Sigma-Aldrich or Fisher Scientific. Crystallization screens were obtained from Hampton Research and Qiagen.

Cloning and Enzyme Purification

cDNAs encoding SbCCR1 (Sobic.007G141200), SbCCR2 (Sobic.004G065600),

Sobic.002G146000, and Sobic.010G066000 from sorghum (Sorghum bicolor) were ligated downstream of the sequence encoding the 6X His tag of pET30a (EMD Millipore) for SbCCR1 or pET45a for Sobic.002G146000 and SbCCR2 and introduced in Escherichia coli Rosetta (DE3) cells (EMD Millipore) via heat shock transformation.

Cultures (1.5 L) of E. coli Rosetta (DE3) strains expressing SbCCR1 or 3-L cultures of E. coli Rosetta (DE3) strains expressing SbCCR2, Sobic.002G146000, or Sobic.010G066000 were grown at 37°C in Luria-Bertani medium prior to expression of the recombinant enzymes. Strain selectivity was provided by supplementing the medium with 50 µg mL-1 kanamycin and 30 µg mL-1 chloramphenicol (SbCCR1) or 100 µg mL-1 ampicillin and 30 µg mL-1 chloramphenicol for all other CCRs. For expression and purification of His-tagged CCRs, all cultures were allowed to reach OD600 = 0.6, brought to their respective induction temperatures (20°C for SbCCR1 or 22°C for all other enzymes), and induced to express by bringing the medium to 0.5 mM IPTG and allowing it to shake for 20 h. Cells were harvested by centrifugation at 5,000g, frozen, and then suspended in 35 mL of 50 mM Tris-HCl supplemented with 300 mM NaCl and 20 mM imidazole at pH 8. Cells were lysed by sonication, and lysates were cleared by centrifugation at 15,000g. The cleared lysates were stirred into 10 mL of Ni-NTA agarose resin (Qiagen), the column was washed with 2 volumes of the lysis buffer, and recombinant enzymes were eluted with buffer containing

50 mM Tris base, pH 8, 300 mM NaCl, and 250 mM imidazole. The eluted sample was concentrated and exchanged into 20 mM Tris-HCl, pH 7.5, then applied to a 10-mL MonoQ

52

column (GE Healthcare), and the enzyme was eluted with a 50 mM step-wise NaCl gradient in the same buffer. All CCRs eluted in approximately 150 mM NaCl, then they were buffer exchanged and concentrated into the appropriate buffer for crystallization or biochemical experiments. Purity was monitored for all protein preparations by SDS-PAGE, and protein concentrations were determined with the method of Bradford using BSA as a standard.

Protein Crystallization and Structure Determination

Crystals of SbCCR1 were grown using the hanging-drop, vapor-diffusion method. For crystallization, purified protein concentrated to 20 mg mL-1 in 20 mM Tris base, pH 7.5, 2 mM

DTT, and 1 mM NADP+ was mixed with an equal volume of reservoir solution and equilibrated against the same solution at 4°C. The reservoir solution contained 100 mM Bis-Tris, pH 6.5, and

25% (w/v) PEG 3350. Crystals of the enzyme typically appeared within 5 d. Adequate cryoprotection was achieved by passing crystals through a small drop of storage buffer/mother liquor mixture (50% of each component by volume) that was brought to a final concentration of

19% (v/v) glycerol. SbCCR1 in complex with NADP+ crystallized in the P32 space group with the asymmetric unit containing two monomers. Diffraction data were collected up to 2.9 Å resolution at the Berkeley Advanced Light Source (beam line 8.2.1). The data were processed with the

HKL2000 package (Otwinowski and Minor, 1997). The statistics for the diffraction data are listed in Table I. Initial phasing of the diffraction data was conducted by molecular replacement with the

PDB coordinates of CCR from Petunia hybrida (PDB identifier 4R1S) using PHENIX Phaser

(Adams et al., 2002). Model building and iterative refinements were conducted using the programs

COOT (Emsley et al., 2010) and PHENIX. The coordinates and diffraction data have been deposited in the PDB with identifier 5TQM.

Substrate Synthesis and Purification

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Feruloyl-CoA, caffeoyl-CoA, and p-coumaroyl-CoA were synthesized according to a method described previously (Walker et al., 2013) but with slight modifications. Briefly, the substrates were prepared enzymatically by the addition of recombinant 4-coumarate ligase from sorghum to a solution containing 50 mM sodium phosphate buffered to pH 7, 800 mM CoA, 2.5 mM ATP, 2 mM ferulic, p-coumaric, or caffeic acid, and 5 mM magnesium chloride. The reactions, ranging in volume from 20 to 50 mL, were gently rocked for no less than 16 h prior to termination. Termination was achieved by thermal denaturation of the enzyme at 80°C for 10 min followed by centrifugation at 15,000g for 30 min. Solutes in the supernatant were pelleted by centrifugation under vacuum. Substrates were purified using ethanol to crystallize the remaining reaction components. The remaining fluid containing pure hydroxycinnamoyl-CoA esters was centrifuged under vacuum to pellet the products, which were stored at -20°C for later use.

Enzyme Kinetic Assays

Wild-type SbCCR1, its mutants, and all other enzymes were purified and used for steady- state kinetic experiments in the presence of the potential substrates p-coumaroyl-, caffeoyl-, and feruloyl-CoA. For experiments using varying concentrations of thioester substrate, the 70-µL reactions contained 50 mM Bis-Tris, pH 6.5, 1 mM NADPH, 1 µg of enzyme, and 1, 2, 5, 10, 20,

50, 100, 200, 500, or 1,000 µM of the thioester substrate. All reactions proceeded at 30°C and were terminated with 30 µL of 17% (v/v) trifluoroacetic acid after various incubation times in the range of 30 s (SbCCR1 and the T154Y SbCCR1 mutant) to 20 min (Sobic.002G146000,

Sobic.010G066000, and the T154A SbCCR1 mutant). The reactions were centrifuged at 21,000g for 30 min at room temperature to pellet protein aggregates. A total of 50 µL of each reaction was injected onto a Luna C18(2) 5 µm, 4.6-mm X 150-mm column (Phenomenex) using the Hitachi

Organizer HPLC unit, and corresponding aldehyde products from each reaction were quantified

54

via peak integration from absorbance profiles at 333 or 346 nm. All reactions were performed in triplicate, and data were processed using Origin version 7.1 software (OriginLab).

Coniferaldehyde Docking

Molecular docking using SbCCR1 and coniferaldehyde was performed using AutoDock

Vina (Trott and Olson, 2010). The files used in the calculations were prepared using AutoDock

Tools 1.5.6 (Trott and Olson, 2010). The search space (x = 14 Å, y = 8 Å, and z = 22 Å) was defined considering the probable location of the phenylpropanoid-binding domain.

Coniferaldehyde was constructed using the program COOT, structurally optimized using

GaussView3 (Gaussian), and validated through comparison with the crystal structure of feruloyl adenylate in complex with a 4-coumarate ligase (PDB identifier 5BSV).

Site-Directed Mutagenesis and Mutant Expression

Two residues of SbCCR1, Thr-154 and Tyr-310, were mutated to Ala (A) and Phe (F), respectively, to test their involvement in binding of feruloyl-CoA. The T154A mutant was generated using the following primers: forward,

5’-TCCATCCGCGCGGTGGCCATGGACCCCAGC-3’, and reverse,

5’-GCTGGGGTCCATGGCCACCGCGCCGATGGA-3’. The Y310F mutant was generated using the following primers: forward,

5’-CGCGGAAGCAGCCGTTCAAGTTCTCGAACCAG-3’, and reverse,

5’-CTGGTTCGAGAACTTGAACGGCTGCTTCCGCG-39. Additionally, mutant T154Y was generated to observe any changes in substrate specificity. The primer sequences used to generate the mutant are as follows: forward, 5’-CATCGGCGCGGTGTACATGGACCCCAGC-3’, and reverse, 5’- GCTGGGGTCCATGTACACCGCGCCGATG-3’. E. coli Rosetta (DE3) cells were

55

transformed with plasmids containing the desired missense mutations in the SbCCR1 cDNA sequence, and proteins were expressed using the procedure described above.

ITC

Isothermal calorimetry titrations were performed for wild-type or mutant SbCCR1s and several ligands in an ITC200 instrument (Malvern Instruments). The protein was prepared by extensive buffer exchange (greater than 10,000 - fold) into the titration buffer, which consisted of

10 mM sodium phosphate buffered to pH 7. The concentration of protein in the calorimetric titration cell was diluted to 50 µM. All titrations were performed at 25°C with a stirring speed of

750 rpm and 27 injections (1.4 mL each). All ligands were brought to a concentration of 1 mM in the titration buffer and injected into the protein solution, and the heat of binding was recorded.

3.5 Results

Global Structure

Recombinant SbCCR1 (Sobic.007G141200) in complex with NADP+ was crystallized in the P32 space group, and its three-dimensional structure was determined at 2.9 Å resolution (Table

I). The asymmetric unit of the crystal lattice was composed of two monomers arranged in a noncrystallographic, 2-fold manner with limited intermolecular interaction. Calculations through

PISA (Krissinel and Henrick, 2007), which evaluates interactions between neighboring monomers in crystal lattices for the purpose of predicting biologically relevant oligomeric states, indicated that SbCCR1 exists as a monomer in solution (solvation free energy gain = 0.8 kcal mol-1 and interface complexation significance score = 0.0).

The overall structure of SbCCR1 was composed of two domains: an N-terminal domain harboring a typical Rossmann fold and a C-terminal domain of mixed a/b-fold, which was similar to the structures of CCRs from the dicotyledonous plants Petunia hybrida and M. truncatula (Pan

56

et al., 2014). The two domains were located on two distinct lobes that sandwiched the adjacent substrate and NADP(H)-binding pockets (Fig. 1).

In order to identify closely related structural homologs in the Protein Data Bank (PDB), a

Dali search (Holm and Sander, 1993) was performed using the atomic coordinates of SbCCR1.

The highest match was with a CCR from P. hybrida (PDB identifier 4R1S), with a Z score of

48.19, followed by a CCR2 from M. truncatula (PDB identifier 4R1U) and a cinnamyl alcohol dehydrogenase2 (PDB identifier 4QTZ) from M. truncatula, having respective Z scores of 43.4 and 41.1. A dihydroflavonol reductase from Vitis vinifera (PDB identifier 2C29) also was highly similar, with a Z score of 39.7, which is further addressed in “Discussion.” The next most closely related structures are methylglyoxal/isovaleral reductase Gre2 from Saccharomyces cerevisiae

(PDB identifier 4PVC), a vestitone reductase from Medicago sativa, and an aldehyde reductase2 from the yeast Sporobolomyces salmonicolor (PDB identifier 1Y1P), all of which had appreciably lower Z scores of 35.2, 34.5, and 32.5, respectively.

A BLAST search (Altschul et al., 1997) to identify proteins with similar amino acid sequences in the PDB revealed that CCR from P. hybrida (PDB identifier 4R1S) showed the highest identity (75%) to SbCCR1, followed by CCR2 from M. truncatula (PDB identifier 4R1U;

71%), cinnamyl alcohol dehydrogenase2 from M. truncatula (PDB identifier 4QTZ; 49%), and dihydroflavonol reductase from V. vinifera (PDB identifier 2C29; 41%). According to the BLAST output, the other enzymes identified with the Dali search showed no significant sequence similarity.

NADPH-Binding Pocket

+ From the early stages of structural refinement, the Fo-Fc map of the SbCCR1/NADP complex revealed clear electron density for one NADP+ molecule buried in a deep pocket on the

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N-terminal domain. This binding region was composed predominantly of a six-stranded, parallel

β-sheet (in the order of β3-β2-β1-β4-β5-β6; Fig. 2A), and its location opposes the putative hydroxycinnamoyl-CoA-binding pocket (Fig. 1). The diphosphate group of the bound NADP+ molecule was within hydrogen-bonding distance from the backbone secondary amines of both Tyr-

42 and Ile-43. In addition, the positive charge on NADP+ appeared to be compensated through the macroelectric dipole moment of the adjacent α-helix (α1), which spans between Tyr-42 and Gly-

55, through N-capping (Pereira et al., 2001). The adenine ring of NADP+ primarily interacted with

Arg-63 of β1 and Asp-89 of β2, while the adenosine ribose was bound solely through a salt bridge between its 29-phosphoryl group and the side chain of Arg-63. In short, the guanidinium group of

Arg-63 contributed to adenine binding through a cation-π interaction with the six-membered heterocyclic ring (Fig. 1), and the carboxyl group of Asp-89 was hydrogen bonded to an N6 hydrogen of the ring. The side chain of Leu-90 also appeared to engage in hydrophobic interaction with the entire ring.

The nicotinamide ring of NADP+ was oriented roughly in parallel to a short stretch of protein backbone, which included Pro-210 through Val-213, presumably providing stability through hydrophobic interactions (Fig. 1). In addition, the nicotinamide carboxamide was in close proximity to the backbone amine of Val-213 and the most proximal internal phosphoryl group of the ligand, which suggests that these interactions also contribute to its binding.

Hydroxycinnamoyl-CoA-Binding Pocket

The substrate-binding domain of the SbCCR1 was surrounded by two groups of α-helices, and the floor of the substrate-binding pocket was largely composed of β-strands. The first helix group included α5, α6, and α7, and the second helix group was composed of α8, α10, and α12 (Fig.

1). The two groups were observed to be on the opposing lobes, surrounding a group of β-strands

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(β8, β10, β11, and β13) that provided structural integrity for the putative phenylpropanoid-binding region. β8 ran perpendicular to β10, β11, and β13 and was sandwiched between the triad of strands and the nicotinamide ring of NADP+. β9 and β12 were in an antiparallel arrangement that formed a solvent-exposed wall distal to the nicotinamide ring.

To investigate specific interactions between SbCCR1 and the phenylpropanoid constituents of hydroxycinnamoyl-CoA substrates, numerous unsuccessful soaking and co- crystallization attempts were made using either feruloyl-CoA or coniferaldehyde. Consequently, molecular docking approaches were employed using these same compounds. The docking calculations positioned the aldehydic group of coniferaldehyde over the A-face of the nicotinamide ring of NADP+ in the trans-configuration (Fig. 2B), which allows pro-R hydride transfer from the

C4 atom of NADPH to the reactive carbonyl of the substrate(s). The distance between the aldehydic C9 atom of coniferaldehyde and the C4 atom of the nicotinamide ring was 2.3 Å. The lipophilic body of the ligand sat atop the side chains of Ile-150 and Val-211. The 3-methoxy and para-hydroxyl groups of the phenylpropanoid ring extended to 2.9 Å of the Thr-154 and Tyr-310 hydroxyl groups, respectively. Met-155, which is completely conserved among known CCRs, appeared to be in a suitable position to provide hydrophobic interaction with the 3-methoxy group on the ring of the ligand.

Isothermal Titration Calorimetry for Wild-Type and Mutant Forms of SbCCR1

Isothermal titration calorimetry (ITC) was used to determine the thermodynamic parameters for the association of potential substrates, products, and substrate/product analogs. For these experiments, ligand solutions were injected into a calorimetric cell containing protein solution, and consequent changes in thermodynamic states of the cell solution were recorded. A large amount of heat was released (ΔH = 26.1 kcal mol-1) when NADPH was used as the titrant.

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This enthalpic change was accompanied by a modest entropic contribution to binding (ΔS = 3.2

-1 -1 cal mol K ), resulting in a Kd of 6.5 mM (Fig. 3A; Table II). No significant thermodynamic changes were observed when NADH was used as the titrant, which indicated that the ligand did not bind to SbCCR1. CoA bound to SbCCR1 with a Kd value of 25 mM, but SbCCR1 did not show significant affinity for ferulic acid (Fig. 3A; Table II). Feruloyl-CoA bound to SbCCR1 with a Kd of 13 mM in the absence of NADP+, with ΔH and ΔS of 28.6 kcal mol21 and 26.4 cal mol-1 K-1, respectively. However, feruloyl-CoA bound only weakly or not at all in the presence of NADP+.

Caffeoyl-CoA displayed slightly higher affinity (Kd = 7.2 mM) but similarly had little affinity for the enzyme-NADP+ complex.

To confirm the involvement in binding of Thr-154 and Tyr-310, which were postulated from the molecular docking results to interact with functional groups on the phenolic ring of feruloyl-CoA through hydrogen bonding, site-directed mutagenesis was employed to generate

SbCCR1 mutants T154A and Y310F. The mutant proteins were expressed at similar levels to wildtype SbCCR1, and both were stable. Based on ITC data analysis, purified T154A and Y310F proteins displayed significantly lower affinity for feruloyl-CoA compared with the wild type (Fig.

3B; Table III), which supports that both residues contribute to binding feruloyl-CoA.

Identification of and Fold-Recognition Modeling for CCR Candidates in Sorghum

The amino acid sequence of SbCCR1 was used as a BLAST (Altschul et al., 1997) query to search the sorghum proteome (Phytozome database; https://phytozome.jgi. doe.gov/) for putative CCRs or CCR-like enzymes. The refined search yielded four additional candidates,

Sobic.004G065600, Sobic.002G146000, Sobic.010G066000, and Sobic.003g116800, all of which contained the currently accepted CCR signature sequence NWYCY. To begin investigating whether these enzymes were true CCRs, the amino acid sequences were first aligned using Clustal

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Omega and then truncated to the N and C termini of SbCCR1 observed in the crystal structure

(Fig. 4). The truncated sequences were submitted to the fold-recognition server Phyre2 using the

SbCCR1 sequence as a control, and the resulting three-dimensional structures of these four candidates were superimposed. Extensive alignment was observed between each candidate and

SbCCR1, with r.m.s.d. values ranging from 0.25 to 0.31 Å. The putative catalytic residues, which included Ser-149, Tyr-183, and Lys-187 of SbCCR1, were entirely conserved (Fig. 4) and in the proper positions for catalysis in all four CCR candidates.

Structural Comparisons between SbCCR1, PhCCR1, and MtCCR2

To structurally compare SbCCR1, PhCCR1 (PDB identifier 4R1S), and MtCCR2 (PDB identifier 4R1U), the atomic coordinates of those three crystal structures were submitted for tertiary structure alignment using the software program COOT. The superimposed structures depicted conservation of the residues involved in phenylpropanoid and NADP(H) binding as well as conservation of the Ser-Tyr-Lys catalytic triad (Figs. 4 and 5). The only significant difference is that Thr-154 of SbCCR1 is replaced with Tyr in PhCCR1. As expected due to isoform difference and the absence of NADP(H) in the MtCCR2 structure, the alignment between SbCCR1 and

MtCCR2 yielded a higher r.m.s.d. value (1.31 Å) than the alignment between SbCCR1 and

PhCCR1 (0.68 Å).

Enzyme Kinetics

Enzyme kinetic assays with the wild-type SbCCR1 were carried out in the presence of various hydroxycinnamoyl-CoAs, including feruloyl-CoA, caffeoyl-CoA, and p-coumaroyl-CoA.

While low activity was observed with caffeoyl-CoA and p-coumaroyl-CoA, much higher activity was observed for feruloyl-CoA (Fig. 6A). In addition, catalytic activities of three of its mutant forms, T154A, T154Y, and Y310F, were assessed with feruloyl-CoA or p-coumaroyl-CoA (only

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T154A and T154Y). Both T154A and Y310F mutants exhibited Vmax values for feruloyl-CoA that were approximately 5-fold lower than that of the wild-type enzyme as well as comparatively reduced catalytic efficiencies and higher Km values (Fig. 6B). However, the T154Y mutant exhibited 4.9- and 144-fold increases in catalytic efficiency for feruloyl-CoA and p-coumaroyl-

CoA, respectively, over those of wild-type SbCCR1 (Table IV).

Among the four additional sorghum genes with similarity to SbCCR1 identified through our BLAST search, Sobic.004G065600, Sobic.002G146000, and Sobic.010G066000 were isolated and could be expressed at levels sufficient for enzyme kinetic assays. As shown in Figure

7 and Table V, Sobic.004G065600 displayed a substantial increase in activity over SbCCR1 with p-coumaroyl-CoA, but its activity in the presence of feruloyl-CoA was relatively diminished.

Because of its demonstrated CCR activity, we named this protein SbCCR2. The enzymes encoded by Sobic.002G146000 and Sobic.010G066000 were far less active overall than SbCCR1 or

SbCCR2 under the assay conditions used and, thus, will be referred to in this report using only their gene identifiers until there is more clarity about their biological roles.

3.6 Discussion

Active Site, Kinetics of Enzyme Reaction, and Catalytic Mechanism

Our kinetic data indicate that SbCCR1 strongly prefers feruloyl-CoA as a substrate over the two other tested hydroxycinnamoyl-CoA thioester compounds (Figs. 6A and 7). Based on this substrate preference, the substrate-binding pocket was reexamined in order to pinpoint specific binding interactions between SbCCR1 and the phenylpropanoid moiety of feruloyl-CoA.

Although it was not the highest matches in our Dali and BLAST searches, dihydroflavonol reductase (DFR) from V. vinifera, which catalyzes the NADPH-dependent reduction of dihydroquercetin, is the only one that offers the ternary complex structure with its substrate

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(Trabelsi et al., 2008). Given that this dihydroflavonol and coniferaldehyde share critical aspects of chemical architecture, and that SbCCR1 contains functional analogs of DFR substrate-binding residues, we deemed the DFR an appropriate model for comparisons. The structural data for this reductase (PDB identifier 2C29) in complex with its substrate indicated that the enzyme uses the polar side chains of two residues (Asn-133 and Asn-227) for binding two hydroxyl groups located at the 3 and 4 positions on the aromatic ring of the substrate (Trabelsi et al., 2008). The backbone

α-carbons of SbCCR1 and the dihydroflavonol reductase superimposed well, yielding an r.m.s.d. value of 1.38 Å. Comparisons of the overlapped structures showed that the side chains of Thr-154 and Tyr-310 of SbCCR1 were at nearly equivalent positions in space to the side chains of Asn-133 and Asn-227 in the dihydroflavonol reductase, which suggest that Thr-154 and Tyr-310 could be their respective functional analogs. Our molecular docking results for coniferyl aldehyde and kinetic profiling of the T154A, T154Y, and Y310F mutants indicated substantial involvement of these two residues in binding feruloyl-CoA. Overall, these structural and kinetic data, coupled with knowledge of the substrate-binding mode of a close structural homolog, strongly suggest that

Thr-154 and Tyr-310 in SbCCR1 are directly involved in the binding of feruloyl-CoA.

In the proposed binding mechanism, the hydroxyl proton of Thr-154 interacted with the

3-methoxy oxygen of the ligand while the para-hydroxyl group established a hydrogen bond with the hydroxyl side chain of Tyr-310, as shown in the docked complex (Fig. 3C). This mechanism is consistent with the previous finding that substrate analogs bearing a 4-methoxy substitution resulted in comparatively less inhibition than endogenous substrate (Baltas et al., 2005). Given the close proximity of Gln-245 to the bound feruloyl-CoA, its side chain also could interact with the para-hydroxyl group of the substrate, which is held in the required conformer through a hydrogen bond with the hydroxyl group of Tyr-310. A Q245L mutant was generated to test whether this

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residue is essential for enzyme activity, but the mutant protein was not expressed at a level sufficient for protein purification. Nevertheless, kinetic profiling of a CCR1 with an amino acid substitution at Gln-245 will be needed to further increase the understanding of binding interactions between CCR1 and hydroxycinnamoyl-CoA substrates in lieu of a complex crystal structure.

New information from this study significantly augments knowledge of the reaction mechanism for CCRs and related reductases. NADPH and feruloyl-CoA associate with their binding pockets, which are located nearest the N- and C-terminal domains, respectively. Tyr-310 binds the para-hydroxyl of feruloyl-CoA, while Thr-154 binds the 3-methoxy group of this molecule (Fig. 8). Considering the apparent overlap of the substrates while they are both bound, they are predicted to follow an ordered, sequential mechanism, where NADPH binds first and is followed by the hydroxycinnamoyl-CoA substrate. In the first catalytic step, pro-R hydride transfer occurs from the C4 atom of NADPH to the reactive thioester carbonyl. The resulting oxyanion is temporarily stabilized by the oxyanion hole established from the side chain hydroxyl groups of

Ser-149 and Tyr-183. Collapse of the tetrahedral intermediate is then followed by C-S bond cleavage and protonation of the CoA thiolate. As indicated by our ITC data, in the presence of

NADP+, there is very low affinity for the CoA ester compounds, which precludes the formation of a nonproductive complex.

CCR Signature Residues and Residue Criteria for Substrate Specificity

To date, all confirmed CCRs share the 181NWYCY185 sequence motif. Although the central

Tyr of this motif (Tyr-183 in SbCCR1) is known to be involved in enzyme catalysis (Jörnvall et al., 1995), the roles for three of the remaining residues appear to be structural in nature by positioning the phenolic side chain of Tyr-183. In SbCCR1, the side chain of Trp-182 is engaged in hydrophobic interactions with nearby Pro-117 and Tyr-185. In addition, the Trp-182 side chain

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was hydrogen bonded to the side chain of Lys-177. Because the side chains of Tyr-185 and Pro-

117 both interact favorably with the Trp ring through face-to-face or face-to-edge interactions

(Samanta et al., 2000), it is likely that substitution of Trp-182 with another residue will destabilize the α-helix (a7) that contains the conserved NWYCY motif, which, in turn, could directly affect both the catalysis and binding of the CoA group. The electron density map of SbCCR1 indeed reveals a face-to-face interaction between the pyrrole portion of the indole ring of Trp-182 and the

Pro-117 side chain, while the orientation of the Tyr-185 side chain facilitates a face-to-edge interaction with the indole side chain of Trp-182. In addition, Cys-184 is in close proximity to

Cys-176, and it has been shown that the two residues contribute to cystine formation under oxidizing conditions (Pan et al., 2014). Given that a 40% loss of activity was observed under these conditions, it seems that reduction of CCR in this region affords flexibility that allows for tighter hydroxycinnamoyl-CoA binding. The role of Asn-181 is less apparent, but it may be involved in the binding of hydroxycinnamoyl-CoA substrates.

The two CCRs that have been identified in several species, CCR1 and CCR2, are currently distinguished by expression differences (Pichon et al., 1998; Lauvergeat et al., 2001) or substrate specificity (Zhou et al., 2010). However, the lack of knowledge about the signature sequence for differentiating the two isoforms based on substrate preference, which is not strictly consistent with their genetic identities, underscores the need for criteria to recognize a functional dichotomy at the protein level (Escamilla-Treviño et al., 2010; Zhou et al., 2010). Convoluting matters is that the fundamental basis for substrate specificity has remained unknown so far. As shown in Figures 6A and 7, our data demonstrate that SbCCR1 strongly prefers feruloyl-CoA over p-coumaroyl- or caffeoyl-CoA. Through analysis of activity data from the mutant enzyme (Fig. 6B) and ITC results

(Fig. 3), Thr-154 in SbCCR1 is clearly involved in directing affinity for feruloyl-CoA. A consistent

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feature of CCRs possessing Thr at this position is that they display a much higher preference for feruloyl-CoA than for other CCRs. Although PhCCR1, MtCCR1, MtCCR2, Sobic.002G146000, and SbCCR2 have a Tyr at this position and display higher activity for feruloyl-CoA, the disparities between their catalytic rates and efficiencies with feruloyl-CoA and p-coumaroyl-CoA or caffeoyl-

CoA are far less than in in vitro observations for PvCCR1a (Escamilla-Trevino et al., 2010) and

SbCCR1. The latter two enzymes have Thr at the equivalent position. Consistent with this observation, the T154Y SbCCR1 mutant exhibited substantially broader substrate specificity and a higher turnover rate. The fact that SbCCR1 is far more active with feruloyl-CoA implies that the other two hydroxycinnamoyl-CoA substrates bind to SbCCR1 in such a way that their reactive thioester bonds are not in position for efficient hydride transfer from NADPH. If this is true, then the physical premise for substrate specificity may not be due only to differences in phenylpropanoid-binding residues but also in the proximity of the conserved Tyr (Tyr-310 of

SbCCR1) to the nicotinamide ring.

Our structurally guided search for other CCRs among sorghum genomic data yielded four more candidates each containing the NWYCY sequence, which were expected to display CCR activity. Sobic.002G146000 is an ortholog of ZmCCR2 (Pichon et al., 1998) and, according to the

MOROHOSHI transcriptome database (Makita et al., 2015), was expressed robustly in sorghum roots, albeit at lower levels than SbCCR1. Sobic.003g116800 also was expressed in roots, but only under nitrogen stress. The remaining two genes, SbCCR2 and Sobic.010G066000, displayed low expression in all conditions and tissues examined and are not likely to play a significant role in the lignification of the secondary cell wall. By employing the same method as for SbCCR1, we were able to purify recombinant SbCCR2, Sobic.002G146000, and Sobic.010G066000. Although

Sobic.002G146000 and Sobic.010G066000 displayed low activity for both feruloyl-CoA and

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p-coumaroyl-CoA, SbCCR2 showed substantially higher activity with p-coumaroyl-CoA relative to SbCCR1. SbCCR2 contains Tyr at the position of Thr-154 in SbCCR1; thus, their activities are coincidentally consistent with those from other CCRs that possess Tyr at the equivalent position to Thr-154. Given these consistencies, it is possible that having Thr at this position drives a strong preference for feruloyl-CoA, whereas the presence of Tyr (or potentially residues other than Thr) affords CCRs broader reductive capacity. p-Coumaroyl-CoA is a precursor to a wide range of phenylpropanoid compounds that have been implicated in plant defense. Therefore, the observed substrate promiscuity of SbCCR2 may be relevant to plant defenses against pathogens, which include the production of defense lignin at the site of attack. Broadening substrate specificity to include significant activity toward p-coumaroyl-CoA, which as a precursor also for flavonoids and stilbenes is quite abundant, will enable the more rapid generation of monolignols. Furthermore, lignin containing p-coumaryl alcohol is more heavily cross-linked, because both C3 and C5 of the phenolic ring can participate in radical coupling reactions with the growing lignin polymer.

Consistent with this hypothesis, defense lignin formed in response to pathogen attack has been shown to have a higher proportion of H units in a number of different species (Lange et al., 1995;

Pomar et al., 2004; Zhang et al., 2007). The activities of Sobic.002G146000 and

Sobic.010G066000 were very low for both feruloyl-CoA and p-coumaroylCoA, leaving open the possibility that these two putative enzymes have different substrates in vivo.

3.7 Conclusions

Reducing lignin content can lower the barrier to the efficient production of renewable fuels and chemicals from lignocellulosic biomass. In view of this goal, CCR activity is necessary for the synthesis of all main monolignols, which makes CCR an attractive target for mutational and transgenic approaches that can modify this step of the pathway. Through molecular docking,

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mutagenesis, and kinetic analyses, we have identified a unique residue, Thr-154, that is critical to the association of the phenylpropanoid portion of feruloyl-CoA, the preferred substrate of

SbCCR1. Based on structural and sequence similarities to CCR1, we also have identified four additional CCRs or CCR-like enzymes in sorghum and, following kinetic analyses of the CCRs encoded by the two most highly expressed SbCCR genes, identified a signature residue for substrate specificity that can be exploited in genome-editing approaches. The detailed structural and other mechanistic knowledge forms the basis for engineering monolignol biosynthetic enzymes that display altered substrate specificity, have reduced substrate or product inhibition, or have greater velocities, so that metabolic flux can be modulated. The potential of this approach is illustrated by the single amino acid substitution T154Y in SbCCR1, leading to faster turnover and broader substrate specificity. Taken together, these findings provide structural information, experimental evidence for binding residue involvement, and information about previously unidentified CCR genes in sorghum.

3.8 Acknowledgement

This work was supported by the National Science Foundation (grant nos. MCB 102114,

CHE 118359, and 1231085 to C.K.), the National Institutes of Health (grant no.

1R01GM11125401 to C.K.), and the M.J. Murdock Charitable Trust (to C.K.); by the Biomass

Research and Development Initiative (grant no. 2011-1006-30358 to W.V.); by the U.S.

Department of Energy’s Office of Energy Efficiency and Renewable Energy, Bioenergy

Technologies Office (grant no. DE-PI0000031 to W.V.); and by the U.S. Department of

Agriculture (National Institute of Food and Agriculture AFRI grant no. 2011- 67009-30026 to

S.E.S. and CRIS project grant no. 3042-21220-032-00D).

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Table 3.1. X-ray diffraction data and refinement statistics for SbCCR1 (PDB identifier 5TQM)

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Table 3.2. Thermodynamic properties of interaction between SbCCR1 and various ligands structures

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Table 3.3. Thermodynamic properties of interaction between wild-type SbCCR1 or mutants T154A and Y310F and feruloyl-CoA

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Table 3.4. Kinetic values for wild-type SbCCR1 in the presence of three hydroxycinnamoyl- CoA substrates

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-

coumaroyl

-

p

CoA or

-

type and mutant of SbCCR1,type and forms SbCCR2,

-

Kinetic values for wild for values Kinetic

Table 3.5. of feruloyl presence in the Sobic.010G066000 and Sobic.002G146000, CoA

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Figure 3.1: Ribbon diagram of the global structure of SbCCR1 in complex with NADP+ and manually docked feruloyl-CoA. The N and C termini of the enzyme are marked N and C, respectively. The dashed curves at both chain termini stand in place of residues that could not be modeled in the experimental electron density map due to a disorder. NADP+, which is depicted with tan carbons in a stick model, was positioned according to its location in the experimental complex. Manually docked feruloyl-CoA is depicted with gray carbons in a stick model as well. During a reaction, the thioester bond of the substrate is positioned just above the nicotinamide ring of NADPH to promote hydride transfer. In this model, both lobes of the enzyme contribute residues to binding CoA, which may be accommodated by a conformational shift upon binding of the substrate. Molecular graphics images were produced using the UCSF

Chimera package.

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Figure 3.2: A, The observed NADP+ in the binding pocket of SbCCR1. NADP+ and all interacting residues are represented as stick models. The backbone of SbCCR1 is represented as a ribbon diagram, and dashed lines represent hydrogen bonds or ionic interactions. All residues that contribute to NADP+ binding are labeled according to their single-letter abbreviations and numbered according to sequence positions. The catalytic triad, composed of

Ser-149, Tyr-183, and Lys-187, is in close proximity to the nicotinamide ring and serves to promote hydride transfer to hydroxycinnamoyl-CoA substrates. B, Coniferaldehyde docked into the putative phenylpropanoid-binding region of SbCCR1. The backbone of SbCCR1 is represented by a ribbon diagram, with protruding side chains that contribute to coniferaldehyde binding modeled as sticks. Coniferaldehyde, which is the product of the reaction with the preferred substrate feruloyl-CoA, is shown in gray. Kinetics experiments with T154A and

Y310F mutants revealed that these two residues are critical for binding the phenylpropanoid portion of feruloyl-CoA. Molecular graphics images were produced using the UCSF Chimera package.

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Figure 3.3: A, ITC curves for wild-type SbCCR1 upon titration with various ligands. As expected, significant enthalpic events were observed when the enzyme was titrated with

NADP+ , NADPH, CoA, and feruloyl-CoA. Surprisingly, reasonable affinity was observed for p-coumaroylCoA and caffeoyl-CoA as well. B, ITC comparison between wild-type SbCCR1 and mutants T154A and Y310F upon titration with feruloyl-CoA. Significant heat release was observed upon titration of the wild-type enzyme (squares) compared with titrations of T154A

(triangles) or Y310F (circles).

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Figure 3.4: Multiple sequence alignment ofthe amino acid sequences for SbCCR1 and several related enzymes.Includedinthe alignment are a CCR1 from P. hybrida, CCR2s from M. truncatula and sorghum, a dihydroflavonol reductase from V. vinifera, and five CCRs or CCR- like enzymesfrom sorghum.Mining of sorghum proteomic data usingthe sequence of SbCCR1 as a query revealedfour additionalCCR candidates. Among the four additional candidates, high

CCR activity could only be confirmed for SbCCR2. Residues that are completely conserved among all enzymes in the alignment are marked with asterisks. Residues highlighted in yellow are in b-strands, while those in red are in a-helices. Residues involved in binding of substrates or cosubstrates are in boldface and underlined. The CCR signature sequence NWYCYis outlined in a box. The N and C termini of CCR candidates from sorghum were truncated to closely match the experimentally observed termini of SbCCR1. The colon (:) and the period

(.) indicate conservation of strongly similar properties and conservation of weakly similar properties, respectively.

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Figure 3.5: Superimposed tertiary structures of SbCCR1 and other CCRs. Depicted are

SbCCR1 (blue), PhCCR1 (tan), and MtCCR2 (gray). Catalytic and phenylpropanoid binding residues, as well as NADP+ from the SbCCR1 and PhCCR1 models, are represented as sticks.

Conserved residues are represented according to residue type and sequence position in

SbCCR1. Molecular graphics images were produced using the UCSF Chimera package.

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Figure 3.6: A, Michaelis-Menten curves for wild-type SbCCR1 in the presence of three hydroxycinnamoyl-CoA substrates. All curves were constructed using initial rate measurements. For each reaction, the concentration of NADPH was held constant at 1 mM and the concentrations of hydroxycinnamoyl-CoA substrates were varied. SbCCR1 is highly active in the presence of feruloyl-CoA but minimally active with caffeoyl- or p-coumaroyl-CoA. B,

Michaelis-Menten curves for wildtype SbCCR1 and two mutant forms, T154A and Y310F, in the presence of feruloyl-CoA. The two mutants were generated and tested for activity.

Significant loss of activity was observed with either mutant. Given the locations of the two residues within the substrate-binding pocket, these data indicate that both residues are involved in binding feruloyl-CoA. The data were processed using Origin Pro 2015.

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Figure 3.7: A, Reaction velocities in the presence of feruloyl-CoA or p-coumaroyl-CoA for several forms of CCR. Sobic.002G146000 and Sobic.010G066000 are represented using the last five numbers of their gene identifiers. B, Thr-154 and Tyr-310 in SbCCR1 are shown binding functional groups of coniferaldehyde. C, Fold-recognition model of the active site of

Sobic.004G065600 (SbCCR3). The bar graph was generated using Excel 2013 (Microsoft).

Molecular graphics images were produced using the UCSF Chimera package.

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Figure 3.8: Proposed catalytic reaction mechanism of SbCCR1. Using the reduction potential of NADPH, SbCCR1 executes the hydrogenation of hydroxycinnamoyl-CoA substrates to form hydroxycinnamyl aldehyde and CoASH. Thr-154 and Tyr-310, which are involved in substrate binding, are shown forming hydrogen bonds to functional groups around the aromatic ring.

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CHAPTER FOUR

CONCLUSIONS

Sulfide metabolism and monolignol biosynthesis are phenomena that require further investigation given their medical and industrial relevance. Nascent discovery of PDOs and lack of knowledge regarding their activity or molecular structures had prevented rationalizations of mutant phenotypes and the molecular etiology of EE. Likewise, the absence of structural information for monolignol biosynthetic enzymes of biofuel crop S. bicolor has precluded rational design of mutant genes that could alter lignin content and thus increase harvest efficiency of cellulosic matter from the plant. Our completed studies involving two PDOs and one CCR provide insight as we search for solutions to these problems.

PDOs have fallen under intense study since recognition of the dioxygenase as the ethe1 gene product and discovery of H2S as a naturally employed vasodilator. To date, few groups have submitted crystallographic information for this class of enzymes to the Protein Data Bank.

Following publication of the first PDO structure (hPDO1) by another group in 2015, our group reported crystal structures and other biochemical information for two PDOs (MxPDO1b and

PpPDO2). As our group published the first complex structure between a PDO and reaction product

GSH, we were able to rationally postulate involvements of residues in the EE disease state while perhaps aiding the developmental process of gene therapeutics. Additionally, the unusual metal-binding character and oligomeric state of CnPDO1 in its crystal lattice may suggest a relationship between metal coordination and oligomerization. Comparisons with any of the glyoxalases II may support this hypothesis given the similarities in metal-binding motifs between the monomeric enzymes and PDOs.

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The structure of a CCR had long been a mystery prior to publication of the Petunia and

Meticago structures in 2016. The former structure, also known as PhCCR1, was published in complex with NADPH and provided insight as to possible docking configurations of the phenylpropanoyl-CoA precursors to their respective monolignols. Our group determined the structure of SbCCR1 in complex with co-product NADP+, providing one of the first CCR structures. Using optimized models of SbCCR1’s primary reaction product, coniferaldehyde, in addition to considering the location of the nicotinamide ring of NADP+, we provided insight on the probable binding configuration of the product within the phenylpropanoid-binding region.

These hypotheses led to successful loss-of-function mutagenesis at two loci and altered function at one of the two, suggesting that lignification in S. bicolor could possibly be altered by employing our published mutation strategies. Our work also led to the discovery of an additional CCR in sorghum, as is shown in our report through activity analyses.

With these newly acquired biochemical and structural data for both types of enzyme, much remains to be understood about PDOs and CCRs. As of this writing, the relationship between metal-binding and oligomerization in PDOs has not been investigated and may lead to understanding of the structural and functional differences with the glyoxalases II. Although CCR has been crystallized in complex with NADP+, there is no crystallographic data for the interaction with its thioester substrates or aldehyde products. Structural knowledge of substrate configurations within CCR binding pockets would likely lead to discovery of new candidate residues for mutation, thereby increasing the likelihood that CCR activity could be modulated to impact lignin production to increase efficiency of ethanol fermentation. Further structural details are needed for both classes of enzymes, particularly in ligand complexes, to increase the range of possibility in practical application.

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