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The Pennsylvania State University

The Graduate School

Department of Biochemistry and Molecular Biology

MECHANISTIC STUDIES INTO BACTERIAL CELLULOSE SYNTHESIS

A Dissertation in

Biochemistry, Microbiology, and Molecular Biology

John B. McManus

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2017

This dissertation of John B McManus was approved* by the following:

Ming Tien

Professor of Biochemistry and Molecular Biology

Dissertation Advisor

Chair of Committee

B. Tracy Nixon

Professor of Biochemistry and Molecular Biology

Hemant Yennawar

Director, X-ray Crystallography Laboratory

Frank L. Dorman

Associate Professor of Biochemistry and Molecular Biology

Squire Booker

Professor of Chemistry

Professor of Biochemistry and Molecular Biology

Scott B. Selleck

Department Head, Biochemistry and Molecular Biology

*Signatures are on file in the Graduate School

ABSTRACT

Cellulose, a polymer of β-1,4-linked glucose units, is synthesized by plants, animals, fungi, and bacteria. It is the major component of the plant cell wall and, thus, is thought to be the most abundant biological molecule on earth. Cellulose synthase, a membrane-bound glycosyltransferase, synthesizes cellulose through the processive addition of glucose, from UDP-glucose, to the non-reducing end of the cellulose polymer.

This enzyme is often found with other proteins in large cellulose synthase complexes.

The primary focus of the present work is a greater understanding of the mechanisms for initiation, elongation, and termination of the cellulose polymer.

Structural studies of cellulose synthase from Rhodobacter sphaeroides, called BcsA-

BcsB, have revealed much regarding the mechanisms for polymer elongation, however the mechanisms for initiation and termination largely remain a mystery.

Two tools were developed for the present work. First was the isolation of a core catalytic subunit of the cellulose synthase complex. The catalytic heterodimer, AcsA-

AcsB, was isolated from Gluconacetobacter hansenii by two methods—product entrapment and affinity chromatography—and then subsequently kinetically characterized. Second was the development of a tool for the size analysis of in vitro- synthesized cellulose. Existing methods of cellulose solubilization and separation by gel permeation chromatography were adapted for this tool.

Initiation of cellulose synthesis can follow either a primer-dependent or a primer- independent mechanism. Using size analysis of in vitro-synthesized cellulose, both

AcsA-AcsB and BcsA-BcsB were shown to follow a primer-independent mechanism for

iii initiation. Reducing end analysis of in vitro-synthesized cellulose from both enzymes indicated that glucose is sufficient to initiate synthesis. Finally, BcsA-BcsB demonstrated

UDP-glucose hydrolase activity, allowing for the construction of a plausible mechanism for self-priming. Here, cellulose synthase hydrolyzes UDP-glucose and subsequently binds the liberated glucose to initiate new cellulose polymer synthesis.

Size analysis also revealed processivity differences between the two enzymes. To investigate the factors underlying processivity, changes in selected residues in the BcsA transmembrane channel were made. In one variant, the processivity and kinetic rates were altered.

Finally, size analysis indicated that the active elongation of the polymer by both

AcsA-AcsB and BcsA-BcsB is faster than the overall turnover numbers. This suggested that the catalytic cycle minimally consists of a fast and a slow phase. With these data, a minimal kinetic mechanism for cellulose synthesis was constructed.

To see if the overall turnover number was equally as slow in vivo, quantitative western blotting was employed to calculate the turnover number in whole G. hansenii cells. The turnover number in vivo was much faster than for purified AcsA-AcsB. Three cellulose synthase complex accessory proteins were investigated to test for their involvement in catalysis. Following this, the turnover number for AcsA-AcsB was measured at each step during purification by quantitative western blotting. This analysis showed a decrease in the turnover number immediately after cell lysis, suggesting that association into the cellulose synthase complex, in whole cells, positively impacts the cellulose synthesis rate of AcsA-AcsB.

TABLE OF CONTENTS

LIST OF ABBREVIATIONS ...... ix LIST OF FIGURES ...... x LIST OF TABLES ...... xii ACKNOWLEDGEMENTS ...... xiii CHAPTER 1: CELLULOSE AND CELLULOSE SYNTHESIS ...... 1 Cellulose Morphology, Crystallinity, and Degree of Polymerization ...... 3 Cellulose Synthesis in Bacteria ...... 6 Regulation of Cellulose Synthesis by cyclic-di-GMP ...... 8 Gluconacetobacter hansenii as a Model for Cellulose Synthesis ...... 9 The Molecular Biology of Cellulose Synthesis in G. hansenii ...... 12 BcsA-BcsB: The Structural Model for the Bacterial CSC Catalytic Core ...... 15 Initiation of Cellulose Synthesis ...... 20 Heparosan Synthase...... 22 Cyclic Glucosyl Synthase ...... 22 STATEMENT OF THE PROBLEM ...... 23 SUMMARY ...... 24 CHAPTER 2: ACSA-ACSB: THE CATALYTIC CORE OF THE CELLULOSE SYNTHASE COMPLEX ...... 28 Introduction ...... 28 Materials and Methods ...... 29 Culture Conditions ...... 29 Cloning of AcsAB-his ...... 30 Protein Concentration Determination ...... 30 AcsA-AcsB Purification by Product Entrapment ...... 31 AcsA-AcsB Purification by Affinity Chromatography ...... 32 Radiometric Measurement of Cellulose Synthase Activity ...... 32 Spectrophotometric Measurement of Cellulose Synthase Activity...... 33 SDS-PAGE and Western Blot Analysis ...... 33 N-terminal Sequencing...... 34

Trichloroacetic Acid-Mediated Cell Lysis ...... 34 Results ...... 34 Purification of AcsA-AcsB by Product Entrapment ...... 34 Processing of AcsAB ...... 38 Purification of AcsA-AcsB by Affinity Chromatography ...... 38 Kinetic Characterization of AcsA-AcsB ...... 42 Discussion ...... 45 AcsA-AcsB is the Catalytic Core of Cellulose Synthase ...... 45 Processing of AcsAB ...... 45 Kinetic Characterization of AcsA-AcsB ...... 46 CHAPTER 3: INITIATION OF CELLULOSE SYNTHESIS ...... 48 Introduction ...... 48 Materials and Methods ...... 50 Preparation of Solvents...... 50 Expression and Purification of BcsA-BcsB...... 51 Modification and Gel Permeation Chromatography of Cellulose in Tetrahydrofuran ...... 51 Dissolution and Gel Permeation Chromatography of Cellulose in Dimethylacetamide / 8% Lithium Chloride ...... 52 Reducing End Modification and Analysis ...... 53 Glucose Quantification by Enzyme-Linked Assay ...... 54 Gel Permeation Chromatography of Glucose and Cellobiose ...... 55 Reducing End Quantification Using Bicinchoninic Acid...... 55 Results ...... 56 Kinetic Characterization of BcsA-BcsB ...... 56 Gel Permeation Chromatography of Cellulose Tricarbanilates ...... 57 Gel Permeation Chromatography of Cellulose Synthesis Time Course ....57 Elongation Rate ...... 63 Reducing End Analysis ...... 64 BcsA-BcsB-Mediated UDP-glucose Hydrolysis ...... 67 Discussion ...... 70 Kinetic Characterization of BcsA-BcsB ...... 70

Product Analysis Demonstrates the Primer-Independent Mechanism for Initiation...... 70 Enzyme Concentration-Independent Rate Measurement ...... 71 A Mechanism for Self-Priming...... 72 CHAPTER 4: PROCESSIVITY AND KINETIC MODELING OF CELLULOSE SYNTHESIS ...... 75 Introduction ...... 75 Materials and Methods ...... 77 Cloning of BcsA-BcsB Variants ...... 77 Kinetic Simulation of Cellulose Synthesis...... 77 Results ...... 78 Kinetic Analysis of BcsA-BcsB Variants ...... 78 Gel Permeation Chromatography of Cellulose from BcsA-BcsB Variants ...... 79 Time Course Analysis of Cellulose from F416A ...... 80 Discussion ...... 82 Processive Synthesis ...... 82 A Minimal Kinetic Mechanism for Cellulose Synthesis ...... 85 CHAPTER 5: ACCESSORY PROTEINS AND WHOLE-CELL CELLULOSE SYNTHESIS ...... 92 Introduction ...... 92 Materials and Methods ...... 94 Determination of in vivo Turnover Number ...... 94 Total Membrane Isolation...... 95 Generation of t-DNA Insertional Mutants ...... 95 Statistical Analysis ...... 96 Cloning, Expression, and Purification of AcsD ...... 96

Cloning, Expression, and Purification of CcpAx ...... 96 Transmission Electron Microscopy ...... 97 Separation of Total Membrane by Sucrose Density Gradient ...... 97 Results ...... 98 Determination of in vivo Turnover Number ...... 98

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The Effects of AcsC, AcsD, and CcpAx on Enzyme Activity in Total Membrane ...... 100

The Effects of AcsD and CcpAx on Purified AcsA-AcsB Activity ...... 105

The Effects of AcsC, AcsD, and CcpAx Insertional Mutations on Cell Morphology ...... 107 Subcellular Localization of CSC Components ...... 108 Cellulose Synthesis Turnover Number upon Cellular Fractionation ...... 110 Discussion ...... 112 Determination of in vivo Cellulose Synthase Turnover Number ...... 112

The Effects of AcsC, AcsD, and CcpAx on AcsA-AcsB ...... 114 Subcellular Localization of CSC Components ...... 116 Cellulose Synthesis Turnover Number upon Cellular Fractionation ...... 116 CHAPTER 6: SUMMARY AND FUTURE DIRECTIONS...... 119 Methodological Contributions ...... 119 Initiation of Cellulose Synthesis Proceeds by a Primer-Independent Mechanism ...... 120 Processivity in Cellulose Synthase ...... 120 A Minimal Kinetic Mechanism for Cellulose Synthesis ...... 121 Cellulose Synthesis and the Cellulose Synthase Complex ...... 122 APPENDICES ...... 124 Appendix A: List of Primers ...... 124 Appendix B: Supplemental Figures ...... 126 Appendix C: Tenua Input...... 131 BIBLIOGRAPHY ...... 134

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LIST OF ABBREVIATIONS

AB Aminobenzamide Acs Acetobacter cellulose synthase BCA Bicinchoninic acid Bcs Bacterial cellulose synthase

BglAx β-glucosidase, Acetobacter xylinum

CcpAx Cellulose complimenting protein, Acetobacter xylinum CM Cytoplasmic membrane

CmcAx Carboxymethyl cellulase, Acetobacter xylinum CSC Cellulose synthase complex Cyclic-di-GMP Cyclic-di-guanosine monophosphate DDM Dodecylmaltoside DEAE Diethylaminoethyl DMSO Dimethyl sulfoxide DOP Degree of polymerization GPC Gel permeation chromatography LDS Lithium dodecyl sulfate MALLS Multi-angle laser light scattering NADH Nicotinamide adenine dinucleotide OM Outer membrane SDS Sodium dodecyl sulfate SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis SH Schramm-Hestrin TCA Trichloroacetic acid TEM Transmission Electron Microcopy TM Total membrane TN Turnover number UDP-glucose Uridine diphosphoglucose

LIST OF FIGURES

Figure 1.1. The Cellulose Polymer ...... 2 Figure 1.2. A Graphical Representation of Confirmed and Predicted Bacterial CSC Genes ...... 8 Figure 1.3. Cellulose Synthesis in Gluconacetobacter hansenii ...... 11 Figure 1.4. Cellulose Synthase from R. sphaeroides ...... 19 Figure 2.1. Isolation of AcsAB by Product Entrapment ...... 36 Figure 2.2. Total Protein from Whole Cells Isolated by TCA-precipitation ...... 37

Figure 2.3. Sequence Alignment between AcsAB and AcsB Homologs ...... 38 Figure 2.4. A Schematic Diagram of the Homologous Recombination used to Generate the Dodecahistidine-tagged acsAB Gene ...... 39 Figure 2.5. A Comparison of Cellulose Synthase Activity in TM from Wild Type and the Histidine-tagged Transformant ...... 40 Figure 2.6. Purification of AcsAB by Cobalt-Affinity Chromatography ...... 42 Figure 2.7. SDS-PAGE Analysis of Purified AcsA-AcsB ...... 43 Figure 2.8. An Enzyme-Coupled Assay for the Continuous Monitoring of Cellulose Synthase Activity ...... 44 Figure 2.9. Steady-State Kinetic Analysis of AcsA-AcsB ...... 44 Figure 3.1. GPC Elution Profiles of Cellulose Tricarbanilates ...... 58 Figure 3.2. GPC Elution Profile of Cellulose from BcsA-BcsB and AcsA-AcsB ...... 60 Figure 3.3. Total Glucose Incorporated into Cellulose from BcsA-BcsB and AcsA-AcsB ...... 61 Figure 3.4. Product Entrapment of Purified AcsA-AcsB and BcsA-BcsB ...... 63 Figure 3.5. GPC Elution Profiles of Cellulose from BcsA-BcsB and Michaelis-Menten Curve for the BcsA-BcsB Elongation Rate ...... 65 Figure 3.6. Reducing End Analysis of Cellulose by Reverse Phase HPLC ...... 66 Figure 3.7. BcsA-BcsB-Dependent Generation of Glucose from UDP-glucose ...... 67 Figure 3.8. BcsA-BcsB Soluble Product Analysis by Gel Permeation Chromatography ...... 69 Figure 3.9. A Proposed Mechanism of New Polymer Initiation by Cellulose Synthase ...... 74 Figure 4.1 BcsA-BcsB Transmembrane Channel Amino Acid Variants ...... 78

Figure 4.2. GPC Elution Profile of Cellulose from BcsA-BcsB Variants ...... 80 Figure 4.3. GPC Elution Profile of Cellulose from the F416A Variant ...... 81 Figure 4.4. Sequence Alignment of Selected Residues from Bacterial Cellulose Synthase A Subunits ...... 83 Figure 4.5. A Minimal Kinetic Mechanism for Cellulose Synthesis ...... 87 Figure 4.6. Simulated Michaelis-Menten Curves for BcsA-BcsB Using Tenua Software ...... 90 Figure 4.7 Simulated Product Distributions from Cellulose Synthase using Tenua Software ...... 91 Figure 5.1. The Calculation of a Turnover Number for Whole Cells ...... 100 Figure 5.2. Figure 5.2. Quantification of Cellulose Synthase Activity in Wild Type (WT) and Insertional Mutants ...... 102 Figure 5.3. Western-Blot Visualization and Quantification of SDS-Polyacrylamide Gels of TM from Wild Type, or acsC, acsD, or ccpAx ...... 103 Figure 5.4. Western-Blot Analysis of acsA Insertional Mutant ...... 104 Figure 5.5. Isolation of AcsA-AcsB and AcsA2-AcsB2 by Product Entrapment ...... 105 Figure 5.6. SDS-Polyacrylamide Gels of Heterologously Expressed and Purified CSC Components ...... 106 Figure 5.7. Transmission Electron Micrographs of G. hansenii Cells...... 107 Figure 5.8. Analysis of G. hansenii TM by Sucrose Density Gradient ...... 110 Figure 5.9. Analysis of acsC Insertional Mutant TM by Sucrose Density ...... 111 Figure B.1. Chromatograms for the Analysis of the Products of Edman Degradation ..126 Figure B.2. SDS-PAGE Analysis of Purified BcsA-BcsB ...... 127 Figure B.3. Steady-State Kinetic Analysis of BcsA-BcsB ...... 127 Figure B.4. GPC Elution Profiles of Native Cellulose Dissolved in DMAc / 8% LiCl .128 Figure B.5. Incubation of Pre-Synthesized Cellulose with Cellulose Synthase ...... 128 Figure B.6. GPC Elution Profiles of AB-Modified Cellulose ...... 129 Figure B.7. Steady-State Kinetic Analysis of BcsA-BcsB Variants ...... 130 Figure B.8. AcsB Quantification by Western-Blot Analysis ...... 130

LIST OF TABLES

Table 2.1. Purification Table for AcsA-AcsB ...... 41

Table 4.1. Kinetic Values for BcsA-BcsB Variants ...... 79

Table 4.2. Kinetic Values for AcsA-AcsB and BcsA-BcsB ...... 89 Table 5.1. The Effect of CSC Proteins on the Rate of Cellulose Synthesis from Purified AcsA-AcsB ...... 106 Table 5.2. Purification Table for Cellulose Synthase by Product Entrapment ...... 112

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ACKNOWLEDGEMENTS

First and foremost, I would like thank my advisor, Ming Tien. This work would not have been possible without constant support and fruitful discussions about experimental design and interpretation.

Thanks to Teh-hui Kao, Ying Deng, and Nita Nagachar for productive conversations and to Ying Deng for generating the insertional mutants and AcsAB-his expression strains used in this work. Thanks to Hui Yang for your diligent and continuing work on the kinetic modeling of cellulose synthesis.

Thank you to Jochen Zimmer for your kind gift of the BcsA-BcsB expression plasmid without which much of this would not have been possible.

Thank you to all my colleagues, especially Joseph Hill for your constant and consistent willingness to help in the laboratory and Bede Portz for everything else.

I would like to acknowledge the Penn State University facilities used in this work including the mass spectrometry facility, the shared fermentation facility, the genomics facility, and the microcopy and cytometry facility. Thank you to Mark Signs and Tania

Laremore, especially. Your willingness to meet with me on a drop-in basis was greatly appreciated. Thank you to Kimberly Martin, you will be missed.

Thank you to all the staff and faculty of the Center for Lignocellulose Structure and Formation for your financial and technical support throughout my graduate career.

Finally, thank you to my committee for taking the time to come to meetings, offer suggestions, and read the following manuscript.

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CHAPTER 1

CELLULOSE AND CELLULOSE SYNTHESIS

“he fell as an oak falls, or a poplar, or a tall pine, that among the mountains shipwrights fell, with whetted axes to be a ship's timber”

-The death of Asius, The Iliad

This passage from The Iliad illustrates the importance wood had as a material in the ancient world. The specificity with which the Greeks could name types of timber demonstrates its indispensable functions to the ancient civilization.

Though wood played a critical role in the development of civilization for thousands of years as both material and fuel, its major component, cellulose, was first identified in 1838 by French chemist, Anselme Payen. He described cellulose as the fibrous material left over from the wood after extraction with treatments of acid, ammonia, water, ether and alcohol (1), and elucidated the chemical formula as C6O5H10, or sugar.

Nearly a century later, the German chemist, Hermann Staudinger, discovered that cellulose is a polymer of glucose molecules bonded by glycosidic linkages at the C1 and

C4 carbons of each glucose unit (2). These glycosidic linkages are exceptionally stable.

The half-life for the rate of spontaneous hydrolysis has been estimated to be on the order of several million years (3), two orders of magnitude longer than the phosphodiester

1 bonds of DNA (3) and four orders of magnitude longer than the peptide bonds of protein

(3). The glycosidic linkages of cellulose are in the β configuration about the anomeric carbon. Each glucose unit is in a successive 180° rotation about the glycosidic bond, relative to the proceeding glucose. The resulting polymer adopts a linear configuration

(4) (Fig. 1.1). The straight, unbranched polymers of cellulose associate through inter- polymer hydrogen bonding and Van der Waals interactions (5). This makes cellulose insoluble in aqueous solutions and imparts a high degree of flexibility, tensile strength, and recalcitrance to digestion (6, 7). Unsurprisingly then, cellulose is a major structural component in a variety of biological systems.

Figure 1.1. The Cellulose Polymer. Cellulose is made up of β-linked glucose units with glycosidic bonds between the 1 and 4 carbons.

The physical properties of cellulose make it an excellent material for use in textiles, timber, pulp and paper, and an array of niche applications. The energy stored in the oxidative potential of the glucose monomers also makes cellulose a very good fuel source and a feedstock for the production of biofuels (8). The bulk of cellulose in nature is plant-synthesized (9), thus, the energy stored in the chemical bonds is derived from the photosynthetic process, making cellulose the most abundant renewable fuel source on earth (10). Its application to renewable fuel has resulted in recent efforts to better understand the chemistry and structure of cellulose and cellulose synthesis.

Cellulose Morphology, Crystallinity, and Degree of Polymerization

Cellulose synthesis not only occurs in plants, but in protists (11), fungi (12), tunicates (13), and bacteria (14). As mentioned, plants comprise the largest global synthesizers of cellulose, producing it on the order of several billion tons per year (15).

On average, cellulose makes up 45% of the lignocellulosic material found in plant cell walls (16, 17).

Cellulose synthesis is carried out by membrane-bound glycosyltransferases called cellulose synthases. These enzymes catalyze the elongation of cellulose by the addition of glucose units from the nucleotide sugar, UDP-glucose, to the non-reducing end of the cellulose polymer (18, 19). As elongation of the polymer occurs, it is translocated across the membrane and the reducing end is pushed out into the extracellular milieu. The polymer crystallizes with other nearby cellulose polymers to form a repeating structure, called a microfibril. A microfibril was originally defined in the literature as the thinnest

3 cellulose unit visible by transmission electron microscopy (20). The ambiguity in this definition means that the dimensions of a microfibril can vary depending on the organism from which it is derived (21). Individual cellulose synthases associate, along with other accessory proteins, to form a cellulose synthase complex (CSC) (22-25). The spatial arrangement of the CSC imparts the morphology of the resultant cellulose fibrils (26-30).

For example, plants, which have clustered, rosette-like CSCs, produce barrel-like fibrils

(31), whereas some bacteria, which have linear CSCs, produce ribbon-like cellulose fibrils (32).

The majority of crystalline cellulose comes in two conformations: cellulose I and cellulose II. The polymers of cellulose I run parallel to each other (i.e. every reducing end pointing in the same direction), while polymers of cellulose II run anti-parallel to each other (33). Cellulose II is the more thermodynamically favorable of the two allomorphs due to an additional hydrogen bond per glucose residue (34). Because of the directionality of enzyme-mediated synthesis, however, cellulose I is the predominant allomorph synthesized by most organisms (35). Two exceptions exist. The alga

Halocystis (36) and the bacterium Sarcina (37) naturally give rise to cellulose II. The treatment of cellulose I fibrils, either with alkali or dye, causes perturbations in the structure that result in the irreversible formation of cellulose II (38, 39). In a process called Mercerization, sodium hydroxide treatment is used to convert the cellulose I fibers of cotton to cellulose II, adding strength and luster to the material (40, 41) Cellulose I is further subdivided into two additional allomorphs: cellulose Iα and cellulose Iβ. The crystal structures of each of these sub-allomorphs have been determined (33, 42).

Diffraction patterns of microfibrils reveal a mixture of both sub-allomorphs, and the

4 ratios of each sub-allomorph synthesized by an organism vary from species to species

(33). Other less common configurations of crystalline cellulose exist. Cellulose III and cellulose IV are two such configurations. These allomorphs are derived from cellulose I and II by various chemical and heat treatments (43)

Crystallinity in cellulose is the degree to which the atoms of the glucan polymers forming cellulose fibrils arrange in a periodic structure (44). The degree of crystallinity varies for cellulose produced by different organisms. Bacterial cellulose exhibits the highest degree of crystallinity (60%-90%) (45). Compare this with Arabidopsis-derived cellulose at 50% (46) and cotton-derived cellulose at only 32% (47). This variation may be due, in part, to the prevalence of other extracellular polysaccharides, such as hemicelluloses and lignin, found in plants. These polymers may disrupt the regularity of the repeating structure, possibly through intercalation (48, 49). However, this cannot be the only determinant, as cotton contains more cellulose as a percent of lignocellulosic material than does Arabidopsis.

The degree of polymerization (DOP) in cellulose is the number of glucose units making up a single glucan polymer. In other words, the DOP is a representation of the length of a glucan polymer. The DOP of cellulose has relevance to industrial applications. For example, paper brittleness and aging is impacted by the cellulose DOP

(50). The DOP is also relevant in the production of biofuels, as this characteristic can affect the release of glucose during chemical or enzymatic digestions of biomass (51).

DOP of a cellulose polymer is different from the length of a cellulose microfibril, an important distinction, as a microfibril may consist of many overlapping cellulose

5 polymers. An average DOP for a single glucan polymer within a population is determined by measuring the total number of glucose units within the population and dividing that number by the total reducing or non-reducing ends OR by dividing the average molecular weight of a population by the molecular weight of anhydroglucose (162 g/mol). This can be achieved by chemical analysis (52), mass spectrometry (53), or viscometry (54). If a

DOP range of a population of polymers is desired, then gel permeation chromatography

(GPC) coupled with multi-angle laser light scattering (MALLS) or column calibration with known standards can be employed (55-57). This method requires either modification of the cellulose or dissolution in ionic liquids (56, 57), as cellulose is insoluble in aqueous solvents. The DOP of cellulose varies depending on the source (51). DOP analysis is an important tool and will be utilized later in this thesis.

Cellulose Synthesis in Bacteria

While plants contribute to the bulk of cellulose in nature (9), bacteria have demonstrated the ability to produce cellulose, which serves as a structural component in biofilms (14, 45, 58-62). It was through homology searches with known bacterial cellulose synthase cDNAs that genes encoding plant cellulose synthases were first identified (63). Bacteria from the genera Escherichia, Pseudomonas, Salmonella,

Gluconacetobacter, Agrobacterium, Azotobacter, and Rhizobium have the ability to synthesize cellulose (64-66). Generally, the core cellulose synthase genes are organized into an operon containing bcsA, bcsB, bcsC, and bcsZ (Fig. 1.2). The bcs nomenclature comes from bacterial cellulose synthase. The typical nomenclature is sometimes replaced

6 with acs or cel along with the subsequent letter to designate the homolog (example: acsA is homologous to bcsA).

BcsA and bcsB encode the proteins forming the catalytic core of the bacterial CSC

(19, 67, 68). BcsA contains both the glycosyltransferase and regulatory domains. Little is known about BcsB’s precise function; however, it is necessary for catalytic activity (58,

69). In some species bcsA and bcsB are fused to form a single open reading frame (67), as is the case for Gluconacetobacter hansenii shown in Figure 1.2 (called acsAB). This phenomenon is a subject of investigation for G. hansenii ATCC23769 in Chapter 2. In other species (X. campestris shown in Figure 1.2) the bcsA gene is split into two open reading frames (70). BcsC is thought to encode a porin-like protein which facilitates translocation of cellulose out of the cell (60). Finally, bcsZ encodes an endoglucanase required for cellulose synthesis (71). Other genes have been identified which are important to cellulose synthesis, yet are not found in all cellulose synthesizing organisms.

For example, bcsD, is found in bacteria that synthesize crystalline cellulose (72).

Figure 1.2. A Graphical Representation of Confirmed and Predicted Bacterial CSC Genes. Homologous genes are colored identically. Genes not organized into an operon are spatially separated. Open reading frames and spacing are not drawn to scale.

Regulation of Cellulose Synthesis by cyclic-di-GMP

The regulation of cellulose synthesis in bacteria is mediated by the secondary messenger molecule, cyclic-di-GMP (65). Cyclic-di-GMP was discovered in conjunction with the activation of cellulose synthesis in vitro. Ross et al. (73) recognized that the addition of a “crude factor” from the supernatant of total membrane isolation resulted in a significant increase in the levels of cellulose synthesis in total membrane (in vitro). This

“crude factor” was later identified as cyclic-di-GMP (74).

Since its discovery, cyclic-di-GMP has been implicated in the regulation of a variety of bacterial functions, including biofilm formation, quorum sensing, motility,

8 pilus formation, and virulence (75-80). These functions generally coincide with either the sessile or motile phases of bacterial growth. High concentrations of cyclic-di-GMP promote activities involved in sessile growth, while low concentrations promote activities involved with motility (75).

The intracellular levels of cyclic-di-GMP are regulated by the actions of diguanylate cyclases, responsible for the formation of the molecule, and phosphodiesterases, which mediate its breakdown (81-83). The relative activities of these two classes of enzymes increase or decrease the concentrations of cyclic-di-GMP in response to various biological stimuli.

The PilZ domain, discovered in conjunction with pili formation (84, 85), is a cyclic-di-GMP binding motif and is found in proteins regulated by cyclic-di-GMP. A

PilZ domain is found in the BcsA peptide of bacterial cellulose synthase and binds cyclic- di-GMP to activate cellulose synthesis (19, 73). This is consistent with the formation of biofilms and promotion of sessile growth. More will be discussed on the molecular mechanism of regulation as it relates to cellulose synthase later in this Chapter.

Gluconacetobacter hansenii as a Model for Cellulose Synthesis

Gluconacetobacter hansenii (formerly Acetobacter xylinum), a gram negative, obligate aerobe, is the premier model for bacterial cellulose synthesis, and is also an excellent surrogate for understanding cellulose synthesis in plants. G. hansenii has been studied extensively since the 1950s beginning with the work of Schramm and Hestrin

(62, 86, 87), however, demonstrations of cellulose synthesis in G. hansenii were reported as early as the 1880s (88). When cultured under static conditions, the bacteria generate a thick pellicle of cellulose, formed at the liquid-air interface of the medium (Fig. 1.3A).

Ensnared in the meshwork of this pellicle are the G. hansenii cells (87). An individual cell is capable of polymerizing up to 200,000 glucose units per second into glucan polymers which aggregate into a ribbon-like cellulose fibril (32, 62) (Fig. 1.3B).

Observation of fibril extrusion under dark-field microscopy reveals that a single cell can synthesize cellulose at a rate of 2 µm per minute (32). This is equivalent to an incorporation rate of 67 glucose molecules per second per cellulose synthase.

G. hansenii is capable of converting up to 50% of the available carbon into cellulose (89). The biological significance of cellulose synthesis in G. hansenii remains unclear; however, the prevailing hypothesis is that the formation of a pellicle at the liquid-air interface provides these aerobic bacteria with increased exposure to oxygen

(30, 62).

The CSC of G. hansenii forms a linear array, composed of cellulose synthase particles, 50-80 nm in length, longitudinally, across the cell surface (26, 28, 90, 91).

Evidence for this comes from freeze-fracture microscopy (Fig. 1.3C) (28, 32), immuno- gold labeling followed by visualization with TEM (91), and fluorescence microscopy

(Fig. 1.3D) (90). The cellulosic ribbon-like fibril extruded by the linear CSC is formed by the association of newly synthesized glucan polymers in a process of hierarchical aggregation which can be characterized in four steps. First, each cellulose synthase particle extrudes a 1.5 nm wide sub-elementary fibril consisting of 10-15 glucan

10 polymers (92). The sub-elementary fibrils then aggregate to form a 3.5 nm wide elementary fibril (65). Elementary fibrils within close proximity crystallize into microfibrils, 6-7 nm in width. Finally, the 40-60 nm wide ribbon forms from the bundling of the microfibrils (32).

Figure 1.3. Cellulose Synthesis in Gluconacetobacter hansenii. (A) The cellulose pellicle formed at the liquid-air interface by G. hansenii. (B) Freeze-fracture image of the cellulosic ribbon formed by G. hansenii (32)*. (C) Freeze-fracture image of CSC particles on the G. hansenii cell surface (black arrows) (32)*. (D) Fluorescence microscopic image of GFP-AcsD arranged into a linear array on a G. hansenii bacterium (90)**.

* Reproduced with permission from R. Malcolm Brown. ** Reproduced with permission from Elsevier Publishing. 11

The Molecular Biology of Cellulose Synthesis in G. hansenii

The genes required for cellulose synthesis in G. hansenii were identified by mutagenesis and observation of the impact on cellulose synthesis in vivo (25, 59, 60, 90,

93, 94). The G. hansenii genes and proteins follow the acs nomenclature, for acetobacter cellulose synthase, rather than the generic bcs.

The six genes described below encode proteins that are thought to be integral to the G. hansenii CSC. While there are other proteins known to impact cellulose synthesis, for example diguanylate cyclase, which regulates intracellular cyclic-di-GMP concentrations, their activities affect a variety of biochemical processes and therefore will not be discussed.

AcsAB encodes a 168 kDa peptide which is cleaved into two subunits, AcsA and

AcsB (58). These two peptides associate to form the catalytic core of the cellulose synthase complex (58). AcsA, the N-terminal half of the AcsAB peptide, was identified as the UDP-glucose binding domain using photo-affinity labeling with azido-UDP- glucose (68). AcsA contains a QxxRW motif, conserved in β-glycosyltransferases (95). It contains the PilZ regulatory domain and the glycosyltransferase domain (25, 58, 96), and is therefore the catalytic and regulatory subunit of the CSC. Membrane topology analysis and comparison with BcsA from R. sphaeroides, indicate that AcsA contains nine transmembrane domains and that it is anchored in the cytoplasmic membrane (19, 97).

The C-terminal half of AcsAB is AcsB. Comparison with BcsB from R. sphaeroides indicates that this protein largely resides in the periplasmic space (19).

AcsC is located just downstream from acsAB and encodes a 138 kDa protein (98).

The start codon of acsC overlaps with the stop codon of acsAB. Together, these genes make up the acsABC operon (60, 98). Homology studies with outer membrane porins suggest that AcsC forms a β-barrel structure through which newly synthesized glucan polymers can exit the cell (97, 99). Secondary structure predictions show that AcsC contains seventeen tetratricopeptide repeats (TPR) (99). TPRs may occur in membrane transport proteins and are known to be involved in binding a diverse set of ligands. (100-

102). The predicted size of the channel formed by AcsC suggests that it accommodates the passage of four glucan polymers (99). Mutations which disrupt acsC yield cells unable to synthesize cellulose in vivo (60), however, this gene is not necessary for cellulose synthesis in total membrane isolated from whole cells (58). This is consistent with its prediction as an exit channel for cellulose.

Originally grouped into the acsABC operon, downstream from acsC, acsD has since been shown to possess its own transcriptional promoter (60, 97). The product of acsD is a 16 kDa protein that forms a homo-octamer, localized to the periplasm (103,

104). The crystal structure of AcsD shows a central pore, formed by the octamer, large enough to accommodate the passage of four glucan polymers (104). This central pore is similar in size to that predicted for the AcsC central pore, and suggests that AcsD functions as a glucan chaperone, guiding polymers synthesized by AcsA-AcsB through the periplasm to AcsC. Mutations that disrupt acsD yield cells with a reduced capacity to synthesize cellulose (25, 60). The cellulosic pellicle produced by these cells in static culture is thin compared to wild type cells and X-ray diffraction analysis reveals that the pellicle is composed primarily of the cellulose II allomorph (25).

Downstream of acsD is bglAx, which encodes a 79 kDa β-glucosidase (105, 106).

Mutations that disrupt bglAx yield cells with a reduced capacity to synthesize cellulose in vivo (60).

Upstream from the acsABC operon is an operon consisting of cmcAx and ccpAx

(60). CmcAx encodes a secreted 45 kDa β-1,4-endoglucanse (107, 108). Mutations in cmcAx yield cells with a reduced capacity to synthesize cellulose in vivo. Microscopy studies on these mutants reveal highly twisted cellulose fibrils (94). Koo et al. (107) showed similar results in in vivo cellulose production levels when anti-CmcAx antibodies were added to G. hansenii cultures. CmcAx may assist in relieving hyper-twisting by cleaving irregularly-packed glucan polymers (94). The addition of heterologously- expressed and purified CmcAx to G. hansenii cultures resulted in increased cellulose production (108).

CcpAx encodes an 8 kDa protein that co-localizes with GFP-tagged AcsD in the linear CSC array (Fig. 1.3D) (90). Mutations which disrupt ccpAx yield cells that are unable to synthesize cellulose in vivo (60, 90). Interestingly, these same mutants, containing GFP-AcsD, show a dispersed linear array when examined by fluorescence microscopy, suggesting that CcpAx is integral to proper CSC formation (90).

BcsA-BcsB: The Structural Model for the Bacterial CSC Catalytic Core

In 2013 the first crystal structure of cellulose synthase (Fig. 1.4) was solved using

BcsA-BcsB from Rhodobacter sphaeroides (19). BcsA-BcsB is homologous to the G.

14 hansenii AcsA-AcsB. Though R. sphaeroides is not known to synthesize cellulose in culture, bcsA and bcsB, when expressed in E. coli and purified, synthesize cellulose (69).

The crystallographic snapshots of BcsA-BcsB reveal important features about how glucose units are added to the cellulose polymer and how that polymer translocates across the cytoplasmic membrane (19, 109, 110).

BcsA and BcsB compose a heterodimer in a one to one stoichiometry (19) (Fig.

1.4A). The BcsA subunit consists of a cytosolic region and a transmembrane region. The cytosolic region contains the glycosyltransferase domain and the cyclic-di-GMP-binding

PilZ domain. The transmembrane region, which anchors BcsA in the cytoplasmic membrane and facilitates translocation of the cellulose polymer, consists of nine amphipathic alpha helices, six of which form an 8 Å wide, 30 Å long channel in which 10 glucose units of the nascent cellulose polymer reside (Fig. 1.4B). The interior side chains of the transmembrane channel interact with the cellulose polymer either by Van der

Waals interactions with the faces of the glucose rings or through hydrophilic interactions with the equatorial hydroxyls of the glucose units. Egress of the glucan polymer into the periplasmic space causes it to kink sideways and align nearly parallel to the periplasmic leaf of the cytoplasmic membrane (Fig. 1.4A) (19).

The non-reducing end (the terminal glucose) of the glucan polymer sits at the base of the transmembrane channel, with the 4-carbon hydroxyl oriented towards the glycosyltransferase domain. The face of the terminal glucose ring interacts with trp383 and the 2-carbon hydroxyl hydrogen bonds with tyr302. A “finger helix” motif, containing the catalytic base, asp343, sits in the glycosyltransferase domain pointing

15 upward, towards the entrance to transmembrane channel. The side chain of asp343 interacts with the 4-carbon hydroxyl of the terminal glucose facilitating nucleophilic attack on the 1-carbon of the donor UDP-glucose (Fig. 1.4C). Mg2+ acts as a Lewis acid, stabilizing the transfer of the electron pair from the 1-carbon of the donor glucose to the phosphate of UDP during glycosidic bond formation. The Mg2+ is coordinated by the side chains of asp246 and asp248. Glycosidic bond formation is thought to proceed through an

SN2-like single displacement mechanism (111). This type of reaction mechanism leads to inversion of stereochemistry about the anomeric carbon (19, 110).

After glycosidic bond formation, a conformational change occurs in which the finger helix motif shifts down into the glycosyltransferase domain. In this position, asp343, as well as thr341, hydrogen bond with the 4- and 2-carbon hydroxyls, respectively, of the newly added glucose (110). The finger helix then shifts back up again, towards the opening of the transmembrane channel. Commensurate with this shift is the translocation of the cellulose polymer by one glucose unit (110). The precise mechanism underlying the finger helix shift remains unclear; however, it may be facilitated individually or by a combination of substrate binding, product release, or the relaxation of the newly added glucose into the 180° rotation relative to its neighboring glucose.

Cyclic-di-GMP-mediated activation of cellulose synthesis occurs by binding of the regulatory molecule to the PilZ domain, resulting in a conformational change that grants UDP-glucose access to the glycosyltransferase domain. In the absence of cyclic-di-

GMP, an unstructured region, called the gating loop, extends across the opening of

16 glycosyltransferase domain, blocking access. The gating loop is held in this position by a salt bridge formed between the backbone of thr511, on the C-terminal end of the gating loop, and arg580 in the PilZ domain (109). In the absence of cyclic-di-GMP, little catalytic activity is detectable (69). Upon binding two cyclic-di-GMP molecules, the gating loop moves away from the opening of the glycosyltransferase domain and toward the cytosolic leaflet of the cytoplasmic membrane. This conformational change occurs when arg580 coordinates with cyclic-di-GMP, thereby breaking the threonine-arginine salt bridge. An arg580 to ala580 substitution results in a constitutively active form of

BcsA-BcsB (109). Upon binding UDP-glucose, the gating loop adopts a third configuration, insertion into the glycosyltransferase domain (109).

The other subunit of the cellulose synthase heterodimer, BcsB, resides primarily in the periplasmic space, save for a single C-terminal alpha helix anchoring BcsB to the cytoplasmic membrane (Fig. 1.4A). While the precise function of BcsB remains unclear,

Omadjela et al. (69) have shown that the expression of BcsA alone results in the loss of catalytic activity. Interestingly, through systematic truncations of BcsB, these workers demonstrate that only the C-terminal transmembrane helix is required to retain catalytic activity in BcsA (69). Structural studies show that BcsB’s helix packs into a groove formed by transmembrane helices 1, 2, and 3 of BcsA, and thus suggest that BcsB may be necessary to maintain the proper structure of BcsA’s transmembrane channel (69). The periplasmic portion of BcsB forms a dome over the exit channel of BcsA. This portion of

BcsB contains two carbohydrate binding domains. The first runs parallel to the periplasmic leaflet of the cytoplasmic membrane above where the glucan polymer kinks

17 to run parallel to the periplasmic leaflet. The second carbohydrate binding domain lies on the surface of BcsB facing out into the periplasm (19).

A Phe426 B Glu463

Arg423 Ser476 BcsB Phe416 Asn412 Glu297

Phe301 Trp383

Tyr302 Trp383

BcsA Asp343

Figure 1.4. Cellulose Synthase from R. sphaeroides. Panel A shows the BcsA- BcsB heterodimer in grey. The BcsA subunit contains the glycosyltransferase domain and the regulatory PilZ domain, both of which reside in the cytoplasm. The transmembrane channel portion of BcsA crosses the cytoplasmic membrane (dotted lines), facilitating the translocation of the glucan polymer (teal) to the periplasmic space. Two cyclic-di-GMP molecules (red) bind to the PilZ domain. The BcsB subunit resides entirely in the periplasm save for a single transmembrane helix anchoring it in the cytoplasmic membrane. Panel B shows first six residues of the glucan polymer (teal) interacting with the amino acid side chains (magenta) of transmebrane channel of BcsA. Panel C shows the non-reducing end of the glucan polymer (replaced here with a galactose) pointing into the glycosyltransferase domain. Aspartate 343 (magenta), which is part of the finger helix domain (green), acts as the catalytic base, deprotonating the 4-carbon hydroxyl for attack on the 1- carbon of UDP-glucose (here replaced with UDP-CH2-glucose) (orange). Magnesium is shown in yellow. Note Trp383 for orientaion. (PDB Entry: 4HG6) (19)

Initiation of Cellulose Synthesis

The synthesis of glycan polymers by processive glycosyltransferases can be divided into three steps: initiation, elongation, and termination (30). The mechanism for initiation is either primer-dependent or primer-independent. Here we will define the primer-dependent mechanism as a requirement for an exogenous compound, which is not the donor substrate used in elongation, before elongation of the glycan polymer can proceed. The primer-independent mechanism requires no such additional compound, and both initiation and elongation can be achieved with only the donor substrate. The crystal structures of BcsA always show a pre-existing polymer in the transmembrane channel

(19, 109, 110). Therefore, the initiation mechanism for cellulose synthesis is still unknown and is a focus of the work presented in this thesis.

Both mechanisms for initiation have been observed in processive glycosyltransferases (112-119). Perhaps the most well characterized of the processive glycosyltransferases are glycogen synthase and starch synthase. Both glycogen and starch synthases are similar to cellulose synthase in their usage of UDP-glucose to catalyze the elongation of glucan polymers (114, 116). However, the major distinction from cellulose synthase lies in the configuration of the glycosidic linkage. The 1,4 linkages catalyzed by starch or glycogen synthase are in the α configuration (120, 121). These glucan polymers are soluble in aqueous solutions.

Glycogen synthase is an well-characterized example for the primer-dependent model of initiation. Elongation of the α-1,4-glucan polymer is achieved upon the addition of glycogen (122) or in the presence of the self-glycosylating enzyme, glycogenin (114).

An abundance of starch synthase isoforms has been isolated from maize (118,

119), rice (116, 117), wheat (123, 124), potato (125), and barley (126), and both primer- dependent and primer-independent isoforms have been identified (116-119, 125, 126).

Briefly, starch synthases are isolated by ammonium sulfate precipitation followed by dialysis and then separation by ion exchange chromatography (116, 119, 125). The fractions from ion exchange chromatography are assayed for activity in the presence and absence of either, typically, glycogen or amylopectin which serve as the exogeneous primer. Those fractions displaying activity in the absence of exogenous primers are deemed to be the primer-independent isoforms. A commonality among primer- independent isoforms appears to be a correlation of activity with citrate concentration, where increased citrate concentration is correlated to increased activity in vitro (116,

119).

Other primer-independent processive glycosyltransferases, which are responsible for synthesizing a diverse collection of glycan polymers, have been isolated (127-130). In a comparative analysis of the catalytic domains of bacterial secreted glycosyltransferases, a phylogenic tree constructed by Simpson et al. (130) found no clustering of those enzymes that follow the primer-dependent mechanism versus those enzymes classified as primer-independent. No unifying motifs are known that can be used to identify primer- dependent versus primer-independent processive glycosyltransferases.

Although the examples mentioned above appear to show glycan synthase activity in the absence of an exogenous primer, it is important to note that their activity may result from the co-purification of an endogenous primer. Indeed, in the case of starch

21 synthase, Schiefer et al. (131) found that the treatment of primer-independent starch synthase with amylase led to the abolition of activity, suggesting the presence of a co- purifying endogenous α-1,4-glucan primer. These authors posit that the observed increased activity, commensurate with increased levels of citrate in vitro, may be a result of increased affinity for such an endogenous primer (131).

Heparosan Synthase

The glycosyltransferase, heparosan synthase is an example of a primer- independent glycosyltransferase. This enzyme catalyzes the polymerization of heparosan by alternatively transferring N-acetylglucosamine and glucuronic acid from the respective donor UDP-sugar to the non-reducing end of the heparosan polymer (128).

Using purified heparosan synthase from Pasteurella multocido, Chavaroche et al. (128) found that increasing the ratio of UDP-N-glucosamine relative to UDP-glucuronic acid resulted in an increase in the rate of polymerization and favored the synthesis of low molecular weight polymers. Anion exchange chromatography identified the presence of a

UV absorbing product, suggesting that UDP remains attached to the heparosan product.

This was confirmed by MALDI-TOF analysis, showing a product containing UDP, glucuronic acid, and N-acetylglucosamine (128). The authors concluded that initiation occurs through the addition of glucuronic acid to the non-reducing end of a UDP-N- acetyl-glucosamine acceptor (128).

Cyclic Glucosyl Synthase

Another example of a primer-independent glycosyltransferase is cyclic glucan synthase (Cgs). Cgs initiates glucan synthesis by the transfer of glucose from UDP-

22 glucose to an unidentified amino acid residue of the protein (132). Elongation then occurs through the processive transfer of glucose units to the non-reducing end of the β-1,2- glucan polymer. Finally, the enzyme generates a cyclic glucan polymer consisting of 17-

22 glucose units by a transglycosylation reaction between the non-reducing end and the protein-anchored reducing end of the polymer (133-135).

Cgs bears several structural similarities to cellulose synthase, including six transmembrane helices (136). Also strikingly similar are the conserved DxD, E/D, and

QxxRW motifs and both enzymes’ requirement of Mg2+ for activity (133).

STATEMENT OF THE PROBLEM

In the absence of a robust plant system, bacterial cellulose synthases provide an accessible handle for the study of the mechanisms of cellulose synthesis. Therefore, isolation and classical in vitro biochemical characterization of a bacterial cellulose synthase are important to this goal. Here, G. hansenii was chosen as the primary model for isolation and characterization of the core catalytic subunit of the bacterial CSC.

While a great deal of progress is being made in the structural investigation of cellulose synthase, there is still much to be learned about the detailed mechanisms of cellulose synthesis. The kinetic mechanism for cellulose synthesis can minimally be divided into initiation, elongation, and termination. Understanding the kinetic mechanism of cellulose synthesis is the basis for understanding how to engineer cellulose synthase such that products of desired physical properties (DOP, crystallinity, morphology, ect.) can be synthesized.

SUMMARY

The goals of this thesis were to characterize the mechanisms of cellulose synthesis. The notable contributions of this work are:

1. The purification and characterization of the catalytic core (AcsA-AcsB) of

the cellulose synthase complex from Gluconacetobacter hansenii.

2. The identification of the post-translational cleavage site of the AcsAB

peptide.

3. Determination of initiation of cellulose synthesis to be by a primer-

independent mechanism.

4. Generation of a BcsA-BcsB (Rhodobacter sphaeroides) variant, which

alters the processivity of the enzyme.

5. Derivation of a minimal kinetic mechanism for cellulose synthesis.

6. Characterization of the roles of three CSC accessory proteins (AcsC,

AcsD, and CcpAx).

7. Determination of a whole-cell turnover number for cellulose synthesis in

G. hansenii.

In Chapter 2 we purified AcsA-AcsB from G. hansenii by product entrapment, a method that takes advantage of the insolubility of cellulose in order to isolate cellulose synthase by centrifugation. We identified the post-translational processing site of the

AcsAB peptide (resulting in the AcsA-AcsB peptide) by N-terminal sequencing. Because we were unable to remove a sufficient amount of AcsA-AcsB from cellulose, product entrapment could not be used to purify cellulose synthase for kinetic characterization. G.

24 hansenii cells were transformed with a dodecahistidine-tagged acsAB gene by our collaborator, Ying Deng. Using this transformant, we purified AcsA-AcsB by affinity chromatography and kinetically characterized the enzyme. We concluded this chapter by showing that AcsA-AcsB is the minimal catalytic core of the cellulose synthase complex

(CSC) in G. hansenii. The findings in this chapter have been published in the following journal article:

McManus JB, Deng Y, Nagachar N, Kao TH, & Tien M (2016) AcsA-AcsB: The Core of the Cellulose

Synthase Complex From Gluconacetobacter hansenii ATCC23769. Enzyme. Microb. Tech. 82:58-65.

In Chapter 3 we used purified AcsA-AcsB and BcsA-BcsB to investigate the mechanism of initiation in cellulose synthase (primer-dependent versus primer- independent). Using gel permeation chromatography (GPC) and reducing-end analysis of cellulose from the products of AcsA-AcsB and BcsA-BcsB, we showed that cellulose synthase follows a primer-independent mechanism for initiation. A comparison of the

GPC profiles of cellulose from BcsA-BcsB and AcsA-AcsB revealed processivity differences between the two enzymes. The GPC profiles also gave us a way to measure the cellulose synthesis rate, independent of enzyme concentration. We found that this rate

(called the elongation rate) is faster than the overall turnover number for both enzymes. It suggests that there may be a slow step in the overall catalytic cycle. Finally, we demonstrated BcsA-BcsB-mediated UDP-glucose hydrolase activity. We concluded this

Chapter by constructing a plausible mechanism for priming.

To investigate the factors underlying processivity, in Chapter 4, we generated variants in BcsA transmembrane channel amino acid residues, which interact with the glucan polymer. Using the GPC methods developed in Chapter 3, we showed both

25 decreased processivity and decreased kinetic rates in one of these variants. We concluded this Chapter with the construction of a minimal kinetic mechanism for cellulose synthesis.

In Chapter 5, we returned to the context of the whole cell (G. hansenii only), and measured the in vivo turnover number for AcsA-AcsB using quantitative western blotting. We found that it was faster than the turnover number for purified AcsA-AcsB and nearly identical to the elongation rate calculated in Chapter 3 for AcsA-AcsB. This suggests that a step or steps in the cellulose synthesis mechanism is more efficient in vivo. To test if there are any effects on catalysis by the actions of CSC accessory proteins, we investigated the impacts three of the CSC accessory proteins (AcsC, AcsD, and

CcpAx) had on cellulose synthesis. Each of these proteins has been shown to affect cellulose synthesis in vivo (25, 59, 90, 93). Using insertional mutants generated by our collaborator, Ying Deng, we measured the rate of cellulose synthesis and the AcsA and

AcsB proteins levels in total membrane (TM). We also examined cells of each of these mutants by transmission electron microscopy. Images showed an expanded periplasm in cells of the acsC insertional mutant. We hypothesize that this is a result of cellulose packing into the periplasmic space. It is consistent with AcsC’s proposed role as an outer membrane exit channel for cellulose. We heterologously expressed and purified AcsD and CcpAx and added them to purified AcsA-AcsB, while measuring the cellulose synthesis rate under saturating substrate conditions. We found no difference in the overall turnover number of AcsA-AcsB with or without the additions of AcsD and CcpAx. We also examined the subcellular localization of AcsA, AcsB, AcsC, and AcsD by sucrose density gradient. Though we were unable to show that the three accessory proteins

26 directly impacted catalysis, an important finding is that CcpAx is necessary for the stability of AcsB in vivo. We concluded the chapter by measuring the turnover number at each step during purification by product entrapment using quantitative western blotting.

Our results show that the turnover number drops to that calculated for purified AcsA-

AcsB immediately after cell lysis. We hypothesize that the formation of the CSC is important to the catalytic cycle for cellulose synthesis.

Part of the findings in this chapter are published in the following journal article:

McManus JB, Deng Y, Nagachar N, Kao TH, & Tien M (2016) AcsA-AcsB: The Core of the Cellulose

Synthase Complex From Gluconacetobacter hansenii ATCC23769. Enzyme. Microb. Tech. 82:58-65.

In the final Chapter, we provide a consolidated review of the findings in this thesis as well as directions for future research.

CHAPTER 2

ACSA-ACSB:

THE CATALYTIC CORE OF THE CELLULOSE SYNTHASE COMPLEX

Introduction

Cellulose is synthesized from Gluconacetobacter hansenii by large multi-protein complexes called cellulose synthase complexes (CSC) (32). The CSC proteins aggregate into a linear array on the surface of the cell (32, 90, 91) and extrude cellulose as a highly crystalline (59) ribbon-like fibril (32).

Our lab’s sequencing of the G. hansenii ATCC23769 genome has enhanced the use of this organism as a model system for cellulose synthesis (98). The catalytic subunit of the CSC in G. hansenii ATCC23769 is encoded by the gene acsAB (Fig. 1.2) (67, 98).

The AcsAB peptide contains the regulatory PilZ domain, the glycosyltransferase domain, and the periplasmic AcsB portion (25, 58, 59, 67, 96-99). Previous work performed in our lab on G. hansenii ATCC23769, using western blotting, suggested that AcsAB was post-translationally processed into three peptides (97). We named these peptides AcsAcat,

AcsAreg, and AcsB (97), and determined, based on the nucleotide sequence, that they contain the glycosyltransferase domain, the PilZ regulatory domain, and the periplasmic

AcsB portion, respectively (97). Heterotrimeric cellulose synthases in bacteria are not without precedent. X. campestris, for example, encodes cellulose synthase in three open reading frames (Fig. 1.2), called bcsA1, bcsA2, and bcsB, for the glycosyltransferase domain, the PilZ domain, and BcsB, respectively (70, 97).

One goal of this thesis is to isolate the minimal catalytic core from the CSC for the investigation of priming in cellulose synthesis. This effort began prior to the publication of the BcsA-BcsB crystal structure (19). In this chapter, we purify cellulose synthase from G. hansenii ATCC23769 by two methods. In doing so, we show that our initial finding of the processing of the AcsAB into a heterotrimer (97) was incorrect, and that AcsAB is processed into heterodimer called AcsA-AcsB. We then elucidate the post- translational processing site. We also kinetically characterize AcsA-AcsB, providing the kinetic values for KM, kcat, and the second order rate constant. Finally, we show that

AcsA-AcsB is the minimal catalytic core of the G. hansenii CSC.

Materials and Methods

Culture Conditions

G. hansenii cells expressing the dodecahistidine-tagged AcsAB were grown at

30°C in a 60 L fermenter in Schramm-Hestrin (SH) media (87) containing 20 mg/L of tetracycline to an absorbance of 1.5 at 600 nm. The cells were harvested by centrifugation. All other cultures were grown in SH media with antibiotics for the appropriate selectable marker at 30°C with shaking at 250 rpm to an absorbance of 0.6 -

0.8 at 600 nm, and the cells were harvested by centrifugation.

Cloning of AcsAB-his

A 3’ flanking fragment of AcsAB was amplified using primers (Appendix A) to

G. hansenii ATCC23769 genomic DNA. The tetC gene was amplified using primers

(Appendix A) to the pUCD2 plasmid. The two PCR fragments and BamHI/EcoRI digested pUC18 were ligated using the In-Fusion HD Cloning Kit (Clontech), and the resulting construct was introduced into wild-type G. hansenii ATCC23769 cells via electroporation. The transformants were selected on SH/tetracycline agar plates.

Protein Concentration Determination

The protein concentration of purified AcsA-AcsB was determined by absorbance at 280 nm using its molar extinction, 195,745 cm−1 M−1 (calculated from the amino acid sequence using ExPASy online software). For all other samples, the protein concentrations were determined by a modified method of Lowry et al. (137) using bovine serum albumin as a standard. In brief, 80 µL of working solution (20 parts 5% sodium carbonate, 10 parts 10% SDS, 8 parts 1 M sodium hydroxide, 1 part 2% sodium potassium tartrate, and 1 part 1% cupric sulfate pentahydrate) was mixed with 80 µL of sample, and developed with 40 µL Folin phenol reagent (diluted 1:10). Samples were analyzed using a Spectramax plate reader.

AcsA-AcsB Purification by Product Entrapment

Product entrapment, first described by Kang et al. (138), was performed as follows. All steps were performed at 4°C, unless otherwise noted. Each gram of harvested cells was resuspended in 10 mL of PMC buffer (20 mM sodium phosphate, pH

7.5, 5 mM MgCl2, 5 mM cellobiose, 1mM PMSF), and lysed in a microfluidizer two times at 20,000 psi. Total lysate was clarified by centrifugation at 2800 × g for 30 min.

Total membrane (TM) was isolated by centrifugation at 250,000 × g for 1.5 h and then each gram of TM was resuspended in 5 mL of solubilization buffer (20 mM sodium phosphate, pH 7.5, 5 mM MgCl2, 5 mM cellobiose, 10% glycerol, 2% Triton) using a glass homogenizer. After 1 h of gentle rotary shaking, the TM was subjected to centrifugation at 100,000 × g for 1 h. A supernatant volume of 6.8 mL was then mixed with 3.4 mL of 60 µM cyclic-di-GMP and 0.2 mL of 100 mM UDP-glucose. The entire mixture was layered on top of 2.6 mL of PMC buffer containing 30% glycerol. The mixture was incubated at 30°C for 11 min and then incubated on ice for 2 h. The mixture was centrifuged in a swing bucket rotor at 50,000 × g for 20 min and the entire supernatant and glycerol cushion were removed. The pellet was resuspended in 1.0 mL of resuspension buffer (20 mM phosphate, pH 7.5, 5 mM MgCl2, 5 mM cellobiose, 10% glycerol, 0.1% dodecylmaltoside) and centrifuged at 16,000 × g for 10 min. The pellet was again resuspended in 0.5 mL resuspension buffer and stored at -80°C.

AcsA-AcsB Purification by Affinity Chromatography

All steps were performed at 4°C unless otherwise noted. TM was collected and solubilized in solubilization buffer as described above for product entrapment. AcsA-

AcsB was purified from solubilized membrane with TALON cobalt-affinity resin

(Clontech). Solubilized membrane (200 mL) was incubated with 10 mL of resin for 1 h and then poured into a column. The resin was washed with 150 mL of WB (20 mM sodium phosphate, pH 7.5, 5 mM MgCl2, 5mM cellobiose, 10% glycerol, 100 mM NaCl,

20 mM imidazole, 0.1% dodecylmaltoside)1. AcsA-AcsB was eluted from the column with 100 mL of WB containing a linear gradient of 20-120 mM imidazole, followed by

50 mL of WB containing 120 mM imidazole. Fractions of 5 mL were collected. Purity of each fraction was determined by SDS-PAGE and visualization with Coomassie-blue stain. Typically, fractions 17 through 30 are combined and concentrated with a 100 kDa cut-off centrifugal concentrator to approximately 1 mL, aliquoted, and stored at -80°C.

Radiometric Measurement of Cellulose Synthase Activity

Unless otherwise noted, activity was measured at 30°C in 40 mM Tris-HCl, pH

8.0, 15 mM MgCl2, 12 μM cyclic-di- GMP, and 5 mM UDP-glucose in a total volume of

300 μL. The stock UDP-14C-glucose was diluted with non-radioactive UDP-glucose to specific activity of 3.7 × 106 Bq/mmol. Reactions were terminated by the addition of 2 mL of 1 M sodium hydroxide. Next, 20 mg of cellulose was added, and the mix was

1 After the initial purification and kinetic characterization of AcsA-AcsB, cellobiose was later excluded from the wash and elution buffers for subsequent purifications for AcsA-AcsB. Its exclusion had no apparent effect on the turnover number for AcsA-AcsB. 32 boiled for 20 min. After cooling to room temperature, 4 mL of 0.5 M HCl was added and the entire mixture was filtered through a glass filter (1.2 μm pore size). The filter was washed with 30 mL water, 10 mL methanol, allowed to dry, and was added to 5 mL

ScintiVerse (Fisher Scientific) scintillation fluid. Radioactive decay was quantified with a

Beckman Coulter LS6500 liquid scintillation counter.

Spectrophotometric Measurement of Cellulose Synthase Activity

Unless otherwise noted, activity was measured at 30°C in 40 mM Tris-HCl, pH

8.0, 15 mM MgCl2, 1 mM CaCl2, 12 µM cyclic- di-GMP, 2 mM phosphoenolpyruvate,

0.5 mM NADH, with 20 U of pyruvate kinase and 20 U lactate dehydrogenase (Sigma-

Aldrich), and 5 mM UDP-glucose in a total volume of 200 µL. The assay mix was pre- incubated at 30°C for 10 min and cellulose synthesis was initiated by the addition of

AcsA-AcsB. Activity was measured by monitoring NADH oxidation by absorbance at

340 nm (69, 139).

SDS-PAGE and Western-Blot Analysis

Protein samples (typically, 20-40 µg per lane or 0.1-1 µg per lane for pure AcsA-

AcsB) were incubated with lithium dodecyl sulfate (LDS) loading buffer (5× LDS loading buffer: 0.225 M tris-HCl, pH 6.8, 50% glycerol, 5% LDS, 0.05% bromophenol blue, 0.25 M dithiothreitol) and incubated on ice for 1 h, or with sodium dodecyl sulfate

(SDS) loading buffer (5× SDS loading buffer: 0.225 M tris-HCl, pH 6.8, 50% glycerol,

5% SDS, 0.05% bromophenol blue, 0.25 M dithiothreitol) and boiled for 10 min.

Samples were loaded onto a 12% polyacrylamide gel and run at 200 V. For western-blot analysis, the proteins were transferred to nitrocellulose at 100 V, 0.5 A for 1 h and immunostained with the indicated primary antibodies followed by a secondary alkaline phosphatase-conjugated anti-rabbit IgG (Sigma Aldrich) incubation. The blots were developed with enhanced chemifluorescence (ECF) reagent (GE Healthcare) for 0.5-2 min and visualized on a fluorescence typhoon scanner.

N-terminal Sequencing

AcsA-AcsB purified by product entrapment was separated by SDS-PAGE and visualized with Coomassie-blue stain. The band corresponding to AcsB (~100 kDa) was excised and sent for sequencing by Edman degradation at the Iowa State University

Protein Facility.

Trichloroacetic Acid-Mediated Cell Lysis

For cell lysis using trichloroacetic acid-acetone and total protein analysis, G. hansenii ATCC23769 cells were grown to an absorbance of 0.8 at 600 nm, collected by centrifugation, and washed in 50 mM phosphate buffer, pH 7.5. Approximately 35 mg of cells were lysed in 1 mL of cold 10% trichloroacetic acid in acetone (wt/vol). Total protein was collected by centrifugation and washed three times with 1 mL of cold acetone. The pellet was resuspended in 1 mL 1× LDS loading buffer, and western-blot analysis was performed as previously described.

Results

Purification of AcsA-AcsB by Product Entrapment

The method of product entrapment had been previously used for purification of both cellulose synthase (68) and callose synthase (140). First, the TM fraction from G. hansenii ATCC23769 was isolated by ultracentrifugation and solubilized in 2% Triton X-

100. Cellulose synthesis was initiated upon addition of the substrate, UDP-glucose, and the activator, cyclic-di-GMP. The enzyme, cellulose synthase, was entrapped with the newly-synthesized insoluble cellulose, which was then isolated by centrifugation. The entrapped proteins were subjected to SDS-PAGE and then visualized by staining with

Coomassie blue (Fig. 2.1A). A previous report indicated that the AcsA protein from

ATCC 53582 was no longer visible by SDS-PAGE analysis after boiling in SDS (68). As such, we compared a preparation of the entrapped protein by SDS-PAGE analysis with either SDS or LDS treatment. Boiling with 1% SDS yielded only a single band at 100 kDa. Incubating on ice with 1% LDS revealed the presence of a second band at 85 kDa

(Fig. 2.1A). LDS was chosen, as it does not precipitate on ice. To confirm that these two protein bands are products of the acsAB gene, we performed western-blot analysis using anti-AcsA1 (raised against residues 129–397 of acsAB), anti-AcsA3 (raised against a synthetic peptide of residues 581–600 of acsAB), and anti-AcsB (raised against residues

610–1550 of acsAB) (Fig. 2.1B). The preparation and specificity of these antibodies are documented (97). The western blot revealed an 85 kDa cross-reactive band when using anti-AcsA1 and anti-AcsA3. The 85 kDa band corresponds to AcsA, which contains the glycosyltransferase and regulatory PilZ domains. The 100 kDa band was cross-reactive

35 with the anti-AcsB antibody and was identified as the primarily periplasmic subunit,

AcsB. Neither visualization with Coomassie-blue stain nor western-blot analysis revealed a 169 kDa band, corresponding to an unprocessed AcsAB, suggesting that processing must have occurred quickly after translation.

Figure 2.1. Isolation of AcsAB by Product Entrapment. Panel A is an SDS- polyacrylamide gel visualized with Coomassie-blue stain showing TM (lane 1), soluble TM (lane 2), and entrapped product after either boiling in SDS (lane 3) or incubating on ice in LDS (lane 4). In Panel B, entrapped product was incubated on ice in LDS and then subjected to western-blot analysis with Anti-AcsA1 (lane 5), Anti-AcsA3 (lane 6), or Anti-AcsB (lane 7) antibodies.

In order to confirm that cleavage is not an artifact of TM isolation and solubilization, we lysed whole cells directly in 10% trichloroacetic acid-acetone. The total protein, along with purified AcsA-AcsB, was subjected to western-blot analysis using anti-AcsB (Fig. 2.2). The western blot revealed a 100 kDa band corresponding to the processed AcsB peptide and no 169 kDa band, representing unprocessed peptide, was present.

Figure 2.2. Total Protein from Whole Cells Isolated by TCA-precipitation. SDS-polyacrylamide gel visualized with Coomassie-blue stain showing TCA-acetone precipitated total protein (lane 1). Total protein (lane 2) and purified AcsA-AcsB (lane 3) were subjected to western-blot analysis using anti-AcsB antibodies, showing that AcsA-AcsB cleavage is not an artifact of protein isolation. The band corresponding to AcsB is indicated by the arrow.

Processing of AcsAB

The cleavage site for AcsA-AcsB was determined by N-terminal sequencing via

Edman degradation of the gel-purified AcsB. The first five amino acids identified were

Ala-Ser-Ala-Pro-Arg (Appendix B, Fig. B.1). Therefore, AcsA-AcsB cleavage occurs between residues 757 and 758. Figure 2.3 shows that the AQA cleavage site is also conserved among cellulose synthases in other bacteria.

G. hansenii 731 RKERVLKGTV KMVSLLALLT FASSAQAASA PRAVAA 1036 G. xylinus 1 M KMVSLIALLV FATGAQAAP. ...IAS 24 G. oboediens 1 M KMVSLIALLV FATGAQAAP. ...IAS 24 G. europaeu 1 MRPRDM KMVSLIALLV FATGAQAAP. ...VAS 29

Figure 2.3. Sequence Alignment between AcsAB and AcsB Homologs. The cleavage site is shown underlined (cleavage occurs after AQA). The other sequences are of AcsB homologs from other cellulose synthesizing bacteria. Note that all three show the methionine from the initiation codon and thus the AcsB homologs are encoded by separate genes.

Purification of AcsA-AcsB by Affinity Chromatography

The AcsA-AcsB cellulose synthase isolated by product entrapment was greater than 95% pure based on Image J analysis of samples separated by SDS-PAGE and visualized with Coomassie-blue stain. However, use of this enzyme preparation for kinetic analysis required removal of the cellulose product associated with this enzyme.

Despite efforts, we were unable to remove the cellulose product without substantial loss

38 of activity. Consequently, our collaborators, Deng et al. (58), transformed G. hansenii with an AcsAB-containing construct and through homologous recombination, generated a

C-terminal, dodecahistidine-tagged AcsB. This resulted in disruption of the downstream

AcsC gene via insertion of a tetracycline resistance marker (Fig. 2.4).

Figure 2.4. A Schematic Diagram of the Homologous Recombination used to Generate the Dodecahistidine-tagged acsAB Gene. A construct containing a homologous flanking region to the 3’ end of AcsAB was transformed into G. hansenii. Single crossover homologous recombination lead to the insertion of a dodecahistidine tag at the C-terminus of AcsB, and insertion of a tetracycline resistance gene.

To assess the impact, if any, of the presence of the histidine tag, we compared the cellulose synthase activity from TM isolated from wild type and that of the histidine- tagged transformant by measuring cellulose synthase activity by the radiometric assay

(Materials and Methods) (139). We found no difference in activity (Fig. 2.5) and therefore concluded that there are no adverse effects on activity from the presence of the histidine tag.

0 specific activity, nmol per minute per mg per minute per specific activity, nmol wild type transformant

Figure 2.5. A Comparison of Cellulose Synthase Activity in TM from Wild Type and the Histidine-tagged Transformant. TM was isolated from G. hansenii wild type cells and G. hansenii cells transformed with a dodecahistidine tag and incubated with UDP-14C-glucose. Activity was assessed by measuring incorporation of 14C-glucose into cellulose.

TM was isolated from the transformant containing the histidine-tagged acsAB gene, solubilized, and subjected to affinity chromatography with TALON cobalt resin.

Fractions from each step of the purification were assayed for cellulose synthase activity by the radiometric assay (described in Materials and Methods) (139) (Table 2.1). The total activity increased upon solubilization of the TM with detergent (Table 2.1). This finding is consistent with work by Hashimoto et al. (141), showing that cellulose synthesis activity increases upon solubilization of TM to varying degrees associated with

40 detergent type. Only a small fraction of the total activity was recovered from the total lysate (Table 2.1).

Table 2.1. Purification Table for AcsA-AcsB.*

Sample Volume Protein Specific Total (ml) (mg/ml) Activity Activity Fold Percent (nmol/min/mg) (nmol/min) Purification Recovered Lysate 600 14.78 3.76 33803 1 100 TM 120 27.40 8.17 14707 2.2 44 TM 1h solubilization 200 16.46 9.21 27645 2.5 82 Solubilized¶ 175 11.48 8.49 22292 2.3 66 Co-TALLON 1.2 1.04 656.71 1182 175 3 *Cellulose synthesis was measured by 14C-glucose incorporation as described in Materials and Methods. ¶After solubilization in detergent, the TM was then centrifuged as described in Materials and Method and then the supernatant was assayed for cellulose synthase activity.

The purified AcsA-AcsB was subjected to SDS-PAGE. The gel revealed two bands that accounted for over 90% of the total band intensity by Image J analysis (Fig.

2.6A). These two bands were confirmed by western blot to be AcsA and AcsB (Fig.

2.6B).

Both product entrapment and affinity purification, when analyzed by SDS-PAGE and visualized with Coomassie-blue stain, show no bands of similar intensity to AcsA and

AcsB corresponding to the expected molecular weights of known CSC proteins Ccpax,

AcsD, CmcAx, or AcsC (Fig. 2.7).

Figure 2.6. Purification of AcsAB by Cobalt-Affinity Chromatography. The TM was isolated as described in Materials and Methods, and AcsA-AcsB was purified by cobalt-affinity chromatography. Panel A shows SDS-polyacrylamide gels, visualized with Coomassie-blue stain, of the TM (lane 1), soluble TM (lane 2), and purified AcsA-AcsB (lane 3). Panel B shows western-blot visualization of purified AcsAB by Anti-AcsA3 (lane 4), Anti-AcsB (lane 5), or Anti-his (lane 6) antibodies.

Kinetic Characterization of AcsA-AcsB

With purified cellulose synthase preparations, the enzyme activity can be monitored in a continuous assay with coupled enzymes, as described in Materials and

Methods (Fig. 2.8). The kd for cyclic-di-GMP was determined by varying the concentration of cyclic-di-GMP at saturating UDP-glucose concentration (5 mM) and measuring the activity. As shown in Fig. 2.9B, the kd was calculated to be 0.18 ± 0.01

μM. Enzyme activity was measured at varying concentrations of UDP-glucose (Fig.

Figure 2.7. SDS-PAGE Analysis of Purified AcsA-AcsB. SDS-polyacrylamide gels visualized with Coomassie-blue stain of the TM (lane 1), soluble TM (lane 2), product- entrapped AcsA-AcsB (lane 3), and affinity-purified AcsA-AcsB (lane 4). Arrows indicate the calculated migrations of AcsC (138 kDa), CmcAx (45 kDa), AcsD (16 kDa), and CcpAx (8 kDa).

2.9A) in the presence of saturating levels of the activator molecule, cyclic-di-GMP (12

μM). A KM of 0.28 ± 0.07 mM was calculated for UDP-glucose and a kcat of 1.72 ± 0.06

−1 −1 −1 s . These values yield a kcat/KM (second order rate constant) of 6100 ± 1600 M s . To demonstrate the reproducibility of AcsA-AcsB activity, four different preparations were measured under saturating substrate conditions. The mean and standard deviation for the rate of cellulose synthesis for these four preparations was 1.60 ± 0.50 s−1.

Figure 2.8. An Enzyme-Coupled Assay for the Continuous Monitoring of Cellulose Synthase Activity. Each turnover results in release of UDP from UDP- glucose. The UDP is converted to UTP by pyruvate kinase and the pyruvate is then reduced by the NADH-utilizing enzyme lactate dehydrogenase. Enzyme activity is measured by the decrease in absorbance at 340 nm associated with NADH oxidation.

2.0 1.8 A B 1.6 1.4

-1 1.2 1.0 0.8

Velocity, s Velocity, 0.6 0.4 0.2 0.0 0 2 4 6 8 10 0 2 4 6 8 10 12 UDP-Glucose, mM cyclic-di-GMP, M

Figure 2.9. Steady-State Kinetic Analysis of AcsA-AcsB. Panel A: effect of UDP- glucose concentration on enzyme velocity. Panel B: effect of cyclic-di-GMP concentration on enzyme velocity. Reaction rates were measured in enzyme-coupled assays by monitoring decrease in absorbance at 340 nm associated with NADH oxidation as described in Materials and Methods. Rates are expressed as velocity (s−1). Data shown is from mean and standard deviation of three separate assays.

Discussion

AcsA-AcsB is the Catalytic Core of Cellulose Synthase

The only protein to co-purify with AcsB-his was AcsA and only AcsA-AcsB came down with the cellulose product during product entrapment. Other proteins found to be essential to cellulose synthesis in vivo: AcsC, CmcAx, AcsD, and CcpAx, (25, 60, 90,

104, 108, 142) were not found by SDS-PAGE analysis (Fig. 2.7). This does not preclude these proteins from being part of the bacterial CSC; however, it suggests that any association into a complex must be relatively weak. Our results show that the minimal components required for cellulose synthesis are AcsA and AcsB. This is in accord with the findings of Du et al. (99) for AcsA-AcsB and Omadjela et al. (69) for BcsA-BcsB, cellulose synthase from Rhodobacter sphaeroides.

Processing of AcsAB

Previous work by Lin et al. (68), using product entrapment on solubilized membrane from G. hansenii ATCC53582, confirmed that AcsA-AcsB is post- translationally modified to yield AcsA and AcsB of molecular masses 83 kDa and 93 kDa, respectively. Using similar entrapment methods to isolate AcsA-AcsB, our work here with G. hansenii ATCC23769, showed that cleavage occurs between residues 757 and 758. These residues are on the carboxyl side of an AQA amino acid triplet.

Examination of BcsB genes from closely-related bacteria reveal that they encode a conserved AQA motif that is preceded by periplasmic targeting sequences (143). AxA

45 sequences are common signal peptidase motifs found in Gram-negative bacteria with cleavage occurring on the C-terminal side of the amino acid sequence (144). Cleavage after an AxA motif suggests that a signal peptidase (signal peptidases are found on the periplasmic side of the cytoplasmic membrane) (144) is responsible. The present results for AcsA are in contrast to our previous work showing that AcsA is further processed into two smaller fragments of 46 kDa and 34 kDa (97). The weakness and variability of these two smaller bands on western blots was the result of AcsA being unstable to boiling in the presence of SDS. When LDS was used as a denaturant under lower temperature conditions, as per Lin and Brown (68), we were then able to detect the 83 kDa band associated with AcsA.

The cleavage site determined in this study occurs upstream of the site identified for ATCC53582 by Saxena et al. (145). The difference in the cleavage sites may be due to the fact that we used a different strain, ATCC23769, in this work. However, another possible explanation is that when these workers isolated the membrane fraction, they employed a step with trypsin treatment prior to entrapment (145). This is consistent with their finding that the cleavage site is located at a lysine residue (145).

Kinetic Characterization of AcsA-AcsB

−1 The kcat value we calculated of 1.72 s was confirmed by the value published by

Du et al. (99) for AcsA-AcsB. These values, however, are lower than the kcat value published for BcsA-BcsB from R. sphaeroides by Omadjela et al. (69) of ~90 s-1.

However, another publication reports a velocity for BcsA-BcsB of ~0.17 s-1 when incubated in the presence of 1 mM UDP-glucose (146).

The KM value for UDP-glucose, determined in this report to be 0.28 mM, is in close agreement with the values previously determined for purified BcsA-BcsB (69) and for the TM fraction from G. hansenii ATCC53582 (147), of 0.5 mM and 0.21 mM, respectively. These values are close to the physiological concentrations of 1–2 mM for

UDP-glucose (148). The Kd value of 0.18 μM determined here for cyclic-di-GMP is an order of magnitude smaller than the value of 1.8 μM, determined for R. sphaeroides

BcsA-BcsB (69). This is nonetheless similar with physiological concentrations of 0.2–2.5

μM for cyclic-di-GMP (149) in Gram-negative bacteria.

In this chapter, we described the purification of AcsA-AcsB by two methods, product entrapment and affinity chromatography. In doing so, we verified that AcsA-

AcsB is the minimal catalytic core of the CSC. We have also shown that the nascent

AcsAB peptide is post-translationally processed into the active AcsA-AcsB heterodimer.

Finally, purification facilitated the kinetic characterization the AcsA-AcsB. With an in vitro system characterized and in hand, we are able to address the primer requirements of cellulose synthase and, along with BcsA-BcsB (R. sphaeroides), investigate processivity in cellulose synthase. Priming and processivity are the focus of Chapters 3 and 4. In

Chapter 5, we will return to the whole cell to explore the influence of other CSC proteins on cellulose synthesis.

CHAPTER 3

THE INITIATION OF CELLULOSE SYNTHESIS

Introduction

Cellulose synthesis can be divided into three steps: initiation, elongation, and termination (30, 150). Elongation of the cellulose polymer occurs through iterative rounds of substrate binding, glycosidic bond formation, UDP release, and polymer translocation (58, 110, 147). Recently, much regarding the mechanism of elongation has been learned through structural studies (19, 109, 110, 151). However, little is known about cellulose initiation and termination. In Chapter 2, we purified and characterized

AcsA-AcsB. We also determined that AcsA-AcsB is the minimal catalytic core of the G. hansenii CSC. In this chapter, we will investigate initiation of cellulose synthesis using purified AcsA-AcsB (G. hansenii) and BcsA-BcsB (R. sphaeroides).

There are two possible mechanisms for the initiation of cellulose synthesis: the primer-dependent mechanism and the primer-independent mechanism (16, 114, 119). In the primer-dependent mechanism, a compound that is not UDP-glucose is required for initiation. In the primer-independent mechanism both initiation and elongation can proceed with UDP-glucose alone.

Attempts to elucidate the mechanism of initiation for cellulose synthesis have been made using membrane extracts from the bacterial system (G. hansenii) and the plant

48 system (cotton). Cooper et al. (152) showed increased cellulose synthase activity in total membrane (TM) isolated from G. hansenii upon the addition of cellodextrins, suggesting that the cellodextrins function as primers. Because this work was performed with bacterial membrane extracts, an alternative interpretation may be that the cellodextrins act as substrates or inhibitors (in the case of cellobiose) for native glucanases, such as

CmcAx (107), thereby increasing the apparent yield of cellulose synthesized by TM.

In 2002, Peng et al. (153) showed sitosterol-β-glucoside as a requirement for the initiation of cellulose synthesis. Hydrophobic glycosides have been shown to function as primers in glycan polymer synthesis (154, 155). The work by Peng et al. (153) was performed using TM extracts from cotton. The authors show the synthesis of sitosterol-

14C-glucan (up to sitosterol-glucotetraose) when cotton membrane extracts were incubated with UDP-14C-glucose. The authors also show the production of sitosterol- glucotriose using yeast membrane extracts expressing a cellulose synthase isoform

(CesA1) from cotton (153). However, these workers were unable to show synthesis of longer oligosaccharides (i.e. cellulose) using the same yeast membrane extracts. Despite these results, it is unlikely that the narrowness and specificity of the transmembrane channel, revealed in the BcsA-BcsB crystal structure (19), could accommodate the passage of a bulky sitosterol group.

The crystal structure of the heterologously-expressed cellulose synthase shows the presence of a pre-existing cellulose polymer in the transmembrane channel of BcsA (19,

109, 110). Thus, studies to determine involvement of primers would not be straight forward. Although we have made efforts to generate enzyme without a pre-existing

49 polymer, none have yet been successful. Therefore, we used product analysis to investigate the priming mechanism for cellulose synthase.

In this chapter, gel permeation chromatography (GPC) is used to measure the degree of polymerization (DOP), or the length, of cellulose synthesized from AcsA-AcsB and BcsA-BcsB. GPC analysis of cellulose synthesized over a time course, by both enzymes, shows that the initiation of cellulose synthesis follows the primer-independent mechanism. Our GPC analysis reveals processivity differences between both enzymes, a characteristic that will be explored further in Chapter 4. We also show the production of new reducing ends from our in vitro incubations. Finally, we provide a plausible mechanism of self-priming by demonstrating BcsA-BcsB-mediated UDP-glucose hydrolase activity and propose a mechanism for initiation.

Materials and Methods

Preparation of solvents

N, N-dimethylacetamide (DMAc), methanol, and pyridine were treated overnight with, and then stored in, molecular sieves (4A). Lithium chloride was baked overnight at

500°C and cooled in a desiccator. DMAc containing 8% lithium chloride was prepared by swiftly weighing out 80 g of lithium chloride and adding it to 1 L of N, N- dimethylacetamide, stirring at 50°C. The solvent was filtered through a 0.2 µm nylon filter and stored under dry argon gas.

Expression and Purification of BcsA-BcsB

The BcsA-BcsB expression plasmid was a kind gift from Jochen Zimmer. BcsA-

BcsB was expressed heterologously in E. coli (Rosetta strain) and purified by affinity chromatography as described by Morgan et al. (19)2. Protein concentration was determined by absorbance at 280 nm using a molar extinction coefficient of 161,925 M-1 cm-1 (ExPASy online software). Aliquots of BcsA-BcsB were stored at -80°C.

Modification and Gel Permeation Chromatography of Cellulose in Tetrahydrofuran

Cellulose carbanilation was carried out by a modified method of Henniges et al.

(156). A volume of 100 μL of phenyl isocyanate in 500 µL of pyridine was added to 1 mg of cellulose under a stream of dry argon gas. Samples were sealed in ampoules and rotated slowly at 70°C for 48 hours. Unreacted phenyl isocyanate was quenched with 1.2 mL of methanol. Samples were dried overnight at 40°C under a stream of dry argon gas, dissolved in tetrahydrofuran, and filtered through 0.2 µm nylon filters (Millipore).

Injections of 200 µL were applied onto a KD-806M size-exclusion HPLC column

(Shodex) at a flow rate of 0.5 mL/min tetrahydrofuran. Molecular weight determination using multi-angle laser light scattering (MALLS) was performed with a DAWN HELIOS

2 After the initial purification and kinetic characterization of BcsA-BcsB, cellobiose was later excluded from the wash and elution buffers for subsequent purifications of BcsA-BcsB. Its exclusion had no apparent effect on the turnover number for BcsA-BcsB. 51

II (Wyatt Technologies) MALLS detector coupled with a t-REx refractometer (Wyatt

Technologies).

Dissolution and Gel Permeation Chromatography of Cellulose in Dimethylacetamide /

8% Lithium Chloride

Cellulose was synthesized as described in Chapter 2 using 5 mM UDP-14C- glucose (specific activity 4.88 x 106 Bq/mmol) in a total volume of 1 mL. All reactions were quenched by the addition of 250 µL of 10% SDS. The cellulose was collected by centrifugation and washed two more times with 1 mL of water each. The cellulose was solubilized by five exchanges of 1 mL of methanol, tumbling gently for 30 minutes each, followed by five exchanges of 1 mL DMAc, tumbling gently for 30 minutes each. The cellulose was dissolved in 300 µL of DMAc containing 8% lithium chloride (wt/vol) for

18 hours, tumbling gently (157). Cellulose solutions were filtered through 0.2 µm nylon filters (Millipore) and stored in a desiccator.

Injections of 100 µL were applied onto a KD-806M size exclusion HPLC column

(Shodex) at a flow rate of 0.1 mL/min in DMAc / 8% lithium chloride. Fractions of 0.5 mL were collected and added to 5 mL of ScintiVerse (Fischer Scientific) scintillation fluid. Radioactive decay was quantified with a Beckman Coulter LS6500 liquid scintillation counter.

Calibration was carried out using Avicel (Sigma-Aldrich) and bacterial cellulose from G. hansenii. The cellulose samples were dissolved in DMAc / 8% lithium chloride

52 as described above. Injections of 100 µL containing approximately 100 µg of cellulose were applied onto a KD-806M size exclusion HPLC column coupled to a DAWN

HELIOS II MALLS detector coupled with a t-REx refractometer. The DOP distribution and retention time of each sample was used to calibrate the KD-806M column (Appendix

B, Fig B.4).

Reducing End Modification and Analysis

Cellulose was synthesized for 2 h, as described in Chapter 2, using 1 mM UDP-

14C-glucose (specific activity 9.73 x 107 Bq/mmol) in a total of volume of 1 mL. All reactions were quenched by the addition of 250 µL 10% SDS. The cellulose was collected by centrifugation and washed two times with 1 mL of water followed by two more washes with 1 mL of DMSO. Cellulose was resuspended in 700 µL DMSO and 300

µL acetic acid. To this reaction mixture was added 46 mg of 2-amino benzamide (2AB).

After the reaction mixture reached 60°C, 64 mg of sodium cyanoborohydride was added

(158). The reaction mixture was tumbled gently at 60°C for 12 hours.

The modified cellulose was collected by centrifugation and washed with 1 mL

DMSO, two times with 1 mL water, once with 1 mL ethanol, and once with 1 mL sterile water. The modified cellulose was resuspended in a sterile-filtered (0.2 µm filter) mixture of 300 µL of 50 mM sodium acetate buffer, pH 5.0, containing 200 U of cellulase

(Worthington) and incubated at 37°C for 24 hours. Following digestion, the mixture was passed through a 3 kDa cut-off spin filter and the entire sample was injected onto a C18 reverse phase HPLC column (Supelco) at a flow rate of 0.5 mL/min. After approximately

9 mL of 10 mM sodium acetate, pH 5.0, had passed through the column, the mobile phase was changed to acetonitrile in 10 mM sodium acetate, pH 5.0 (1:9). Twenty fractions of 0.5 mL were collected and added to 5 mL of ScintiVerse (Fischer Scientific) scintillation fluid. Radioactive decay was quantified with a Beckman Coulter LS6500 liquid scintillation counter.

Glucose Quantification by Enzyme-Linked Assay

Cellulose was synthesized using 3 µM BcsA-BcsB with 5 mM UDP-glucose in a total volume of 300 µL as described in Chapter 2. At the indicated time points, the reaction mixture was passed through a 3 kDa cut-off filter to remove protein and cellulose.

Glucose was quantified by a modified procedure of Tsuge et al. (159). An enzyme master mixture of 100 µL containing 100 mM sodium phosphate, pH 6.5 (purged with pure oxygen for 15 min), 0.01% o-dianisidine, 30 U of glucose oxidase (Sigma-Aldrich) and 10 U of horseradish peroxidase (Sigma-Aldrich) was added to 100 µL of sample to a total final volume of 200 µL. Incubations were at 30°C. The free glucose is converted by glucose oxidase to gluconic acid, releasing hydrogen peroxide which is used by horseradish peroxidase to oxidize o-dianisidine. The oxidation of o-dianisidine is measured by an increase in absorbance at 460 nm and quantified using a molar extinction coefficient of 11,300 M-1 cm-1. The oxidation of one glucose molecule results in the oxidation of two o-dianisidine molecules. Two oxidized o-dianisidine molecules dimerize to form the chromophore. Therefore, one mole of chromophore is produced for every one

54 mole of glucose oxidized. The incubation continued until a cessation in the increase of absorbance, thus ensuring full conversion of glucose.

Gel Permeation Chromatography of Glucose and Cellobiose

Cellulose was synthesized using 3 µM BcsA-BcsB with 5 mM UDP-14C-glucose

(specific activity 4.88 x 106 Bq/mmol) as described in Chapter 2. The cellulose was collected by centrifugation, the supernatant was passed over 1 mL of DEAE-cellulose

(Sigma Aldrich), and the resin was washed with 2 mL of water to remove unreacted

UDP-glucose. The flow-through was dried under reduced pressure and the resulting residue was dissolved in 500 µL of water. The sample was loaded onto a 145 cm by 1.5 cm column packed with P-2 Biogel resin (Sigma). Separation of the analytes was carried out in water, collecting fractions of 2 mL. Samples of 0.5 mL were added to 5 mL

ScintiVerse (Fischer) scintillation fluid. Radioactive decay was quantified with a

Beckman Coulter LS6500 liquid scintillation counter.

Reducing End Quantification Using Bicinchoninic Acid

Stock solutions were prepared and stored in the dark for up to three months as follows:

Solution A Solution B 0.512 M Na2CO3 5.0 mM CuSO4·5 H2O 0.288 M NaHCO3 12.0 mM L-serine 5.0 mM Bicinchoninic acid

Solutions A and B were mixed in a 1:1 ratio immediately prior to assaying to create a working solution. Volumes of 0.4 mL of working solution were added to 0.1 mL of sample and vortexed. Samples were developed at 80°C for 30 minutes, cooled in a water bath at 25°C for 10 minutes, and measured by absorbance at 560 nm (52).

Results

Kinetic Characterization of BcsA-BcsB

Purified BcsA-BcsB yielded two major bands when analyzed by SDS-PAGE and visualized by Coomassie-blue stain. The molecular weights of the bands were approximately 85 kDa and 90 kDa, corresponding to BcsB and BcsA, respectively (69)

(Appendix B, Fig. B.2). Image J analysis indicated that BcsA and BcsB accounted for over 97% of the band intensity when analyzed by SDS-PAGE and visualized with

Coomassie-blue stain. The steady-state kinetic parameters for BcsA-BcsB were determined by the enzyme-linked coupled assay for UDP described for AcsA-AcsB in

Chapter 2. Enzyme activity was measured at varying concentrations of UDP-glucose

(Appendix B, Fig. B.3) in the presence of saturating levels of the activator molecule, cyclic-di-GMP (10 μM). A KM of 0.83 ± 0.04 mM was calculated for UDP-glucose and a

−1 kcat of 0.51 ± 0.01 s was determined. These values yield a kcat/KM (second order rate constant) of 614 ± 33 M−1 s−1. To demonstrate the reproducibility of BcsA-BcsB activity, four different preparations were measured under saturating substrate conditions. The mean and standard deviation for the rate of cellulose synthesis for these four preparations was 0.50 ± 0.16 s−1.

Gel Permeation Chromatography of Cellulose Tricarbanilates

Fig. 3.1 shows GPC profiles of cotton, Avicel, and cellulose synthesized from

AcsA-AcsB and BcsA-BcsB. Each sample was modified and dissolved in THF as described in Materials and Methods. The detection of cellulose by differential refractive index was coupled with MALLS. This provided information on the molecular weight independent of column calibration. A specific refractive index (dn/dc) for cellulose tricarbanilates in THF of 0.169 mL g-1 (55) was used to calculate the molecular weight, which was then converted to DOP. The DOP values were obtained by dividing the polymer molecular weights by the molecular weight of an anhydroglucose tricarbanilate

(518 g/mol).

These results show cotton is the largest with a DOP of 7,800 at the peak, consistent with the published values in the range of 1,700-19,000 (160, 161). Eluting last is Avicel, with a DOP of 200 at the peak, consistent with the published values of 100-

300 (162). Eluting between the cotton and Avicel samples are cellulose synthesized by

AcsA-AcsB and BcsA-BcsB, with DOP values of 2,900 and 1,600 at the peaks, respectively. The large increase in refractive index in each of the chromatograms at an elution volume of approximately 10 mL is due to unreacted modifying reagent.

Gel Permeation Chromatography of Cellulose Synthesis Time Course

Due to the difficulty in detecting shorter glucan polymers and the requirement of relatively larger amounts of cellulose for modification, synthesis using UDP-14C-glucose

57 and direct dissolution in DMAc / 8% LiCl was employed. This method decouples detection from MALLS. Cellulose synthesized by AcsA-AcsB and BcsA-BcsB for varying lengths of time (Fig. 3.2) was dissolved in DMAc / 8% LiCl and subjected to

GPC analysis.

Figure 3.1. GPC Elution Profiles of Cellulose Tricarbanilates. Cellulose samples (as labeled in the figure) were modified as described in Materials and Methods. For cellulose synthesized from BcsA-BcsB or AcsA- AcsB, 0.5 µM of enzyme with 5 mM UDP-glucose in a reaction volume of 1.0 ml was used as described in Materials and Methods. Reactions were allowed to proceed for 12 h. A KD-806M column was used with tetrahydrofuran as the mobile phase at a flow rate of 0.5 ml/min. The figure shows the molecular weight distribution, converted to DOP on the right axis, above the associated refractive index detection signal.

In order to calibrate the GPC column for DOP calculation, Avicel and bacterial cellulose (from G. hansenii cultures) were dissolved in DMAc / 8% LiCl and subjected to

GPC analysis coupled with MALLS and refractive index detection (Appendix B, Fig.

B.4). A specific refractive index (dn/dc) for cellulose in DMAc / 8% LiCl of 0.0575 mL

58 g-1 (163) was used to calculate the molecular weight which was then converted to DOP.

The DOP values were determined by dividing the polymer molecular weights by the molecular weight of an anhydroglucose (162 g/mol). The DOP distribution for each of these samples was transcribed onto the GPC elution profiles of the cellulose synthase time courses and used to generate a calibration curve (Fig. 3.2, right axis).

Cellulose synthesized from BcsA-BcsB for 2-30 minutes increased in size, eventually reaching a DOP upper limit of 11,700. From 60-240 minutes, the maximum size did not increase, however, the overall incorporation of glucose continued (Fig.

3.3A), consistent with the synthesis of new polymers (Fig 3.2A). The same trend

(increase in the maximum DOP followed by an increase in the overall glucose incorporated) was observed for AcsA-AcsB (Fig. 3.2B and Fig. 3.3B) except that the maximum DOP was 23,100. Thus, both enzymes exhibited a processivity limit. The DOP at the peak of the elution profiles for AcsA-AcsB and BcsA-BcsB were 3,000 and 1,500, respectively. These values are nearly identical to those determined by carbanilation and

GPC analysis in THF.

Examination of the DOP at the peak of the two-minute time point elution profiles gave us a means of calculating the turnover number for both AcsA-AcsB and BcsA-BcsB independent of enzyme concentration. For BcsA-BcsB (DOP 800) and AcsA-AcsB (DOP

3,000), turnover numbers of 6.3 s-1 and 25 s-1 were calculated, respectively. These numbers approximate the rates of polymer elongation and are higher than the turnover numbers determined for BcsA-BcsB of 0.5 s-1 and for AcsA-AcsB of 1.7 s-1 by steady- state kinetic analysis.

500 10000 A 400 1000

300 240 min 120 min 100 60 min 200 30 min 10 min 10 5 min 100 2 min 1

0 0.1 300 B 10000 250 1000

200 Polymerization of Degree 100

Cellulose (nmol glucose)Cellulose (nmol 150 10 100 1 50

0 0.1 4 6 8 10 12 Elution Volume, ml

Figure 3.2. GPC Elution Profile for Cellulose from BcsA-BcsB (A) and AcsA- AcsB (B). Aliquots were removed from enzymatic reaction mixtures at times as specified in the figures. Cellulose was synthesized from 0.5 µM BcsA-BcsB or AcsA- AcsB with 5 mM UDP-14C-glucose in a reaction volume of 1.0 ml as described in Materials and Methods. The underivatized cellulose was dissolved in DMAc / 8% LiCl as described in Materials and Methods. Fractions (0.5 ml) were collected and quantified by scintillation spectroscopy. The degree of polymerization, measured by MALLS for bacterial cellulose and Avicel is shown as bold lines above the elution profiles and is measured on the right axis. A calibration curve is drawn through these two lines.

1400

1200 A B

1000

800

600

400

Cellulose (nmol glucose) (nmol Cellulose 200

0 0 50 100 150 200 250 3000 50 100 150 200 250 300 Time, min

Figure 3.3. Total Glucose Incorporated into Cellulose from (A) BcsA-BcsB and (B) AcsA- AcsB. The moles of glucose from each fraction in the elution profiles in Figure 3.2 were added together to determine the total amount of glucose incorporated into cellulose at each time point.

The continual appearance of small polymers over time is consistent with new polymer synthesis. This, in turn, would be consistent with primer-independent initiation of synthesis. However, it may also be attributed to contaminating glucanase activity. To test for the presence of such activity, we synthesized cellulose with UDP-14C-glucose using BcsA-BcsB. We isolated the cellulose and incubated it with either buffer, AcsA-

AcsB, or BcsA-BcsB for 16 hours, then analyzed the cellulose by GPC. We found no difference in the degree of polymerization for any incubation (Appendix B, Fig. B.5) indicating that neither enzyme preparation contained contaminating glucanase activity.

The difference between the elongation rate and the overall turnover number may be attributed to a large fraction of inactive cellulose synthase in the synthesis incubation mixtures. To test for this, we performed product entrapment (Chapter 2) on purified preparations of both enzymes. We reasoned that only active enzyme will be entrapped and thus found in the pellet. Here, 0.5 µM cellulose synthase was incubated for 30

61 minutes with 5 mM UDP-glucose in a total volume of 0.5 mL. Cellulose was collected by centrifugation for 15 min at 16,000 × g and 4°C. To confirm that enzyme does not pellet without synthesizing cellulose, an incubation containing no UDP-glucose was prepared.

To ensure that inactive enzyme does not pellet with cellulose and active enzyme, an incubation mixture, quenched with 100 mM EDTA after 30 min, was subsequently spiked with an additional 0.5 µM of enzyme. To the supernatant (final volume of 600 µL) of each sample was added 150 µL 5 × LDS sample buffer (Chapter 2) and the pellet was taken up in 750 µL of 1× LDS buffer. Identical volumes of each sample were analyzed by western blot (Fig. 3.4). All band intensities were quantified using Image J software.

For AcsA-AcsB, we found that the pellet contained 48% of the combined supernatant and pellet band intensities (Fig 3.4A, lane 1). No enzyme was found in the pellet of the incubation mixture containing no UDP-glucose (Fig 3.4A, lane 2). No increase in the amount of pelleted enzyme was observed with samples spiked with additional enzyme at the end of the incubation. (Fig 3.4A, lane 3). The second two results confirm that cellulose itself does not pellet enzyme. Rather, only enzyme which has synthesized cellulose is pelleted.

For BcsA-BcsB we found that the pellet contained 84% of the combined supernatant and pellet band intensities (Fig 3.4B, lane 1). A small amount of enzyme was found in the pellet of the incubation mixture containing no UDP-glucose (17% of the combined supernatant and pellet band intensities) (Fig 3.4B, lane 2). No increase in the amount of pelleted enzyme was observed with samples spiked with additional enzyme at the end of the incubation (Fig 3.4B, lane 3).

Figure 3.4. Product Entrapment of Purified AcsA-AcsB (panel A) and BcsA-BcsB (panel B). An amount of 0.25 nmol of either AcsA-AcsB or BcsA-BcsB was incubated with 2.5 μmol UDP-glucose (1 and 3) or 0 μmol UDP-glucose (2) at 30°C for 30 minutes in a total volume of 0.5 mL. After 30 minutes, either 0 nmol (1 and 2) or 0.25 nmol of AcsA-AcsB (panel A) or BcsA-BcsB (panel B) with 50 µmol EDTA (3) was added to the reaction and cellulose was collected by centrifugation at 16,000 × g for 15 minutes. The supernatants were removed and pellets were resuspended in the same volume as the supernatants. Identical volumes of each supernatant (back bars) and pellet (grey bars) were analyzed by western blotting using rabbit anti-histidine antibody and band intensities were quantified by Image J software.

Elongation Rate

To further characterize the elongation rate, we measured the size distribution of cellulose produced at varying substrate concentrations by GPC. Here, both AcsA-AcsB and BcsA-BcsB were characterized. Elongation rates were measured at varying UDP- glucose concentrations. Cellulose at each incubation was collected, solubilized, and separated by GPC as described in Materials and Methods (Fig. 3.5A and B). To improve resolution, fractions of 0.25 mL were collected and measured instead of fractions of 0.5 mL. The elongation rates were calculated, as before, from the DOP at the peak for each

63 elution profile. The elongation rates were fit to the Michaelis-Menten equation using a

-1 -1 nonlinear regression (Fig. 3.5B and D). This yielded a kcat of 4.1 s and 25 s , for BcsA-

BcsB and AcsA-AcsB, respectively. The KM for UDP-glucose was 0.29 mM and 0.06 mM for BcsA-BcsB, and AcsA-AcsB, respectively. The second order rate constant was

14,000 M-1 s-1 and 417,000 M-1 s-1 for BcsA-BcsB and AcsA-AcsB, respectively. The second order rate constants for elongation of the glucan polymer by both enzymes are greater than that for the overall catalytic cycle (600 M-1 s-1 and 6,100 M-1 s-1 for BcsA-

BcsB and AcsA-AcsB, respectively). This indicates that the elongation of the glucan polymer is more efficient than the overall catalytic cycle.

Reducing End Analysis

Further evidence for a primer-independent mechanism came from reducing end analysis. During cellulose synthesis, the 4-carbon hydroxyl of the non-reducing end attacks the 1-carbon of the UDP-glucose, while the reducing end of the polymer is pushed out into the extracellular milieu. To investigate the synthesis of new reducing ends (and therefore new cellulose polymers), cellulose was synthesized by BcsA-BcsB and AcsA-AcsB using UDP-14C-glucose and modified with 2-aminobenzamide (2AB).

During modification, the amine of 2AB forms a Schiff base with the 1-carbon of the reducing end of cellulose. This bond is made permanent by reduction with sodium cyanoborohydride (158). The 14C-labeled cellulose was digested with cellulases and the products were separated by reverse-phase HPLC, which facilitated the separation of glucose from 2AB-glucose. Prior to injection, non-radiolabeled 2AB-glucose was added to each sample and detection was carried out by fluorescence at 420 nm.

Figure 3.5. GPC Elution Profiles of Cellulose from AcsA-AcsB (A) and BcsA- BcsB (C), and Michaelis-Menten Curves for the AcsA-AcsB (B) and BcsA- BcsB (D) Elongation Rates. Cellulose was synthesized from 0.5 µM enzyme with varying concentrations of UDP-14C-glucose in a reaction volume of 1.0 ml for two minutes (AcsA-AcsB) and four minutes (BcsA-BcsB) as described in Materials and Methods. (A and C) Cellulose was collected, solubilized, and separated by GPC as described in Materials and Methods. (B and D) Effect of UDP-glucose concentration on elongation rate. Elongation rates were measured and reported as velocity (s−1).

A major radioactive peak, corresponding to glucose (Fig. 3.6), elutes at approximately 2.5 mL. A radioactive peak, co-eluting at approximately 13 mL with the internal 2AB-glucose standard (Fig. 3.6), is present. The second fluorescent peak is a result of unreacted 2AB. The presence of radio-labeled reducing ends is a clear indicator of the primer-independent mechanism in which glucose, from UDP-glucose, is initiating polymer synthesis. The appearance of new reducing ends may also be a byproduct of the

65 modification procedure. To test for this, we synthesized cellulose with UDP-14C-glucose using BcsA-BcsB. We either solubilized the cellulose immediately after synthesis or modified it with AB and then solubilized the cellulose. We analyzed both samples by

GPC. We found no difference in the degree of polymerization for either sample

(Appendix B, Fig. B.6) indicating that modification with AB is not a source of new reducing ends.

8 x 105 CPM Flourescence 1.0

0.8 300 0.6

200 0.4

100 0.2

0 0.0 5 CPM 6 x 10 Flourescence 1.0

0.8

Radioactivity, CPM Radioactivity,

Fluorescence, Relative 300 0.6

200 0.4

100 0.2

0 0.0 0 5 10 15 20 Elution volume, ml

Figure 3.6. Reducing End Analysis of Cellulose by Reverse Phase HPLC. Cellulose was synthesized by (panel A) AcsA-AcsB- or (panel B) BcsA-BcsB-containing reaction mixtures containing 1 mM UDP-14C-glucose and 0.5 µM enzyme in a total volume of 1.0 ml and then was modified with AB2, digested with cellulase and separated in 0 or 10% acetonitrile as described in Materials and Methods. Fractions of 0.5 mL were collected and quantified by scintillation spectroscopy. An internal standard of AB2- modified glucose was measured by fluorescence (emission at 420 nm). Note the split axis in the figure.

BcsA-BcsB-Mediated UDP-glucose Hydrolysis

Based on the detection of 14C-labeled reducing ends, we reasoned that glucose, derived from UDP-glucose hydrolysis, may be initiating new polymer synthesis. In order to detect cellulose synthase-mediated UDP-glucose hydrolysis, we utilized glucose oxidase. In this assay, free glucose is oxidized by glucose oxidase to yield hydrogen peroxide and gluconic acid. Horseradish peroxidase, using hydrogen peroxide, oxidizes o-dianisidine. The oxidation of o-dianisidine is measured by an increase in absorbance at

460 nm.

2.5 20

2.0 15

M 1.5 UDP, (+)cyclic-di-GMP UDP, (-)cyclic-di-GMP 10 glucose, (+)cyclic-di-GMP 1.0 glucose, (-)cyclic-di-GMP

UDP, UDP, mM

glucose, 5 0.5

0.0 0 0 50 100 150 200 250 300 350

Time, minutes

Figure 3.87 BcsA-BcsB-Dependent Generation of Glucose from UDP- glucose. Incubations contained 3 µM BcsA-BcsB and 5 mM UDP-glucose in a total volume of 0.3 ml as described in Materials and Methods with (filled markers) or without (open markers) 10µM cyclic-di-GMP.

Due to interference from light scattering by cellulose, a continuous assay was not possible. Instead, we took time points, removing cellulose and protein by filtering incubations through a 3 kDa cut-off centrifugal filter before measuring glucose. In the presence of cyclic-di-GMP and BcsA-BcsB, free glucose increased over time. The pattern (but not net amount) is identical to the increase in free UDP (Fig. 3.7, closed symbols). In the absence of cyclic-di-GMP, the rates of UDP and glucose liberation at 10 minutes were 1.2% and 10% that of the rates of liberation in the presence of cyclic-di-

GMP, respectively (Fig 3.7, open symbols). No glucose was detectable when this assay was performed for AcsA-AcsB.

The detection of free glucose led us to ask whether larger oligomers were also released from the enzyme. Cellodextrins are oligomers of two to seven β-1,4 linked glucose units. Cellodextrins of six glucose units or less are varyingly soluble in water

(164). In order to test for the presence of soluble cellodextrins generated by BcsA-BcsB, the soluble fraction of the reaction mixture was analyzed by size-exclusion chromatography. Following synthesis, the supernatant was passed over DEAE-cellulose to remove unreacted UDP-14C-glucose. The remaining products were separated by size- exclusion chromatography with internal standards for glucose (Fig. 3.8, eluting at 190 mL) and cellobiose (Fig. 3.8, eluting at 174 mL). Our results showed only the presence of glucose co-eluting with the glucose standard. No peak is present for cellobiose (Fig. 3.8).

The larger species, eluting at 158 mL, is calculated to have a molecular weight of 467 g/mol based on the elution peaks of glucose and cellobiose. This peak likely corresponds to remaining UDP-glucose, whose molecular weight is 566 g/mol.

6 1.0

5 soluble products from BcsA-BcsB Internal Standards 0.8 4 0.6 3 0.4 2 0.2 glucose, nmol 1

Absorbance, 560 Absorbance,nm 560 0 0.0

0 50 100 150 200 Elution Volume, mL

Figure 3.8. BcsA-BcsB Soluble Product Analysis by Gel Permeation Chromatography. Cellulose was synthesized by reaction mixtures containing 5 mM UDP-14C-glucose and 0.5 µM BcsA-BcsB in a total volume of 1.0 ml. Cellulose was removed by centrifugation and the supernatant was passed over 1 mL of DEAE- cellulose, washed two more times with 1 mL water each. The flow-through was dried under reduced pressure, the residue was dissolved in 0.5 mL water and spiked with glucose and cellobiose. The entire sample was loaded onto a 145 cm by 1.5 cm column packed with P-2 resin (biorad) and fractions of 2 mL were collected and quantified by scintillation spectroscopy. The internal standards were measured spectrophotometrically by the bicinchoninic acid assay, detailed in Materials and Methods.

Discussion

Kinetic Characterization of BcsA-BcsB

-1 Our finding of a kcat of 0.5 s for BcsA-BcsB is in contrast with the findings of

-1 Omadjela et al. (69) who determined a kcat value of 90 s . However, another publication from the Zimmer lab (109) shows a velocity of 0.47 s-1 for BcsA-BcsB under saturating substrate conditions for BcsA-BcsB. We have also shown a mean and standard deviation for the rate of cellulose synthesis under saturating UDP-glucose conditions of four preparations of BcsA-BcsB to be 0.50 ± 0.16 s−1. Our product entrapment of purified

BcsA-BcsB, essentially entrapment of active enzyme, confirms that at least 84% of the enzyme is active. This number is only a minimal estimate of active enzyme; the percent of active enzyme may be greater than 84%.

Product Analysis Demonstrates the Primer-Independent Mechanism for Initiation

Using two different experimental approaches, our results show that bacterial cellulose synthase initiates synthesis by a primer-independent mechanism. The first approach, GPC analysis of cellulose from both the AcsA-AcsB and BcsA-BcsB, gives details on the DOP of the products. It shows an increase in the polymer length of cellulose from 2 to approximately 10 minutes. At 30 minutes and longer, the products increase in amount (total glucose incorporated) but not maximum polymer length. Thus, cellulose synthase has a processivity limit. Our analysis also shows that new polymers of shorter length continue to appear over time. Both of these features are characteristic of a

70 primer-independent mechanism for initiation. Another feature apparent from both the carbanilation and unmodified cellulose GPC elution profiles is the DOP distribution difference of cellulose from AcsA-AcsB versus BcsA-BcsB. AcsA-AcsB produces cellulose of a greater peak and maximum DOP than BcsA-BcsB. Thus, AcsA-AcsB is a more processive enzyme. More will be discussed on processivity of cellulose synthase in

Chapter 4.

Second, we analyzed the reducing ends of the cellulose polymers synthesized by

BcsA-BcsB and AcsA-AcsB. We synthesized cellulose with UDP-14C-glucose, modified the reducing ends, and separated them from the rest of the incorporated glucose. By this method, we found 14C-glucose incorporated into the reducing end. This can only be accounted for by the initiation of new polymers using UDP-glucose.

Enzyme Concentration-Independent Rate Measurement

Determination of the DOP at the two- and five-minute time points yielded a turnover number of 25 s-1 for AcsA-AcsB and 6.3 s-1 for BcsA-BcsB. Again, these values

-1 are much faster than kcat values determined from steady-state kinetics of 1.7 s for AcsA-

AcsB and 0.51 s-1 for BcsA-BcsB. These results indicate that there may be two phases in cellulose synthesis.

In order to avoid confusion, the enzyme concentration-independent rates calculated by GPC analysis are referred to as the elongation rate. The elongation rate is the rate at which the enzyme incorporates glucose into an actively elongating glucan polymer.

When referring to the overall rate at which UDP is released from the enzyme we will use the term turnover number. The rate given by the turnover number includes the rates of initiation, elongation, and termination. Both rates are to be expressed in the unit ‘per second’ (s-1). Our analysis of the elongation rate for BcsA-BcsB while varying UDP- glucose concentrations indicates a second order rate constant of 14,000 M-1 s-1. This number is larger than the second order rate constant of the overall catalytic cycle for

BcsA-BcsB but is similar to the second order rate constant for another processive inverting glycosyltransferase, hyaluronan synthase, which is approximately 13,000 M-1 s-

1 (165). This suggests that the rate-limiting step in catalysis is not elongation (processive synthesis), but perhaps initiation of synthesis.

A Mechanism for Self-Priming

The presence of UDP-glucose hydrolase activity provides a mechanism by which cellulose synthase can generate glucose, as a primer, to initiate new polymer synthesis.

UDP-glucose hydrolase activity is not unique and has been demonstrated in other glycosyltransferases (115, 128, 166). The absence of glucose in AcsA-AcsB suggests that this enzyme is more efficient at priming. That is, it does not release glucose into solution.

Because our size exclusion analysis of soluble cellodextrins did not show the presence of any cellobiose, we hypothesize that once the cellulose polymer becomes two or more glucose units, backsliding into the active site is unlikely.

McNamara et al. (151) have suggested that, based on a structural comparison between BcsA-BcsB and the sodium-dependent galactose-transferase, SGLT, glucose is

72 sufficient to initiate new polymer synthesis. Modeling of the enthalpies of binding performed by our collaborator, Hui Yang3, for BcsA-BcsB with glucose and SGLT with galactose give values of -219 kJ/mol and -156 kJ/mol, respectively (Yang, unpublished).

In our model (Fig. 3.9), polymer elongation involves attack of the 4-carbon hydroxyl group of the glucan polymer non-reducing end to the anomeric carbon of UDP- glucose. Once the new glycosidic bond is formed, the polymer must translocate the length of one glucose unit (0.5 nm), release UDP and then bind UDP-glucose to continue the catalytic cycle (Fig. 3.9A). It is critical that translocation occur by only one glucose unit. A second translocation step would result in termination of the synthesis of the cellulose polymer within the channel. That is, the non-reducing end would be too far into the transmembrane channel to attack the UDP-glucose. The enzyme would then have to enter an initiation phase prior to re-entering the elongation phase. (Fig. 3.9B). Modeling by collaborator Hui Yang suggests that once the attack position is vacated by the non- reducing end of the glucan polymer, water takes its place (Yang, unpublished). Initiation of a new glucan polymer would then begin with UDP-glucose hydrolysis. After this, the liberated glucose enters the transmembrane channel becoming the reducing end of a new glucan polymer (Fig. 3.9B).

3 Penn State University, Department of Chemical Engineering. 304 S. Frear Building, University Park, PA 16802 73

Figure 3.9. A Proposed Mechanism of New Polymer Initiation by Cellulose Synthase. Panel A shows polymer elongation in which (I) the non-reducing end of the polymer (which associates with trp383 (W) in BcsA-BcsB), attacks the 1-carbon of UDP-glucose. (II) The extended polymer must then translocate one glucose unit and release UDP. (III) The newly added glucose is positioned to begin the cycle again. Panel B shows the proposed mechanism initiation. (I) The glucan polymer translocates one addition glucose unit, allowing water to take the place of the 4-carbon hydroxyl of the non-reducing end. (II) UDP-glucose is hydrolyzed by water (deprotonated by the catalytic base; asp343 in BcsA-BcsB) to yield glucose. (III) The newly liberated glucose is positioned to begin a new cellulose polymer. Note that the ‘OH’ indicates the reducing end of the cellulose polymer.

CHAPTER 4

PROCESSIVITY AND A MINIMAL KINETIC MECHANISM FOR CELLULOSE

SYNTHESIS

Introduction

Processivity is the maximum DOP to which a processive enzyme can elongate its polymeric product (167). In Chapter 3, we showed, by GPC analysis, that AcsA-AcsB produces glucan polymers of a greater maximum DOP than those produced by BcsA-

BcsB. Thus, AcsA-AcsB is a more processive enzyme than BcsA-BcsB. Understanding processivity in cellulose synthase is of central interest to industrial applications. The recalcitrance of paper to degradation, for example, is impacted by the DOP of its constituent cellulose (50). In textiles, the depolymerization of cellulose by acid treatment reduces the durability and tensile strength of cotton fabric (95). In the biofuels industry, the DOP of glucan polymers affects the efficiency of biomass digestion during extraction treatments with chemicals and enzymes (51, 168). Engineering plants to synthesize cellulose of lower DOP could improve digestion efficiency (8).

DNA polymerases are processive enzymes which provide a basis for understanding the principles underlying processivity of enzymes (169, 170). For example, the E. coli enzyme, DNA Pol I, generates fragments of 10-50 nucleotides before dissociating from the template strand (171). On the other hand, DNA Pol III, also from E. coli, forms a multiprotein complex that synthesizes fragments several thousand

75 nucleotides in length between priming events (172). The complex accessory proteins, exonuclease ε and sliding clamp β, increase DNA Pol III’s processivity; DNA Pol III α subunit alone synthesizes only 11 nucleotides between priming events (172-174). A comparison between the structures of different DNA polymerases gives a structural basis for their processivity. Those enzymes fully enclosing the template strand have a higher processivity than enzymes with partial enclosure around the template (175). DNA pol III is one such example: the α subunit forms the classic hand-like structure, a partial enclosure around the template strand (176), and the sliding clamp β fully encircles the

DNA template (177, 178).

Unlike DNA polymerase, cellulose synthase does not utilize a template strand.

The transmembrane channel of BcsA, through which the polymer translocates, makes extensive contact with the cellulose polymer (19, 109, 110). Therefore, to probe the role of this channel on enzyme kinetics and on processivity, we changed residues shown to interact with the glucan polymer in the transmembrane channel of BcsA-BcsB (19). In this chapter, we kinetically characterize these variants. In one such variant, we found reduced processivity as well as a reduced turnover number and elongation rate. Finally, at the end of this chapter we construct a minimal kinetic mechanism for cellulose synthesis.

Materials and Methods

Cloning of BcsA-BcsB Variants

For each variant, two fragments were generated by PCR amplifying, using primers to the BcsA-BcsB expression plasmid (Appendix A). One fragment is upstream from the variation site and one fragment is downstream from the variation site. Both fragments contained complementary sequences (3’ end for fragment one and 5’ end for fragment two) overlapping at the variation site. The BcsA-BcsB expression plasmid was digested using BmgBI and AgeI or StuI (for the V551A variant only) and gel purified. The two fragments, along with digested BcsA-BcsB plasmid, were ligated using the In-Fusion HD

Cloning Kit (Clontech). XL1blue cells were transformed with the ligated plasmid.

Colonies were selected for sequencing (Penn State Genomics Facility) and Rosetta cells were transformed with successful variant plasmids. All variants used alanine to replace the original amino acid, except for the tryptophan 383 variant, which was to phenylalanine.

Kinetic Simulation of Cellulose Synthesis

Kinetic simulations were performed by our collaborator, Hui Yang, using a derivative of KINSIM software, called Tenua (179).

Results

Kinetic Analysis of BcsA-BcsB Variants

The catalytic cycle of cellulose synthesis involves attack of 1-carbon of UDP- glucose by the 4-carbon hydroxyl group of the non-reducing end of cellulose synthase.

After the addition of the new glucose, the polymer has to translocate one glucose unit in order to continue elongation. Like DNA polymerase, we hypothesize that the binding affinity of the enzyme for the glucan polymer relates to the processivity of the enzyme.

We reasoned that the affinity for the glucan polymer would be most affected by residues associating with the glucan polymer. Selected amino acid residues of the transmembrane channel shown to interact with the cellulose polymer were altered. Individual variants were generated as described in Materials and Methods. Figure 4.1 shows the residues that were changed.

Figure 4.1. BcsA-BcsB Transmembrane Channel Amino Acid Variants. The glucan polymer is shown in teal and the first ten glucose residues (those which reside in the transmembrane channel) are numbered. All the labeled amino acid residues were changed to alanine, except for Trp383, which was changed to phenylalanine. Mino acid changes resulting in inactive BcsA-BcsB are colored red and amino acid changes resulting in active BcsA-BcsB are colored green.

Only variants N412A, F416A, Q463A, and V551A were active. The steady-state kinetic parameters were determined by the enzyme-linked assay for UDP (Chapter 2). Enzyme activity was measured at varying concentrations of UDP-glucose (Appendix B, Fig B.4).

Table 4.1 shows the kcat and KM values and the second order rate constants for each variant.

Table 4.1. Kinetic Values for BcsA-BcsB Variants.

-1 -1 -1 Mutation kcat, s KM, mM kcat/KM, M s

N412A 0.184 0.56 328 F416A 0.072 0.62 116 Q463A 0.313 0.80 392 V551A 1.155 0.66 1747

Gel Permeation Chromatography of Cellulose from BcsA-BcsB Variants

To probe processivity, cellulose synthesized by each variant was analyzed by

GPC analysis, as described in Chapter 3. Cellulose synthesis was monitored by the enzyme-linked assay described in Chapter 2. Each reaction was quenched with 2% SDS after 30 nmol of UDP was liberated to ensure that the same amount of cellulose was synthesized by each variant. The cellulose was dissolved in DMAc / 8% LiCl and subjected to GPC analysis (described in Chapter 3) (Fig. 4.2). The DOP at the peak of the elution profiles for wild type and variants N412A, Q463A, and V551A was 1,505, with a

79 maximum DOP of 11,700. The DOP at the peak for variant F416A was 760, with a maximum DOP of 5,900.

14 10000 12 1000 10

8 100 Wild Type 6 N412A 10 F416A 4 Q463A V551A 1 2

Cellulose (nmol glucose)

Degree of Polymerization Degree

0 0.1 4 6 8 10 12

Elution volume, ml

Figure 4.2 GPC Elution Profile of Cellulose from BcsA-BcsB Variants. Cellulose was synthesized from 0.5 µM of each BcsA-BcsB variant using 5 mM UDP- 14C-glucose in a reaction volume of 0.2 ml. Cellulose synthase activity for each mutant was monitored by UDP release using the enzyme-linked assay described in Chapter 2. When each reaction had consumed 30 nmol of UDP-glucose, the reaction was quenched with 2% SDS. The underivatized cellulose was dissolved in 8% LiCl (wt/vol) in dimethylacetamide as described in Chapter 3. Fractions (0.5 ml) were collected and quantified by scintillation spectroscopy. The degree of polymerization, measured by MALLS for bacterial cellulose and Avicel (Chapter 3) is shown as bold lines above the elution profiles and is measured on the right axis. A calibration curve is drawn through these two lines.

Time Course Analysis of Cellulose from F416A

Because of the apparent reduced processivity of F416A compared with wild type, we examined the elongation rate. This was done, as for wild type, described in Chapter 3, by GPC analysis of cellulose synthesized over a time course by F416A (Fig. 4.3).

Cellulose synthesized from F416A for 2-5 minutes increased in size, eventually reaching a DOP maximum of 5,900 (the front of the elution profile). From 30-300 minutes, the maximum size did not increase, however, the overall incorporation of glucose continued. From 30-300 minutes, the DOP at the peaks was 800. Examination of the DOP at the peak of the two-minute time point elution profile reveals a DOP of 200, equivalent to an elongation rate of 1.6 s-1. Examination of the peak at the five-minute time point elution profile reveals a DOP of 400, equivalent to an elongation rate of 1.3 s-

1. Like wild-type BcsA-BcsB, the overall turnover number for F416A is slower than the elongation rate.

120 10000

100 300 min 1000 80 150 min 30 min 5 min 100 60 2 min 10 40

20 1

Cellulose (nmol glucose) (nmol Cellulose

Degree of Polymerization Degree

0 0.1 6 8 10 12

Elution volume, ml

Figure 4.3. GPC Elution Profile of Cellulose from the F416A Variant. Aliquots were removed from enzymatic reaction mixtures at times as specified in the

81 figures. Cellulose was synthesized from 0.5 µM of the F416A variant with 5 mM UDP- 14C-glucose in a reaction volume of 1.0 ml as described in Chapter 2. The underivatized cellulose was dissolved in 8% LiCl (wt/vol) in dimethylacetamide as described in Chapter 3. Fractions (0.5 ml) were collected and quantified by scintillation spectroscopy. The degree of polymerization, measured by MALLS for bacterial cellulose and Avicel (Chapter 3) is shown as bold lines above the elution profiles and is measured on the right axis. A calibration curve is drawn through these two lines.

Discussion

Processive Synthesis

In Chapter 3 we showed, by GPC analysis, that cellulose produced from AcsA-

AcsB has a DOP at the peak and maximum that was greater than the DOP at the peak and maximum for cellulose produced from BcsA-BcsB. Thus, AcsA-AcsB has a greater processivity than BcsA-BcsB. The processivity of cellulose synthase, like that of DNA polymerase (180), can be minimally explained by the binding constant of the enzyme to the polymeric product. The equilibrium constant for a substrate to an enzyme is the ratio of the binding rate constant, k3 to the dissociation rate constant k-3 given for cellulose synthase in mechanism.

The equilibrium constant is a ratio of the on and off rates. The greatest interaction of

BcsA with the glucan product is through the amino acid residues in the transmembrane channel (19, 109, 110), and thus we changed these residues in order to probe the effects on processivity.

For the W383F variant, we reasoned that changing the residue from tryptophan to phenylalanine (rather than alanine) would be conservative and thus might allow BcsA-

BcsB to retain activity while altering processivity. However, this was not the case; the

W383F variant was not active. The crystal structure of BcsA-BcsB shows that the tryptophan ring associates, through Van der Waals interactions, with the ring of the attacking glucose (the non-reducing end). The nitrogen of the tryptophan ring also forms a hydrogen bond with the oxygen of the glucose ring in the UDP-glucose substrate (110).

This hydrogen bond is not possible in the W383F variant. Trp383 is conserved in bacterial and plant cellulose synthases (110) (Fig. 4.4). Our results, as well as bioinformatics and structural data, suggest that it may serve a necessary role in glycosidic bond formation. Both the F301A and the W317A variants were also inactive. Both the phe301 and the trp317 residues interact with the second glucose unit of the glucan polymer via Van der Waals and hydrophilic interactions, respectively. Phe301 is conserved across bacterial cellulose synthases (Fig 4.4). Trp417 is not conserved across bacterial cellulose synthases (Fig 4.4).

R. sphaeroides 296PENEMFYGKIH306 378QQRGRWATGMMQ389 412NSMSFWFFPL421 G. hansenii PEGNLFYGVVQ GQRVRWARGMLQ SAMTSFLFAV E. coli NEGTLFYGLVQ GQRIRWARGMVQ NAMFHFLSGI Pseudomonas sp. NEGELFYGLVQ NQRIRWARGMAQ NAMLHFFYGL Agrobacterium sp. SENEMFYGIIQ GQRSRWAQGMMQ SSTLFWLFPF

Figure 4.4. Sequence Alignment of Selected Residues from Bacterial Cellulose Synthase A Subunits. Trp383 and phe301 in BcsA from R. sphaeroides interact with the first and second glucose residues of the glucan polymer and are conserved across bacterial cellulose synthases. Trp417 and phe416 in BcsA from R. sphaeroides interact with the second and third glucose residues of the glucan polymer, respectively, and are not conserved across bacterial cellulos synthases.

Interestingly, when attempting to purify inactive variants, only the monomer containing the histidine affinity tag is purified, when examined by SDS-PAGE analysis.

For BcsA-BcsB, this is the BcsA subunit. Our attempts to generate glucan-free AcsA-

AcsB resulted in inactive enzyme and our results showed only AcsB purifying. Recall from Chapter 2 that the AcsB subunit contains the histidine tag for the homologous expression system. There are myriad explanations why catalytically-inactive heterodimers dissociate. One such explanation may be that the association of the two subunits (A and B) is dependent upon the presence of the glucan polymer. This would suggest that the polymer either physically ‘glues’ the two subunits together, or causes conformational changes in either subunit which promote dimerization.

The V551A variant showed an increased overall turnover number but no apparent shift in the DOP distribution of cellulose. Val551 lies at the opening of the transmembrane channel (glucose residue 10) where the glucan polymer exits BcsA. The

N412A and Q463A variants showed a decrease in the overall turnover number but no apparent shift in the DOP distribution of cellulose. One interpretation of these observations is that the rate of polymer translocation is decreased but the net rate for the dissociation of the glucan polymer remains unchanged. The reasons underlying the observations for these variants remain unclear.

Only the F416A variant showed a decrease in the overall turnover number (0.07 s-

1) and the processivity. This prompted us to measure the elongation rate of the variant, which was also reduced compared to wild type (1.6 s-1 versus 4.1 s-1). Phe416 (in BcsA-

BcsB) is not conserved in all bacterial cellulose synthases (Fig. 4.4). Our results suggest

84 that this residue is not necessary for cellulose synthesis but does impact the affinity of the enzyme for the glucan polymer. It supports our hypothesis that certain residues that interact with the glucan polymer affect the enzyme’s processivity. McNamara et al. (151) postulate that the transmembrane channel must not bind the glucan polymer so tightly as to hinder translocation, however, must still prevent backsliding into the cytosol or premature release of the polymer into the extracellular space. Polymers of sufficient molecular weight, the authors continue, may be torn from the enzyme by sheering forces or may stall synthesis due to increasing interactions with the extracellular milieu (151). It is conceivable then, that an equilibrium between the affinity of the enzyme for the glucan polymer and the increasing extracellular forces, in part, contribute to the processivity of cellulose synthase

Finally, it is known that the digestion efficiency of cellulosic biomass is impacted by the DOP of the constituent cellulose (8). A practical application of our findings involves targeting residues in the transmembrane channel of cellulose synthases from plants with biomass feedstock applications. This is a new a way of altering the DOP of cellulose.

A Minimal Kinetic Mechanism for Cellulose Synthesis

Figure 4.5 shows our proposed mechanism for cellulose synthesis. In this mechanism, the 4-carbon hydroxyl group of the non-reducing end of the glucan polymer attacks the anomeric carbon of UDP-glucose. Once the new glucose is added, the polymer has to process into the channel the length of one glucose unit (0.5 nm), release UDP and then

85 bind UDP-glucose to continue the catalytic cycle (110). As we proposed in Chapter 3, procession by at least two, rather than one, glucose unit would result in the termination of synthesis for that glucan polymer. As a consequence, the enzyme enters the priming cycle. In the priming cycle, cellulose synthase hydrolyzes UDP-glucose then bind the liberated glucose to re-enter the elongation cycle. The following paragraphs address each step in the kinetic mechanism (Fig. 4.5). The net rates are valid under initial velocity and the species shown in each mechanism are defined in the figure legend for Figure 4.5.

Figure 4.5. A Minimal Kinetic Mechanism for Cellulose Synthesis. Elongation starts with the cellulose polymer properly position in the channel for nucleophile attack of UDP-G by the non-reducing end (E-Cx). UDP-G then binds to E-Cx at a second order rate constant (k5). Attack of the 4-hydroxyl to the UDP-G forms the glycosidic bond. This rate (k6) is unknown, but is not rate limiting in the fast phase. The newly- added glucose then translocates into the channel by one glucose unit (k4). This we have assigned as the rate limiting step in the elongation cycle and is measured as 4.1 s-1 (BcsA-BcsB) and 25 s-1 (AcsA-AcsB). Alternatively, the cellulose polymer can translocate by more than one glucose unit (k3) which results in polymer release. Upon release, the enzyme then needs to re-initiate. UDP-G then binds to E at a second order rate constant (k1). Hydrolysis of UDP-G to yield glucose occurs at a rate given by k2. The known rates are tabulated (Table 4.2)

In Chapter 3, we measured the elongation rate for BcsA-BcsB by varying the

UDP-glucose concentration and fitting the rates to the Michaelis-Menten equation. The

-1 -1 rate of 25 s and 4.1 s gives us a minimum value for k’4 in mechanisms 3 for AcsA-

AcsB and BcsA-BcsB, respectively. The net rate constant, k’6, for mechanism 2 is a first order rate for glycosidic bond formation. Because this step is not rate limiting for the fast cycle (Fig. 4.5), its value is not important. The net rate constant, k’4, for mechanism 3 is a first order rate for an unknown process, but one in which the enzyme becomes

87 catalytically competent once again. We have assigned the rate limiting step for the fast cycle to mechanism 3.

The second order rate constant, 417,000 M-1 s-1 for AcsA-AcsB and 14,000 M-1 s-1 for

BcsA-BcsB, was obtained by fitting the elongation rates determined in Chapter 3 to the

Michaelis-Menten equation, gives us the value for the net rate constant, k’5, for mechanism 4.

The net rate constant, k’1, in mechanism 5 is a second order rate constant that is unknown, but, must be smaller than the second order rate constant for the overall reaction

(6,100 M-1 s-1 for AcsA-AcsB or 600 M-1 s-1 for BcsA-BcsB), obtained by fitting initial velocities at varying substrate concentrations to the Michaelis-Menten equation (Chapters

2 and 3). Under saturating substrate conditions, the rate limiting step for the slow cycle

(Fig 4.5) is shown in mechanism 6 and is given by the net rate constant, k’2. Mechanism 6 includes UDP-glucose hydrolysis and binding of the liberated glucose by cellulose synthase.

In our minimal mechanism, the second translocation step, or polymer release, is defined by the net rate, k’3, for mechanism 1. By dividing the total amount of glucose by the DOP

88 in the time course elution profiles from Chapter 3 (Fig. 3.2) the number of reducing ends

-3 -1 -3 -1 can be determined. A rate for k’3 of 1.2 × 10 s (BcsA-BcsB) and 0.3 × 10 s (AcsA-

AcsB) was determined for both enzymes. Thus, the processivity of a cellulose synthase is directly influenced by the relative rate of elongation k’4 versus strand release k’3. This is corroborated by the ratio of these two rates (k4/k3) for AcsA-AcsB (335,000) versus

BcsA-BcsB (10,000). The measured kinetic values for both enzymes are given in Table

4.2.

Table 4.2. Kinetic Values for AcsA-AcsB and BcsA-BcsB.

-1 -1 -1 -1 -1 -1 -1 k3, s k4, s k5, M s kcat, s kcat/KM, M s

AcsA-AcsB 0.3 × 10-3 s-1 25 417,000 1.7 6,100 -3 -1 BcsA-BcsB 1.2 × 10 s 4.1 14,000 0.5 600

Next, we simulated our kinetic mechanism of cellulose synthesis using Tenua software4. Having experimentally determined four of the six rate constants (Table 4.2) only k2 and k6 were unknown. The simulation can be used to estimate k2. The value of k6 is not important since it is not the rate-limiting step in elongation. The validity of the mechanism and the rate constant is demonstrated by the ability of Tenua to simulate the steady-state kinetics. Simulations were run at varying UDP-glucose concentrations and the resulting rates were fit to the Michaelis-Menten equation (Figure 4.6). The simulations mimic the experimental values remarkably well. The kinetic values for Tenua simulation can be found in Appendix C.

4 Simulations were performed by Hui Yang. The input into Tenua for simulations is listed in Appendix C. 89

Having been able to simulate steady-state kinetics, we then attempted to simulate processivity. This was problematic because that involves thousands of steps. However, we were able to set up a mock simulation that allowed us to determine the effect on the trends of cellulose DOP by changing some of the rate constants. Rate constants 100 times higher than the actual rates were used such that Tenua would provide a simulation on the size distribution (Fig. 4.7A and B). The effect of increasing k3 decreases the DOP (Fig.

4.7A) whereas increasing k4 increases the DOP (Fig. 4.7B). Thus. the processivity of a cellulose synthase is directly influenced by the relative rate of elongation k4 versus strand release k3.

Figure 4.6. Simulated Michaelis-Menten Curves for (A) AcsA-AcsB and (B) BcsA- BcsB Using Tenua Software. Rates were simulated for cellulose synthesase at varying UDP-glucose concentrations using Tenua software. The simulated rates are shown as open circles and the experimentally determined rates are shown as filled circles. Rates are expressed as velocity (s-1).

250 120

100 200

80 150 60 100

40 Cellulose,nmolglucose Cellulose,glucosenmol 50 20

0 0 value value 140 5000 500 50

140 5000 500 50 2 min 4

3 3 2 min k

k 5 min 120 5 min 120 10 min 10 min 100 100 30 min 30 min 60 min 80 60 min 80 120 min 120 min 60 240 min 60 240 min 40

40 Cellulose,glucosenmol Cellulose,glucosenmol 20 20 0

0 Increasing the Increasing

Increasing the Increasing 140 5000 500 50 90 5000 500 50 80 120

70 100 60 80 50 40 60

30 40

20 Cellulose,glucosenmol Cellulose,glucosenmol 20 10 0 0 5000 500 50 5000 500 50 Degree of Polymerization Degree of Polymerization

Figure 4.7 Simulated Product Distributions from Cellulose Synthase using Tenua Software. Because Tenua can only accommodate 100 steps, the rate constants were multiplied by 100 for the purpose of illustrating the importance of the k4 to k3 ratio in determining the processivity of the enzyme. The Right Panel shows that increasing k4 results in increased DOP. The left shows that increasing k3 results in decreased DOP. The Tenua input is available in Appendix C.

CHAPTER 5

THE CELLULOSE SYNTHASE COMPLEX: ACCESSORY PROTEINS AND

WHOLE-CELL CELLULOSE SYNTHESIS

Introduction

Gram negative bacteria are contained in a cell envelope that is made up of two phospholipid bilayers: an inner membrane, called the cytoplasmic membrane, and an outer membrane (181). The aqueous region between the two membranes, the periplasmic space, contains a thin peptidoglycan cell wall (181). The cell envelope is responsible for communication with the external environment, transport of nutrients into the cell, and export of metabolites out of the cell (181-183).

In G. hansenii, cellulose is synthesized by large multi-protein complexes which form a linear array both spanning and running longitudinally across the cell envelope (32,

90). This linear array is called the cellulose synthase complex (CSC) and extrudes cellulose as crystalline ribbon-like fibril which elongates into the extracellular milieu (32,

60). Synthesis, crystallization, and transport of the glucan polymers across the cell envelope is facilitated by the CSC. Mutagenesis studies were the first to identify the genes (proteins) required for cellulose synthesis (60, 67, 90, 94, 96, 145). Those identified in G. hansenii were AcsA, AcsB, AcsC, AcsD, CcpAx, CmcAx, and BglAx (58,

68, 90, 94, 99, 103, 104, 107, 142).

In work described in Chapter 2, we isolated the minimal catalytic core, AcsA-

AcsB, from the CSC. In Chapter 3 we describe work discovering an elongation rate that

92 was faster than the overall turnover number. The difference between these rates can be explained by a slow step in the overall catalytic mechanism. In Chapter 4, we presented a minimal mechanism which posits that the slow step in catalysis is initiation. To test if the turnover number is equally as slow in vivo, in this chapter, we measure the turnover number for AcsA-AcsB in whole cells. We find that it is faster in whole cells. We then investigate the influence of CSC accessory proteins on the cellulose synthesis rate and cellulose translocation in whole cells, total membrane (TM), and with purified enzyme.

Specifically, we examine the roles of AcsC, AcsD, and CcpAx, each of which has been shown to impact cellulose synthesis in whole cells (25, 60, 90). Much of what is known about AcsC has been learned through homology studies, and it is thought that

AcsC facilitates the transport of cellulose through the outer membrane (59, 99). AcsD forms a homo-octamer localized to the periplasm (103). The crystal structure of AcsD shows a central pore large enough to facilitate the transport of four glucan polymers

(104). This suggests that AcsD aides in crystallization of the glucan polymers (99, 104).

CcpAx complexes with AcsD and appears to be important in the maintenance of the CSC

(90).

Finally, we calculate the turnover number for AcsA-AcsB at each step during product entrapment purification (Chapter 2) to identify at which step the cellulose synthesis rate declines.

Because R. sphaeroides is not known to produce crystalline cellulose in vivo, we characterized only the G. hansenii (AcsA-AcsB) system.

Materials and Methods

Determination of in vitro Turnover Number

G. hansenii ATCC23769 cells were grown, shaking (250 rpm) at 30°C, to an absorbance of 0.8 - 1.1 at 600 nm. Cultures contained 0.01% cellulase (Worthington).

The cells were collected from 50 mL of culture by centrifugation at 6,000 × g for 5 minutes. The cells were washed twice with 50 mL of media, resuspended in 50 mL of media, and poured into a 250 mL flask. After being allowed to recover for 5 min while shaking at 30°C, samples for cellulose and AcsA-AcsB quantification were taken at 0,

10, 20, and 30 min while shaking (250 rpm) at 30°C.

For cellulose quantification, 1 mL of cells was collected in triplicate and boiled for 10 min. The insoluble material was collected by centrifugation at 16,000 × g for 5 min and washed two times with 1 mL of water, once with 1 mL of ethanol, and once with 1 mL of sterile water. The insoluble material was resuspended in a sterile-filtered solution of 300 µL of 50 mM sodium acetate buffer, pH 5.0, containing 300 U of cellulase

(Worthington) and incubated at 25°C for 24 hours. Following digestion, the mixture was passed through a 3 kDa cut-off spin filter (Millipore). Glucose, released from cellulose, was quantified by the glucose oxidase assay, as described in Chapter 3.

For AcsA-AcsB quantification, 3 mL of cells were collected in triplicate at 0 min.

Cells were collected by centrifugation at 16,000 × g for 5 min and resuspended in 1 mL of 50 mM sodium phosphate, pH 7.0. Cells were lysed by sonication. A total of 80 µL of lysate was added to 20 µL of 5 × LDS loading buffer and incubated on ice for 1 h.

Samples of 10 µL were separated by SDS-PAGE and analyzed by western blot using anti-AcsB (Chapter 2). Known quantities of purified AcsA-AcsB were used to generate a standard curve. Band intensities were quantified using Image J software.

Cell number was determined by colony count. Cells were grown, shaking (250 rpm) at 30°C, to an absorbance of 1.0 at 600 nm. Cultures contained 0.01% cellulase

(Worthington). Serial dilutions of cells were prepared in sterile water containing 50 mM sodium phosphate, pH 7.0. Samples of 100 µL were removed from each dilution and plated onto SH media and grown at 30°C for 3 days. Plates containing between 20 and

200 colonies were used to determine cell number per mL of the initial culture.

Total Membrane Isolation

TM was isolated for use in sucrose density gradient separation and enzyme assays as described by Myers et al. (184) and stored in 10 mM Tris-HCl, pH 8.0, with 20% glycerol at -80°C. Note that this method is different than the large scale preparations for enzyme purification as described in Chapter 2.

Generation of t-DNA Insertional Mutants

T-DNA insertional mutants for acsC, acsD, and ccpAx were generated by our collaborators, Deng et al. (60). Primers for the acsD insertional mutation are listed in

Appendix A. Primers for the acsC and ccpAx insertional mutations are documented (60).

Statistical Analysis

To determine statistical significance in the activity and protein levels reported in

Figures 5.2 and 5.3, a pairwise comparison between the means of the wild type control and each mutant was performed using Welch’s t-test (two tailed, unequal variance).

Cloning, Expression, and Purification of AcsD

The gene encoding AcsD was amplified from G. hansenii genomic DNA by PCR using primers for AcsD (Appendix A). The 0.5 kb fragment was ligated into a NdeI/XhoI- digested modified pET21a vector, which added a C-terminal hexahistidine tag preceded by a thrombin cleavage sequence to the AcsD peptide, and expressed in BL21(DE3) cells. The protein was purified by Ni-NTA chromatography (185) and quantified using a molar extinction coefficient of 26,470 cm-1 M-1 at 280 nm.

Cloning, Expression, and Purification of CcpAx

The fragment chosen for cloning was based on the active peptide identified by the work of Sunagawa et al. (90). The ccpAx gene was obtained by PCR (Appendix A). The

186 bp fragment was cloned into the modified pET21a vector also with a hexahistidine tag at the C-terminus. Expression and purification methods were identical to those used for AcsD. CcpAx was quantified by Lowry assay (Chapter 2).

Transmission Electron Microscopy5

Cells were fixed with 2% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2 containing 3 mM CaCl2 at room temperature. The cells were then post-fixed with 1%

OsO4 for one hour at 4°C, followed by en bloc staining in 2% aqueous uranyl acetate for

30 min. The cells were dehydrated with a graded ethanol series and embedded in Eponite

12 (Ted Pella). The sample blocks were sectioned at 80 nm thick with a Leica UC6 ultramicrotome and the grids were double stained with uranyl acetate and lead citrate.

The cells were examined and imaged with a JEOL 1200 EXII TEM.

Separation of Total Membrane by Sucrose Density Gradient

TM was resuspended in 6 mL of 10 mM Tris-HCl, pH 8.0. and carefully layered on top of 34 mL of 25% - 55% sucrose containing 10 mM Tris, pH 8.0. The entire gradient was subjected to centrifugation in a swing bucket rotor at 110,000 × g for 17 hours at 4°C. Fractions of 3 mL (13 total fractions) were removed from the top of the gradient and stored at -80°C. The purity of the cytoplasmic membrane (CM) was assessed by marker enzyme assays using succinate dehydrogenase. The assay was performed using a modified procedure described by Anwar et al. (186). The 200 μL reaction mixture contained 60 mM sodium phosphate, pH 7.2, 10 mM potassium cyanide,

5 µg phenazine methosulfate, 25 mM sodium succinate, and 20 µg of dichlorophenol indophenol (DCIP). The reaction was initiated upon the addition of 10 µL of each

5 Staining, sectioning, and microscopy was performed by the Pennsylvania State University Microscopy and Cytometry Facility. 97 fraction and monitored by decrease in the absorbance of dichlorophenol indophenol

(DCIP, extinction coefficient 13,000 M-1 cm-1) at 600 nm at 30°C.

Results

Determination of the in vivo Turnover Number

In our kinetic mechanism for cellulose synthesis (Chapter 4), we showed that catalysis can be broken up into two phases, and hypothesized that the slow phase includes initiation. Here, we hypothesize that the slowness of the overall turnover number (relative to the elongation rate) may be due to an inefficiency in one of the catalytic steps in vitro.

Therefore, we measured the in vivo rate of cellulose synthesis to test if it is equally as slow in whole cells. To do this, whole cells were grown to mid-log in the presence of excess cellulase, washed, and then inoculated into fresh media as described in Materials and Methods. Aliquots from this shaking culture were removed at 0, 10, 20 and 30 min and then the cellulose content and the AcsA-AcsB content were determined. The time scale of 30 min is short enough not to have large changes in the cellulose synthase content due to cellular growth because the double time for G. hansenii is 7 hours (187).

By the time 10 minutes had elapsed, cellulose was already visible in the media.

Cellulose content of the cultures was measured by digestion with a mixture of endo- and exo-glucanase and β-glucosidase followed by quantification with glucose oxidase (Fig. 5.1C) as described in Materials and Methods. AcsA-AcsB levels were assessed by quantitative western blotting using anti-AcsB. Homologously-expressed and purified AcsA-AcsB (Chapter 2) was used a standard for the western blots (Fig. 5.1A and

B). AcsB was chosen for quantitation over AcsA due to concerns over AcsA’s instability

(Chapter 2).

From mid-log cultures, aliquots were removed and the cell number determined by colony count as described in Materials and Methods. Quantification of both AcsB content and cell number allowed us to determine the number of AcsB molecules per cell. We calculated 1.5 × 104 ± 7.8 × 103 AcsA-AcsB molecules per cell. Similarly, the number of glucose molecules incorporated per cell was found by dividing the number of glucose molecules incorporated per milliliter of culture by the number of cells per milliliter of culture. We calculated 3.8 × 105 ± 2.0 × 105 glucose units incorporated into cellulose per second per cell. This is in close agreement with the results of Schramm and Hestrin (62) of 2.0 × 105 glucose molecules per second per cell.

The in vivo turnover number was calculated by dividing the number of moles of cellulose produced per second per milliliter of cells by the number of moles of AcsA-

AcsB per milliliter of cells. Rates of 25 ± 5 s-1, 22 ± 11 s-1, and 24 ± 8 s-1 were calculated for three separate biological samples. The mean and standard deviations for each biological replicate represents three technical replicates. The whole-cell cellulose synthesis rate, approximately 24 s-1, is 14-fold higher than the steady-state rate of 1.7 s-1 determined for purified AcsA-AcsB (Chapter 2) and nearly identical to the elongation rate of 25 s-1 determined by GPC (Chapter 3). Our measured rate of whole-cell turnover number is relatively close to the value of 67 s-1 determined by the microfibril extrusion rate observed under dark field microscopy (32).

Figure 5.1. The Calculation of a Turnover Number for Whole Cells. Cells were grown to mid-log in 0.01% cellulase, harvested, washed with media, and resuspended in media. Samples were taken at 0, 10, 20, and 30 minutes. Cellulose synthase was quantified from whole cells (A, lanes 4-6) by western blotting with anti-AcsB using purified AcsA-AcsB as standards (A, lanes 1-3). Band intensities were quantified by Image J analysis (B). Standards are shown as filled circles and samples as filled diamonds. Moles of cellulose were determined by digestion with cellulase and quantification by glucose oxidase (C) as described in Chapter 3. Error bars represent the standard deviation for three technical replicates.

The Effects of AcsC, AcsD, and CcpAx on Enzyme Activity in Total Membrane

Genome sequencing has shown that acsAB is part of an operon that includes acsC

(98). Mutations in acsC, acsD, and ccpAx decrease or eliminate the amount of cellulose synthesized in vivo (25, 60, 90). We reasoned that these proteins may impact cellulose synthesis and, as such, we further explored the effects of AcsC, AcsD, and CcpAx on

AcsA-AcsB activity and expression.

100

Our collaborators, Deng et al. (60), generated knock-out mutants for acsC, acsD, and ccpAx using Tn5 transposon insertional mutagenesis. Previous publications show that knock-outs in acsC, acsD, and ccpAx significantly lowered, if not totally eliminated, cellulose production in G. hansenii cultures (25, 60, 90). It is not known, however, whether the impact of these proteins is on catalysis or on the translocation and crystallization of the cellulose by the CSC, which is located in the cytoplasmic membrane across the periplasm to the exterior of the cell. This translocation must occur through the periplasm and the outer membrane.

Therefore, we measured the cellulose synthase specific activity in TM by 14C- glucose incorporation, described in Chapter 2. For the acsC insertional mutant, the in vitro specific activity was not statistically different, when compared to that of wild type

(Fig 5.2). This is consistent with AcsC’s proposed role as an exit porin (99). TM from the acsD insertional mutant showed a slightly reduced specific activity (Fig. 5.2). TM from the ccpAx insertional mutant showed a significant reduction in the specific activity

(p-value <0.01), showing only 33 ± 4% of wild type (Fig. 5.2).

We next examined the levels of the AcsA and AcsB protein in these three mutants by western-blot analysis (Fig. 5.3A and 5.3B). The AcsA and AcsB levels in the TM fraction were similar to that in wild type for the acsC and acsD insertional mutants (Fig.

5.3C). Interestingly, the ccpAx insertional mutant had no impact on the AcsA protein levels, but significantly decreased the AcsB levels relative to wild type (p-value <0.01).

101

Figure 5.2. Quantification of Cellulose Synthase Activity in Wild Type (WT) and Insertional Mutants. The TM from each of the insertional mutants (as designated in the figure with a Δ before the name of the gene) and from wild type (WT) were isolated. Cellulose synthase activity was measured by 14C-glucose incorporation as described in Chapter 2. Each assay contained a total of 120 µg of TM. Results show mean and standard deviation from three biological replicates (mean value for wild type was 25.0 ± 2.6 nmol/min/mg). *Statistically significant difference in activity when compared to WT using Welch’s t-test, p-value <0.05. **Statistically significant difference in activity when compared to WT using Welch’s t-test, p-value <0.01

102

Figure 5.3. Western-Blot Visualization and Quantification of SDS-Polyacrylamide Gels of TM from Wild Type, or acsC, acsD, or ccpAx insertional mutants. A total 30 μg of TM protein was loaded in each lane. Panel A: Western blot, visualized with anti-AcsA3, of TM from wild type (lane1), acsC insertional mutant (lane 2), acsD insertional mutant (lane 3), and ccpAx insertional mutant (lane 4). Panel B: Western blot, visualized with anti-AcsB, of TM from wild type (lane 6), acsC insertional mutant (lane 7), acsD insertional mutant (lane 8), and ccpAx insertional mutant (lane 9). The three blots in each panel are of three biological replicates. Molecular mass markers and purified AcsA-AcsB are indicated with labels in lanes 5 and 10. Panel C: The band intensities of blots shown in Panel A and B were quantified using Image J and plotted as a percent of wild type for AcsA (black) and AcsB (grey). The insertional mutants are as labeled with a Δ. **Statistically significant difference in protein levels when compared to WT using Welch’s T-test, p-value < 0.01.

We also tested the acsA insertional mutant which showed 20 ± 4% the specific activity of wild type (Fig. 5.2). When compared to wild type, the acsA insertional mutant shows a statistically significant (p-value <0.01) reduction in specific activity, but not elimination of activity. However, western-blot analysis revealed no band present for

AcsA or AcsB (Fig. 5.4).

103

Figure 5.4. Western-Blot Analysis of acsA Insertional Mutant. Panel A, visualized with anti-AcsA3, is of a blot containing pure AcsA-AcsB (lane 1), wild-type TM (lane 2), and acsA insertional mutant TM (lane 3). Results show no AcsA band present for the acsA insertional mutant, indicated by the arrow. Panel B, visualized with anti- AcsB, is of a blot containing pure AcsA-AcsB (lane 4), wild-type TM (lane 5), and acsA insertional mutant TM (lane 6). Results show no AcsB band present for the acsA insertional mutant as indicated by the arrow.

G. hansenii possesses two additional cellulose synthase genes (98). These two gene products may be responsible for the residual cellulose synthesis activity measured in the acsA insertional mutant. As such, we performed product entrapment (Chapter 2) on the acsA insertional mutant. SDS-PAGE analysis of the pellet and visualization with

Coomassie-blue stain showed two bands (Fig. 5.5). Each band was excised and the proteins identified by mass spectrometry. Mass spectrometry analysis revealed that the proteins are products of the second cellulose synthase gene, acsAB2.

104

Figure 5.5. Isolation of AcsA-AcsB and AcsA2-AcsB2 by Product Entrapment. SDS-polyacrylamide gel visualized with Coomassie-blue stain showing protein isolated by product entrapment from wild type (lane 1) and from the acsA insertional mutant (lane 2). Product entrapment was performed as described in Chapter 2.

The Effects of AcsD and CcpAx on Purified AcsA-AcsB Activity

To further evaluate whether AcsD and CcpAx has any effect on in vitro cellulose synthase activity, both proteins were expressed heterologously in E. coli and purified by

Ni-affinity chromatography (Fig. 5.6A and B). Both proteins were added separately or together to a final concentration of 1 µM to assay mixtures containing 0.25 μM of purified AcsA-AcsB. The rate of cellulose synthesis was measured by the enzyme coupled assays monitoring NADH oxidation (Chapter 2). In all cases, there was no apparent effect on the turnover number for AcsA-AcsB under saturating substrate conditions (Table 5.1).

105

Figure 5.6. SDS-Polyacrylamide Gels of Heterologously Expressed and Purified CSC Components. AcsD (Panel A) and CcpAx (panel B) were expressed and purified as described in Materials and Methods. The purified protein was then subjected to SDS-PAGE analysis and visualized with Coomassie-blue stain.

Table 5.1. The Effect of CSC Proteins on the Rate of Cellulose Synthesis from Purified AcsA-AcsB.

Additions velocity, s-1

None 1.69 ± 0.09 AcsD 1.67 ± 0.04 CcpAx 1.61 ± 0.04 AcsD + CcpAx 1.78 ± 0.01

The Effects of AcsC, AcsD, and CcpAx Insertional Mutations on Cell Morphology

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We reasoned that if activity was not significantly impacted in the TM of the acsC and acsD insertional mutants, a possible explanation would be that these mutations interfered with the transport of the cellulose polymers out of the cell. To examine this possibility, we utilized TEM images. In comparison to wild type (Fig. 5.7A) the acsC insertional mutant had a noticeably expanded periplasm layer (Fig. 5.7C). The acsD insertional mutant, which makes lower levels of cellulose in vitro (13), did not appear to have an expanded periplasm (Fig. 5.7D). The ccpAx insertional mutant also does not appear to have an expanded periplasm (Fig. 5.7B). Attempts to quantify cellulose from the periplasm of the acsC insertional mutant were not successful.

Figure 5.7. Transmission Electron Micrographs of G. hansenii Cells. Whole cells from G. hansenii (A) wild type, (B) ccpAx insertional mutant, (C) acsC insertional mutant, and (D) acsD insertional mutant were fixed and stained as described in Materials and Methods. The expanded periplasm in the acsC insertional mutant cells is indicated by arrows. Subcellular Localization of CSC Components

107

Our product entrapment results from Chapter 2 did not show any other CSC proteins co-purifying with AcsA-AcsB. To investigate the potential interaction of AcsA,

AcsB, AcsC, and AcsD, the localization of these proteins was probed with western blotting and by activity assays (radiometric assay described in Chapter 2) after separation of the TM by sucrose density gradient ultracentrifugation.

The TM fraction of wild type cells was separated into the cytoplasmic membrane

(CM) and outer membrane (OM) over a sucrose density gradient as described in Material and Methods. Two opaque bands were visible after partitioning, and 13 fractions of 3 mL each were removed for analysis. Protein concentration was determined, by Lowry assay

(Chapter 2), for each fraction (Fig. 5.8A). Two opaque bands were primarily located in fractions 3-5 and fractions 10-12. The CM is localized in the lower density fractions and this was confirmed by the marker enzyme assay for succinate dehydrogenase (186) showing most of the activity in fractions 4-6 (Fig. 5.8C). Negligible succinate dehydrogenase activity in the lower band indicates little or no contamination of the CM in the OM fraction. In addition to the marker enzyme assay, each fraction was also subjected to SDS-PAGE and the proteins were visualized with Coomassie-blue stain (Fig

5.8B).

A previous study by Bureau et al. (147) and comparison with BcsA-BcsB from R. sphaeroides (19) predict that AcsA is anchored in the CM and AcsB, while anchored in the CM, is primarily located in the periplasm. This is also consistent with the substrate

UDP-glucose being found in the cytoplasm. Here, only the OM fractions catalyzed cellulose synthesis with little to no cellulose synthase activity detected in the CM fraction. A second, smaller peak corresponding to cellulose synthase activity can be seen

108 at fraction 8 (Fig. 5.8C). This activity is likely due to AcsA-AcsB in a membrane fraction of intermediate density.

Western blotting was used to identify the subcellular localization of the components of the CSC (Fig. 5.8D). Our results indicate AcsA, AcsB, and AcsC being primarily localized to the OM. This is consistent with the localization of cellulose synthase activity with the OM. AcsD is found throughout the entire gradient. This is consistent with AcsD being localized to the periplasm (103).

One possible interpretation of these results is that association of AcsA-AcsB with

AcsC causes partitioning of AcsA-AcsB to the OM. To test this, we fractionated TM from the acsC insertional mutant by sucrose density gradient. The results were identical to those found for wild type, with cellulose synthase activity again partitioning with the

OM (Fig. 5.9A). western-blot analysis reveals that both AcsA and AcsB again partition primarily with the OM (Fig. 5.9B).

109

-1

A -1 C 0.18 0.25 4 cellulose synthase

mol min 0.16 succinate dehydrogenase

mol min 0.20 3 0.14

-1 0.12 0.15 2 0.10 0.08 0.10

protein, mg mL mg protein, 1 0.06

0.04 0.05 0 0.02 260 130 0.00 0.00 100

72 cellulose synthase activity, 2 3 4 5 6 7 8 9 10 11 12 52 260

succinate dehydrogenase activity, 42 130 Anti-AcsC 35 100 Anti-AcsB

25 72 Anti-AcsA3

15 15 Anti-AcsD

10 B D

Figure 5.8. Analysis of G. hansenii TM by Sucrose Density Gradient. A total of 6 mL of TM was applied to 34 mL of a 25% to 55% sucrose gradient and separated by centrifugation. A total of 13 fractions of 3 mL each was removed and analyzed. Panel A shows protein concentration for each fraction as determined by Lowry assay described in Chapter 2. Panel B shows SDS-PAGE analysis of each fraction, visualized with Coomassie-blue stain. Panel C shows cytoplasmic marker enzyme, succinate dehydrogenase, activity (open circles) and cellulose synthase activity (closed circles). Panel D shows western-blot analysis of each fraction with the indicated primary antibodies on the right.

Cellulose Synthesis Turnover Number upon Cellular Fractionation

Our results do not indicate that AcsC, AcsD, or CcpAx have an impact on the turnover number for AcsA-AcsB. Therefore, we measured the turnover number at each step of product entrapment (Chapter 2) in order to track the decline in the turnover number from 25 s-1 in whole cells to 1.7 s-1 in purified AcsA-AcsB. We chose to use product entrapment, rather than affinity purification, because the strain expressing the histidine-tagged AcsA-AcsB does not express AcsC.

110

Activity was measured by 14C-glucose incorporation (Chapter 2) and AcsA-AcsB was measured by quantitative western blotting (Appendix B, Fig. B.8), using anti-AcsB, as described in Materials and Methods for the in vivo cellulose synthesis rate.

Our results (Table 5.2) indicate that the turnover number is 1.3 s-1 immediately after cell lysis (total lysate). In TM, the turnover number increases to 2.0 s-1, further increasing to 2.7s-1 after solubilization of the TM. A turnover number of 2.2 s-1 was measured in soluble membrane. The turnover number for entrapped AcsA-AcsB of 1.9 s-1 is nearly identical to AcsA-AcsB purified by column chromatography.

Figure 5.9. Analysis of acsC Insertional Mutant TM by Sucrose Density Gradient. A total of 6 mL of TM was applied to 34 mL of a 25% to 55% sucrose gradient and separated by centrifugation. A total of 13 fractions of 3 mL each was removed and analyzed. Panel A shows cytoplasmic marker enzyme, succinate dehydrogenase, (open circles) and cellulose synthase activity (closed circles). Panel B shows western-blot analysis of each fraction with the indicated primary antibodies.

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Table 5.2 Purification Table for Cellulose Synthase by Product Entrapment.

Volume Total Activity Total AcsA-AcsB Sample Protein (mg/mL) Specific Activity (nmol/min/mg) Turnover Number (s-1) (mL) (nmol/min) (nmol) Lysate 150 6.6 7.0 6982 89.9 1.3 TM 9 26.3 12.6 2982 25.4 2.0 TM 1h solubilization 16 13.2 18.6 3927 24.6 2.7 Solubilized 13 8.4 16.7 1822 13.9 2.2 Entrapped 2.5 n/a n/a 927 7.9 1.9 protein

Discussion

Determination of in vivo Cellulose Synthase Turnover Number

The turnover number for AcsA-AcsB in whole cells was determined over a short time period such that this time period did not result in cell doubling. This short time period assured that the quantification of AcsA-AcsB content by western blot provided an accurate value. The turnover number in whole cells (24 s-1) is much higher than the turnover number of the purified enzyme (1.7 s-1). The turnover number of cellulose synthase in G. hansenii whole cells was also measured by Brown et al. (32). Their value of 67 s-1 is also much greater than the turnover number of the purified enzyme. Brown et al. (32) measured the elongation rate of the cellulose ribbon from whole cells under dark field microscopy (33 nm per second). By dividing this rate by the length of a single glucose molecule (0.5 nm) a turnover number of 67 s-1 was calculated. Our measurements are also supported by Schramm and Hestrin (62) who calculated a rate of 2.0 × 105 glucose molecules incorporated into cellulose per cell per second. Their measurement is close to our rate of 3.8 × 105 ± 2.0 × 105 glucose molecules incorporated per cell per second.

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The turnover number for AcsA-AcsB, determined in whole cells, is 14-fold higher than the turnover number for purified AcsA-AcsB. It is nearly identical to the elongation rate for AcsA-AcsB calculated by GPC (25 s-1). Given our kinetic mechanism (Chapter

4), the two steps which are most likely to be affected in vivo versus with purified AcsA-

AcsB are priming (k’6) or polymer release (k’1). This means that the value for k’6 is potentially greater or the value k’1 for is potentially reduced in vivo (compared with purified enzyme). Given that the in vivo turnover number is nearly identical to the elongation rate, it is plausible that either of these steps are altered such that the steps associated with elongation then become rate limiting for the catalytic cycle. Such a change in one or more of the rate constants for these steps could proceed with the assistance of CSC accessory proteins.

The CSC is known to contain multiple accessory proteins necessary for cellulose synthesis in vivo (25, 58, 60, 90, 93). The processive enzyme, DNA polymerase, also complexes with multiple protein subunits, affecting both processivity and rate (172-175,

178). The E. coli polymerase, DNA Pol III α subunit, alone, synthesizes polymers of 11 nucleotides in length (173, 174). When complexed with the β (sliding clamp) and ε subunits, its processivity increases several hundred-fold (several thousand nucleotides incorporated between priming events) (172). The rate-limiting step for E. coli DNA Pol I is shown to be dissociation from the template strand (188). Therefore, by increasing the processivity, the overall rate of DNA synthesis increases (173, 174). The increase in the in vivo turnover number could similarly be explained by complexing of AcsA-AcsB with

CSC accessory proteins.

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To test for the effects of accessory proteins on the cellulose synthesis rate, we investigated the impact of the CSC proteins AcsC, AcsD, and CcpAx on cellulose synthesis by both mutagenesis studies and biochemical studies using purified proteins.

Our investigations did not ultimately yield evidence for the influence of these proteins on the turnover number of AcsA-AcsB. One important finding in this investigation, however, is the effect CcpAx has on the apparent stability of the AcsB protein.

The Effects of AcsC, AcsD, and CcpAx on AcsA-AcsB

Mutants for acsC, acsD, or ccpAx have shown reduced or no cellulose synthesis activity in vivo (60, 93). Thus, these proteins have been implicated in cellulose synthesis and they are thought to be part of the CSC. In our work, the TM from the ccpAx insertional mutant showed a significant reduction in specific activity as compared to TM from wild type. The specific activity in TM from the ccpAx insertional mutant is similar to that observed in the TM from the acsA insertional mutant. Western-blot analysis of the ccpAx insertional mutant TM showed a significantly reduced level of AcsB protein, explaining the reduced specific activity for this mutant. The apparent reduction in the level of AcsB, relative to AcsA, cannot be due to lower expression levels of AcsB. This is because these two proteins are translated as a single peptide (Chapter 2). The more likely explanation is rapid turnover of AcsB in the absence of CcpAx by mechanisms unknown.

TM from the acsD insertional mutant showed a slight reduction in specific activity when compared to TM from wild type. No reduction in the AcsA or AcsB protein levels, as measured by western blotting, was observed. This would suggest that AcsD

114 affects the catalytic activity of AcsA-AcsB, however, the addition of purified AcsD to purified AcsA-AcsB did not impact the turnover number. The addition of purified CcpAx to purified AcsA-AcsB did not impact the turnover number. The combination of both

CcpAx and AcsD added to purified AcsA-AcsB also did not affect the turnover number.

We saw no statistically significant reduction in the specific activity in the TM from the acsC insertional mutant compared with TM from wild type. TEM imaging of the acsC insertional mutant cells reveals an expanded periplasm, which is not present in wild type cells, the acsD insertional mutant cells, or the ccpAx insertional mutant cells. We hypothesize that this is due to the accumulation of cellulose in the periplasm. This hypothesis is consistent with AcsC being an outer membrane porin necessary for egress of cellulose to the extracellular milieu.

TM from the acsA insertional mutant shows 25% of the specific activity of wild type TM, despite showing no AcsA protein by western-blot analysis. The residual activity is likely due to the presence of two additional genes for cellulose synthase (60). Product entrapment and SDS-PAGE analysis on the acsA insertional mutant shows two bands, the proteins from which were confirmed by identification with mass spectrometry to be products of the acsAB2 gene. These two proteins are of a different apparent molecular weight than either AcsA or AcsB. They are also not visible, by SDS-PAGE analysis visualized with Coomassie-blue stain, in the product-entrapped protein from wild type

(Fig. 5.5). The amino acid sequences of the two additional cellulose synthases, AcsAB2 and AcsAB3, show 40% and 46% sequence identity, respectively, to AcsAB (60). Each of these additional acsAB genes have an associated acsC isoform within a single operon

(60). It’s somewhat perplexing then, that in vivo cellulose synthesis is not observed in the

115 acsA insertional mutant. The biological function of these two additional cellulose synthases remains unclear.

Subcellular Localization of CSC Components

Our results with the cellulose synthase activity being primarily localized in the

OM differs from that of Bureau et al. (147). This may be due to a trypsin treatment these workers employed prior to OM and CM separation (147). It is possible that, while degrading a range of membrane-spanning proteins, trypsin may also disrupt the interactions between the OM cellulose synthase components. The apparent partitioning of cellulose synthesis activity with the OM may be due to the association of AcsA-AcsB with one of these membrane-spanning proteins. Association with AcsC could explain this. However, our fractionation results from the acsC insertional mutant showed that cellulose synthase activity and AcsA and AcsB continued to partition primarily with the

OM.

Cellulose Synthesis Turnover Number upon Cellular Fractionation

Our results in this chapter show that the turnover number for AcsA-AcsB decreases from 24 s-1 in vivo to 1.3 s-1 immediately after lysis (total lysate). Our results also do not indicate that AcsC, AcsD, or CcpAx are responsible for the higher turnover number for AcsA-AcsB in whole cells.

In Chapter 2, we showed that AcsA-AcsB does not co-purify with any of the known CSC proteins either by product entrapment or affinity chromatography. This shows that CSC proteins do not associate with AcsA-AcsB in soluble membrane, and it

116 suggests that the association between AcsA-AcsB and other CSC proteins is relatively weak. The disruption of the CSC almost certainly occurs during cell lysis. This conclusion comes from three observations. First, cellulose synthesized by whole cells predominantly takes the form of the cellulose I allomorph (33, 189), while cellulose synthesized from TM predominantly takes the form of cellulose II (147). The ordered

CSC in whole cells, which synthesize glucan polymers unidirectionally (in the form of a ribbon), produces cellulose I. Cellulose II, the more thermodynamically favored allomorph, is produced by the randomly oriented (disrupted CSC) AcsA-AcsB molecules in the TM. It has also been suggested that AcsD plays a role in the formation of cellulose

I (72). Secondly, our sucrose density gradient results do not indicate that AcsC or AcsD, which is found throughout the gradient, are associating with AcsA-AcsB in TM. Finally, the turnover number for AcsA-AcsB in total lysate (1.3 s-1) is close to the turnover number in TM (2.0 s-1), suggesting that the conditions under which AcsA-AcsB is catalyzing cellulose synthesis are similar in both fractions.

One possible explanation behind the fragility of the CSC is that its association is contingent on cell integrity. Work by Deng et al. (59) shows that insertional mutations in genes coding for the enzymes lysine decarboxylase and alanine racemase alter the morphology and crystallinity of cellulose produced in vivo. Both of these enzymes have been implicated in the assembly of the peptidoglycan cell wall (59, 190). Indeed, the cell morphology in both of these mutants was visibly different by light microscopy imaging compared to wild type (59). This is likely due to distortions in the architecture of the peptidoglycan layer (191). Their findings suggest that perturbations in cell morphology, or perhaps the integrity of the peptidoglycan layer, impact the organization of the CSC.

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Recently, the plant cellulose synthase, CesA8 from Populus tremula, was expressed in

Pichia pastoras and purified (192). For CesA8, reconstitution into proteoliposomes is necessary to achieve activity (192). This is in contrast to bacterial cellulose synthase (69).

Also in contrast to bacterial cellulose synthase, the reconstituted proteoliposomes synthesize microfibrils (192). The formation of microfibrils are dependent upon a minimal ordering of the cellulose synthase enzymes (30, 92). One interpretation may be that spontaneous organization of CesA8s into a homomeric complex (enabled by the presence of a lipid bilayer) facilitate cooperativity between the enzymes which is necessary to achieve activity.

Like DNA polymerase, cellulose synthase may rely on accessory proteins and/or simply organization into the CSC (in the case for the plant enzyme) to maximize a step

(or steps) in the catalytic cycle, thus increasing the apparent turnover number in whole cells. Disruption of the CSC during cell lysis effectively removes the weakly associating accessory proteins from participating with AcsA-AcsB in cellulose synthesis. This results in the decrease in turnover number for AcsA-AcsB which is apparent after cell lysis.

Though a complete CSC has yet to be isolated from any cellulose-producing organism, understanding the role the protein-protein interactions play in the catalytic cycle for cellulose synthesis would greatly benefit from the purification of such an intact CSC.

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CHAPTER 6

SUMMARY AND FUTURE DIRECTIONS

The central focus of this work was to characterize the mechanisms of enzymatic cellulose synthesis. In this chapter, we present a consolidated review of the contributions to this goal. Moreover, we address areas for future investigations, which are afforded by the findings and methodological developments made in this thesis.

Methodological Contributions

The development of a system for the homologous expression of AcsA-AcsB from

G. hansenii (Chapter 2) was the initial goal of this work. It provides a tool for continuing in vitro and in vivo biochemical work. In purifying AcsA-AcsB we also corrected our earlier interpretation of AcsAB processing (97), reducing the active form from a heterotrimer to a heterodimer.

The major methodological contribution was the development of a technique for analyzing cellulose synthesized in vitro (Chapter 3). Though methods previously existed for the measurement of cellulose DOP, they require relatively larger amounts of material compared to what can be cost-effectively synthesized by purified protein. Therefore, we adapted the solubilization and separation methods for cellulose to fit our needs.

Specifically, we used synthesis of cellulose with UDP-14C-glucose (for detection), followed by solubilization in DMAc / 8% LiCl by solvent exchange (157), and finally analysis by gel permeation chromatography (GPC) (56, 57). This method allows for the measurement of the distribution of a cellulose population using relatively small amounts of sample.

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Initiation of Cellulose Synthesis Proceeds by a Primer-Independent Mechanism

In Chapter 3, through the use of GPC analysis, we showed that both AcsA-AcsB

(G. hansenii) and BcsA-BcsB (R. sphaeroides) operate by a primer-independent mechanism. GPC analysis showed that cellulose synthesized in vitro, from both enzymes, increased in size, eventually reaching a maximum (the processivity limit for the enzyme).

Once the maximum size of cellulose had been reached, cellulose synthase continued to incorporate glucose into cellulose; this can be accounted for by the appearance of new small species. Next, we used reducing end analysis to show that radio-labeled glucose was present on the reducing ends of in vitro-synthesized cellulose. This indicated that glucose initiates synthesis. Finally, using the glucose oxidase-linked enzyme assay, we demonstrated UDP-glucose hydrolase activity in BcsA-BcsB. This allowed us to construct a plausible mechanism for priming. Our mechanism proposes that cellulose synthase hydrolyzes UDP-glucose and subsequently binds the liberated glucose to initiate new polymer synthesis. Future investigations into initiation may benefit from transient- state kinetics studies using rapid-quench techniques (193).

Processivity in Cellulose Synthase

A comparison of the GPC product profiles from BcsA-BcsB and AcsA-AcsA revealed processivity differences between the two enzymes (Chapter 3). In Chapter 4, we made point mutations in selected residues which were part of the transmembrane channel of BcsA-BcsB in order to probe factors which affect processivity. We found one mutant

(F416A) with reduced processivity, overall turnover number, and elongation rate compared to wild type BcsA-BcsB. Because of the relevance of cellulose DOP to both

120 material and biofuels applications (8, 50, 51, 95, 168), this is an area predisposed to continuing research. In Chapter 4, we recognized that certain channel residues impact an enzyme’s processivity. Future work should seek to understand how the identity of different residues (acidic, basic, or hydrophobic) affect processivity, where along the length of the transmembrane channel the greatest affects become apparent, and identifying other domains within the enzyme which can be implicated in processivity; with the ultimate goal of synthesizing cellulose of a desired DOP. By swapping whole domains of enzymes from organisms known to produce cellulose of high DOP into expression systems such as BcsA-BcsB, the amino acid residues important to processivity can be systematically identified. The separation methods adapted herein are applicable to future studies.

A Minimal Kinetic Mechanism for Cellulose Synthesis

GPC analysis also gave us a means to calculate the turnover number independent of the enzyme concentration. In Chapter 3, rates of 6.6 s-1 and 27 s-1 were calculated for

BcsA-BcsB and AcsA-AcsB, respectively. These numbers were larger than the overall turnover number calculated for either enzyme, suggesting that there may be a slow cycle in catalysis. Further evidence for this hypothesis was gathered by comparing the activation energies for the overall incorporation of glucose to the activation energy for the elongation of the glucan polymer. Further refinement of the BcsA-BcsB elongation rate (refined to 4.1 s-1) and calculation of the second order rate constant for elongation was attained by measuring the elongation rates at varying UDP-glucose concentrations and fitting them to the Michaelis-Menten equation.

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Based on our findings, we constructed a kinetic mechanism for cellulose synthesis. Developing software which is capable of modeling many thousands of steps will allow for the refinement of the cellulose synthase kinetic mechanism and prediction of the product profiles for different cellulose synthases. Development of this type of modeling software ties into the future work described for understanding processivity, wherein the product profiles can be predicted for engineered cellulose synthases.

Cellulose Synthesis and the Cellulose Synthase Complex

The homologous expression system for AcsA-AcsB, developed in Chapter 2, gave us the opportunity to compare the kinetics of cellulose synthesis in vivo versus in vitro. In

Chapter 5 we determined the whole-cell turnover number to be 25 s-1, which is much higher than the turnover number for purified AcsA-AcsB (1.7 s-1). It is unknown which step or steps in the kinetic mechanism are affected. The difference in turnover number, however, may be due to a factor which is removed during purification. Next, we explored the roles of three CSC proteins (AcsC, AcsD, and CcpAx) to see if they had any impact on the overall turnover number. For this, we used insertional mutants generated by our collaborator, Ying Deng (59, 60). Our results did not yield any evidence implicating these three proteins in the catalytic cycle of cellulose synthase. However, we did find that

CcpAx, in addition to its importance to the integrity of the CSC (90), effects the stability of AcsB. As a consequence of the ccpAx insertional mutation, AcsB is degraded, resulting in the near abolition of cellulose synthesis activity from AcsA. The precise mechanism behind the degradation of AcsB is unknown. Future work on CSC accessory proteins should seek to provide a greater understanding of the roles AcsC, AcsD, and CcpAx in cellulose assembly and crystallization through understanding the assembly of the CSC.

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Using the antibodies in our possession in combination with crosslinking techniques and western blotting, the protein-protein contacts can be uncovered (97, 103, 194). By quantitative western blotting, the stoichiometry of the proteins in the G. hansenii CSC can be determined (24).

Immediately upon cell lysis we found that the overall turnover number for AcsA-

AcsB drops from 24 s-1 in vivo to 1.3 s-1. Based on this, we hypothesize that CSC integrity is important in the catalytic mechanism of cellulose synthesis. Whether the in vivo turnover number can be attributed to the precise configuration of the CSC or a specific accessory protein which is effectively removed upon cell lysis is unclear. The identity of any such cofactors also remains unclear. The immediate future work should identify which factors may be implicated in the in vivo turnover number. This is not a simple matter in light of the tenuous association between the CSC proteins. Successful measurement of the in vivo turnover number for knock-out mutants in CSC accessory proteins may offer a clue as to whether certain accessory proteins are important in catalysis. Careful analysis of the DOP of cellulose synthesized in vivo (from wild type G. hansenii and the insertional mutants), and in vitro (from purified AcsA-AcsB) may help to identify the step in the kinetic mechanism which is different in vivo compared with in vitro synthesis. It is the recommendation of this author that such analysis be carried out by the carbanilation and GPC techniques described in Chapter 3. Ultimately, the grand goal is to develop a method for the isolation of an intact cellulose synthase complex for biochemical and structural studies.

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APPEDICES

Appendix A: List of Primers

Name Chapter Sequence Description AcsAB his F 2 CGACTCTAGAGGATCCGTCGGC Forward primer for AcsAB flanking CGATCATGTCAGTGA region for the AcsAB-his construct. AcsAB his R 2 CTAGTGGTGATGATGGTGATGG Reverse primer for AcsAB flanking TGGTGATGATGGTGATGTG region for the AcsAB-his construct. ACTTGCGCCTCTCATCCTCA His TetC-F 2 CATCATCACCACTAGTTCTCATG TetC gene forward primer the TTTGACAGCTTATCA AcsAB-his construct.

His TetC-R 2 CCATGATTACGAATTCGGTTGGT TetC gene reverse primer the AcsAB- TTGCGCATTCACA his construct.

F301/416frag 4 GAGCTCGTGGTGGTGTTCGATG Fragment 1 forward primer for BcsA- 1F CCGACCACGTC BcsB F301A, W383A, N412A, F416A, W417A, Q463A, and V551A mutants. F301/416frag 4 CCGCCCACGACGAGGAGGACC Fragment 2 reverse primer for BcsA- 1R GACCGGT BcsB F301A, W383A, N412A, F416A, W417A, and Q463A. V551frag2R 4 GGGCGAGAAGGCGCGGCGTCA Fragment 2 reverse primer for BcsA- GG BcsB V551A mutant. F301Frag1R 4 GTGGATCTTGCCGTAGGCCATC Fragment 1 reverse primer for BcsA- TCGTTCTCGGG BcsB F301A mutant. F301Frag2F 4 CCCGAGAACGAGATGGCCTACG Fragment 2 forward primer for BcsA- GCAAGATCCAC BcsB F301A mutant. W383Frag1R 4 CATCATGCCGGTGGCGAAGCGG Fragment 1 reverse primer for BcsA- CCGCGCTGCTG BcsB W383A mutant. W383Frag2F 4 CAGCAGCGCGGCCGCTTCGCCA Fragment 2 forward primer for BcsA- CCGGCATGATG BcsB W383A mutant. N412Frag1R 4 CCAGAAGCTCATCGAGGCGAGG Fragment 1 reverse primer for BcsA- TAGCACAGGCG BcsB N412A mutant. N412Frag2F 4 CGCCTGTGCTACCTCGCCTCGAT Fragment 2 forward primer for BcsA- GAGCTTCTGG BcsB N412A mutant. F416Frag1R 4 CAGCGGGAAGAACCAGGCGCT Fragment 1 reverse primer for BcsA- CATCGAGTTGAG BcsB F416A mutant. F416Frag2F 4 CTCAACTCGATGAGCGCCTGGT Fragment 2 forward primer for BcsA- TCTTCCCGCTG BcsB F416A mutant. W417Frag1R 4 CACCAGCGGGAAGAACGCGAA Fragment 1 reverse primer for BcsA- GCTCATCGAGTT BcsB W417A mutant. W417Frag2F 4 AACTCGATGAGCTTCGCGTTCTT Fragment 2 forward primer for BcsA- CCCGCTGGTG BcsB W417A mutant.

124

Q463Frag1R 4 CGCAAACAGCGCGTTCGCCACG Fragment 1 reverse primer for BcsA- AGGAAGCTCAC BcsB Q463A mutant. Q463Frag2F 4 GTGAGCTTCCTCGTGGCGAACG Fragment 2 forward primer for BcsA- CGCTGTTTGCG BcsB Q463A mutant. V551Frag1R 4 GCCCACGACGAGGAGGGCCGA Fragment 1 reverse primer for BcsA- CCGGTCGCCGGG BcsB V551A mutant.

V551Frag2F 4 CCCGGCGACCGGTCGGCCCTCC Fragment 2 forward primer for BcsA- TCGTCGTGGGC BcsB V551A mutant.

AcsD-LF 5 CGACTCTAGAGGATCCCTGGCG Forward primer for left flanking GGTGTATCTACTGCTGA region of AcsD for the AcsD insertional mutant. AcsD-LR 5 TGTCAAACATGAGAACAGAAGC Reverse primer for left flanking GCGTTCAGTTCGATCT region of AcsD for the AcsD insertional mutant. AcsD-RF 5 CTGCGCATCGTGCATGAAAACC Forward primer for the right flanking T region of AcsD for the AcsD insertional mutant. AcsD-RR 5 CCATGATTACGAATTCTATATTG Reverse primer for the right flanking GGGTGTTGCCAGCCCAT region of AcsD for the AcsD insertional mutant. AcsDTetCF 5 TTCTCATGTTTGACAGCTTATCA Forward primer for the tetC gene for the AcsD insertional mutant. AcsDTetCR 5 ATGCACGATGCGCAGGGTTGGT Reverse primer for the tetC gene for TTGCGCATTCACA the AcsD insertional mutant. ccpAx F 5 GGTGGCATATGACCAAGACAGA Forward primer for the ccpAx ccpAx- CACGAATTCT his construct. ccpAx R 5 CACCACTCGAGTTAGGATTCTTC Reverse primer for the ccpAx ccpAx- TTCGTTTTCACG his construct. AcsD F 5 TGGTGGCATATGACAATTTTTGA Forward primer for the AcsD-his GAAAAAACCG construct. AcsD R 5 CCACCACTCGAGTCAGGTCGCG Reverse primer for the AcsD-his GAACTG construct.

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Appendix B: Supplemental Figures

Figure B.1. Chromatograms for the Analysis of the Products of Edman Degradation. The arrows indicate the peaks corresponding to the major amino acid present in each cycle.

126

Figure B.2. SDS-PAGE Analysis of Purified BcsA-BcsB. SDS-polyacrylamide gel visualized with Coomassie-blue stain of BcsA (~89 kDa) and BcsB (~84 kDa).

0.5

0.4

-1 0.3

0.2

Velocity, s Velocity,

0.1

0.0 0 2 4 6 8 10 12 UDP-glucose, mM

Figure B.3. Steady-State Kinetic Analysis of BcsA-BcsB. Effect of UDP-glucose concentration on enzyme velocity. Rates are expressed as velocity (s−1). Data shown is from mean and standard deviation of three separate assays.

127

1.2 10000

1.0 1000 100 0.8 Bacterial 10 0.6 cellulose Avicel 1 0.4 0.1

0.2 0.01

Degree of Polymerizaiton of Degree

Refractive Index, Relative Index, Refractive

0.0 0.001 4 6 8 10 12 Elution Volume, mL

Figure B.4. GPC Elution Profiles of Native Cellulose Dissolved in DMAc / 8% LiCl. Cellulose from G. hansenii cultures and Avicel were dissolved directly in DMAc / 8% LiCl as described in Chapter 2. The samples were applied to a KD-806M column coupled to a DAWN HELIOS II multi angle light scattering detector and a t-REx refractometer used for molecular weight determination. The DOP distributions (circles) are shown above their associated elution peak. DOP distributions were used for column calibration as described in Chapter 3.

60 No Addition AcsA-AcsB BcsA-BcsB 40

Cellulose(nmol glucose) 0 6 8 10 12 Elution Volume, ml

Figure B.5. Incubation of Pre-Synthesized Cellulose with Cellulose Synthase. Cellulose was synthesized from 0.5 µM BcsA-BcsB with 5 mM UDP-14C-

128 glucose in a reaction volume of 1.0 ml as described in Chapter 3. To determine whether AcsA-AcsB (open circles) and BcsA-BcsB (filled circles) preparations contained cellulase activity, the cellulose was incubated with these enzymes for 16 hours in the absence of substrate. The figure shows the GPC elution profiles of the cellulose in DMAc / 8% LiCl as described in the legend.

160 modified cellulose 140 unmodified cellulose 120 100 80 60 40

Cellulose (nmol glucose) 20 0 6 7 8 9 10 11

Elution Volume, mL

Figure B.6. GPC Elution profiles of AB-modified cellulose. Cellulose was synthesized from 0.5 µM of BcsA-BcsB with 1 mM UDP-14C-glucose in a reaction volume of 0.5 ml for 30 min as described in Chapter 3. To determine whether modification caused hydrolysis, the cellulose was modified with aminobenzamide (AB) as described in Chapter 3. Cellulose preparations were dissolved in DMAc / 8% LiCl. The figure shows the GPC elution profiles of the cellulose in DMAc / 8% LiCl as described in the legend.

129

F416A Q463A 0.08 0.35

0.30 0.06 0.25

-1

-1 0.20 0.04 0.15

Velocity, s Velocity, Velocity, s Velocity, 0.10 0.02 0.05

0.00 0.00 0 2 4 6 8 10 12 0 2 4 6 8 10 12 UDP-glucose, mM UDP-glucose, mM V551A N412A 0.20 1.2

1.0 0.15

-1 0.8 -1 0.10 0.6

Velocity, s Velocity, 0.4 s Velocity, 0.05 0.2

0.0 0.00 0 2 4 6 8 10 12 0 2 4 6 8 10 12 UDP-glucose, mM UDP-glucose, mM

Figure B.7. Steady-State Kinetic Analysis of BcsA-BcsB Point Mutants. Effect of UDP-glucose concentration on enzyme velocity. Rates are expressed as velocity (s−1).

A B 1 2 3 L 4 5 6 7 8 6000 260 5000 130 100 4000

72 3000

2000

52 Intensity Band 1000 42 0 35 0.0 0.2 0.4 0.6 0.8 1.0 AcsB, pmol

Figure B.8. AcsB Quantification by Western-Blot Analysis. Cellulose synthase was quantified from total lysate (A, lane 4), total membrane (A, lane 5), total membrane (solubilized 1h) (A, lane 6), soluble membrane (A, lane 7), and entrapped protein (A, lane 8) by western blotting with anti-AcsB using purified AcsA-AcsB as standards (A, lanes 1-3). Band intensities were quantified by Image J analysis (B). Standards are shown as filled circles and samples as filled diamonds.

130

Appendix C: Tenua Input

EA0 + UG <-> UGEB0 <-> EB1 + U ; EB1 <-> EA1 ; EA1 + UG <-> UGEB1 <-> EB2 + U ; EB2 <-> EA2 ; EA2 + UG <-> UGEB2 <-> EB3 + U ; EB3 <-> EA3 ; EA3 + UG <-> UGEB3 <-> EB4 + U ; EB4 <-> EA4 ; EA4 + UG <-> UGEB4 <-> EB5 + U ; EB5 <-> EA5 ; EA5 + UG <-> UGEB5 <-> EB6 + U ; EB6 <-> EA6 ; EA6 + UG <-> UGEB6 <-> EB7 + U ; EB7 <-> EA7 ; EA7 + UG <-> UGEB7 <-> EB8 + U ; EB8 <-> EA8 ; EA8 + UG <-> UGEB8 <-> EB9 + U ; EB9 <-> EA9 ; EA9 + UG <-> UGEB9 <-> EB10 + U ; EB10 <-> EA10 ; EA10 + UG <-> UGEB10 <-> EB11 + U ; EB11 <-> EA11 ; EA11 + UG <-> UGEB11 <-> EB12 + U ; EB12 <-> EA12 ; EA12 + UG <-> UGEB12 <-> EB13 + U ; EB13 <-> EA13 ; EA13 + UG <-> UGEB13 <-> EB14 + U ; EB14 <-> EA14 ; EA14 + UG <-> UGEB14 <-> EB15 + U ; EB15 <-> EA15 ; EA15 + UG <-> UGEB15 <-> EB16 + U ; EB16 <-> EA16 ; EA16 + UG <-> UGEB16 <-> EB17 + U ; EB17 <-> EA17 ; EA17 + UG <-> UGEB17 <-> EB18 + U ; EB18 <-> EA18 ; EA18 + UG <-> UGEB18 <-> EB19 + U ; EB19 <-> EA19 ; EA19 + UG <-> UGEB19 <-> EB20 + U ; EB20 <-> EA20 ; EA20 + UG <-> UGEB20 <-> EB21 + U ; EB21 <-> EA21 ; EA21 + UG <-> UGEB21 <-> EB22 + U ; EB22 <-> EA22 ; EA22 + UG <-> UGEB22 <-> EB23 + U ; EB23 <-> EA23 ; EA23 + UG <-> UGEB23 <-> EB24 + U ; EB24 <-> EA24 ; EA24 + UG <-> UGEB24 <-> EB25 + U ; EB25 <-> EA25 ; EA25 + UG <-> UGEB25 <-> EB26 + U ; EB26 <-> EA26 ; EA26 + UG <-> UGEB26 <-> EB27 + U ; EB27 <-> EA27 ; EA27 + UG <-> UGEB27 <-> EB28 + U ; EB28 <-> EA28 ; EA28 + UG <-> UGEB28 <-> EB29 + U ; EB29 <-> EA29 ;

131

EA29 + UG <-> UGEB29 <-> EB30 + U ; EB30 <-> EA30 ; EA30 + UG <-> UGEB30 <-> EB31 + U ; EB31 <-> EA31 ; EA31 + UG <-> UGEB31 <-> EB32 + U ; EB32 <-> EA32 ; EA32 + UG <-> UGEB32 <-> EB33 + U ; EB33 <-> EA33 ; EA33 + UG <-> UGEB33 <-> EB34 + U ; EB34 <-> EA34 ; EA34 + UG <-> UGEB34 <-> EB35 + U ; EB35 <-> EA35 ; EA35 + UG <-> UGEB35 <-> EB36 + U ; EB36 <-> EA36 ; EA36 + UG <-> UGEB36 <-> EB37 + U ; EB37 <-> EA37 ; EA37 + UG <-> UGEB37 <-> EB38 + U ; EB38 <-> EA38 ; EA38 + UG <-> UGEB38 <-> EB39 + U ; EB39 <-> EA39 ; EA39 + UG <-> UGEB39 <-> EB40 + U ; EB40 <-> EA40 ; EA40 + UG <-> UGEB40 <-> EB41 + U ; EB41 <-> EA41 ; EA41 + UG <-> UGEB41 <-> EB42 + U ; EB42 <-> EA42 ; EA42 + UG <-> UGEB42 <-> EB43 + U ; EB43 <-> EA43 ; EA43 + UG <-> UGEB43 <-> EB44 + U ; EB44 <-> EA44 ; EA44 + UG <-> UGEB44 <-> EB45 + U ; EB45 <-> EA45 ; EA45 + UG <-> UGEB45 <-> EB46 + U ; EB46 <-> EA46 ; EA46 + UG <-> UGEB46 <-> EB47 + U ; EB47 <-> EA47 ; EA47 + UG <-> UGEB47 <-> EB48 + U ; EB48 <-> EA48 ; EA48 + UG <-> UGEB48 <-> EB49 + U ; EB49 <-> EA49 ; EA49 + UG <-> UGEB49 <-> EB50 + U ; EB50 <-> EA50 ; EA50 + UG <-> UGEB50 <-> EB51 + U ; EB1 <-> EA0 + T1 ; EB2 <-> EA0 + T2 ; EB3 <-> EA0 + T3 ; EB4 <-> EA0 + T4 ; EB5 <-> EA0 + T5 ; EB6 <-> EA0 + T6 ; EB7 <-> EA0 + T7 ; EB8 <-> EA0 + T8 ; EB9 <-> EA0 + T9 ; EB10 <-> EA0 + T10 ; EB11 <-> EA0 + T11 ; EB12 <-> EA0 + T12 ; EB13 <-> EA0 + T13 ; EB14 <-> EA0 + T14 ; EB15 <-> EA0 + T15 ; EB16 <-> EA0 + T16 ; EB17 <-> EA0 + T17 ; EB18 <-> EA0 + T18 ; EB19 <-> EA0 + T19 ; EB20 <-> EA0 + T20 ;

132

EB21 <-> EA0 + T21 ; EB22 <-> EA0 + T22 ; EB23 <-> EA0 + T23 ; EB24 <-> EA0 + T24 ; EB25 <-> EA0 + T25 ; EB26 <-> EA0 + T26 ; EB27 <-> EA0 + T27 ; EB28 <-> EA0 + T28 ; EB29 <-> EA0 + T29 ; EB30 <-> EA0 + T30 ; EB31 <-> EA0 + T31 ; EB32 <-> EA0 + T32 ; EB33 <-> EA0 + T33 ; EB34 <-> EA0 + T34 ; EB35 <-> EA0 + T35 ; EB36 <-> EA0 + T36 ; EB37 <-> EA0 + T37 ; EB38 <-> EA0 + T38 ; EB39 <-> EA0 + T39 ; EB40 <-> EA0 + T40 ; EB41 <-> EA0 + T41 ; EB42 <-> EA0 + T42 ; EB43 <-> EA0 + T43 ; EB44 <-> EA0 + T44 ; EB45 <-> EA0 + T45 ; EB46 <-> EA0 + T46 ; EB47 <-> EA0 + T47 ; EB48 <-> EA0 + T48 ; EB49 <-> EA0 + T49 ; EB50 <-> EA0 + T50 ; EB51 <-> EA0 + T51 ;

Values used for Tenua Simulation of Steady-State Kinetics.

k1, M-1 s-1 k2, s-1 k3, s-1 k4, s-1 k5, M-1 s-1 k6, s-1 AcsA-AcsB 200 0.035 0.00021 25 500,000 90 BcsA-BcsB 20 0.015 0.001 5 14,000 10

Values used for Tenua Simulation of Mock Distributions.

k1, M-1 s-1 k2, s-1 k3, s-1 k4, s-1 k5, M-1 s-1 k6, s-1 3000 1.5 0.4 (varied) 5 (varied) 1,500,000 100

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John B. McManus

Education

Ph.D., Biochemistry and Molecular Biology The Pennsylvania State University August 2010 – August 2017

B.S., Biochemistry and Molecular Biology The Pennsylvania State University August 2003 – August 2007

Work Experience

Sapphire Energy, San Diego, CA, 2009 – 2010

Charles River Labs, Malvern, PA, 2007 – 2008

Department of Biochemistry, Penn State University, University Park, PA, 2006 – 2007

Agricultural Research Service, Beneficial Insects Research Laboratory, Newark, DE, 2002 – 2004

Teaching Experience

Department of Biochemistry, Penn State University Park, PA, 2011, 2014

Awards and Grants

Paul M. Althouse Outstanding Teaching Assistant Award, 2011

NRC Research Associate Program Postdoctoral fellow, 2017

Publications

Iyer PR, Liu YA, Deng Y, McManus JB, Kao TH, Tien M. Processing of cellulose synthase (AcsAB) from Gluconacetobacter hansenii 23769. Arch. Biochem. Biophys. 2013;529:92-8.

Nagachar N, and McManus JB. Microbial Cellulose Synthesis. Microbial Factories: Biofuels, Waste Treatment. 2015; 1: 203-216, Springer Publishing

McManus JB, Deng Y, Nagachar N, Kao TH, Tien M. AcsA-AcsB: The core of the cellulose synthase complex from Gluconacetobacter hansenii ATCC23769. Enzyme Microb. Technol. 2016; 82:58-65