The Pennsylvania State University

The Graduate School

Department of Biochemistry and Molecular Biology

BIOCHEMICAL CHARACTERIZATION OF THE PROTEINS INVOLVED IN

GLUCONACETOBACTER HANSENII CELLULOSE SYNTHESIS

A Dissertation in

Integrative Biosciences

by

Radhakrishnan Iyer Prashanti

 2012 Radhakrishnan Iyer Prashanti

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2012

The dissertation of Radhakrishnan Iyer Prashanti was reviewed and 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

Nicole R. Brown

Associate Professor of Wood Chemistry

Charles T. Anderson

Assistant Professor of Biology

Peter J Hudson

Program Chair, Integrative Biosciences

Director, The Huck Institutes of the Life Sciences

* Signatures are on file in the Graduate School

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ABSTRACT

Gluconacetobacter hansenii is a Gram-negative bacterium, considered as the model organism for studying the process of cellulose biogenesis. This is due to its unique ability to synthesize and secrete copious amounts of cellulose as an extracellular polysaccharide, in its growth medium. G. hansenii is therefore an ideal bacterium to study cellulose as a material as well as cellulose synthesis as a biological process. We have therefore employed this bacterium as our subject for our inquiry into the biochemistry of the process of cellulose synthesis.

In this work, the main area of focus is towards understanding the bacterial cellulose synthesis and secretion complex, in terms of its component proteins, their structure, organization and their interactions. Two parallel approaches were used to understand and gain insights into the cellulose synthesis complex. One was to heterologously express and purify the proteins that are known to be involved in cellulose synthesis for structural studies or for generation of antibodies. Another method was to isolate the cellulose synthase complex from the G. hansenii cells and dissect its component proteins to reveal as-yet-unknown constituents of this complex.

The cellulose synthase operon encodes for three proteins, AcsAB, AcsC and

AcsD. AcsD protein was heterologously expressed and purified. The pure protein was employed for structural characterization as well as for antibody generation. Using the specific anti-AcsD antibody, the subcellular localization of this protein was identified to be the periplasmic space. Studies of AcsD using gel filtration, analytical ultracentrifugation and dynamic light scattering revealed that it exists as an octamer in solution. Structural characterization of AcsD using small angle-X-ray scattering reveals

that, in accordance with its crystal-structure, the protein forms a complex of a tetramer of dimers that assumes a cylindrical conformation with a central pore.

The predicted non-membrane regions of the cytoplasmic membrane-bound cellulose synthase (AcsAB) protein were heterologously overexpressed and purified using affinity methods. Specific antibodies generated against these regions revealed that the protein, though encoded by a single gene, is actually processed into three polypeptides AcsA (45 kDa), AcsB1 (34 kDa) and AcsB2 (95 kDa). Western blot of the fractions from a sucrose density centrifugation combined with sequence-based analysis revealed that the AcsB2 protein is localized in the periplasmic region of the bacterial cell.

The genome of G. hansenii 23769 was sequenced to provide a database for mass- spectrometry based-proteomic studies of cellulose synthesis. The completed genome is now a public database in NCBI. These studies involved Multidimensional Protein

Identification Tool (MudPIT) analysis of the total membrane (TM), outer membrane

(OM) and the cytoplasmic membrane (CM) fractions of the cells for the comparison of the proteomic profile of the three compartments. This revealed that the AcsB, AcsC and the AcsD were largely concentrated in the OM whereas the CM compartment contains lower abundance of AcsAB protein.

Using blue native polyacrylamide gel electrophoresis (BN-PAGE), the protein complexes in solubilized G. hansenii TM were isolated and the complex containing the proteins involved in cellulose was located using the specific antibodies against AcsD and

AcsA. As revealed by LC-MS analysis of the antibody cross-reacting gel-band, this complex also contains phosphoglucomutase, glucose-6-phosphate isomerase and UDP- glucose pyrophosphorylase. These proteins were known to be involved in cellulose iv

biosynthesis pathway, but this work presents evidence for the existence of these proteins in association with the proteins involved in the cellulose synthesis and secretion complex.

In addition to using gel-based methods for identifying the components of the cellulose synthesis complex, zymography was used to demonstrate in-vitro cellulose synthesis using detergent solubilized membranes. This study directs towards a greater efficiency of the detergent dodecyl maltoside compared to Triton-X 100, in solubilization of the TM, while retaining the enzyme-activity.

In summary, we have used a combination of traditional and modern biochemical approaches to study the protein components of the cellulose synthesis machinery. Our work has resulted in a sequenced genome, structural analysis and localization of AcsD, and identification of processing of the AcsAB. We have also presented evidence that the proteins involved in the cellulose biosynthetic process, indeed exist as a complex and have identified other proteins relevant to the process of cellulose synthesis, to be components of this complex. Based on our findings, we have proposed our model for the bacterial cellulose synthesis and extrusion complex.

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TABLE OF CONTENTS

LIST OF ABBREVIATIONS ...... x LIST OF FIGURES…………………………...... xi LIST OF TABLES ...... xiii ACKNOWLEDGEMENTS ...... xiv CHAPTER I BIOCHEMISTRY OF CELLULOSE SYNTHESIS ...... 1 Cellulose and its impact on our lives ...... 1 Chemical structure of cellulose ...... 4 Physical properties of cellulose ...... 6 G. HANSENII AS THE MODEL ORGANISM FOR CELLULOSE SYNTHESIS ………………… ...... 9 Bacterial cellulose: properties and uses ……...... 11 Visualization of cellulose-synthesizing complexes ...... 13 Uridine diphosphate glucose (UDP-glucose) ...... 15 Cyclic diguanylate (c-di-GMP) ...... 16 In vitro cellulose synthesis in presence of the regulator ….…...... 19 Identification of the genes involved in cellulose synthesis ...... 20 BACTERIAL CELLULOSE SYNTHASE OPERON……………...... 22 acsAB ………...... 23 AcsC ...... 25 AcsD...... 26 acs operon is flanked by genes that modulate cellulose synthesis ...... 27 dgc and pdeA genes ...... 28 Cellulose biosynthetic pathway and mechanism of cellulose synthesis ..….… 30 STATEMENT OF THE PROBLEM ...... 33 SUMMARY ...... 34 CHAPTER II WHOLE GENOME SEQUENCING OF GLUCONACETOBACTER HANSENII 23769...... 38 INTRODUCTION ...... 38 STEPS IN GENOME SEQUENCING……..…...... 40 General outline of sequencing protocol ...... 40

reparation of single stranded DNA library ...... 41 Amplification of the library by emulsion PCR ...... 41 Sequencing by synthesis (Pyrosequencing) ...... 41 Paired-end library generation ...... 43 SOLid sequencing ...... 45 ANALYSIS OF THE SEQUENCE USING SOFTWARE TOOLS ...... 46 Assembly ...... 46 Generation of contigs ...... 46 Scaffolds ...... 47 Gene prediction and annotation ...... 48 MATERIALS AND METHODS ……..……….…...... 49 Isolation of genomic DNA ..…...... 49 Processing and analysis of the sequenced data using software tools …...... 51 RESULTS ……….…...... 55 Assembly metrics ...... 55 Finishing, annotation and databank entry ...... 55 Genome features ……….……...... 56 Features relevant to cellulose synthesis …...... 56 Proteomic analysis of the membrane compartments ……...... 57 DISCUSSION ….…...... 60 CONCLUSIONS …………………………………………………..……….... 62 CHAPTER III LOCALIZATION OF THE ACSD PROTEIN IN THE PERPIPLASM OF G. HANSENII CELLS …….…...... 63 INTRODUCTION ...... 63 MATERIALS AND METHODS ………...... …...….……65 Bacterial strains and culture conditions …...... …66 AcsD cloning, expression and protein purification ……...... 66 Protein expression and purification ….…….…...... 68 Antibody preparation …...…...... ….. 69 Preparation of membrane fractions ...... 69 Preparation of periplasmic and cytoplasmic fractions ...... 70

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Detection of AcsD using Western blot ….…...... 72 RESULTS ……...... 72 Purification of AcsD ...... 73 Specificity of anti-AcsD antibody …...... …..... 73 Subcellular fractionation ...... 74 Detection of AcsD in the periplasmic fraction ...... 75 DISCUSSION...... 77 CONCLUSION ……...... 79 CHAPTER IV DETERMINATION OF THE SOLUTION-STRUCTURE OF ASCD …………...... 81 INTRODUCTION...... 81 Principles behind structural analysis by SAXS…...... 84 MATERIALS AND METHODS.…...... 85 AcsD overexpression and purification …...... …. 85 Sample preparation for AUC ...... 86 DLS experiment …………...... 86 Analytical ultracentrifugation ………...... …..... 86 Gel filtration ……………...... 88 Sample preparation for SAXS ...... 88 Data acquisition ………..…...... 89 SAXS data analysis …...... 90 RESULTS ………...... 92 DLS experiment …...... ………. 92 Gel filtration ……………..…...... 94 Sedimentation velocity experiments ……...... ….. 95 Determination of the AcsD structure using SAXS…...... ……..99 DISCUSSION ……….……..……….…………….…...... 107 CONCLUSION ……….……..……….……...... …. . 109 CHAPTER V BIOCHEMICAL CHARACTERIZATION OF ACSAB, THE CELLULOSE SYNTHASE PROTEIN ...... 110 INTRODUCTION …...... 110

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MATERIALS AND METHODS ...... 110 Cloning and heterologous expression of the acsAB gene regions …..…...... …113 Expression of the AcsAB soluble regions ...... 114 Protein purification using denaturing method ...... 115 Antibody generation, purification and Western blot ...... 117 Sucrose density gradient centrifugation …...... … 118 RESULTS ……...... 118 Predicting the soluble regions of the AcsAB protein …...... 118 Western blot using anti-AcsAB1 and anti-AcsAB2 antibodies ...... 119 Western blot using anti-peptide antibodies ...... 119 Sucrose density gradient centrifugation …...... … 122 DISCUSSION ...... 124 CONCLUSIONS ...... 128 CHAPTER VI ISOLATION OF THE CELLULOSE SYNTHASE COMPLEX USING ELECTROPHORETIC TECHNIQUES ...... 129 INTRODUCTION ...... 129 MATERIALS AND METHODS …...... 131 Solubilization of the TM proteins …………...... 131 Native gel electrophoresis of the solubilized TM proteins ……...... 132 Zymography .…...... 133 Blue native polyacrylamide gel electrophoresis (BN-PAGE) ……….….…...... 133 RESULTS ……...... 139 Zymography for in vitro cellulose synthase activity ….………...... 139 Selection of an efficient detergent for BN-PAGE ……….……...... 139 BN-PAGE of DDM-solubilized TM …….……...... ,...... 140 DISCUSSION ……...... 144 CONCLUSIONS …...... 147 CHAPTER VII SUMMARY AND FUTURE DIRECTIONS ...... ….…. 148 REFERENCES ...... 153

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ABBREVIATIONS Acs: Acetobacter cellulose synthase

AUC: Analytical ultracentrifugation

BN-PAGE: Blue native polyacrylamide gel electrophoresis

Brij 58: Polyethylene glycol hexadecyl ether c-di-GMP: Cyclic di-guanosine monophosphate

CM: Cytoplasmic membrane

DDM: Dodecylmaltoside

EDTA: Ethylene diamine tetraacetate

DLS: Dynamic Light Scattering

IPTG: Isopropyl thiogalactopyranoside

Ni-NTA: Nickel nitriloacetate

OM: Outer membrane

MudPIT: Multidimensional Protein Identification Tool

PCR: Polymerase chain reaction

SDS-PAGE: Sodium dodecyl sulphate electrophoresis

SAXS: Small angle X-ray scattering

SDS: Sodium dodecyl sulphate

TM: Total membrane

UDP-glucose: Uridine diphosphate glucose

WC: Whole cells

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LIST OF FIGURES

Figure 1.1 Chemical structure of cellulose …………...... 5

Figure 1.2 Turnover of c-di-GMP in bacterial cells ...... 18

Figure 1.3 Structure of cellulose synthase operon in related strains

of Acetobacter ………………………………………………………………. 22

Figure 2.1 Steps involved in a genome sequencing project...... 39

Figure 2.2 Generation of a DNA library by paired-end method ………...... 44

Figure 2.3. Agarose gel electrophoresis of genomic DNA …………...... 50

Figure 2.4 Subsystem catalogue of the genes ………...... 59

Figure 3.1 Cloning of the acsD gene …...... 66

Figure 3.2 Overexpression and purification of recombinant AcsD ….…...... 69

Figure 3.3 Determining the specificity of the anti-AcsD antibody ……..….. 73

Figure 3.4 Subcellular fractionation and detection of AcsD ……………...... 76

Figure 3.5 The amino acid sequence of AcsD …………………………..….. 77

Figure 4.1 SAXS experiment ………………………………….………….…. 83

Figure 4.2 Anion exchange chromatography of AcsD …………………….... 87

Figure 4.3 DLS analysis of AcsD ...... 92, 93

Figure 4.4 Gel filtration of AcsD …...... 94

Figure 4.5a Sedimentation coefficient distributions: An overlay showing normalized distribution plots for AcsD…………..……...... 96

Figure 4.5b,c Continuous sedimentation coefficient distribution c(s) ………..98

Figure 4.6a Experimental scattering profiles ..…………….….…….…….... 99

Figure 4.6b Buffer subtracted scattering curves for AcsD ………………..... 100

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Figure 4.6c Guinier plot ..…………………………………………………...... 101

Figure 4.7 Plot for pair distribution function derived using GNOM ………….. 102

Figure 4.8 Representative bead models of AcsD generated by GASBOR...... 104,105

Figure 4.9: Comparison of the contours of the solution structure and the crystal structure of AcsD ………………………………………….………….…106

Figure 5.1 Depiction of the heterologously expressed regions of AcsAB protein ...... 113

Figure 5.2 Agarose gel of amplified products of acsAB gene regions ……………..116

Figure 5.3 SDS-PAGE of heterologously-expressed AcsAB polypeptides ……….. 121

Figure 5.4 Western blot using specific polypeptide and peptide antibodies ………. 121

Figure 5.5 Graph for Molecular weight determination of processed

AcsA polypeptide …………………………………………………….... 123

Figure 5.6 Western blot of fractions from sucrose density gradient of TM …. ……125

Figure 5.7a Analysis of AcsB2 sequence using Signal P (Gram negative) ...... 125

Figure 5.7b Lipo P prediction of a signal sequence in the AcsB polypeptide …….. 126

Figure 6.1. Zymogram of detergent-solubilized G. hansenii TM ……………….... 138

Figure 6.2 Comparison of the second dimension gel profiles ……………………... 141

Figure 6.3 BN PAGE of G. hansenii TM…..………...... 142

Figure 7.1 Working model for cellulose synthesis complex ………………………. 152

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LIST OF TABLES

Table 2.1 Proteins relevant to cellulose biosynthesis identified by MudPIT. …. ….. 59

Table 3.1 Marker enzyme assays for sub-cellular fractions ….…...... 74

Table 3.2 Protein identification by LC-MS of trypsin-digested 17kDa gel band…... 77

Table 6.1a Composition of polyacrylamide gradient BN-gel …………………….. 135

Table 6.1b Buffers for BN-PAGE ………………………………………………..... 136

Table 6.2 Proteins detected after LC-MS of the BN-gel band ………………...... 143

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ACKNOWLEDGEMENTS

When I think about my life as a graduate student, I feel like I have been looking at the world of research, standing on the shoulders of giants. I am fortunate to have had mentorship from outstanding scientists, within and outside Penn State. I am thankful to my advisor Dr. Ming Tien whose academic experience and able guidance have been invaluable for me. He has always motivated me in order to bring out the best in me. I would like to extend my sincere thanks to Dr. Tracy Nixon, Dr. Nicole Brown and Dr.

Charles Anderson, for being part of my thesis committee. I am extremely thankful to Dr.

Nicole Brown for being very supportive and for introducing me to the world of cellulose synthesis. I am very grateful to Dr. Charles Anderson for accepting to be a part of my committee at the very last minute. This kind gesture by him, will always be remembered.

I have had the good fortune of working with Dr. Tracy Nixon, whose easy grasp of structural analysis of proteins, have helped me through my learning curve on this subject. His patience and dedication while teaching me and training me during the SAXS experiments, are something I will remember forever and would like to imbibe in my own teaching methods. It is a dream-come-true, to be guided by the experts in the field of plant cell wall themselves, Dr. Candace Haigler and Dr. Daniel Cosgrove. Through the platform of the CLSF, they have given me the opportunity to present my work in front of varied audiences in numerous conference presentations. Their kindness, encouragement and faith in me, has helped me more than they know.

I would like to acknowledge Dr. Teh-Hui Kao and Dr. Jeffrey Catchmark for having fruitful discussions with me over the years. I also want to thank Dr. Yara

Yingling, Dr. James Kubicki, Dr. Alan Esker and Dr. Linghao Zhang for lifting-up my

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spirits before all my presentations and treating me like their own student. A heartfelt thanks goes to Laura Ullrich and Liza, who have been such a good friends. Laura took care of all the arrangements for my presentations and even made room reservations for my thesis defense.

I am really thankful to my colleagues who have enriched my research through their inputs. My initial training in the Tien lab was by Dr. Shin Sato, who trained me in the area of fungal biochemistry, while I was a rotation student in this lab. He trained me in techniques that I now routinely use in my work, and am extremely indebted to him for initiating me to lab work. I am very grateful to Dr. Scott Geib who was instrumental in teaching me the ropes of bioinformatic techniques required for genome sequencing. This work would not have been possible without his guidance. My dear friends and senior graduate students Camille Stephens and Tatyana Sysoeva have always been a good support throughout my graduate life. I am greatly thankful to them for teaching me crucial biochemistry skills. I would also like to thank everyone in Tein lab for the being a very supportive group of friends.

The road to my graduate career was a winding one. But I would have not taken this path without the love for learning which was instilled in me, very early in my life.

My family and my teachers had envisioned my life in academics, much before I knew about myself. My teachers have been instrumental in motivating me to perform better than I thought I am capable of. I thank my teachers, Mrs. Kumar, Mrs. Sophy Verghese,

Mrs. Sreeja Prakash, Mrs. Aparna Kulkarni, Mrs. Vimala Srinivasan, Mrs. P. Katre, Mrs

D. Majumdar, Mrs.Kulkarni, Dr. DN Mishra, Dr. A. Maniyar, Dr. T. Sheikh, Dr. Rama

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Kannan, Mrs. Aparna Rajagopalan and Mrs Radhika. You saw in me what I did not know about myself.

In my life I have had the privilege of some of the greatest teachers. My first teachers were my parents, uncles, aunts and grandparents. Through their innumerable story-telling sessions, reading time and family discussions, I was initiated into my informal education. My parents are extremely ambitious for me and have provided me with the best of education at the expense of their own comfort. I am grateful to my dear sister Priya for teaching me through her example, to be dedicated to a cause and to achieve it against all odds. My parents chose an equally loving and caring family for me to get married into. The affection and blessings showered on me by my mother- and father-in-law, is the greatest treasure in my life. Without the encouragement and motivation from my in-laws (parents-, uncles-, aunts-, sisters- and brothers-) and my husband, this Ph.D. would have been just a dream. I have learnt the art of enjoying research from my husband, who has taught me, that enthusiasm and passion are the biggest skill-sets of a good scientist. My conversations with my family members were my source of rejuvenation and strength throughout my graduate life. It is their goodness of heart that has translated into my good fortune. To this wonderful family of amazing people, I dedicate this thesis.

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CHAPTER I

BIOCHEMISTRY OF CELLULOSE SYNTHESIS

Cellulose and its impact on civilization

Cellulose is a homopolysaccharide that is the most abundant naturally-occurring macromolecular polymer on earth, being produced at the rate of 180 billion tons per year

(Brown and Montezinos 1976). Cellulose is the major component of the plant cell wall where it is embedded in a matrix of lignin, hemicelluloses and pectin. On average, cellulose comprises around 45% of the plant lignocellulosic biomass but the content of this polymer varies with the plant type (Matrone, Ellis et al. 1946; Meier 1964;

Toyoshima, Onda et al. 1990). In woody plants, cellulose constitutes almost half the weight of the biomass (Meier 1964), whereas in grasses the content of cellulose is roughly 20-30% (Matrone, Ellis et al. 1946; Toyoshima, Onda et al. 1990). The secondary cell walls of cotton, with their cellulose content of almost 100% are the purest form of cellulose in nature (Itoh 1990; Haigler 2007). The predominance of cellulose in the plant kingdom makes it the major constituent of biomass, an abundant renewable resource and a significant contributor to the global carbon cycle.

Its distribution together with its unique properties, has made the use of cellulose widespread. Humans have been exploiting the predominance of this natural material for centuries and have invented several applications for cellulose-derived materials, making cellulose-based products a quintessential part of our daily lives. Its unique properties of high tensile strength, flexibility and recalcitrance, make it an ideal material for lumber, paper, textiles and many commodities. The extreme dependence of mankind on cellulosic products continues in present day society. This has put a severe burden on forestry and

agriculture and has lead to long-term adverse effects on the planet like deforestation and loss of indigenous flora.

An obstacle in the use of plant-based cellulosic material, is that for many of its applications, a pure form of cellulose is desired. But, the lignin and hemicellulosic network of plant cell walls are not easily removed or circumvented. Furthermore, conversion of cellulose is impeded by the inherent recalcitrance of cellulose. Purification of cellulose therefore, involves several steps of harsh alkaline treatments and/or the use of sulfites to digest the lignin and free the cellulose away from cell wall polymers. This process of purifying cellulose from plant material is a highly energy-demanding, water- consuming and polluting process (Canadian Environmental Protection Act Priority

Substances List Assessment 1991). A new term, "paper pollution", has been coined to refer to the environmental hazards of the pulp-milling process, which is the third largest industrial contributor to pollution, causing land, water and air pollution (Teruyama, Itoh et al. 1990). The complete abolishment of the use of cellulosic materials is not likely, thus, sustainable and environmentally-friendly methods for cellulose purification are of relevance.

In addition to its traditional uses in paper and lumber industries, at present the major focus on cellulose is due to its potential use as the starting material for bioethanol production. Cellulose is envisioned as a future source for biofuels, in the form of cellulosic ethanol which is touted to substitute petroleum as a transport fuel (Sticklen

2008). Cellulosic is derived from non-edible portions of renewable feed stocks like corn stover and straw, or from non-food sources like agricultural wastes, bagasse, sugarcane and wood (2007). In addition to being environment-friendly, it provides a greater net

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energy benefit and a lower green house emission than corn-based ethanol (Energy

Conservation Board). Current research on cellulose is largely directed towards genetically-engineering plants in order to obtain large quantities of cellulosic biomass without using much land space. Such approaches include but are not restricted to, augmenting the rate of cellulose synthesis in the plant cell walls (Andersson-Gunneras,

Mellerowicz et al. 2006), making cellulosic material less crystalline (Abramson,

Shoseyov et al. 2010) and modifying the chemistry of other wall polymers like lignin and hemicellulose with the aim of weakening their matrix and making cellulose more accessible (Ragauskas, Williams et al. 2006; Chen and Dixon 2007; Abramson, Shoseyov et al. 2010).

Finding alternatives to the process of chemical pulping, altering the properties of cellulose and enhancing its biosynthesis require an in depth understanding of the plant cell wall architecture as well as knowledge of the biochemical pathways leading towards the cell wall polymer synthesis and incorporation. Specifically, this endeavor necessitates a thorough inquiry into the structure, mechanical properties and the process of cellulose biogenesis. My dissertation work, though not directly applicable to the production of biofuels, is directed towards understanding the cellulose biosynthetic machinery. The enzymes and structural proteins contributing to the biosynthesis of cellulose, have a profound impact in the chemistry and morphology of the final product. Thus, mastering the technology to derive enhanced cellulose production in plant cells requires that we acquire adequate information on the process of cellulose biosynthesis. This would not only help to understand how nature produces this polymer, but also enable us to modify or engineer the properties of cellulose for various applications.

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Chemical structure of cellulose

The properties of any polymer are largely dictated by its chemical constituents and structure. As shown in Figure 1.1, cellulose is a linear polymer of β-1,4-linked glucose residues. This makes the basic unit of cellulose, a dimer glucose, called cellobiose, where each glucose is rotated at an angle of 180C with respect to its neighboring residue. As shown in Figure1.1b, the straight chains of cellulose, have a directionality conferred upon by the anomeric carbon. The end of the cellulose chain with the unlinked anomeric carbon is the reducing end and one with an exposed C-4 is the non-reducing end. This directionality is a critical factor in determining the mechanism of polymer synthesis and extension, which will be discussed in this chapter in the section on

“Cellulose biosynthetic pathway and mechanism of cellulose synthesis”.

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Figure 1.1 Chemical structure of cellulose a) Glucose is a six-carbon sugar with two forms based on the orientation of the hydroxyl group of the anomeric carbon, C-1. The α-form has the hydroxyl group on the opposite side of the ring from the -CH2OH group and ß-glucose has the hydroxyl group on the same side as the -CH2OH group. b) Cellobiose is a dimer of ß-D-glucose where C-1 of one sugar is linked to the C-4 of the other by an acetal linkage. c) Cellulose is a linear polymer of ß-1,4-linked glucose units.

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Another feature of cellulose conferred upon it by the β-1,4 linkage, that differentiates cellulose from other glucan polymers like starch (α-1,4 linked branched, helical polymer) and callose (β-1,3 linkage), is the formation of extended, straight, unbranched chains with hydroxyl groups at C-2, C-3 and C-5 positions available for formation of intra- and inter-chain hydrogen bonds and van der Waals interactions

(Brown and Montezinos 1976; Montezinos and Brown 1976). These forces govern hierarchical associations of cellulose fibers to highly energy-minimized, para-crystalline forms that are insoluble yet very flexible and possess the tensile strength equivalent to steel (Niklas 1992), making it the strongest organic molecule on density basis

(Yamanaka, Watanabe et al. 1989).

Physical properties of cellulose

The basic unit of a cellulose fiber aggregate is an termed as a microfibril, a name that was originally given for the thinnest strands of cellulose observed under electron microscope (Hakoshima, Itoh et al. 1990). Cellulose microfibrils are classified based on their unit size, spectral properties, degree of polymerization and orientation of fibers. The size of the microfibrils, varies from 2-10 nm in plants to 30nm in Spirogyra (Hahne,

Herth et al. 1983; Hausser and Herth 1983).

The cellulose chains can be arranged in different orientations in the microfibril, giving rise to different crystalline forms (Brown and Montezinos 1976; Montezinos and

Brown 1976; Montezinos and Brown 1976) of cellulose, known as cellulose I and cellulose II allomorphs (Kaihoh, Itoh et al. 1990). Majority of the cellulose in nature occurs as the cellulose I allomorph (Chiou, Chen et al. 1990; Kaihoh, Itoh et al. 1990).

Using specific silver staining for reducing ends and cellobiohydrolase-mediated

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digestion, it was shown that all the glucan chains in cellulose I are oriented parallel to each other, with reducing ends of all the chains pointed in one direction (Herth 1983).

Anti-parallel arrangement of the cellulose chains gives rise to the cellulose II allomorph, the most thermodynamically stable form, which is a rare occurrence in nature (Brown

1996). Other than the marine alga Halocystis (Mulisch, Herth et al. 1983) and Gram positive bacterium Sarcina (Canale-Parola 1970), which produce the cellulose II allomorph naturally, this form of cellulose is known to be produced exclusively in cases where the ordered arrangement of cellulose fiber is perturbed due to dye addition (Chang and Itoh 1990), mercerization (Chanzy and Roche 1975), strong alkaline treatment or mutations in cellulose producing proteins (Kuga, Takagi et al. 1993; Chen and Brown

1996). An extra hydrogen bond for each glucose residues contributes to the extreme stability of cellulose II (Itoh, Oneil et al. 1984).

Their distinctly specific spectral signatures help in differentiating the allomorphs and sub-allomorphs of cellulose. Cellulose I can be further categorized into two allomorphs, Iα and Iβ (Kaihoh, Itoh et al. 1990; Ohchi, Itoh et al. 1990). Fiber diffraction and 13C-NMR studies on these sub-allomorphs, revealed that cellulose Iα has a single chain, triclinic unit cell, which further confirms the parallel chain model and Iβ is a monoclinic unit cell with two chains (Brown and Montezinos 1976; Chiou, Chen et al.

1990). Based on a crude estimates, (Kaihoh, Itoh et al. 1990), Acetobacter cellulose and cotton cellulose contain approximately 60-70% Iα and Iβ respectively. Thus the Iα form is predominant in prokaryotic cellulose, in which cellulose biosynthesis is part of the cell cycle and the Iβ form predominates in plant cellulose where secondary cell wall cellulose is produced after cell division is completed and the wall therefore possesses a

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complex architecture. Electron diffraction pattern of the entire length of microfibrils of the alga Microdictyon tenuis revealed regions that were purely Iα or Iβ or a mixture of the two forms (Chiou, Chen et al. 1990). This led to the conclusion that all naturally- occurring celluloses are a mixture of Iα and Iβ-allomorphs in varying ratio (Chiou, Chen et al. 1990). Since Iα-cellulose is meta-stable and more reactive than Iβ cellulose (Chiou,

Chen et al. 1990), the presence of both forms in different ratios accounts for the differential reactivity of native celluloses obtained from different sources, leading to substantial variation in the crystal packing and hydrogen bonding patterns, that influences their physical properties (Chiou, Chen et al. 1990).

Though there is considerable interspecies variation in the composition and size of cellulose, the allomorphic distribution and dimensions of the microfibrils remain consistent for a given species at a particular stage of its life cycle. Electron micrograph images show that the dimensions of the microfibrils are reflected by the number of chains in each microfibril, which is in turn determined by the number and pattern of the cellulose synthesizing complex subunits involved in the synthesis (Brown and

Montezinos 1976; Zaar 1979; Okuda, Tsekos et al. 1994; Reiss, Katsaros et al. 1996).

The structural information of cellulose is crucial for its characterization as well as for speculating on and developing models for the mechanisms of its synthesis. Similarly, the process and factors involved in cellulose biosynthesis, leave an imprint on the final morphology and pattern of cellulose formed.

G. HANSENII AS THE MODEL ORGANISM FOR CELLULOSE SYNTHESIS

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Other than plants, may species across the various kingdoms of life possess cellulose biosynthetic ability. Cellulose synthesis is a rare but existent phenomenon in animals, belongibg to the family of urochordates (Toyoshima, Onda et al. 1990). The mycetozoan Dictyostelium discoidum (Tsuji, Itoh et al. 1990) and the protozoans of

Acanthamoeba species (Wanibe, Yokoyama et al. 1990) also exhibit cellulose synthetic abilities. Although not a component of the wall, cellulose synthesis as part of primary metabolism is observed in bacterial species such as Acetobacter xylinum (Ross, Mayer et al. 1991), Rhizobium leguminosarum (Kitagawa, Kanamori et al. 1990; Mukai, Toba et al. 1990), Klebsiella pneumoniae (Nomura, Harino et al. 1990), Sarcina ventricle(Ross,

Mayer et al. 1991), Agrobacterium tumifaciens (Matthysse, White et al. 1995),

Salmonella typhimurium (Hatta, Baba et al. 1990) and Escherichia coli (Hatta, Baba et al.

1990) and cyanobacteria (Ayaki, Fujikawa et al. 1990). Some prevalent organisms that have been used a model systems for studies on cellulose synthesis are Gossypium hirsutum (cotton), Oryza sativa (rice), Arabidopsis thaliana (plant), Physcomitrella

(moss), Valonia (algae), and Acetobacter xylinum, Gluconacetobater hansenii (bacteria).

Among the bacterial species, organisms of the Acetobacter / Gluconacetobacter genus stand out due to their ability to synthesize and extrude copious amounts of highly pure ribbons of cellulose. One among these is G. hansenii, that forms the subject of study in this dissertation. In a recent taxonomic shuffle (Lisdiyanti, Navarro et al. 2006), some strains of Acetobacter xylinum were placed under the genus Gluconacetobater. Therefore, the present day nomenclature of some of the cellulose producing strains of Acetobacter xylinum is Gluconacetobacter hansenii. This change in nomenclature has resulted in the strain used in this work, ATCC 23769 being classified as G. hansenii.

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G. hansenii, a Gram negative, obligate aerobic bacterium has been considered for several years, as an archetype for cellulose synthesis-related studies. A single cell can polymerize 200,000 glucose molecules per second (Hestrin and Schramm 1954), which are extruded in the form of a 100 nm wide, flat ribbon of cellulose along the longitudinal axis of the cell (Brown, Willison et al. 1976; Akiyama, Yamada et al. 1990) and remain attached to the cells during cell division (Marx-Figini 1982; Ring 1982). When cultured under static conditions, the cellulose released forms a thick mat that accumulates at the air-liquid interface of the culture medium (Schramm and Hestrin 1954). This visible film, known as a pellicle, floats on the surface and completely covers the culture medium

(Schramm and Hestrin 1954; Schramm and Hestrin 1954). It is composed of cellulose fibers enmeshed with bacterial cells (Schramm and Hestrin 1954). When cultivated under agitated conditions, the increased oxygen tension facilitates faster growth of cells and the cellulose produced is observed as round balls (Schramm and Hestrin 1954).

Cellulose production is directly proportional to the cell density of the culture and as much as 50% of the available carbon is assimilated into the cellulose (Schramm and

Hestrin 1954; Kamide, Matsuda et al. 1990). The bacterial population contains several strain variants that overproduce cellulose and form a thicker aggregate in shaking culture and a thicker film in the stationary medium (Williams and Cannon 1989). These populations are transient and revert back to their cellulose synthesizing ability upon transfer into a static culture. Several generations of sub-culturing is required to obtain permanent cellulose non-producing mutants (Valla and Kjosbakken 1982).

There have been many attempts to understand the utility of an extracellular polysaccharide matrix. The most common notion is that the pellicle functions as a

10

flotation device to keep the aerobic cells in close proximity to the atmosphere (Schramm and Hestrin 1954; Cook and Colvin 1980). Another interesting observation is that when, co-cultured with molds and other bacterial species on fruits, the cellulosic film provided competitive advantage to the Acetobacter cells over others in the ability to colonize the substrate and protected the cells from being invaded by other species (Williams and

Cannon 1989). When observed under an electron microscope, these pellicles displayed a regular arrangement of tunnel-like lacunae along the surface of microfiber aggregates, suggesting the possibility of higher order in the organization of cellulose microfibrils

(Thompson, Carlson et al. 1988).

Bacterial cellulose: properties and uses

G. hansenii cellulose fibers contain microfibrils with average diameter of 20-40Å

(Akiyama, Yamada et al. 1990). Bacterial cellulose is 64% 1α (Kaihoh, Itoh et al. 1990;

Yoshida, Morisaki et al. 1990). Being an extracellular, inert and highly pure form of cellulose, it is an easy material to isolate and study as compared to the plant cellulose.

Other obvious advantages of using a bacterial system versus a plant system are: non- requirement of specialized culture conditions, faster growth cycle and ease of mutant generation and isolation.

The unique combination of high mechanical strength, extreme flexibility, in addition to its insolubility, biocompatibility, elasticity, mechanical strength and resistance to degradation, has made bacterial cellulose an ideal material of choice for biomedical applications (Ross, Mayer et al. 1991; Czaja, Kawecki et al. 2004; Czaja, Romanovicz et al. 2004). Bacterial cellulose is an anisotropic network of fibers, with a high degree of hydration, making it a suitable scaffold for seeding epithelial cells after severe skin injury

11

and therefore the material of choice to serve as an artificial skin graft for burn victims

(Czaja, Kawecki et al. 2004; Czaja, Romanovicz et al. 2004). This skin-substitute could be used in future, in lieu of an autograft (Cheung, Neikirk et al. 1990). Native and modified bacterial cellulose have been used as scaffolds for tissue engineering of cartilage (Yamaguchi, Ohsawa et al. 1990), blood vessels (Tobita, Kusama et al. 1990) and cardiac valves (Kuno, Kamisaki et al. 1990). Bacterial cellulose also finds use in electronic paper (Shah and Brown 2004) and acoustic diaphragms in earphones (Becker,

Itoh et al. 1990). Thus, bacterial cellulose synthesis has implications much beyond its assumed role as a model to study plant cellulose synthesis and could be a potential replacement for the latter in many of its uses. Since, many of its applications are in the biomedical and specialty material sectors, the knowledge of the mechanism of bacterial cellulose synthesis would be an important contribution to the basic as well as applied sciences.

It was using A. xylinum, that the major breakthroughs in the study of cellulose synthesis and characterization were achieved. Some of these milestones, which are discussed in detail in subsequent sections, are listed as follows,

1. The first successful isolation and cloning of a cellulose synthase gene (Saxena,

Lin et al. 1990). The first plant gene encoding cellulose synthase was identified based in its homology to the bacterial gene (Pear, Kawagoe et al. 1996).

2. Demonstration of high rates of in vitro cellulose synthetic activity (Glaser 1958; Aloni,

Delmer et al. 1982)

3. Identification of a four gene operon encoding for proteins involved in cellulose synthesis (Wong, Fear et al. 1990)

12

4. Determination of conserved residues within the catalytic domains of cellulose synthase protein (Saxena, Lin et al. 1990; Saxena, Henrissat et al. 1995)

5. Identification of c-di-GMP as a regulator for cellulose synthesis (Ross, Weinhouse et al. 1987)

6. Demonstration of Calcofluor as a dye to bind to and alter cellulose properties (Haigler,

Brown et al. 1980)

7. Model for cellulose biogenesis as a coupled process of polymerization and crystallization (Benziman, Haigler et al. 1980)

8. Electron microscopic observation of a linear array of complexes involved in active cellulose synthesis (Brown, Willison et al. 1976; Akiyama, Yamada et al. 1990)

9. Simulation of the assembly of higher plant cell walls (Yamaguchi, Ohsawa et al. 1990)

Some of the above-mentioned findings will be elaborated in the subsequent sections.

Visualization of the cellulose-synthesizing complexes

Biosynthesis of cellulose has been studied using various tools and from different perspectives. One of the earliest techniques employed to characterize cellulose synthesis was microscopy. Roelofsen (Roelofsen 1958) proposed as early as 1958 that enzyme complexes located at the growing tip of the cellulose chain, were involved cellulose synthesis and polymerization. Using freeze- fracture electron microscopy, Brown et. al.

(Brown, Willison et al. 1976) observed a single linear array of particles involved in cellulose synthesis on the outer membrane (OM) of A. xylinum. The site of emergence of the cellulose fibers on the surface of the cells is called a terminal complex (TC), named so because freeze-fracture electron microscopic analysis revealed the globular protein complexes on plant cell walls at the termini of cellulose fibrils (Brown and Montezinos

13

1976; Montezinos and Brown 1976)as predicted by Roelofsen (Roelofsen 1958;

Montezinos and Brown 1976).

After the discovery of the proteins involved in cellulose synthesis, it was possible to use immunological techniques to ascertain that the TC observed by microscopic methods in the past were indeed cellulose synthases (Kimura, Laosinchai et al. 1999).

Sodium dodecyl sulfate-solubilized freeze fracture replica labeling (SDS-FRL) was used to visualize the A. xylinum TC labeled with antibodies generated against cellulose synthase proteins (Kimura, Chen et al. 2001). The gold-labeled antibodies revealed that a linear row of 12-nm particles was localized in the inner side of the OM, which correspond to the ring-shaped pores observed in the exoplasmic side of the OM. These antibodies convincingly proved that the TCs contained proteins involved in cellulose biogenesis.

TCs show a great diversity in their arrangement in different organisms. Freeze- fracture studies on plant and algal cell walls revealed that the TCs were organized in the form of a six-membered rosette in land plants and green algae (Kiermayer and Sleytr

1979), (Giddings, Brower et al. 1980). In many algae, the TCs are arranged in the form of larger, rectangular arrays synthesizing cellulose ribbons (Katsaros, Reiss et al. 1996;

Reiss, Katsaros et al. 1996). Linear arrays of TCs are observed in prokaryotic bacteria and certain red and brown algae (Zaar 1979). The correlation between the size of a complex and the dimensions of the emerging cellulose were calculated by Herth (Herth

1983). These studies were corroborated by evidence presented by Okuda et. al. (Okuda,

Tsekos et al. 1994), by reviewing different types of TC arrangement and cellulose sizes.

It has been observed, as could be predicted intuitively, that the cellulose chains emerging

14

from linear TCs have a flat ribbon-like morphology, where as those from plant rosettes are cylindrical in shape (Itoh 1990; Itoh and Kimura 2001). Thus, it can inferred that the diversity of microfibril dimensions, seen across species and kingdoms, arises from the number of cellulose chains constituting the fibril, which in turn is governed by the number and arrangement pattern of the TCs. Factors that determine the ordering of TCs into a linear or a rosette pattern, can be identified only through biochemical and structural analysis of the proteins constituting them.

Nucleotide derivatives that drive cellulose synthesis: Uridine diphosphate glucose

(UDP-glucose) and cyclic di guanosine monophosphate (c-di-GMP)

A significant contribution to the understanding of cellulose synthesis was the

Nobel-winning discovery of sugar-nucleotides as “high energy molecules” by Leloir et al.

(Caputto, Leloir et al. 1950; Murai, Saito et al. 1990). Leloir et al. (Leloir, Olavarria et al.

1959; Leloir and Goldemberg 1960; Leloir 1961)elucidated the role of sugar-nucleotide

UDP-glucose in the synthesis of glycogen and thereby showed that biological polysaccharide synthesis reactions were not the reversal of degradation reactions, as was assumed previously (Caputto, Leloir et al. 1950; Leloir and Cardini 1957; Murai, Saito et al. 1990). Using glycogen synthesis as an example it was shown that all other polysaccharide synthesis reactions are in fact transfer reactions where the sugar from the sugar-nucleotide is transferred to the polymer which increases in length with each addition and the enzymes catalyzing these reactions were termed as glycosyl transferases

(Leloir 1961). In case of bacterial cellulose, the UDP-glucose for cellulose synthesis is provided by UDP-glucose pyrophosphorylase (Swissa, Aloni et al. 1980).

Cyclic diguanylate (c-di-GMP) : the unique activator of cellulose synthesis

15

The first-ever demonstration of in vitro cellulose synthesis activity was achieved as early as 1958 by Glaser (Glaser 1958), in particulate membrane fractions prepared from G. hansenii cells. However, it was in 1982 that high rates of synthesis in cell-free extracts were obtained by the Benziman group (Aloni, Delmer et al. 1982). However the most importan contribution of this work was that it led to the discovery of c-di-GMP

(Ross, Weinhouse et al. 1987).

Cyclic nucleotides (cAMP and cGMP) have been known to be crucial components of prokaryotic and eukaryotic signal transduction pathways (Karpen 2004).

Cyclic di-GMP was included in the list of second messengers after its biological role was elucidated by Benziman and co-workers, as the factor that allosterically activates cellulose synthesis in A. xylinum (Ross, Mayer et al. 1990). Eventually, several workers revealed the involvement of c-di-GMP in regulation of a wide array of cellular functions that influence virulence, pilus formation, cell cycle, antibiotic secretion and biofilm formation (Dow, Fouhy et al. 2006; Fouhy, Lucey et al. 2006; Jenal and Malone 2006;

Ryan, Fouhy et al. 2006; Cotter and Stibitz 2007; Pratt, Tamayo et al. 2007; Tamayo,

Pratt et al. 2007; Wolfe and Visick 2008). In general, it is noted that c-di-GMP is involved in quorum sensing and favors switching of a motile, planktonic lifestyle to a sessile on (Romling, Gomelsky et al. 2005). At low concentrations. c-di-GMP promotes a motile phenotype and at high concentrations it stimulates a sessile, biofilm-associated lifestyle (Cotter and Stibitz 2007; Pratt, Tamayo et al. 2007; Wolfe and Visick 2008).

Thus, the discovery of c-di-GMP is a milestone not just in the field of cellulose synthesis but also in all areas of bacterial signaling (Romling, Gomelsky et al. 2005; Ryan, Fouhy et al. 2006). Intracellular concentrations of c-di-GMP are maintained by the action of two

16

types of enzymes, diguanylate cyclases (Dgc) and phosphodiesterases (Pde) (Tal, Wong et al. 1998; Ausmees, Mayer et al. 2001; Ryan, Fouhy et al. 2006). Under cellular conditions, Dgc and Pde coordinately control c-di-GMP levels (Figure 1.2) to affect a target protein which acts as a switch to control the bacterial behaviour (Jenal and Malone

2006; Pratt, Tamayo et al. 2007; Tamayo, Pratt et al. 2007). In case of Acetobacter, the target protein is cellulose synthase which is activated by c-di-GMP, thereby promoting the process of cellulose synthesis (Ross, Mayer et al. 1990).

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Figure 1.2 Turnover of cyclic di-GMP in bacterial cells The cellular levels of c-di- GMP are maintained by the concerted action of two enzymes: diguanylate cyclase (DGC) and phosphodiesterase (PDE). These activity of these enzymes are regulated based on the environmental conditions. The DGCs have a conserved GGDEF motif in their active site (Ausmees, Mayer et al. 2001) and produce cyclic di-GMP by cyclization of two molecules of GTP (Paul, Weiser et al. 2004; Ryjenkov, Simm et al. 2006). Degradation of cyclic di-GMP molecules into two GMPs is mediated by PDE. These enzymes contain an EAL or a HD-GYP motif in their active site. The enzymes with EAL domain linearize the c-diGMP to form 5’pGpG, (Schmidt, Ryjenkov et al. 2005; Tamayo, Tischler et al. 2005) which is further cleaved to form two GMP molecules by non-specific PDEs (Christen, Christen et al. 2005; Romling, Gomelsky et al. 2005). The HD-GYP domain- containing proteins break the phosphodiester linkage in cyclic di-GMP first to form 5’- pGpG which is further cleaved to into two (Leloir, Olavarria et al. 1959; Leloir and Goldemberg 1960; Leloir 1961) GMPs by the same enzyme (Ryan, Fouhy et al. 2006).

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In vitro cellulose synthesis in presence of its regulator

A systematic work by Bureau and Brown in 1984 (Bureau and Brown 1987), showed for the first time that the cellulose synthetic activity was localized in the CM of

Acetobacter. The TM of the bacterium was separated into CM and OM fractions by sucrose-density gradient centrifugation after solubilizing the membrane fractions with lysozyme and trypsin(Bureau and Brown 1987). A cellulose synthesis assay was performed by incubating the membrane preparations in presence of 14C-labelled UDP- glucose and Mg2+. The insoluble radioactive product was separated by filtration and measured using liquid scintillation. The product was characterized by enzymatic hydrolysis using cellobiohydrolase and endoglucanase, methylation analysis followed by gas chromatography, high performance gel permeation chromatography and X-ray diffraction. The degree of polymerization of the resultant product was 5270 and the crystalline nature determined by X-ray diffraction was found to be that of cellulose II

(Bureau and Brown 1987). The Km of this reaction which followed Michaelis-Menten kinetics, was 2.0 x 10-4 mM and the Vmax was found to be 52.4 nmol glucose incorporation per mg of protein per minute. Similar values were obtained for digitonin- solubilized whole membrane fractions, by Aloni et. al. (Aloni, Delmer et al. 1982). The cellulose synthase activity observed predominantly in the CM had an optimum temperature of 30ºC and pH of 8.3. The enzyme activity in the membrane-bound and digitonin-solubilized form was found to be Mg-dependent, and inhibited by uridine mono- (Ki = 0.7mM), di- and triphosphates (Ki = 0.14mM) and guanyl nucleotides: pGpG and GpG (Ross, Mayer et al. 1990). Following the lead of cell-free cellulose synthesis in bacteria, in vitro cellulose synthesis was achieved in cotton fibers (Okuda, Li et al. 1993),

19

(Kudlicka, Brown et al. 1995), aspen cell suspension cultures (Colombani, Djerbi et al.

2004) , mung bean (Kudlicka and Brown 1997) and blackberry (Lai-Kee-Him, Chanzy et al. 2002).

Identification of c-di-GMP as the regulator (Ross, Weinhouse et al. 1987) and demonstration of in vitro activity (Bureau and Brown 1987) facilitated purification of proteins (Lin, Brown et al. 1990; Mayer, Ross et al. 1991) and determination of the cellulose synthase genes (Wong, Fear et al. 1990). Cellulose synthase was purified using the product entrapment method (Lin, Brown et al. 1990), that was successfully employed to isolate chitin synthase by Kang et. al. (Kang, Elango et al. 1984). This technique involves incubation of detergent-solubilized membranes in a reaction mixture as described above, and subsequent centrifugation to obtain the cellulose synthase entrapped within an insoluble cellulosic pellet. When the reaction mixture lacks either UDP-glucose or c-di-GMP, no synthase activity is retrieved in the pellet, proving the identity of the enzyme recovered from the pellet to be a cellulose synthase. Moreover, treatment with cellulase released 50% of the enzyme activity in the soluble portion. This also reflects the high affinity of the enzyme for the product formed.

Identification of the genes involved in cellulose synthesis

Using the product entrapment method, up to 350-fold purification of the enzyme could be obtained which was further used for isolating a highly pure synthase protein.

The pure enzyme was found to be composed of an 83 and a 93kDa polypeptide (Lin

1989). These polypeptides were used variously for antibody generation for immunological and localization studies (Chen and Brown 1996) as well for development of radiolabelled probes to identify protein binding characteristics and molecular weight

20

determination (Lin, Brown et al. 1990). But most importantly, the peptide sequences derived (Mayer, Ross et al. 1991) were used to design oligonucleotide probes to clone and sequence the gene encoding for cellulose synthase (Saxena, Lin et al. 1990) and deduce the operon harboring it (Wong, Fear et al. 1990).

Figure 1.3 Structure of cellulose synthase operon in related strains of Acetobacter Cellulose synthase genes have been variously named as acsA (Acetobacter cellulose synthase, axCesA (for Acetobacter xylinum cellulose synthase) and bcsA (bacterial cellulose synthase). G. hansenii strains ATCC 23769 and ATCC 53582 possess a single open reading frame for the cellulose synthse gene (acsAB) (Kawano, Tajima et al. 2002). In the strain 1306-3, the gene contains two open reading frames (acsA and acsB) characterized by Wong et al., whereas in G. xylinus strain NBRC 3222, the cellulose synthase gene contains three open reading frames (Ogino, Azuma et al. 2011). In the strain ATCC 1306-3, in which the operon structure was first studied, the initiation codon (97 bp upstream of the cellulose synthase gene and the termination codon (26 bp downstream of acsD gene) for the operon, are indicated by the triangle and stem-loop structure respectively.

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THE BACTERIAL CELLULOSE SYNTHASE OPERON

Genes involved in the synthesis of bacterial polysaccharides are usually organized as an operon that encodes for proteins mediating the various steps in the synthetic process

(Vazquez, Moreno et al. 1999; Whitney, Hay et al. 2011). In case of cellulose synthesis, the enzyme cellulose synthase converts UDP-glucose to cellulose in a single step (Lin,

Brown et al. 1990). This enzyme is encoded as part of an operon, referred to as the

Acetobacter cellulose synthase (acs) operon (Wong, Fear et al. 1990). The operon structure of cellulose synthase was elucidated by Wong et. al. (Wong, Fear et al. 1990) by genetic complementation of cellulose non-producing mutants of the strain A. xylinum

1306-3. The operon was further characterized by Saxena et al. (Saxena, Kudlicka et al.

1994) in 1994 to elucidate the function of the protein encoded by each gene of the operon, using site-directed insertional mutagenesis of ATCC53582 strains (Saxena,

Kudlicka et al. 1994). As shown in Figure. 1.3, the Acetobacter cellulose synthase (Acs) operon consists of four genes that were found to be transcribed into a polycistronic mRNA, with the site of transcription initiation located 97 bp upstream of the acsA gene.

These genes encode for four proteins: AcsA (84.4kDa). AcsB (85.3kDa), AcsC (141kD) and AcsD (17.3kDa). Sequence comparisons with initiation codons of the acs, ald and alh genes revealed that a highly conserved GGACGNG sequence is located 2-6 bases 5' of the AcsA start site (Wong, Fear et al. 1990). Based on similar homologous regions in the sequences upstream of the three genes, it was inferred that the transcription initiation site is represented by the sequence CATCGCTG which is located between -11 bp and -4 bp upstream of acsA. The transcription termination site is a 26 bp region at the 3' end of the acsD gene containing an inverted repeat sequence that has the potential to form a stem-

22

loop structure which serves as the signal for transcription termination in bacteria. In the

strains ATCC53582 and ATCC 23769, the acsA and acsB genes are fused to form one

gene (Kawano, Tajima et al. 2002), while in other strains like NBRC 3288 (Ogino,

Azuma et al. 2011), the ORF is split into three genes (Figure 1.3).

AcsAB

Although acsA and acsB were initially considered as two separate genes, it was

found later that depending on the strains of Acetobacter used for study, acsA and acsB

were either found as two separate ORFs or a single gene referred to as acsAB (Figure

1.3) In the Acetobacter xylinum strains ATCC23769 (now changed to G. hansenii) and

ATCC53582, cellulose synthase is encoded by a single gene encoding a 168 kDa protein,

whereas in 1306-3, BPR 2001, JCM 7664, there are two genes encoding for the different

regions of the protein.

Comparing the operon of catalytic A. xylinum ATCC 53582 with that of A.

xylinum ATCC 1306-3 revealed that the AcsA and AcsB polypeptides share ~81%

similarity to the N-terminal and C-terminal of the AcsAB protein (Saxena, Kudlicka et al.

1994). Using hydrophobic cluster analysis (HCA), where secondary structure prediction

of a protein is combined with the alignment to homologous proteins, conserved residues

in cellulose synthase protein were identified (Saxena, Brown et al. 1995). Cellulose

synthase belongs to the glycosyl transferase family 2 (GT2) and contains a DXXD motif

and another single highly-conserved aspartate residue and followed by QXXRW motif

(Saxena, Brown et al. 1995; Saxena and Brown 1997). Collectively this signature motif

of GTs is referred to as the D,D,D, Q/RXXRW motif . Attempts to replace the aspartate

residues by site-directed mutagenesis, resulted in loss of catalytic activity, providing the

23

reason behind the asparatate being conserved across species . This motif is conserved not only in cellulose synthases of all cellulose-producing organisms but is also common to

GTs like chitin synthase, hyaluronan synthase and glycosyl ceramide synthase (Saxena,

Brown et al. 1995; Saxena, Henrissat et al. 1995). Close examination of the deduced amino acid sequence shows that this sequence is found in the AcsA or the N-terminal half of the AcsAB protein. Using photoaffinity labeling of the purified protein with (-32P)- azido-UDP-glucose, Lin et al. (Lin, Brown et al. 1990)identified this protein to be the 83 kDa subunit of cellulose synthase (Lin, Brown et al. 1990). Based on this and the identification of the DXXD motif to be crucial for binding UPD-glucose, the acsAB gene was considered to encode for the catalytic domain of the protein.

The C-terminus of the AcsAB protein or the AcsB protein is presumed to contain sites for c-di-GMP binding (Kimura, Chen et al. 2001), thus serving as the regulatory domain of the protein. Kimura et al (Kimura, Laosinchai et al. 1999) used the 93 kDa polypeptide obtained from product entrapment to generate antibodies and localize this protein in the membrane fraction of the A. xylinum cells. However, after the discovery of

PilZ domain as the cyclic di-GMP binding motif (Amikam and Galperin 2006; Ryjenkov,

Simm et al. 2006) the function of AcsB as the regulatory domain was disproved

(Amikam and Galperin 2006). This is because of the fact that based on alignment studies, the PilZ domain is found in the C-terminus of the AcsA protein and in the center of the

AcsAB protein (Consortium 2012). Thus, currently the exact role played by the AcsB protein is unknown.

The amino acid sequence of the AcsAB protein shows 11 transmembrane domains (Saxena, Kudlicka et al. 1994). Data from this prediction further confirms the 24

results of the in vitro cellulose synthesis studies described earlier (Bureau and Brown

1987) and proves that AcsAB forms a integral membrane protein. The cytoplasmic localization of the cellulose synthetic activity and thereby the AcsAB, was also demonstrated to be localized in the cytoplasmic membrane of A. xylinum cells using sucrose density gradient centrifugation for separation of membrane fractions and subsequent assay of the fractions with radiolabelled UDP-(14C)-glucose as substrate

(Bureau and Brown 1987).

AcsC

The acsC gene codes for a 138kDa polypeptide. GTG in lieu of ATG, is the start codon in acsC gene and this codon overlaps the termination codon of the acsAB gene.

Though the exact role played by the gene product of the acsC has not been experimentally proved, its sequence homology to bacterial membrane channels and porins, suggests that the protein is involved in extrusion of the cellulose chains. Sequence based-prediction tools also show that it contains an N-terminal signal sequence for OM localization. Further the sequence reveals that the protein contains seven tetratricopeptide repeat (TPR, COG4783) motifs, which constitute approximately 20% of the protein, and a conserved motif for post-translational modification and protein turnover (COG3118)

(Marchler-Bauer, Anderson et al. 2005). The presence of TPR domains in many proteins is critical for their role in membrane transport and occurs in multiple copies in many proteins involved in binding with other proteins or ligands (Das, Cohen et al. 1998;

Blatch and Lassle 1999). Hence, the presence of a TPR motif in AcsC may in fact, be crucial for its role as the probable OM pore for cellulose secretion. This is further exemplified in the significant homology of this protein to VirB10 from A. tumefaciens

25

(47% similar, 23% identical) and Tra2 region of E. coli protein Trb1 (49% similar, 29% identical), which are known to interact with other proteins and form pore structures for secretion of macromolecules (Saxena, Kudlicka et al. 1994). From mutagenesis studies, it was established that AcsC is required for in vivo cellulose synthesis but not for the in vitro cellulose production (Saxena, Kudlicka et al. 1994). It is evident that a protein whose function is to form a pore in the membrane of the cells for extrusion of the cellulose fibers, would not be necessary when the cellulose is produced under cell-free conditions.

AcsD

The acsD gene encodes for a 17.3kDa protein whose role in the process of cellulose synthesis is largely unknown. Saxena et. al. (Saxena, Kudlicka et al. 1994) characterized this protein using TnphoA-mediated site-directed insertions of kanamycin

Genblock in the acsD gene (AcsD::Km). Unlike the wild type cells, the kanamycin resistant cells produced vastly reduced amounts of cellulose, under static as well as agitated culture conditions(Saxena, Kudlicka et al. 1994). Though the cellulose pellicle produced under static growth conditions by the AcsD::Km cells was very thin compared to the thick cellulosic mat of the wild-type cells, it was composed of cellulose II allomorph. Similar to wild-type cells, the cells from the stationary culture of AcsD::Km showed a linear array of intra-membranous particles (Saxena, Kudlicka et al. 1994).

However, under agitated culture by AcsD-deficient cells was a mixture of both cellulose I and cellulose II allomorphs, as revealed by X-ray diffraction analysis high-magnification observation of the product (Saxena, Kudlicka et al. 1994). AcsD protein is also very unique because it is the only protein encoded by the operon whose crystal and solution

26

structure has been deduced (Hu 2008; Hu, Gao et al. 2010). The localization and structure of AcsD and its possible role in the crystallization of cellulose ribbons are discussed in detail in Chapter III and IV of this dissertation.

The acs operon is flanked by genes that modulate cellulose synthesis

Other then the proteins encoded by the acs operon, other proteins are shown to be involved in the process of cellulose synthesis (Koo, Song et al. 1998; Koo, Song et al.

1998; Kawano, Tajima et al. 2008)The acs operon is flanked 5’ and 3’ ends by genes encoding an endoglucanase (cmcax) and a β-glucosidase (bglxA) (Standal, Iversen et al.

1994). Surprising though it may seem, expression of these cellulases has been shown to augment the rate and quantity of cellulose production (Tonouchi, Thara et al. 1995; Koo,

Song et al. 1998).

The cmcax gene upstream of the acs operon encodes for an endoglucanase belonging to GT family 8. Cmcax is an abbreviation for carboxymethylcellulase from

Acetobacter xylinum. The protein has a molecular weight of 42 kDa and shows an N- terminal 21 amino acid signal sequence for secretion. Overexpression of this protein, as well as its addition to the culture medium has shown to enhance cellulose production after incubation for 3 days, but this effect was not observed if the enzyme was added after

7 days (Kawano, Tajima et al. 2002). Though Acetobacter culture reaches stationary phase after five days, with cellulose production peaking at the third day, the endoglucanase expression was found to be elevated after five days of culture (Kawano,

Tajima et al. 2008), and this seems to contradict its role as an enhancer of cellulose synthesis (Kawano, Tajima et al. 2002). The cellulose hydrolyzing activity of endoglucanase, but not its ability to bind cellulose, serves to enhance cellulose production

27

(Tonouchi, Thara et al. 1995). It was shown that addition of Cmcax to cultures causes dispersion of cellulose fibers as shown in TEM images (Tonouchi, Thara et al. 1995).

Haigler et. al. (Haigler 1982) proposed that since the rate-determining step in cellulose polymerization and crystallization is the assembly of the microfibers, disruption of this assembly by endoglucanase causes accelerated cellulose synthesis (Haigler 1982).

Presence of a protein in plants, homologous to endoglucanase (KORRIGAN), in close proximity to the cellulose biosynthetic proteins, (Nicol, His et al. 1998; Robert, Bichet et al. 2005)further emphasizes the significance of the role of cellulose hydrolyzing activity in the process of cellulose synthesis.

The bglxA gene downstream of the acs operon encodes for a glucosidase belonging to GT family-3 (Tajima, Nakajima et al. 2001). It has been suggested that the

-glucosidase in A. xylinum functions to condense glucose units in the media to form a gentiobiose that serves activate the endogluconase activity in the cultures, which in turn accentuates cellulose production (Kawano, Tajima et al. 2008). In general, glucosidases hydrolyze cellobiose and smaller cello-oligosaccharides to produce glucose units and also catalyze the reverse reaction of addition of residues to cellulose chains. However, many glucosidases also function as transglycosidases (Kono, Kawano et al. 1999). This implies that BglxA might serve to maintain steady levels intracellular glucose and cello- oligosaccharides as substrates for cellulose synthase.

Dgc and PdeA

The Acetobacter genome contains three homologous cdg (cyclic diguanylate) operons that are each composed of a pdeA gene upstream of the dgc gene (Tal, Wong et al. 1998). These encode for three orthologous isozymes of Dgcs and Pdes with 28

homologous GGDEF and EAL domain organizations (Tal, Wong et al. 1998). The N- termini of the Dgc and PdeA proteins also contain oxygen-sensitive domains (Tal, Wong et al. 1998; Ryan, Fouhy et al. 2006). In addition to this, the cdg1 operon contains oxygen-regulated transcription activator gene (cdg1a) (Ryan, Fouhy et al. 2006).

Presence of oxygen-sensitive motifs in the proteins which control the turnover of the activator of cellulose synthesis (c-di-GMP), suggests that oxygen tension plays a crucial role in regulation of this process (Schmidt, Ryjenkov et al. 2005; Tamayo, Tischler et al.

2005; Ryan, Fouhy et al. 2006). As expected, inactivation of the Dgc gene, leads to decreased cellulose production (Tal, Wong et al. 1998; Delmer 1999). Cyclic-di-GMP binds to the PilZ domain of this protein (Amikam and Galperin 2006), which contains the conserved RxxxR and D/NxSxxG amino acid motifs. Although the PilZ domain functions as the effector of c-di-GMP, the mechanism of c-di-GMP-dependent regulation is largely unknown. Structural studies with PilZ domains of Vibrio cholerae and Pseudomonas aeruginosa have revealed that it undergoes a drastic conformational change upon binding to c-di-GMP (Benach, Swaminathan et al. 2007; Cotter and Stibitz 2007; Ramelot, Yee et al. 2007). X-ray crystal structure of the dimeric PilZ bound c-di-GMP, directs us to the possibility that the molecular surface created upon binding is available for binding to other target proteins, thereby translating the inter-domain changes induced by ligand- binding to the downstream regulatory effects (Benach, Swaminathan et al. 2007).

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The cellulose biosynthetic pathway and mechanism of cellulose synthesis

The steps involved in the pathway of cellulose biosynthesis were deduced using tracer studies using 14C-glucose The major steps in the pathway are as follows:

1) Transport of glucose across the cell membrane.

2) Phosphorylation of glucose to glucose-6-phosphate by glucokinase.

3) Isomerization of glucose-6-phosphate to glucose-1-phosphate by phosphoglucomutase.

4) Conversion of glucose-1-phosphate to UDP-glucose by UDP-glucose phosphorylase.

5) Polymerization of glucose into cellulose chains by the action of cellulose synthase.

Cellulose synthesis is an irreversible, energy consuming reaction, wherein all the enzymes except cellulose synthase are shared by other pathways in the cell. Being the unique enzyme in the cellulose synthesis pathway that catalyzes the committed step, cellulose synthase activity is the primary candidate for strict regulatory control.

Although the pathways leading to cellulose synthesis have been efficiently identified, the mechanism of glucose addition to the growing cellulose chain is still shrouded in mystery. It is known that UDP-glucose binds to the globular region of cellulose synthase, which is presumed to be cytoplasmic based on inferences from prediction tools

(http://www.cbs.dtu.dk/services/TMHMM/). But some key questions remain unanswered.

Are there more than one binding sites for UDP-glucose in the protein? How does the inversion of one glucose residue with respect to the other, take place? Is there a primer involved? How does the allosteric regulator cyclic di-GMP affect binding and catalysis?

Is the polymer elongated from the reducing end or the non-reducing end?

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Several hypothesis have been formulated to explain the mechanism of glycosyl residue incorporation into the cellulose chain. Though, identification of the mechanism underlying the process of cellulose synthesis remains a major challenge in the field, it is clearly known that the enzyme, cellulose synthase utilizes UDP-glucose and not cellobiose as the natural substrate (Ross, Mayer et al. 1991). This aspect of cellulose biosynthesis, wherein the repeating unit of the polymer is not the same as the monomers that bind to the synthesizing enzyme, is the most intriguing of all questions in the area of cellulose biogenesis.

Several mechanisms have been proposed to determine the series of events that account for the formation of β-1,4 linkages, release of the product from the enzyme active site, and extrusion of the result in polymer. Saxena et al.(Saxena, Brown et al. 1995) proposed that at the enzyme active site, UDP-glucose molecules bind to two distinct pockets, such that they are rotated at an angle of 180C with respect to one another and this dimer is continuously fed to the growing cellulose chain from the reducing end.

Delmer (Delmer 1999) modified this model to one in which the catalytic site is large enough to allow the rotation of glucose units, thereby facilitating inversion of each glucose within the binding pocket, during polymerization (Delmer 1999). In an extension to this model, the catalytic subunits from two enzymes are proposed to be organized to form a dimerized active site participating in the addition of two glucose residues at a time

(Albersheim, Darvill et al. 1997).

Many workers have proposed the involvement of a lipid-intermediate (Carpita and

Vergara 1998). The lipid portion of this intermediate tethers to the membrane while glucose residues are added to form a chain of cellulose in the cytoplasm . Once a 31

threshold length of the chain is reached, it is flipped to transport the chains outside the cells (Cooper and Manley 1975). Evidence for the presence of a lipid-intermediate was provided by Han and Robyt (Han and Robyt 1998) based on pulse-chase experiment with

14C-labelled UDP-glucose, to reveal that the elongation of cellulose chain occurs from its reducing ends. According to their model, nucleophilic addition of glucose from a lipopyrophosphoryl-glucosyl intermediate, to the cellulose chain, which is linked by a α- linkage to a lipid pyrophosphate, results in a β-1,4 linkage. This model accounts for the elongation of the cellulose chains from the reducing ends as seen in other bacterial polysaccharides like O-antigen polysaccharide and cell wall peptidomurein (Han and

Robyt 1998; Han and Robyt 1998)

Glucan polymers like starch and glycogen have a glycoprotein precursor, so it is logical to assume that cellulose synthesis could also require a protein-linked glycan. This assumption is supported by a pulse-chase experiment done in Acetobacter whole cells, by

Swissa et. al. (Swissa, Aloni et al. 1980). In spite of several initial evidences into the nature of the cellulose synthase or the polymer itself, a glycosylated cellulose synthase has not been isolated and the type of glycosylation is not known. Similarly, the enzymes involved in the lipopyrophosphorylation of UDP-glucose have not been isolated from bacterial membranes. Thus, the details of the exact mechanism of processive addition of glucose units and formation of the cellulose chain are still obscure and subject to speculation.

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STATEMENT OF THE PROBLEM

The details of the mechanism of cellulose synthase-catalyzed reaction are largely unknown and in all probabilities common to both the plant and bacterial systems. Once the cellulose is synthesized, it serves diverse roles in the two kingdoms. In plants, cellulose is incorporated as part of the cell wall, whereas in bacteria, it is released outside the cells as an extracellular polysaccharide. Due the differences in the cell wall architecture and the site of deposition, another essential inquiry specific to the bacterial system is, the process of extrusion of the polymer. The lack of understating the mechanism of synthesis as well extrusion is due to the dearth of information regarding the structure of the protein components of the cellulose synthetic machinery. There is considerable evidence that the proteins encoded by the cellulose-synthesis operon are involved in the synthesis and extrusion of the polymer (Saxena, Kudlicka et al. 1994).

However, there is no experimental evidence to prove their association with one another to form the cellulose synthesis and extrusion complex. If there is indeed there is a set of proteins that serve to extrude the synthesized polymer outside the cells, then such a protein complex is yet to be identified. Biochemical and structural characterization of the proteins should therefore be the starting point of any further attempts to understand the whole system of cellulose synthesis.

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SUMMARY

My work uses the two-pronged approach of characterizing the known proteins encoded by the operon as well as sequencing the genome to identify other proteins that contribute towards cellulose synthesis. The major research contributions from my work on characterizing cellulose synthesis are:

 Sequencing the genome of G. hansenii ATCC 23769.

 Localization and solution structure determination of the protein involved in crystallization of bacterial cellulose.

 Identification of the in vivo processing of cellulose synthase protein leading to cleavage into two polypeptides.

 Developing a procedure for in-vitro bacterial cellulose synthase assay using zymogram method.

 Heterologous expression and solubilization of the non-membrane bound regions of the cellulose synthase protein.

Chapter II of this dissertation describes the whole genome sequencing of G. hansenii that exists as a draft genome in the public domain of the National Center for Biotechnology

Information (NCBI). The genome sequence is made possible because of a combination of

454-titanuium FLX sequencing and SOLid sequencing methods. This is the first fully sequenced genome of a cellulose producing Acetobacter species. Therefore, it provides a good reference database for mapping the genomes of other related bacterial species and strains of Acetobacteriaceae. The sequenced genome was significant for our needs due to its contribution as the reference database towards our proteomic and mass spectrometric

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analysis of cellulose synthetic proteins. A concise version of this chapter has been published in the following journal article:

Iyer PR, Geib SM, Catchmark J, Kao T-H, Tien M. Genome Sequence of a Cellulose-

Producing Bacterium, Gluconacetobacter hansenii ATCC 23769. (2010) Journal of

Bacteriology 192:16, 4256-4257.

In Chapter III, we investigated the localization of the AcsD protein. At the time of our study, exact role of this protein, its structure and its localization were not known and could not be inferred from its sequence. Being the smallest protein encoded by the operon, it was easy to clone and express the gene and purify the protein heterologously.

This facilitated antibody generation against this protein as well as its structural characterization as described in Chapter IV. The antibody against AcsD was used to locate the protein in the different cellular compartments. Cytoplasmic membrane (CM), outer membrane (OM), cytoplasm and periplasm were isolated from the cells and their purity ascertained using marker enzyme assays. When proteins from these subcellular fractions were subjected to a western blot using anti-AcsD antibody, it was found that the protein is localized in the periplasmic region of the cell. Though, a simple experiment, this work has put the missing piece in the puzzle of secretion machinery in its right place.

A modified form of Chapter III has been published in the journal article:

Iyer PR, Catchmark JM, Brown RM, Tien M. Biochemical localization of a protein involved in synthesis of Gluconacetobacter hansenii cellulose (2010). Cellulose 18 (3):

739-747.

In Chapter IV, we tried to further characterize the AcsD protein, in terms of its structure. Around this time, Hu et. al. (Hu 2008) studied and released the crystal structure

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of this protein. We therefore attempted to solve its solution structure. This work is an addition to the crystal structure because the proteins is studied under the conditions in which it exists in the cellular environment. We found using gel filtration, analytical ultracentrifugation and small angle X-ray scattering (SAXS) that the protein indeed exists as an octamer in solution, as shown in its crystal structure.

Chapter V of this dissertation describes an important finding with regards to the processing of the cellulose synthase (AcsAB) protein. Although, encoded by a single gene, this protein is processed into three parts, as revealed by Western blots using specific antibodies against different regions of the AcsAB protein. Our attempts at purifying the polypeptides and the exact location of the cleave have not seen much success. However, based on the molecular weight, we have proposed the sequence of the resultant products.

The sixth chapter of this thesis presents a consolidated view of attempts at isolation of the cellulose synthase complex from the Acetobacter cells. Though not purified to its most elemental components, we have a partially purified complex of proteins that associate with one another and can be purified using in-gel methods. Native polyacrylamide gels were used to obtain a core complex of proteins that retained their activity after detergent solubilization followed by electrophoresis and chromatography. Using blue native gels, we have isolated the complex, that contains all the proteins encoded by the cellulose synthase operon as well as other proteins relevant to the pathway.

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CHAPTER II

WHOLE GENOME SEQUENCING OF GLUCONACETOBACTER HANSENII ATCC 23769

INTRODUCTION

The synthesis of cellulose occurs by a polymerization reaction catalyzed by the enzyme cellulose synthase, which utilizes UDP-glucose as the substrate (Aloni 1983;

Ross, Mayer et al. 1991). Since glucose units are polymerized to form the cellulose chains, the chemical composition of the cellulose thus produced is the same in all cellulose-producing organisms (Haworth 1932). However, the morphological properties of the polymer are unique to each species (Brown, Willison et al. 1976; Herth 1983; Itoh and Brown 1984) and these differences in crystallinity and dimensions of the polymer are largely attributed to the great diversity in the pattern and organization of the complexes that serve as sites of synthesis and extrusion of cellulose (Brown, Willison et al. 1976;

Giddings, Brower et al. 1980; Herth 1983; Itoh and Brown 1984; Tsekos 1999). although it is known that these complexes harbor the cellulose synthase protein itself (Kudlicka and Brown 1997; Itoh and Kimura 2001), their other proteins components of these complexes are yet to be completely identified. It is speculated that in addition to the cellulose synthase, these complexes contain the proteins that contribute to the process of cellulose extrusion (Saxena, Kudlicka et al. 1994; Delmer 1999).

Our primary goal is to characterize the process of cellulose synthesis in G. hansenii and in doing so, we aim to determine the composition and organization and of

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the cellulose synthesis complex in this bacterium. In our attempts to isolate and characterize the components of the cellulose synthesis complex using mass spectrometry

(MS) and sequencing techniques, we were hindered by the absence of a completely sequenced genome. So far, advances in studying the biogenesis of bacterial cellulose have been restricted to the proteins that are encoded by the acs- operon and its neighboring genes (Saxena, Lin et al. 1990; Saxena and Brown 1995; Tajima, Nakajima et al. 2001). Therefore, it is not possible to know whether more proteins other those encoded by the Acetobacter cellulose synthesis operon (acs) participate and exert an effect on the process of cellulose synthesis. Public databases (NCBI, EXPASY) are replete with many versions of cellulose synthesis-related proteins that were discovered at different time points by different authors (Lin, Brown et al. 1990; Wong, Fear et al. 1990;

Mayer, Ross et al. 1991; Nakai, Moriya et al. 1998). This further complicates the identification of newer proteins using MS-based techniques. We therefore sequenced the genome of the cellulose producing species Gluconacetobacter hansenii ATCC 23769, to explore the genetic blue-print further and unravel more factors contributing to this process.

The sequenced genome was used for proteomic studies using mass spectrometry.

In this chapter, I will also be describing the results from the MudPIT (Multidimensional

Protein Identification Tool) analysis of the G. hansenii membrane compartments. This analysis provided a complete proteomic profile of the total membrane (TM), cytoplasmic membrane (CM) and the outer membrane (OM). We have specifically concentrated on the comparison of the proteins related to cellulose synthesis in all of these compartments.

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Figure 2.1 Steps involved in a genome sequencing project. Sequencing the genome of an organism involves both biological as well as computational aspects. The biological work consists of isolation of the genomic DNA and fragmenting it into several pieces of suitable sizes for subsequent amplification to generate several clonal copies of each fragment (Duan 2010) which are sequenced using pyrosequencing platforms . These sequences, recorded in a file, are ready for further computational analysis(Duan 2010). The first step is to align and arrange the random sequences in a proper orientation in order to reconstruct the original full-length genomic DNA. This is called assembling the genome. The assembled genome is mapped to that of a neighboring organism, if available. This step can be skipped if the genome is sequenced de novo (Jarvie and Harkins 2008; Duan 2010). The sequence is then converted to coherent information through gene finding (Delcher, Harmon et al. 1999) and annotation . The files generated from the assembly and annotation are submitted to the databank , where the project becomes accessible to public.

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STEPS IN GENOME SEQUENCING

Genome sequencing employs a combination of molecular biology, instrumentation and computational techniques, to deduce the sequence of the genomic

DNA of an organism (Koonin 2001; Lesk 2007). This process essentially comprises of sequencing short fragments of genomic DNA, assembling them as one large scaffold, determining the open reading frames (ORFs), annotating these genes and finally converting it into a data repository (Duan 2010). This entire process is therefore regarded as the journey of a DNA sample from a test-tube to database (Lesk 2007). The steps involved in the process of sequencing are shown in Figure 2.1. Though sequencing as a technique requires interdisciplinary skills (Lesk 2007), our endeavor in this direction was motivated by the desire to probe further into the biochemistry of cellulose synthesis.

General outline of sequencing protocol employed in the 454-sequencing platform

The 454-sequencing technology was developed by CuraGen (Andrew 2003) for high-throughput DNA sequencing at a low cost. Presently owned by Roche , the 454- sequencing technology employs large-scale pyrosequencing method to sequence large stretches of DNA at a rapid rate . The latest version of the GS-FLX Titanium platform is capable of sequencing 600 megabases of DNA within 10 hours . The general workflow consists of DNA fragmentation , library generation, emulsion PCR , sequencing and analysis of the sequence (Duan 2010), all which are described in the subsequent section.

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Preparation of single stranded DNA library

The DNA is sheared into random fragments by nebulization . Pieces that are between 500-800bp are then selected. After DNA repair, short adaptors are ligated to the

3’ and 5’ ends of the blunt-ended DNA fragments . These adaptors serve as universal primers for amplification of the DNA fragment as well as sequencing .

Amplification of the library by emulsion PCR

The fragmented DNA ligated to a common primer/adaptor are immobilized on

DNA-capture beads such that each bead carries one unique single stranded DNA fragment. The DNA fragments amplified by emulsion PCR (emPCR) by emulsifying the beads in a mixture containing all the reagents required for PCR . Amplification generates single-molecule replicates of each fragment called “polonies' on the surface of each bead

(Mitra, Shendure et al. 2003), that contains millions of clonal copies of a particular library . Each clonal library-bound bead is segregated from the other by depositing each bead in a well of a titanium-coated Pico Titer Plate (PTP) .

Sequencing by synthesis (Pyrosequencing)

The amplified clonal fragments are sequenced by the 454-platform using the

"sequencing by synthesis" method (Nyren 2007). Here, each base incorporated in the growing DNA strand, is detected from the pyrophosphate released at each step of incorporation. Hence, the name "pyrosequencing" (Ronaghi, Karamohamed et al. 1996;

Ronaghi, Uhlen et al. 1998). Each well of the PTP with beads containing the clonal library, is loaded with DNA polymerase, ATP sulphurylase and luciferase (Ronaghi,

Karamohamed et al. 1996; Ronaghi, Uhlen et al. 1998). In each cycle, four nucleoside triphosphates are added to the reaction sequentially, in a predetermined order. When the

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complementary nucleotide is linked to the primer, the reaction catalyzed by DNA polymerase releases a pyrophosphate (PPi). ATP sulphurylase stoichiometrically converts this released PPi into ATP in presence of adenosine 5’ phosphosulphate .

ATP sulphurylase 2- AMP-SO3H + PPi ATP + SO4

ATP quantitatively fuels the reaction catalyzed by luciferase that converts luciferin to light- producing oxyluciferin (Gould and Subramani 1988).

Luciferase Luciferin + ATP ADP + Oxyluciferin

The unincorporated nucleotides and ATP are continuously degraded by the enzyme apyrase and the system is ready for another nucleotide addition. As the complementary strand is extended, each new incorporated base is recorded by the charge coupled device camera (CCD) which records the bioluminescence. The intensity of the chemiluminescent signal generated is proportional to the number of bases incorporated .

The 454 data analysis software converts the signal intensity after incorporation of a base in each well, into the type and number of nucleotide. Signals from all the wells are recorded simultaneously to generate sequences from all of the libraries . These collections of short fragments (700-800bp) of sequences are called as "reads" . Read length is the number of contiguous sequences that can be determined in one sequencing attempt (Lesk 2007). The read length generated after every sequence round, determines the limits for the subsequenct assembly procedure (Pop and Salzberg 2008). Since the

DNA is initially fragmented to generate short stretches that can be efficiently sequenced

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by the pyrosequecing technology, the reads thus generated have to be re-organized like pieces from a jigsaw puzzle (Lesk 2007). The presence of several copies of the genes helps to paste the sequences wherever they are missing, but some regions of the genome are too GC rich to be sequenced efficiently (Szybalski 1993). Such regions in the sequence contribute to “gaps” (Lesk 2007). The process of sequencing to close these gaps is called "finishing" (Lesk 2007). These gaps are either resolved using primer-walking

(Strauss, Kobori et al. 1986; Kaiser, MacKellar et al. 1989) or more rounds of sequencing

(Jarvie 2006) and mapping (Lesk 2007) .

Paired-end library generation

A recommended method for closing the gaps is to complement the sequencing results obtained from the de novo sequencing with a paired-end library, and sequencing the fragments using the "sequencing by ligation" principle (Jarvie and Harkins 2008). A paired-end library is generated by shearing the DNA and selecting for either 3 kb, 8 kb or

20 kb fragments which will determine the distance between the paired-end tags (Jarvie and Harkins 2008). The clonal amplification steps using emPCR are similar to the ones previously described (2007). The process of paired-end library generation is explained in detail in Figure 2.2.

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Figure 2.2 Generation of DNA library by paired-end method. The genomic DNA is methylated to prevent EcoR1 mediated cleavage and subjected to fragmentation. The DNA fragments are ligated on the 3' and 5' ends to biotinylated-hairpin adaptors that contain non- methylated EcoRI sites and Mme1 site. Exonuclease digestion removes all the DNA species which do not contain hairpin adaptors. EcoRI-mediated digestion, removes the terminal hairpin structures of the adaptors and exposes the cohesive ends on either side of the fragments, which self-ligate. This enables circularization of the DNA fragment around the two remaining portion of the two adaptors (now called linkers). The 8kb circular DNA contains a 44-mer linker between the fragmented DNA. The DNA circles are linearized by nebulization into approximately 250bp fragments. Those fragments that contain the biotinylated linker (Bio) are selected for, by immobilizing them to a streptavidin bead (SA). These fragments contain a 44bp linker flanked by 100bp DNA on either side . The two 100 bp fragments were located 8kb apart in the original DNA sample. These fragments are again linked to longer paired-adaptors that provide priming regions for subsequent amplification as well as sequencing (Jarvie and Harkins 2008).

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SOLid sequencing

The sequence of the paired-end library is determined by the "Sequencing by ligation" method (2007; Pop and Salzberg 2008) . This procedure exploits the property of DNA ligase, which can repair single-strand cleavage by adding nucleotides complementary to the template strand and thereby extending one strand of DNA (Lehman 1974). This technology was commercialized by Applied Biosystems in 2007 and adapted in a platform called “Support

Oligonucleotide Ligation Detection” (SOLid) (Duan 2010).

A paired-end library, is sequenced both from the forward and the reverse directions, using the DNA ligase and it provides 35 bp reads from either ends of the DNA (Pop and

Salzberg 2008). In this method, the DNA library is immobilized on a bead using the universal paired-end primer and this serves as the template. A set of eight color-coded di-base (two nucleotide) probes, compete for ligating to the template strand and extend the sequencing primer.

Each di-base is represented by one of the four colors: blue, red, green or yellow . In the first round of sequencing all 16 possible the probes are added to the reaction. When the 3' end of the probe is complementary to the sequence immediately adjacent to the sequencing primer, annealing occurs. DNA ligase mediates the ligation and the unbound probes are washed away.

The un-extended reactions are prevented from further annealing cycles by selective de- phosphorylation. The probes are cleaved with silver nitrate to remove the fluorescent tag and the last three bases. The ligation reaction is repeated to obtain another 5-mer that contains two nucleotides at 3' end, that are complementary to the template. A round of 15 cycles generates a

75 bp sequence, after which the extended sequences are melted off the template and another primer is hybridized, such that it is one base closer to the bead. The same pool of probes is used again to determine the sequence. The process of resetting the primer and sequencing is repeated

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five times. This ensures that each nucleotide in the sequence is probed twice and therefore the chance of sequencing errors is minimized. For sequencing from the reverse side, a 3' hydroxylated primer is ligated to the adaptor of the template and 5' phosphorylated probes are annealed to the primer. Subsequent steps in sequencing are performed similar to that of the forward template .

The result is displayed as a color-space sequence of di-bases. Since the adaptor sequence is known, the 1st base pair of the probe, annealed to the primer can be inferred. From here the entire color-coded sequence can be converted to a nucleotide sequence. The di-base system serves as a built-in accuracy check for the sequencing. This method, unlike the pyrosequencing technique, is not hampered by homopolymer repeat regions .

ANALYSIS OF SEQUENCE DATA USING SOFTWARE TOOLS

Assembly

Sequence assembly involves aligning and merging the reads generated by the sequencer in order to reconstruct the original sequence. If we look at the entire genome as a book then, these reads are likes shreds of all the pages from that book. The process of assembly is analogous to taking these shreds of several copies of a book and trying to paste the pieces of paper back together to regenerate one copy of the original book.

Generation of contigs

The assembler developed by Roche, to merge the reads generated by the 454-

sequencing platform is called as GS (Genome Sequencer) assembler or Newbler (New

Assembler) . Newbler can assemble reads generated by a single or multiple platforms .

When a genome is sequenced by both 454 and paired-end methods, the assembler

identifies reads as either linker-positive (454-pyrosequencing) or linker-negative (paired-

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end, SOLid sequencing) (2010). As a first step, the linker-negative reads are assembled into a de novo shotgun assembly (Staden 1979). These are catalogued into contigs which are long and contiguous sequences of DNA obtained by aligning and merging shorter reads, by identifying the best sequence overlaps between them (Jarvie 2006; Jarvie and

Harkins 2008). The contigs can be filtered on the basis of minimum sequence length desired, thereby eliminating very short and redundant sequences. The contigs generated by merging overlapping regions of DNA, result in sequences that are aligned without any regard to their order or position (Jarvie 2006; 2010).

Scaffolds

Although shotgun sequencing (Staden 1979) affords superiority in terms of longer read lengths (2007), it is through the paired-end library reads that the contigs can be properly ordered and oriented into larger concatenated assemblies, called scaffolds

(Jarvie and Harkins 2008). This is done in the second round of assembly, where the assembler considers the linker-positive reads and identifies matching regions between the reads and the known contigs, using pair-wise alignment. When the end-tags of a paired- end read map uniquely to two different contigs, then these contigs are linked into a scaffold, if the known distance between the paired-ends (8kb) is reflected in the sequence contained in the two contigs (Jarvie and Harkins 2008) .

The assembly by Newbler thus produces two levels of sequence data

(Jarvie 2006). Scaffolds are larger stretches of sequences, where the contigs are oriented and ordered based on the uniquely mapped paired-end halves (2010). Based on the size of the insert generated when both the halves of a paired-end map to the same contig, the insert size is determined and this is used to derive the distance between adjacent contigs

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(2010). This information is essential to find out if the gap is real or an artifact of sequencing. The gaps between contigs are represented by a series of Ns in the sequence

(2010).

If the gaps in the assembly are unresolved even after multiple approaches and rounds of shotgun sequencing, then such gaps are filled by primer walking (Strauss,

Kobori et al. 1986), a procedure in which primers are designed to the nucleotide sequences flanking the gap, and the DNA is amplified by PCR (Strauss, Kobori et al.

1986). The amplified portion is sequenced and manually aligned to the gap region using an alignment software (Tamura, Dudley et al. 2007). If the gap is not completely closed, the process of primer-design for the newly generated ends and amplification is continued until the complete sequence length covering the gap, is obtained (Lesk 2007). Based on the final application of the sequenced genome, some persistent and difficult-to-sequence regions are left behind as gaps. For most biological applications, a genome that is 90% gapless, is considered to be sufficient for further inquiry (Lesk 2007).

Gene prediction and annotation

Once the genome has been assembled into consenses contigs or scaffolds, it is ready for gene predicitions and annotation (Lesk 2007; Aziz, Bartels et al. 2008; Pop and

Salzberg 2008). Annotation of a sequence catalogues the sequences and gives access to the information contained in it, which would otherwise be just a long linear list of nucleotide bases. The method of predicting a gene involves using a program like

GLIMMER (Gene Locator and Interpolated Markov Modeler) to scan the sequence and identify ORFs based on predictions from a variable Markov model algorithm (Delcher,

Harmon et al. 1999). All possible reading frames in both strands are scanned individually.

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Those candidate gene sets that score beyond the threshold set by the algorithm are predicted as coding sequences (Salzberg, Delcher et al. 1998; Delcher, Harmon et al.

1999).

MATERIALS AND METHODS

Isolation of genomic DNA

The genetic material of G. hansenii was isolated using the Genomic DNA purification kit A1120, Promega, Wisconsin). The procedure given in the instruction manual accompanying the kit (2010) was modified slightly to accommodate the culture duration and conditions for G. hansenii. Briefly, the bacterial cell culture was started by inoculating 5 ml of Schramm-Henstrin (SH) medium with cells from a frozen glycerol stock. After 1 day of growth, the culture was supplemented with 0.1% (w/v) cellulase

(from Aspergillus niger, Sigma-Aldrich). After 72 hours of growth, the cell pellet was obtained by centrifugation at 16,000 × g for 2 minutes and was re-suspended in 600 µl of

"nuclei lysis solution" and incubated at 80°C for 5 minutes. The solution was cooled to room temperature and 3µl of ribonuclease was added to the suspension and incubated at

37°C for 30 minutes. The sample was again brought to room temperature and 200 µl of

“protein precipitation solution” was mixed with it by vortexing at high-speed for 20 seconds. After incubation on ice for 5 minutes, the sample was centrifuged at 16,000 × g for 3 minutes. The supernatant containing the DNA was transferred to a microcentrifuge tube containing 600 µl of isopropanol. Upon gentle mixing, thread-like strands of DNA become visible. This DNA was recovered by centrifugation at 16,000 × g for 2 minutes.

The supernatant was decanted and 600 µl of 70% ethanol was added to the DNA pellet to wash it by gently inverting the tube several times. The residual ethanol was removed after

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centrifugation. The DNA pellet was allowed to air-dry for 15 minutes. DNA rehydration solution (100 µl) was added to the tube and incubated at 65°C for 1 hour. The rehydrated

DNA was stored at 4°C.

The concentration of the isolated DNA was determined by reading the absorbance at 260 nm, of 1µl of the sample using a Nanodrop spectrophotometer. The samples submitted for the de novo 454-sequencing and for paired-end sequencing contained 25 μg of DNA, and 22 μg of DNA respectively, as measured by Nanodrop spectrophotometer .

Lane 1 Lane 2

20 kb 10 kb

Figure 2.3 Agarose gel electrophoresis of genomic DNA. 5µl of the genomic DNA isolated from the G. hansenii cells was analyzed on a 1% agarose gel. The genomic DNA can be seen as a clear band (indicated by an arrow, Lane 1) that migrates above the 20kb band in the “Fermentas Gene-ruler 1kb (SM#0333)” molecular weight ladder (Lane 2). The large molecular weight of the isolated DNA and the absence of smaller bands indicated that the DNA isolation procedure yielded a high-quality, non-fragmented genomic DNA sample suitable for the purposes of sequencing.

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Processing and analysis of the sequenced data using software tools

The sequence data output from the 454-sequencer was provided in the form of a

.sff (Standard Flowgram Format) file, which contained light information on the intensity of signal for each read. The sequence contained in this file were assembled using the GS assembler, Newbler. Newbler was run in a 64-bit version in the Linux operating system, using a command-line interface. The command to access a file and assembles its reads is:

“runAssembly aceto.sff”. The output directory created was given a name automatically by the software which shows the time and date of the assembly performed. For instance, the assembly folder generated after combined assembly of the 454- and paired-end reads, was called P_2010_03_01_09_04_14_runAssembly”. This directory contained the following sub directories. Containing the following files after completion of assembly:

Newblermetrics.txt: Each Newbler metrics file provides information about the number of scaffolds, and the number of contigs.

Contigs.fna: Fasta file of all reads

Contigs.qual: quality scores of bases within a contig

Scaffolds,fna: fasta file of all contigs within a scaffold. Gaps represented by a series of

Ns

Scaffolds.qual: quality file of all contigs in a scaffold.

The 454 reads and paired-end reads were assembled individually, in addition to assembling the two reads together. The contigs and the scaffolds generated by these

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assemblies were carefully analyzed. The largest contigs and scaffold was identified using the Newbler metrics file. Since the largest scaffold was as large as the bacterial genome size, the gaps (Ns), in this scaffold were examined. The Newbler assembler, often adds a gap of 21 nucleotides and these were located and, being artifacts of the assembly process, were deleted. The remaining large and small gaps were closed by PCR.

Primer walking: Primers were designed for sequences flanking the gap regions. PCR was performed according to standard procedures using HF Phusion Taq polymerase

(Thermo Scientific) PCR reagents. The PCR product from each reaction was subjected to agarose gel electrophoresis and for verifying the size of the amplified product. Along with 5 µl specific primers, 5 µl of PCR products were sent to the "Nucleic acid

Sequencing Center", in University Park, PSU.

The sequences obtained were aligned with the contigs using MEGA 4.1 (Beta)

(Tamura, Dudley et al. 2007) and the gaps were closed by substituting the "Ns" with the sequenced region. Since some gaps were very large, several rounds of primer design,

PCR and sequencing were done to fill in the nucleotides that were missing. The gaps that were persistent and could not be closed in spite of several rounds of PCR were left in the sequence as a series of Ns. The fasta file of the largest scaffold, in which several gaps had been replaced with meaningful sequences, was now considered as a high quality draft assembly (Chain, Grafham et al. 2009) and this was used as the dataset for subsequent steps of gene prediction and annotation.

Annotation: The high quality draft assembly was annotated using multiple platforms.

Firstly, we used the Glimmer software tool for gene-predictions (Delcher, Harmon et al.

1999), in order to obtain a private genome repository for analyzing our in-house

52

proteomic data. The assembled genome was also uploaded and analyzed using the Rapid

Annotation Service Tool (RAST) (Aziz, Bartels et al. 2008), for obtaining a genome viewer to study the operon structure and features of the regions flanking the operon and to enable comparative analysis of the genome and the genes with those of the related species. However, in order to generate a public genome database, we submitted all the files in the format described by NCBI to the NCBI-owned, Prokaryotic Genomes

Automatic Annotation Pipeline (PGAAP) pipeline .

Sample preparation for MudPIT analysis

Membrane fractions were obtained as described in the Materials and Methods section of chapter III. A total of 1 mg of protein from each membrane compartment was subjected to in-solution digestion, using the methods provided by the "Proteomics Core facility", Hershey, PSU. Briefly, Protein concentrations of the samples from each membrane compartment were determined using the Bradford method (Bradford 1976).

The volume of sample from each compartment, containing 1 mg of protein, was subjected precipitation by mixing with 100% (w/v) trichloroacetic acid to the final concentration of 30% (v/v) and freezing the sample. After thawing, these samples were centrifuged at 10,000 x g for 25 minutes to obtain the proteins in the pellet. The protein pellet was washed in 80 % ice-sold acetone and allowed to dry in the fume-hood. The pellet was resuspended in 100 µl of 100 mM Tris-Cl buffer, pH 7.8, containing 6 M Urea.

To this suspension, 5 µl of reducing agent (30 mg dithiothreitol in 100 mM Tris- Cl, pH

7.8) was added to obtain a final concentration of 10 mM DTT. The reduction was allowed to proceed for 1 h. The reduced protein sample was alkylated by adding 20 µl of alkylating reagent (36 mg of iodoacetamide in 100 mM Tris-Cl buffer, pH 7.8), to obtain

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a final concentration of 10 mM iodoacetamide. The protein sample was alkylated for 1 h at room temperature. The sample was again reduced as before, to remove any unconsumed iodoacetamide. To finally digest the protein, 100 µl of Trypsin is added to the sample from a stock of 20 ng/ µl. The sample is allowed to incubate for 16 hours at

37°C. The digested sample was dried in a vacuum concentrator and reconstituted in 100

µl distilled water. The drying and reconstitution was repeated three times and the sample is sent for MudPIT analysis to the Proteomics Core facility, Hershey, PSU. This strategy of multidimensional chromatography involves, use of biphasic column made of polysulfoethyl sspartamide, which is packed at its distal end with a reverse-phased resin, such as C18 resin. The proximal end of the column is packed with a strong-cation exchange (SCX) resin. The digested peptide mixtures are introduced onto the SCX resin at the rate of 1ml/ min, and fractions are eluted with a stepped-salt gradient. The elutions from the SCX flow into the C18-column, from where the fractions are eluted into the mass spectrometer, by applying a gradient of acetonitrile. After regeneration of the reverse phase resin, another fraction is released from the SCX-resin using an increased salt gradient. This cycle is repeated until the SCX-resin is exhausted. The elutions from the C18-column are mixed with a flow of MALDI matrix solution and spotted onto a stainless steel MALDI target plate. MALDI target plates (15 per experiment) were analyzed in a data-dependent manner on an ABI 4800 MALDI TOF-TOF. After acquisition of MS/MS spectra for all the spots in all the plates, protein identification and quantitation were performed using the Paragon algorithm as implemented in Protein Pilot

3.0 software (version 2.01), from ABI/MDS-Sciex. All the identified peptides were enlisted in an excel file, only identifications with a ProteinPilot Unused Score of > 1.3

54

(>95% confidence interval) were accepted. The MudPIT analysis and subsequent MS/MS analysis was performed by Anne Stanley, Proteomics Core Facility, Hershey, PSU).

RESULTS

Assembly metrics

A combinatorial sequencing approach generated 489,201 reads containing

162,0766,26 bp from the shotgun library and 195,088 reads containing 592,174,90 bp from 8-kb paired-end library. Together these reads contained a total of 221,294,116 bp.

These reads were assembled using the Newbler assembler, producing 88 large contigs

(>500 bp) and a chromosome-sized scaffold of 3,646,142 bp with an average coverage of

50.5X. This scaffold contained exclusively chromosomal DNA and no plasmid sequences.

Finishing, annotation and databank entry

The gaps in the large scaffold were filled by primer-walking and subsequent sequencing of the PCR products. The resulting high-quality draft assembly, consisting of a large scaffold with 71 contigs, was annotated using the Prokaryotic Genomes

Automatic Annotation Pipeline (PGAAP) service of the National Institute of

Biotechnology Information (NCBI). The gene predictions in the pipeline were made using Genemark and Glimmer (Salzberg, Delcher et al. 1998; Delcher, Harmon et al.

1999). The genome can be accessed from NCBI. It has been given the accession number

PRJNA43711. The genome viewer can also be accessed from the RAST database (Aziz,

Bartels et al. 2008).

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Genome features

The chromosomal sequence of G. hansenii 23769 contains 3,547,122 bp, with a G

+ C content of 59%. The genome contains a total of 3,351 genes, of which 3,308 are protein-encoding genes, which account for 84% of the genome. All the genes contain the prefix "GXY". There are 43 genes for transfer RNAs and 2 ribosomal RNA loci.

Genome features were also analyzed using the RAST subsystem technology version 4.0 (Aziz, Bartels et al. 2008), which reconstructs metabolic networks and the resulting annotation data is viewable through the SEED-viewer. Based on the RAST analysis, the genome contains 14 genes encoding enzymes involved in fatty acid synthesis, 17 heme and siroheme biosynthesis related coding sequences and bacterial chemotaxis associated cheA, cheB and cheR genes (Aziz, Bartels et al. 2008). All major subsystems are depicted in the pie graph in Figure 2.4.

Features relevant to cellulose synthesis

The genome contains the genes encoding proteins involved in cellulose synthesis in an operon consisting of acsAB (GXY_04277), acsC (GXY_04282), and acsD

(GXY_04292), as previously shown by Saxena et. al. (Saxena, Kudlicka et al. 1994) and

Wong et. al. (Wong, Fear et al. 1990). Interestingly, there are two additional copies of acsAB, GXY_08864 and GXY_14452 which share 40% and 46% identity, respectively, with the acsAB in the operon. Three sets of the acsAB and AcsC protein were reported for the strain ATCC 7664 (Umeda, Hirano et al. 1998), but for the strain used in the present study (ATCC 23769), only two of these three sets of acsAB genes have been reported (Saxena and Brown 1995). There are two acsC copies, GXY_08869 and

GXY_014472, in the genome, which have 28% and 30% identity to the acsC gene in the

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operon. Each copy of acsAB is adjacent to a copy of acsC. The distance between acsAB,

GXY_08864 and acsC, GXY_08869 is 17bp. acsAB GXY_14452 and acsC GXY_14472 are separated by 3299 base pairs. However, acsD is only present in the operon and is not duplicated elsewhere in the genome. The genome also contains three copies of the diguanylate cyclase genes, as reported by Tal et. al. (Tal, Wong et al. 1998) at loci GXY_

01169, GXY_016414 and GXY_01393.

Proteomic analysis of the membrane compartments

In addition to identifying unique features of the genome, we have utilized this genome for proteomic studies of the G. hansenii membrane compartments. MudPIT is a specialized form of LC/MS analysis in which is suited for the analysis of membrane proteins. The MudPIT analysis of the TM, OM and CM compartment generated enormous amount of proteomic data. The

All the files containing the raw data for peptides as well as the proteins, along with important prameters like coverage, confidence and scores in the supplementary files are contained in the storage disk provided with the dissertation (MudPIT/ TM/

TM_protein.xls, MudPIT/ TM/ TM_peptide.xls, MudPIT/ TM/ TM_protein.xls, MudPIT/

CM/ CM_peptide.xls, MudPIT/ OM/ OM_protein.xls, MudPIT/ OM/ OM_peptide.xls).

The MudPIT results were analyzed only in the context of the proteins that are involved in the cellulose synthesis pathway. Table 2.1 lists the all the cellulose- biosynthetic proteins in the TM, CM and OM fractions of these cells. Among the Acs proteins, AcsC is seen in the OM compartment and only one unique peptide corresponding AcsD is seen in the CM and the OM compartment. TM contains four unique peptides of AcsD. Our results in chapter III identify the localization of AcsD In

57

the periplasmic space, so its presence in the TM but not in the CM and OM fractions confirms our results. However AcsAB, which was considered to be a CM-bound protein, was not detected in the CM. Peptides from both the N- and C-termini of this protein were detected in the TM, but were conspicuously absent in the CM. Surprisingly, the peptides from the C-termini alone were detected in the OM. This indicates that the C-terminus of the protein is localized in association with the OM compartment of the cell. This result is corroborated with further evidences in Chapter VI of this thesis. Other proteins relevant to cellulose synthesis that were detected in the CM compartment are phosphoglucomutase, the enzyme that converts glucose 6-phosphate to glucose 1- phosphate, and a novel cellulose biosynthesis protein of unknown function.

Proteins TM CM OM

AcsAB 15 0 13

AcsC 2 0 3

AcsD 4 1 1

Cellulose biosynthesis protein of 4 2 2 unknown function

Phosphoglucomutase 18 14 0

Table 2.1 Distribution of the proteins relevant to cellulose biosynthetic pathway across the membrane compartments The number of peptides detected for each protein, with a confidence interval of 95% or above, are shown for each compartment. For the AcsAB protein, all of the 13 peptides detected in the OM aligned to the C-terminal portion of the protein. Of the 15 peptides identified for the AcsAB in the TM compartment, two were from the C-terminal portion of the protein.

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tion tion

can be canbe classifiedbased various on criteria. The

G. hansenii

enesencoded by

ochemical ochemical demonstration of the protein function is required to confirm the

Theg

Figure 2.4 Subsystem 2.4 Figure catalogue of genes the pie diagram distributes the genes based on their roles in the cellular metabolism. This classification is based on that the assump all the encoded genes are expressed and pathways. the of metabolic existence a bi

59

DISCUSSION

Characterization of any biochemical process requires the complete knowledge of not just the proteins participating in the pathway but also the other factors that regulate, interact and contribute to the process indirectly. Biochemical techniques like electrophoresis, cross-linking and chromatography can be used to isolate the proteins that interact with the operon-encoded cellulose synthase and related proteins. However, in order to identify these isolated proteins, we need to use tools like mass spectrometry and protein sequencing. These techniques in turn need a database of sequences to serve as a reference. Thus, to identify all the components of the cellulose secretion machinery, it was important to have a database of a sequenced genome.

The relevance of a sequenced genome is that, it allows us to look at all the factors influencing the cellulose-producing lifestyle of Acetobacter, instead of myopically concentrating on only the proteins that are encoded by the acs operon. We have used the sequenced genome as a reference database and determined the proteomic profile of the membrane compartments of this bacterium by MudPIT (Multidimensional Protein

Identification Tool) analysis. Although, a large amount of proteomic data has been generated using MudPIT, we have solely searched for the cellulose-synthesis related proteins and found some insightful results. The assumption so far in the field of cellulose synthesis is that the AcsAB protein is an integral membrane protein (Bureau and Brown

1987), localized in the CM of the bacterial cells. This was based on the detection of cellulose biosynthetic activity in the CM of the A. xylinum cells. We found that all the peptides corresponding to cellulose synthase were identified in the OM fraction. These peptides, when aligned to the whole-length protein of 1550 amino acid residues, match to

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the N-terminal end of the protein. None of these peptides match to the catalytic domain of the protein. This finding is relevant because, we have presented evidence for the association of the N-terminal end of the cellulose synthase protein to the OM compartment. This result will be further discussed in Chapter V. AcsC protein is localized in the OM, as expected from its N-terminal signal sequence for translocation to the OM. The peptides corresponding to AcsC are absent in the analysis of CM fractions.

In addition to the Acs proteins, phosphoglucomutase, a cytoplasmic enzyme involved in the UDP-glucose synthesis is found associated with the TM and CM compartments. Our results direct us towards the distribution and organization of the proteins involved in the cellulose synthesis complex. The organization of this complex was further studied and is described in the subsequenct chapters.

The proteomic data obtained for the membrane compartments, can be analyzed further, with respect to other pathways in the bacterial cells. Other than proteomic studies this sequenced genome is also being used for identifying protein-protein interactions contributing to cellulose synthesis using Yeast-two-hybrid system (Fields and Song 1989;

Iyer, Burkle et al. 2005) and studies involving random mutagenesis that influence cellulose production (both are unpublished work, by Deng, Y. and Kao, T-H ). The results from MudPIT and other mass spectrometric analysis for identifying the proteins components of the cellulose synthase complex, are elaborated in the Chapter VI.

In the wake of the genomic era, an additional tool to understand the significance and evolution of cellulose biosynthetic ability, is through phylogenetic analysis. In a recent search for the phylogenetic origin of horizontally transferred genes in plants, 15 genes were found to be common between Bacteria and Plantae (Price, Chan et al. 2012).

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It was shown that the plants acquired the genes for thiamine-pyrophosphate-dependent pyruvate decarboxylase family protein, through a horizontal gene transfer event from species of Proteobacteria. Among these bacterial species were G. hansenii 23769 and the cellulose non-producing G. diazotrophicus, which in an ancient HGT event, has contributed to the acquisition of the thiamine pyrophosphate dependent pyruvate decarboxylase gene, by the plant genome. This protein is involved in an alcohol fermentation pathway (Price, Chan et al. 2012). The maximum likelihood phylogenetic tree included sequences from the G. hansenii genome as well of as those of G. diazotrophicus, and A. pasteurensis. Thus, in this study, the genome sequence has contributed towards attempts at understanding the evolution of the plant photosynthetic machinery.

CONCLUSIONS

We have sequenced the genome of G. hansenii 23769. this genome is can be accessed using the accession number PRJNA43711, in NCBI. Using this genome, we have obtained the proteomic profile of the membrane compartments of this bacterium.

We have identified that although peptides from the full-length AcsAB protein were detected in the MS-analysis of the TM, the C-terminus of the AcsAB protein is associated with only the OM compartment. The N-terminus of this protein was not identified as in either the CM or OM. AcsC is present in the OM as expected and AcsD tends to associate largely with the OM.

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CHAPTER III

LOCALIZATION OF THE ACSD PROTEIN IN THE PERPIPLASM OF

G. HANSENII CELLS

INTRODUCTION

The acs operon has been shown to encode for the proteins which participate in the process of cellulose synthesis (Wong, Fear et al. 1990). The roles of acs-operon encoded proteins have been determined by using site-directed mutagenesis (Saxena, Kudlicka et al. 1994) and inferred by sequence analysis (Saxena, Lin et al. 1990; Saxena, Brown et al.

2001) and limited biochemical characterization (Bureau and Brown 1987; Lin, Brown et al. 1990; Chen and Brown 1996). As mentioned in Chapter I of this dissertation, AcsAB harbors the active site for glucose polymerization and is localized in the cytoplasmic membrane (CM) (Bureau and Brown 1987; Lin, Brown et al. 1990). Even though the role of AcsC has not been proven experimentally, the sequence shows a 21 amino N-terminal signal for outer membrane (OM) localization (Gattiker, Michoud et al. 2003). The presence of several tetratricopeptide repeat domains (Gattiker, Michoud et al. 2003) and the homology of AcsC to several bacterial porins (Saxena, Kudlicka et al. 1994) suggests that it serves as the pore in the outer membrane (OM) of G. hansenii cells through which cellulose is secreted (Saxena, Kudlicka et al. 1994).

Using mutants developed by TnphoA-mediated site-directed insertions, both

AcsA and AcsC proteins were found to be required for cellulose synthesis in vivo

(Saxena, Kudlicka et al. 1994). Upon disruption of the AcsD gene by insertional mutagenesis, the mutant produced has a partial Cel- phenotype on agar plates and produces 40% less cellulose compared to the wild-type cells (Saxena, Kudlicka et al.

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1994). Also, the cellulose produced was composed of both the cellulose I and cellulose II allomorphs (Saxena, Kudlicka et al. 1994). It was therefore inferred that AcsD is required for maximal cellulose production and influences the crystalline nature of the cellulose produced (Saxena, Kudlicka et al. 1994). Since in vitro cellulose synthesis was unhampered by the absence of AcsD, it could be gathered that this protein was not an essential requirement for cellulose biogenesis but was involved in the structural assembly of the final product. This lead to the speculation that this protein could be either localized in extracellular compartment, tethered to the OM or in the periplasmic space (Endler,

Sanchez-Rodriguez et al. 2010). In both these scenarios, the protein is capable of acting on the cellulosic chains emerging from the CM-bound cellulose synthase. However, unlike AcsAB or AcsC, the sub-cellular localization of AcsD has neither been proved experimentally nor can be inferred from its sequence. Though the structure of AcsD has been recently elucidated by X-ray diffraction pattern (Hu 2008), absence of information about the localization of the protein makes it difficult to understand its interactions with the other proteins encoded by the operon, as well as the precise mechanism by which the protein contributes to the process of cellulose extrusion.

In this study, sub-cellular fractions of G. hansenii cells were probed using Western

Blot with antibodies developed against AcsD. We found that AcsD resides in the periplasmic region of the bacterial cells. If indeed the acs operon-encoded proteins form a complex that mediates the synthesis and secretion of the polymer, then knowledge of the organization of the protein components of this complex and their interaction, is crucial to understanding the mechanism of cellulose synthesis and extrusion. Therefore, elucidating the sub-cellular localization of AcsD is a step towards speculating on the model of a

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cellulose biosynthesis system.

MATERIALS AND METHODS

Materials

Ni–NTA resin was purchased from Qiagen (Cat#301210). p-nitrophenyl phosphate

(N1127) and L-malate were products of Sigma–Aldrich. PCR primers were ordered from

Integrated DNA technologies.

Bacterial strains and culture conditions

G. hansenii ATCC 23769 cells were cultured in Schramm-Hestrin medium (SH)

(Schramm and Hestrin 1954) at 30°C in a rotary shaker. All cellular incubations were performed in SH media in the presence of 0.1% (w/v) cellulase (Trichoderma reesei cellulase, Sigma–Aldrich).

AcsD cloning, expression and protein purification

Single colony PCR (Woodman 2008) was used for amplification of AcsD gene using primers (5'- ggatccacaatttttgagaaa -3' and 5'-ctcgagggtcgcggaact-3'). The primers encode for restriction sites BamH1 and Xho1 respectively. The PCR product was analyzed in a 1% agarose gel (Figure 3.1a) from which the band corresponding to the

471bp fragment was provided. The gene was ligated into pGEM-T vector and was transformed into JM109 cells. Transformation cultures were plated onto LB (Luria–

Bertani) plates supplemented with 50 µg/mL ampicillin (LB-Amp plate) and 0.5mM

IPTG and 80 lg/mL X-gal. White colonies from the plate were picked and used for plasmid isolation. The plasmid was purified using the Qiagen plasmid mini-prep procedure. Presence of the restriction sites for BamH1 and Xho1 enabled digestion of pGEM-T-ligated gene (Figure 3.1b) as well as the pET-21a vector, with which the gene

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was subsequently ligated. The ligated gene was transformed in competent BL-21 DE3 cells. Ligations and transformations were performed using the procedure given in the pGEM vector manual (pGEM-T and pGEM-T Easy vector systems, Technical manual

No.042, Promega). The transformants were identified by plating 100µl of the transformation culture on an LB-ampicillin plate. The presence of the AcsD gene insert in pET-21a was verified by sequencing the ligated vector using specific primers designed for amplification. a b 1 2 1 2 3

pGEM (cloning vector)

1kb acsD acsD gene 506 bp (gene 500 471 bp insert) bp

Figure 3.1 Agarose gel electrophoresis of PCR product and pGEM ligated AcsD a) Amplification of AcsD gene from A.xylinum cells: The gene was obtained from A. xylinum culture by colony PCR. Lane 1 shows the PCR product and lane 2 shows the molecular weight ladder. b) Cloning of the acsD gene: The amplified PCR product was cloned into pGEM-T vector and restriction digested by BamH1 and EcoR1 enzymes. b) The restriction digested gene and pGEM vector are observed as bands between 400-500 bp and 3 kb respectively in Lane 1. The digested product was ligated into pET-21a and transformed into BL-21 (DE3). Lane 2 and 3 contain the molecular weight standards.

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Protein expression and purification

The AcsD protein was heterologously expressed and purified using standard procedures. Briefly, a single colony was picked from the LB-Ampicillin plate, used for identification of transformants, and inoculated in 5 mL LB medium supplemented with

2.5µl of 100 mg/mL ampicillin. The culture was incubated at 37°C for 16 h, with vigorous shaking at 200 rpm. This overnight grown culture was used as the inoculum for

1 L of LB medium supplemented with 500µl of ampicillin, and was again incubated under similar condition as before. The absorbance of the culture was monitored at 30 min intervals until an absorbance at 600 nm reached 0.6. At this stage, IPTG was added to a final concentration of 1mM and the culture was allowed to continue for an additional 4h.

Cells were harvested by centrifugation at 1,500 x g for 30 min and the cell pellet was frozen overnight at -20°C. The frozen cell pellet was thawed, resuspended in lysis buffer composed of 50mM NaH2PO4 pH 8.0 and 300mM NaCl and sonicated for 5 min with 15 s pulses. The lysate was centrifuged at 2,300 x g for 30min to obtain the protein in the supernatant. Protein purification was performed as per the instructions in the Qiagen handbook for Ni-NTA columns (The QiaExpressionist Handbook). Protein concentration was quantified by the Bradford assay using bovine serum albumin as a standard

(Bradford 1976).

Antibody preparation

Purified AcsD (1 mg) was sent to Covance research products for preparation of polyclonal antibodies from rabbits. The antibodies were affinity purified by transferring pure AcsD onto a nitrocellulose membrane using a modification of the method described by Robinson et al. (Robinson, Anderton et al. 1988). Briefly, the pure protein is subjected

67

to SDS–PAGE and transferred onto a nitrocellulose membrane. The band corresponding to the AcsD protein is detected by staining with Ponceau S stain (0.25% Ponceau S, 40% methanol, 15% acetic acid). The band visible on the nitrocellulose membrane is cut out and destained in TBS buffer (10 mM Tris, pH 8.0, 150 mM sodium chloride, 0.05%

Tween-20). The strip is incubated for 1 h in 6 mL crude serum diluted with 4 mL of TBS buffer. The non-specifically bound proteins are removed by three washes in TBST (TBS buffer containing 0.5% Tween-20). The bound antibody is eluted by dipping the strips for

10 min in 3mL 0.2 M glycine. The nitrocellulose strip is washed three times in TBST. To adjust the pH of the strip to 7.5, 100 µl of 3M Tris-Cl pH 8.0, is added to the strip and the whole process starting from the 1 h incubation, is repeated 6 times to obtain affinity- purified antibodies.

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a b 1 2 3 1 2 3 4

17 kDa

Fig. 3.2 Overexpression and Purification of recombinant AcsD a) Lane 1 contains proteins from un-induced cell pellet. Induction with IPTG causes overexpression of the AcsD protein as shown in Lane 2. The molecular weight of the protein band is a little above 17 kDa as the heterologously expressed protein contains six histidines at its C-terminal. b) AcsD was purified by Ni-Sepharose chromatography. The Coomassie blue stained SDS–PAGE gels shows the flow through from loading the crude cell extract (lane 1). The column was washed (lane 2) and then eluted with imidazole (lane 4). Lane 3 shows the molecular weight markers. Purified AcsD was then used to obtain rabbit polyclonal antibodies.

Preparation of membrane fractions

The cell pellet was obtained by centrifugation of a 48-hour grown culture. The total membrane fraction (TM) containing the CM and OM (along with components entrapped in the periplasm) from G. hansenii cells were obtained using the procedure described by Myers and Myers (Myers and Myers 1992) as modified by Ruebush et al.

(Ruebush, Brantley et al. 2006). Briefly, cell pellet from the culture at mid-log phase was obtained by centrifugation at 1500 x g for 20 min. The cells were resuspended in 24 mL of TS buffer (25% sucrose in 10 mM Tris–Cl pH 8.0) per gram wet weight. The suspension was constantly stirred at room temperature for 15 min, after which sequential

69

additions of the following components were made at 15 min intervals: one-tenth volume of lysozyme (0.64 mg/ml lysozyme), one-tenth volume of EDTA to obtain a final concentration of 5mM (20 mg/ ml), a final concentration of 0.3% (w/v) Brij58, a final concentration of 12 mM MgCl2 from a 15 mM M stock and finally a few crystals of deoxyribonuclease. The resulting suspension was centrifuged at 1500 g for 30 min to remove cell debris and whole cells. The supernatant composed of membrane fractions was centrifuged for 2 h at 177,500 x g (Ruebush, Brantley et al. 2006) (50,000 rpm) in a

Beckman Ti-70 rotor to obtain the TM pellet. The pellet was resuspended in 10 ml of 10 mM Tris–Cl buffer pH 8.3 and the protein content of this pellet was measured by Lowry et al. (Lowry, Rosebrough et al. 1951).

To isolate the CM and OM, 6 mL of TM was loaded on a 25–55% sucrose step-gradient tube and spun at 82,500 x g (25,000 rpm) for 17 h in a Beckman SW 28 rotor. The OM was obtained as a thick band around the 55% sucrose concentration and the CM band was obtained at the density of 35%. The fractions were collected as one ml aliquots using a 1 mL pipette. The TM, CM and OM were stored as 10% glycerol stocks at -70°C.

Preparation of periplasmic and cytoplasmic fractions

The periplasmic and cytosolic fractions were isolated by modification of methods described by Thomas et al. (Thomas, Daniel et al. 2001) and Streeter and Le Rudulier

(Streeter and Le Rudulier 1990). Cells were pelleted by centrifugation of 50 mL of actively growing culture at 1,500 x g for 10 min at 4°C. The cell pellet was resuspended in 7.5 mL TES buffer (50 mM Tris pH 8.0, 20% sucrose and 0.1 mM EDTA) containing

0.8 mg/mL lysozyme and incubated at room temperature for 10 min. Cells were harvested by centrifugation as before and resuspended in 2 mL of 5 mM ice-cold

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magnesium chloride. After incubation for 10 min on ice, centrifugation at 8,000 × g yielded periplasm in the supernatant and spheroplast in the pellet. The spheroplasts were resuspended in 2 mL HEPES-saline buffer (50 mM HEPES, 150 mM KCl), pH 7.2 and sonicated for 15 min at 15% pulse to release the cytoplasm, which was obtained in the supernatant after centrifugation at 250,000 × g for 30 min.

The purity of the membrane fractions was assessed by assaying for associated marker enzyme levels for each cellular compartment. Succinate dehydrogenase activity was assayed according to the methods described by Anwar et al. (Anwar, Brown et al. 1983).

The reaction mixture contained 60 mM sodium phosphate (pH 7.2), 10 mM potassium cyanide, 10 µg phenazime methosulfate, 20 µg dichlorophenol-indolphenol (DCIP), 25 mM sodium succinate and cellular fraction containing 100 µg protein in a total volume of

1 mL. The decrease in absorbance at 600 nm was monitored for 10 min at 25°C and specific activity calculated using extinction coefficient of DCIP, e = 13 mM-1 cm-1 (Fox,

Borneman et al. 1990). Alkaline phosphatase activity was assayed by modification of the procedure described by Garen and Levinthal (Garen and Levinthal 1960). The increase in

410 nm absorbance corresponding to hydrolysis of p-nitrophenyl phosphate (PNP) to p- nitrophenol was monitored in a reaction mixture composed of 0.1 mL of the cellular fraction, 1 mM PNP and 1 M Tris–Cl buffer pH 8.0. The change in absorbance was recorded for 5 min and the specific activity was calculated using the extinction coefficient (e) of p-nitrophenol as 16.7 mM-1 cm-1 (Halford 1971). For malate dehydrogenase (de Maagd and Lugtenberg 1986), the decrease in 340 nm absorbance due to oxidation of NADH, was monitored in a 1.1 mL assay mixture composed of 50 mM N-

2-hydroxyethylpiperazine-N'-2'-ethanesulfonic acid (HEPES) pH 7.2, 0.3 mM NADH,

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100 µL of the subcellular fraction. The reaction was initiated by addition of 25 µL of 10 mM oxaloacetate. Specific activity of the enzyme was calculated using extinction coefficient for NADH, e = 6.2 mM-1 cm-1.

Detection of AcsD using Western blot

The protein content in all the fractions was determined by using the method described by Lowry et al. (Lowry, Rosebrough et al. 1951). A total of 10 µg of the protein was loaded into wells of a 15% polyacrylamide gel and blotted into a nitrocellulose membrane. Western blotting was carried out using 1:300 dilution of anti-

AcsD as the primary antibody and a 1:10,000 dilution of the anti-rabbit IgG conjugated with alkaline phosphatase as the secondary antibody. 5-Bromo-4-chloro-3- indolylphosphate/nitro blue tetrazolium (BCIP/NBT) substrate was used for visualization of antibody-bound protein bands.

RESULTS

Purification of AcsD

The entire length of the 471bp acsD gene was amplified by colony PCR. The resultant DNA was sequenced, ligated to pET-21a vector such that the 3' end of the gene encoded for 6 histidine residues. The vector containing the acsD gene was transformed into BL-21 (DE3) cells. Addition of IPTG enabled enhanced expression of AcsD with a histidine-tag at the C-terminus as shown in Figure 3.2a. Sonication and subsequent centrifugation of the cell pellet in lysis buffer, released the protein in the supernatant fraction. The protein is therefore not strongly membrane-associated and is a soluble protein. This soluble fraction was used for purification of AcsD using Ni-affinity column.

Purity (greater than 95%) was assessed by subjecting the protein to SDS–PAGE followed

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by Coomassie blue staining (Fig. 3.2b). In addition to having the correct molecular weight (17kDa), the purified protein was identified as AcsD by cross-reactivity with the anti-histidine tag antibody on Western blot.

Specificity of anti-AcsD antibody

The purified AcsD protein was used to obtain polyclonal antibodies from rabbit.

The specificity of the antibody is shown in Fig. 3.3. Only one cross-reactive band was observed in Western blot when the anti-AcsD antibody was used to probe the whole cell extracts of the G. hansenii (Figure 3.3). Since the anti-AcsD antibody is highly specific, it was suitable for localization studies of AcsD in G. hansenii cellular compartments.

1 2 3

Figure 3.3 Determining the specificity of anti-AcsDantibody. Total cell extracts were subjected to SDS–PAGE and proteins visualized by Coomassie blue (lane 2) Lane 1 is molecular weight marker. Lane 3 shows the band obtained after the whole cell proteins were transferred to nitrocellulose membrane and visualized by Western blot with the antibody.

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Subcellular fractionation

We isolated the membrane fraction (and subsequently separated it into CM and

OM), the periplasmic fraction and the cytosol. The relative purity of the periplasmic fraction, the TM fractions and the cytoplasm fractions was assessed by marker enzyme assays for each of the respective fractions (Table 1). The cytoplasmic marker enzyme, malate dehydrogenase, exhibited an activity ratio of 6.25 between the cytoplasm and the periplasm. An activity ratio of 34 was observed for the CM marker enzyme, succinate dehydrogenase for the CM to periplasm. Our results from the marker enzyme assays, shown in Table 3.1 indicate that for all four fractions (CM, OM, periplasm and cytoplasm), there is little contamination between the fractions (ratio of specific activities is at least 5.6). More importantly, our results show that the periplasmic sample is largely free of cytosolic and membrane components.

Detection of AcsD in the periplasmic fraction

Upon establishing the purity of each fraction, we then performed experiments to determine the localization of AcsD. First, the periplasm, cytoplasm and TM fractions were subjected to SDS–PAGE (Fig. 3.4). The protein profile shown in Fig. 3.4a shows that the each fraction has a unique protein band profile, indicating the lack of similarity between the fractions and of the purity of each subcellular fraction.

All the cellular fractions were subjected to SDS–PAGE, followed by electrotransfer to a nitrocellulose membrane, which was analyzed by Western blotting with anti-AcsD antibody (Fig. 3.4b). A single band at 17 kDa corresponding to the molecular weight of

AcsD, was observed in periplasm. An intense band was also detected in the TM fraction

(Figure 3.4b). Because TM fraction preparations will also contain proteins from the

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periplasm (entrapped by protein-protein interactions), our results remain consistent with

AcsD localized in the periplasm. This was further confirmed when the TM fraction was separated into its component parts of the OM and the CM using sucrose-density ultracentrifugation. Western blot analysis again revealed localization in the periplasm.

Neither the OM nor the CM fraction contained appreciable amount of AcsD when compared to the periplasmic fraction (Fig. 3.4).

The 17 kDa band corresponding to the one obtained in the Western blot, in the lane containing the periplasmic fraction was excised from the Coomassie-stained gel. This band was digested with trypsin and analyzed by LC–MS. (The procedure for trypsin- digestion is elaborated in the Chapter VI of this thesis, which mainly focuses on the MS- related work.) The results of the MS analysis (Table 3.2), clearly indicated that the 17kDa band is AcsD. The TM fraction also showed a similar size band which would not be surprising due to entrapment of the periplasmic fraction during membrane isolation procedure.

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Table 3.2 Protein identification by LC-MS of trypsin-digested 17kDa gel band.

Entry Description Mascot MW Peptides Coverage Score (%)

ACSD_ACEX Cellulose 353 17432 R.DVDAEDLNAVPR. 9.2 Y synthase operon Q (91) protein D R.WVTSQAGAFGDY 11.8 VVTR.D (149)

The individual scores for the peptides are indicated with brackets along with the peptide sequence. Individual scores >42 indicate identity or extensive homology (p<0.05).

MW CM OM Cytop. Peri. CM OM Cytop. Perip. TM 72 CM OM Cytop. Perip. TM TM

52

42

34

26

17

Figure 3.4 Subcellular fractionation and detection of AcsD. a) SDS-PAGE profile of proteins from the subcellular fractions of G. hansenii. Each lane contained 10 µg of protein from the specified cellular compartment and subjected to SDS-PAGE. The band in periplasmic fraction, that was excised and sent for MS-analysis is indicated by an arrow. Cytop: Cytoplasm. Perip: Periplasm. b) Western blot of the cellular compartments using anti-AcsD antibody. Using anti-AcsD antibody, the proteins separated by SDS-PAGE were transferred onto a nitrocellulose membrane and Western blotted using anti-AcsD antibody. Bands corresponding to the AcsD protein can be seen only in the periplasmic and TM fractions.

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MTIFEKKPDFTLFLQTLSWEIDDQVGIEVRNELLREVGRGMGTRIMPPPC

QTVDKLQIELNALLALIGWGTVTLELLSEDQSLRIVHENLPQVGSAGEPS

GTWLAPVLEGLYGRWVTSQAGAFGDYVVTRDVDAEDLNAVPRQTIIM

YMRVRSSAT

Figure 3.5 The amino acid sequence of AcsD The N-terminal domain shows twin lysin residues (bold font) that indicate that the protein is possible candidate for secretion into the extracellular space using Sec-dependent transport.

DISCUSSION

Cellulose is synthesized by G. hansenii in the form of an extracellular ribbon of crystalline microfibrils. It has been shown that the enzyme machinery forms a linear complexes arranged along the longitudinal axis of the cells (Brown and Montezinos 1976). The biochemistry behind the process of cellulose synthesis and secretion is yet to be completely understood. However, it has been shown that the polymerization of glucose occurs in a single- enzymatic step from UDP-glucose to cellulose, catalyzed by cellulose synthase (Swissa, Aloni et al. 1980). This enzyme is encoded by the first gene of the acs operon, called acsAB (Wong, Fear et al. 1990; Saxena, Kudlicka et al. 1994). The other proteins (AcsC and AcsD) encoded by the operon are presumed to be instrumental in the assembly and export of the polymer across the cell membranes (Saxena, Kudlicka et al. 1994). Since, both AcsAB and AcsC are membrane- associated (Saxena, Kudlicka et al. 1994), to determine the organization of the cellulose synthesis and extrusion system, it was essential to know if the AcsD protein is a membrane protein or a cytoplasmic protein.

Determination of the role of AcsD in cellulose synthesis was first addressed by

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mutagenesis studies by Saxena et. al. (Saxena, Kudlicka et al. 1994). These workers showed that cells that lacked a functional AcsD, could still produce cellulose. However, these mutants exhibited a lowered rate of cellulose synthesis in vivo. But the in vitro rate (with isolated membrane fractions) is not altered in the absence of this protein. The cellulose that is made by the mutants lacking AcsD, however has altered crystallinity, being a mixture of both cellulose I and cellulose II allomorphs (Saxena, Kudlicka et al. 1994). This forces us to conclude that the protein comes into contact with the nascent cellulose chains and influences their assembly. This also indicates that the protein acts downstream of cellulose synthase in the process of cellulose extrusion.

Prior to our work, no other localization studies have been done for AcsD. Using conventional Western blot analysis and subcellular fractionation, we have shown for the first time that AcsD is localized in the periplasm. AcsD sequence lacks the known signals such as the twin arginine motif (RR) required for transportation of proteins to the periplasm (Yahr and

Wickner 2001; Palmer 2007). Examination of AcsD sequence (Figure 3.5) shows that the N- terminal of the protein contains twin-lysine residues, which is reminiscent of several proteins transported by the Sec-dependent pathway (Eitan 2007). However, we cannot conclude that the protein is transported by this pathway, based on the presence of the lysine residues alone.

Interestingly, commonly-used methods for computational prediction of periplasmic proteins like pSORTb (Gardy, Laird et al. 2005) provide no clue about the cellular localization of

AcsD. However, there has been evidence of other proteins devoid of such signals peptides like the Brucella abortus catalase (Sha, Stabel et al. 1994) which have been localized in the periplasm. Like AcsD, the periplasmic localization of this protein also cannot be deciphered using pSORTb. Based on the organization of proteins in other non-protein efflux systems

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such as the AcrA/AcrB/TolC system (Gerken and Misra 2004), we presume that AcsD protein serves as a periplasmic protein channel for transport of the newly-synthesized cellulose chain.

Our view is corroborated by evidence of weak interactions between of AcsD with cello- oligomers (Hu, Gao et al. 2010). A schematic consistent with this arrangement is shown in

Chapter IV.

It has been proposed previously that cellulose synthesis is a cell-directed process and several levels of regulations operate between the polymerization of glucose residues and emergence of twisted ribbons of cellulose (Benziman, Haigler et al. 1980; Haigler 1982).

Localization of AcsD in the periplasm together with its role in defining the crystalline character of the emerging cellulose fibers, leads us to the conclusion that the most probable role of AcsD to provide a channel through the periplasm for the nascent cellulose strand. For it to function as a channel necessitates that some part of AcsD be associated with AcsAB and/ or AcsC. Further studies on the interactions between the proteins would clearly accentuate the evidences presented herein for periplasmic localization of AcsD.

CONCLUSION

We have successfully cloned and heterologously expressed the AcsD protein. The specific antibody raised against this protein was used to determine its sub-cellular localization in the G. hansenii cells. The AcsD protein is localized in the periplasm. This work is the first attempts towards understanding the organization of the proteins of the cellulose synthesis complex, across the bacterial membrane.

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CHAPTER IV

DETERMINATION OF THE SOLUTION-STRUCTURE OF ASCD

INTRODUCTION

The previous chapter of this thesis describes our attempts at determining the sub- cellular localization of AcsD. Using specific antibodies against AcsD, we have found that the protein is localized in the periplasm of the Gram negative bacterium G. hansenii. This chapter describes our studies with the pure AcsD protein in the determination its structure.

Hu et al.(Hu 2008; Hu, Gao et al. 2010) elucidated the crystal structure of this protein at the time that our study was initiated. These workers found that the protein crystallizes as an octamer (Hu 2008; Hu, Gao et al. 2010). They also succeeded in obtaining the solution-structure of this protein, while our attempts in this area were underway (Hu, Gao et al. 2010). The AcsD protein forms an octameric complex. The octameric assembly is composed of a tetramer of dimers and forms a cylindrical structure with a 4-fold axis of symmetry and a central pore. Each AcsD monomer is oriented in the complex, in such a way that the N-termini of all the monomers are directed towards the center, and the C-termini radiate outside the cylinder. Four monomers in the upper layer of this octameric complex interact with four monomers in lower layer with the two layers shifted at an angle of 50ºC with respected one another . This creates four dimer-dimer interfaces that create four spiral interstices in the wall of the cylinder. Each of these passageways serve as a site of interaction for a glucan chain (Hu 2008; Hu, Gao et al.

2010). 80

The octamer is cylindrical in conformation and is composed of two stacked tetrameric complexes. The cylinder has a height of 62 Å, an outer diameter of 90 Å and an inner diameter of 65 Å. The top layer consisting of four monomers, is twisted at angle of 50ºC with respect to the bottom layer. The N-termini of each monomer in the octameric AcsD cylinder, form four inner passageways Each of these passageways is a site for association of a cellopentaose chain. There are two equally-probable and opposite orientations in which each cellopentaose can align with respect the dimeric interface.

(Hu, Gao et al. 2010).

This preliminary crystallographic analysis paved the way for our work on the solution-structure determination of this protein. Though, it can be argued that the crystal structure provides enough material to study the protein, our attempt at solution-structure determination was directed towards studying the oligomerization behavior of this protein under conditions close to the cellular environment. In doing so, we wanted to enquire if the oligomerization was indeed a preferred state of this protein, or was it a consequence of the best orientation assumed by the protein in its unit cell, during the crystal lattice formation. We determined the molecular weight of the predominant species of AcsD protein that exists in solution by employing analytical ultracentrifugation (AUC), dynamic light scattering (DLS) and gel filtration. All these experiments, described herein, indicated that the AcsD protein assembles as an octamer in solution.

We studied the solution-structure of this protein using small angle X-ray scattering (SAXS) and found that the octamer associates in the form of a dimer of tetramers with a central pore. Putting together the periplasmic localization and the cylindrical structure of this protein, a model for cellulose secretion complex is described.

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Since solution structure determination of a protein using SAXS is not a routine experiment in biochemistry laboratories, the underlying principles and the methods followed for the experiment and analysis, will be briefly described in the following part of this section.

Principles behind structural analysis by SAXS

Biological SAXS experiments are performed by exposing a solution of macromolecules to a high-intensity of collimated X-ray photon beam (Koch, Vachette et al. 2003). The incident X-rays are scattered by the electrons of the macromolecule at different angles. The elastic scattering pattern of the radiation contains information about the distribution of electron densities within the macromolecule. Therefore, the scattering pattern of this radiation is registered in the form of circles of finite width and plotted in as a function of the scattering angle, Q (Putnam, Hammel et al. 2007).

Q =4/*sin(/2) Equation 4.1

This scheme of events is depicted in Figure 4.1

The ability of a macromolecule like protein, to scatter X-rays depends on the concentration of the protein in solution, which determines the electron density of the protein. The difference between the electron density of the proteins and that of the aqueous buffer it is contained in, is referred to, as the excess scattering length density or contrast. The average electron density of protein and water are ~0.44 e-/Å3 and ~0.33 e-

/Å3, respectively (Putnam, Hammel et al. 2007).

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Figure 4.1 Small angle X-ray scattering experiment: The sample in the exposure capillary is irradiated by X-rays. The incident X-rays are scattered by the electrons in the sample to an angle of 2. The detector converts the 2D scattering into a 1D scattering profile, by means of radial integration.*This figure is adapted from Putnam et al. (Putnam, Hammel et al. 2007).

This results in a very minimal contrast, which is only 10% of what can be

achieved if the protein were in vacuo (Putnam, Hammel et al. 2007). However, in vacuo

analysis of protein structure is both impractical as well as far removed from cellular

conditions. A measurable signal from the protein molecule against an almost equally

intense signal of the aqueous buffer, is obtained by subtracting the scattering due to

buffer from the total scattering data for the protein in the buffer. The data obtained from

the SAXS experiment is thus a 1D-scattering curve that represents the scattering of a

single particle averaged over all orientations. From this scattering profile, it is possible to

calculate the aggregation state and dimensions of the macromolecule.

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Data analysis: The important parameters that can be deduced from the scattering curve are: the radius of gyration (Rg) and the pair distribution function P(r). The Rg also known as the second moment of inertia, refers to the mass distribution of a macromolecule around its center of gravity. Analysis of the SAXS scattering curve at low intensities can be used for approximation of the Rg of a protein assuming that the scattering due to proteins is to be equal to that of a spherical particle. This was calculated by the French scientist Andre Guinier (Guinier 1955). The Guinier approximation is represented in the form of a plot of natural logarithm of the measured intensities against Q2. In case, of the proteins, Guinier approximation is derived in the region near the beam-stop where the Q x Rg < 1.3. The extrapolated Y-intercept of this plot gives the value of I(0), which is the intensity at zero scattering angle, called the forward scattering (Guinier 1955).

I(Q) = I (0) exp {(-1/3)Rg2Q2}3 Equation 4.2

The Fourier transform of the I(Q) scattering curve yields the pair distribution function P(r) function (Glatter 1977), which describes the paired-set of distances between all of the electrons in a protein. Since this function describes all the paired-distances of electrons in the macromolecule, small changes in the relative positions of these electrons leads to large changes in P(r) distribution, thereby detecting conformational changes in the protein. The Rg of a molecule can be calculated from the P(r) function. Unlike the Rg obtained from the Guinier plot, the one derived from P(r) function is not restricted to low angles of scattering, but takes into consideration all of the data in real space. Using

Debye equation (Debye 1915) the P(r) functions can be related to the distribution of a single macromolecule to the scattering intensity as a function of scattering angle, Q

(Glatter 1977).

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Dmax I(0) = 4 0 P(r) dr Equation 4.3

As a corollary to this, if the shape and oligomerizarion of the protein are known a priori, then the scattering profile of the molecule can me calculated and P(r) distribution can be computed from the atomic position in the model. The point of intersection of the

P(r) distribution curve with the X-axis is the Dmax, the maximum diameter of the particle. All the available and known parameters of the protein, such as the scattering profiles, oligomerization state, symmetry and stoichiometry, are fed into a program called

GASBOR (Svergun 2001b), which reconstructs the 3D shape of the protein in the form of a bead-model. Since there are multiple structures that can have the same scattering profile, the modeling program narrows down on the appropriate structure by discarding all values of the scattering that do not match the limits set by Dmax. Using the prior knowledge of symmetry of the molecule the GASBOR program creates a model of sphere of diameter equal to the Dmax and populates it with number of beads equal to the number of amino acids in the protein. The adaptation of these principles for determining the structure of AcsD will be discussed in the subsequent sections.

MATERIALS AND METHODS

AcsD overexpression and purification: The procedures for protein expression and purification have been described in Chapter III. The protein was further purified in a

High trap QHP column of resin volume 5ml (GE Healthcare), to obtain a highly pure sample for DLS and AUC experiments (Figure 4.2). The protein obtained after Ni-NTA purification (25 ml of 0.5 mg /ml) was dialyzed against 50 mM Tris Cl pH 8.4 (Buffer

A). The protein was loaded into the column in Buffer A and eluted with a linear gradient of 0 to 100% buffer B (50 mM Tris-Cl pH 8.4, 1M NaCl) at the rate of 1ml per min for 85

100 minutes. The AcsD protein elutes as a major peak (900 mAU) at approximately 90 ml with a smaller shoulder (200 mAU). The sample collected at the peak (1.3 mg/ ml) and shoulder were subjected to SDS-PAGE and staining (Figure 4.2 b).

Sample preparation for AUC: AcsD purified using Ni-NTA affinity column (Figure

3.2), was in the elution buffer, which contained 50 mM NaH2PO4, 300 mM NaCl and 25 mM imidazole. pH 8.2. This protein was dialyzed thoroughly in the same buffer without the imidazole. 1 ml of AcsD at a concentration of 2 mg/ml was submitted for sedimentation velocity (SV) ultracentrifugation. A 35 ml sample of the dialysate buffer

(50mM NaH2PO4 and 300mM NaCl, pH 8.2) was also supplied.

Analytical ultracentrifugation: Using Sednterp program, the following physical constants for the protein were calculated MW = 17,375 Da, 20° =0.7408 ml/g. The buffer density and viscosity were calculated to be 1.0176 g/ml and 0.010576 poise at

20°C, respectively. Sedimentation velocity experiment was conducted at 20°C and

50,000 rpm using interference optics with a Beckman-Coulter XL-1 analytical ultracentrifuge. Double sector synthetic boundary cells equipped with sapphire windows were used to match the sample and reference menisci. The rotor was equilibrated under vacuum at 20°C and after an equilibration period of ~1 hour at 20°C the rotor was accelerated to 50,000rpm. The interference scans were acquired at 60 second intervals for approximately 7 hours. Sample dilutions for the analysis were prepared as shown in

Table 6.1

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a

1 2 3 4

Figure 4.2 Anion exchange chromatography of AcsD a) The major peak at 90 ml elution volume is followed immediately by a small shoulder. The peaks are eluted at 60% buffer B. b) The fractions collected at the peak of the elution and the shoulders were subjected to SDS-PAGE. Lanes 1contains the molecular weight ladder. Lane 2 is an empty lane. Lane 3 and 4 contain 20 μl of the peak fraction collected at the shoulder.

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Sample preparation for DLS: AcsD protein concentration was measured using the absorbance at 280 nm using extinction coefficient of 26470 M-1 cm-1. The concentrations used for DLS experiment were obtained by dilution from a stock of 8 mg/ml in 50 mM

Tris Cl pH 8.4 with 5% glycerol.

DLS experiment: All measurements were carried using the Viscotek-DLS equipment in

The X-ray crystallography facility, University Park. The Omnisize software linked to the instrument automatically generated a correlation curve for the sample based on its scattering profile. The size-range for calculating the hydrodynamic radius (Rh) was restricted between 1nm to 1000nm. Based on the Rh value the MW of the proteins is estimated by the software assuming the molecule is a perfect sphere. The data obtained from these measurements are used to arrive at the approximate, and not the exact molecular weight, of the protein.

Sample preparation for SAXS: AcsD protein (50 ml containing 6.5 mg of protein) was subjected to anion exchange, as described above. The protein fraction collected at the peak was concentrated a using a 10K concentrator (Amicon) to obtain a final concentration of 5.2 mg in a volume of 200 μl. Half the sample volume (100 μl ) was further purified by gel filtration. The other half was use for in-line gel filtration with

SAXS experiment. Protein concentrations were measured by reading the absorbance at

280 nm and using the extinction coefficient of 23760 M-1 cm-1.

Gel filtration: Gel filtration was carried out using a Superdex 200 column (24 ml, GE

Healthcare). The sample (100 μl of 26 mg AcsD) was loaded using a 100 μl loop. The buffer (50 mM Tris pH 8.4, 300mM NaCl and 5% glycerol) was passed through the column at the flow rate of 0.5 ml/min, for the duration of 48 minutes to obtain allow

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elution of the protein in 1 column volume of buffer. The 2ml volume protein eluted at the peak (1.2 AU), was collected and concentrated to 1 ml protein at 3 mg/ml.

SAXS experiment: The AcsD protein was purified using anion exchange and gel filtration chromatography was used for the SAX experiment. Three kinds of samples were used for the analysis, pure AcsD, pure AcsD incubated at room temperature for 24 hours and .

Data acquisition: All the SAXS experiments were performed at the Advanced Photon

Source (APS), Argonne National Laboratory (ANL) using the Biophysics Collaborative

Access Team (BioCAT) undulator beamline 18-ID. The scattering profile was obtained at very low angle of the beam (1.03 Å). These beams are directed towards an exposure capillary into which the sample is drawn. The measurements are carried out by 100 µl of

AcsD sample between the capillary and the sample tube at the rate of 50 µl/s. 10-20 exposures were taken for each sample. Each exposure lasted for 1 s. Measurements of empty capillary as well as capillary with buffer were taken to serve as blanks. The empty capillary was exposed for 20 shots and 10 shots each of the buffer and the protein sample were taken. The scattering profile due to the capillary, buffer and protein are shown in

Figure 4.6. The scattering due to buffer and the capillary has to be subtracted from the solution scattering profile. This is achieved by subtracting the scattering due to the capillary, from that of the protein and then subtracting the capillary scattering from the buffer, using the equation:

(Protein - Buffer)-Capillary - x (Buffer - Protein) x is a fraction which is approximately equal to 0.993 and is proportional to the amino acid composition, concentration of the protein and the partial specific volume. Since the

89

sequence of the protein and the concentration are known, we can calculate the partial specific volume (υ = ml/g), which gives us the fraction of the total volume occupied by the protein alone.

SAXS data analysis The scattering data were analyzed using IGOR software and the programs contained in the ATSAS package. Rg was calculated using the Guinier program using the IGOR BioCAT macros. The P(r) distribution was calculated using

GNOM software (Semenyuk 1991). The P(r) distribution was used by the GASBOR program (Svergun 2001b) for generating 50 bead models. With the a priori knowledge of the crystal structure, the model was built with an imposed symmetry of p42. The models were superimposed for comparison using Supcomb (Kozin 2001) and the average structure was arrived at using the DAMAVER (Volkov 2003) program. The average structure was further refined using the damfilt software and this structure was overlaid with the existing crystal structure of the AcsD protein.

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RESULTS

DLS experiment: This technique was valuable in monitoring the aggregation pattern of the protein over a range of concentrations. We observed that the protein showed a consistent molecular weight between 140 kDa and 153 kDa, across concentrations from

0.5 mg/ml to 8 mg/ml (Figure 4.3). The molecular weight of the major and minor species, for AcsD at different concentrations, are given in Figure 4.3. It can be seen from the graph and the accompanying data, that the mass distribution remains consistent across various concentrations of the protein, giving a value for molecular weight between 141-

153 kDa. This signifies the presence of a species equal to an octamer. DLS gave us our first indication to probe further into the oligomeric associations of AcsD.

This preliminary technique was valuable in monitoring the aggregation pattern of the protein over a range of concentrations. We observed that the protein showed a consistent molecular weight between 140 kDa to 153 kDa, across concentrations from 0.5 mg /ml to

8 mg/ml (Figure 4.3). It can be seen from the graph and the accompanying data, that the mass distribution remains consistent across various concentrations of the protein, giving a value for molecular weight between 141- 153 kDa. This signifies, presence of a species equal to an octamer. DLS gave us our first indication to probe further into the oligomeric associations of AcsD.

91

AcsD concentration: 0.5 mg/ml

Peak % Area Rh (nm) Position Std Dev % RSD MW (kD) 1 97.7 4.93 5.01 0.35 7.1 143.28 2 1.8 22.42 21.38 1.16 5.2 5137.64 3 0.5 180.71 169.82 17.64 9.8 7.11E+05

AcsD concentration: 1 mg/ml

Peak % Area Rh (nm) Position Std Dev % RSD MW (kD)

1 98.3 4.91 5.01 0.23 4.6 141.95

2 1.7 34.1 32.36 1.77 5.2 1.38E+04

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AcsD concentration: 8mg /ml

Peak % Area Rh (nm) Position Std Dev % RSD MW (kD) 1 97.6 5.12 5.01 0.42 8.3 156.62 2 1.8 52.59 48.98 5.82 11.1 3.85E+04 3 0.6 153.01 153.11 21.59 14.1 4.80E+05

Figure 4.3 DLS analysis of AcsD: The plot of mass distribution of the AcsD protein is displayed after selction for the range between 1 nm – 1000nm. The amplitude of the hydrodynamic radius of the protein sphere is indicated at the peak. Based on this Rh value, the molecular weight of the protein is estimated by the software (Omnisize). a. At the concentration of 0.5 mg/ml, the protein has shydrodynamic radius of 4.93 and a molecular weight of 143.28 kDa. b. At a concentration of 1 mg/ml, the protein has a Rh value of 4.91, corresponding to a molecular weight of 141.95 kDa. c. At a very concentration of 8 mg/ml the DLS analysis gives a Rh of 5.12 nm and molecular weight of 156.625 kDa.

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Gel filtration

The AcsD purified by Ni- NTA was further purified by gel filtration, for purposes of structural studies (SAXS) as well as to determine the aggregation behavior of the protein. The elution profile of the protein (Figure 4.4), shows a major peak which elutes at 13.2 ml. When analyzed by a semilog plot with standard protein elutions, this corresponds to the molecular weight of 136 kDa. This value again gives us an octameric complex of AcsD. We proceeded to determine the accurate molecular weight of this complex by sedimentation velocity experiment using the pure protein.

Figure 4.4 Gel filtration profile of AcsD. The plot of absorbance (in milli absorbance units) versus elution volume shows that the protein elutes at a volume of 13.2 ml. The inset shows a semi-log plot of elutions of protein with known molecular weights.

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Sedimentation velocity experiments

The data from the SV run were analyzed using three modeling programs, viz.

DcDt, Sedfit, and Sedphat. The DcDt + program provides a model independent sedimentation distribution, g(s*) analysis using a time derivative of the concentration profile. An overlay of the normalized distribution plots for the three highest concentrations of AcsD is shown in Figure 4.5. The protein concentrations were determined by integration of the g(s*) profile. The three concentrations used were 0.29 mg/ml, 0.71 mg/ml and 1.12 mg/ml. the lowest concentration was not used since the concentrations derived from integrating the g (s*) was only 70% of the estimated values.

The three curves overlay with one another. This indicates that the over the range of concentrations covered by the samples, there are no reversible reactions. If there are larger aggregates formed, there would have been a noticeable shift in the sedimentation coefficient towards a higher value upon increasing the concentration of the sample.

However, we observe that in the curve for the highest concentration of the protein (1.21 mg/ml), although there is a slight increase in the amount of material sedimenting at a higher S value, the lack of a prominent shift in the peak, indicates that this is a reversible aggregation. The protein predominantly exists as a single species in solution.

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Figure 4.5 Sedimentation coefficient distributions calculated from a sedimentation velocity experiment a) An overlay showing the normalized distribution plots for AcsD The sedimentation velocities have been corrected to standard conditions of water as solvent and temperature 20C. Protein concentrations shown in the figure were derived by integration of g (s*) profile. The plots superimpose, suggesting matching behavior of the sample over a range of concentrations. There is no indication of self association as there is not a considerable shift towards a higher S value. The arrow shows the slight increase in the sedimentation coefficient in the highest concentration of AcsD used (1.21 mg/ml), but this is not a prominent shift, obviating the possibility of formation of irreversible aggregates.

Analysis 2: Sedfit program

Sedfit, version 11.71 was used as the direct boundary modeling program for individual concentrations of AcsD, to generate continuous sedimentation coefficient distribution c(s) plots. The c(s) analysis was done at a resolution of 0.05S, using maximum entropy regularization with 95% confidence limit. As shown in Figure 4.5b, the c(s) distribution plots are sharpened relative to other analysis methods, because the broadening effects of 96

diffusion are removed by use of an average value for the frictional coefficient that is obtained as a fitted parameter. The plot is consistent with the g(s*) data from DcDt+, in that the prominent peak is at 6.75 S.

The plot of c(s)data at a scale of 10X that of the full scale plot (Figure 4.5c), shows that there is a small amount of material (1.5-3%) sedimenting slower than the peak and approximately 15% of the material sedimenting faster than the main peak. This is in agreement again with the g(s*) results as the change in the amount of a higher order aggregate is too small to suggest that it is a irreversible aggregate formed as a consequence of higher concentration.

Using model-based numerical solutions to the Lamm equation, multiple sets of the sedimentation velocity data were analyzed by the boundary modeling program for global analysis, Sedphat version 6.50. Data analysis by Sedphat program was done with a model of a hybrid local continuous distribution and a single global discrete species. This model was used because, in the sedimentation behavior of AcsD, there are two species in solution. There is a main non-interacting species that we are interested in characterizing, along with smaller aggregates that should be accounted for, in order to not bias the analysis of the species of interest. The values for the globally fitted parameters are a sedimentation coefficient of 6.75 S, and a molecular weight of 145 kDa with a of 95% confidence range of 141 to 149. The best fit value for the molecular weight is in close agreement with the expected value of 139 kDa for an octamer of AcsD. This confirms that AcsD exists as an octamer in solution.

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b c

Figure 4.5 b) Continuous sedimentation coefficient distribution c(s). Analysis of sedimentation plot of three continuous sedimentation coefficient distribution c(s), normalized to the concentrations derived from integration of the c(s) plot. The three concentrations used are 0.275 mg/ml (green curve), 0.699 mg/ml (blue curve) and 1.116 mg/ml (black curve). The values for sedimentation coefficients have all been corrected to standard conditions, S(20,w). A small amount of material sediments at approximately 9 S (indicated by arrow), but the major peak is at roughly 6.5 S. c) 10X magnification of the normalized c(s) plot This magnification allows discerning the minor peaks. We observe that 1.5-2.5% of material (shown in encircled peak), sediments slower than the main peak and 14 - 20% of the material (peak indicated by arrow) sediments at a faster rate. The main peak is at approximately 6.5S

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Determination of the AcsD structure using SAXS

Since the existence of a stable octameric complex of AcsD had been demonstrated by more than one method, we further explored the structure of AcsD in solution using small angle X-ray scattering. As a first step in this experiment, the protein scattering profile was obtained along with the scattering profile of the buffer it was contained in, and the capillary that held the sample during exposure. An example of scattering curves obtained for the empty capillary, buffer and the AcsD protein is shown in Figure 4.6a. a

Figure 4.6a: Experimental scattering profiles: The scattering curve is plotted with intensity in a logarithmic scale as the X-axis as a function of momentum transfer (Q). The units of Q are the inverse of wavelength units. So it is measured as Å-1 The scattering curve of the empty capillary (green) and buffer (blue) drop more rapidly than that of the protein (red), due to the greater Rg value of the latter.

99

b

Figure 4.6b The buffer subtracted scattering curves for AcsD: The background corrected scattering curves of a fresh sample of AcsD (red) and a 24-hour old sample of AcsD (blue).

The background-subtracted scattering curves for a fresh sample of AcsD in buffer containing glycerol land TCEP and a 24-hour sample AcsD is shown in Figure 4.6b.

Irrespective of the incident wavelength the scattering due to a molecule remains consistent except at very high and very low values of Q, where anomalous scattering occurs (Stuhrmann 1981). This is observed in the plot of I (Q) vs Q where, at very low scattering angles there is aggregation of the fresh AcsD sample. The linear portion of this curve is used for Guinier analysis (Guinier 1955) and the Guinier plot is displayed in

Figure 4.6c. As can be noted in the plot, the radius of gyration, Rg value derived from this analysis, using the equation 4.1, is 43.025.

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c

Figure 4.6c. A represention of the linear region of the fresh AcsD sample used for deriving the Guinier distribution.

In addition to obtaining the Rg value using the linear portion of the scattering

curve, the P(r) function was obtained by Fourier transformation of the scattering data

using the GNOM program. The representative pair-distribution plot shown in Figure 4.7,

gives the Dmax value at 175. This Dmax defined as the largest linear dimension of the

particle (Putnam, Hammel et al. 2007). The Dmax of 175, amino acids residue number of

1256 (=8 x 156), and symmetry of p42, was imposed for reconstructing the 3D structure

of AcsD octamer, using the GASBOR program (Svergun 2001b). This program generates

several bead models of all possible structures that can be obtained by populating the

sphere of diameter equal to 175 Å with 1256 atoms with a symmetry of 4 x 2. This value

of p42 symmetry was used based on the information obtained from the crystal structure

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which is tetramer of dimers. Some of these representative models are shown in Figure

4.8a.

Figure 4.7: Plot for pair distribution function derived using GNOM: The scattering curve of AcsD was Fourier transformed by GNOM program (Semenyuk 1991) to obtain the pair distribution function. The P(r) function is zero at r=0 and r > Dmax. The largest value of r at which P(r) is zero is considered Dmax.

The C-terminal end of the proteins are found extending in all structures and all possible orientation of these chains are observed in all the models. The protein associates in form of a cylinder with an upper and lower layer, surrounding a central pore. The average structure was derived from 50 independent models, using the DAMVER program

(Figure 4.8). DAMFILT software finds the regions in the protein structures and identifies regions of highest densities and selects for the filtered structure

(www.emblhamburg.de/ExternalInfo/Research/Sax/). This structure when superimposed with the available crystal structure of the protein, agrees well with the latter (Figure

4.8c). However, when the filtered structure is superimposed with the average structure, the former is of a greater dimension (Figure 4.8d). This is due to the fact that it is the 102

representation of all the models hence contains all the orientation of the atoms and therefore a much larger distribution of particles is observed.

From all the structures obtained using the SAXS analysis, the protein associates in a manner similar to that shown in the crystal structure. When the two structures were compared as shown in Figure 4.9, the tilt observed between the two tetrameric stacks can also be seen in the SAXS structure in both the top view and the side view. Our data confirm that the protein exists as an octameric complex in-solution and similar to the crystal structure, the essential features of the protein, central pore and the twist in the molecule, are seen in our analysis as well.

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a b

Side view

Top view

Figure 4.8: Representative bead models of AcsD generated by the GASBOR program : 3D shape reconstruction of AcsD was by imposing a p42 symmetry (derived from crystal structure) and residues of amino acids in the AcsD protein. a) Two of the many globally distributed bead models are shown. inside a sphere of diameter equal to the Dmax (175) and number of beads equal to the number of amino acid residues. b) Average structure (pink beads) and the filtered structure of the protein (blue beads). 104

c b d

a

d

c) Filtered structure of the AcsD octamer is (blue beads) superimposed with the crystal structure (red ribbon). d) The average structure is superimposed with the filtered structure and crystal structure shown in (b) to show the comparatively large dimensions of the former, since it takes into account all possible orientations of the protein in space.

*The ribbon structure of the AcsD crystal was obtained from the pdb database (Hu Gao et al 2010)

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a

b

Figure 4.9: Comparison the contours of the solution structure and the crystal structure of AcsD: The solution structure is represented in blue burred image and the crystal structure is represented in bead model with each monomeric chain depicted in a different color a. Top view: The central pore in the crystal structure is seen as a cavity in the solution structure. It can be seen that the lower layer of tetramers is slightly offset against the upper layer. The tilt in the alignment of the two layers is indicated by arrows. b. Side view: The slight slant in the alignment of the dimeric subunits in the crystal structure is reproduced in the solution structure. The structures were made using Sculptor software (sculptor.biomachina.org). The crystal structure file (2z9e) was downloaded from the protein databank (www.rcsb.org).

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DISCUSSION

The cylindrical octamer organized as a tetramer of dimers with a central hole, is the structure arrived at, by us in our SAXS analysis. In addition to SAXS, all our characterization techniques, for the determination of in-solution oligomerization and for structural analysis of the AcsD protein, converge towards identifying that the protein exists as an octamer in solution. This structure is composed of residues arranged in two layers, in such a way that the interfaces between the dimers form four tilted passageways that are shown to interact with four glucan chains. The AcsD structure determined from our SAXS analysis shows, these spiral passageways in form of notches between the dimeric units shown in the Figure 4.9. The presence of these tilted interaction sites for glucan chains are presumed to contribute to the spinning the glucan chains and assembling them together. The involvement of AcsD in forming the passage for cellulose extrusion has been suggested previously (Hu, Gao et al. 2010). The structure of the octameric complex, with a central pore, confirms the assumption that AcsD could serve in the passage of the glucan chain outside the bacterial cells.

It is known that the there are no homologues of this protein in the plant kingdom.

However, AcsD protein sequence is the most conserved of all the acs operon-encoded protein, among all the cellulose-synthesizing bacteria. Thus, the contribution of this protein towards the unique properties should be explored further. With our initial finding of the periplasmic localization together with this structural characterization, we propose that the AcsD protein, serves as the channel in the periplasm through which the newly- 107

synthesized cellulose fibers pass as they are enter the periplasm (Figure 7.1, Chapter VII).

The twists in the internal walls of the AcsD cylinder that interact with the cellulose fibers,

The structure of this protein not only enables passage of the glucan chains through the aqueous periplasm but also imposes a spin in the glucan chains and serve to assemble the fibers. It has also been shown that cellulose synthesis, assembly and crystallization are cell-directed process and occur simultaneously (Haigler, Malcolmbrown et al. 1980). It has also been observed that the bacterial cellulose has an inherent twist in its structure

(Colvin 1961).Our model serves to explain the mechanism behind this twist in bacterial cellulose as well as corroborating the observation that deletion of this protein results in a lower yield of cellulose and an alteration in its crystalline nature (Saxena, Kudlicka et al.

1994; Hu, Gao et al. 2010). Our model agrees well with the earlier finding by Saxena et al. (Saxena, Kudlicka et al. 1994) that, in the absence of this protein, cellulose synthesis by the CM-bound AcsAB is unaffected and cellulose extrusion from the OM pore, is still possible with the passageway through periplamic space is lost. This means that the glucan chains are not assembled well to facilitate their extrusion through the porin. This results in extrusion of a mass of unassembled fibers leading to loss in crystallinity and some of the fibers tend to accumulate in the periplasmic space. Consequently, instead of being bundled and directed towards the OM, this results in reduction of the amount of cellulose extruded.

Considering that all the Acs proteins are encoded by genes, which are part of a single operon, the octamerization of the AcsD, poses a question about the oligomerization status of the other Acs proteins. Since the antibodies against all the four proteins have

108

been generated, as part of my work for this thesis, the quantification of these proteins, can inform us about the stoichiometry of these proteins in the G. hansenii cells.

CONCLUSIONS

We have shown that the AcsD protein exists as an octamer in solution and assumes a cylindrical structure, with a central pore. Although the crystal structure of this protein has been determined before our work, we have obtained a fair idea of how the protein exists under solution conditions. The various arrangement of the end chains in the structure suggests the possibility that the protein is capable of assuming different conformations in solution, a fact that cannot be seen in the crystal structure of the protein.

However, the arrangement of the monomers around a central pore in the form of two stacked tetrameric rings, is in agreement with the crystal structure of the protein.

Facilities that were used for this work:

1. Use of the Advanced Photon Source at Argonne National Laboratory was supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357 and project ID APS ESAF 18-ID-2010.

2. Analytical Ultracentrifugation was performed at the Biotechnology Bioservices Center in the University of Connecticut. The samples were shipped, on ice, and all the analysis was done by Lary, J. W.

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CHAPTER V

BIOCHEMICAL CHARACTERIZATION OF ACSAB PROTEIN,

THE CELLULOSE SYNTHASE OF G. HANSENII 23769

INTRODUCTION

Bacterial cellulose synthase is composed of a catalytic domain and (Lin, Brown et al. 1990) a regulatory domain (Mayer, Ross et al. 1991). The regulatory domain binds to cyclic di GMP, which has been shown to be the allosteric activator of this protein (Ross,

Weinhouse et al. 1987; Mayer, Ross et al. 1991). The catalytic domain binds to the substrate UDP-glucose and polymerizes glucose units processively to form a cellulose chain. This subunit contains the D,D,D and QXXRW motif (Saxena, Brown et al. 1995;

Saxena and Brown 1997; Saxena and Brown 2000; Saxena, Brown et al. 2001), which is a characteristic feature, conserved in all glycosyltransferases that use a nucleotide sugar as a glycosyl donor (Saxena, Brown et al. 1995). In most bacterial species, the different subunits of cellulose synthase are encoded by two or more genes. In Enterobacteriaceae species like Salmonella typhimurium (Hatta, Baba et al. 1990) and Escherichia coli

(Hatta, Baba et al. 1990; Perna, Plunkett et al. 2001), the ORFs bcsA and bcsB (bcs: bacterial cellulose synthase) encode the catalytic and regulatory subunits of the protein, respectively. In case of Gluconacetobacter, it has been shown that G. xylinus NBRC

3288 (Ogino, Azuma et al. 2011) contains three ORFs coding for the cellulose synthase, while the strains JCM7664 (Umeda, Hirano et al. 1999) and 1306-3(Wong, Fear et al.

1990) contain acsA and acsB as two distinct ORFs encoding for cellulose synthase.

However, in the strains ATCC 23769 and ATCC 53582, the genes are fused to form one open reading frame (ORF).

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Cellulose synthase purified using product entrapment from A. xylinum 53582, was shown to be composed of 93 kDa and 83 kDa polypeptides (Lin 1989). Photolabeling studies using radioactive UDP-glucose as the substrate demonstrated that the 83 kDa polypeptide is the substrate-binding domain of the enzyme (Lin, Brown et al. 1990). The gene encoding this catalytic subunit of the protein was identified by sequencing the protein (Wong, Fear et al. 1990) and analysis of the cellulose-deficient mutants (Wong,

Fear et al. 1990). Based on biochemical and sequencing data, the 93kDa protein was proposed to be the cyclic di GMP-binding protein and was suggested to be associated with the CM (Bureau and Brown 1987; Saxena, Kudlicka et al. 1994). However, using freeze-fracture labeling techniques with antibodies raised against the 93 kDa protein, this protein is localized in the protoplasmic fracture (PF) face of the OM and concluded to be a CM protein (Kimura, Chen et al. 2001).

Mayer et al. (Mayer, Ross et al. 1991) showed that in strain 1306-21, cellulose synthase is composed of three major peptides of molecular weight 90, 67 and 54 kDa.

The gene encoding the 90 kDa polypeptide was cloned and expressed in E. coli (Mayer,

Ross et al. 1991). It was found in Western blots using these antibodies generated against these polypeptides that the 67 and 54 kDa peptides were cleavage products of the 90 kDa polypeptide, proving that there is a possibility of post-translational processing of the cellulose synthases. AcsB/ BcsB was found to bind to the activator cyclic diGMP, which led to the general agreement that the B subunit is the regulatory domain (Amikam and

Benziman 1989; Mayer, Ross et al. 1991). However, recently this regulatory role for

AcsB has been questioned (Amikam and Galperin 2006).

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In case of the strain of Acetobacter ATCC 23769, which has been the center of our work, the acs operon harbors a single gene (acsAB) encoding cellulose synthase. This feature of fused-ORF is also seen in the other two cellulose synthase genes in the genome which are not part of an operon. The polypeptide sequence of the translated product shows that the encoded protein contains 1550 amino acids and has a molecular weight of

168,161 kDa. This protein is predicted to contain 10 transmembrane helices (TMHs) and also two large globular non-membrane-bound regions (Figure 5.1).

In this work we have heterologously expressed and purified the non-membrane bound regions of the protein. Using specific antibodies against these regions, we have found that the protein is processed into three polypeptides. The polypeptide of 45 kDa containing the catalytic domain localizes in the CM, but the larger polypeptide (95 kDa) localizes in the OM. We have also shown that this polypeptide does not contain the PilZ domain. Instead, the short stretch of 34 kDa region between the AcsA and AcsB subunits harbors the regulatory, cyclic di-GMP-binding PilZ domain. In addition to determining this processing pattern, we have also determined the organization of the Acs proteins in the membrane compartment.

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1 100 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1550

CesA 133 364

355 471 555 653 Glycosyltransferase PilZ

129 397 610 1550 AcsAB2

AcsAB 581 600 721 740 P1 P2 Figure 5.1 Depiction of the heterologously expressed regions of the AcsAB protein The polypeptide sequence of the AcsAB protein was analyzed using the TMHMM software tool (Krogh, Larsson et al. 2001) in order to identify the transmembrane-bound and non-membrane-bound regions. The AcsAB protein sequence is depicted in the form of a black line with the transmembrane helical regions as white boxes. The conserved cellulose synthase (CesA), glycosyltransferase and PilZ domains, identified using the Uniprot software (Consortium 2012), are indicated in grey boxes. The sequences between 129 - 397 and 568 -1510, do not contain any membrane spanning domains. Polypeptide sequences selected from these non-membrane spanning regions were heterologously expressed and used for antibody generation, are shown as AcsAB1 and AcsAB2. The peptide regions, P1 and P2 were selected for synthesis and antibody generation. The numbers flanking the lines and boxes, show the position of N- and C- terminal position residues of the polypeptides with respect to the full-length protein.

MATERIALS AND METHODS

Cloning and heterologous expression of the acsAB gene regions

Two major non-membrane-bound regions of the AcsAB protein were identified

using prediction tools, TMHMM (Krogh, Larsson et al. 2001) and HMMTOP (Tusnady

and Simon 1998; Tusnady and Simon 2001) as shown in Figure 6.1. The procedures for

cloning and transformation of the gene regions coding for the non-membrane-bound parts

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of the protein (depicted in Figure 5.1), was performed using the procedures described in

Chapter III for AcsD expression. Using the primer pairs (5'-gctagcatgttccagacgatcgcgccg-

3' and 5’-ctcgagccgctggcccccatgacag-3’), containing the Nhe1 and Xho1 restriction site, the acsAB1 was amplified from G. hansenii cells, by colony PCR (Figure 5.2). Similarly primers, 5'-gctagcgccgcggtaaagatgtcatgg-3' and 5’-ctcgagcgacttgcgcctct-3’, were used to amplify the acsAB2 gene fragment. The PCR products was ligated into the pGEM-T easy vector and digested with Nhe1 and Xho1 restriction enzymes, to enable cloning into the pET-21a vector (Novagen).

Expression of the AcsAB soluble regions

Expression vectors were transformed into the BL-21 (DE3) cells and plated on

LB-ampicillin plates to isolate colonies containing the plasmid. A single colony was inoculated from this plate into a culture volume of 5ml in a shaker flask and cultured overnight at 30C and 220 rpm. This culture was used to inoculate a liter of LB 50 μg/ml of ampicillin. The cultures were grown at 37 C at 220 rpm until the absorbance at 600 nm reached 0.4-0.6. At this point, the protein expression was induced by adding IPTG to a final concentration of 1 mM and continuing the culture for 3h. Samples of induced and un-induced culture (20 μl) were subjected to SDS-PAGE to confirm protein expression

(Figure 6.3). The cells were harvested by centrifugation at 5,000 × g for 15 min at 4 C.

The cell pellet was resuspended in 5 ml of lysis buffer (100mM NaH2PO4, 10 mM Tris-

Cl, 8 M Urea pH 8.0) per gram of wet weight and lysed by stirring for 60 min at room temperature. The lysate was centrifuged at 2,500 × g for 30 min at room temperature to pellet the cell debris. For SDS-PAGE, 20 μl of the supernatant was boiled in 5 × SDS- sample buffer and subjected to electrophoresis. 114

Protein purification using denaturing method

The protein concentration of the supernatant obtained after cell-lysis, was determined by Bradford (Bradford 1976). The supernatant was mixed with 50% Ni-NTA slurry at a concentration of 1ml of resin per 10 mg of protein. The lysate was allowed to equilibrate with the resin by incubation on a rotary shaker for 60 min at room temperature. The lysate-resin mixture was loaded onto a column which was washed with an equal volume of wash buffer (100 mM NaH2PO4, 10 mM Tris-Cl, 8 M Urea pH 6.3). and proteins were eluted by passing 0.5 ml of elution Buffer 1 (100 mM NaH2PO4, 10 mM Tris-Cl, 8 M Urea pH 5.8) four times and elution Buffer 2 (100 mM NaH2PO4, 10 mM Tris-Cl, 8 M Urea pH 4.5) four times through the column. All the fractions (flow through, wash and elutions) were analyzed by SDS-PAGE and saved in -20C.

Expression and purification of AcsAB2 was performed in a similar manner (Figure 6.3 b).

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1 2 3 4

3000 bp 2443 bp bp

1636 bp

1000 bp 828 bp 500 bp 5

06 bp

Figure 5.2 Agarose gel of amplified products of acsAB gene regions. PCR amplification of the 828 bp acsAB1 (Lane 2) and 2443 bp acsAB2 (Lane 2) gene sequences was done from G. hansenii cells using colony PCR. Molecular weight ladder was loaded in lane 1 and lane 3 is an empty lane.

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a b

Figure 5.3 SDS-PAGE of heterologously-expressed AcsAB polypeptides a) SDS-PAGE analysis of with protein fractions from AcsAB1 expression and purification: 1: Uninduced cell pellet; 2: Induced cell pellet; 3, 4: Flow through after Ni-NTA purification; 5, 6: Washes from Ni-NTA column; 7, 8, 9: Elutions containing pure AcsAB1 at 31kDa; 10: Molecular weight ladder. b) SDS-PAGE analysis of with protein fractions from AcsAB2 expression and purification: 1: Molecular weight ladder; 2: Uninduced cell pellet, 3: Empty lane; 4: Induced cell pellet. 5, 6: Flow through after Ni-NTA purification; 7: Wash from Ni-NTA column; 8, 9: Elutions containing pure AcsAB2.

Antibody generation, purification and Western blot

The polypeptides were subjected to SDS-PAGE and the bands were cut from the

gel and sent to Covance research products for antibody generation in rabbits. Antibodies

were also generated against two peptide regions in the protein, that were predicted to be

antigenic as well as non-homologous to any other protein in the cells. The anti-AcsAB1

and anti-AcsAB2 antibodies were affinity purified as described in Chapter III. Peptide

antibodies were affinity purified by using Sulfolink agarose G-resin following the

procedures given in the Sulfolink immobilization kit for peptides (Thermo Scientific,

Pierce Research products). All antibodies were stored as 1ml aliquots at -70C. Western

blots were performed using standard procedures using 20 µg of TM or whole cell

proteins. 117

Sucrose density gradient centrifugation

TM was isolated as described in Chapter III. A total of 12 mg of TM protein was loaded onto a 35-ml 25%-55% discontinuous sucrose gradient. The gradient was centrifuged for a duration of 16 hours at 177,500 × g. The fractions from the gradient were collected manually and stored at -70C as 3 ml aliquots. In order to locate the Acs proteins in these fractions, 50 μl of each fraction was boiled in 10 μl of 5× SDS-sample buffer, and subjected to SDS-PAGE followed by Western blotting using anti- AcsAB1, anti-AcsAB2, anti-C and anti-D antibody.

RESULTS

Predicting the soluble regions of AcsAB protein

The amino acid sequence of AcsAB was analyzed by the TMHMM software

(http://www.cbs.dtu.dk/services/TMHMM/). The software predicted two major non- membrane-bound regions and ten TMHs in the protein. The outcome of this prediction is shown in Figure 5.1. Based on the predictions, there are two major regions devoid of

TMHs. These are between amino acids 129 - 397 and 568 - 1510. The region from 129 -

397 contains the conserved QRVRW motif for glycosyltransferases (Saxena, Brown et al.

2001) as well as the domain (DXD, D) (Saxena, Brown et al. 1995; Saxena and Brown

1997; Saxena, Brown et al. 2001), residues involved in substrate binding.

We amplified the portions of the acsAB gene, which encode for the predicted non-membrane bound parts of the protein and heterologously expressed and purified these polypeptides. The Coomassie-stained gel showing the SDS-PAGE analysis of the crude and purified proteins is shown in Figure 5.3. The two heterologously expressed 118

regions of the protein are a 31 kDa AcsAB1, that contains the regions between amino acids 129 - 397 and a 102 kDa-AcsAB2, which contains amino acid sequence from 610 –

1550. Both these polypeptides were expressed with a C-terminal hexa-histidine tag.

These purified polypeptides were then used as antigens to obtain polyclonal antibodies to be used in Western blots, to locate cellulose synthase proteins in the G. hansenii whole cell and TM.

Western blot using anti-AcsAB1 and anti-AcsAB2 antibodies

The antibodies against AcsAB1 and AcsAB2 were expected to cross-react with a single 168 kDa-protein, in Western blot of G. hansenii whole cell and TM proteins.

However, as shown in Figure 5.4a, the anti-AcsAB1 antibody identifies a protein with molecular weight between 45-52 kDa, in the Western blot. The precise molecular weight of the this polypeptide was calculated to be 46 kDa, by plotting the logarithm of the molecular weight against the distance of migration of the protein bands, using the commercial molecular weight marker as the standard (Figure 5.5). The anti-AcsAB2 antibody binds to a 95 kDa band in whole cells. In case of the AcsB2 fragment, the molecular weight directly matched to the 95 kDa band, as seen in Figure 5,4b, and therefore a graph was not used to calculate the molecular weight. This indicates that the

AcsAB protein is cleaved into at least two polypeptide fragments of molecular weights

46 kDa (AcsA) and 95 kDa (AcsB). However, their molecular weight does not add up to

168 kDa, which is the calculated molecular weight of the entire protein encoded by acsAB gene.

Western blots using anti-peptide antibodies

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In order to further understand the cleavage pattern of the protein, antibodies were generated against two synthesized-peptides corresponding to the selected regions of

AcsAB protein (Figure 5.1). The anti-peptide antibody was generated in order to find if there are other processed fragments of this protein that had been left undetected by the anti-AcsAB1 and anti-AcsAB2 antibodies. The two synthesized peptides correspond to twenty amino acid-regions between residues 581- 600 and 721 - 740, in the full-length protein (shown as blue lines in Figure 5.1). One of the peptides (P1), spans the region between 581 to 600, in the AcsAB protein, that lies between the expressed-AcsAB1 (129

- 397) and AcsAB2 (610 - 1550) polypeptides. The other peptide (P2), spans the region from 721 to 740 within the AcsAB2 polypeptide (610 - 1550). The antibody generated these peptides (anti-P1P2) was expected to potentially identify the sequence not identified by the anti-AcsAB1 and anti-AcsAB2 antibodies. Both peptides were injected in one rabbit and therefore, the anti-serum contains antibodies against both the peptides.

The antiserum was affinity purified to using both P1 and P2 peptides.

When the affinity purified anti-P1P2 antibody was employed in a Western blot against whole cellular proteins, the predominant bands seen on the membrane were of molecular weights in 95 kDa and 34 kDa, as shown in the Figure 5.4c. This antibody does not cross-react with heterologously expressed AcsAB1 but detects the AcsAB2 band in the Western blot. This indicates that the 95 kDa protein recognized by anti-AB2 and anti-P1P2 antibody are the same polypeptide, but the 34 kDa band recognized by anti-

P1P2 antibody lies in between the 45 kDa and the 95 kDa polypeptide sequences.

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a b c

Figure 5.3 Western blot using specific polypeptide and peptide antibodies a) Anti-AcsAB1 antibody recognizes a band between 42 kDa and 52 kDa in both whole cells (Lane 2) and TM (Lane 3). Lane 1 is the molecular weight ladder. Lane 4 contains the heterologously-expressed and purified AcsAB1. b) Anti-AcsAB2 antibody binds to a protein of molecular weight 95 kDa, in both whole cells (Lane 1) and TM (Lane 2). This band. Lane 1 contains the molecular weight ladder. Lane 4 contains the heterologously-expressed AcsAB2 protein as a positive control. c) Anti-peptide antibodies recognize a band at 95 kDa and a 34 kDa band in whole cells (lane 2). The anti-peptide antibodies do not bind to pure AcsAB1 protein (lane 3) but recognize the 95 kDa AcsAB2 band (lane 4).

3 y = -0.0305x + 2.8835 2.5 R2 = 0.9928

2

1.5 Log MW 1

0.5

0 0 10 20 30 40 50 60 Distance (mm)

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Figure 5.5 Graph for Molecular weight determination of processed AcsA polypeptide. Standards loaded in the gel were from the commercially available molecular weight ladder (Fermentas Spectra MulticolorBroad Range Protein ladder). The molecular weights of the ladder were 260k Da, 135 kda, 95 kDa, 72 kDa, 53 kDa, 42 kDa, 34 kDa, 26 kDa, and 17 kDa. Substituting the value of x in the line equation for the distance of migration of the cellular AcsA band (30mm) (lanes 1 and 2 in Figure 5.4a), we get the number 1.967, which corresponds to the molecular weight of 46 kDa. The distance of migration of the pure AcsA protein (45.5 kDa) was also calculated by substituting the value of x in the equation. The calculated molecular weight of the pure protein (31.3 kDa) matched that of the molecular weight derived from the protein sequence (31.5 kDa).

Sucrose density gradient centrifugation

Cellulose synthase protein has been localized previously to the CM compartment of the bacterial cells (Bureau and Brown 1987; Kimura, Chen et al. 2001). We wanted to explore if both AcsA as well as AcsB polypeptides were localized in this membrane compartment. We therefore subjected the TM to sucrose density gradient in order to separate the CM and OM fractions. After the high-speed ultracentrifugation at 82,500 x g, the TM is fractionated into CM and OM compartments which distribute themselves in the sucrose gradient based on their density. The CM was a slightly yellow band at 35% density and OM is seen as a denser, white band at 45% density. When aliquots from the sucrose density gradient were collected, majority of the CM band was in second aliquot and the major portion of the OM was in aliquots 8 and 9.

When all the fractions were subjected to Western blot, AcsA and AcsC are seen predominantly in the CM and OM respectively, as was expected. AcsD is distributed throughout the gradient. AcsB is also found through-out the length of the gradient, but a greater amount of the protein, as indicated by the thickness of the cross-reacting band in the Western blot, is in the fractions where AcsC is present. This implies that the AcsB is not in the same membrane compartment as AcsA. This protein is either periplasmic with a strong association with the OM,

122

through its interaction with AcsC, or it is tethered to the CM and exists as a peripheral membrane protein with a large portion exposed to the periplasmic side of the OM.

a

b

Figure 5.6 Western blot of fractions from sucrose density gradient of TM a. Representation of the discontinuous sucrose density gradient: The numbers indicate the fractions collected from the various layers of the density gradient. OM proteins formed a very dense white band (7 and 8) at 45 -55% density of sucrose and CM formed a yellowish band (3and 4) at 35% sucrose density. Each fraction was 3 ml in volume. b. Western blot of fractions from the gradient: Fractions from the sucrose density gradient were collected and saved in 3ml aliquots. A sample of 50 μl each fraction of the sucrose gradient

123

was subjected to SDS-PAGE. Western blot using specific antibodies against Acs A, B,C, D proteins, shows that the AcsC protein is seen predominantly in the OM compartment (lanes 7 and 8) and AcsA in CM fraction (lanes 2, 3). A dense band of AcsB was observed in the fractions in which AcsC was present, indicating the association of the AcsB protein with the OM. TM (control lane) contains a 10 μg TM protein to serve as positive control.

DISCUSSION

Our results using antibodies against the different regions of cellulose synthase show that there are predominantly three polypeptides generated by the processing of this protein. These polypeptides are of molecular weight 46 kDa, 34 kDa and 95 kDa. We have named these processed polypeptides as AcsA, AcsB1 and AcsB2 respectively. Since such a processing of cellulose has not been shown before, we searched for the cellulose synthse sequences from other bacterial species to locate subunits of the protein that are similarly processed. We found that in

Escherichia coli str B171 (Perna, Plunkett et al. 2001; Aziz, Bartels et al. 2008) and

Xanthomonas campestris sp. (Thieme, Koebnik et al. 2005; Aziz, Bartels et al. 2008), the cellulose synthase is encoded by three ORFs. The three ORFs encode 3 polypeptide regions of the protein. The first of the three genes codes for the catalytic subunit containing the D, D, D and QXXRW motifs. The protein encoded by the second gene contains the PilZ domain and the third gene codes for the largest polypeptide whose function is unknown, though it is classified as a BscB protein.

We further explored the sequence of G. hansenii cellulose synthase in the light of the calculated molecular weights of the polypeptides and tried to find if the protein is processed in a manner identical to the ones from the E. coli (Perna, Plunkett et al. 2001) and X. campestris

(Thieme, Koebnik et al. 2005). Heterologously expressed AcsAB1 is a 31 kDa protein that spans

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residues between 129 – 397 of the 1550 amino acid-long protein sequence derived from the acsAB gene. The antibody raised against this protein recognized a 46 kDa band in Western blot.

Figure 5.7a Analysis of AcsB2 sequence using Signal P Gram negative (Petersen, Brunak et al. 2011) The signal P-output is in the form of a plot of probability scores versus the position of the amino acid residue in the sequence. A high S-score indicates that the corresponding amino acid is part of a signal peptide, and low scores indicating that the amino acid is part of a mature protein. A C-score is significantly high at the cleavage site. Y-score is a derivative of the C-score combined with the S-score resulting in a better cleavage site prediction than the raw C-score alone.The cleavage site is assigned from the Y-score where the slope of the S-score is steep and a significant C-score is found. Based on this prediction, we can say that the signal peptide is between amino acid residues 725 and 740 with a predicted cleavage between QAA-SAP.

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A A

A A P S A V Q R K

A Q

725 735 745 755 765 775 785 Amino acid residues

Figure 5.7b Lipo P prediction of a signal sequence in the AcsB polypeptide (Rahman, Cummings et al. 2008) When the sequence of AcsB (642 - 1550) polypeptide derived from our calculations, was analyzed by the LipoP software, the signal peptide (SP1) was predicted to be between residues 736 - 745. The Y-axis shows probability of the signal peptide region in logarithmic scale and the X-axis is the number of amino acid residue in the sequence of the polypeptide. Since this software can predict only signal peptides in the first 30 amino acid stretch, when the entire sequence starting from 694th amino acid residue was analyzed, it could not detect the signal peptide.

Based on the results from the Western blot using the anti-peptide and anti-AcsAB2 antibodies, we conclude that AcsB1 is the regulatory domain containing from residues 409 to

693, with a calculated molecular weight of 32 kDa. This subunit harbors the PilZ domain (572 -

647) (Amikam and Galperin 2006; Ryjenkov, Simm et al. 2006) that binds to the c-di-GMP.

AcsB2 portion of the protein extends from amino acid residue 694 to 1550, with an exact molecular weight of 91,633 kDa. These are predicted sites of the processing of the AcsAB protein, based on molecular weight and cleavage pattern of similar proteins from other organisms

(Perna, Plunkett et al. 2001; Aziz, Bartels et al. 2008) and experimental evidences are required for exact identification of the N- and C- termini of each polypeptide.

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In order to determine the exact site of cleavage and the sequences of these resultant polypeptides we tried to isolate the 95 kDa band from the G. hansenii whole cells and determine its sequence by N-terminal sequencing methods. However, we were unable to isolate sufficient protein for required for the sequencing. Since the exact role of AcsB2 polypeptide is not known, we used the specific antibodies generated against the AcsB2 region of the protein to locate this protein in the membrane compartments. We found that this protein co-localizes with the AcsC protein in the OM compartment. Our results are in consensus with the freeze-fracture study by

Kimura et al. (Kimura, Chen et al. 2001), which revealed that the 93 kDa-polypeptide associates with the PF face of the OM. Though it was concluded from this finding that the protein is localized in the CM, we believe that their results are directing towards the possibility of the

AcsB2 region to be in association or exposed to the OM.

To corroborate our finding from the sucrose density gradient, we analyzed AcsB2 polypeptide to find if there is a possibility of this polypeptide to exist in the periplasm or in association with the OM-bound AcsC. When the sequence was analyzed using Signal P

(Petersen, Brunak et al. 2011) (Figure 5.7a) and Lipo P (Rahman, Cummings et al. 2008) (Figure

5.7b) software tools, we found that both the prediction software identify a signal peptide of 25 kDa in the sequence between the residues 725 to 740, with a cleavage site between residues 740 -

741. This implies that the AcsB2 polypeptide is either anchored to the CM with a large portion exposed to the periplasm or is released into the periplasm. Another analysis software PSLpred

(Bhasin, Garg et al. 2005) predicts the substrate binding regions (133- 364) and PilZ domain

(555- 653) of the AcsAB sequence to be cytoplasmic, but identifies the of AcsB2 region (694 to

1550), to be periplasmic. More studies are required to exactly predict the nature of the signal peptide and verify that it serves as a translocation signal. However, all the predictions

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corroborate our finding, that the AcsB2 polypeptide is largely associated with the OM with a major portion in the periplasm.

The existence of more than one polypeptide subunits in the cellulose synthases from all bacterial species (Perna, Plunkett et al. 2001; Thieme, Koebnik et al. 2005; Ye, Lan et al. 2010), proves that the functional cellulose synthase enzyme requires catalytic and regulator domains to be harbored in distinct poylpeptides. Therefore, the functional and evolutionary significance of the fused gene should be explored in G. hansenii strains 23769 and 53582 (Lin, Brown et al.

1990; Kawano, Tajima et al. 2002) given the fact that the enzyme still requires three polypeptide subunits. There is much to be understood regarding the structure of the cellulose synthase and the localization and organization of all the processed subunits of this proteins and how they come together to contribute to the functioning of this enzyme. But, our data from this work leads us to explore the cellulose synthase sequence in a new light, for structural studies as well as organization of the cellulose synthase complex.

CONCLUSIONS

1. The cellulose synthase in the strain G. hansenii 23769, encoded by a single gene is processed into three polypeptides of molecular weights 45 kDa, 34 kDa and 95 kDa. The sizes of these polypeptides suggest that the AscA contains the substrate binding and catalytic regions. AcsB1 harbors the PilZ region and AcsB2 contains no conserved motifs in the sequence, and therefore its role in the process of cellulose synthesis is yet to be understood.

2. Though catalytic activity of the cellulose synthase resides in the CM as shown before, the

AcsB2 portion of the protein is predominantly exposed to the periplasm.

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CHAPTER VI

ISOLATION OF THE CELLULOSE SYNTHASE COMPLEX USING

ELECTROPHORETIC TECHNIQUES

INTRODUCTION

The bacterial cell envelope is a unique structure that performs several crucial functions. Being the outermost boundary of the bacterial cells, it serves as a barrier that protects the cell from external stress (Seltmann and Holst 2002; Talaro 2007). It also provides the site for signal transduction (Greie, Hebestreit et al. 2003; Galperin 2005) and synthesis reactions (Seltmann and Holst 2002; Talaro 2007). It performs the major task of communicating with the surrounding milieu through selective transport of substances to and from the cells (Seltmann and Holst 2002; Talaro 2007). The cell membrane is a selectively permeable structure with specific carrier-mechanisms for import of nutrients

(Erni 2001) and export of metabolic products (Seltmann and Holst 2002; Talaro 2007).

Cellulose is one such metabolic product, which is released in to the extracellular environment by the cells of the Gram negative bacterium, G. hansenii (Schramm and

Hestrin 1954). While it is not known why bacteria secrete cellulose, it is well known that microbes secrete polysaccharides (Whitney, Hay et al. 2011) supposedly for support of the microbial community (Ma and Wood 2009). The cell envelope of a Gram negative bacterium (Brock 1999), is composed of two concentric phospholipid bi-layers, the CM and the OM, separated by an aqueous, peptidoglycan-containing periplasmic space (Filloux, Bally et al. 1990; Seltmann and Holst 2002; Saier 2006). Transport of molecules in these bacteria therefore requires proteins that span the membrane compartments and provide a passageway across these three layers (Filloux, Bally et al.

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1990; Saier 2006). The transport systems required for toxins (Sheps, Zhang et al. 1996), ions (Stintzi, Barnes et al. 2000; Greie, Hebestreit et al. 2003; Debut, Dumay et al. 2006;

Dumay, Debut et al. 2006) and proteins (Delepelaire and Wandersman 2001) have been extensively studied and categorized into several types based on the structure and organization of the secretion or assimilation system (Boos and Eppler 2001; Greie,

Hebestreit et al. 2003; Quinaud, Ple et al. 2007). However, much remains to be understood about the synthesis and secretion mechanism of bacterial polysaccharides like cellulose.

The cellulose synthase AcsAB was initially considered as an integral membrane protein in the CM (Saxena, Lin et al. 1990; Saxena, Henrissat et al. 1995). But our work, described in the previous chapter, has shown that AcsAB is processed into three polypeptides (Chapter V) which span the CM and the periplasm. We have also shown that AcsD is a periplasmic, octameric-protein with a central pore (Iyer 2010). Based on its N-terminal signal sequence and secondary structure-predictions, AcsC is the outer- membrane porin (Saxena, Kudlicka et al. 1994). It contains seven TPR motifs

(Consortium 2012) that are considered as regions for protein-proteins interactions and serve as multi-protein scaffold in polysaccharide export complex (Whitfield and

Mainprize 2010; Whitney, Hay et al. 2011). The hypothesis that the Acs proteins form a complex, has been proposed in the past (Chen and Brown 1996; Endler, Sanchez-

Rodriguez et al. 2010). The existence of a secretion system for other polysaccharides

(Whitney, Hay et al. 2011), also provides us clues towards the nature and composition of the cellulose secretion complex. However, such a complex has so far, not been isolated and characterized.

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In order to explore the exact composition, structure and organization of the cellulose synthesis and extrusion machinery, I initiated studies to purify this complex. I also aimed to identify the core components of the cellulose synthase complex. We have used a combinatorial approach of membrane protein solubilization using mild detergents, and subsequent isolation by gel electrophoresis. We have compared the efficiency of two detergents, dodecyl maltoside (DDM) and Triton X-100, in solubilizing total membrane proteins of G. hansenii and identified that DDM is more efficient in isolating the cellulose synthase complex as well in preserving its enzymatic activity. Our results from blue-native gel (BN-PAGE) shows that all the proteins encoded by the acs-operon are associated in a complex. This experiment also reveals that other proteins involved in the cellulose biosynthetic pathway, from glucose 6-phosphate to UDP-glucose are also associated with this complex.

MATERIALS AND METHODS

Bacterial culture conditions

A G. hansenii cell pellet was obtained from a 60 L culture of the bacterial cells in

SH medium (Schramm and Hestrin 1954), grown for 48 h at 30C. The culture preparation was carried out in the “Shared Fermentation Facility”, Huck Institute of the

Life Sciences. After 48 h of culture, under agitated conditions, 320 g of cell paste was obtained by centrifugation. This cell paste was stored at -70°C. All the membranes used in this study were prepared with this frozen cell paste as the source of Acetobacter cells.

Solubilization of the TM proteins

TM preparations were obtained using the procedure described by Ruebush et al.

(Ruebush, Brantley et al. 2006) and were stored in 10mM Tris-Cl buffer, pH 8.2 in 10% 131

glycerol at -70°C. Isolation of the TM and its separation into CM and OM fractions were performed according Chapter III. Protein concentrations were determined by method described by Bradford (Bradford 1976).

For BN-PAGE experiments, membrane proteins were solubilized using the methods of Wittig et al. (Wittig, Braun et al. 2006). TM aliquots containing 20 mg of protein (6.6 ml) were diluted five-fold in 10 mM Tris-Cl, pH 8.6 and centrifuged at

100,000 × g for 30 min. The pellet obtained was resuspended in solubilization buffer (50 mM sodium chloride, 2 mM amino hexanoic acid and 1 mM imidazole pH 8.2) to obtain a protein concentration of 10 mg/ ml. Dodecylmaltoside was added to obtain a final concentration of 1g per g protein. With Triton X-100, it was added to obtain a final concentration of 1.5 g per g protein. The samples were allowed to solubilize for 10 minutes, on ice. Glycerol was added to a final concentration of 5% (w/v). Coomassie G-

250 was added to give a protein to dye ratio of 8:1. When DDM was used as the detergent, 50 µl of the Coomassie stock was added and 100 µl was added, if Triton X-100 was used. A similar procedure, as described above, was used when 20 mg of CM proteins were subjected to BN-PAGE. In all the detergent-mediated solubilizations, DDM (final concentration 1% w/v) and Triton X-100 (final concentration 1.5% w/v) were used in concentrations much above their CMC values of 0.01% and 0.015%, respectively .

Native gel electrophoresis of the solubilized TM proteins

For zymogram experiments, proteins were solubilized using the same final concentrations of detergent as described above. Solubilized TM, containing 120 µg protein in final volume of 40 µl, was subjected to native polyacrylamide gel

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electrophoresis (Harwood 2000) at 4°C, 5-10 mA constant current until the dye front reached the bottom of the gel.

Zymography

After electrophoresis, individual lanes from the native gel were incubated for 16 hours at room temperature, the lanes were incubated in 10 ml of TME (50 mM Tris-Cl pH 8.0, 10 mM MgCl2 and 1 mM EDTA) containing final concentrations of 20 µM UDP- glucose, 1 µM c-di GMP, 20 mM MgCl2 and 5 mM CaCl2. The composition of the reaction mixture was adapted from the buffer for an in vitro cellulose synthase assay, by

Mayer et al. (Mayer, Ross et al. 1991). For negative control, the gel lane was incubated in a reaction mixture devoid of UDP-glucose. After overnight incubation, the gel lanes were washed for 10 minutes in deionized water followed by staining in 10ml of 0.01%

Calcofluor for 10 minutes. Enzyme activity was confirmed by visualization of

Calcofluor-bound cellulose as a green-fluorescent band under UV light (Monheit, Cowan et al. 1984).

Blue native polyacrylamide gel electrophoresis (BN-PAGE)

The procedure for BN-PAGE was adapted from the method described by Wittig et. al. (Wittig, Braun et al. 2006). This procedure is briefly described in this section.

Gel composition: Acrylamide solution used for the first dimension of BN-PAGE was prepared by mixing 48 g of acrylamide and 1.5 g of bis-acrylamide in 100 ml of water. This is referred to as AB-3 mix (Hjerten 1962). The composition of the gel buffers is provided in Table 6.1b. The gel was assembled using custom-made glass plates (16 cm x 17.5 cm) of and spacers (0.1cm). A 4-15% gradient gel was poured between the glass plates using a gradient maker. Once the gradient gel was polymerized, the 3.5% stacking

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gel was poured and a custom-made comb with 5 wells of dimensions 2 cm x 2.5 cm x

0.1 cm, was placed in it. After polymerization, the comb is removed and the wells are overlaid with 1X gel buffer (Table 6.1b). The gel is covered with wet paper towels and plastic wrap and saved in 4°C until further use.

Electrophoresis conditions: The anode chamber and cathode chambers were filled with respective buffers. The composition of these buffers is provided in Table 6.1b.

A volume of 500 µl of solubilized TM sample was loaded into each well. BN-PAGE was conducted at 4°C with the power supply set at 100 V until the samples entered the gel.

Electrophoresis was continued beyond this point at a voltage of 500 V with current limited to 15 mA. The undiluted cathode buffer (Table 6.1b) was replaced by the diluted cathode buffer after the dye front reached one-third of the total gel length.

Electroblotting of proteins from first dimension BN gel: After electrophoresis, the gel was carefully removed from the cast and the lanes were separated from one another using a gel cutter. The individual lanes were then subjected to a second dimension denaturing PAGE or Western blot. Western transfer was carried out at a constant current of 30 mA for 16 hours.

Second dimension SDS-PAGE: The gel strip from the first dimension was soaked for 60 min in 100 mM Tris-Cl pH 8.0 containing 1% SDS and 1% mercaptoethanol. The strip was washed briefly with water, placed horizontally between two gel plates, and assembled such that the gel strip was towards the bottom of the plate.

This was done to ensure ease of pouring a gel without inducing air bubble-formation. The

4% acrylamide solution was poured first to form a layer of stacking gel immediately above the native gel piece and after polymerization of this gel, the resolving gel was cast.

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After polymerization, the cast was turned the right way up, to obtain the BN-gel strip on the top. The gel was subjected to electrophoresis at initially a constant voltage of 100 V and the voltage was increased to a 300 V after the dye front reached the resolving gel.

Once the dye front reached the bottom of the resolving gel, the electrophoresis was stopped and the gel was stained overnight with a solution of colloidal Coomassie stain

(Neuhoff, Arold et al. 1988).

Table 6.1a: Composition of polyacrylamide gradient BN-gel*

Stacking gel Gradient Resolving gel

3.5% acrylamide 4% acrylamide 15% acrylamide

AB-3 mix 0.44 ml 1.5ml 4.45 ml

Gel buffer 3X 2.0 ml 6.0 ml 5 ml

Glycerol - - 3.0 g

Water 3.4 ml 10.4 ml 2.55ml

10%APS 50 µl 100 µl 100 µl

TEMED 5 µl 10 µl 10 µl

Total volume 6 ml 18 ml 15 ml

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Table 6.1b: Buffers for BN-PAGE*

Cathode Dilute cathode Anode Gel buffer buffer buffer buffer

Tricine (mM) 50 50 - - Imidazole (mM) 7.5 7.5 25 75 Coomassie blue G-250 0.02 0.002 - - (%) 6-aminohexanoic - - - 1.5 acid (M)

*Compositions of the gel and buffers are adapted from the methods described by Wittig et al. (Wittig, Braun et al. 2006).

In-gel digestion of gel bands and spots: The bands visible after the first dimension BN-PAGE or the stained gel spots from second dimension PAGE, were analyzed by LC/MS. In-gel digestions were done according to the instructions provided by the Proteomics core facility, Hershey, PSU. Briefly, the procedure involved excision of a spot or band from the gel and dicing it using a clean unused razor blade into approximately 1-3 mm pieces. The pieces from individual spots/ band were taken in a labeled Eppendorf tube and destained with 200 µl of 100 mM ammonium bicarbonate in

50% acetonitrile, for 10 min at 37ºC. The gel pieces were dehydrated by incubation in 50

µl of acetonitrile for 10 minutes followed by air-drying in a laminar hood for 30 minutes.

Proteins in the gel were reduced in 100 µl of 10 mM dithiothreitol in 25 mM ammonium bicarbonate, pH 8.0, for 30 minutes, at room temperature. The reduced proteins were alkylated by incubation for 30 minutes, in a solution of 100 µl of 20 mM iodoacetamide in 25mM ammonium bicarbonate (pH 8.0). The gel pieces were again dehydrated as

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described above, prior to adding a 20µl of 200 ng/µl of Promega sequencing grade trypsin in 25 mM ammonium bicarbonate. The gel pieces were allowed to soak the trypsin by keeping the tubes at 37ºC for 30 minutes, after which more ammonium chloride was added to completely cover the gel slices, and the tubes were incubated at

37ºC. After 16 h of incubation, the tubes were spun for 30 seconds to collect all the gel pieces at the bottom and the trypsin-containing solution was transferred into an autosampler vial. The gel pieces were then sonicated in a solution of 50µl of 50% acetonitrile and 5% formic acid, to extract the digested peptides. This extract was added to the first extract in the vial. The samples were dried in a vacuum concentrator to a final volume of approximately 10µl and submitted for LC-MS analysis in the Proteomics and mass spectrometric core facility, University Park, PSU.

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a b

1 2 1 2 3

Figure 6.1. Zymogram of detergent-solubilized G. hansenii TM. a. DDM-solubilized TM were subjected to electrophoresis under non-denaturing conditions in a gel composed of 3% stacking and 6% resolving acrylamide. After incubation in a reaction buffer containing UDP-glucose and c-diGMP, staining with Calcofluor results in a fluorescent smear at the top of resolving gel (Lane 1). The staining of the gel in colloidal Coomassie stain shows protein throughout the gel length (Lane 2). b. Electrophoresis of the DDM-solubilized TM in a 4-15% gradient acrylamide gel results in a more discrete band after Calcofluor staining (Lane 2). Similar band of lower intensity (indicated by arrow) is observed when an equivalent amount of TM protein is solubilized with Triton X-100 (Lane 2). However, when the gel lane is incubated in a reaction mixture without UDP-glucose, no fluorescent band is seen in the same region after Calcofluor staining (Lane 1).

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RESULTS

Zymogram of in vitro cellulose synthesis

Native-gel electrophoresis of solubilized TM was performed with a 3% stacking and 6% resolving gel (Figure 6.1a). A total of 120 µg of TM protein was loaded in each well. After electrophoresis, one lane of the gel was incubated with UDP-glucose and cyclic di-GMP in Tris-buffer pH 7.0, as described in Materials and Methods After overnight incubation, at room temperature, the gel was stained with Calcofluor which visualized a cellulose band (Lane 1). The adjacent lane from the gel (Lane 2) was visualized for protein with Coomassie. The two visualization methods are consistent with a large moleccular complex actively catalyzing cellulose synthesis after electrophoretic separation..

Due to the instability of the 3% portion of the gel, the electrophoresis was again performed with a 4-15% gradient gel (Figure 6.1b). Unlike the smear obtained in a 6% gel, proteins migrated in a more discrete pattern in the 4-15% gradient gel. No cellulose was detected in the absence of UDP-glucose (lane 1). Solubilization of the same amount of protein using Triton X-100 results in a fainter band after zymogram reaction and

Calcofluor staining (lane 3).

Selection of an efficient detergent for BN-PAGE

The zymogram described above suggested that an active cellulose-synthesizing complex is stable to electrophoresis. As such, we used BN-PAGE techniques to attempt identification of the proteins involved in this complex. In order to identify the better detergent of the two, both TM and CM were solubilized with DDM and Triton X-100, as described in the "Materials and Methods". The comparative images of all four second

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dimension denaturing gel are shown in Figure 6.2. There are six major complexes in these gels as indicated by the five rows formed by the vertically aligned spots. The most number of spots are obtained in the gel containing TM proteins solubilized with DDM

(Figure 6.2a). When this gel is compared to that of Triton X-100 solubilized TM (Figure

6.2b), the number and intensity of spots was found to be greatly reduced, however, the separation of the proteins still follows the same pattern, in the form of six major complexes. When the second dimension gel with CM proteins, is compared to the that of the TM, several spots in each vertical row are absent, indicating that those spots correspond to OM proteins in TM. The triton-X-100 solubilized CM does not show any discernable protein spot. We aimed to identify the profile of CM and OM proteins by analyzing the spots in the CM and TM gels. MS-based analysis of the trypsin-digested spots from the second dimension gel were uninformative, due to abundance of keratin and/ or very low confidence of proteins detected. Although the second dimension SDS-

PAGE did not give any information using MS-analysis, we used the gel profiles as indicators to identify DDM as the more efficient detergent.

BN-PAGE of DDM-solubilized TM

We performed BN-PAGE of DDM-solubilized TM. A representative gel from the first dimension of a BN-gel is shown in Figure 6.3. Adjacent second and third lane from the same gel were visualized by Western blot using anti-AcsA and anti-AcsD antibody.

Both Western blots showed a band in the same position of the blots.

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a b

a

c d

Figure 6.2 Comparison of the second dimension gel profiles a.TM solubilized with DDM, b: CM solubilized with DDM, e. TM solubilized with Triton X-100 and d. CM solubilized with Triton X-100. The orientation of the gel from BN PAGE is indicated by the horizontal arrow with the arrowhead pointing towards the bottom of the BN-gel. The vertical arrows indicate the proteins that migrate along a linear path, indicating that these proteins are part of a complex

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d

b

1 2 3

a

Figure 6.3 BN PAGE of G. hansenii TM a) Lane 1 of the first dimension BN-gel shows several bands. a) When the BN-gel lane is lined up with the Western blotted lanes from the same gel, it can be seen that the cross-reacting bands in the Western blot align against each other, as well as with, one of the bands in the first dimension gel. The BN-PAGE band aligning with the bands in the Western blots, was subjected

to In-gel digestion followed by LC/MS analysis. The proteins identified by LC-MS are

listed in the Table 6.2. This band contains proteins relevant to the cellulose-biosynthetic

pathway (Swissa, Aloni et al. 1980), in addition to other proteins in the cell. Of the

operon-encoded proteins, only cellulose synthase is detected by this method. AcsD

protein was detected in the Western blot, but not detected in MS analysis. Since

antibodies are more sensitive and accurate than MS-based detection methods, presence

of AcsD in the gel band is well-supported. Thus, other than AcsC, (for which the

antibody was not available at the time of this work), we observe all the proteins that were

known to be involved in cellulose synthesis, in this BN-gel band. 142

Accession number Protein name PLGS score Coverage (%)

putative EFG85019.1 phosphoribosylaminoimidazole 6785.32 20.89 carboxylase catalytic subunit S-adenosyl L-homocysteine EFG83091.1 5711.04 32.33 hydrolase EFG82973.1 hypothetical protein GXY 15499 4678.76 33.70

putative glutamate synthase EFG85152.1 1918.27 46.30 NADPH large chain precursor

EFG85151.1 putative oxidoreductase 1630.00 52.90

EFG85173.1 outer membrane protein OmpA 1345.23 47.19

UTP glucose 1 phosphate EFG84008.1 964.02 36.64 uridylyltransferase

6 phosphogluconate EFG85627.1 813.62 41.14 dehydrogenase like protein

580.85 EFG84192.1 phosphoglucomutase 40.87

BAC82543.1 cellulose synthase subunit AB 469.35 13.35

EFG85976.1 putative phosphoketolase 436.035 17.12

EFG85920.1 chaperone clpB 421.8425 55.23

EFG83882.1 succinate CoA transferase 266.56 41.70

EFG85957.1 hypothetical protein GXY 00199 225.61 20.3

import inner membrane translocase EFG84948.1 170.62 10.13 subunit Tim44 Table 6.2 Proteins detected after LC-MS of the BN-gel band The band in the BN-PAGE gel (shown in figure 6.2b) was in-gel digested and analyzed by LC-MS. The G. hansenii 23769 genome was used as the reference database. PLGS (Protein Lynx Globak server) is a statistical measure of accuracy of the peptide assignement with a higher score indicating a higher confidence of proteins identity (Rosenegger et al. 2010). Proteins that are known to be involved in the cellulose synthesis pathway are indicated in bold font.

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DISCUSSION

Protein-protein interaction studies heavily rely on the availability of specific identification techniques like Western blot (using antibodies against proteins of known sequences), protein sequencing and mass-spectrometry. All these techniques in turn require a database of known protein sequences and therefore by extension, require a fully sequenced genome of the organism. In order to generate a genomic database for proteomic studies, sequenced the genome of A. xylinum 23769 (described in Chapter II ).

Other than genome sequencing, another important contribution to all our experiments described in this chapter, is the generation of antibodies against the acs- operon-encoded proteins. Since we are specifically interested in the proteins contributing to cellulose biogenesis antibodies were made against AcsA, AcsB, AcsC and AcsD proteins. Our intention was to identify all the protein components of the cellulose synthase complex; the more abundant and the unknown ones by mass spectrometry and the known targets using Western Blot.

Since the acs operon-encoded proteins are predominantly membrane-localized, as shown in Chapter I and Chapter IV, we have used selective detergent solubilization to mildly separate the membrane proteins without disrupting the protein-protein interactions in complexes. It has been shown that BN-PAGE allows high degree of separation of membrane proteins under native conditions. The detergents enable solubilization of the proteins in a manner that the integrity of protein complexes is not compromised, but individual complexes are separated from one-another (Schagger and von Jagow 1991;

Wittig, Braun et al. 2006). Coomassie G-250 dye in the sample buffer remains tightly bound to the proteins and imparts a negative charge to the complexes and serves to

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maintain the otherwise hydrophobic membrane protein- complexes in a soluble form

(Schagger and von Jagow 1991; Camacho-Carvajal, Wollscheid et al. 2004). Both Triton

X-100 and DDM have been used as detergent of choice for BN-PAGE of membrane proteins (Wittig, Braun et al. 2006). We found that when TM proteins that are solubilized with DDM were subjected to BN-PAGE, distinct bands corresponding to each protein complex can be seen (Figure 6.2a). Using specific antibodies against the AcsD and AcsA proteins, we located the band in this BN-gel, that contained the cellulose synthase complex. MS analysis of the BN-gel band, identified proteins from cytoplasmic as well as membrane compartments (Table 6.2). The cytoplasmic enzymes detected in the MS- analysis are phosphoglucomutase, UDP-glucose phosphorylase and cellulose synthase

(Swissa, Aloni et al. 1980). Phosphoglucomutase is the enzyme that moves the phosphate group in glucose 6-phosphate from C-6 to C-2, to form glucose 1-phosphate. Glucose 1- phosphate is converted to UDP-glucose, by the enzyme UDP-gluocose phosphorylase.

UDP-glucose is the nucleotide sugar substrate for the CM-bound cellulose synthase, that poymerizes the sugar into cellulose chains (Swissa, Aloni et al. 1980).

Our finding suggests that the cellulose synthase complex contains, in addition to the proteins encoded by the acs operon (AcsA and AcsD proteins detected by Western blot), proteins involved in the synthesis of the substrate, UDP-glucose. This implies that in G. hansenii, the cellulose complex is composed of proteins involved in synthesis as well as secretion of the polymer. This feature is shared by the alginate synthase complex, which contains, all the cytoplasmic proteins involved in synthesis of nucleotide sugar as well as membrane proteins involved in alginate synthesis as part of a large secretion complex.

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Other than the proteins known to be involved in cellulose biogenesis, some other proteins are also seen in this band. At this point we can only speculate about their contribution to the cellulose synthesis pathway. OmpA protein has recently been shown reduce cellulose production on hydrophobic surfaces through induction of stress response system, in Escherichia coli cells (Ma and Wood 2009). The relevance of this proteins in

G. hansenii cellulose synthesis is yet to be understood.

In second dimension gels, it can be seen that there are proteins migrate as five major complexes in the TM and CM gels. From comparing the profiles of these gels, it can be seen that for both TM as well as CM solubilization, more protein spots are visible on the gel, when the TM is solubilized with DDM. We found that the in-gel digestion and subsequent LC-MS of the spots selected from the gels, gave us very poor quality data with very low confidence. None of the proteins detected in these spots corresponded to proteins known to be involved in cellulose synthesis. Thus, MS-based analysis of the BN-

PAGE data has been the biggest bottle-neck to our complete understanding of the cellulose synthase complex.

Another form of native gel technique employed to characterize the cellulose synthase complex was the zymogram method. We have used zymograms as a means of visualizing the cellulose synthetic activity of the detergent-solubilized TM. This assay detects the protein of interest by the virtue of its enzymatic activity by incubating the gel with a suitable substrate and using a product-specific stain as an indicator (Lantz and

Ciborowski 1994; Martinez, Alarcon et al. 2000). When detergent-solubilized TM were subjected to native gel electrophoresis and incubated in the cellulose synthesis enzyme reaction mixture, the band gives fluorescence after staining with the cellulose-binding dye,

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Calcofluor (Figure 6.1). There are several proteins in this band that are not associated with cellulose synthesis and are merely present due to their comparable molecular weights. But, more importantly, our results prove that bacterial cellulose synthase activity can be assayed by zymogram method, without the use of radioactive reagents. Here again, DDM serves as a detergent of choice, as solubilization of the same concentration of protein (120 µg) with this detergent gives a brighter fluorescence after Calcofluor staining, than that obtained after solubilization with Triton X-100.

It has also been shown previously that membrane fractions from A. xylinum when incubated in presence of cyclic di GMP and UDP-glucose, synthesize cellulose (Bureau and Brown 1987) and in vitro cellulose synthesis is possible even after inactivation of

AcsD and AcsC (Saxena, Kudlicka et al. 1994). With this study have proved that, these proteins are nevertheless structurally associated with cellulose synthesis complex.

CONCLUSIONS

An efficient method for solubilization of the total membrane has been developed, that ensures retention of the cellulose synthase enzymatic activity in vitro as well as maintains the protein-protein interactions of the Acs complex. Our work has shown that: i) BN-PAGE of DDM-solubilized membranes is an efficient method for isolation of the minimal complex of proteins related to the cellulose biosynthetic pathway. ii) Cellulose synthesis and secretion machinery is composed of a protein complex spanning the cytoplasm and the membrane compartments. This complex is composed of proteins encoded by the operon and those involved in the pathway from glucose 6- phosphate to cellulose.

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CHAPTER VII

SUMMARY AND FUTURE DIRECTIONS

My doctoral work has opened up several areas where research can be continued to obtain more insights into the biochemistry of cellulose synthesis. One important contribution to the study of bacterial cellulose synthesis, is the generation of antibodies against proteins encoded by the cellulose synthase operon. The antibodies have contributed towards some very insightful discoveries in the course of my work.

The antibodies against different regions of the cellulose synthase protein, revealed the nature of processing of the cellulose synthase protein in vivo. We have shown that the protein is processed into three polypeptides. The 45 kDa N-terminal polypeptide, AcsA, contains the catalytic domain of the protein with substrate binding (Saxena and Brown

1997; Saxena and Brown 2000) and conserved glycosyl transferase motifs (Saxena and

Brown 1997; Saxena and Brown 2000). The PilZ motif is contained in the 34 kDa polypeptide and is considered as the regulatory region of the enzyme. We have named it

AcsB1, to distinguish it from the C-terminal 95 kDa subunit, AcsB2, whose function is unknown.

These antibodies have also led to the identification of the subcellular localization of the protein subunits. We have found that, although AcsA is CM-bound, the AcsB2 protein is largely periplasmic in nature and contains a signal peptide (Rahman,

Cummings et al. 2008) which enables the transport of AcsB2, to this subcellular compartment. MudPIT analysis and sucrose density gradient centrifugation, also provide evidence for the association of this protein to the OM. With these findings, we have shown that, the cellulose synthase in G. hansenii 23769, is composed of multiple

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subunits, similar to the cellulose synthase in other bacterial species (Perna, Plunkett et al.

2001; Thieme, Koebnik et al. 2005). Thus, in spite of being encoded by a single gene, the cellulose synthase from this organism contains the same multi-subunit architecture like that of other bacterial species, which are encoded by more than one gene (Hatta, Baba et al. 1990; Ogino, Azuma et al. 2011).

This finding directs us to enquire into the evolutionary significance of the single gene in G.hansenii. This is also crucial for understanding the structure of this protein.

With the heterologously expressed AcsAB1 region, I have attempted solubilization and crystallization trials, with initial success. The methods which led to crystal formation were not amenable to scale-up. However, with the knowledge of the exact region of cleavage of AcsAB protein, a new set of primers can be designed, that would amplify the gene regions that code for the processed fragments of the protein. This would mimic the cellular polypeptide in G. hansenii and could be a better candidate for structure determination.

The Anti-AcsAB1 antibody can also be used for quantitation of this protein, for purposes of arriving at the stoichiometry of AcsA and AcsD proteins in the cell, as well as deriving the enzyme kinetics for cellulose synthase. In addition to the anti-cellulose synthase antibodies, the anti-AcsD antibody helped to determine the periplasmic localization of the AcsD protein.

Other than the antibodies, a major contribution of my work has been sequencing the genome of G. hansenii 23769. This sequenced genome has been the quintessential reference database for our proteomic experiments. MudPIT analysis of the TM, OM and

CM, has given us the proteomic profiles of these membrane compartments. These

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proteome data are available for further enquiry into the components of these compartments. We have solely concentrated on the cellulose synthesis-related proteins in this proteome data and have found that the proteins involved in cellulose synthesis are distributed in the CM and OM compartments unequally. We find a greater signal for the

AcsB protein from the OM compartment confirming our findings from Chapter V.

This genome has also been used for identification of proteins from BN-PAGE bands. It has also contributed towards identification of binding partners in the cellulose synthesis and extrusion complex. We now know that similar to the other polysaccharide extrusion systems (Pecina, Pascual et al. 1999; Svanem, Skjak-Braek et al. 1999;

Vazquez, Moreno et al. 1999; Keiski, Harwich et al. 2010; Whitney, Hay et al. 2011), the complex contains cytoplasmic proteins involved in the biochemical pathway from the precursor of the sugar-nucleotide substrate to the cellulose. This complex also contains structural proteins which span the membrane compartments, serving as passageways for assembly and release of the cellulose fibers.

Combining the significant findings from all the work described in my dissertation, a model for the bacterial cellulose synthesis and extrusion complex, as shown in Figure

7.1. According to this model, the AcsA protein is the CM-bound cellulose synthase that contains its catalytic region exposed towards the cytoplasm, where it binds to the substrate UDP-glucose. This UDP-glucose is formed in close proximity to the synthase because our data suggest that it is associated with phosphoglucomutase and UDP-glucose phosphorylase. These enzymes catalyze the formation of glucose 1-phosphate and UDP- glucose respectively. Their close association with the cellulose secretion machinery serves to channel cellular glucose towards the cellulose-synthase complex. Cellulose

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synthase is activated by binding of cyclic di-GMP to the PilZ domain. Our findings show that this PilZ domain is also exposed to the cytoplasmic side of the CM. This is in agreement with the known cytoplasmic distribution of the cyclic di-GMP to which the

PilZ domain binds (Amikam and Galperin 2006). This, so far, is the putative organization of the elements involved in the synthesis of cellulose. The mechanism of transport of the synthesized cellulose into the periplasm is unknown and yet to be determined. However, once in the periplasm, data suggest that the cellulose fibers are channeled into the pore formed by the AcsD octamer via the AcsB protein that serves as a scaffold between the

CM and the OM. The passageway through the AcsD pore which has interaction sites for cellulose chains, assembles the cellulose fibers into a bundle. This bundle is twisted due to the tilted nature of dimer-dimer interfaces that serve as interaction sites. This twist in the fibers condenses them and serves to crystallize the chains into microfibrils. The assembled microfibrils are extruded through the porin-like AcsC protein, located in the

OM. Many of the findings that have contributed to this model have been made as part of my work towards this dissertation. This model can be further refined with the knowledge of the stoichiometry of the constituent proteins and their structural characterization.

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Figure 7.1 Working model for the cellulose synthesis complex This model shows cytoplasmic proteins phosphoglucomutase (blue box) and UDP-glucose pyrophosphorylase (purple box) in association with the CM-bound AcsA and AcsB1. UDP-glucose is produced from glucose 1-phosphate by phosphoglucomutase and UDP- glucose pyrophosphorylase. This serves as the substrate for cellulose synthesis by AcsA. Both AcsA and AcsB1 contain transmembrane helical and globular region. The globular region in AcsA exposed to the cytoplasm, contains the UDP-glucose-binding site, whereas the cytoplasm-exposed region of the AcsAB1 contains the cyclic di-GMP- The AcsB2 protein is tethered to the CM but largely exposed to the periplasm with close contact with the AcsC protein. The cellulose fibers produced by the AcsA protein are directed into the pore formed by octameric AcsD, which is localized in the periplasm. These microfibrils are assembled in the periplasm and are extruded through the OM- porin, AcsC.

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Publications 1. Sato S, Feltus FA, Iyer PR, Tien M. (2009) The first genome-level transcriptome of the wood-degrading fungus Phanerochaete chrysosporium grown on red oak. Curr Genet. 55 (3): 273-86. 2. Iyer PR, Geib SM, Catchmark J, Kao TH, Tien M (2010) Genome sequence of a cellulose- producing bacterium, Gluconacetobacter hansenii ATCC 23769. J Bacteriol. 192 (16):4256-7. 3. Iyer PR, Catchmark J, Brown NR, Tien M. (2011) Biochemical localization of a protein involved in synthesis of Gluconacetobacter hansenii cellulose. Cellulose 18 (3): 739-47. Presentations 1. “Biochemistry of bacterial cellulose synthesis: How do bacteria spin the yarn?” 243rd ACS National Meeting, Division of Cellulose and Renewable Materials, San Diego, California (March 25-29, 2012). 2. “Isolation and characterization of proteins involved in Gluconacetobacter hansenii cellulose synthesis” Division of Cellulose and Renewable Materials, 241st ACS National Meeting & Exhibition in Anaheim, California (March 27-31, 2011). 1. “Biochemical characterization of the cellulose synthase complex from Gluconacetobacter hansenii” Energy Frontier Research Center, The Pennsylvania State University (November, 2010). 2. Poster “Biochemical and structural characterization of AcsD, a protein involved in bacterial cellulose synthesis”. Graduate Exhibition. The Pennsylvania State University (March, 2010). 3. “Whole genome sequencing of Gluconacetobacter hansenii 23769” Energy Frontier Research Center Retreat, The Pennsylvania State University (May, 2010). 4. Poster :“Protein-protein interactions in Acetobacter xylinum cellulose synthase complex” 12th Annual Environmental Chemistry Student Symposium, The Pennsylvania State University (March 27 – 28, 2009). 5. Poster :“Study of proteins involved in bacterial cellulose synthesis” Graduate Exhibition, The Pennsylvania State University (March 29, 2009). 6. “Characterization of Acetobacter xylinum Cellulose Synthase” Institute of Biological Engineering, Santa Clara (March 19 – 21 2009). Awards and Scholarships 1. Awarded second position in poster presentation Environmental Chemistry Student Symposium (2010) 2. Awarded first position in poster presentation in Environmental Chemistry Student Symposium (2009) 3. Recipient of the Huck Institutes of Life Sciences Fellowship at The Pennsylvania State University (2006-2007) 4. Recognized and awarded as the “Best Outstanding Student” by Srimad Andavan College of Arts and Sciences, Tiruchirappalli, India (2001) 5. Awarded Merit scholarship for the best academic performance by Department of Biochemistry, Srimad Andavan College of Arts and Sciences, Trichy, India (2001). 6. Won Second Prize in “Chemquiz’97”, an inter-college chemistry quiz event organized by the Pune University Department of Chemistry, Pune, India (1997).