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

ii

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

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

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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

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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.

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

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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

ix

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

x

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

xiv

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

xv

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

xvi

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.

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

2

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

3

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.

4

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 180C 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”.

5

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.

6

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

7

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 13

C-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

8

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

9

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.

10

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

11

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

12

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)

13

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

14

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

15

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

16

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

17

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).

18

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).

19

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 14

C-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),

20

(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

21

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.

22

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-

23

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

24

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 (-32

P)-

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

25

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-(14

C)-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

26

(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

27

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

28

(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

29

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).

30

The cellulose biosynthetic pathway and mechanism of cellulose synthesis

The steps involved in the pathway of cellulose biosynthesis were deduced using

tracer studies using 14

C-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?

31

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 180C 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

32

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.

33

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.

34

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

35

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

36

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.

37

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

38

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.

39

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.

40

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.

41

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

42

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 quantitatively fuels the reaction catalyzed by luciferase that converts luciferin to

light- producing oxyluciferin (Gould and Subramani 1988).

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

ATP sulphurylase

AMP-SO3H + PPi ATP + SO42-

Luciferase

Luciferin + ATP ADP + Oxyluciferin

43

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.

44

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).

45

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

46

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-

47

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

48

(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.

49

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

50

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

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.

51

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

52

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

53

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

54

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

55

(>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).

56

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

57

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

58

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

unknown function 4 2 2

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.

59

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60

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

61

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).

62

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.

63

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.

64

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

65

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

66

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

acsD gene

471 bp506 bp

1 2

acsD gene

471 bp506 bp

1 2

acsD gene

471 bp506 bp

1 21 2 3

pGEM

(cloning

vector)

acsD

(gene

insert)

1kb

500

bp

1 2 3

pGEM

(cloning

vector)

acsD

(gene

insert)

1 2 3

pGEM

(cloning

vector)

acsD

(gene

insert)

1kb

500

bp

1kb

500

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.

67

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

68

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.

69

a b

1 2 3 1 2 3 1 2 3 4

17 kDa

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

70

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

71

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,

72

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

73

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 31 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.

74

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

75

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.

76

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

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).

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.

Entry Description Mascot

Score

MW Peptides Coverage

(%)

ACSD_ACEX

Y

Cellulose

synthase operon

protein D

353 17432 R.DVDAEDLNAVPR.

Q (91)

R.WVTSQAGAFGDY

VVTR.D (149)

9.2

11.8

CM OM Cytop. Perip. TM

34

52

17

26

42

72

MW CM OM Cytop. Peri.

CM OM Cytop. Perip. TM

TM

77

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

78

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

79

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.

80

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).

81

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.

82

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-/Å

3 and ~0.33 e

-

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

83

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.

84

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)Rg2Q

2}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).

85

I(0) = 4 0 Dmax

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

86

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

87

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.

a

1 2 3 4

88

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

89

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

90

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.

91

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.

92

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

93

AcsD concentration: 8mg /ml

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.

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

94

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.

95

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.

96

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 20C. 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

97

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.

98

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

99

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.

100

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.

101

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

102

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

103

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.

104

a b

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).

Top view

Side view

105

c

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)

b

a

d

106

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).

107

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-

108

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

109

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.

110

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).

111

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).

112

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.

113

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

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

129 397

AcsAB

610

581 600 721 740

1550 AcsAB2

114

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 30C 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.

115

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 -20C.

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

b).

116

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.

5

06 bp

1000 bp

3000 bp

1636 bp

828 bp

2443 bp bp

500 bp

1 2 3 4

117

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 -70C. Western

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

proteins.

118

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 -70C 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

119

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

120

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.

121

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).

y = -0.0305x + 2.8835

R2 = 0.9928

0

0.5

1

1.5

2

2.5

3

0 10 20 30 40 50 60

Distance (mm)

Lo

g M

W

122

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,

123

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

124

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

125

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.

126

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.

A

A

A

Q

P S

Q

A

R

A

A

V

K

725 735 745 755 765 775 785 Amino acid residues

127

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

128

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.

129

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.

130

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.

131

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 30C. 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%

132

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

133

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

134

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.

135

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

136

Table 6.1b: Buffers for BN-PAGE*

Cathode

buffer

Dilute cathode

buffer

Anode

buffer Gel buffer

Tricine (mM) 50 50 - -

Imidazole (mM) 7.5 7.5 25 75

Coomassie blue G-250

(%) 0.02 0.002 - -

6-aminohexanoic

acid (M) - - - 1.5

*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

137

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.

138

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).

b

1 2 3 1 2

a

139

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

140

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.

141

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

b

d

a b

a

c

a

d

a

142

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.

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.

a

1 2 3

143

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.

Accession number Protein name PLGS score Coverage (%)

EFG85019.1

putative

phosphoribosylaminoimidazole

carboxylase catalytic subunit

6785.32 20.89

EFG83091.1 S-adenosyl L-homocysteine

hydrolase 5711.04 32.33

EFG82973.1 hypothetical protein GXY 15499 4678.76 33.70

EFG85152.1 putative glutamate synthase

NADPH large chain precursor 1918.27 46.30

EFG85151.1 putative oxidoreductase 1630.00 52.90

EFG85173.1 outer membrane protein OmpA 1345.23 47.19

EFG84008.1 UTP glucose 1 phosphate

uridylyltransferase 964.02 36.64

EFG85627.1 6 phosphogluconate

dehydrogenase like protein 813.62 41.14

EFG84192.1 phosphoglucomutase 580.85

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

EFG84948.1 import inner membrane translocase

subunit Tim44 170.62 10.13

144

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

145

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.

146

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,

147

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.

148

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

149

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

150

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

151

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.

152

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.

153

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VITA

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).