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University of Massachusetts Amherst University of Massachusetts Amherst ScholarWorks@UMass Amherst ScholarWorks@UMass Amherst Doctoral Dissertations Dissertations and Theses July 2016 Conversion of Cellulose to Ethanol by the Biofuels Microbe Conversion of Cellulose to Ethanol by the Biofuels Microbe Clostridium Phytofermentans: Quantification of Growth and Role Clostridium Phytofermentans: Quantification of Growth and Role of an Rnf-Complex in Energy Conservation of an Rnf-Complex in Energy Conservation Jesús G. Alvelo-Maurosa University of Massachusetts Amherst Follow this and additional works at: https://scholarworks.umass.edu/dissertations_2 Part of the Environmental Microbiology and Microbial Ecology Commons, and the Microbial Physiology Commons Recommended Citation Recommended Citation Alvelo-Maurosa, Jesús G., "Conversion of Cellulose to Ethanol by the Biofuels Microbe Clostridium Phytofermentans: Quantification of Growth and Role of an Rnf-Complex in Energy Conservation" (2016). Doctoral Dissertations. 642. https://scholarworks.umass.edu/dissertations_2/642 This Open Access Dissertation is brought to you for free and open access by the Dissertations and Theses at ScholarWorks@UMass Amherst. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of ScholarWorks@UMass Amherst. For more information, please contact [email protected].

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Page 1: Conversion of Cellulose to Ethanol by the Biofuels Microbe … · CONVERSION OF CELLULOSE TO ETHANOL BY THE BIOFUELS MICROBE . CLOSTRIDIUM PHYTOFERMENTANS: QUANTIFICATION OF GROWTH

University of Massachusetts Amherst University of Massachusetts Amherst

ScholarWorks@UMass Amherst ScholarWorks@UMass Amherst

Doctoral Dissertations Dissertations and Theses

July 2016

Conversion of Cellulose to Ethanol by the Biofuels Microbe Conversion of Cellulose to Ethanol by the Biofuels Microbe

Clostridium Phytofermentans: Quantification of Growth and Role Clostridium Phytofermentans: Quantification of Growth and Role

of an Rnf-Complex in Energy Conservation of an Rnf-Complex in Energy Conservation

Jesús G. Alvelo-Maurosa University of Massachusetts Amherst

Follow this and additional works at: https://scholarworks.umass.edu/dissertations_2

Part of the Environmental Microbiology and Microbial Ecology Commons, and the Microbial

Physiology Commons

Recommended Citation Recommended Citation Alvelo-Maurosa, Jesús G., "Conversion of Cellulose to Ethanol by the Biofuels Microbe Clostridium Phytofermentans: Quantification of Growth and Role of an Rnf-Complex in Energy Conservation" (2016). Doctoral Dissertations. 642. https://scholarworks.umass.edu/dissertations_2/642

This Open Access Dissertation is brought to you for free and open access by the Dissertations and Theses at ScholarWorks@UMass Amherst. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of ScholarWorks@UMass Amherst. For more information, please contact [email protected].

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CONVERSION OF CELLULOSE TO ETHANOL BY THE BIOFUELS MICROBE

CLOSTRIDIUM PHYTOFERMENTANS: QUANTIFICATION OF GROWTH AND

ROLE OF AN RNF-COMPLEX IN ENERGY CONSERVATION

A Dissertation Presented

by

JESÚS G. ALVELO MAUROSA

Submitted to the Graduate School of the

University of Massachusetts Amherst in partial fulfillment

of the requirements for the degree of

DOCTOR OF PHILOSOPHY

May 6, 2016

Department of Microbiology

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© Copyright by Jesús G. Alvelo Maurosa 2016

All Rights Reserved

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CONVERSION OF CELLULOSE TO ETHANOL BY THE BIOFUELS MICROBE

CLOSTRIDIUM PHYTOFERMENTANS: QUANTIFICATION OF GROWTH AND

ROLE OF AN RNF-COMPLEX IN ENERGY CONSERVATION

A Dissertation Presented

by

JESÚS G. ALVELO MAUROSA

Approved as to style and content by:

_______________________________________

Susan B. Leschine, Chair

_______________________________________

James F. Holden, Member

_______________________________________

Samuel P. Hazen, Member

_______________________________________

Yasu Morita, Member

____________________________________

Steven J. Sandler, Department Head

Department of Microbiology

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DEDICATION

To my parents, Jesús M. Alvelo Santiago and Alba I. Maurosa Colón. To my aunt Norma

Maurosa and to my sisters Cristina M. Alvelo and María M. Alvelo. Sin ellos soy nada.

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The youth has the right to defend their country with the weapons of knowledge.

Pedro Albizu Campos

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vi

ACKNOWLEDGMENTS

I would like to thank Dr. Susan B. Leschine for being a constant source of

inspiration, guidance, patience and support through my formation as a scientist throughout

my years. I thank Dr. Weimin Gu and Tom Warnick for being mentors and giving me great

advice during my first year in Dr. Leschine Lab. My gratitude extends for the sharing of

equipment and materials to Dr. Sarah Hensley, Begum Topcuoglu, Dr. Lucy Steward, Dr.

Pablo Visconti and Dr. Ana Maria Salicioni for their support and help.

I also want to thank our collaborators Dr. Jeffery Blanchard, Dr. Samuel P. Hazen

and Scott Lee for their support and help in this dissertation. I thank Dr. Roberto Orellana,

Dr. Jennifer Lin and Dr. Burcu Unal for their unconditional support, scientific discussions

and friendship. I also thank Dr. Wilmore Webley and Dr. Klaus Nusslein for their

examination on my qualifier exam and their advice. I would also like to thank to Dale

Callaham for his mentorship and guidance on my path to learning microscopy. Also, I want

to thank the administrative personnel in the Department of Microbiology Maryanne Wells,

Sue Ladford, Lorell Perreault and Ruthie Allies.

I am grateful for other students and colleges from the department and other

programs for their friendship and support, making things enjoyable, especially William

Rodriguez, Marcos Manganare, Jennifer Hayashi and Julia Puffal. Moreover, I thank Dr.

Susan Roberts and Shana Passonno for their financial support through the Institute of

Cellular Engineering and to Dr. Sandra Petersen and Dr. Heyda Martinez for their financial

support through IMSD and NEAGEP. I thank my committee members, Dr. James Holden,

Dr. Samuel Hazen and Dr. Yasu Morita for their additional guidance and support.

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ABSTRACT

CONVERSION OF CELLULOSE TO ETHANOL BY THE BIOFUELS MICROBE

CLOSTRIDIUM PHYTOFERMENTANS: QUANTIFICATION OF GROWTH AND

ROLE OF AN RNF-COMPLEX IN ENERGY CONSERVATION

MAY 6, 2016

JESÚS G. ALVELO MAUROSA,

B.S., UNIVERSIDAD DE PUERTO RICO, RECINTO DE RÍO PIDERAS

Ph.D., UNIVERSITY OF MASSACHUSETTS AMHERST

Directed by: Professor Susan B. Leschine

The anaerobic mesophilic bacterium Clostridium phytofermentans grows and

ferments multiple plant-based substrates into ethanol as the main product of fermentation.

The capacity of C. phytofermentans to convert plant biomass into ethanol, propanol, and

short-chain fatty acids is strongly attractive for industry. Specific physiological capabilities

of C. phytofermentans allow the microbe to generate high amounts of ethanol compared to

acetate. However, little is known about membrane energetics in C. phytofermentans, or its

role in energy conservation and production of high levels of ethanol during fermentation

of plant biomass substrates.

In the first research project presented in this dissertation, we examined C.

phytofermentans growth on three insoluble plant-based substrates: Whatman #1 filter

paper, wild-type Sorghum bicolor and the S. bicolor reduced lignin double mutant, bmr-

6/bmr-12. Cells were visualized and quantified employing a novel dual-fluorescent

staining protocol, combining Fluorescent Brightener 28, a blue fluorescent dye that stains

cellulose and chitin, and SYTO 9, a green fluorescent dye that stains nucleic acid. Our

results demonstrated this protocol allows the visualization and differentiation of C.

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viii

phytofermentans cells from plant debris using epifluorescence microscopy. Our results also

showed greater cell growth and substrate attachment on reduced lignin substrates.

Moreover, ethanol production increased over time while cell numbers decreased when

cultured with either sorghum substrate. These results suggested that only a portion of the

cell population remained active during growth, possibly due to sporulation and reduced

metabolic activity.

The second project presented in this dissertation involved the characterization of a

bacterial microcompartment (BMC) in C. phytofermentans. Transmission electron

microscopy (TEM) of thin sections of C. phytofermentans cells grown with fucose or

rhamnose as growth substrate showed the presence of polyhedral structures, similar in

appearance to known BMCs. Visualization of these intracellular structures was possible

due to various modifications of a TEM sample embedding protocol. Use of picric acid and

potassium ferricyanide resulted in enhanced contrast of BMC images while stabilizing cell

wall structure during sample preparation.

A third and final project described in this dissertation concerned the development

of a model for C. phytofermentans membrane energetics and energy conservation that

might provide an explanation for high levels of ethanol production by this microbe.

Genomic and biochemical analyses indicated the presence of a membrane-bound sodium-

and proton-translocating pyrophosphatase and three ATPases, including a Na+ V-type

ATPase, a Na+ F-type ATPase and a possible H+ ATPase that had not previously been

characterized. Furthermore, results showed that Rnf ferredoxin:NAD+ oxidoreductase

activity in inverted membrane vesicles was driven by a proton electrochemical gradient.

Supporting data showed that cell growth was inhibited by both sodium and proton

ionophores. Taken together, these results suggested that C. phytofermentans is capable of

conserving energy through ferredoxin oxidoreductase activity of the Rnf-complex,

resulting in increased levels of NADH for ethanol production.

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TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS ................................................................................................. vi

ABSTRACT ...................................................................................................................... vii

LIST OF FIGURES .......................................................................................................... xii

CHAPTER

1. INTRODUCTION ...........................................................................................................1

1.1 Central Objectives and Goals.............................................................................1

1.2 Organization of the Dissertation ........................................................................5

2. LITERATURE REVIEW AND RESEARCH APPROACH ..........................................6

2.1 Microbial Conversion of Plant Biomass into Biofuels ......................................6

2.2 Metabolism of Complex Carbohydrates by Human Gut Microbiota .................7

2.3 Clostridium phytofermentans: model organism .................................................8

2.3.1 Carbohydrate breakdown and ethanol production ..............................8

2.3.2 Cell enumeration in the presence of insoluble plant biomass

substrates ........................................................................................10

2.3.3 Bacterial Microcompartments ...........................................................15

2.3.4 Bioenergetics .....................................................................................18

2.4 Summary and Research Approach ...................................................................20

3. DIRECT IMAGE-BASED ENUMERATION OF CLOSTRIDIUM

PHYTOFERMENTANS CELLS GROWING ON INSOLUBLE PLANT

SUBSTRATES ......................................................................................................22

3.1 Abstract: ...........................................................................................................22

3.2 Introduction ......................................................................................................23

3.3 Materials and Methods .....................................................................................25

3.3.1 Bacterial strain and culture conditions. .............................................25

3.3.2 Dual-staining procedure. ...................................................................26

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x

3.3.3 Fluorescence microscopy and image processing. .............................26

3.3.4 Cell quantification. ............................................................................27

3.3.5 Determination of fermentation products. ..........................................27

3.3.6 Data analysis .....................................................................................27

3.4 Results ..............................................................................................................28

3.4.1 Visualization of C. phytofermentans cells growing on

insoluble substrates using the dual-staining procedure. .................28

3.4.2 Cell morphology and substrate interactions visualized by 3D

imaging. .........................................................................................29

3.4.3 Cell growth and ethanol production during fermentation of

insoluble substrates by C. phytofermentans. ..................................29

3.5 Discussion ........................................................................................................32

4. IMAGING INTRACELLULAR PROTEIN-BOUND

MICROCOMPARTMENTS IN BACTERIAL CELLS ........................................40

4.1 Abstract: ...........................................................................................................40

4.2 Introduction: .....................................................................................................41

4.3 Materials and Methods: ....................................................................................43

4.3.1 Bacterial strain and conditions: .........................................................43

4.3.2 Transmission Electron Microscopy: .................................................43

4.4 Results and Discussion: ...................................................................................44

5. MEMBRANE ENERGETICS AND ENERGY CONSERVATION IN THE

CELLULOLYTIC ETHANOLOGEN CLOSTRIDIUM

PHYTOFERMENTANS........................................................................................49

5.1 Abstract: ...........................................................................................................49

5.2 Introduction: .....................................................................................................50

5.3 Materials and Methods .....................................................................................53

5.3.1 Protein sequence alignment, comparison and structure

modeling ........................................................................................53

5.3.2 Bacterial strain and growth conditions .............................................53

5.3.3 Effects of ionophores on cell growth ................................................54

5.3.4 Preparation of inverted membrane vesicles ......................................54

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xi

5.3.5 ATP hydrolysis and ATP synthesis determinations..........................54

5.3.6 Rnf-associated ferredoxin oxidoreductase activity ...........................55

5.4 Results ..............................................................................................................55

5.4.1 Predicted structures of the Na+ binding sites of the F-type and

V-type ATPase c-rings ...................................................................55

5.4.2 Structural model of the pyrophosphatase Na+, H+ binding

sites ................................................................................................56

5.4.3 Effects of ionophores on C. phytofermentans growth ......................56

5.4.4 ATP synthesis in inverted membrane vesicles is driven by a

Na+ gradient ...................................................................................57

5.4.5 An ATP driven proton gradient supports NADH-dependent

ferredoxin reduction .......................................................................57

5.5 Discussion ........................................................................................................58

6. DISCUSSION AND FUTURE DIRECTIONS .............................................................73

REFERENCES ..................................................................................................................77

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xii

LIST OF FIGURES

Figure Page

1.1 CO2 emissions by sector in 2012. The transportation system combined with

generation of electricity and heat contributes 2/3 of the total global

CO2 emissions. (International Energy Agency, 2014. “CO2 Emissions

from Fuel Combustion”). ...................................................................................4

2.1 Cellulolytic bacterium from soil growing on a plate of Noble Agar medium

containing Congo red and supplemented with gelatin. (Henricks,

1995). ...............................................................................................................14

2.2 Diagrammatic representation of the metabolic functions associated with the

Pud (propanediol utilization) BMC and the conversion of 1,2-

propanediol (red box) into propionate or into propanol via the

oxidiation of NADH by PduQ. Propanol is short-chain fatty acid with

the potential to be an alternate liquid transportation fuel ................................17

3.1 Diagrammatic overview of dual-staining protocol. ....................................................35

3.2 Differential interference contrast (DIC) and epifluoresence micrographs of C.

phytofermentans cultured for 3 days with filter paper, wild-type S.

bicolor (WT Sorgum), or lignin double mutant S. bicolor (bmr-6/bmr-

12 Sorghum). Cultures were prepared for microscopy using the dual-

staining procedure. For each sample, one field is shown imaged by

DIC microscopy (A), and fluorescence microscopy at 385-400 nM

showing cellulose fibers stained with Calcofluor White (B) and at 480

nM showing Syto 9 stained cells (C). The epifluorescence images in

rows B and C are merged in the row D images. Stained culture

preparations were diluted with sterile basal culture medium prior to

slide preparation: the filter paper culture was dilute 1:3 and sorghum

cultures were diluted 1:1. .................................................................................36

3.3 Two-dimentional rendering of a 3D image of C. phytofermentans cultured for

five days in filter paper medium. The culture was prepared for

microscopy using the dual-staining procedure. Multiple focal plane

images at a dept of 1 µm were obtained and used to generate a 3D

images. .............................................................................................................37

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xiii

3.4 Cell growth (A) and ethanol production (B, C) by C. phytofermentans, and

cell numbers (D), cell attachment to substrate (E) and ethanol

concentration (F) after three days of growth with filter paper (),

wild-type S. bicolor (), or lignin double mutant S. bicolor bmr-

6/bmr-12 () as growth substrate. The concentration of ethanol in

cultures is expressed as mM (B, F) and the amount of ethanol

produced per cell is expressed as nmoles per cell (C). Error bars

represent standard deviation of the mean (n=3). Statistically signficant

differences are represented with asterisks; one asterisk for P ≤ 0.05,

two asterisks for P ≤ 0.01, and three asterisks for P ≤ 0.001. ..........................38

3.5 Correlation of ethanol production with cell number during growth. Ethanol

production is given as a function of the cell number in cultures with

filter paper (A), wild-type S. bicolor (B) and lignin double mutant S.

bicolor bmr-6/bmr-12 (C) as growth substrate. ...............................................39

4.1 Transmission electron micrograph of C. phytofermentans cells from cultures

grown with glucose as growth substrate. Polyhedral structures were

not observed in cells from glucose-grown cultures. ........................................46

4.2 Transmission electron micrograph of C. phytofermentans cells from cultures

grown in fucose as growth substrate. Some cells contained polyhedral

structures (arrow), presumably bacterial microcompartments. ........................47

4.3 Transmission electron micrograph of C. phytofermentans cells from cultures

grown in rhamnose as growth substrate. Multiple polyhedral

structures, presumably bacterial microcompartments, were observed in

thin sections of C. phytofermentans grown in rhamnose culture

medium. ...........................................................................................................48

5.1 Model of the C. phytofermentans Na+ F-type ATPase c-ring. An 11 c-ring

homo-mer was developed including the location of amino acids

involved in Na+ binding. The model was developed using SWISS

MODEL and 3D rendered using VMD 1.9.1. ..................................................62

5.2 Alignment of F-type ATPase amino acid sequences. Conserved amino acids

involved in Na+ are marked with an asterisk. The C. phytofermentans

protein contains the same conserved amino acids found in other F-type

ATPases, V63, E65 and Q32 with the exception of T66. The sequence

alignment was performed with microbes know to possess Na+ F-type

ATPases: I. tartaricus, F. nucleatum and C. paradoxum. Sequences

were aligned using PROSMAL3D and visualized by JalView. ......................63

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xiv

5.3 A model of the C. phytofermentans Na+ V-type ATPase c-ring protein. A 10-

c-ring homo-mer was developed containing the location of amino

acids involved in Na+ binding. The model was developed using

SWISS MODEL and 3D rendered using VMD 1.9.1. .....................................64

5.4 Alignment of V-type ATPase amino acid sequences. Conserved amino acids

involved in binding Na+ are marked with an asterisk. C.

phytofermentans contains the same amino acids as known microbes

with Na+ V-type ATPases: E. hirae and E. faecalis. C. asparagiforme

and C. bolteae proteins have the greatest similarity to that of C.

phytofermentans. Sequences were aligned using PROSMAL3D and

visualized by JalView. .....................................................................................65

5.5 Model of the pyrophosphatase protein of C. phytofermentans. The model

shows conserved amino acids T87 (magenta spheres), F91 (purple

spheres), M117 (yellow spheres) and D143 (red spheres) involved in

binding H+ and Na+. The model was developed using SWISS

MODEL and 3D rendered using VMD 1.9.1. ..................................................66

5.6 Alignment of the C. phytofermentans pyrophosphatase amino acid sequence

with other pyrophosphatase sequences. ...........................................................68

5.7 Effect of ionophores on growth of C. phytofermentans in defined medium M6

with glucose as substrate. Cell growth was inhibited by both ETH

2120 and TCS when added at the time of inoculation or at mid-

exponential phase of growth. ...........................................................................69

5.8 Effect of ionophores on ATP synthesis in inverted membrane vesicles. ATP

synthesis was initiated by the addition of ADP to reaction mixtures.

ATP was measured by luminescence using a luciferin-luciferase assay

kit. ATP synthesis was significantly reduced in reactions containing

the Na+ ionophore ETH 2120. .........................................................................70

5.9 NADH-dependent ferredoxin reduction assay of Rnf oxidoreductase activity

in inverted membrane vesicles. Reaction mixtures contained

ferredoxin, NADH, and inverted membrane vesicles in buffered

solution. Reactions were initiated by addition of ATP at 60 seconds. In

some reactions, ETH 2120 or TCS was added at a concentration of 10

µM. Ferredoxin reduction was measured at 430 nM. ......................................71

5.10 Metabolic model illustrating the role of membrane energetics in energy

conservation in C. phytofermentans. The Rnf complex utilizes

electrons from reduced ferredoxin to reduce NAD+ to NADH while

translocating H+ across the membrane generating an electrochemical

gradient used by a H+ V-type ATPase. A pyrophosphatase (PPase)

augments the H+ and Na+ electrochemical gradients driving ATP

synthesis by the Na+ V-type and Na+ F-type ATPases, ...................................72

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1

CHAPTER 1

INTRODUCTION

1.1 Central Objectives and Goals

Global temperature has been on the rise due to increased concentrations of atmospheric

CO2 (Allen et al. 2009). In order to limit the increase in global temperature to 2o C, 1,200

gigatons (Gt) of CO2 or less needs to be produced from 2015 to 2100 (Fuss et al., 2014).

Emissions from use of fossil fuels and cement production have contributed to increased

concentrations of CO2 with nearly 40 Gt produced in 2015. Strong public policy and human

will are essential if we do not want to go above the 2o C benchmark. Greater effort will be

required the longer the wait.

The transportation sector is responsible for 23% of total world CO2 emissions (Fig.

1.1). Bioethanol is an important renewable energy source that could substitute for

petroleum-based fuels in land transportation and reduce CO2 emissions. First generation

bioethanol from edible food crops, such as corn and sugarcane, has dominated the market

but land use management and increased food prices have created controversy (Graham-

Rowe, 2011). Second generation bioethanol derived from lignocellulosic biomass has the

potential for sustainable production and a decreased impact on global food supply

(Somerville et al., 2010). Nonetheless, recalcitrance of lignocellulosic biomaterial, due

primarily to its lignin content, presents a challenge for bioethanol production inasmuch as

the pretreatment of woody biomass requires hydrolytic enzymes, acid, and heat, which

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2

makes the process complex and costly (Lynd 2005, Ragauskas 2006, Chen 2007 and

Sanderson 2011).

An alternative approach for the conversion of lignocellulosic biomass to bioethanol is

consolidated bioprocessing (CBP), where a microbe capable of producing lignocellulose-

decomposing enzymes also ferments the resulting carbohydrates to ethanol as a

fermentation product (Lynd 2005). Clostridium phytofermentans, which has emerged as a

model CBP microorganism (Jin et al., 2011, Tolonen et al., 2011, Zuroff et al., 2013, Yee

et al., 2014 and Petit et al., 2015), directly converts all major components of plant biomass

(cellulose, hemicellulose, pectin, starch) into fermentation products, primarily ethanol. In

order to better appreciate the potential of C. phytofermentans as a CBP microbe, strategies

to examine the microbe interacting with plant biomass are required. Additionally, a deeper

understanding of the physiology of C. phytofermentans is needed to evaluate the potential

of this microbe for biofuels production; e.g., the role of intracellular protein-bound

bacterial microcompartments (BMCs) in fermentation product formation and the function

of membrane electrochemical gradients in energy conservation and ethanol production.

The goals of this dissertation are to: 1) develop a method to enable visualization and

quantification of microbial cells interacting with plant biomass, 2) generate a method that

allows consistent visualization of BMCs to facilitate studies of the function and distribution

of BMCs in cells and 3) using structural bioinformatics and biochemical assays, develop a

testable model for energy conservation in C. phytofermentans involving membrane-bound

ATPases, an Rnf complex and ion gradients across the membrane.

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3

This dissertation project addresses the following questions:

i- How does the lignin content of plant biomass feedstocks affect the growth

of C. phytofermentans?

ii- What preparation procedures are required to visualize bacterial

microcompartments using transmission electron microscopy?

iii- What mechanisms are utilized by C. phytofermentans to conserve energy,

and how is energy conservation related to fermentation product formation?

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Figure 1.1 Figure 5.5 Model of the pyrophosphatase protein of C. phytofermentans. The

model shows conserved amino acids T87 (magenta spheres), F91 (purple spheres), M117

(yellow spheres) and D143 (red spheres) involved in binding H+ and Na+. The model

was developed using SWISS MODEL and 3D rendered using VMD 1.9.1.

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1.2 Organization of the Dissertation

The research questions described above are addressed as separate chapters in this

dissertation. Chapter 3 has appeared as a first-author publication in Applied and

Environmental Microbiology; some of the research presented in Chapter 4 has been

published in PLoS ONE as part of a research collaboration; the research described in

Chapter 5 has been prepared for publication in Frontiers in Microbiology, also as a first-

author publication. Chapter 1 describes the objectives of the research. A literature review

with background information and the research approach are given in Chapter 2. In Chapter

3 a microscopy method to visualize C. phytofermentans cells in the presence of plant

biomass is presented along with its application in investigations of cell growth kinetics on

insoluble plant biomass substrates which contributed to a co-authored publication (Lee et

al., 2012) and it was recently published on another peer reviewed journal (Alvelo-Maurosa

et al., 2015). Chapter 4 describes transmission electron microscopy methodology for

visualizing bacterial microcompartments in C. phytofermentans cells grown with fucose or

rhamnose as substrate. Chapter 4 results were a contribution to a peered review publication

(Petit et al., 2013) Chapter 5 describes research using structural bioinformatics and

biochemical assays to evaluate ion gradients involved in membrane energetics, which are

tied to the activities of ATPases and an Rnf complex. A model for membrane energetics in

C. phytofermentans also is presented in Chapter 5 and will be submitted to a peer review

journal. Chapter 6, the final chapter of this dissertation, includes a discussion of the

significance of the research and concluding remarks.

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CHAPTER 2

LITERATURE REVIEW AND RESEARCH APPROACH

2.1 Microbial Conversion of Plant Biomass into Biofuels

Lignocellulosic plant biomass is composed of proteins, lignin and polysaccharides

(Leschine, 2005.). The prevalent polysaccharide in plant cell walls is cellulose, a complex

chain-like system composed of over 15,000 beta-(1,4)-linked glucose molecules (Brett CT,

2000). Cellulose enforces strength in the cell wall to maintain turgor pressure.

Hemicelluloses (glucomannans and xylans) are composed of multiple monomers including

D-xylose, L-arabinose, D-mannose, D-glucose, D-galactose and D-glucuronic acid

(Leschine, 1995, Leschine, 2005). Three monolignols compose lignin: coniferyl, synapyl,

and p-coumaryl alcohols, which form guaiacyl, syringyl, phenylpropanoids and p-

hydroxyphenyl units of lignin (Bonawitz and Chapple, 2010). Lignin is cross linked to

cellulose and other polymers, such as hemicellulose, which provides hydrophobicity and

strength to the plant cell wall, and also reduces accessibility of polysaccharides to

enzymatic degradation and penetration by pathogens (Grabber, 2005 and Bonawitz, 2010).

Microbial interactions with plant biomass are central to technologies for the

conversion of lignocellulosic feedstocks into useful products. For example, ethanol is an

important renewable energy source that may replace petroleum-based transportation fuels

and reduce CO2 emissions (Sticklen 2008). Development of more sustainable processes for

second generation biofuels derived from lignocellulosic biomass is limited by the

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recalcitrance of the substrate, primarily due to its lignin content, and often requires

expensive pretreatment and use of hydrolytic enzymes (Sanderson 2011, Ragauskas et al.,

2006 and Chen et al., 2007). An alternative strategy to convert lignocellulosic feedstocks

into biofuels is consolidated bioprocessing (CBP), where microbes capable of producing a

complex set of glycoside hydrolases decompose plant biomass and ferment the products in

a single step, for a more cost-effective process with savings in capital and operational costs

(Lyng et al., 2005).

The use of various wildtype and genetically engineered microbes as CBP catalysts

has been shown to increase bioethanol yield (Olson et al., 2012, Peralta-Yahya, 2010 and

Lynd et al., 2008). Furthermore, reducing the lignin content of lignocellulosic biomass

allows greater accessibility of microbes to plant carbohydrates, such as hemicellulose and

cellulose, and higher yields of bioethanol (Chen & Dixon 2007, and Himmel, 2007).

Studies have shown that microbes are capable of improved CBP performance and greater

saccharification of plant biomass with a reduced lignin content, resulting in higher yields

of bioethanol and other fermentation products (Lee et al., 2012 and Yee 2012).

2.2 Metabolism of Complex Carbohydrates by Human Gut Microbiota

The human intestine is home to the gut microbiome, composed of approximately

3.9 x 1013 bacteria (Sender et al., 2016), with a constantly changing diversity due to the

host diet (Bäckhead et al., 2005). The human gut microbiome can be viewed as a large

reactor where microbial breakdown of complex polysaccharides occurs. These compounds,

referred to as “dietary fiber,” are derived from major components of the human diet such

as fruits, cereals and vegetables. Humans and most other animals lack the enzymatic

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capacity required to digest cellulose and many other components of plant cell walls, and

instead they rely on plant biomass-decomposing microbial communities to provide

nutrition from plant fiber. Dietary fiber serves as a source of growth substrates for intestinal

microbes that ferment these components of plant biomass to short chain fatty acids, which

are tied to colonic and systemic health (Wong et al., 2006, Flint et al., 2012, Sticklen 2008,

Sanderson 2011 and Ragauskas et al., 2006).

Plant polysaccharides such as pectin, cellulose, hemicellulose and starches are

degraded by carbohydrate-active enzymes (CAZymes,) produced by the gut microbiota (El

Kaoutari et al. 2013). CAZymes generated by the microbiome cleave oligosaccharides and

polysaccharides into monosaccharides that are then fermented. These anaerobic digestions

produce short-chain fatty acids such as butyrate, acetate and propionate that contribute to

important physiological processes (Cantarel, Lombard and Henrissat 2012). For example,

butyrate is an important energy source for colonocytes and promotes differentiation of

colonic regulatory T cells that have multiple functions including the capacity to fight off

cancer cells (Furusawa et al., 2013). It has also been found that patients with type-2 diabetes

have a decrease in butyrate-producing bacteria due to dysbiosis, which may lead to an

increased probability of colon-rectal cancer (Qin et al., 2012).

2.3 Clostridium phytofermentans: model organism

2.3.1 Carbohydrate breakdown and ethanol production

Clostridium phytofermentans is a cellulolytic anaerobic bacterium isolated from

soil near a water reservoir in Massachusetts (Warnick, Methé, and Leschine 2002). The

microbe is capable of hydrolyzing and fermenting plant components such as cellulose and

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hemicellulose and producing ethanol as its main product. The C. phytofermentans genome

encodes 161 CAZymes, with 116 glycoside hydrolases, including hemicellulases,

pectinases, and amylases among others (Tolonen et al., 2011 and Petit et al., 2015). With

its versatility for plant saccharification and high level of ethanol production, C.

phytofermentans differentiates itself form other characterized microbes as an ideal

candidate to better understand the mechanism of decomposition of different types of plant

biomass substrates.

It has been determined that C. phytofermentans produces different amounts of

ethanol depending on the type of plant biomass substrate, pre-treatment method, particle

size, pH and temperature (Jin et al., 2011 and Lee et al., 2012). The lignin content of plant

biomass plays a significant role in ethanol production. Lee et al. 2012 reported that C.

phytofermentans produces higher levels of ethanol when grown with three different types

of brown midrib sorghum as substrate compared to growth with wildtype sorghum. Brown

midrib sorghum contains less lignin than wildtype sorghum, suggesting that a reduction in

the lignin content of plant biomass allows more accessibility to cellulose and

hemicellulose, making it more digestible. However, prior to research described in Chapter

3 of this dissertation, cell growth kinetics on plant biomass substrates, and the impacts of

microbe-biomass interactions on biofuel production were unknown due to a lack of

methods for visualizing and quantifying microbes in the presence of plant biomass

substrates.

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2.3.2 Cell enumeration in the presence of insoluble plant biomass substrates

The enumeration of cellulolytic bacteria was first described by Hungate (Hungate

RE, 1950) who reported the quantification of cellulolytic colonies on plates of cellulose

agar media. Some bacterial strains grew in liquid media with cellulosic substrates but did

not grow on plates of agar media; thus, quantification of growth for some strains was not

possible. Various other methods yielded inconsistent results, mostly due to the use of

different media components rather than the presence of cellulose itself (Bryant, 1959). By

1960, Kistner had developed a water-cooled agar solidification apparatus to generate a

uniform film of agar along the walls of rubber-stoppered roll bottles (Kistner, 1960). The

surface of the agar was scanned using an “Apparatenfabriek van Dooren” counting device

and cellulolytic colonies were quantified with consistency, Kistner being the first

researcher to do so. Another research group developed a fluorescent antibody technique,

the first researchers to do so with a cellulolytic microbe, to enumerate cellulolytic cocci

microbes from high dilution samples from sheep rumen fluid using fluorescence

microscopy (Jarvis, 1967). Numbers for cellulolytic cocci in sheep rumen fluid were

consistent; however, the researchers did not use cellulosic substrates in their media when

quantify cells. Still, the work of Jarvis et al. to develop the first fluorescent technique for

visualizing specific microbes in mixed populations was remarkable. By the end of 1960’s

two methods for quantifying cellulolytic bacteria were in use: a ‘direct’ method which

relied on the growth of cellulolytic bacteria and production of zones of clearing in

cellulose-containing agar media, and an ‘indirect’ method in which an agar medium

provided growth conditions for multiple species of microbes, and then, colonies were hand-

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picked and transferred to specific media. When compared, these methods provided very

similar numbers (Gylswyk, 1970).

For over a decade, few advances in methods to enumerate cellulolytic bacteria

occurred. Most attempts involved staining a cellulosic substrate in order to detect cellulose

breakdown, but none of these techniques focused on direct cell counts (Smith, 1977 and

Mahasmeh, 1980). A method developed by Teather (Teather, 1982) involved staining β-

D-glucans in plates of agar media with Congo red. This technique allowed detection of β-

D-glucan hydrolysis on plates of agar media, but it was often still complemented by

measurements using the roll bottle technique. In 1994, an enzyme-linked immunosorbent

assay (ELISA) for detecting Clostridium aldrichii growing on lignocellulosic substrates

was reported (Brigmon, 1994). This assay detected cellulolytic microbes in both pure and

mixed cultures at cell number of 105 cells ml-1 in a period of 36 hours. In 1995, Hendicks

et al., developed a new solid medium that contained Noble agar, gelatin and Congo red in

order to visualize and quantify cellulolytic microbes on plates of a cellulose agar medium

(Hendricks 1995). This method enabled the identification of cellulose-utilizing bacteria on

plates of agar media by the appearance of zones of clearing around colonies. (Fig 2.1).

In 2004, Zhang and Lynd reported measurements of the growth of Clostridium

thermocellum in batch cultures with Avicel as growth substrate (Zhang et al., 2004). These

researchers employed an indirect ELISA method for measurement of concentrations of

cellulase protein and distinct C. thermocellum cell mass determinations. Cell mass and

cellulase protein concentration increased exponentially through 20 hours of growth.

Interestingly, Zhang and Lynd also reported that cell mass concentration was the highest

after 22 hours of growth, and then declined. In 2006, this group of researchers measured

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cell growth and specific cellulose hydrolysis rates in both batch and continuous cultures

and concluded that cellulose hydrolysis is higher in growing cultures of C. thermocellum

as compared to purified cellulase preparations, demonstrating the importance of “enzyme-

microbe synergy.” (Lu, 2006).

Various researchers have used qPCR to estimate the number of cellulolytic bacteria

in ruminants, soils and lakes (Picard et al., 1992., Koike and Kobayashi 2001, Mosoni et

al., 2007., McDonald et al., 2009and de Menezes et al., 2012). Cell numbers have been

estimated by measuring the abundance of a gene, such as the 16S rRNA gene, rrs (Mosoni,

2007; Koike, 2001). In some instances, traditional techniques, such as growth on solid

media in roll tubes or most-probable number (MPN) determinations were combined with

qPCR results to estimate cell numbers. Yet none of these studies focused on the growth of

individual cellulolytic bacteria in media containing cellulose or another lignocellulosic

substrate. In 2009, researchers from Italy quantified bacterial biomass in wastewater sludge

by using fluorescent dyes and flow cytometry (Foladori et al., 2009). By using dyes, such

as Dead/Live Staining Kit, SYBR-Green I, and Propidium Iodide, Foladori was able to

visualize microbes in both raw sludge samples and in diluted samples. The ratio of viable

to non-viable bacteria was determined using flow cytometry. Cell aggregates were counted

as one cell, and total numbers were estimated, but final numbers were calculated using the

volume of each particle. The report by Foladori is unique in employing a technique to

directly count cells in a medium containing cellulose fibers.

In an effort to solve the problem of quantifying cellulolytic bacteria in media

containing insoluble plant biomass substrates such as cellulose, we employed a dual-

fluorescent staining technique along with epifluorescence microscopy that allowed us to

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differentiate and enumerate C. phytofermentans cells (Alvelo-Maurosa et al., 2015), as

discussed in Chapter 3 of this dissertation. The protocol involved staining bacteria using

SYBR-Green from the Live/Dead Cell Staining Kit and the use of Fluorescent Brightener

28 to stain cellulose. By using a series of dilution ratios and vortex speeds, we quantified

growth in media containing cellulose (ball-milled Whatman #1 filter paper), wildtype

sorghum and a double-lignin mutant bmr-6/bmr-12 sorghum. This method facilitated

measurements of cell attachment to substrate and cell-ethanol ratios. Also, our research

provided 3D images of C. phytofermentans cells growing on cellulosic substrates. To our

knowledge, this is the first extensive study involving direct cell counts of a microbe

growing on insoluble lignocellulosic substrates.

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Figure 2.1 Cellulolytic bacterium from soil growing on a plate of Noble Agar medium

containing Congo red and supplemented with gelatin. (Henricks, 1995).

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2.3.3 Bacterial Microcompartments

Bacterial microcompartments (BMCs) are protein polyhedral bodies that sequester

diverse metabolic pathways (Cameron et al., 2013). The structure of BMCs is similar to

those of viral capsids, where hexameric and pentameric protein clusters are combined to

form a protein shell. Enzymes of the BMCs are encapsulated by shell proteins possibly to

prevent exposure to detrimental environmental factors such as oxygen (Fan et al, 2012).

However, BMC shells do have pores, 4-7 Å in diameter, which allow passage of

metabolites (Kerfeld and Erbilgin 2015). Various types of BMCs with different functions

and generating various metabolites are known; e.g., 1,2-propanediol (PDU), ethanolamine

(EUT), ethanol (ETU), choline (CUT), glycyl radical propanediol (GRP), planctomycete

and verrucomicrobia metabolosome (PVM) and CO2 fixation (carboxysome) (Kerfeld and

Erbilgin 2015).

BMCs have been found in 23 bacterial phyla and their functions vary in different

environments (Axen et al., 2014). For example, in humans, some members of the gut

microbiome catalyze trimethylamine produced from choline via a glycyl radical enzyme

(GRE)-associated microcompartment (Craciun et al., 2012). PDU and EUT

metabolosomes are involved in the virulence of some pathogens such as Escherichia coli

O157:H7 and Salmonella enterica (Harvey et al., 2011 and Kendall et al, 2012). Members

the Planctomycetes phylum are capable of breaking down fucose, rhamnose and fucoidan

in aerobic conditions (Erbilgin et al., 2014).

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Genomic, transcriptomic, biochemical and microscopic evidence has indicated the

presence of a PDU BMC in C. phytofermentans, which is involved in the conversion of

1,2-propanediol to propionate and propanol during growth on the deoxyhexose sugars (also

known as methyl-pentoses), fucose or rhamnose (Petit et al., 2015) (Fig. 2.2). The presence

in C. phytofermentans of a BMC for the production of 1, 2-propanediol and propionate

from deoxyhexose sugars, indicates the potential of this microbe for production of multiple

biofuels from a broad range of cellulosic substrates.

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Figure 2.2 Diagrammatic representation of the metabolic functions associated with the Pud

(propanediol utilization) BMC and the conversion of 1, 2-propanediol (red box) into

propionate or into propanol via the oxidiation of NADH by PduQ. Propanol is short-chain

fatty acid with the potential to be an alternate liquid transportation fuel

(Adapted from Yeates, Crowley, and Tanaka, 2010)

http://people.mbi.ucla.edu/yeates/microcompartments.html

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2.3.4 Bioenergetics

A characteristic of C. phytofermentans, which is central to its industrial relevance,

is its propensity for ethanol production; ethanol is the main product of fermentation of

multiple plant biomass substrates (Petit et al. 2015 and Warnick et al., 2002). NADH

reduces Acetyl-CoA to form ethanol, generating NAD+, which is required in the glycolytic

pathway. Formation of acetate, another fermentation product, from Acetyl-CoA yields

ATP by substrate level phosphorylation. Since yields of ethanol are higher than acetate we

hypothesized that the microbe was capable of conserving energy and forming ATP in other

ways. Microarray analyses revealed numerous highly expressed genes involved in

membrane energetics and energy conservation (Petit et al., 2015). For example, the genome

of C. phytofermentans encodes three trans-membrane ATPases. A Na+ F-type

(Cphy_3741) and a Na+ V-type (Cphy_3076) ATPase have been identified by sequence

alignment and enzyme assays (as discussed in Chapter 5 of this dissertation). Additionally,

a third ATPase (Cphy_3445), with unknown ion specificity, has also been identified

through genome and microarray analyses.

ATPases are membrane-bound ion pumping channels that may contribute to the

formation of trans-membrane ion gradients or the formation of adenosine triphosphate

(ATP) from adenosine diphosphate (ADP) and inorganic phosphate (Pi). Membrane-bound

ATPases are essential for cellular energetics and are found in all three domains of life

(Mulkidjanian et al., 2007). F-type ATPases can be found in bacteria, and in the

chloroplasts and mitochondria of eukaryotic cells. Structural features of F-type ATPases,

such as the c-ring, a unit involved in ion binding, different from those of the V-type

ATPases. ATPases may be capable of pumping H+, Na+, K+ and/or Ca2+, which results in

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the generation of an electrochemical gradient (Kühlbrandt, 2004 and Toyoshima, 2004).

ATPases are versatile proteins required for the energy conservation in bacteria and may

play an important role in fermentative anaerobes.

The genome of C. phytofermentans also encodes an Rnf complex (Cphy_0211-

0216) with an integral ferredoxin oxidoreductase. The Rnf complex utilizes electrons from

ferredoxin and reduces NAD+ to NADH while translocating either Na+ or H+. The Rnf

complex was first identified as a protein complex involved in the nitrogen fixation process

of Rhodobater capsulatus (Schmehl et al., 1993). In the acetogen Acetobacterium woodii,

the Rnf complex catalyzes the reduction of NAD+ to NADH using ferredoxin as electron

donor while extruding a sodium ion (Müller et al., 2008). In contrast, the Rnf-complex of

Clostridium ljungdahlii has been identified as proton-translocating and essential for

autotrophic growth. (Tremblay et al., 2012). An Rnf complex has been linked to energy

conservation in A. woodii during caffeate respiration (Hess et al., 2013). Genomic analyses

have revealed the presence of an Rnf-complex in other Clostridiales (Clostridium,

Eubacterium), in Thermoanaerobacter, Thermanaerovibrio and in some Achaeans (Biegel

et al., 2011 and Schlegel et al., 2012).

In an analysis of the genome and transcriptome of C. phytofermentans, Petit et al.

(2015) proposed a membrane energetics model that includes a sodium-pumping Rnf-

complex and three sodium ATPases, but this model was not tested experimentally. We

hypothesized that the Rnf complex of C. phytofermentans contributes to ethanol formation

by producing NADH and that it also contributes the formation of a Na+ or H+ ion gradient,

which drives trans-membrane ATP synthase(s). Research related to this hypothesis and

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characterization of the C. phytofermentans Rnf complex and membrane energetics are

presented in Chapter 5 of this dissertation.

The genome of C. phytofermentans also encodes a membrane-bound

pyrophosphatase (PPase) (Cphy_1812) that couple’s energy of pyrophosphate (PPi)

hydrolysis to the translocation of Na+ or H+ across the cell membrane (Luoto et al., 2013).

PPi is an important energy source for the phosphorylation of fructose-6-phosphate and

conversion of phosphoenolpyruvate to pyruvate (Petit et al., 2015). C. phytofermentans

also possess glycolytic enzymes such an adenylate kinase, which exclusively use PPi and

can increase ATP yield of glycolysis (Mertens et al., 1993). We hypothesize that the

membrane-bound pyrophosphatase in C. phytofermentans may be involved in generating a

Na+/H+ trans-membrane gradient and also may use the Na+/H+ gradient to produce PPi for

glycolysis while reducing the need for acetate-dependent ATP synthesis.

2.4 Summary and Research Approach

This dissertation examines the physiology of C. phytofermentans in plant biomass

breakdown, the formation of a bacterial microcompartment and the role of membrane

energetics in ethanol production. Although it is known that ethanol production by C.

phytofermentans varies with growth substrate, measurements of cell growth and

observations of cell-substrate interactions in plant biomass substrates have not been

possible due to the inability to differentiate cells from insoluble plant biomass substrates.

In Chapter 3 of this dissertation a dual-fluorescent stain method to visualize C.

phytofermentans cells in the presence of plant biomass is presented along with the

application of this method in investigations of cell growth kinetics on insoluble plant

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biomass substrates. Using this staining method and fluorescence microscopy we present

results of analyses of cell growth dynamics with three different plant biomass substrates:

Whatman #1 filter paper, wildtype Sorghum bicolor and a reduced lignin mutant bmr-

6/bmr-12 Sorghum bicolor. Cell growth kinetics are correlated with ethanol production

over a period of 5 days.

Chapter 4 describes transmission electron microscopy methodology for visualizing

bacterial microcompartments (PDU metabolosomes) in C. phytofermentans cells grown

with fucose or rhamnose as substrate. These methods were used to confirm the presence of

BMCs in C. phytofermentans (Petit et al., 2013).

In Chapter 5 of this dissertation, a membrane energetics model for C.

phytofermentans is presented. Genomic analyses, structural bioinformatics and

biochemical assays were used to identify three membrane-bound ATPases, including two

Na+-translocating ATPases, an F-type and a V-type, and a third unknown type of ATPase,

possibly H+-translocating. A dual Na+/H+ pyrophosphatase was identified by sequence

alignment and structural bioinformatics. Finally, a possible H+-pumping Rnf complex with

ferredoxin oxidoreductase activity was identified. We propose that C. phytofermentans

utilizes these membrane-bound protein complexes to generate electrochemical gradients,

produce ATP (possibly reducing acetate production), generate PPi for glycolysis and

NADH for ethanol formation.

Chapter 6, the final chapter of this dissertation, includes a discussion of the

significance of the research and concluding remarks.

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CHAPTER 3

DIRECT IMAGE-BASED ENUMERATION OF CLOSTRIDIUM

PHYTOFERMENTANS CELLS GROWING ON INSOLUBLE PLANT

SUBSTRATES

3.1 Abstract

A dual fluorescent dye protocol to visualize and quantify Clostridium

phytofermentans ISDg (ATCC 700394) cells growing on insoluble cellulosic substrates

was developed by combining Calcofluor White staining of growth substrate with cell

staining using the nucleic acid dye, Syto 9. Cell growth, cell-substrate attachment, and

fermentation product formation were investigated in cultures containing either Whatman

#1 filter paper, wild-type Sorghum bicolor, or an S. bicolor reduced lignin double mutant,

bmr-6/bmr-12, as growth substrate. After three days of growth, cell numbers in filter paper

cultures were 6.0- and 2.2-fold higher than cell numbers in cultures with wild-type sorghum

and lignin double mutant sorghum, respectively. However, cells produced more ethanol

per cell when grown with either sorghum substrate than with filter paper as substrate.

Ethanol yields of cultures were significantly higher with lignin double mutant than with

wild-type sorghum or filter paper as substrate. Moreover, ethanol production correlated

with cell attachment in sorghum cultures: 90% of cells were directly attached to the lignin

double mutant sorghum substrate, while only 76% of cells were attached to wild-type

sorghum substrate. With filter paper as growth substrate, ethanol production was correlated

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with cell number; however, with either wild-type or mutant sorghum, ethanol production

did not correlate with cell number, suggesting that only a portion of the microbial cell

population was active during growth on sorghum. The dual-staining procedure described

here may be used to visualize and enumerate cells directly on insoluble cellulosic

substrates, enabling in-depth studies of interactions of microbes with plant biomass.

3.2 Introduction

Microbial decomposition of plant biomass is central to nutrient cycling in numerous

varied environments and this process plays a key role in the cycling of carbon on the planet.

In anoxic environments rich in decaying plant material, diverse communities of interacting

microbes are responsible for the degradation of cellulose, complex polysaccharides, and

other abundantly produced plant cell wall components. Given that most of these substrates

are insoluble, microbial decomposition of plant biomass occurs exocellularly, and

breakdown products may become available to other community members forming a basis

for multifarious interactions that occur in these environments (Leschine 1995).

Anoxic decomposition of plant biomass is also tied to health and nutrition in animals by

way of gastrointestinal tract microbial communities. Most animals lack the enzymatic

capacity required to digest cellulose and many other components of plant cell walls, and

instead rely on plant biomass-decomposing microbial communities to provide nutrition

from plant fiber. Ruminants, a group of herbivorous mammals, degrade forage in a

specialized foregut organ, the rumen. The microbial community housed in the rumen plays

an essential role in development and health of the ruminant by decomposing and

fermenting plant materials, and forming products, such as volatile fatty acids, that serve as

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essential nutrients (Jami, Israel et al. 2013). In humans, complex plant carbohydrates, also

known as dietary fiber, serve as substrates for intestinal microbes that ferment these

components of plant biomass to short chain fatty acids, which are tied to colonic and

systemic health (McDougall GJ 1996, Wong et al., 2006, Flint, et al., 2012, Koropatkin, et

al., 2012 and Larsbrink et al., 2014).

Microbial interactions with plant biomass are also central to technologies for the

conversion of lignocellulosic feedstocks into useful products. For example, ethanol is an

important renewable energy source that may replace petroleum-based transportation fuels

and reduce CO2 emissions (Sticklen 2008, Slade et al., 2009, Dwivedi et al., 2015).

Development of more sustainable processes for second generation biofuels derived from

lignocellulosic biomass is limited by the recalcitrance of the substrate, primarily due to its

lignin content, and often requires expensive pretreatment and use of hydrolytic enzymes

(Ragauskas, Williams et al. 2006, Chen and Dixon 2007, Sanderson 2011). An alternative

strategy to convert lignocellulosic feedstocks into biofuels is consolidated bioprocessing,

where microbes capable of producing a complex set of glycoside hydrolases decompose

plant biomass and ferment the products in a single step, for a more cost-effective process

with savings in capital and operational costs (Lynd et al., 2005, Jin, Balan et al., 2011,

Tolonen et al., 2011, Zuroff et al., 2013, Yee et al., 2014 and Petit et al., 2015). However,

details of microbe-biomass interactions and the impacts on cell growth and fermentation

are unknown due to a lack of methods for visualizing and quantifying microbial cells in

the presence of plant biomass.

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Here we present a simple and effective method for visualizing and quantifying

microbes interacting with insoluble plant substrates. The method involves separately

staining cells and substrate with two different fluorescent dyes. To evaluate our method,

we used the cellulolytic microbe Clostridium phytofermentans that produces ethanol as its

primary fermentation product and grows using a wide array of simple and complex

carbohydrate components of plant biomass as substrates (Warnick et al., 2002 and Petit et

al., 2015). We demonstrate that the dual-staining procedure may be used to determine

characteristics of C. phytofermentans growth and substrate attachment with insoluble

cellulosic substrates.

3.3 Materials and Methods

3.3.1 Bacterial strain and culture conditions

C. phytofermentans ISDg (ATCC 700394) was cultured using the anaerobic

techniques of Hungate in a modified form of medium GS-2C (Warnick, Methe et al. 2002)

containing the following (g/l): yeast extract, 6.0; urea, 2.1; KH2PO4, 2.9 K2HPO4 1.5;

Na2HPO4, 6.5; trisodium citrate dihydrate, 3.0; L-cysteine hydrochloride monohydrate,

2.0; and resazurin, 1; with pH adjusted to 7.0 using KOH. This basal medium was

supplemented with 0.3% (wt/vol) of the specific substrate (Whatman #1 filter paper, wild-

type Sorghum bicolor, or brown midrib (bmr) reduced lignin double mutant S. bicolor bmr-

6/bmr-12) ball-milled as described by Leschine and Canale-Parola (Leschine and Canale-

Parola 1983). Cultures were incubated at 30oC under anaerobic conditions (100% N2) as

described by Hungate (Hungate et al., 1966).

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3.3.2 Dual-staining procedure

Samples were prepared following a procedure similar to that described previously

(Zuroff et al., 2014). Prior to staining, each culture was diluted 1:1 with basal medium by

adding 1 ml of culture to 1 ml sterile basal medium and vortexed at maximum speed for 15

seconds. This diluted culture was further diluted 1:1 (50 µl in 50 µl of sterile basal medium)

and vortexed. Eight µl of a 10-µg/ml solution of Calcofluor White (Fluorescent Brightener

28, Sigma-Aldrich) were added to the diluted culture sample and incubated for 8 min in

the dark. Subsequently, 0.3 µl of SYTO® 9 (LIVE/DEAD BacLight Bacterial Viability

Kit, Thermo Fisher Scientific) were added to the sample and incubated for 7 min in the

dark. A volume of 16 µl of stained sample was placed on a slide and covered with a 22 x

22 mm cover slip. Figure 1 provides a diagrammatic representation of the dual-staining

protocol.

3.3.3 Fluorescence microscopy and image processing

A Nikon Eclipse E600 fluorescence microscope, equipped with a Diagnostic

Instruments Spot-RT CCD camera and a 40x objective was used to obtain images. For each

sample, differential interference contrast and epifluorescence micrographs were obtained.

Images were acquired and processed using SPOT Advanced imaging software. Cellulosic

growth substrates stained with Calcofluor White dye was visualized at 385 nm and cells

stained with SYTO 9 dye were visualized at 480 nm. For three-dimensional (3D) imaging,

multifocal plane images of 1 µM depth were obtained and combined to generate an image

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21 µM deep. Subsequently, the Z-stack was de-convolved using Autoquant 2.1 (Media

Cybernetics) and the three-dimensional image was rendered using Imaris 6.0 (Bitplane)

software.

3.3.4 Cell quantification

Micrographs were taken at ten different locations on each slide. A custom-written

script applied image dimensions, and cell counts were performed manually using Image J

(http://rsb.info.nih.gov/ij/). Substrate-attached cells and free cells were separately counted.

Vegetative and sporulating cells were included in cell counts. Cell numbers are the average

of triplicate determinations.

3.3.5 Determination of fermentation products

Non-gaseous fermentation products were determined by HPLC. Ethanol and

acetate concentrations in culture supernatant fluids were measured using a BioRad Aminex

HPX-87H 300x7.8 mm column at 30oC with 5 mM H2SO4 as mobile phase at a flow rate

of 0.60 ml/min in a Shimadzu LC-20AD HPLC with a RID-10A Refractive Index Detector.

3.3.6 Data analysis

Data analysis was performed using Prism 6.0 (GraphPad Software) One-way

ANOVA with a Tukey’s post-test to measure variance. Error bars represent standard

deviation of the mean.

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3.4 Results

3.4.1 Visualization of C. phytofermentans cells growing on insoluble substrates using

the dual-staining procedure.

Cellulolytic bacteria, such as C. phytofermentans, are known to adhere to their

cellulosic substrates and many species form biofilms on this nutritive surface (Alonso et

al., 2008, Wang et al., 2011, Young et al., 2012, Dumitrache et al., 2013 and Zuroff et al.,

2014). However, differentiating cellulolytic microbial cells from their insoluble growth

substrates, enumerating cells, and quantifying cell population growth are major challenges.

Therefore, the use of light microscopy and fluorescent dyes to distinguish plant-

decomposing microbes from their growth substrates is appealing since it would enable

researchers to measure microbial growth on plant biomass in terms of an increase in cell

number while also observing microbe-substrate interactions. Representative images

showing cell-substrate interactions are presented in Fig. 2. A nucleic acid dye, Syto 9,

(LIVE/DEAD BacLight Bacterial Viability Kit; (Boulos et al., 1999) was used to stain

bacterial cells and Calcofluor White, a cellulose/chitin dye, stained plant material and filter

paper. In order to demonstrate the difficulty of visualizing cells growing on insoluble plant

substrates by light microscopy, differential interference contrast (DIC) micrographs were

also obtained (Fig. 2A). However, when viewed by fluorescence microscopy at 385-400

nm, the filter paper and plant fibers were readily visualized due to the intense blue

fluorescence of Calcofluor White staining, but bacterial cells were not apparent (Fig. 2B).

When samples were illuminated at 480 nm, bacterial cells fluoresced green due to Syto 9

staining of their nucleic acids (Fig. 2C). The two fluorescent images (Fig. 2B and 2C) were

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merged in Fig. 2D to demonstrate cell-substrate differentiation with bacterial cells stained

green by Syto 9 and the paper and plant fiber stained blue with Calcofluor White.

3.4.2 Cell morphology and substrate interactions visualized by 3D imaging.

In order to visualize interactions of C. phytofermentans cells with an insoluble

growth substrate, cultures in filter paper medium were stained using the dual-staining

procedure and three-dimensional images were obtained as illustrated in Figure 3. Green

fluorescing rod-shaped cells of varying lengths were observed, some with swellings at one

end, presumably due to endospore formation as described previously by Warnick et al.

(Warnick, Methe et al. 2002). Some C. phytofermentans cells were aligned parallel to

cellulose fibers as observed by Zuroff et al. (Zuroff, Gu et al. 2014), while many cells

appeared to be attached perpendicularly to finer threads of the cellulose substrate with the

putative endospores located opposite the site of attachment as observed in Clostridium

thermocellum (27). In the XY cross-sectional planes, cells were visualized within the

substrate. Such 3D images indicate the potential of the dual staining protocol to image

bacterial cells interacting with insoluble plant biomass substrates.

3.4.3 Cell growth and ethanol production during fermentation of insoluble substrates

by C. phytofermentans.

Measurements of growth of bacteria on their insoluble substrates are problematic

for several reasons. Viable counts on plates of agar media (Hungate 1950, Kistner 1960

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and Hendricks et al., 1995) are complicated by the propensity of cellulose-decomposing

cells to adhere to their substrate, frustrating efforts to accurately dilute cultures. Also,

viable counts are time-consuming and challenging for fastidious anaerobes. Our method,

which allows rapid, direct enumeration of cellulolytic bacterial cells, is facilitated by the

use of two fluorescent dyes that enhance the contrast between microbes and cellulosic

substrates.

Growth of C. phytofermentans on filter paper, wild-type S. bicolor, or lignin double

mutant S. bicolor bmr-6/bmr-12 was measured (Fig. 4A). With filter paper as substrate,

cell numbers increased over five days, whereas cell numbers on both wild-type and lignin

double mutant sorghum reached a maximum at day 3 with 1.1x108 cells/ml and 2.7x108

cells/ml, respectively (Fig. 4A). However, the ethanol concentration in cultures with all of

the three substrates steadily increased for 5 days (Fig. 4B). Consistent with previously

reported results (Lee et al., 2012), cultures with lignin double mutant sorghum as substrate

produced significantly more ethanol than cultures grown on wild-type sorghum.

Interestingly, cultures with filter paper as substrate produced significantly less ethanol than

cultures with an equivalent amount of either wild-type or lignin double mutant sorghum

(Fig. 4B), indicating the important chemical differences in these substrates. When filter

paper served as substrate, after day 2, ethanol production per cell decreased (Fig. 4C) as

cell numbers increased (Fig. 4A). Since both vegetative and sporulating cells were included

in cell counts, reduced ethanol production per cell may reflect cell differentiation and the

onset of sporulation, which would suggest that sporulating cells are less metabolically

active. Unexpectedly, ethanol production per cell in cultures grown with either wild-type

or lignin double mutant sorghum increased after three days of growth (Fig. 4C) when cell

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numbers were decreasing (Fig. 4A). Possibly, with an early onset of sporulation, a portion

of the microbial cell population was less metabolically active during the first three days of

growth on sorghum, and, upon completion of sporulation, mature spores, which were not

counted, were lost from the recorded cell counts. These results also suggest that limiting

sporulation may be an effective strategy to increase metabolic activity and ethanol

production in C. phytofermentans.

Figure 4 also includes a direct comparison of cell number (Fig. 4D), substrate

attachment (Fig. 4E) and ethanol production (Fig. 4F) after three days of C.

phytofermentans growth on the three different biomass substrates. Although cell growth on

filter paper exceeded growth on either sorghum substrate (Fig. 4D), there was no

statistically significant difference in the percentage of cells attached to filter paper (88%)

and to lignin double mutant sorghum (89%) (Fig. 4E). However, cell attachment to wild-

type sorghum was significantly lower than to either filter paper or lignin double mutant

sorghum (Fig. 4E). The increased microbial cell attachment to the lignin double mutant

sorghum as compared to wild-type sorghum may reflect greater access to cellulose and

other polysaccharide components of biomass in the lignin double mutant sorghum. While

cell numbers in cultures with either sorghum substrate were low compared to cultures

growing on filter paper (Fig. 4D), the concentration of ethanol produced with wild-type

sorghum (17.5 mM) or lignin double mutant sorghum (20.7 mM) was significantly greater

than that produced with filter paper as substrate (13.7 mM) after three days of growth (Fig.

4F), possibly due to a higher content of readily fermentable polysaccharides in the sorghum

substrates.

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These experiments indicated that the C. phytofermentans growth cycle varies

greatly depending on the cellulosic substrate. With filter paper as growth substrate, ethanol

production was strongly correlated with cell number (Fig. 5A). Surprisingly, little or no

correlation was found between cell number and ethanol production with either wild-type

sorghum (Fig. 5B) or lignin double mutant sorghum (Fig. 5C) as growth substrate. As

shown in Figure 4, ethanol production occurred later in the growth cycle on both sorghum

substrates when C. phytofermentans cell numbers were decreasing, indicating that some

cells remained active while the overall cell population decreased, possibly due to

sporulation.

3.5 Discussion

The dual-staining procedure described here is a simple and efficient method to

visualize and enumerate cells in the presence of insoluble plant biomass substrates. This

method provides direct total cell counts, and may replace or complement complex and

indirect methods for measuring microbial growth on cellulosic substrates (Yang et al.,

2009). Also, growth characteristics of co-cultures and other complex communities may be

assessed using this method if community members are morphologically distinguishable.

Importantly, the dual staining procedure enables studies of interactions of microbial cells

with insoluble substrates such as plant biomass feedstocks. It is possible to employ this

method to study cell-substrate interactions using 3D visualization with little impact on the

sample. Also, coupling the dual-staining procedure with image processing techniques, cell

counts may be rapidly performed enabling high-throughput analyses. A possible limitation

of this method might be interference due to extracellular polymeric substances (EPS)

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produced by biofilm formation on substrates, making cell visualization more difficult;

however, EPS formation did not pose a problem in the current study.

Our observation of higher ethanol production by C. phytofermentans during

fermentation of lignin double mutant sorghum as compared to wild-type sorghum is

consistent with the findings of Lee et al (Lee et al., 2012). Also, correlating with higher

ethanol production, we observed higher microbial cell attachment to the lignin double

mutant sorghum as compared to wild-type sorghum. This may be attributed to the fact that

the lignin content of plant biomass adversely affects substrate attachment, decomposition,

fermentation, and subsequently, ethanol production (Lynd et al., 2002, Fu et al., 2011, Lee

et al., 2012 and Lacayo et al., 2013).

Using the dual-staining procedure, we examined C. phytofermentans growth on

different insoluble substrates. According to company literature, Whatman #1 filter paper is

prepared from cotton linters that are treated to achieve an alpha cellulose content of at least

98%, and as such, it is a highly processed, relatively pure form of cellulose. We observed

very different growth characteristics on purified cellulose (filter paper) as compared to

growth on wild-type and mutant sorghum. Both cell numbers and ethanol concentration

increased for five days when filter paper served as substrate. However, with wild-type and

mutant sorghum as substrate, cell numbers reached a maximum after three days of growth

and then decreased while ethanol concentration continued to increase for five days. With

filter paper as growth substrate, ethanol production was correlated with cell number but,

unexpectedly, little or no correlation was found between cell number and ethanol

production with either wild-type or mutant sorghum as growth substrate. This result

indicated that only a portion of the microbial cell population remained active during growth

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on sorghum, possibly due to differentiation and the onset of sporulation in some cells,

resulting in reduced metabolic activity and lower ethanol production.

The dual-staining procedure described here is an effective means for quantifying

microbial cells growing on plant biomass substrates. Due to its simplicity, quantitative data

may be obtained rapidly and efficiently. In the dual-staining procedure, individual

fluorescent dyes are added at separate times. Syto 9, a nucleic acid dye, stains cells and

emits a green fluorescence that is observed by fluorescence microscopy. Cellulosic

biomass substrates are stained by Calcofluor White, which emits a blue fluorescence when

exposed to UV light. We demonstrated that visualization and enumeration of microbial

cells growing on different cellulosic substrates may be performed by obtaining

micrographs and counting cells using Image J. Statistical analyses may be used to evaluate

differences in cell growth and substrate attachment, which in turn, may be correlated with

fermentation product formation. Finally, use of the dual-staining procedure for three-

dimentional visualization of microbes, such as C. phytofermentans, growing on insoluble

plant substrates is a useful tool for gaining insight into plant-microbe interactions in a broad

range of applications from biofuels to human health.

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Figure 3.1 Diagrammatic overview of dual-staining protocol.

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Figure 3.2 Differential interference contrast (DIC) and epifluoresence micrographs of C.

phytofermentans cultured for 3 days with filter paper, wild-type S. bicolor (WT Sorgum),

or lignin double mutant S. bicolor (bmr-6/bmr-12 Sorghum). Cultures were prepared for

microscopy using the dual-staining procedure. For each sample, one field is shown imaged

by DIC microscopy (A), and fluorescence microscopy at 385-400 nM showing cellulose

fibers stained with Calcofluor White (B) and at 480 nM showing Syto 9 stained cells (C).

The epifluorescence images in rows B and C are merged in the row D images. Stained

culture preparations were diluted with sterile basal culture medium prior to slide

preparation: the filter paper culture was dilute 1:3 and sorghum cultures were diluted 1:1.

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Figure 3.3 Two-dimentional rendering of a 3D image of C. phytofermentans cultured for

five days in filter paper medium. The culture was prepared for microscopy using the dual-

staining procedure. Multiple focal plane images at a dept of 1 µm were obtained and used

to generate a 3D images.

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Figure 3.4 Cell growth (A) and ethanol production (B, C) by C. phytofermentans, and cell

numbers (D), cell attachment to substrate (E) and ethanol concentration (F) after three days

of growth with filter paper (), wild-type S. bicolor (), or lignin double mutant S. bicolor

bmr-6/bmr-12 () as growth substrate. The concentration of ethanol in cultures is

expressed as mM (B, F) and the amount of ethanol produced per cell is expressed as nmoles

per cell (C). Error bars represent standard deviation of the mean (n=3). Statistically

signficant differences are represented with asterisks; one asterisk for P ≤ 0.05, two asterisks

for P ≤ 0.01, and three asterisks for P ≤ 0.001.

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Figure 3.5 Correlation of ethanol production with cell number during growth. Ethanol

production is given as a function of the cell number in cultures with filter paper (A), wild-

type S. bicolor (B) and lignin double mutant S. bicolor bmr-6/bmr-12 (C) as growth

substrate.

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CHAPTER 4

IMAGING INTRACELLULAR PROTEIN-BOUND MICROCOMPARTMENTS

IN BACTERIAL CELLS

4.1 Abstract:

Bacterial microcompartments (BMCs) are polyhedral organelles that harbor

metabolic enzymes and are bounded by a protein shell. Three loci with genes encoding

shell proteins of BMCs were identified in the genome of Clostridium phytofermentans, a

soil bacterium that directly converts biomass to biofuels. One of the BMC loci was

homologous to a BMC-encoding locus in the human gut commensal microbe Roseburia

inulinivorans and was hypothesized to be involved in the conversion of fucose and

rhamnose to propanol and propionate in C. phytofermentans. In order to demonstrate the

presence of BMCs within cells of C. phytofermentans using transmission electron

microscopy (TEM), it was necessary to develop a fixation, embedding and staining

protocol that would image BMCs while preserving the structure of the bacterial cell

envelope, the cell membrane and wall. A procedure involving cell fixation with sodium

cacodylate buffer containing glutaraldehyde and picric acid and staining with osmium

tetroxide and potassium ferrocyanide was developed. Thin sections of C. phytofermentans

cells examined using TEM revealed the presence of polyhedral structures, presumably

BMCs, in C. phytofermentans cells grown with fucose or rhamnose but not with glucose

as growth substrate.

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4.2 Introduction:

Bacterial microcompartments (BMCs) are polyhedral organelles with protein shells

that harbor metabolic enzymes. The first BMCs to be identified were the carboxysomes

(Shively et al., 1973). Carboxysomes contain enzymes that fix CO2 including ribulose-1,5-

bisphosphate carboxylase/oxygenase (RuBisCO). Other varieties of BMCs are found in

multiple microbes; some are implicated in human health such as 1,2-propanediol- and

ethanolamine-producing BMCs involved in the pathogenicity of Salmonella enterica

(Joseph et al., 2006, Klumpp et al., 2007, Sriramulu et al., 2008, and Harvey et al., 2011).

Moreover, it has been found that microbes in the Planctomycetes phylum are capable of

encoding BMCs capable of breaking down rhamnose, fucose and fucoidan in the presence

of oxygen (Lage et al., 2014).

Clostridium phytofermentans is a soil bacterium that directly converts all major

components of plant biomass to ethanol as its primary product of fermentation (ref:

Warnick et al., 2002). Analysis of the genome sequence of C. phytofermentans revealed

the presence of three loci with genes encoding shell proteins of BMCs (Petit et al., 2013 &

2015). Transcriptional profiling analyses indicated that one of the three loci is expressed

during growth of C. phytofermentans on fucose or rhamnose (Petit et al., 2013). This BMC

locus has homology to a BMC-encoding locus in the human gut microbe, Roseburia

inulinivorans, a member of the Lachnospiraceae family (phylum Firmicutes, class

Clostridia), which also includes C. phytofermentans (Scott et al., 2006 & 2011). In R.

inulinivorans, BMCs encoded by this locus are believed to be involved in the metabolism

of 1,2-propanediol, an intermediate in the conversion of fucose to propionate and propanol

(Scott et al., 2006 and Petit et al., 2013). In S. enterica 1,2-propanediol is produced during

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growth on either fucose or rhamnose and secreted, but may be taken up and converted into

propionate and propanol by enzymes encoded by genes within a BMC locus (Obradors et

al.,1988, Baldona et al., 1988, Buan et al., 2004, Penrod and Roth 2006, Cheng et al., 2011,

Crowley et al., 2010, Harvey et al., 2011 and Chen et al., 2012).

Fucose and rhamnose are deoxyhexose sugars (also known as methyl-pentoses)

found in plant cell walls, in bacterial cells, in some insect species, and on the surface of

mammalian cells as components of glycans and glycopeptides. In plant cell walls, fucose

and rhamnose form a component of pectin, highly hydrated, branched, heterogeneous

polysaccharides, notably in rhamnogalacturonans (Leschine et al., 2005). Fucose is

commonly found on plant and mammalian cell surfaces as N- and O-linked glycosylated

proteins (glycans); for example, fucose is highly abundant in the intestinal tracts of

mammals (Robbe et al., 2003) where it plays an important role in signaling of and

colonization by members of the gut microbiome. (Pacheco et al., 2012). Fucosylated

proteins of the intestinal epithelium also play a role in maintaining the intestinal

microbiome during illness in the host (Pickard et al., 2014).

C. phytofermentans is a microbe capable of converting multiple components of

plant biomass into ethanol, making it an attractive tool for the bioethanol industry. In this

study, we developed a procedure for visualizing BMCs within cells of C. phytofermentans

grown with fucose or rhamnose as substrate. A TEM embedding protocol was developed

for stabilizing the cell wall and improving BMC detection in transmission electron

micrographs. Visualization of BMCs within cells of C. phytofermentans correlated with

the expression of several genes homologous to those found in the BMC locus of R.

inulinivorans, and also with expression of fucose dissimilatory enzymes and an ATP-

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binding cassette transporter as dominant transcripts. Taken together, the data are consistent

with the conclusion that the C. phytofermentans BMC locus encodes a microcompartment

involved in the fermentation of fucose and rhamnose (Petit et al., 2013).

4.3 Materials and Methods:

4.3.1 Bacterial strain and conditions:

C. phytofermentans was cultured in a modified form of an anaerobic medium

containing the following (g/l): yeast extract, 6.0; urea, 2.1; KH2PO4, 4.0; Na2HPO4, 6.5;

trisodium citrate dihydrate, 3.0; L-cysteine hydrochloride monohydrate, 2.0; and resazurin,

1; with pH adjusted to 7.0 using KOH. This medium was supplemented with 0.3% (wt/vol)

of the specific substrate (glucose, fucose or rhamnose) added as a filter-sterilized solution

to the sterile medium. Triplicate liquid cultures were incubated at 30°C under anaerobic

conditions (100% N2) as described by Hungate (Hungate et al., 1950). Growth was

determined spectrophotometrically by monitoring changes in optical density at 660 nm.

4.3.2 Transmission Electron Microscopy:

C. phytofermentans cells, cultured with fucose or rhamnose as growth substrate,

were grown until mid-exponential phase, fixed for 1 hr at room temperature with 2.5%

glutaraldehyde, 0.03% picric acid, and 0.5% NaCl in 0.05 M cacodylate buffer and washed

three times in 0.05 M cacodylate buffer containing 0.5% NaCl. Gel pellets were formed by

adding 3% agarose, incubated in ice for 15 min followed by post-fixation with 1% osmium

tetroxide, and 1.5% potassium ferrocyanide in 0.05 M cacodylate buffer for 2 hrs. The gel

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pellets were washed with water and stained by submerging in 2% uranyl acetate for 1 hr,

followed by three additional water washes and subsequent dehydration by increasing

ethanol concentration. Samples were embedded in 26% Epon-812, 15.5% Araldite-502,

57% dodecenyl succinic anhydride and 1.5% epoxy accelerator DMP-30, and polymerized

at 60°C overnight. Thin sections (70 nm) were cut using a Reicher-Jung Ultracut E

microtome; sections were stained with lead citrate, and observed using a JEOL 100CX

transmission electron microscope at 100 kV.

4.4 Results and Discussion:

Cell wall structure was stabilized by using a cell fixation and embedding protocol

that incorporated picric acid and potassium ferrocyanide. As a control, thin sections of C.

phytofermentans cells grown using glucose as substrate were prepared; BMCs were not

observed within cells cultured on glucose (Fig. 4.1). However, BMCs structures were

visualized in C. phytofermentans cells grown in a culture medium containing fucose (Fig.

4.2) or rhamnose (Fig. 4.3). The diameter of the BMCs in cells grown with either fucose

or rhamnose ranged from 30 to 50 nm. Although potassium ferrocyanide and osmium

tetroxide enhanced the visualization of BMCs and the overall appearance of C.

phytofermentans cells in micrographs, the mechanisms behind the enhanced contrast of

proteins are unknown. It is suspected that reduction of osmium by potassium ferrocyanide

plays a role (Bozzola and Russell, 1998). The addition of picric acid at the initial fixation

step most likely enhanced of the preservation of cell envelope structure. The embedding

protocol developed for visualizing BMCs efficiently and effectively enhanced the contrast

of samples while preserving cell wall structure.

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The protocol developed in this study enabled visualization of BMCs within cells of

C. phytofermentans grown with fucose or rhamnose as substrate. The presence of BMCs

within cells of C. phytofermentans correlated with the expression of several genes

homologous to those found in the BMC locus of R. inulinivorans. Also, the appearance of

BMCs in C. phytofermentans cells cultured on fucose correlated with expression of fucose

dissimilatory enzymes and an ATP-binding cassette transporter as dominant transcripts.

Results presented here, along with findings presented in Petit et al., 2013 (Petit et al., 2013),

indicate that the C. phytofermentans BMC locus encodes a microcompartment involved in

the fermentation of fucose and rhamnose.

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Figure 4.1 Transmission electron micrograph of C. phytofermentans cells from cultures grown

with glucose as growth substrate. Polyhedral structures were not observed in cells from glucose-

grown cultures.

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Figure 4.2 Transmission electron micrograph of C. phytofermentans cells from cultures grown in

fucose as growth substrate. Some cells contained polyhedral structures (arrow), presumably

bacterial microcompartments.

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Figure 4.3 Transmission electron micrograph of C. phytofermentans cells from cultures grown in

rhamnose as growth substrate. Multiple polyhedral structures, presumably bacterial

microcompartments, were observed in thin sections of C. phytofermentans grown in rhamnose

culture medium.

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CHAPTER 5

MEMBRANE ENERGETICS AND ENERGY CONSERVATION IN THE

CELLULOLYTIC ETHANOLOGEN CLOSTRIDIUM PHYTOFERMENTANS

5.1 Abstract:

Clostridium phytofermentans is a microbe from forest soil that ferments a wide array of

plant polysaccharides and produces ethanol as its primary fermentation product. Realizing the

industrial potential of this microbe as a catalyst for biofuels production requires a full

understanding of the metabolic pathways and associated membrane energetic mechanisms related

to ethanol production. Genome and transcriptome analyses of C. phytofermentans have revealed

numerous highly expressed genes involved in membrane energetics and energy conservation that

are potentially related to ethanol production, including an Rnf complex (Cphy_0211-0216), a

membrane-bound sodium- and proton-translocating pyrophosphatase (Cphy_1812), and three

ATPases. Identification of sodium-translocating F-type (Cphy_3735-3742) and V-type

(Cphy_3070-3077) ATPases was confirmed by the presence of conserved sodium binding motifs

in the c subunits of these enzymes. In support of this annotation, ATP synthase activity was

inhibited by the sodium ionophore, ETH 2120. Structural information for the third ATPase (V-

type ATPase; Cphy_3440-3447) was not available and its ion specificity was not directly

determined. Integral ferredoxin oxidoreductase activity of the Rnf-complex was assayed by using

inverted membrane vesicle preparations and following reduction of ferredoxin by NADH. Activity

required ATP, and was inhibited by the proton ionophore TCS, but not by the sodium ionophore

ETH 2120, indicating that the reaction was driven by a proton electrochemical gradient, generated

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through the activity of the previously uncharacterized V-type ATPase. These results indicate that

C. phytofermentans conserves energy by generating both sodium and proton electrochemical

gradients across the membrane. In support of this conclusion, C. phytofermentans growth was

inhibited by both the proton ionophore TCS and the sodium ionophore ETH 2120. Based on these

observations, we conclude that C. phytofermentans is able to conserve energy through ferredoxin

oxidoreductase activity of the Rnf-complex, resulting in increased levels of NADH for ethanol

production.

5.2 Introduction:

Clostdridium phytofermentans is a mesophilic anaerobe that ferments multiple plant

substrates, including cellulose, hemicellulose, pectin, and starch, and produces ethanol as its

primary fermentation product (Warnick et al 2002). Additionally, fermentation of fucose and

rhamnose into propanol and propionate makes C. phytofermentans a strong candidate for the

industrial conversion of plant biomass into various bioproducts including liquid transportation

fuels (Petit et al., 2013; Petit et al., 2015). C. phytofermentans also has been deployed as a catalyst

in an assay to detect plant biomass phenotypic variations as a measure of plant feedstock quality

(Lee et al., 2012). The effectiveness of this assay is tied to a property of the microbe, that ethanol

production is proportional to feedstock quality (digestibility), and recognition of the ease and

accuracy of ethanol concentration measurements. Technologies for improved ethanol productivity

also have been developed, including a symbiotic bioprocessing of cellulose to ethanol involving

coculturing C. phytofermentans with Saccharomyces cerevisiae (Zuroff et al., 2013) and a

metabolic engineering strategy to improve ethanol tolerance (Tolonen et al., 2015)

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Genomic and transcriptomic analyses have indicated that C. phytofermentans utilizes the

Embden-Meyerhof-Parnas pathway to metabolize sugars to pyruvate, which is converted to

Acetyl-CoA, the precursor for acetate and ethanol formation. (Petit et al., 2015). Acetyl-CoA may

be reduced to ethanol in reactions coupled to the oxidation of NADH, or converted to acetate by

phosphate acetyl transferase and acetate kinase, along with formation of ATP. However,

production of ethanol as the primary product of fermentation suggests that C. phytofermentans

may also use a chemiosmotic mechanism to meet energy (ATP) requirements. We hypothesized

that the ferredoxin:NAD+ oxidoreductase activity of an Rnf complex, which had been identified

in analyses of the C. phytofermentans genome, transcriptome, and proteome (Petit et al., 2015 and

Tolonen et al., 2011 & 2013), contributes to a trans-membrane electrochemical gradient supplying

energy for ATP synthesis while also replenishing the pool of NADH, providing reductant for

ethanol production.

The specific ion dependence of the C. phytofermentans Rnf oxidoreductase activity has not

previously been reported. In Clostridium ljungdahlii, autotrophic growth is dependent on an Rnf–

driven proton gradient (Tremblay et al., 2012), while the acetogens Acetobacterium woodii and

Eubacterium limosum, along with several other bacteria, use an Rnf complex to generate a sodium

gradient, which contributes to ATP synthesis in these microbes (Hess et al., 2013; Hess et al.,

2016; Jeong et al., 2015; Müller et al., 2008).

The genome and transcriptome study mentioned above (Petit et al., 2015) also identified

genes for three ATPases among highly expressed genes likely to participate in energy conservation

in C. phytofermentans. Based on sequence alignment comparisons, an F-type (Cphy_3735-3742)

and a V-type (Cphy_3070-3077) ATPase were predicted to be sodium-translocating membrane

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complexes. A third ATPase (V-type ATPase cluster Cphy_3440-3447) was not highly similar to

any characterized ATPase complexes and its ion specificity was not predicted (Petit et al., 2015).

Additionally, highly expressed pyrophosphate-dependent glycolytic enzymes also have been

detected in C. phytofermentans, and it was hypothesized that utilization of these energetically

efficient enzymes (pyrophosphate- rather than ATP-dependent) may indirectly contribute to

ethanol production by increasing the ATP yield of glycolysis and decreasing the demand for ATP

generation via acetate production (Petit et al., 2015). Interestingly, a membrane-bound sodium-

and proton-translocating pyrophosphatase (Cphy_1812) has been identified in the genome of C.

phytofermentans (Luoto et al, 2012). This enzyme may couple the energy of pyrophosphate

hydrolysis to the formation of sodium and proton electrochemical gradients across the membrane

to drive ATP synthesis. Conversely, this membrane-integral enzyme may catalyze the synthesis of

pyrophosphate, reducing the demand for ATP, by using the energy of an electrochemical gradient,

for example, the gradient generated through the activity of the Rnf complex.

The objective of this study is to advance understanding of the metabolic pathways and

associated membrane energetic mechanisms related to ethanol production by C. phytofermentans.

Recent genomic and transcriptomic analyses have identified several proteins that are likely

involved in membrane energetics and energy conservation in this microbe (Petite et al., 2015).

Structural bioinformatics and biochemical assays were used to characterize the ATPases and Rnf

complex. Based on the results of these studies, we generated a metabolic model for energy

conservation in C. phytofermentans involving both a sodium and proton electrochemical gradient.

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5.3 Materials and Methods

5.3.1 Protein sequence alignment, comparison and structure modeling

The website Uniprot (UniProt, 2015) was used for protein sequence identification. Protein

sequence alignments were performed using Clustal Omega (Sievers et al., 2011, Goujon et al 2010

and McWilliam et al., 2013) and PROSMAL 3D (Pei et al., 2008). Protein structure homology-

modelling was done using SWISS-MODEL (Bordoli et al.,2009). Molecular visualization of the

modelled proteins was visualized and rendered using Visual Molecular Dynamics (Humphrey et

al., 1996).

5.3.2 Bacterial strain and growth conditions

C. phytofermentans ISDg (ATCC 700394) was cultured in a defined medium, M6 (2.0 g/l

NaH2PO4, 10.0 g/l K2HPO4, 1.0 g/l (NH4)2SO4, 1.0 g/l L-cysteine hydrochloride monohydrate, 20

ml/l XT solution (5.0 g/l xanthine and 5.0 g/l thymine in 0.06 N NaOH) 10 ml/l AA1 solution (5.0

g/l of each of the following amino acids: alanine, arginine, histidine, isoleucine, leucine,

methinonine, proline and valine), and 10 ml/l Bach's trace element (BTE) solution [49]. Resazurin

(1 mg/l) was added as an oxidation/reduction indicator. After autoclaving, 30 ml/l CPV5 solution

(20 mg/l p-aminobenzoic acid, 1 mg/l biotin, 30 mg/l folinic acid, 80 mg/l nicotinamide, 5 mg/l

pantethine, 2 mg/l pyridoxal hydrochloride, 30 mg/l riboflavin, and 10 mg/l thiamine) was added.

This medium was supplemented with 0.5% (wt/vol) of glucose. Cultures were incubated at 30oC

under anaerobic conditions (100% N2) as described by Hungate (Hungate 1950).

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5.3.3 Effects of ionophores on cell growth

Cells were inoculated into anaerobic culture tubes containing 10 ml M6 media, 25 µM final

concentration of the Na+ ionophore ETH 2120 or the protonophore 3,3’,4’,5-

tetrachlorosalicylanilide (TCS). Cell growth was measured every 12 hours by optical density at

600 nm.

5.3.4 Preparation of inverted membrane vesicles

Ten milliliters of cell culture were inoculated into 1 L of M6 medium, and incubated at

30°C for 5 days. Cells were harvested by centrifugation at 10,000 x g at 8oC for 45 minutes. After

centrifugation, cell pellets were sonicated for 45 minutes and homogenized using a glass tissue

grinder. Cell homogenates were resuspended in 5 ml of Buffer A (50 mM Tris-HCl with 2 mM

dithiothreitol (DT) at pH 8.0 in N2), centrifuged at 100,000 x g at 8oC for 45 minutes. Supernatant

fluid was removed; cell pellets were resuspended in 2 ml of Buffer A, followed by homogenization

using a glass tissue grinder. Washing steps were performed three times. After the final wash, the

membrane-associated fraction was resuspended in 2 ml of Buffer A, frozen in liquid N2 and stored

at -80oC.

5.3.5 ATP hydrolysis and ATP synthesis determinations

For ATP hydrolysis, 15 µl of a preparation of C. phytofermentans inverted membrane

vesicles (IMVs) was added in 1 ml of Buffer B (100 mM MES buffer, 100 mM Tris-HCl buffer,

5 mM MgCl2, 200 mM KCl, pH 7.8 in N2) and the reaction was initiated by the addition of Na2ATP

(2.5 mM, final concentration) at 30oC. Phosphate levels were measured using Abcam’s Phosphate

Assay Kit (Colorimetric). ATP synthesis was carried in the same conditions as indicated above.

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IMVs were added to Buffer B and the reaction was initiated by the addition of Na2ADP (2.5mM,

final concentration). ATP was measured using a luminometer via the luciferin-luciferase assay at

560 nm excitation and operated at 30oC.

5.3.6 Rnf-associated ferredoxin oxidoreductase activity

Reactions were performed at 30oC in a N2 atmosphere in anaerobic cuvettes sealed with

rubber stoppers. Reaction mixtures contained 1 ml of Buffer C (100 mM Tris-HCl, pH 7.8), 5 mM

MgCl2, 2 mM NaCl, 2 mM dithiothreitol (DT), 30 µM ferredoxin, 5 mM NADH and 10 µl IMVs).

The reaction was started by the addition of 2 mM of ATP final concentration. The ionophores ETH

2120 and 3,3’,4’,5-tetrachlorosalicylanilide (TCS) were added at a final concentration of 10 µM

each. Ferredoxin reduction was measured at 430 nm.

5.4 Results

5.4.1 Predicted structures of the Na+ binding sites of the F-type and V-type ATPase c-rings

To gain insight into the ion specificity of the C. phytofermentans ATPases, structural

models of the F-type ATPase and V-type ATPase c-rings were created by homology modeling.

The structure of the F-type c subunits from Fusobacterium nucleatum was used as a template to

compare with the c-ring from Cphy_3741. The c-ring of Enterococcus hirae, which contains V-

type c subunits, was used as a template to compare with the Cphy_3076 c ring. The resulting model

for Cphy_3741 consists of 11 c-subunits, with an ion-binding site in between each pair of subunits.

The C. phytofermentans F-type c-ring sequence was aligned with Ilyobacter tartaricus, F.

nucleatum and Clostridium paradoxum c-rings (Fig. 3). Each binding site contains the specific site

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for Na+ ion binding: Gln32, Glu65 and Val63. The fourth binding site for Na+, Ser66, is replaced

by threonine in the C. phytofermentans F-type c-ring sequence. The Cphy_3076 3D protein model

consists of 10 c-subunits and has a 53.12% identity with E. hirae (Mizutani et al., 2011). The C.

phytofermentans V-type c-ring sequence was aligned with E. hirae, E. faecalis, Clostridium

aspargiforme and Clostridium bolteae (Fig. 4). Results of this analysis indicate that the C.

phytofermentans V-type c-ring ion-binding groups are highly conserved and consistent with other

microbes that contain sodium-binding groups in V-type ATPases. Each binding site contains

specific amino acid sequences that bind Na+: Leu61, Thr64, Gln65, Gln110 and Glu139.

5.4.2 Structural model of the pyrophosphatase Na+, H+ binding sites

A C. phytofermentans membrane-bound pyrophosphatase (-PPase) structural model was

generated by homology modeling. The structure of the -PPase dimer from Bacteroides vulgatus

was used as a template to compare with C. phytofermentans ion binding subunits. The resulting

model consists of a dimer from the cytosolic side of the membrane. In order to identify the ion-

binding sites, the C. phytofermentans -PPase sequence was aligned with that of B. vulgatus,

Clostridium lentocellum, Prevotella dentalis and Prevotella melaninogenica. Results indicate that,

in the C. phytofermentans -PPase, each binding site contains the specific amino acid subunits

involved in the binding of Na+ and H+, Thr87, Phe91, Met117 and Asp143.

5.4.3 Effects of ionophores on C. phytofermentans growth

In order to identify the specific ion(s) involved in generating an electrochemical gradient

in C. phytofermentans, we examined growth in the presence of sodium and proton ionophores.

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Growth in a defined medium supplemented with glucose as an energy source was inhibited by both

the Na+ ionophore ETH 2120 and the H+ ionophore TCS. The ionophores inhibited cell growth

during the initial phases of exponential growth. Cell morphology did not change and cell lysis did

not occur when cells were exposed to the ionophores (Data not shown).

5.4.4 ATP synthesis in inverted membrane vesicles is driven by a Na+ gradient

ATP synthesis in inverted membrane vesicles was examined with and without the Na+

ionophore ETH 2120 or the H+ ionophore TCS. ATP synthesis was initiated by the addition of

ADP to reaction mixtures. A control sample generated ATP at a rate of 17 µmol/min. The rate of

ATP synthesis in the presence of TCS (23 µmol/min) was similar to, or greater than that in control

samples. In the presence of ETH 2120, the rate of ATP synthesis was significantly reduced to 3

µmol/min. No ATP synthesis was observed in the absence of ADP or PO4.

5.4.5 An ATP driven proton gradient supports NADH-dependent ferredoxin reduction

In order to determine the specific ion dependence of the C. phytofermentans Rnf

oxidoreductase activity, NADH-dependant ferredoxin reduction was measured in inverted

membrane vesicles. Energy for this reaction was supplied by an electrochemical ion gradient

driven by ATP hydrolysis, catalyzed by a membrane-bound ATPase. Thus, we assayed Rnf-

complex activity “in reverse” as described by Hess et al., 2013.

Inverted membrane vesicles, ferredoxin and NADH were added to reaction mixtures and

activity was initiated by the addition of ATP. Ferredoxin reduction did not occur in the absence of

ATP. Interestingly, ferredoxin reduction was significantly reduced when the protonophore TCS

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was added to reaction mixtures; however, when the sodium ionophore ETH 2120 was added,

ferredoxin reduction was significantly enhanced. These results indicated that the integral

ferredoxin oxidoreductase activity of the Rnf-complex is driven by a proton electrochemical

gradient, which was generated through the activity of the previously uncharacterized V-type

ATPase.

5.5 Discussion

Analysis of the genome of C. phytofermentans revealed genes that encode various

membrane-bound proteins, which were hypothesized to play a role in membrane energetics and

energy conservation (Petit et al., 2015). In order to test the hypothesis, we first analyzed the

sequence alignments and modeled one of the V-type ATPases, the F-type ATPase, and the

membrane-bound pyrophosphatase. A Na+-binding V-type ATPase c-ring protein model for

Cphy_3076 was generated using the E. hirae c-ring homologue (Fig. 5.1). The C. phytofermentans

Na+ V-type ATPase c-ring shares 53.12% sequence identity with a Na+ V-type ATPase from E.

hirae. The model generated a c-ring with 10 c-subunits including an ion-binding site for each

subunit (Fig. 5.1) (Mizutani et al., 2011). The c-ring features include conserved amino acid

residues, Leu61, Thr64, Gln65, Gln110 and Glu139, which bind Na+ and were perfectly aligned

with E. hirae, E. faecalis, C. aspargiforme and C. bolteae c-rings (Fig 5.2) (Murata et al., 2005).

The sequence of the c-ring of C. phytofermentans F-type ATPase was modeled using the Na+ F-

type ATpase c-ring (Cphy_3741) of Fusobacterium nucleatum. with a sequence identity of

63.10%, as a template (Schulz et al., 2013) (Fig. 5.3). The model yielded an 11-subunit c-ring with

an ion-binding site in each subunit (Fig 5.3). Sequence alignment of the F-type ATpase of C.

phytofermentans with I. tartaricus, F. nucleatum, and C. paradoxum revealed three of four amino

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acids that bind Na+: Gln32, Glu65 and Val63 (Fig 5.4). The fourth binding ion Ser66 is replaced

by threonine, which retains the same hydroxyl groups of serine that binds Na+. The acetogen A.

woodii contains an F/V-hybrid rotor ring that translocates Na+ (Matthies et al., 2014). Interestingly,

the c2/3 subunit of the hybrid rotor that binds Na+ in A. woodii, contains the same amino acid

sequence as C. phytofermentans F-type ATPase. Thus, we conclude that C. phytofermentans has a

Na+ F-type ATPase. In Petit et al there is a mention of a possible second V-type ATPase cluster

(Cphy_3440-3447) however it sequences does not match any known sequence of ATPases.

A membrane-bound ion-pumping pyrophosphatase (PPase) (Cphy_1812) that was a

moderately expressed in C. phytofermentans (Petit et al., 2015 and Luoto et al., 2012) was also

modeled using a H+ PPase from the plant Vigna radiata as template (Lin et al., 2012) (Fig 5.6).

Results indicate a 49.13% sequence identity between the proteins. C. phytofermentans -PPase

sequence was aligned with B. vulgatus, P. dentallis, C. lentocellum, and P. melaninogenica.

Results indicate that in the C. phytofermentans -PPase each binding site contains the specific amino

acid subunits involved in binding both Na+ and H+, Thr87, Phe91, Met117 and Asp143 (Fig 6).

Membrane-integral PPases translocate H+, Na+ or both during pyrophosphate (PPi) hydrolysis. C.

phytofermentans possess glycolytic enzymes such an adenylate kinase, which exclusively use PPi,

increasing the ATP yield of glycolysis (Mertens, 1993). The membrane-bound pyrophosphatase

of C. phytofermentans may use a Na+ and H+ electrochemical gradient to generate PPi, reducing

the demand for ATP generation via acetate production.

Experiments aimed at examining the effects of ionophores on C. phytofermentans growth

yielded the surprising result that growth with glucose as substrate was inhibited by both the sodium

ionophore ETH 2120 and the protonophore TCS. When ionophores were added at the time of

inoculum or at mid exponential phase (Fig. 5.7). In C. ljungdahlii cell growth is inhibited by TCS

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but not ETH 2120 (Tremblay et al., 2012). The opposite occurs in Clostridium paradoxum, which

possess a Na+ F-type ATPase; a sodium ionophore affects growth while the protonophore TCS

does not (Ferguson et al., 2006). Inhibition of cell growth by ionophores follows a pattern: C.

ljundahlli possesses an H+ Rnf complex and a H+ F-type ATpase, while C. paradoxum has a Na+

F-type ATPase. In contrast, C. phytfofermentans relies on both a H+ and Na+ electrochemical

gradient for growth, as occurs in F. nucleatum (Schulz et al., 2013).

To determine whether a H+ or Na+ electrochemical gradient is involved in ATP synthesis,

we measured the ATP synthase activity of inverted membrane vesicles. Activity was not inhibited

by the protonophore TCS but was significantly reduced by the sodium ionophore ETH 2120 (Fig.

5.8). This result indicates that C. phytofermentans possess a sodium-pumping V-type ATPase

and/or F-type ATPase.

Since a protein structure for the Rnf-complex is not available, we were unable to make a

protein model comparison of the A. woodii, and C. ljungdahlii Rnf complexes with that of C.

phytofermentans. Previous studies have determined that the C. phytofermentans Rnf-complex

genes (Cphy_0211-0216) are 44-61% similar to A. woodii (Petit et al., 2015). To investigate the

activity of the C. phytofermentans Rnf-complex, we measured ATP- and NADH-dependent

reduction of ferredoxin using inverted membrane vesicles. In this assay, activity of the Rnf-

complex is the reverse of that in cells, using electrons from NADH to reduce ferredoxin. The

reaction was initiated in the addition of ATP and ferredoxin reduction was measured at 430 nm

(Fig 5.9). Results showed an ATP dependence for ferredoxin reduction. Interestingly, Ferredoxin

reduction was significantly reduced in the presence of the protonophore TCS and enhanced in the

presence of sodium ionophore ETH 2120, suggesting a proton electrochemical gradient coupling

between an ATPase and the Rnf-complex. Based on these results, we conclude that the integral

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ferredoxin oxidoreductase activity of the Rnf-complex is driven by a proton electrochemical

gradient, which, in our assay, was generated through the activity of the previously uncharacterized

H+ V-type ATPase (Cphy_3440-3447) (Hess et al., 2012 and Tremblay et al., 2012).

With protein modeling and experimental results reported here, we developed a metabolic

model to illustrate the role of membrane energetics in energy conservation in C. phytofermentans,

and to offer an explanation for how this microbe is able to generate high levels of ethanol while

maintaining adequate pools of ATP (Fig 5.10). The Rnf complex utilizes electrons from reduced

ferredoxin to reduce NAD+ to NADH while translocating H+ across the membrane, generating an

electrochemical gradient that may be used by the previously uncharacterized H+ V-type ATPase.

A membrane-bound pyrophosphatase (PPase) contributes to ATP formation by augmenting H+

and Na+ electrochemical gradients that may drive ATP synthesis by the Na+ V-type and Na+ F-

type ATPases. We propose that the Rnf complex contributes to ethanol production by producing

NADH required to reduce Acetyl-CoA to ethanol.

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Figure 5.1 Model of the C. phytofermentans Na+ F-type ATPase c-ring. An 11 c-ring homo-mer was developed including the location

of amino acids involved in Na+ binding. The model was developed using SWISS MODEL and 3D rendered using VMD 1.9.1.

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Figure 5.2 Alignment of F-type ATPase amino acid sequences. Conserved amino acids involved in Na+ are marked with an asterisk.

The C. phytofermentans protein contains the same conserved amino acids found in other F-type ATPases, V63, E65 and Q32 with the

exception of T66. The sequence alignment was performed with microbes know to possess Na+ F-type ATPases: I. tartaricus, F.

nucleatum and C. paradoxum. Sequences were aligned using PROSMAL3D and visualized by JalView.

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Figure 5.3 A model of the C. phytofermentans Na+ V-type ATPase c-ring protein. A 10-c-ring homo-mer was developed containing the

location of amino acids involved in Na+ binding. The model was developed using SWISS MODEL and 3D rendered using VMD 1.9.1.

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Figure 5.4 Alignment of V-type ATPase amino acid sequences. Conserved amino acids involved in binding Na+ are marked with an

asterisk. C. phytofermentans contains the same amino acids as known microbes with Na+ V-type ATPases: E. hirae and E. faecalis.

C. asparagiforme and C. bolteae proteins have the greatest similarity to that of C. phytofermentans. Sequences were aligned using

PROSMAL3D and visualized by JalView.

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Figure 5.5 Model of the pyrophosphatase protein of C. phytofermentans. The model shows conserved amino acids T87 (magenta

spheres), F91 (purple spheres), M117 (yellow spheres) and D143 (red spheres) involved in binding H+ and Na+. The model was

developed using SWISS MODEL and 3D rendered using VMD 1.9.1.

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Figure 5.6 Alignment of the C. phytofermentans pyrophosphatase amino acid sequence with other

pyrophosphatase sequences.

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Figure 5.7 Effect of ionophores on growth of C. phytofermentans in defined medium M6 with

glucose as substrate. Cell growth was inhibited by both ETH 2120 and TCS when added at the

time of inoculation or at mid-exponential phase of growth.

0 1 0 2 0 3 0 4 0 5 0 6 0

0 .0

0 .2

0 .4

0 .6

0 .8

1 .0

1 .2

C o n tro l

E T H 2 5 u M

T C S 2 5 u M

T C S M id

H o u rs

OD

60

0n

m

E T H M id

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Figure 5.8 Effect of ionophores on ATP synthesis in inverted membrane vesicles. ATP synthesis

was initiated by the addition of ADP to reaction mixtures. ATP was measured by luminescence

using a luciferin-luciferase assay kit. ATP synthesis was significantly reduced in reactions

containing the Na+ ionophore ETH 2120.

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Figure 5.9 NADH-dependent ferredoxin reduction assay of Rnf oxidoreductase activity in

inverted membrane vesicles. Reaction mixtures contained ferredoxin, NADH, and inverted

membrane vesicles in buffered solution. Reactions were initiated by addition of ATP at 60

seconds. In some reactions, ETH 2120 or TCS was added at a concentration of 10 µM.

Ferredoxin reduction was measured at 430 nM.

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Figure 5.10 Metabolic model illustrating the role of membrane energetics in energy conservation in C. phytofermentans. The Rnf

complex utilizes electrons from reduced ferredoxin to reduce NAD+ to NADH while translocating H+ across the membrane generating

an electrochemical gradient used by a H+ V-type ATPase. A pyrophosphatase (PPase) augments the H+ and Na+ electrochemical

gradients driving ATP synthesis by the Na+ V-type and Na+ F-type ATPases,

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CHAPTER 6

DISCUSSION AND FUTURE DIRECTIONS

This dissertation has presented three research projects with the goal of 1) whether a

correlation between cell growth and ethanol production existed in the presence of different plant

biomass substrates, 2) visualizing and characterizing bacterial microcompartments produced by C.

phytofermentans when fucose and rhammnose gets fermented, and 3) generating a bioenergetics

model of C. phytofermentans using genomic and biochemical techniques that links energy

conservation of membrane proteins and their role in ethanol production.

The first research project described the development of a dual-fluorescent staining protocol

in order to visualize and quantify C. phytofermentans cells in plant biomass using epifluorescent

microscopy. After the protocol was developed, we analyzed the correlation of cell growth and

ethanol production in different plant feedstocks. We found that C. phytofermentans cell growth

cycle varies depending on the cellulose substrate. With filter paper, cell growth and ethanol

production was strongly correlated. However, no correlation was found between cell growth and

ethanol production in either wild-type sorghum or the reduced lignin double mutant strain

production in the presence of lignocellulosic substrate. The dual-staining protocol allows for a

simple and efficient method for visualizing and quantifying microbial growth on cellulosic

substrate.

The second research project was the first to characterize and visualize the bacterial

microcompartments (BMCs) when a microbe is grown in fucose and rhammose and produces 1,2-

propanediol (PDU) and propionate. BMCs are protein polyhedral bodies that encapsulate different

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types of metabolic pathways. Their functions vary depending on the type of environment the

microorganism is found. We were able to visualize BMCs inside of C. phytofermentans by doing

thin-section transmission electron micrograph. In the dehydration and polymerization process we

modified the typical TEM embedding protocol by adding potassium ferrycianide and picrid acid.

These compounds, although it is not known exactly how, were able to enhance the contrast of

BMCs and stability the cell membrane when the thin-section occurred. Understanding different

BMCs will increase our knowledge of the different metabolites produced and their role. This

project also determined that the employment of potassium ferrycianide and picrid acid allows for

higher image contrast and a better preservation of the integrity of cell membrane.

The third research project generated a bioenergetics model of membrane energy

conservation by characterizing membrane bound ATPases, a ferredoxin oxidoreductase Rnf-

complex and a membrane bound pyrophosphatase. Using protein homologs and sequence

alignments, we were able to generate 3D representative structures of a Na+ F-type ATPase, a Na+

V-type ATPase and a Na+/H+ pumping pyrophosphatase along with their predicted ion binding

sites. We also found that C. phytofermentans cell growth gets inhibited in the presence of either

proton or sodium ionophores, indicating that the microbe relies on both ions for cell growth. We

also employed a ferredoxin reduction assay to generate a coupling system an ATPase and the Rnf-

complex. The coupling assay was inhibited in the presence of the proton ionophore but it was

enhanced in the presence of sodium ionophore. Our results indicate that C. phytofermentans uses

both H+ and Na+ ions for the conservation of energy through the membrane. This research is the

first one every that studies the energy conservation through the membrane of a biofuels microbe

and proposes an energy pathway through the membrane that is link to the production of ethanol.

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I demonstrated that visualization and quantification of a microbe growing in different plant

biomass substrates can be achieved with the use of two fluorescent dyes. To my knowledge, this

is the first time direct cell count of a microbe in plant biomass substrate. However, this method

has certain limitations. Future versions of this project will be developed to include a transgenic

cell line of C. phytofermentans with a GPF tag. With this process we will only have to rely on one

fluorescent dye, calcofluor white, and minimize photo bleaching effects and dye interaction with

biofilm, making cell quantification and visualization faster. This project would also expand into

examining C. phytofermentans growth in different types of plant biomass such as sugarcane

bagasse, different temperatures and in an agitated environment.

With the modification of the fixation, dehydration and embedding protocol from a TEM

biological sample we achieved the first high contrast imaging of BMCs generated in the presence

of fucose and rhmanose. While precise visualization of BMCs is achieved with this method, it is

very difficult to quantify and visualize BMCs inside of a cell due to two-dimensional limitations

of TEM micrographs. In order to better understand the development of BMCs inside of C.

phytofermentans a future project will consist in generating a transgenic cell line of C.

phytofermentans where a fluorescent tag can be placed in the BMCs. This process will allow us to

further examine the dynamics of BMC generation during cell growth phases and to quantify the

amount of BMCs per cell. The present C. phytofermentans BMC project can also be expanded to

modify certain pathways within the BMCs to achieve higher production of 1,2-propanediol and

propionate.

The development of a bioenergetics model is a highly iterative process. The model was

built using both genomic and biochemical data and it has shed light on how different membrane

bound proteins and its ions contribute to the formation of ethanol. While the model was built with

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complementary genomic and biochemical data, additional information of other proteins and their

genes is necessary to improve its accuracy. Future versions of the model should include

biochemical analysis of the H+/Na+ pyrophosphatase in different pH conditions. Moreover, a

modification in the rnfB section that can hinder the electron transfer of the Rnf-complex should

provide us a better understanding of its NADH production contributes to ethanol formation.

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