production of functional recombinant chloroform reductive

1

PRODUCTION OF FUNCTIONAL RECOMBINANT CHLOROFORM REDUCTIVE DEHALOGENASE

Rabeya Rahmatullah

A thesis in fulfilment of the requirements for the degree of

Doctor of Philosophy

School of Biotechnology and Biomolecular Sciences Faculty of Science

The University of New South Wales Sydney, Australia

October 2020

I

Abstract Organohalides are recalcitrant, toxic environmental pollutants. The presence of these

compounds at elevated levels in soil and aquifers has led to a focus on methods to

remediate these sites. Reductive dehalogenase enzymes (RDases) found in organohalide

respiring bacteria (OHRB) utilise organohalides as electron acceptors for cellular energy

and growth, producing lesser-halogenated compounds that may be more biodegradable

and less toxic. Consequently, microbial reductive dehalogenation via organohalide

respiration represents a promising solution for clean-up of organohalide pollutants and

has been successfully applied in bioremediation of contaminated sites. Therefore, an

understanding of the structure–function relationship of RDases is of considerable

interest. Dehalobacter sp. UNSWDHB is an OHRB capable of respiring highly toxic

chloroform (CF) and converting it to dichloromethane (DCM). TmrA has been identified

as the key RDase responsible for this conversion and different strategies for functional

expression of recombinant TmrA is the focus of this thesis.

There have been several efforts to express recombinant RDases in different host

organisms. In this study, TmrA was recovered from inclusion bodies expressed in E. coli

and refolded in the presence of FeCl3, Na2S and cobalamin. TmrA has been previously

expressed in a soluble and functional form in the corrinoid-producing Bacillus

megaterium. The specific activity estimated for the recombinant TmrA was 11-fold

lower than the activity of the native TmrA. In this study different culture conditions were

screened to improve the specific activity of the recombinant TmrA produced in B.

megaterium. The screenings showed that lowering the concentration of xylose for

induction of TmrA expression increased the specific activity. TmrA was then expressed

in a soluble and active form in Shimwellia blattae. Co-expression with two different

chaperone proteins from the original host did not increase the level of soluble

expression however activity assays showed that removing the TAT signal from TmrA

increases the dechlorination activity compared to when the TAT signal is present. Finally,

TmrA was expressed in a soluble and active form in the H2-oxidizing C. necator H16, a

II

novel host for the expression of RDases. In summary, a range of strategies to

successfully produce recombinant TmrA are presented in this thesis

III

Acknowledgements I would like to express my deepest appreciation to my Supervisor, Christopher Marquis.

Thank you for taking me on as your student even though I came from an entirely

different academic background. I struggled so much with my health and life in general

during my stay in Sydney, it would not have been possible for me to continue my PhD

journey without a kind and compassionate Supervisor like you. Thank you for being so

understanding and patient with me all the time. I am grateful for your endless trust and

motivation which has made my time in your lab a wonderful experience. And special

thanks for your constant supply of chilies throughout the years!

I am deeply indebted to Hélène Lebhar, an awesome mentor, and an amazing friend.

Thank you for all your help in the lab, for being patient with me when I was making silly

mistakes. Thanks for being my shoulder to cry on and my go to person when I needed a

hug!

I would like to thank all the awesome lab mates I have had the pleasure of working with

the past four years. Priyanka, Bat, Miriam, Natalie, Carolin, Nga, Eve and Ismat, you have

all made my time in and out of the lab a fun and enjoyable one. Thanks for all the love

and care you have so generously showered me with. Special thanks to Keaghan for

teaching me all I know about PowerPoint and for always being there to listen to my crazy

rambling when I was having difficult days. I am super grateful to Sam for keeping me

sane during the final months of my PhD. Thank you for all your help, encouragement,

and hugs! I am also indebted to Hasti for helping me format my thesis.

I would like to thank the Australian Government for funding my studies through the

Endeavour Postgraduate Scholarship. I would like to express my gratitude to UNSW for

paying the tuition fee gap and the Postgraduate Research Student Support which funded

my attendance at the FEMS conference in Scotland. I would also like to thank the School

of BABS for offering the BABS Postgraduate Travel Fund and Top-Up Scholarship.

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And at last but not the least, I would like to thank my family for their continuous love

and support throughout my journey. I would especially like to thank my sisters, Radia

and Raisa, for being my rock and my biggest cheerleaders!

V

List of Abbreviations BM Bacillus megaterium

bp base-pairs

cDNA complementary DNA

CF (or TCM) chloroform (or trichloromethane)

CHAPS 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate

CN C. necator

CV column volume

GC gas chromatography

DCM dichloromethane

DEAE diethyl-aminoethyl

DNase deoxyribonuclease

DTT dithiothreitol

%dO2 dissolved oxygen concentration

EDTA ethylenediaminetetraacetic acid

EI electron impact

EPR electron paramagnetic resonance

FF fast flow

FGN fructose-glycerol-nitrogen

FN fructose-nitrogen

IPTG isopropyl-β-D-thiogalactopyranoside

kb kilo base pairs

kDa kilo dalton

KPi potassium phosphate

g relative centrifugal force RCF

h hour

HGT horinzontal gene transfer

VI

LB Luria broth

LC-MS/MS liquid chromatography tandem mass spectrometry

MBH membrane bound hydrogenase

MBP maltose binding protein

min minute

MPD 2-methyl-2,4-pentanediol

mRNA messenger RNA

MS mass spectrometry

NAD+ nicotinamide adenine dinucleotide

NADH nicotinamide adenine dinucleotide, reduced

NADPH nicotinamide adenine dinucleotide phosphate

Ni-NTA nickel-nitrilotriacetic acid

OD600 optical density measured at 600 nm wavelength

OHRB organohalide respiring bacteria

OHR organohalide respiration

PCR polymerase chain reaction

RDase (or RdhA) reductive dehalogenase

rdh reductive dehalogenase gene

RH regulatory hydrogenase

RNase ribonuclease

rRNA ribosomal RNA

SDS sodium dodecyl sulphate

SDS-PAGE sodium dodecyl sulphate polyacrylamide gel electrophoresis

SH soluble hydrogenase

TAT twin arginine translocation

TB Terrific broth

TD-PCR touchdown polymerase chain reaction

TF trigger factor

U unit

WCW wet cell weight

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TABLE OF CONTENTS

Abstract .............................................................................................................................. I

Acknowledgements .......................................................................................................... III

List of Abbreviations ......................................................................................................... V

TABLE OF CONTENTS ....................................................................................................... VII

LIST OF FIGURES .............................................................................................................XIV

CHAPTER ONE: Introduction ............................................................................................. 2

1.1 Organohalides: two sides of a coin ........................................................................................... 3

1.2 Chloroform: the good, the bad and the ugly ............................................................................ 5

1.3 Degradation of organohalides: abiotic and biotic ..................................................................... 8

1.4 Organohalide respiring bacteria (OHRB)................................................................................. 13

1.5 Environmental bioremediation and OHRB ............................................................................. 15

1.6 Key enzyme in reductive dehalogenation: reductive dehalogenases (RDases) .................... 17

1.7 Evolution of organohalide respiration and RDases ................................................................ 18

1.8 Structural characteristics of RDases ........................................................................................ 20

1.9 Types of RDases ....................................................................................................................... 23

1.10 Organisation of RDase gene cluster ...................................................................................... 24

1.11 Transcriptional regulation of RDase genes ........................................................................... 26

1.12 Dehalogenation reaction mechanisms of RDases ................................................................ 28

1.13 Electron transport chain in OHRB ......................................................................................... 30

1.14 Corrinoid cofactors in OHRB ................................................................................................. 34

1.15 Substrate range and specificity of RDases ............................................................................ 37

1.16 Challenges in producing native RDases ................................................................................ 39

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1.17 Heterologous expressions of RDases: limitations and successes ......................................... 49

1.18 Dehalobacter UNSWDHB...................................................................................................... 58

1.19 RDase of Dehalobacter UNSWDHB (TmrA) .......................................................................... 59

1.20 Research aims and thesis outline .......................................................................................... 60

1.21 References ............................................................................................................................. 62

CHAPTER TWO: Recovering functional chloroform reductive dehalogenase from

inclusion bodies in Escherichia coli ................................................................................. 80

2.1 INTRODUCTION ....................................................................................................................... 81

2.2 EXPERIMENTAL PROCEDURES ................................................................................................ 83

2.2.1 Bioreactor fermentation ................................................................................. 83

2.2.2 SDS-PAGE ........................................................................................................ 84

2.2.3 SDS-MPD based system .................................................................................. 84

2.2.4 Aerobic purification and anaerobic cofactor reconstitution method ............ 86

2.2.5 Dechlorination activity assay .......................................................................... 87

2.3 RESULTS ................................................................................................................................... 89

2.3.1 Large-scale heterologous production of TmrA in E. coli ................................. 89

2.3.2 Recovery and refolding of TmrA from inclusion bodies: SDS-MPD based system

................................................................................................................................. 90

2.3.3 Aerobic purification and anaerobic cofactor reconstitution method ............ 92

2.4 DISCUSSION ............................................................................................................................. 95

2.5 CONCLUSIONS ......................................................................................................................... 98

2.6 REFERENCES ............................................................................................................................ 99

CHAPTER THREE: Production of functional recombinant chloroform reductive

dehalogenase in Bacillus megaterium: improving existing findings ............................. 101

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3.1 INTRODUCTION ..................................................................................................................... 102

3.2 EXPERIMENTAL PROCEDURES .............................................................................................. 104

3.2.1 Preliminary screening ................................................................................... 104

3.2.2 Second screening: full factorial design ......................................................... 105

3.2.3 SDS-PAGE ...................................................................................................... 106

3.2.4 Western blot analysis ................................................................................... 106

3.2.5 Determination of protein concentration ...................................................... 107

3.2.6 Dechlorination activity assay ........................................................................ 107

3.2.7 Large-scale culture and cell lysis ................................................................... 107

3.2.8 Purification of NHis-TmrA ............................................................................. 108

3.3 RESULTS ................................................................................................................................. 109

3.3.1 Primary screening of culture conditions ....................................................... 109

3.3.2 Second screening of culture conditions ....................................................... 111

3.3.3 Purification of NHis-TmrA ............................................................................. 116

3.4 DISCUSSION ........................................................................................................................... 118

3.5 CONCLUSIONS ....................................................................................................................... 122

3.6 REFERENCES .......................................................................................................................... 123

CHAPTER FOUR: Heterologous expression of chloroform reductive dehalogenase TmrA

in Shimwellia blattae: co-expression with chaperone proteins ................................... 126

4.1 INTRODUCTION ..................................................................................................................... 127


4.2.1 Plasmid construction .................................................................................... 130

X

4.2.2 Preparing competent S. blattae cells and transformation ........................... 135

4.2.3 Heterologous expression of CF reductive dehalogenase in S. blattae ......... 137

4.2.4 Cell lysis ......................................................................................................... 138

4.2.5 Dechlorination activity assay ........................................................................ 138

4.2.6 SDS-PAGE ...................................................................................................... 138

4.2.7 Transcript analysis ........................................................................................ 139

4.3 RESULTS ................................................................................................................................. 141

4.3.1 Anhydrotetracycline-inducible expression of TmrA in S. blattae ................. 141

4.3.2 IPTG-inducible expression of TmrA in S. blattae .......................................... 145

4.3.3 Co-expression of TmrA with chaperone proteins ......................................... 147

4.3.4 Transcript analysis ........................................................................................ 150

4.3.5 Dechlorination activity of TmrA expressed in S. blattae .............................. 153

4.4 DISCUSSION ........................................................................................................................... 154

4.5 CONCLUSIONS ....................................................................................................................... 157

4.6 REFERENCES .......................................................................................................................... 158

CHAPTER FIVE: Cupriavidus necator as a chloroform reducing organism: a molecular

biology approach ........................................................................................................... 161

5.1 INTRODUCTION ..................................................................................................................... 162


5.2.1 C. necator H16 strain confirmation .............................................................. 164

5.2.2 Chloroform and dichloromethane tolerance test ........................................ 164

5.2.3 Vitamin B12 uptake assay .............................................................................. 165

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5.2.4 Expression of native Ni-Fe hydrogenase in C. necator ................................. 166

5.2.5 Cell lysis ......................................................................................................... 167

5.2.6 Hydrogenase activity assay ........................................................................... 167

5.2.7 Hydrogenation-dechlorination activity assay ............................................... 167

5.2.8 Gene cloning ................................................................................................. 168

5.2.9 Transformation of C. necator ........................................................................ 172

5.2.10 Recombinant expression of TmrA in C. necator ......................................... 174

5.2.11 Cell lysis ....................................................................................................... 175

5.2.12 SDS-PAGE .................................................................................................... 175

5.2.13 Dechlorination activity assay ...................................................................... 175

5.2.14 Transcript analysis ...................................................................................... 176

5.3 RESULTS ................................................................................................................................. 177

5.3.1 C. necator chloroform and dichloromethane tolerance test ....................... 177

5.3.2 C. necator vitamin B12 uptake assay ............................................................. 179

5.3.3 Expression of native Ni-Fe hydrogenase in C. necator ................................. 179

5.3.4 Hydrogenation-dechlorination activity assay using cell lysates of C. necator

and B. megaterium ................................................................................................ 180

5.3.5 IPTG-inducible expression of TmrA in C. necator ......................................... 181

5.3.6 Rhamnose-inducible expression of TmrA in C. necator................................ 182

5.4 DISCUSSION ........................................................................................................................... 185

5.5 CONCLUSIONS ....................................................................................................................... 189

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5.6 REFERENCES .......................................................................................................................... 190

CHAPTER SIX: General discussion and concluding remarks .......................................... 196

6.1 Reductive dehalogenases in organohalide respiration ........................................................ 197

6.2 Escherichia coli: the recombinant protein expression factory ............................................. 198

6.3 Bacillus megaterium: the big beast ....................................................................................... 198

6.4 Shimwellia blattae: the vitamin B12-producing gut bacteria from cockroach ...................... 199

6.5 Cupriavidus necator: the “knallgas” bacteria ........................................................................ 200

6.6 Comparing the four expression systems .............................................................................. 201

6.7 Standout findings of this study .............................................................................................. 203

6.8 Future research ..................................................................................................................... 204

6.9 References ............................................................................................................................. 206

Appendices ...................................................................................................................... 208

A.1 Vector Maps .......................................................................................................................... 209

A.2 Sequence of genes used in the experiments ....................................................................... 215

A.2.1 tmrA without TAT-signal .................................................................................... 215

A.2.2 tmrA with TAT-signal ......................................................................................... 216

A.2.3 tf104 ............................................................................................................... 217

A.2.4 tf161 ............................................................................................................... 218

A.3 Composition of culture media and buffers .......................................................................... 219

A.4 Dechlorination activity assay calculation data ...................................................................... 220

A.5 Screening of recombinant B. megaterium for expression of TmrA using an anti-His

monoclonal antibody in a Western blot assay ........................................................................... 224

A.6 LC-MS/MS Results to confirm identity of TmrA in excised gel bands .................................. 229

XIV

LIST OF FIGURES Figure 1.1. Global chlorine cycle ....................................................................................... 4

Figure 1.2. Model of reductive dehalogenase maturation in cells ................................. 21

Figure 1.3. Dimeric structure of PceA from S. multivorans ............................................ 23

Figure 1.4. Organisation of RDase gene clusters ............................................................ 25

Figure 1.5. (a) Organocobalt adduct mechanism ............................................................ 28

Figure 1.6. (b) Radical mechanism for reductive dehalogenation .................................. 29

Figure 1.7. (c) Reductive dehalogenation by cobalt–halogen bond formation .............. 30

Figure 1.8. Putative composition, organisation, and function of a quinone-dependent

electron transport chain during catabolic reductive dehalogenation ............................ 32

Figure 1.9. Hypothetical model of organohalide respiration complex ........................... 34

Figure 1.10. The structures of cobalamin-B12 and norspuedocobalamin-B12 ............... 35

Figure 1.11. Full length amino acid sequence of TmrA................................................... 60

Figure 2.1. TmrA fusion constructs with affinity (NHis and CHis) and solubility tags MBP

(Maltose binding protein) and TF (trigger factor), encoded in pOPIN expression vector

......................................................................................................................................... 82

Figure 2.2. Fermentation of BL21(DE3)pLysS cells transformed with pOPIN-TmrA-CHis,

pOPIN-TmrA-MBP and pOPIN-TmrA-TF .......................................................................... 89

Figure 2.3. SDS-PAGE analysis of heterologously expressed TmrA fusion proteins ....... 90

Figure 2.4. Purification of TmrA-CHis by affinity chromatography and anion exchange

chromatography .............................................................................................................. 91

Figure 2.5. Ni-NTA column purification of TmrA-MBP .................................................... 91

XV

Figure 2.6. Purification of TmrA-CHis and TmrA-TF by affinity chromatography ........... 92

Figure 2.7. CF dechlorination .......................................................................................... 94

Figure 3.1. TmrA fusion construct used in B. megaterium expression using pPT7 vector

....................................................................................................................................... 103

Figure 3.2. SDS-PAGE of the soluble protein fractions from cell pellets obtained from the

second screening conditions ......................................................................................... 112

Figure 3.3. Dechlorination of CF with B. megaterium cell lysates ................................ 112

Figure 3. 4. Effect of different culture conditions on specific activity .......................... 113

Figure 3.5. Prediction profiler generated with JMP software. ..................................... 113

Figure 3.6. SDS-PAGE of the soluble protein fractions from cell pellets obtained from

cultures with reduced xylose concentrations ............................................................... 114

Figure 3.7. Effect of culture conditions on specific activity .......................................... 115

Figure 3.8. Prediction profiler generated with JMP for induction with different xylose

concentrations. ............................................................................................................. 115

Figure 4.1. tmrAB operon of Dehalobacter sp. UNSWDHB .......................................... 128

Figure 4.2. PCR products in Gibson assembly of tmrA and pASK-IBA63c-plus ............. 142

Figure 4.3. SDS-PAGE analysis of insoluble and soluble fractions of cell lysate of S. blattae

expressing TmrA ............................................................................................................ 143

Figure 4.4. PCR products for Golden Gate cloning and subsequent colony PCR .......... 144

Figure 4.5. SDS-PAGE analysis of soluble fraction of cell lysates of S. blattae expressing

TmrA .............................................................................................................................. 145

Figure 4.6. PCR products used in Gibson assembly and tmrA from colony PCR. ......... 146

Figure 4.7. SDS-PAGE of cell lysate of S. blattae ........................................................... 147

XVI

Figure 4.8. PCR products in Golden Gate assembly of tf104 into vectors pASK-IBA63c-

plus and pBBR1MCS-2 ................................................................................................... 148

Figure 4.9. PCR products in Golden Gate assembly of tf161 into vectors pASK-IBA63c-

plus and pBBR1MCS-2 ................................................................................................... 149

Figure 4.10. SDS-PAGE analysis of S. blattae cells containing TmrA and chaperone

proteins cloned into pBBR1MCS-2 ................................................................................ 149

Figure 4.11. Transcript analysis of tmrA_full gene expressed in S. blattae .................. 150

Figure 4.12. Transcript analysis of tf104 and with tmrA_full genes cloned in pBBR1MCS-

2 vector and expressed in S. blattae ............................................................................. 151

Figure 4.13. Transcript analysis of tf161 and with tmrA_full genes cloned in pBBR1MCS-

2 vector and expressed in S. blattae ............................................................................. 151

Figure 4.14. Transcript analysis of tf104 and with tmrA_full genes cloned in pASK-IBA63c-

plus vector and expressed in S. blattae. ....................................................................... 152

Figure 4.15. Transcript analysis of tf161 and with tmrA_full genes cloned in pASK-IBA63c-

plus vector and expressed in S. blattae ........................................................................ 152

Figure 4.16. Dechlorination of CF to DCM by S. blattae strains extracts ...................... 153

Figure 4.17. The different plasmid constructs and S. blattae transformants used in the

expression tests............................................................................................................. 155

Figure 5.1. Layout of the Deutz microplates with different concentrations of CF and DCM

for tolerance test. ......................................................................................................... 165

Figure 5.2. OD600 of C. necator grown with various concentrations of chloroform (CF) at

different time points. .................................................................................................... 178

Figure 5.3. OD600 of C. necator grown with different concentrations of dichloromethane

(DCM). ........................................................................................................................... 178

XVII

Figure 5.4. Vitamin B12 uptake assay ............................................................................ 179

Figure 5.5. Reduction of NAD+ to NADH (observed at 340 nm) coupled with H2 oxidation

by soluble hydrogenase from C. necator ...................................................................... 180

Figure 5.6. Transcript analysis of tmrA expressed in C. necator grown in rich media. 181

Figure 5.7. Gene cloning and subsequent colony PCR .................................................. 182

Figure 5.8. SDS-PAGE of rhamnose-inducible expression of TmrA in C. necator ......... 183

Figure 5.9. CF dechlorination assay by A. C. necator .................................................... 184

Figure A1.1. Vector map of plasmid used in E. coli experiments. ................................ 209

Figure A1.2.Vector map of plasmid used in E. coli experiments. ................................. 210

Figure A1.3. Vector map of plasmid used in E. coli experiments. ................................ 211

Figure A1.4. Vector maps of plasmids used for recombinant expression of TmrA in B.

megaterium .................................................................................................................. 212

Figure A1.5. Vector maps of plasmids used in S. blattae experiments. ....................... 213

Figure A1.6. Vector map of the plasmid used in recombinant TmrA expression in C.

necator. ......................................................................................................................... 214

Figure A.4.1. DCM standard curve for dehalogenation activity calculation. ................ 220

Figure A.4.2. CF standard curve for dehalogenation activity calculation. .................... 221

Figure A.5.1. Western blot images of screening conditions ......................................... 224



Figure A.5.4. Western blot images of screening conditions.. ....................................... 227

XVIII

Figure A.5.5. Western blot image of pre- and post-induction samples from 1 L culture of

B. megaterium. .............................................................................................................. 228

Figure A.1. LC-MS/MS result for insoluble TmrA-Chis expressed in E. coli. .................. 231

Figure A.2. LC-MS/MS result for insoluble TmrA-Chis expressed in E. coli (continued).

....................................................................................................................................... 232

Figure A.3. LC-MS/MS result for insoluble TmrA-MBP expression in E. coli. ................ 233

Figure A.4. LC-MS/MS result for insoluble TmrA-MBP expression in E. coli (continued).

....................................................................................................................................... 234

Figure A.5. LC-MS/MS data for pASK_TmrA expressed in S. blattae (soluble cell fraction).

....................................................................................................................................... 235

Figure A.6. LC-MS/MS data for pASK_TmrA expressed in S. blattae (insoluble cell

fraction). ........................................................................................................................ 235

Figure A.7 LC-/MS/MS data for pBBR_TmrA expressed in S. blattae (insoluble cell

fraction). ........................................................................................................................ 236

XIX

LIST OF TABLES

Table 1.1 Different biological dehalogenation mechanisms and the enzymes involved 10

Table 1.2 Properties of (partially) purified native RDases .............................................. 40

Table 1.3 Purification methods used for native RDases ................................................. 47

Table 1.4 Heterologous expression of RDases ................................................................ 52

Table 3.1 Culture conditions for primary screening: factors and levels ....................... 105

Table 3.2 Culture conditions for second screening: factors and levels ........................ 106

Table 3.3 Primary screening conditions generated by JMP software .......................... 110

Table 3.4 Second screening conditions ......................................................................... 111

Table 4.1 List of primers used in plasmid construction for soluble expression of TmrA

with and without chaperone proteins in S. blattae and subsequent transcript analysis

studies. .......................................................................................................................... 133

Table 4.2 List of S. blattae transformants created and used in recombinant protein

expression experiments ................................................................................................ 136

Table 5.1 Sample and controls used in the Hydrogenation-dechlorination assay ....... 168

Table 5.2 TD-PCR thermal cycling conditions for linearizing pKRrha plasmid .............. 170

Table 5.3 List of primers used in plasmid construction for rhamnose-inducible expression

of TmrA in C. necator and transcript analysis studies ................................................... 171

Table 5.4 Different culture conditions tested for the IPTG-inducible expression of TmrA

in C. necator .................................................................................................................. 174

Table 5.5 Culture conditions for screening of rhamnose-inducible expression of TmrA in

C. necator ...................................................................................................................... 175

1

Table A.4.1 Raw data for standard curve construction ................................................ 220

Table A.4.3 Raw data for dechlorination activity assay for E. coli samples .................. 221

Table A.4.4 Raw data for dechlorination activity assay for B. megaterium samples ... 222

Table A.4.5 Raw data for dechlorination activity assay for S. blattae samples ............ 222

Table A.4.6 Raw data for dechlorination activity assay for C. necator samples ........... 223

2

CHAPTER ONE: Introduction

3

1.1 Organohalides: two sides of a coin

Organohalides are organic compounds containing one or more carbon-halogen covalent

bonds which can be saturated, unsaturated or cyclic and are one of the largest groups

of chemicals found in the environment (Smidt and Vos, 2004). They are important

components of the halogen biogeochemical cycles (Figure 1.1.). From only 24 in 1968,

the number of identified natural organohalides continue to increase at a rate of 100–

200 per year and till 2014 more than 5000 natural compounds have been identified

which are produced by a multitude of living organisms such as bacteria, fungi,

microalgae, sponges, higher plants, insects and animals or by abiotic processes like

volcanoes, forest fire and other geothermal activities Gribble (2009, 2012, 2015). More

than half of the described natural organohalides contain chlorine, almost half contain

bromine, several hundred contain both chlorine and bromine (Field, 2016), 182 contain

iodine (Wang et al., 2014a) and about 30 contain fluorine (Sato, 2013). However,

organohalides have received much attention because of their anthropogenic origin and

widespread use in daily life and large industrial sectors alike. The negatively charged

halogen renders organohalides thermal and chemical stability and makes them suitable

to be used as refrigerants, water disinfectants, pesticides, solvents, plastics, propellants,

fire retardants and extinguishers, pharmaceuticals and much more. It is estimated that

chlorine is required at some stage of production in 85% of the pharmaceuticals produced

globally (Sasson, 2009). The highest amounts of organohalides produced in the industry

are chloroethanes, chloroethenes, chlorobenzenes, chlorophenols and polychlorinated

biphenyls (Horton et al., 2002).

Organohalides are produced in great quantities; global production of chlorinated

compounds was 56 million metric tonnes in 2010, production of brominated and

fluorinated compounds in 2008 were 0.563 and 5.6 million metric tonnes respectively

(UNEP, 2013). However, organohalides are recalcitrant pollutants and their massive

usage, improper handling and disposal poses threat to human health and the

environment. Because of their high persistency and mobility, organohalides have even

been detected in areas like the Arctic region and Mount Everest where such synthetic

chemicals have never been used (Fawell and Hunt, 1988). Alarming levels of

4

anthropogenic organohalides have been identified in deep ocean environments which

depicts the extent of organohalide pollution (Jamieson et al., 2017).

Figure 1.1. Global chlorine cycle adapted from Atashgahi et al., (2018). Solid and dotted lines represent biological and chemical processes, respectively.

In the most recent list of 126 Priority Pollutants by US EPA, 72 (57%) are organohalides

(EPA, 2014) and more than half of the 1337 identified National Priority List (NPL) sites

are contaminated by organohalides (EPA, 2016). Their contamination ranges from

groundwater to ozone layer depletion (Velders et al., 2007). The persistence of

organohalides depends on the number, type and position of the halogen substituents

and the higher degree of halogenation results in higher biological recalcitrance (Leys et

al., 2013). The halogen moiety of organohalides contribute to their increased lipid

solubility which increases the possibility of membrane/fat tissue incorporation resulting

in bioaccumulation (Häggblom and Bossert, 2003). This facilitates their movement

through the food chain (El-Shahawi et al., 2010) resulting in increasing levels of

organohalides in human and wildlife (Johnson-Restrepo et al., 2005, Solomon and Weiss,

2002). This can lead to the damage of their endocrine, reproductive, immune and

5

nervous systems, leading to allergies, cancer and even death (Lee et al., 2006). Many

organohalides are classified as carcinogens or suspected carcinogens and long-term

exposure may result in developmental defects and chronic illnesses in human (El-

Shahawi et al., 2010). Hence the production and sale of many organohalides are banned

worldwide (Wang et al., 2016) and it is no surprise that now there is more interest in the

destruction of organohalides than their creation. The clean-up of these compounds from

contaminated sites is now a priority.

1.2 Chloroform: the good, the bad and the ugly

Chloroform, also known as trichloromethane (TCM), is a colourless, volatile, non-polar

liquid with a density of 1.48 kg m-3 and is slightly soluble in water. It is produced both

naturally and anthropogenically and is considered as the most abundant organohalide

in the atmosphere (Cappelletti et al., 2012). It is produced naturally in certain species of

fungi, termites, marine macro and microalgae, in soil and peat (Grøn et al., 2012,

Haselmann et al., 2000a, Haselmann et al., 2000b, Hoekstra et al., 1998, Khalil et al.,

1990, Nightingale et al., 1995, Scarratt and Moore, 1999, Simmonds et al., 2010). The

global estimate of atmospheric release of chloroform is 700–820 Gg y−1 of which 90% is

natural in origin (Gribble, 2009, Laturnus et al., 2002), the largest source being offshore

seawater followed by soil processes (McCulloch, 2003). The current emission rate from

anthropogenic sources is estimated to be 60–73 Gg y−1 (Field, 2016).

Chloroform revolutionized medical science and was considered as the major anaesthetic

agent for about a century from its introduction in 1847 till after the Second World War

(Davidson et al., 1982). But its use as an anaesthetic ceased in the early 20th century

because of the fatalities rising from its toxicity (Hutcheon, 2010). Chloroform was widely

used in drug products like cough syrups, antihistamines and decongestants but because

of its proven carcinogenicity in laboratory animals the U.S. Food and Drug

Administration banned its use as an additive in human drug and cosmetic products in

1976 (Rosenthal, 1987). Based on the evidence, EPA classified chloroform as a probable

human carcinogen, International Agency for Research on Cancer classified it as a

possible human carcinogen (Group 2B), and National Toxicology Program described it as

6

a substance that may reasonably be anticipated to be carcinogenic in humans. Its

application as an industrial solvent, however, increased twice as much in 1990 than in

1980 (Cappelletti et al., 2012). Nowadays, more than 90% of the total production of

chloroform is used in the manufacture of refrigerant chlorodifluoromethane (HCFC-22)

while the remaining chloroform has diverse applications in production of

fluoropolymers (e.g. polytetrafluoroethylene, fluorinated ethylene-propylene and

others), pesticide formulation, industrial solvent and adhesive, fire extinguishers, dry

cleaning spot remover, intermediate in the preparation of dyes.

Chloroform can be released into the environment as a consequence of its manufacture,

use and water chlorination practices; important sources being pulp and paper mills,

water treatment plants, chemical manufacturing plants, and waste incinerators

(McCulloch, 2003). Spills and leaks from storage and waste sites may result in

chloroform entering water and soil. Abiotic and biotic dechlorination of carbon

tetrachloride which is another common industrial solvent also causes accumulation of

chloroform at contaminated sites (Grostern et al., 2010). Because of inappropriate

disposal practices, chloroform is a pollutant of concern in all developed countries where

it is produced in bulk amounts, particularly USA, Europe and Japan (Rossberg et al.,

2006). It is in the recent list of 126 priority pollutants by EPA, 11th in the CERCLA Priority

List of Hazardous Substances and present in 351 out of 1337 pollutant sites of the

National Priority List in the USA (EPA, 2016). Chloroform is a recalcitrant environmental

pollutant which under purely hydrolytic conditions has an estimated half-life of 3100

years (Mabey and Mill, 1978). When exposed to the environment, chloroform gradually

leaches from soil to groundwater because of its high density, low soil adsorption and

water solubility and remain as subsurface solvent pools known as dense non-aqueous

phase liquid (DNAPL) within subsurface water systems (Lee et al., 2012). Chloroform is

also an inhibitor of microbial processes essential for biogeochemical cycling such as

methanogenesis (Weathers and Parkin, 2000) and organohalide respiration of

chlorinated ethanes and ethenes (McMurdie et al., 2011). Groundwater contaminated

with less than 2.5 mM (0.3 mg l-1) chloroform can inhibit microbial dechlorination of

chlorinated ethenes and cause accumulation of partially dechlorinated toxic

intermediates like cis-dichloroethene and vinyl chloride (Bagley et al., 2000, Duhamel et

7

al., 2002). Perchloroethene (PCE) dechlorination by organohalide respiring bacteria can

be completely inhibited by less than 1 mg l-1 of chloroform (Maymó-Gatell et al., 2001).

Perchloroethene and trichloroethene contaminated sites are frequently observed to be

co-contaminated by chloroform (Grostern et al., 2010). Thus, bioremediation of

contaminated sites where chloroform is also present may require prior removal of the

chloroform (Bagley et al., 2000).

Co-metabolic transformation of chloroform by methanogenic, acetogenic and sulfate-

reducing cultures have been observed (Bouwer and McCarty, 1983, Chung and

Rittmann, 2007, Chung and Rittmann, 2008, Freedman et al., 1995, Guerrero‐Barajas

and Field, 2005, Gupta et al., 1996, Koons et al., 2001, Olivas et al., 2002, Weathers and

Parkin, 1995). However, these processes require costly primary growth substrates such

as volatile fatty acids, methanol or dichloromethane and in some cases the addition of

extraordinarily high amounts of cyanocobalamins or other reduced metallo-coenzymes

are necessary (Becker and Freedman, 1994, Guerrero‐Barajas and Field, 2005), which

renders the process expensive. Metabolic or growth-linked transformation is thus

preferred over co-metabolic transformation for bioremediation because of its higher

efficiency and sustainability (Grostern et al., 2010). Dehalobacter sp. CF,

Desulfitobacterium sp. PR and Dehalobacter sp. UNSWDHB are organohalide respiring

bacteria which have been reported to be capable of chloroform respiration by utilizing

chloroform as terminal electron acceptors in respiration (Jugder et al., 2016b) and have

potential for bioremediation of chloroform (co-)polluted sites (Ding et al., 2014,

Grostern et al., 2010, Lee et al., 2012).

8

1.3 Degradation of organohalides: abiotic and biotic

Resistance to chemical and biological degradation is a quality that has made

organohalides useful in industrial applications but at the same time is also the reason

that makes them recalcitrant environmental pollutants (Häggblom and Bossert, 2003).

Despite their recalcitrance, degradation of these compounds does occur in natural

environments. Removal of halogens from these organohalides produce dehalogenated

compounds that are generally less toxic and have less probability to bioaccumulate

which makes them more susceptible to further microbial breakdown. However, it is

crucial to mention that sometimes the degradation product of dehalogenation can be

even more toxic than their precursors; for example, a degradation product of TCE or PCE

by TceA reductive dehalogenase from Dehalococcoides mccartyi 195 is vinyl chloride

(VC) which has a higher toxicity than the precursor compounds (Duhamel et al., 2002).

Spontaneous abiotic dehalogenation reactions usually happen through chemical

reduction by redox coupled reactions where concentration of reactive minerals is high

(Häggblom and Bossert, 2003). Naturally occurring minerals such as iron sulphide,

pyrite, magnetite and green rusts can degrade organohalides abiotically (Butler and

Hayes, 1999, Tobiszewski and Namieśnik, 2012). Abiotic transformation can also be

mediated by zerovalent iron, corrinoids and chemical oxidation using permanganates or

persulfates (Arnold and Roberts, 2000, Elsner et al., 2008, Glod et al., 1997, Hrapovic et

al., 2005, Krone et al., 1989, Tsitonaki et al., 2010). The degradation product of abiotic

dechlorination by β-elimination is acetylene which is considered to be benign to the

environment (Liang et al., 2007).

One of the key factors that determines the fate of organohalides in the environment is

their microbial degradation and a critical step to that is the cleavage of carbon-halogen

bonds (Häggblom and Bossert, 2003). The natural production and anthropogenic release

of organohalides to the environment has probably been the driving force for the

evolution of an unusually high microbial capacity to dehalogenate different classes of

organohalides (Smidt and Vos, 2004). Microbes generally use halogenated compounds

in four ways: as a carbon source and oxidisable substrate; in fermentative metabolism;

as an electron acceptor in respiration; and in cometabolic transformation related to

9

metabolic processes (Janssen et al., 2001). The inclination of microbes to use

organohalides as energy sources for growth is dependent on their energetic properties

such as bond strength, ionic potential, redox potential and Gibbs free energy of

formation. The use of microbes as the means of remediation is considered more cost

effective and sustainable than other mechanisms and hence this approach has been

studied extensively which has led to the detection and elucidation of a variety of

dehalogenases and dehalogenation mechanisms (Fetzner, 1998, Janssen et al., 1994). A

summary of different dehalogenation mechanisms is represented in Table 1.1.

10

Table 1.1 Different biological dehalogenation mechanisms and the enzymes involved

Name Definition Reaction Mechanism Enzymes involved

Hydrolytic dehalogenation

A halogen on an organic molecule is replaced by a hydroxyl group (Sims et al., 1991)

R-X + H2O → R-OH + HX 2-Haloacid

dehalogenase, haloalkane

dehalogenase

Thiolytic dehalogenation

Chlorine atoms are displaced by the nucleophilic attack of the thiolate anion of glutathione which is catalysed by glutathione S-transferases (Wang et al., 2010)

Glutathione S-transferase

Intramolecular substitution

Intramolecular nucleophilic substitution of a halogen by an adjacent hydroxyl that functions in halohydrins to yield epoxides (van Pée and Unversucht, 2003)

Haloalcohol dehalogenases

11

Dehydrohalo-genation

Elimination of hydrochloric acid causes the formation of a double bond (Imai et al., 1991)

Dehydrohalogenase

Dehalogenation by hydration

Addition of water to the double bond that yields an unstable intermediate from which malonate semialdehyde is formed (van Hylckama Vlieg and Janssen, 1991)

cis-3-chloroacrylicacid dehalogenase, trans-3-chloroacrylicacid dehalogenase

Dehalogenation by methyl transfer

The methyl group of chloromethane is transferred onto tetrahydrofolate and inorganic chloride; and methyl tetrahydrofolate are released (Meßmer et al., 1996)

CH3Cl + FH4 → CH3-FH4 + H+ +Cl-

Methyltransferase

Oxidative dehalogenation

Incorporation of either one or both oxygen atoms from O2 into the substrate (Häggblom and Bossert, 2003)

Monooxygenase, dioxygenase, peroxidase

12

Reductive dehalogenation

The removal of a halogen substituent from a molecule with simultaneous addition of electrons to the molecule (Mohn and Tiedje, 1992). There are two types of reductive dehalogenation:

i. Hydrogenolysis: formation of one mole of HX per two electrons

ii. Dehaloelimination: two moles of HX are formed per two electrons where the presence of two vicinal halogens results in the formation of an unsaturated C=C bond

i. RX + H+ → RH + X-

ii.

Reductive dehalogenases

13

1.4 Organohalide respiring bacteria (OHRB)

The occurrence of a large number of naturally synthesized organohalogens suggest that

natural rather than the anthropogenic sources of organohalides governed the

emergence of organohalide respiring bacteria (OHRB). This is also supported by the

global distribution of dehalogenating bacteria in polluted as well as pristine

environments (Atashgahi et al., 2018). OHRB use organohalides as a terminal electron

acceptor in anaerobic respiration and conserve energy from this process (Leys et al.,

2013). The mechanism of energy conservation via reductive dehalogenation is an

unresolved mystery (Buckel and Thauer, 2013). OHRB use molecular hydrogen or other

oxidizable compounds as electron donors and organohalides as electron acceptors in

the respiratory chain where none of the participating compounds themselves provide

energy but yields from the electric potential difference between the participating redox

couples (Adrian and Löffler, 2016). The redox potentials of most of the organohalides

range between +410 to +470 mV. Considering that 70 kJ is required to generate one

mole of ATP in a living cell, such an energy yield would likely result in the formation of

2– 2.5 ATP molecules per molecule of chloride ion released (Schink and Friedrich, 1994).

Hence biomass yields from organohalide respiration are generally low and depending

on the bacteria and the substrate used, may range from 0.3 to 5.6 g dry weight per mole

of chloride produced (Ding et al., 2014). This is potentially because only two moles of

H+ per mole of H2 oxidised are released and three moles of H+ are required to generate

sufficient proton-motive force to produce one mole of ATP (Schumacher and Holliger,

1996).

Organisms that could use organohalides as electron acceptors in an energy-yielding

process were found in the 1980s and Desulfomonile tiedjei was the first isolated OHRB

(DeWeerd et al., 1990). Since then, there has been major advances in sequencing

technology and till February 2020, the number of OHRB genome accessible via

NCBI/Genbank database has reached 75. The average genome size ranges from 2.6 to

3.1 Mb with an average GC content of about 0.44 to 0.45 (Richardson, 2013). OHRBs are

strict anaerobes that are slow-growing and are sensitive to pH and light. They inhabit a

wide range of terrestrial and aquatic environments and the currently known OHRB

14

belong to distinct phylogenetic groups of deeply branching Chloroflexi, Gram-negative

Proteobacteria and Gram-positive Firmicutes (Adrian and Löffler, 2016). To date,

bacteria belonging to the genera Anaeromyxobacter, Comamonas, Geobacter,

Dehalobacter, Dehalococcoides, Dehalogenimonas, Desulfitobacterium, Desulfoluna,

Desulfomonile, Desulfovibrio, Desulfuromonas, Shewanella, and Sulfurospirillum have

shown experimental evidence of reductive dehalogenation (Ahn et al., 2009, Chen et al.,

2013, Christiansen and Ahring, 1996, Holliger et al., 1998a, Krumholz, 1997, Lohner and

Spormann, 2013, Moe et al., 2009, Sanford et al., 2002, Sun et al., 2001, Sung et al.,

2003).

Depending on whether organohalide respiration is their only energy-gaining metabolism

or not, the known OHRB can be divided into facultative and obligate groups (Maphosa

et al., 2010). The facultative OHRB have a more versatile metabolism, growing on a wide

range of electron acceptors, and include Geobacter, Desulfuromonas,

Anaeromyxobacter, Desulfomonile, Desulfovibrio, Desulfoluna, Sulfurospirillum,

Comamonas, Shewanella and Desulfitobacterium (Adrian and Löffler, 2016). On the

other hand, obligate OHRB are restricted to organohalide respiration for energy

conservation and growth and include Dehalococcoides, Dehalogenimonas, and

Dehalobacter (Adrian and Löffler, 2016). The niche specialization of the two groups of

OHRB is also visible in the range of electron donors used; while the facultative OHRB can

use various electron donors, usually the organic end products of primary fermenters,

along with H2 and formate; the obligate OHRB are almost always restricted to H2. In

general, organisms with a diverse metabolism have larger genomes than organisms

inhabiting more restricted ecological niches. Complying with this concept, the obligate

organohalide-respiring D. mccartyi and D. lykanthroporepellens have very small

genomes with an average size of 1.4 Mbp (which is amongst the smallest found in free-

living bacteria) (Kube et al., 2005), whereas facultative OHRB generally have larger

genomes ranging from 3.2 to 6.5 Mbp (Kruse et al., 2016). Furthermore, genome

sequencing reveals that in general, obligate OHRB have large number of genes encoding

reductive dehalogenase homologues (11 to 36 in number) while facultative OHRB carry

a lower number of reductive dehalogenase-encoding genes, only one to seven for

currently available OHRB genomes (Kruse et al., 2016).

15

1.5 Environmental bioremediation and OHRB

The widespread use and disposal of organohalides have rendered them the most

common groundwater contaminants throughout the developed world. Hence, they

have become a primary focus of remediation since the 1980s (Steffan and Schaefer

2016). Early active remediation approaches involved technologies such as pump and

treat or air sparging (Honetschlägerová et al. 2019). In the early 1990s bioremediation

emerged as a potential alternative because of the low cost, low energy consumption

and low environmental impact. Initial bioremediation approaches involved aerobic

cometabolic treatments but the discovery of OHRB brought about a revolutionary

change in bioremediation of organohalide-contaminated sites (Steffan and Schaefer

2016). In situ bioremediation by enhanced reductive dehalogenation has become a

widely applied remediation approach and is accomplished by biostimulation and/or

bioaugmentation to stimulate microbial reductive dehalogenation by organohalide

respiration (Atashgahi et al. 2017).

Biostimulation is possibly the most widely used approach for bioremediation of

organohalide-contaminated aquifers. Biostimulation promotes the growth and activity

of dechlorinating microbes in contaminated environments by nutrient supplementation

and/or adjustment of environmental conditions. In most cases the materials added are

selected to provide a source of H2 and the most common approach is the addition of a

carbon source that can be fermented to H2. A wide variety of organic compounds (e.g.

ethanol, lactate, butyrate, propionate, formate, cyclodextrin, gamma poly-glutamic

acid) have been used to provide energy and electron donors for the dehalogenation

processes (He et al. 2003, Azizian et al. 2008, Wu et al. 2016, Blanford et al. 2018, Sheu

et al. 2018). Direct injection of H2 and electrolytic production of H2 in situ have also been

applied (Ma et al. 2006, Lohner and Tiehm 2009, Lohner et al. 2011, Zhang et al. 2001).

Water-soluble substrates like sodium lactate, molasses, alcohols and glycerol, and a

range of insoluble substrates like emulsified vegetable oils, polylactates, and plant-

based materials are being marketed as commercial electron donor products (Steffan and

Schaefer 2016). Recently thermally enhanced in situ biostimulation has also been

proposed as a sustainable bioremediation strategy for chlorinated ethenes (Slenders et

16

al. 2010, Ni et al. 2016). Němeček et al., (2018) demonstrated substantial improvement

of chlorinated ethene degradation by the combination of increased temperature and

supplementation with a fermentable substrate (whey). Zero-valent iron (ZVI) has a high

reduction potential capable of inducing various dechlorination pathways to reduce the

accumulation of toxic by-products like vinyl chloride and chemical reduction using ZVI

has become one of the most commonly used in situ remediation technologies for

organochlorides (Fu, Dionysiou, and Liu 2014, Guan et al. 2015, Han et al. 2016). The

combination of ZVI and biostimulation is now being considered to be a promising

bioremediation strategy for organochlorines (Sheu et al. 2016, Yang et al. 2018, Wu et

al. 2020).

Bioaugmentation is the practice of adding cultured microbes to speed up the rate of

contaminant degradation. Bioaugmentation studies with D. mccartyi has demonstrated

complete dehalogenation of chlorinated compounds in situ (Harkness et al. 1999, Ellis

et al. 2000, Major et al. 2002, Lendvay et al. 2003, Schaefer et al. 2010). In the United

States alone, more than 1000 applications of bioaugmentation cultures have been

carried out to increase reductive dehalogenation rates and prompt groundwater

remediation (Steffan and Schaefer 2016). Biostimulation together with

bioaugmentation with dechlorinating enrichment cultures containing D. mccartyi

populations and their non-dechlorinating partners has also been showed to be a

successful strategy (Rahm et al. 2006, Perez-de-Mora et al. 2014).

Successful biostimulation and bioaugmentation strategies rely upon molecular tools for

qualitative and quantitative assessment of the dechlorinating bacterial community

(Lebron et al. 2011). Monitoring the presence and abundance of specific biomarker

genes has become a valuable procedure for remediation strategy (Ritalahti et al. 2006,

Holmes et al. 2006, Munro et al. 2017, Nijenhuis, Stollberg, and Lechner 2018, Němeček

et al. 2018). To date, microbial monitoring pertaining to bioremediation have mainly

focused on quantitative PCR (qPCR) based tracking of key OHRB and their rdhA genes

(Lendvay et al. 2003, Scheutz et al. 2008, Lee et al. 2008, Scheutz et al. 2010, Ritalahti et

al. 2010, Révész et al. 2014) and occasionally of related non-dechlorinating microbial

groups (Perez-de-Mora et al. 2014). Recently the potential of developing inexpensive

17

and field deployable detection devices using the novel molecular method called Loop

mediated isothermal amplification (LAMP) has been explored (Stedtfeld et al. 2014,

Kanitkar et al. 2016, Stedtfeld et al. 2016, Kanitkar, Stedtfeld, Hatzinger, Hashsham, and

Cupples 2017, Kanitkar, Stedtfeld, Hatzinger, Hashsham, Cupples, et al. 2017).

1.6 Key enzyme in reductive dehalogenation: reductive dehalogenases (RDases)

The common feature of the broad range of OHRB is the presence of the reductive

dehalogenases. They are a group of metalloprotein enzymes that catalyse reductive

dehalogenation and are the only known terminal respiratory enzymes to be dependent

on a corrinoid cofactor (Dobbek and Leys, 2016). Unlike other corrinoid-dependent

enzymes, reductive dehalogenases do not catalyse carbon rearrangement or methyl

group transfer reactions, rather they catalyse an elimination reaction which makes them

a novel class of cobamide-containing oxidoreductases (Banerjee and Ragsdale, 2003). In

2017 the Nomenclature Committee of the International Union of Biochemistry and

Molecular Biology (NC-IUBMB) classified RDases as EC 1.21.99.5 under the sub-subclass

EC 1.97.1 for oxidoreductases that do not belong in the other subclasses.

The physiologically active mature respiratory RDases are associated with the

cytoplasmic membrane and are located in the exoplasm of the bacterial cells (periplasm

in Gram-negative bacteria) (John et al., 2006, Nijenhuis and Zinder, 2005, Reinhold et

al., 2012); though studies reveal that in some respiratory RDases the membrane

association is weak (John et al., 2006, Suyama et al., 2002). Most of the respiratory

RDase enzymes are known to be oxygen-sensitive with a half-life of approximately one

hour to several hours (Schubert and Diekert, 2016) but oxygen insensitivity has also been

reported for respiratory (Goris et al., 2014) and non-respiratory RDases (Chen et al.,

2013, Payne et al., 2015). Studies demonstrate that pure enzymes are active over a

temperature range of 20 to 55°C but have the highest specific activity at 37-42°C

(Maillard et al., 2003, Neumann et al., 1996) and have the ability to catalyse reactions

at pH 5.5-8.7 with optimum activities at 7.2 to 8.1 (El Fantroussi et al., 1998).

18

1.7 Evolution of organohalide respiration and RDases

Organohalides generally occur naturally in low concentrations in diverse environments,

which explains the widespread distribution of OHRB. rdh genes have been identified

from most natural environments as well as from bacteria associated with other

eukaryotes, even humans (Brouwer et al., 2011). The phylogenetic depth (Hug et al.,

2013b), molecular dating (McMurdie et al., 2011), and distribution of rdh genes deep in

the sub-seafloor (Kawai et al., 2014) suggest that organohalide respiration is an

evolutionary old process. However, the origin of organohalide respiration is still not well

understood; there is evidence in support of the contribution of both vertical and

horizontal gene transfer (HGT) towards the distribution of rdh genes across taxonomic

groups. The distribution of the rdh gene family is not even across the prokaryotic tree of

life, for instance RDases are only present in Dehalococcoidia class but not in any other

members of the phylum Chloroflexi (Hug et al., 2013a, Kaster et al., 2014, Wasmund et

al., 2014). This interspersed distribution of rdh genes makes it difficult to determine if

RDases represent an ancient enzyme family that exhibited a strong pattern of gene loss

through prokaryotic evolution or followed a more taxonomically limited adaptation and

spread through gene duplication and gene transfer (Hug, 2016).

Horizontal gene transfer can be an important evolutionary mechanism for

microorganisms exposed to toxic organohalides to adopt acquired catabolic pathways

from even phylogenetically distinct species to adapt and thrive in contaminated

ecosystems (Jugder et al., 2016a). The available evidence on HGT in rdh genes implies

that it exists widely among various genera of OHRB (Krajmalnik-Brown et al., 2007,

Maillard et al., 2005, McMurdie et al., 2007, McMurdie et al., 2011). Multiple

mechanisms of HGT have been proposed, including transposons, insertion elements,

genomic islands, recombinases and phage-mediated gene movement (Hug, 2016,

Jugder et al., 2016a).

Most of the rdh genes in Dehalococcoides mccartyi strains are located in the two ‘high

plasticity regions’ at either side of the origin of replication, separated by a highly

conserved core region common to all strains (Kube et al., 2005, McMurdie et al., 2009).

These regions serve as evolutionary hotspots with frequent horizontal gene transfers

19

assisting in rapid evolution of D. mccartyi strains, although no mechanism has been

identified yet (McMurdie et al., 2009). Some D. mccartyi genomes harbor prophages

that may play a role in HGT (Pöritz et al., 2013, Zhao et al., 2017). Five out of 24

sequenced strains of D. mccartyi encode CRISPR-Cas (clustered regularly interspaced

short palindromic repeats—CRISPR) systems that protect the host from invading mobile

elements like plasmids, phages, and transposons (Molenda et al., 2019). The role of

these phages and CRISPR-Cas systems in assisting or hindering lateral transfer of rdh

genes among D. mccartyi strains is yet to be identified. HGT has also been reported in

Dehalobacter strains which are also metabolically restricted OHRB (Kruse et al., 2013,

Maphosa et al., 2012). D. lykanthroporepellens BL-DC-9T has 25 rdh genes and 13 of

them are adjacent to insertion sequence elements which implies horizontal acquisition

from an unknown host or hosts (Mukherjee et al., 2014, Siddaramappa et al., 2012).

Fascinating evidence for HGT of rdh genes was the identification of a circularised

transposon containing two identical insertion sequences (ISDha1) surrounding the

pceABCT gene cluster in D. hafniense strain TCE1 (Maillard et al., 2005). The pceABCT

gene cluster of D. hafniense TCE1 shares 100 % sequence identity with those from D.

restrictus and Desulfitobacterium strains PCE-S and Y51, suggesting a more recent

horizontal acquisition rather than vertical inheritance (Maillard et al., 2005). The

distribution of rdh genes in facultative OHRB such as Comamonas, Geobacter and

Shewanella, belonging to phylogenetic groups that mostly comprise non‐OHRB has been

attributed to HGT (Atashgahi et al., 2016). Another example of HGT in opportunistic

OHRB is the loss of OHR capacity of strains of Desulfitobacterium when grown on

alternative electron acceptors. It is suggested that they acquire rdh genes from closely

related obligate OHR strains of Dehalobacter via HGT (Duret et al., 2012). G. lovleyi strain

SZ contains mobile genetic elements such as integrase and transposase genes close to

rdh operon which points towards potential horizontal gene transfer events (Sanford et

al., 2016).

20

1.8 Structural characteristics of RDases

Despite having high sequence variability RDases have similar biochemical features.

RDases generally consist of two subunits: RdhA and RdhB (Futagami et al., 2008). RdhA

is the catalytically active subunit with a molecular weight of ~50-67 kDa (Magnuson et

al., 2000, Neumann et al., 1996, Neumann et al., 1998, van de Pas et al., 1999). All

characterized RDases contain two [4Fe–4S] clusters (either two 4Fe–4S or one 4Fe–4S

and one 3Fe–4S) (Maillard et al., 2003) that are considered to be associated with

electron transfer from the physiological donor to the active site (Fincker and Spormann,

2017). Apart from the single exception of the 3-chlorobenzoate RDase of D. tiedjei,

which contains a heme cofactor (Ni et al., 1995), all RDase enzymes purified and

characterized to date have been shown to harbor a cobalt-containing porphyrin-derived

corrinoid cofactor which is inferred to play an important role in substrate activation and

reduction (Fincker and Spormann, 2017).

RDases have a twin-arginine (TAT) signal sequence (RRXFXK) at the N-terminus (Berks,

1996, Bruschi and Guerlesquin, 1988). This signal is required for the recognition of the

precursor by the TAT export machinery in the membrane (Palmer and Berks, 2012b).

The membrane-integral TAT translocase exports folded and in most cases cofactor-

containing proteins across the cytoplasmic membrane. The two iron–sulfur clusters and

the corrinoid cofactor of the RDases are synthesized in the cell cytoplasm, catalyzed by

a specific set of enzymes and are transferred into the RDase apoprotein possibly in a

fully assembled form (Figure 1.2) (Schubert and Diekert, 2016). The correct folding and

consequent TAT-dependent export of the enzymes is dependent upon the proper

incorporation of the cofactors (Sargent, 2007) and impairment of the proper cofactor

biosynthesis generally leads to the accumulation of the enzyme precursor in the

cytoplasm followed by the degradation of the unfolded apoprotein (Siebert et al., 2002).

The second subunit RdhB (8-10 kDa) is a hydrophobic protein with two to three

transmembrane helices. The rdhB gene can be located either upstream or downstream

of the rdhA gene. The RdhB proteins have 66 to 116 amino acids with minimal amino

acid sequence similarity. However, all of them encode relatively hydrophobic proteins

suggesting that they may have similar three-dimensional structures (Neumann et al.,

21

1998, van de Pas et al., 1999). Most RdhBs have two conserved glutamic acid residues

in the exoplasmic loop which might play role in the RdhA-RdhB interaction (Schubert et

al., 2018). RdhB is believed to function as a membrane anchor for the RdhA, though

whether that is their sole function or not is yet to be determined (Schubert et al., 2018).

Figure 1.2. Model of reductive dehalogenase maturation in cells (adapted from Schubert and Diekert, 2016). RR is the twin arginine motif of the Tat signal peptide, cob/cbi are corrinoid biosynthesis genes, isc/suf are iron sulfur cluster biosynthesis genes.

All RDases were thought to be monomeric until the crystal structure of PceA from

Sulfurospirillum multivorans revealed its dimeric structure (Bommer et al., 2014). In

contrast, the NpRdhA structure is clearly monomeric. Both crystallized RdhAs contain

two 4Fe–4S clusters less than 10 A° apart and are located close to the surface of the

enzymes; the proximal cluster (relative to the corrinoid) is in direct van derWaals

interaction with the edge of the corrin ring. Norpseudovitamin-B12 is the cofactor of

22

PceA and it is deeply buried inside the internal substrate binding pocket (Figure 1.3). The

corrinoid cofactor for NpRdhA is cobalamin B12 and in both the RDases the cobalamin is

bound in the same base-off conformation. The coordinative binding of the lower bases

is restricted by steric hindrance (Keller et al., 2018). Spectroscopic analyses of other

RDases using electron paramagnetic resonance spectroscopy also showed the cobamide

cofactor in the base-off state (Schumacher et al., 1997, van de Pas et al., 1999). The

structural analysis of the substrate-bound enzyme PceA revealed that the substrate,

TCE, accesses the active site only via a narrow “letterbox” opening followed by a

hydrophobic channel that leads to a tight substrate-binding pocket (Bommer et al.,

2014). The shape, access channel and most of the residues that line the active site are

distinct for the two RdhA structures. This demonstrates the evolutionary distance

between these enzymes and the contrast in their substrate specificity. PceA reduces

small, hydrophobic chlorinated ethenes whereas NpRdhA reduces aromatic

dibromophenols. There are, however, a few residues that are conserved; a tyrosine is

found to point directly towards the cobalt ion in both the RdhA structures. The phenol

group is within hydrogen bonding distance of a second conserved and positively charged

residue (Lys in Pce; Arg in NpRdhA). This RdhA Tyr-Lys/Arg motif is most probably

involved in proton transfer coupled to substrate reduction and can be identified in most

RdhA proteins. In NpRdhA, mutations in either residue suppress enzyme activity (Payne

et al., 2015). Structural studies showed no evidence of involvement of the cobamide

cofactor’s nucleotide loop in the catalytic cycle of RDases, rather it plays a role in

cofactor binding. RDases bind the cofactor with a network of hydrogen bonds deep

inside the protein hence, the varied structure of the cobamide nucleotide loop might

interfere with the incorporation and correct positioning of the cofactor (Keller et al.,

2018).

Interestingly, despite the lack of any obvious sequence similarity both the enzymes

demonstrated the highest structural resemblance with a human B12-processing enzyme

CblC, which is responsible for catalysing the reduction of cyanocobalamin and

alkylcobalamin to form adenosyl- or methylcobalamin (McMurdie et al., 2011). The

similarity in fold and cofactor position as detected from a structural superimposition all

strongly suggest a common ancestry between RDases and CblC.

23

Figure 1.3. Dimeric structure of PceA from S. multivorans (adapted from Bommer et al., 2014). The grey and green regions represent two monomers of PceA. The spheres depict the proximal and distal Fe-S clusters and the purple sticks represent the cobalamin cofactor, norpseudo-B12.

1.9 Types of RDases

As described in the previous section, respiratory and non-respiratory RDase enzymes

have the same cofactor content and principal architecture, however, the non-

respiratory RDases lack the TAT motif and are located in the cytoplasm (Chen et al.,

2013, Payne et al., 2015). The suggested functions for these non-respiratory RDases

encompass re-oxidation of respiratory cofactors, detoxification of substrates, removal

of halogens from organics to enable subsequent catabolism (Chen et al., 2013, Payne et

al., 2015, Wasmund et al., 2016).

Recently a third type of potential RDase has been identified in marine uncultured

Desulfatiglans-related deltaproteobacterium, which is possibly a hybrid of RdhA and

RdhB (Jochum et al., 2018). The putative rdh gene encoding this third type of RDase has

no TAT signal peptide, no associated rdhB and has an exclusive feature of encoding three

transmembrane helices (TMHs) in the N-terminal region (Jochum et al., 2018). The C-

24

terminus carries the two binding motifs for Fe-S clusters and faces the inner side of the

cytoplasmic membrane, which is likely due to the absence of the TAT signal peptide

(Atashgahi, 2019). Like most RdhBs, the third putative RDase has a similar short

cytoplasmic loop between helix 1 and 2 that contains the two conserved glutamic acid

residues; the cytoplasmic localization of the C-terminus of the putative RDase may

facilitate such an interaction with this loop, which leads to the prediction of the putative

RDase being a hybrid of RdhA and RdhB (Atashgahi, 2019). So far three putative hybrid

rdh genes have been Identified in Dethiosulfatarculus sandiegensis, Desulfatiglans SAG2

and strain NaphS2 (Jochum et al., 2018); but genomic studies suggest that many other

similar genes exist in the genomes of pure cultures along with metagenome-assembled

genomes (Atashgahi, 2019).

1.10 Organisation of RDase gene cluster

Despite their phylogenitically diverse origin, reductive dehalogenases share conserved

domains and motifs of rdh genes (Jugder et al., 2015b). The minimal rdhAB gene cluster

is composed of the rdhA genes encoding for the catalytic subunit and the rdhB gene

coding for the membrane anchor protein and is frequently accompanied by a variable

set of accessory genes, the exact functions of most of them are still unknown (Magnuson

et al., 2000, Neumann et al., 1998, van de Pas et al., 1999). Although there is evidence

of several solitary rdhA genes, the presence of an associated rdhB gene warrants the

involvement of the encoded RdhA protein in OHR (Key et al., 2017, Key et al., 2016, Low

et al., 2015, Maillard and Willemin, 2019). There are significant variations in the

composition of rdh gene clusters among OHRB which is closely related to the phylogeny

of OHRB; obligate OHRB show lower diversity in terms of accessory genes in the genome

than facultative OHRB (Maillard and Willemin, 2019).

Apart from the genes encoding for the catabolic subunit and the membrane anchor

protein, the most commonly occurring accessory genes are the three different types of

transcriptional regulators (CRP/FNR, MarR, two-component system) and rdhT. The

transcriptional regulators are described in the following section. rdhT is transcribed in a

monocistronic manner and it encodes a trigger-factor like protein, RdhT, which is

25

proposed to act as a chaperone protein for proper folding of the RdhA apo-protein and

its subsequent exportation to the periplasm via the TAT secretion pathway (Morita et

al., 2009). The absence of RdhT during heterologous expression of RdhAs may explain

the difficulties encountered when attempting expression of soluble and active RdhA

(Morita et al., 2009). In some cases, the rdhT gene is adjacent to the rdhAB operon (such

as in D. hafniense Y51), whereas in other OHRBs, it is located elsewhere in the genome

and is not found in any rdhA related operons; for example, the tmrAB operon of

Dehalobacter sp. UNSWDHB (Figure 1.4).

Figure 1.4. Organisation of RDase gene clusters (adapted from Jugder et al., 2015). Examples of different organisations of gene clusters for five species of OHRB are presented here.

26

1.11 Transcriptional regulation of RDase genes

Understanding the transcriptional regulation of genes involved in OHR is of great

interest to researchers because of the potential it presents as a promising alternative to

uncover the substrate specificity of the reductive dehalogenases that are yet to be

characterized. In the past two decades there has been over 70 studies focusing on the

transcriptional regulations of rdh genes and they reveal that rdh gene transcription

patterns follow two distinct trends: they either respond strongly to specific

organohalides (e.g. Desulfitobacterium sp.) or transcription occurs independently

regardless of the organohalides present/absent (e.g. Dehalococcoides and

Dehalogenimonas sp.) (Maillard and Willemin, 2019).

Transcriptional regulators belonging to three distinct protein families: CRP/FNR, MarR,

two-component system (TCS), have been identified in close vicinity of many rdh gene

clusters, suggesting a tight regulation of their expression. CRP/FNR like regulators are

found in Dehalobacter sp. and Desulfitobacterium sp. (Kruse et al., 2016),

Sulfurospirillum sp. have regulators belonging to TCS (Goris et al., 2017, Goris et al.,

2014) whereas Dehalococcoides sp. and Dehalogenimonas sp. harbor a combination of

TCS and MarR-type regulators (Siddaramappa et al., 2012, Wagner et al., 2013).

The CRP/FNR family was named after the cyclic AMP (cAMP) receptor protein (CRP) and

the fumarate nitrate regulatory protein (FNR). RdhK is used as a general name to

describe this type of regulators involved in OHR (Kruse et al., 2016). RdhK regulators are

characterized by a sequence of 230–250 amino acids, an effector-binding domain

located at the N-terminal and a C-terminal helix-turn-helix (HTH) DNA-binding motif

(Kruse et al., 2016). The regulatory function is carried out by the binding of allosteric

effector molecules to the N-terminal domain followed by a signal transmission to the

DNA-binding domain (Körner et al., 2003). To date, experimental evidence has been

gained for only a small number of RdhK regulators such as CprK from D. dehalogenans

strain JW/IU-DC1; CprK1, CprK2 and CprK4 from D. hafniense strain DCB-2 (Maillard and

Willemin, 2019). Crystal structures of CprK and CprK1 proteins have been revealed and

has improved understanding on the nature of the interactions and the conformational

changes that occur upon binding of the effector (Joyce et al., 2006, Levy et al., 2008).

27

MarR-type regulators are made of a winged helix-turn-helix (HTH) motif; they mediate

contact between the dimeric regulator and short palindromic sites in the target DNA

(Wilkinson and Grove, 2006). MarR-type regulators for aromatic compound metabolism

act as repressors, the repressor is released from the promoter upon binding of the

aromatic ligand and leads to the induction of the regulated gene(s) (Wagner et al., 2013).

Several marR homologous genes are found in the vicinity of rdh gene clusters in

Dehalococcoides sp. (Wagner et al., 2013) and D. lykanthroporepellens (Siddaramappa

et al., 2012). RdhR was proposed as a general term to represent MarR-type proteins

involved in the regulation of OHR (Kruse et al., 2016).

A typical two-component system is composed of a sensory histidine kinase (HK)

component and a response regulator (RR) component. The HK is capable of auto-

phosphorylation on a conserved histidine residue, the phosphoryl group is then

transferred to a conserved asparagine residue on the RR component which drives

conformational change in the RR protein to activate the output response (Jacob-

Dubuisson et al., 2018). The HK and RR components of TCS potentially involved in OHR

are referred to as RdhS and RdhP, respectively (Kruse et al., 2016). TCS regulators have

been identified in the direct vicinity of rdh gene clusters in Sulfurospirillum sp.,

Dehalococcoides sp. and in D. lykanthroporepellens (Maillard and Willemin, 2019). The

HK proteins in D. mccartyi strains CBDB1 and 195 are reported to be cytosolic (Kube et

al., 2005, Seshadri et al., 2005) but those in Sulfurospirillum sp. are predicted to be

membrane-associated (Goris et al., 2014).

Other than the functionally characterized regulatory proteins, many putative proteins

have also been observed in OHRB genomes. The reductive dehalogenase C subunit

(RdhC) of the Rdh complex, found in all classes of OHRB, are homologous to NirI/NosR-

type transcriptional regulators and is composed of five transmembrane domains, a flavin

mononucleotide-binding domain and a C-terminal polyferredoxin-like domain which

leads to the prediction of being involved in transcriptional regulation (Cuypers et al.,

1992, Futagami et al., 2008, Wagner et al., 2013).

28

1.12 Dehalogenation reaction mechanisms of RDases

The complete catalytic cycle of the corrinoid-dependent reductive dehalogenation is

enigmatic up to date. It has been generally accepted that cob(I)alamin is the reactive

species at the active site of an RDase. Three different reaction mechanisms for RDases

involving Co(I) as a low-potential electron donor or nucleophile, have been proposed: a)

Organocobalt adduct mechanism, b) Radical mechanism and c) Cobalt-halide adduct

mechanism. The three mechanisms are briefly explained using PCE as an example.

In mechanism (a), a nucleophilic attack of Co(I) to the halogenated carbon forms an

organocobalt adduct [trichlorovinyl-Co(III) corrinoid] resulting in a halide elimination

(Krasotkina et al., 2001). Fe–S cluster delivers an electron leading to the reduction of the

organocobalt adduct and formation of a trichlorovinyl anion, which, when protonated,

results in TCE as the end product. The transfer of a second electron from an Fe–S cluster

to Co(II) regenerates Co(I) (Figure 1.5). Computational and mechanistic studies of PCE

dehalogenation with corrinoids in solution where organocobalt adducts have been

identified, has led to the development of this mechanism (Lesage et al., 1998, McCauley

et al., 2003). However, it has not been supported experimentally.

Figure 1.5. (a) Organocobalt adduct mechanism (adapted from Fincker and Spormann, 2017).

Mechanism (b) is demonstrated in Figure 1.6, where Co(I) serves as an electron donor

which leads to the formation of radical anion followed by trichlorovinyl radical and the

elimination of a chloride ion (Holliger et al., 1998b). The radical is then further reduced

by an electron transferred from the Fe-S cluster and subsequently protonated to

generate TCE. The Co(II) is reduced back to Co(I) by an electron from other Fe-S clusters.

A variation of this mechanism has also been proposed where before being converted to

29

TCE, the radical anion intermediate is first protonated to a trichloroethane radical

(Fincker and Spormann, 2017). The presence of a radical intermediate is yet to be

supported by experimental evidence, however, there has been enzymatic studies where

a mechanism of Co(I) donating an electron to form a radical species leading to

dehalogenation has been observed (Schmitz et al., 2007, Ye et al., 2010).

Figure 1.6. (b) Radical mechanism for reductive dehalogenation (adapted from Fincker and Spormann, 2017).

A third dehalogenation pathway (mechanism (c), Figure 1.7) has been proposed based

on the EPR spectroscopy of ortho-bromophenol RDase of N. Pacificus which suggests a

direct interaction of the cobalt ion with the halogen substituent in the substrate (Payne

et al., 2015). The reaction starts with the attack of the Co(I) on the halide of the

substrate, leading to the formation of a cobalt–halogen bond, the exact chemical nature

of which is still unknown. The carbon-halogen bond is then cleaved either homolytically

or heterolytically. In the heterolytic cleavage mechanism protonation yields the

dehalogenated product and a transient Co(III)–halogen adduct. The Co(III)–halogen

adduct is then reduced to Co(I) via Co(II)–halogen and a halide by two electrons from

the Fe–S cluster(s). Alternatively, homolytic cleavage of the carbon-halogen bond causes

the formation of a radical and a Co(II)–halogen adduct. Reduction of the radical results

in a trichlorovinyl anion, which after protonation forms the dehalogenated product. A

second electron transfer from the Fe-S cluster releases the halide from the Co(II)–halide

adduct and regenerates Co(I). (Payne et al., 2015) suggested that depending on the

chemical structure of the substrate, both variations of the cobalt-halide adduct

30

mechanism could exist. Aromatic substrates would likely follow the homolytic cleavage

while heterolytic cleavage would be favoured for aliphatic substrates. So far neither

route has been proven experimentally, however, the formation of a cobalt–halogen

bond is supported by the data from an electron density modelling study in D. mccartyi

CBDB1 suggesting that the first step of reductive dehalogenation is the transfer of an

electron from Co(I) to an apical halogen site (Cooper et al., 2015). No mechanism is yet

clearly conclusive and more structural and mechanistic studies are needed to

understand the mechanism(s) of reductive dehalogenation in OHRB and the relationship

with OHR

Figure 1.7. (c) Reductive dehalogenation by cobalt–halogen bond formation (adapted from Fincker and Spormann, 2017).

1.13 Electron transport chain in OHRB

In OHRB electrons are transferred from H2 or other organic compounds to organohalides

via membrane-associated electron transport chains (Löffler and Edwards, 2006, Mohn

and Tiedje, 1992). This process may couple with oxidative phosphorylation to support

the cell growth of OHRB; however, biological reductive dehalogenation is not necessarily

linked to chemiosmosis. Genomic and proteomic studies reveal that there is no uniform

organization of electron transfer chains among the diverse OHRB, rather diverse modes

of coupling reductive dehalogenation to proton translocation have evolved (Schubert et

al., 2018). The electron transfer paths can be classified into (i) those using quinones for

electron and proton shuttling within the membrane and (ii) those with a quinone-

independent electron transfer chain.

31

There is strong evidence that in many OHRB quinones have an important role as electron

carriers to RDase. The presence of genes encoding the enzymes of menaquinone

biosynthesis and/or different menaquinones have been identified in D. restrictus, S.

multivorans, D. tiedjei, D. dehalogenans (Dairi et al., 2011, Goris et al., 2014, Holliger et

al., 1998a, Kruse et al., 2013, Kruse et al., 2015, Louie and Mohn, 1999b, Scholz-

Muramatsu et al., 1995). Reduced 2,3-dimethyl-1,4-naphthoquinone, a menaquinone

analog, could be used to drive PCE reduction in cell suspensions of D. restrictus

(Schumacher and Holliger, 1996). An inhibitory effect of quinone antagonist 2-n-heptyl-

4-hydroxyquinoline N-oxide (HQNO) has been observed in D. restrictus, S. multivorans,

D. tiedjei (Louie and Mohn, 1999b, Scholz-Muramatsu et al., 1995, Schumacher and

Holliger, 1996). UV spectroscopy revealed that membrane-bound menaquinones were

reduced with the addition of H2 and re-oxidized upon the addition of PCE (Schumacher

and Holliger, 1996). Similar results were obtained in D. dehalogenans with the addition

of dithionite and 3-chloro-4-hydroxyphenylacetate (Kruse et al., 2015). However, the

mode of electron transfer from (mena)quinone pool to RDase remains unanswered.

Proteomic analysis of S. multivorans has identified a putative quinol dehydrogenase in

proximity to the gene encoding PceA (Goris et al., 2014). Similar quinol dehydrogenase

encoding genes have been identified in proximity of RDase genes in S. halorespirans,

Sulfurospirillum sp. SL2–1 and D. tiedjei (Cao et al., 2016, Goris et al., 2017).

The midpoint redox potential of menaquinone (−74 mV) (Thauer et al., 1977) is

considerably higher than the Co(II)/Co(I) redox couple (about −370 mV) of the enzyme-

bound cobamide cofactor in the RDase, which gives rise to the question “how do

electrons at the quinol level reduce Co(II) in the corrinoid cofactor to the Co(I) state?”.

Two mechanisms have been suggested to explain the generation of low redox potential

electrons from quinols: a reverse electron flow driven by the proton gradient (Miller et

al., 1996) and electron bifurcation (Buckel and Thauer, 2013, Peters et al., 2016).

Electron bifurcation is the split of an electron pair by flavoproteins into one electron

having a more negative reduction potential and the other with a more positive reduction

potential than that of the electron pair (Buckel and Thauer, 2018). RdhC, which has been

identified in the genomes of a range of OHRBs and harbors a conserved sequence motif

for covalent binding of flavin mononucleotide (FMN) cofactor, has been suggested to

32

play a role in electron bifurcation (Buttet et al., 2018). A postulated composition of a

quinone-dependent electron transport chain is shown in Figure 1.8.

Figure 1.8. Putative composition, organisation, and function of a quinone-dependent electron transport chain during catabolic reductive dehalogenation (adapted from Fincker and Spormann, 2017).

Menaquinone is the electron carrier coupling H2 oxidation to organohalide reduction.

The reduction of menaquinone with electrons derived from H2 by a hydrogenase

consumes cytoplasmic protons. The quinol-oxidizing component is still unknown. For

thermodynamic considerations metabolic energy in the form of a reverse electron flow

(REF) or a quinone-based electron bifurcation may be involved (red arrow). Black and

red arrows depict exergonic and endergonic reactions, respectively.

Unlike Desulfitobacterium, Sulfurospirillum, and Dehalobacter species, there is no

evidence for the presence of quinones or genes encoding quinones in Dehalococcoides

and Dehalogenimonas species (Key et al., 2017, Key et al., 2016, Kube et al., 2005, Kublik

et al., 2016, Moe et al., 2009, Molenda et al., 2016, Schipp et al., 2013, Seshadri et al.,

2005, Siddaramappa et al., 2012). Moreover, 1,2,3-trichlorobenzene reductive

dehalogenation activity in D. mccartyi CBDB1 was not inhibited by HOQNO, nor was OHR

driven by reduced quinones (Jayachandran et al., 2004; Kublik et al., 2016). Also, the

presence of protonophores did not inhibit the dehalogenation activity in D. mccartyi

strains CBDB1 and 195 (Jayachandran et al., 2004, Nijenhuis and Zinder, 2005). These

33

findings strongly support the concept of a quinone-independent respiratory chain in D.

mccartyi and Dehalogenimonas species.

Of the five different putative multi-subunit hydrogenases encoded in D. mccartyi strains

CBDB1 and 195 (Hup, Ech, Hyc, Hym and Vhu), only Hup hydrogenase is predicted to be

localized in the periplasm and to be responsible for transmitting electrons from H2 to

the organohalide-respiratory chain (Kube et al., 2005, Seshadri et al., 2005). The Hup

hydrogenase operon consists of four genes: the large [NiFe]-containing catalytic subunit

(HupL), the small membrane-bound subunit (HupS), the Fe–S cluster-containing subunit

(HupX), and a gene predicted to be involved in hydrogenase maturation (Fincker and

Spormann, 2017). A complex formation between HupSLX, a RDase, and a complex iron–

sulfur molybdoenzyme (CISM) in D. mccartyi strain CBDB1 has been extracted and

shown to catalyze hydrogen-dependent reduction of 1,2,3-trichlorobenzene (Hartwig et

al., 2017, Kublik et al., 2016). The multi-protein reductive dehalogenase complex has

been referred to as the organohalide respiration complex (OHR complex) (Seidel et al.,

2018). The CISM is predicted to act as a critical electron-channeling module between

Hup hydrogenase and the RDase (Schipp et al., 2013). CISM is composed of two proteins:

the organohalide respiration involved molybdoenzyme (OmeA) and its putative

membrane-integrated anchor OmeB (Schubert et al., 2018). Both the Hup and the CISM

are each missing a subunit that the other contains (Mansfeldt et al., 2014), suggesting a

synergistic interaction. The predicted mechanistic model for the complex is that

electrons enter the complex through HupL and are then transferred via the iron-sulfur

clusters in HupS, HupX and OmeA to the RdhA which finally reduces the organohalides

(Seidel et al., 2018). It is still unknown how protons are translocated across the

membrane in the OHR complex. OmeB has a glutamate residue in helix eight which can

possibly be involved in proton pumping (Zinder, 2016). Another possible mechanism

may be an additional protein such as HppA, interacting with the OHR complex to

generate the proton motif force needed for ATP synthesis (Seidel et al., 2018). A

hypothetical model of the organohalide respiration complex is presented in Figure 1.9.

34

Figure 1.9. Hypothetical model of organohalide respiration complex. This figure adapted from (Seidel et al., 2018). The cubes indicate Fe-S clusters, the rhomboid structure indicates the corrinoid cofactor, the blue and green dots represent molybdenum and nickel ions, respectively.

1.14 Corrinoid cofactors in OHRB

Cobalamins are corrinoids consisting of a contracted tetrapyrrole ring system attached

to a nucleotide loop (Lenhert and Hodgkin, 1961). They vary in their structure by

containing different upper and lower axis ligands. Physiologically functional cobalamins

use a methyl group or a 5′-deoxyadenosyl moiety as the upper ligand and one of the 17

known lower ligands which are purine, benzimidazoles and phenols (Renz et al., 1987,

Rondon et al., 1997, Yan et al., 2018). Cobalamin-containing enzymes bind the cofactor

in two different conformations: the base-on mode where the lower base of the

nucleotide loop establishes a coordinative bond with the central cobalt ion or the base-

off conformation where the lower base is displaced from the cobalt (Banerjee and

Ragsdale, 2003, Bridwell-Rabb and Drennan, 2017, Gruber et al., 2011). The structure of

cobalamin-B12 and norpseudocobalamin-B12 is shown in Figure 1.10.

35

Figure 1.10. The structures of cobalamin-B12 (left) and norspuedocobalamin-B12 (right). The figures were adapted from Ludwig and Matthews 1997 and Keller et al. 2014, respectively. R represents an axial ligand which can be cyano-, methyl-, adenosyl- or water groups. The bound hydrogen atom at position 176 marked by a dashed circle is replaced by a methyl group in pseudo‐B12.

Corrinoids are important cofactors for the growth of OHRB, while some can synthesize

their own e.g. S. multivorans (Goris et al., 2015, Keller et al., 2014), S. halorespirans

(Goris et al., 2017), D. hafniense strains Y51, JH1, Viet1 and PCE1 (Choudhary et al., 2013,

Reinhold et al., 2012, Yan et al., 2018), Geobacter lovleyi (Wagner et al., 2012),

Geobacter sulfurreducens (Yan et al., 2012), Dehalobacter strains TCA1, CF, DCA and

UNSWDHB (Sun et al., 2002, Tang et al., 2016, Wang et al., 2017, Wong et al., 2016);

others are dependent on exogenous supply and hence are known to thrive within

mutualistic anaerobic microbial communities rather than in pure culture (Maphosa et

al., 2012). The lack of intact corrinoid synthesis pathway in many species can possibly be

correlated to the energy cost associated with de novo corrinoid synthesis which involves

more than 30 enzymatic steps (Moore et al., 2013) and thus favours a salvaging strategy

rather than de novo synthesis.

Though the underlying mechanism is still unknown, evidence suggests that disparity in

the structure of cobamide cofactors can influence the reductive dehalogenation of

RDases (Keller et al., 2018, Keller et al., 2014, Yan et al., 2018, Yan et al., 2013, Yan et

36

al., 2012, Yan et al., 2016, Yi et al., 2012). Studies show that the lower ligand are

modulators of corrinoid-dependent enzyme functions such as RDases, thus signifying

the importance of understanding enzyme-specific corrinoid cofactor requirements to

predict and if possible, manipulate their catalytic activity (Degnan et al., 2014, Keller et

al., 2014). Cobamide structure can be regulated by guided biosynthesis which is the

feeding of building blocks such as lower base precursors to the growing cells of a

cobamide-producer (Keller et al., 2018, Keller et al., 2014, Mok and Taga, 2013). Guided

cobamide biosynthesis is an effective tool to modulate the structure of cobamide

cofactors in OHRB and to investigate cofactor selectivity of RDases in the native

organism or the host for heterologous production (Schubert et al., 2019).

The obligate OHRB D. mccartyi is a cobamide auxotroph (Löffler et al., 2013). Three types

of corrinoid are functional in D. mccartyi strain 195: 5,6-dimethylbenzimidazolyl-

cobamide, 5-methylbenzimidazolyl-cobamide and 5-methoxybenzimidazolyl-cobamide

(Kruse et al., 2019). However, studies have shown that non-functional corrinoids such

as 5-hydroxybenzimidazolyl-cobamide or 7-adeninyl-cobamide can be remodelled to

functional ones by replacing the lower ligand when 5,6-dimethylbenzimidazole is

provided in the media (Men et al., 2014, Yi et al., 2012). A probable explanation for the

inhibitory effect of specific lower bases on reductive dehalogenation of RDases might be

the incompatibility of the lower base in the cobamide cofactor with the cobamide-

binding site of the particular RDase (Keller et al., 2018).

S. multivorans does not have genes for benzimidazole biosynthesis (Goris et al., 2014)

but it can utilise exogenous dimethylbenzimidazole (DMB) for cobamide biosynthesis.

The presence of DMB in the media can influence the organism's ability to perform

organohalide respiration and elevated levels of DMB can impair the PCE-dependent

growth, PceA activity, and PceA maturation in S. multivorans (Keller et al., 2014). Keller

et al., (2018) showed that S. multivorans can efficiently incorporate exogenously applied

5-methylbenzimidazole (5-MeBza), 5-hydroxybenzimidazole (5-OHBza), and 5-

methoxybenzimidazole (5-OMeBza) as lower bases into norcobamides instead of

adenine which is the original lower base of norpseudo-B12. While 5-OHBza and 5-

OMeBza had no negative effect on the enzyme activity, 5-MeBza reduced the enzyme

37

activity by half, although none of them affected the PCE-dependent growth. Structural

analysis of the norcobamides by NMR spectroscopy revealed a regioselectivity in the

utilization of these precursors for norcobamide biosynthesis.

Guided cobamide biosynthesis experiments in Desulfitobacterium hafniense strain DCB‐

2 showed that reductive dehalogenation of 3‐chloro‐4‐hydroxy‐phenylacetate by RdhA6

and 3,5‐dichlorophenol by RdhA3 was unaffected when exogenous benzimidazoles

(MeBza, Bza, OHBza or OMeBza), azabenzimidazoles (5‐azaBza, 4‐azaBza) and 4,5‐

dimethylimidazole (DMB) were incorporated by the organism into cobamides suggesting

the versatility of both enzymes in cobamide utilisation (Schubert et al., 2019). They also

studied the effect of the alteration of cobamide lower ligand precursors on recombinant

RDases produced in Shimwellia blattae. The recombinant PceAY51 was non-selective

towards different cobamides, however the activity of DcaA of Desulfitobacterium

dichloroeliminans was completely inhibited in cells utilising 5,6‐dimethylbenzimidazolyl,

but considerably increased in cells that incorporated 5‐methoxybenzimidazole into the

cobamide cofactor. This finding suggests that since cobamide cofactor-binding core is

structurally conserved among RDases (Bommer et al., 2014, Payne et al., 2015), it is likely

that only minimal structural changes are sufficient to alter cobamide preferences.

1.15 Substrate range and specificity of RDases

OHRB usually harbor multiple RDases in their genome and the combined substrate

specificities of the RDases encoded in an organism’s genome regulates the range of

halogenated compounds they can utilise as electron acceptors. Characterized RDases

show substrate specificity for different classes of organohalides, including aliphatic

chloro- and bromo- ethenes and ethanes, aromatic polychlorinated and brominated

phenols, and chlorinated benzenes (Fincker and Spormann, 2017). Additionally, the

range of substrates tested have mainly been limited to organohalides present at the

contaminated sites where the microorganisms have been isolated from, so the

possibility of the characterized RDases being capable of utilizing other substrates cannot

be overlooked. Microbial reductive deiodination has also been observed in nature by

Desulfitobacterium chlororespirans (Cupples et al., 2005) and Desulfoluna spongiiphila

38

(Ahn et al., 2009). Dehalococcoides mccartyi strain CBDB1 showed deiodination of

iodinated contrast media (ICM) and 2,3,5- triiodobenzoic acid without correlated cell

growth suggesting decoupling of electron transport from proton translocation (El-

Athman et al., 2019). No RDase tested to date has been observed to dehalogenate

fluorinated organic compounds however, recently an organohalide-respiring microbial

community capable of C‒F bond cleavage via reductive defluorination of two per- and

polyfluoroalkyl substances (PFASs), perfluoro (4-methylpent-2-enoic acid) and 4,5,5,5-

tetrafluoro-4-(trifluoromethyl)-2-pentenoic acid has been reported (Yu et al., 2019).

Substrate specificity of RDases does not depend on their phylogenetic origin or the

similarities between amino acids sequences of RDases. PceA, a tetrachloroethene-

dechlorinating enzyme can be found in five different genera: Shewanella,

Dehalococcoides, Dehalobacter, Sulfurospirillum, and Desulfitobacterium (Lohner and

Spormann, 2013, Magnuson et al., 1998, Maillard et al., 2003, Neumann et al., 1996,

Tsukagoshi et al., 2006). While having an affinity for the same substrate, the PceAs from

these genera share a maximum 29.8 % pairwise identity (Hug et al., 2013b). On the other

hand, CfrA and DcrA enzymes from Dehalobacter strains CF and DCA share 95 %

sequence identity at the amino acid level but their substrate ranges do not overlap (Tang

and Edwards, 2013).

RDases can catalyse dehalogenation of structurally related organohalides at differing

rates, though they usually have high substrate selectivity for only one or two

compounds. Initially it was believed that aliphatic and aromatic organohalides are

dehalogenated by two different types of RDases, however several RDases have showed

the capability to remove halogens from both aromatic and aliphatic halogenated

compounds, such as PcbA1, PcbA4 and PcbA5 from Dehalococcoides mccartyi strains

catalyze chlorine removal from both PCBs and PCE (Wang et al., 2014b). Substrate

specificity in aromatic RDases generally depends on the position of the organohalide

substituent. For example, CprA from Desulfitobacterium sp. PCE-1 and

Desulfitobacterium dehalogenans have a preference for ortho-dechlorination (van de

Pas et al., 2001, van de Pas et al., 1999), while on the other hand, CprA5 from

Desulfitobacterium sp. PCP-1 prefers meta-dehalogenation (Thibodeau et al., 2004).

39

1.16 Challenges in producing native RDases

Reductive dehalogenases can be an important tool for bioremediation of recalcitrant

organohalides. Identification and characterization of reductive dehalogenases is

required for the understanding of their structure, reaction mechanisms and substrate

specificity thus, paving the way to developing effective bioremediation strategies.

Research into how organohalides are used as an energy or carbon source is also

important for deciphering microbial biochemical interactions in the environment

(Schubert et al., 2018). Over 2000 putative reductive dehalogenase genes have so far

been identified but only a handful have been characterized (Adrian and Löffler, 2016). A

single OHRB may have several RdhA enzymes which make it difficult to identify a specific

substrate range for a single rdhA gene product and further hinder the functional

assignment (Mac Nelly et al., 2014, Schubert and Diekert, 2016). It is difficult to predict

substrate specificity based on gene sequence (Hug et al., 2013b) and it has been

concluded that sequence similarity and substrate specificity are generally not correlated

(Buttet et al., 2013). Thus, functional assays require the isolation and purification of the

native enzyme which has so far proven to be very difficult and to date only 20 RDases

have been purified partially or to apparent homogeneity (Table 1.2).

Purification of RDases from the native hosts and the subsequent characterization is

hindered by several factors, the major ones being the slow growth of the OHRB and

associated low cell yield (Hug et al., 2013b), especially the obligate OHRB which cannot

be cultured with alternative electron acceptors. The small cell sizes (<0.5 µm in some

cases), the oxygen sensitivity of the organisms and the low water solubility of the

organohalides serving as the terminal electron acceptors are also important obstacles.

The enzymes themselves also impose restrictions to the process because of their

membrane association, oxygen intolerance and requirement for cobalamin and Fe-S

cluster cofactors, thus making the production of the enzymes from native sources

impractical in large scale (Maphosa et al., 2012).

.

40

Table 1.2 Properties of (partially) purified native RDases

Enzym

e an

d

Acce

ssion

No

.

Host Organism Mo

lar mass

(kDa)

pH

Op

tima

Tem

p

Op

tima (°C

)

Oxygen sensitivity (half-life in minute)

Sub

strate

Km (µM)

Vm

ax (n

mo

l/min

/

mg)

Pro

du

ct (s)

Oth

er

sub

strates

Cofactor (s)

Inh

ibito

rs

References

3CBA-RDase I4C3I3

Desulfomonile tiedjei DCB-1

64 and 37

7.2-7.5

38 Sensitive but not deter-mined

3-CBA ND 71 BA NA Heme ND Ni et al., 1995

PceA W6EQP0_SULMO

Sulfurospirillum multivorans (formerly Dehalospirillum multivorans)

57 8 42 120 PCE 200 158400 c-1,2-DCE

TeIE 1 corrinoid; 2 × [4Fe-4S]

Cyanide; high conc. of 1,2-c-DCE, chloromethanes

Neumann et al., 1996 Ye et al.; 2010

TCE 240 158400 c-1,2-DCE

1,2-DBE 162 55320 VB TBE 49 77520 VB 2,3-DCPE 893 23520 2-CPE 2,3-DBPE 40 81600 NA

PCE-RDase Q3Z9N3

Mixed enrichment culture of Dehalococcoides mccartyi 195 (formerly D. ethenogenesis 195)

51 6.9-7.5


PCE ND 20500 TCE 2,3-DCPh Corrinoid; Fe/S?

Iodo-ethane; sodium cyanide; sodium sulfite; sodium dithionite; cuprous chloride; zinc chloride

Magnuson et al., 1998 Löffler et al. 2013 Fung et al. 2007

TCE-RDase Q3ZAB8

Mixed enrichment culture of Dehalococcoides mccartyi 195 (formerly D. ethenogenesis 195)

61 6.9-7.5


TCE ND 8000-12000

VC NA Corrinoid; Fe/S?

Iodopropane; sodium cyanide; sodium Azide; sodium sulfite; sodium dithionite;

Magnuson et al., 1998 Löffler, et al. 2013

1,1-DCE ND 8000-12000

VC

c-1,2-DCE

ND 12100 VC

41

t-1,2-DCE

ND 4000-6000

VC cuprous chloride; zinc chloride VC ND 4000-

6000 Ethene

PCE-RDase Q848J2

Desulfitobacterium frappieri strain PCE-S

65 7.2 50 50 PCE 10 39000 c-1,2-DCE

NA 1 corrinoid, 2 × [4Fe-4S]

propyl iodide; high conc. of TCE

Miller et al., 1998 Ye et al.; 2010

TCE 4 41400 c-1,2-DCE

1,2-DBE 36 2220 VB and ethene

TBE 8 3960 VB 2,3-DCPE 319 1380 NA 2,3-DBPE 13 4920 NA

3-C-4-HPA- RDase P81594

Desulfitobacterium hafniense

46.5 NR NR 100 3-C-4-HPA

ND 6192 4-HPA ND 1 corrinoid; 12 Fe; 13 S

ND Christiansen et al., 1998

CprA Q9XD04

Desulfitobacterium dehalogenans

48 8.2 52 90 3-C-4-HPA

20 28000 NA 3-CPh; 4-CPh; 2,5-DCPh; 2-F-4-CPh; PCE; TCE

1 corrinoid; 1 × [4Fe-4S]; 1 × [3Fe-4S]

ND van de Pas et al., 1999

2-B-4-CPh

ND 24300 NA

2,3-DCPh ND 15500 NA 2,4-DCPh ND 4200 NA 2,6-DCPh ND 800 NA PentaCPh

ND 200 NA

2-CPh ND 200 NA 3-C-4-HBA-RDase Q8RQC9

Desulfitobacterium chlororespirans

50 6.8 60 77 3-C-4-HBA

12.4 14800 4-HBA TCA; 2,4-DCPh; 2,3,4-TCPh; 2,3,5-TCPh; 2,3,4,5-TeCPh; 2,3,4,6-TeCPh;

1 corrinoid, 2 × [4Fe-4S]

propyl iodide; KCN; sodium sulfite, EDTA; 4-HBA; 3-HBA; 3,6-dichlorosalicylate; 3-CBA; 3,5-DCBA; 2-CPh; 2,4-Dichloropropene; 2-Chloro-3-hydroxyphenol,

Krasotkina et al., 2001 Loffler et al., 1996

3,5-DC-4-HBA

12.3 37900 3Cl-4-HBA, 4-HBA

3-C-4-HPA

310 5500 4-HPA

2,3-DCPh 334 10200 3-CPh 2,6-DCPh 1,46

0 5100 2-CPh

42

2,4,6-TCPh

160 12600 2,4-DCPh, 4-CPh

2,3,5,6-TeCPh

2-chlororesorcinol; 2-Chloro-4-hydroxyphenol, chlohydroquinone; 3,6-Dichloro-2-hydroxybenzoate

PentaCPh

390 1940 2,3,4,5-TeCPh, 3,4,5-TCPh

TeC-BPD ND 120 TC- and DC-BPD

PceA Q93SG6

Clostridium bifermentans DPH1

35.7(monomer)

70(dimer)

7.5-8.0

35 1200 PCE 12 73 c-1,2-DCE

TCE, c-1,2-DCE; t-1,2-DCE, 1,1-DCE; 1,2-DCP; 1,12-TCA

corrinoid propyl iodide Okeke et al., 2001

CPRD Q939S8

Desulfitobacterium sp. PCE-1

48 NR NR Sensitive but not deter-mined

3-C-4-HPA

ND 4.01 4-HPA 2-CPh; 3-CPh; 4-CPh; 2,3-DCPh; 2,4-DCPh; 2,5-DCPh; 2,6-DCPh, PCE, TCE, DCE

Corrinoid Iodopropane; N2O

van de Pas et al., 2001

PCERD Q9APN4

Desulfitobacterium sp. PCE-1

48 NR NR Sensitive but not deter-mined

PCE ND 5.51 TCE 2-CPh; 3-CPh; 4-CPh; 2,3-DCPh; 2,4-DCPh; 2,5-DCPh; 2,6-DCPh, TCE, DCE

Corrinoid Iodopropane; N2O

van de Pas et al., 2001

PCE/TCERD Q8GJ31

Desulfitobacterium sp. TCE-1

59 NR NR Sensitive but not

PCE ND 10.01 TCE 3-C-4-HPA; 2,3-DCPh, DCE

Corrinoid?

Iodopropane; N2O

van de Pas et al., 2001 TCE ND 10.01 DCE

43

deter-mined

PceA Q8L172

Desulfitobacterium sp. Y51

58 7.0-7.5

25-37 330 PCE 105.7

164.4 ± 27.2

c-DCE HCA; PentaCA 1,1,1,2-TeCA; 1,1,2,2-TeCA

Corrinoid; FE/S?

propyl iodide; sulfite

Suyama et al., 2002

TCE 535.3

811.3 ± 39.0

c-DCE

CrdA Q8GFE2

Desulfitobacterium hafniense PCP-1(formerly D.frappieri)

33.8 7.0 <45 <60 2,4,6-TCPh

18.3± 2.8

28.8± 1.5

2,4-DCPh

2,3,4,5-TeCPh; 3,4,5-TCPh; 2,3,5-TCPh; 2,3,4-TCPh; 2,3,6-TCPh; 2,4,5-TCPh; 2,3-DCPh; 2,4-DCPh; PCE

Corrinoid; no Fe/S

Sulfite; sodium azide; nitrate; iodopropane

Boyer et al., 2003

PentaCPh

26.8± 2.9

882.8± 33.6

2,3,4,5-TeCPh

PceA Q8GJ27

Dehalobacter restrictus

60±1 8.1 NR 280 PCE 20.4 15000 c-DCE TCFE; TCM; HCA; TeCA; TCA and 1,1,1-TC-2,2,2- TFA


Iodopropane; ammonium

Maillard et al., 2003 TCE 23.7 20280 c-DCE

CprA5 Q6V7J3

Desulfitobacterium hafniense PCP-1(formerly D.frappieri)

57 6.8-7.0

50 110 3,5-DCPh 49.3 ±3.1

188.1 ± 5.2

3-CPh PentaCPh; 3,4,5-TCPh; 2,4,6-TCPh; 2,3,4,5-TeCPh; 2,3-DCPh;

Corrinoid; FE/S?

Sulfite; KCN; iodopropane; NaN3

Thibodeau et al., 2004

44

2,3,5-TCPh; 2,3,6-TCPh; 2,4,5-TCPh; 2,4-DCPh

VcrA Q69GM4

Dehalococcoides mccartyi VS

62 NR NR 5±3 VC ND 350 Ethene NA Corrinoid? 2 × [4Fe-4S]?

ND Müller et al., 2004 1,1-DCE ND 350-

390 VC to ethene

c-DCE ND 350-390

VC to ethene

1,2-trans-DCE

ND 350-390

VC to ethene

TCE ND 18.5 cDCE

PentaCPh- CprA3 Q8RPG3

Desulfitobacterium hafniense PCP-1

47 7.0 55 128 PentaCPh

46.7 1145 ± 33.1

2,3,4,5-TeCPh

2,3,5,6-TeCPh; 2,3,4,5-TeCPh; 2,4,6-TCPh; 2,3,6-TCPh; 2,3,4-TCPh

Corrinoid; Fe/S?

Iodopropane; sulfite; KCN; NaN3

Bisaillon et al., 2010

BhbA K4MLZ9

Comamonas sp. 7D-2

117 7.5 25-30 not oxygen sensitive

3,5-DB-4-HBA

37.6 56569.2

4-HBA 3,5-DC-4-HBA; 2-C-4-HBA; 2,6-DBPh; bromoxynil

no corrinoid, 2 × [4Fe-4S]?

Hg2+; Zn2+; Cd2+; Cu2+; Al3+; Co2+; Mn2+; sodium dithionite; sodium thiosulphate

Chen et al., 2013

3-B-4-HBA

148.9

55241.9

4-HBA

TmrA T0I1B4

Dehalobacter sp. UNSWDHB

44.5 7.2 45 Sensitive but not deter-mined

CF 154 ± 41

1250 ± 0.09

DCM 1,1,2-TCA; 1,1,1-TCA; 1,1-DCA


High conc. CF Jugder et al., 2017

45

Abbreviations: BA, benzoate; n-CBA, n-chlorobenzoate; PCE, tetrachloroethene; TCE, trichloroethene; c-DCE, cis-1,2-dichloroethene; t-DCE, trans-1,2-dichloroethene; 1,1-DCE, 1,1-dichloroethene; n,n-DBE, n,n-dibromoethene; TBE, tribromoethene; n,n-DCPE, n,n-dichloropropene; n,n-DBPE, n,n-dibromopropene; 2-CPE, 2-chloropropene; VC, vinyl chloride; TeIE, tetraiodoethene; n,n-DCPh, n,n-dichlorophenol; VB, vinyl bromide; 3-C-4-HPA, 3-chloro-4-hydroxyphenylacetate; 4-HPA, 4-hydroxyphenylacetate; n-B-n-CPh, n-bromo-n-chlorophenol; n-CPh, n-chlorophenol; n,n-DCPh, n,n-dichlorophenol; n,n,n-TCPh, n,n,n-trichlorophenol; PentaCPh, pentachlorophenol; 2-F-4-CPh, 2-fluoro-4-chlorophenol; 3-B/C-4-HBA, 3-bromo/chloro-4-hydroxybenzoate; 3,5-DB/C-4-HBA, 3,5-dibromo/chloro-4-hydroxybenzoate; 4-HBA, 4-hydroxybenzoate; 3-C-4-HPA, 3-chloro-4-hydroxyphenylacetate; 4-HPA, 4-hydroxyphenylacetate; TeC-BPD, tetracholobiphenyldiol; TC-BPD, trichloro-biphenyldiol; DC-BPD, dichloro-biphenyldiol; CF, chloroform; DCM, dichloromethane; n,n,n-TCPh, n,n,n-trichlorophenol; PentaCA, pentachloroethane; n,n,n,n-TeCA, n,n,n,n-tetrachloroethane; HCA, hexachloroethane; n,n,n-TCA, n,n,n-trichloroethane; n,n-DCA, n,n-dichloroethane; TCM, tetrachloromethane; n,n,n,n-TeCA, n,n,n,n-tetratchloroethane; n,n,n,n-TeCPh, n,n,n,n-tetrachlorophenol; 3-C-4-HBA, 3-chloro-4-hydroxybenzoate; 3,5-DC-4-HBA, 3,5-dichloro-4-hydroxybenzoate; TCFE, trichlorofluoroethene; n,n,n,-TC-n,n,n-TFA, n,n,n-trichloro-n,n,n,-trifluoroethane; 2,6-DBPh, 2,6-dibromophenol; NA, not available; ND, not determined; NR, not recorded.

46

Various endeavours to produce native RDases have been reported since mid-1990s, they

involve the use of different cell lysis methods followed by different solubilization

techniques and chromatographic separation, mostly resulting in partially purified

proteins with varied specific activity (Jugder et al., 2016a). The important steps involved

in the purification of native enzymes that has so far been isolated are summarized in

Table 1.3.

47

Table 1.3 Purification methods used for native RDases

Enzyme Organism Cell lysis method

Membrane solubilization

Chromatography steps Recovery (%)

Purification factor

References

3CBA-RDase Desulfomonile tiedjei DCB-1 French Press Yes (1% triton X-100)

IEC→HAC→HIC 7 123.1 Ni et al., 1995

PceA Sulfurospirillum multivorans (formerly Dehalospirillum multivorans)

Chemical Lysis No IEC→HIC→ SEC 64 106 Neumann et al., 1996

PCE-RDase Mixed enrichment culture of Dehalococcoides mccartyi 195 (formerly D. ethenogenesis 195)

French Press, tissue homogenizer

Yes (0.1% triton X-100)

HIC→HIC 17 75 Magnuson et al., 1998

TCE-RDase Mixed enrichment culture of Dehalococcoides mccartyi 195 (formerly D. ethenogenesis 195)

French Press, tissue homogenizer

Yes (0.1% triton X-100)

HIC→HIC 4 24 Magnuson et al., 1998

PCE-RDase Desulfitobacterium frappieri strain PCE-S

Chemical lysis No IEC→SEC→ IEC 75 165 Miller et al., 1998

3-C-4-HPA- RDase

Desulfitobacterium hafniense

French Press Yes (50 mM CHAPS)

IEC→HIC 18 11.3 Christiansen et al., 1998

CprA Desulfitobacterium dehalogenans

Sonication Yes (1% triton X-100)

IEC→IEC→ IEC 46 90 van de Pas et al., 1999

3-C-4-HBA-RDase

Desulfitobacterium chlororespirans


IEC→IEC→ IEC 32.3 181 Krasotkina et al., 2001

PceA Clostridium bifermentans DPH1 Ultrasonic disrupter

No IEC→SEC 42.2 13.2 Okeke et al., 2001

CPRD Desulfitobacterium sp. PCE-1


IEC→IEC→ IEC 0.3 136.3 van de Pas et al., 2001

PCERD Desulfitobacterium sp. PCE-1


IEC→IEC→ IEC 1 275.5 van de Pas et al., 2001

PCE/TCERD Desulfitobacterium sp. TCE-1


IEC→IEC→ IEC 58.8 100.1 van de Pas et al., 2001

PceA Desulfitobacterium sp. Y51 French Press No HAC→HIC→ CFC 0.2 2.55 Suyama et al., 2002

48

CrdA Desulfitobacterium hafniense PCP-1(formerly D.frappieri)

French Press Yes (0.1% triton X-100)

HIC→SEC 8.3 3.4 Boyer et al., 2003

PceA Dehalobacter restrictus Ultrasonication Yes (1.2% triton X-100)

IEC→SEC 53 22.3 Maillard et al., 2003

CprA5 Desulfitobacterium hafniense PCP-1(formerly D.frappieri)

French Press Yes (0.1% triton X-100)

IEC→HIC 6.8 12.7 Thibodeau et al., 2004

VcrA Dehalococcoides mccartyi VS

French Press Yes (2 mM CHAPS)

IEC→SEC 5 4.5 Müller et al, 2004

PentaCPh- CprA3

Desulfitobacterium Hafniense PCP-1

French press Yes (3% triton X-100)

IEC→HIC 2.3 11.3 Bisaillon et al., 2010

BhbA Comamonas sp. 7D-2 Sonication Yes (0.5% triton X-100)

IEC 12 35.8 Chen et al., 2013

TmrA Dehalobacter sp. UNSWDHB Bead beating, ultracentrifuge

Yes (1% triton X-100)

IEC 38 23 Jugder et al., 2017

Ion exchange chromatography (IEC), Hydrophobic interaction chromatography (HIC), Size exclusion chromatography (SEC), Hydroxyapatite chromatography (HAC), Chromatofocusing chromatography (CFC). Recovery (%): % of total activity recovered.

49

1.17 Heterologous expressions of RDases: limitations and successes

With the continued advancement of genetic engineering tools, production of

recombinant proteins in microbial systems has become an attractive alternative to

protein extraction from natural sources. Escherichia coli is the most extensively used

prokaryotic host for heterologous expression of protein; efforts have been made over

the last two decades to produce RDases heterologously in E. coli. So far, no respiratory

RDase has been expressed recombinantly in E. coli in a soluble and catalytically active

form. Though some of these efforts resulted in the over-expression of the protein in

interest, they were catalytically inactive, aggregated as inclusion bodies in the cytoplasm

(Jugder et al., 2018, Kimoto et al., 2010, Neumann et al., 1998, Sjuts et al., 2012, Suyama

et al., 2002). The formation of inactive RDases was mainly credited to the fact that E.

coli is incapable of synthesizing corrinoids de novo and hence there were insufficient

amounts of cobamide cofactor in the cells (Neumann et al., 1998, Suyama et al., 2002).

Experiments using different solubility tags and reconstitution of corrinoid cofactors and

Fe-S clusters also failed possibly because of the use of the improper and insufficient

cobalamin cofactor and incomplete reconstitution of the two Fe-S clusters (Jugder et al.,

2018, Sjuts et al., 2012). The first success story reported in E. coli was the over-

expression of vinyl chloride reducing VcrA from Dehalococcoides mccartyi strain VS and

subsequent in vitro reconstitution by the incorporation of a corrinoid cofactor and two

iron-sulfur clusters, thus producing a functional protein (Parthasarathy et al., 2015).

Collin (2017) over-expressed active catabolic reductive dehalogenase, NpRdhA, a

phylogenetically distinct and aerobic RdhA from Nitratireductor pacificus strain pht-3B

in a vitamin-B12 transporter btuB over-expressing E. coli. Nakamura et al., (2018) over-

expressed tetrachloroethene reducing PceA from Geobacter sp. in E. coli, purified and

denatured the protein under aerobic conditions. Consequently, they refolded PceA in

the presence of FeCl3, Na2S and cobalamin under anaerobic conditions and the

reconstituted protein demonstrated dechlorination of tetrachloroethene. Very recently,

Halliwell et al., (2021) over-expressed NpRdhA in E. coli HMS174(DE3), they also co-

expressed NpRdhA with a vitamin-B12 transporter btuB in E. coli HMS174(DE3). (Halliwell

et al., 2021) The same group expressed NpRdhA in a commercial strain of Vibrio

50

natrigenes, Vmax™, a host that has not been previously reported as an expression host

for RDases. V. natrigenes cannot synthesis vitamin B12 de novo, rather it salvages

cobalamin/cobinamide via an aerobic pathway (Agarwal et al., 2019).

The production of catalytically active RDases is made complex by their need of

cobamides and Fe–S clusters; the synthesis and proper insertion of the cofactors hinders

recombinant expression approaches. Hence, the corrinoid-producing Gram-negative

gammaproteobacterium Shimwellia blattae and the Gram-positive Bacillus megaterium

have been identified as suitable hosts for heterologous production of functional RDases

(Mac Nelly et al., 2014, Payne et al., 2015). B. megaterium produces 5,6-

dimethylbenzimidazolyl cobamide, the standard-type B12 cofactor, and S. blattae

synthesizes pseudo-B12, adeninyl cobamide de novo (Mac Nelly et al., 2014, Wolf and

Brey, 1986). The PceA from Desulfitobacterium hafniense strain Y51 and RdhA3 of strain

DCB-2 were produced in Shimwellia blattae where co-expression of the respective

chaperone proteins PceT and RdhT improved over-expression of the functional enzymes

(Mac Nelly et al., 2014). Raising the intracellular corrinoid level by adding exogenous

corrinoid hydroxocobalamin and the precursor of the corrinoid’s lower ligand base (5,6-

dimethylbenzimidazole) to the growth media and using glycerol as the growth substrate

for S. blattae also increased the formation of active RDases (Mac Nelly et al., 2014). A

similar protocol was followed to produce functional DcaA from D. dichloroeliminans

DCA1 in S. blattae; co-expression of dedicated chaperone DcaT resulted in increased

dechlorination activity (Kunze et al., 2017). Halliwell et al., (2021) expressed active

NpRdhA in S. blattae.

The NpRdhA was first heterologously expressed in Bacillus megaterium by using C-

terminal His-tagged NpRdhA and inducing its xylose-inducible promoter (Payne et al.,

2015). The TmrA protein of Dehalobacter strain UNSWDHB has also recently been

expressed in a catalytically active form in Bacillus megaterium with an N-terminal His-

tag and devoid of the TAT signal peptide to exclude any requirement for co-expression

of chaperone protein to facilitate membrane dislocation of TmrA (Jugder et al., 2018).

These efforts now allow the production of reductive dehalogenase preparations in mg-

scale which makes the structural and functional studies possible. A summary of the

51

attempted and successful heterologous expression of RDases that have been reported

so far is presented in Table 1.4.

52

Table 1.4 Heterologous expression of RDases

Enzyme Native Host Expression Host

Vector used

Affinity tag(s)

Inducer Length of Induction

Other Experimental Features

Specific Activity (nmol/

min/mg) of recom-

binant protein

Specific Activity (nmol/

min/ mg) of

purified native

protein

Comments References

PceA Sulfurospirillum multivorans (formerly Dehalospirillum multivorans)

E. coli BL21 (DE3)/pLysS

pET 11d - 0.4 mM IPTG

1.5-4 h Expression of pceB was attempted Vitamin B12 was added to the growth media crude extracts of D. multivorans were added to the growth medium

Inactive 158,400 (PCE)

PceA was expressed only in the presence of pUBS 520 which provided high levels of E. coli tRNA4Arg

Neumann et al., 1998

PceA Desulfitobacterium hafniense strain Y51

E. coli BL21 (DE3)/pLysS

pET32b(+)

Trx (thioredoxin protein) Tag, S-Tag, N-His, C-His

1.0 mM IPTG

NM Solubilized fusion protein was used to raise antibody

Inactive 164.4 ± 27.2 (PCE)

Attempt to express pceA and pceB using vector pBluescript II KS+ in E. coli JM109 was unsuccessful

Suyama et al., 2002

811.3 ± 39.0 (TCE)

PceA1/PceA2

Environmental sample

E. coli BL21 And other strains (not mentioned)

pQE6, pBAD, pCold

His-tag IPTG NM PceA1 was synthesized in vitro

Inactive - IPTG treatment decreased the yield of expressed protein PceA1 synthesized in vitro was active

Kimoto et al., 2010

PceA Dehalobacter restrictus

E. coli BL21 (DE3)

pOPIN, pET50b(+),

N-His, SUMO, MBP,

0.1 mM IPTG

12-16 h TAT signal sequence was initially included then excluded

Inactive 15,000 (PCE)

Only TF fusion proteins resulted in

Maillard et al., 2003

53

pCOLD TF

NusA, Strep-tag II, TF

pCOLD vector contained a cold shock promoter (cspA) which allowed induction of gene expression at lower temperature Fe-S and cobalamin cofactors were reconstituted

20,280 (TCE)

stable and soluble expression Low induction temperature is essential for soluble expression

Sjuts et al., 2012

PceA Desulfitobacterium hafniense strain Y51

Shimwellia blattae

pASK-IBA63c-plus

N-Strep-tag II

20 ng/ml anhydrotetracycline

6 h TAT signal sequence was included Co-expressed with chaperone protein pceT Glycerol was used as carbon and energy source

0.50 ± 0.11 (PCE)

164.4 ± 27.2 (PCE)

First functional recombinant RDase NStrep-PceA produced the highest amount of protein NStrep-PceA co-expressed with PceT had the best solubility Addition of 5,6-DMB and OH-B12 significantly increased the specific activity Reductive dehalogenation activity was also detected in the absence of PceT Purification of the protein was not possible

Suyama et al., 2002 Mac Nelly et al., 2014

811.3 ± 39.0 (TCE)

RdhA3 Desulfitobacterium hafniense strain DCB-2

Shimwellia blattae

pASK-IBA63c-plus

N-Strep-tag II

20 ng/ml anhydro

6 h Co-expressed with chaperone protein RdhT

0.15±0.03 (2,3-DCP)

- Maximum reductive dehalogenation was observed with

Mac Nelly et al., 2014

54

tetracycline

0.40±0.09 (2,4-DCP)

NStrep-RdhA3-RdhT-CStrep after 18 h of cultivation after induction in DMB and OH-B12 amended media Reductive dehalogenation was also detected in the absence of chaperone protein

0.41±0.08 (3,5-DCP)

0.24±0.07 (2,4,5-TCP) 0.17±0.03 (2,4,6-TCP) 0.03±0.01 (PCE)

NpRdhA Nitratireductor pacificus pht-3B

Bacillus megaterium

pPT7 C-His 0.1% xylose

overnight at 17 °C, 180 rpm

Cultures were supplemented with

50 M ammonium iron(II) sulphate and 1

M B12

NM - Purified under aerobic conditions

Payne et al., 2014

VcrA Dehalococcoides mccartyi strain VS

E. coli BL21 (DE3)

pLIC-HMK

N-His tag, MBP-tag, TEV protease Cleavage site

1 mM IPTG

3 h TAT signal sequence was removed Corrinoid and Fe-S cofactors were reconstituted

1050 (VC, with MBP tag);

350 (VC) Change in culture conditions (media, temperature, concentration, and length of induction) did not result in soluble expression of protein Inclusion bodies were purified to apparent homogeneity by His-Tag affinity chromatography First report of refolding inclusion bodies into active reductive dehalogenase

Müller et al, 2004 Parthasarathy et al., 2015

2250 (VC, no MBP)

55

DcaA Desulfitobacterium dichloroeliminans DCA1

Shimwellia blattae

pASK-IBA63c-plus

Strep-tag

200 ng mL-1 anhydrotetracycline

18 h Co-expressed with chaperone protein dcaT Glycerol was used as growth substrate

2.5 - The best result (increased amount of protein and specific activity) was obtained when dceA and dceT genes were cloned onto the same plasmid, the induction time was increased to 18 h and the media contained

OMeBZA (10M)

Kunze et al., 2017


E. coli BL21 over-expressing btuB

pPT7 His 0.25 M IPTG


Co-expressed with btuB-pLysS plasmid Cultures were supplemented with 50 μM ammonium iron(II)sulphate, 1 μM B12

- Purified under aerobic conditions Specific activity comparable to NpRdhA expressed in B. megaterium

Collin, 2017

TmrA Dehalobacter UNSWDHB

Bacillus megaterium

pPT7 N-His 0.5% xylose


TAT signal sequence was removed Cultures were supplemented with 50 µM ammonium iron(II) sulphate and 1 μM B12

110 1250 ± 0.09 (TCM)

The enzyme was purified via two liquid chromatographic steps

Jugder et al., 2017, 2018

PceA Geobacter sp. E. coli BL21 (DE3)

pET23b pCold TF

Strep, TF

IPTG NM Corrinoid and Fe-S cofactors were reconstituted

NM - Cell lysis and protein purification was done aerobically whereas the cofactor reconstitution steps were performed under anaerobic conditions.

Nakamura et al., 2018

56


E. coli HMS174(DE3)

pET30 C-His 1 mM IPTG

NM Cultures were supplemented with 50 μM ammonium iron(II) sulphate and 1 μM B12

24 ± 0.8 x 103 (crude extract) 170 ± 3 x 103 (purified protein)

- Purified under aerobic conditions

Halliwell et al., 2021


E. coli HMS174(DE3)

pET30 and pET3a

C-His 1 mM IPTG

NM Co-expressed with BtuB Cultures were supplemented with 50 μM ammonium iron(II) sulphate and 1 μM B12

35 ± 0.2 x 103 (crude extract) 200 ± 8 x 103 (purified protein)




Bacillus megaterium

pPT7 C-His 0.1% xylose


4.1 ± 0.1 x 103 (crude extract) 290 ± 15 x 103 (purified protein)




Shimwellia blattae

pASK-IBA63c-plus

C-His 20 ng/mL anhydrotetracycline


13 ± 0.3 x 103 (crude extract) 130± 6.7 x 103 (purified protein)




Vibrio natrigenes Vmax™

pET30 C-His 1 mM IPTG


18 ± 1.1 x 103 (crude extract from TB media) 20 ± 0.3 x 103 (crude extract from 2YT media)



57

180 ± 8 x 103 (purified protein from TB media) 260 ± 11 x 103 (purified protein from 2YT media)

NM: Not Mentioned; OMeBza: 5-methoxybenzimidazole

58

1.18 Dehalobacter UNSWDHB

The genus Dehalobacter belongs to the phylum Firmicutes, class Clostridia, order

Clostridiales and family Peptococcaceae (Holliger et al., 1998a). Strain UNSWDHB of

Dehalobacter sp. was isolated in Australia from subsurface soil microbial community

from a chloroform contaminated aquifer near Sydney (Lee et al., 2012, Wong et al.,

2016). D. UNSWDHB is a gram negative, obligate anaerobic and organohalide-respiring

bacterium. Phase-contrast and fluorescence microscopy of the pure culture shows

uniform cell morphology, cells are planktonic, and rod shaped. Transmission electron

microscopy (TEM) reveals a cell dimension of 2 µm length and 0.3–0.5 µm diameter

along with the presence of a lateral flagellum.

Wong et al. (2016) carried out extensive studies on the isolation and characterization of

Dehalobacter sp. UNSWDHB. The strain uses molecular hydrogen as the sole external

electron donor and can reductively dechlorinate trichloromethane, 1,1,2-

trichloroethane, 1,1,1-trichloroethane and 1,1-dichloroethane which are utilised as

electron acceptors. Unlike other OHRB, it has a high tolerance to trichloromethane

(TCM) because it does not affect the RDase and can dechlorinate up to 4 mM TCM to

dichloromethane (DCM) at a rate of 0.1 mM per day. It can dechlorinate 1,1,2-

trichloroethane (1 mM, 0.1 mM per day) to vinyl chloride (VC) and 1,2-dichloroethane

(1,2-DCA) at a ratio of 1:1.7. 1,1,1-trichloroethane and 1,1-dichloroethane are slowly

utilized and cannot be removed completely under growth conditions, with initial

dechlorination rates of 5 and 1 mM per day. Enzyme kinetic experiments also confirm

strong substrate affinity for TCM and 1,1,2-trichloroethane (Km = 30 and 62 µM

respectively) and poor substrate affinity for 1,1,1-trichloroethane and 1,1-

dichloroethane (Km = 238 and 837 µM respectively). Cell yields per mole of chloride

released in response to TCM, 1,1,2-TCA, 1,1,1-TCA and 1,1-DCA were recorded to be

(6.3±0.5) x 1012, (4.3±0.5) x 1012, (3.4±1.6) x 1012 and (6.3±0.6) x 1012 respectively.

D.UNSWDHB has a genome size of ca. 3.2 Mb, GC content of 44.9%, 3041 genes along

with 17 rdhA genes (Deshpande et al., 2013). The strain lacks the proteins involved in

corrinoid salvage or transport and unlike other Dehalobacter sp. UNSWDHB can grow

59

without exogenous supply of cobalamin which confirms the presence of a fully

functional cobalamin synthesis pathway (Wong et al., 2016). The genome of strain

UNSWDHB encodes a complete Wood–Ljungdahl pathway (WLP) and with the addition

of TCM, the WLP proteins show a comparatively high abundance though the functions

of these pathways are still unresolved (Jugder et al., 2016d). Additionally, the TCM-

induced upregulation of genes encoding nine hydrogenases, ATP synthase, NADH-

ubiquinone oxidoreductase, proton-translocating respiration complex I, a proton-

pumping inorganic pyrophosphatase, pyruvate phosphate dikinase and a complete

menaquinone biosynthesis pathway reveal the metabolic diversity of the strain

UNSWDHB (Jugder et al., 2016b).

1.19 RDase of Dehalobacter UNSWDHB (TmrA)

The UNSWDHB strain genome includes 17 full-length and three truncated rdhA genes

distributed in 16 gene clusters (Deshpande et al., 2013). The TCM RDase (TmrA)

expressed by strain UNSWDHB (Accession Number EQB21016) is encoded by 1368

nucleotides transcribed into 456 amino acids and has a predicted molecular weight of

50 kDa. The full-length amino acid sequence of TmrA is shown in Figure 1.11. TmrA

shows 95% amino acid identity with CfrA (RDase of Dehalobacter strain CF) and CtrA

(RDase of Desulfitobacterium strain PR) with six important substitutions or deletions

which hinders its ability to efficiently dechlorinate 1,1,1-TCA or 1,1-DCA like the other

two (Wong et al. 2016). The tmrA gene cluster contains genes encoding the membrane

anchor protein (TmrB) and a putative transcriptional regulator protein (TmrC) while the

gene encoding the trigger factor protein (TmrT) is located downstream of tmrABC. A

proteomic study revealed significant upregulation of TmrA, TmrB and TmrC during TCM

respiration (Jugder et al. 2016b). Jugder et al., (2017) purified native TmrA from

Dehalobacter sp. UNSWDHB which displayed a specific activity of (1.27 ± 0.04) × 103

units mg of protein-1 in vitro. The UV-visible spectrometric analysis revealed the

presence of a corrinoid and two [4Fe-4S] clusters.

60

Figure 1. 11. Full length amino acid sequence of TmrA. The framed region consisting of the TAT signal peptide (highlighted in yellow) and a predicted transmembrane domain (bold font) was removed prior to gene cloning in some experiments described in this dissertation.

1.20 Research aims and thesis outline

In recent years there has been some success in recombinant expression of functional

RDases though there is much more to be done to improve available tools and explore

new tools for functional recombinant expression of RDases for structural and functional

studies and potentially for implementation in bioremediation tasks. This project aims to

produce functional recombinant chloroform-reducing dehalogenase, TmrA, from

Dehalobacter sp. UNSWDHB in different host systems, building on work previously

undertaken in our lab. Expression in different host systems will allow the comparison of

the efficiency of expression in each system and provide valuable guidelines for future

recombinant RDase production projects.

Chapter 2 describes the refolding and reconstitution of TmrA from inclusion bodies

expressed in E. coli into catalytically active forms.

Chapter 3 focuses on the improvement of soluble expression and specific activity of

TmrA expressed in B. megaterium by manipulating the culture conditions using factorial

design experiments.

Chapter 4 investigates the heterologous expression of TmrA in S. blattae. As described

earlier in this chapter, this corrinoid producing host has previously been shown to

successfully generate functional recombinant dehalogenases in a limited range of

61

studies. This required the development of a number of molecular cloning tools, and the

development of a bioprocess to obtain soluble and catalytically active TmrA.

Chapter 5 focuses on attempts to engineer Cupriavidus necator, a Gram-negative,

hydrogen-oxidizing bacteria capable of growing at the interphase of anaerobic and

aerobic environments, into a chloroform-reducing bacterium. The use of this host to

express functional RDases has not previously been described in the literature.

Chapter 6 brings together the data and information obtained in the previous chapters

and discusses how this thesis has deepened our understanding of heterologous

expression of RDases and in turn the reductive dehalogenation process using chloroform

dehalogenation to dichloromethane as a model. The methods and insights gained from

the work presented in this thesis provide a platform for future protein structure-

function studies, protein engineering for examining substrate specificity and improved

activity and potentially engineering studies for eventual bioremediation applications.

62

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CHAPTER TWO: Recovering functional chloroform reductive dehalogenase from inclusion bodies in Escherichia coli

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2.1 INTRODUCTION

Dehalobacter sp. UNSWDHB is a gram negative, obligate anaerobic and organohalide-

respiring bacterium which was isolated in Australia from a chloroform contaminated

aquifer near Sydney (Wong et al., 2016). Since its isolation (Lee et al., 2012) and genome

sequencing (Deshpande et al., 2013), several studies have been carried out to decipher

the biochemical properties of the strain and corresponding reductive dehalogenases

and their potential role in bioremediation. Wong et al., (2016) conducted preliminary

biochemical studies on Dehalobacter sp. UNSWDHB. Jugder et al. (2016) reported a

proteomic and transcriptomic analysis of the organism in response to chloroform

following a short period of starvation, identifying a range of genes upregulated in

response to chloroform presence. The same authors also purified the native TmrA from

the UNSWDHB strain and undertook characterisation of the enzyme, including an

estimation of the specific activity of TmrA (Jugder et al., 2017). Finally, they reported

successful expression of functional purified recombinant TmrA in Bacillus megaterium

(Jugder et al., 2018). This was the first membrane associated respiratory RDase to be

heterologously expressed in a soluble and active form.

E. coli has proven to be a successful expression host for a vast number of protein classes,

for overexpression in soluble and functional forms. Because of the wealth of available

knowledge and success stories, E. coli is the first choice for recombinant expression for

most proteins. Several attempts have been made to express RDases in E. coli but no

respiratory RDase has been expressed in a soluble and catalytically active form so far.

The endeavour of Jugder et al., (2018) to express the RDase of the UNSWDHB strain in

a soluble and functional form in E. coli was similarly unsuccessful. He removed the TAT

signal and the predicted membrane spanning regions from the tmrA gene with the

intention of directing the expression of the periplasmic RDases toward the cytoplasm.

Rhamnose-inducible heterologous expression of TmrA under the control of an

adjustable rhaBAD promoter was not tightly regulated and resulted in insoluble

expression. The use of IPTG-inducible pOPIN vector system with a T7 inducible promoter

and affinity/solubility fusion tags (N/C-terminal His, SUMO, Trx, NusA, MBP, and TF)

resulted in overexpression of TmrA as inclusion bodies. Despite attempts to generate

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soluble protein by induction at a lower temperature, the use of different growth media

and IPTG concentration, and cofactor supplementation, there was no soluble expression

of TmrA. The final attempt was to recover TmrA from inclusion bodies by purification of

denatured TmrA followed by refolding as was performed by (Parthasarathy et al., 2015)

with the vinyl chloride-reducing dehalogenase VcrA from Dehalococcoides mccartyi

strain VS. Several refolding methods were performed such as refolding by flash or slow

dilution, dialysis, on-column refolding, and chemical cofactor reconstitution but all

attempts were unsuccessful. The failure to express soluble and functional RDases in E.

coli (Jugder et al., 2018, Kimoto et al., 2010, Neumann et al., 1998, Sjuts et al., 2012,

Suyama et al., 2002) has been attributed to the inability of E. coli to synthesize corrinoids

de novo and therefore there were insufficient amounts of cobamide cofactor in the cells

(Neumann et al., 1998; Suyama et al., 2002).

In this chapter we aimed to refold TmrA from inclusion bodies expressed in E. coli into a

functional form capable of dechlorinating chloroform to dichloromethane. We used the

pOPIN-TmrA-CHis, pOPIN-TmrA-MBP and pOPIN-TmrA-TF constructs (Figure 2.1)

described in Jugder et al., (2018) for this purpose.

Figure 2.1. TmrA fusion constructs with affinity (NHis and CHis) and solubility tags MBP (Maltose binding protein) and TF (trigger factor), encoded in pOPIN expression vector. The recognition sequence 3C allows the Human Rhinovirus 3C Protease to cleave off the fusion proteins encoded in pOPIN expression vector.

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2.2 EXPERIMENTAL PROCEDURES

Initially a non-conventional refolding method described by Roussel et al. (2013) for

recovering TmrA from inclusion bodies was attempted. The method is based on the

association of the ionic detergent Sodium Dodecyl Sulphate (SDS) with a co-solvent

featuring a diol-type pattern, the 2-Methyl-2,4-pentanediol (MPD). A suitable

concentration ratio of SDS and MPD leads to the refolding of the protein previously

denatured with SDS concentrations above the critical micelle concentration (CMC)

(Michaux et al., 2016). The second method pursued in this study included an aerobic

chromatographic step followed by the refolding of TmrA under anaerobic conditions in

the presence of FeCl3, Na2S and cobalamin to facilitate cofactor reconstitution as

described by (Nakamura et al., 2018).

2.2.1 Bioreactor fermentation

The constructs pOPIN-TmrA-Chis and pOPIN-TmrA-MBP were selected for the SDS-MPD

based protein refolding system and for the second refolding method, pOPIN-TmrA-Chis

and pOPIN-TmrA-TF constructs were chosen. The reasoning behind selecting these

constructs is described in the Discussion section. To produce sufficient biomass, batch

fermentation of E. coli BL21(DE3)pLysS cells containing pOPIN-TmrA-CHis, pOPIN-TmrA-

MBP and pOPIN-TmrA-TF was carried out in a laboratory-scale 7 L glass bioreactor

(Applikon, The Netherlands). Cell growth was monitored by measuring OD600nm. 5 L of

2YT media with 50 μg/mL ampicillin and 17 μg/mL chloramphenicol was inoculated with

an overnight pre-culture at an OD600 of 0.2. The culture was maintained at 37 °C, 500

rpm with dissolved oxygen concentration (%dO2) regulation at 30%. Protein expression

was induced at OD600 of 1.2 with 0.5 mM IPTG for 3 hours. Cells were harvested by

centrifugation at 8,000 g and 4 °C for 10 min, washed with 1X PBS solution, and stored

at -20 °C for further processing. Pre- and post-induction samples were collected to

confirm TmrA expression by SDS-PAGE.

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2.2.2 SDS-PAGE

Samples collected before and after induction were normalised according to an OD600

of 1.0 and checked for soluble expression using BugBuster Protein Extraction Reagent

(Merck Millipore) following the manufacturer’s instructions. Proteins were separated on

a Bolt 4-12% Bis-Tris Plus polyacrylamide precast gel (Thermo Scientific). SeeBlue Plus2

Prestained Standard (Thermo Scientific) was used as the molecular weight marker and

the gel was stained with GelCode Blue Stain Reagent (Thermo Scientific).

2.2.3 SDS-MPD based system

All the buffers used purged with N2 gas and unless mentioned otherwise, the procedures

were carried out inside an anaerobic chamber (Baker Ruskinn Concept).

2.2.3.1 Cell Lysis

Cell pellets obtained from the 5 L fermentation (pOPIN-TmrA-Chis and pOPIN-TmrA-

MBP) were thawed and resuspended in a lysis buffer (50mM Tris–HCl pH 8, 17mM NaCl,

PMSF tablets, 250 μg/mL lysozyme) with a wet cell weight (WCW) to buffer ratio of 1:5

for 20 minutes at 25 °C. Cells were further broken by addition of 1 mg/mL of sodium

deoxycholate for 60 min at 37 °C with constant shaking, and 25 μg/mL of DNase I (Sigma

Aldrich D5025) for 60 minutes at 25 °C. The cell suspension was then centrifuged at

14,000 g for 20 minutes at 4 °C. The resulting pellet was resuspended in a washing buffer

(2 M urea, 20 mM Tris-HCl pH 8, 500 mM NaCl, 2% Triton X-100) and centrifuged at

14,000 g for 20 minutes at 4 °C.

2.2.3.2 Purification of TmrA from insoluble fraction

The inclusion bodies were solubilized with 50 mL of solubilization buffer (50mMTris–HCl

pH 8, 17mM NaCl, 8M urea), centrifuged at 5,000 g, 4 °C for 10 minutes, and filtered

through a 0.22 μm membrane. Affinity chromatography and anion exchange

chromatography were carried out as two separate processes for TmrA-CHis. Only affinity

chromatography was done for TmrA-MBP. The inclusion body preparation was loaded

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onto a 1 mL Ni-NTA Superflow Cartridge using a peristaltic pump (Gilson Minipuls 2)

calibrated to a flowrate of 1 mL/min. The column was previously equilibrated with 20

CV of buffer A (100 mM Tris-HCl pH 8, 150 mM NaCl, 10% glycerol, 8 M urea). Gradient

elution of the protein was performed with elution buffer containing varying

concentrations of imidazole (buffer A + 10/50/250 mM imidazole).

Anion exchange chromatography was performed using a HiTrap™ DEAE FF 1 mL column.

The column was equilibrated with 10 CV of start buffer (100 mM Tris, pH 8, 10% glycerol)

after which the inclusion body preparation was loaded at a flowrate of 1 mL/min. The

protein was eluted with 5 CVs of an elution buffer containing 1 M NaCl.

2.2.3.3 Cofactor reconstitution

Prior to cofactor reconstitution, the concentrated refolded protein solution was reduced

with 5 mM β-mercaptoethanol and incubated at room temperature for 30 minutes for

complete reduction. Protein concentration was determined spectroscopically at 280 nm

using a molar extinction coefficient value of 72,140 M-1 cm-1 (calculated from the

primary amino acid sequence using ProtParam-ExPaSy). Anoxic solutions of 250 mM

FeCl3 and 100 mM Na2S were added at five-fold and ten-fold molar excess over protein

to the solution which was then incubated at 30 °C for 90 minutes followed by

centrifugation at 5,000 g for 5 minutes. Cyanocobalamin was then added to the

supernatant at a final concentration of 10 mg/mL and incubated at room temperature

for 30 minutes.

2.2.3.4 Refolding of denatured TmrA fusion proteins

The eluted fractions from section 2.2.3.3 were loaded onto PD-10 columns for buffer

exchange. The columns were previously equilibrated with a buffer containing 150 mM

NaCl, 50 mM Tris-HCl pH 8, and 120 mM SDS. SDS-unfolded protein was then diluted 1:1

in a refolding buffer (50 mM Tris-HCl pH 8, 150 mM NaCl, 3 M MPD). The protein solution

was incubated at room temperature for 24, 48, 72 and 96 h and then stored at -20 °C to

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stop the refolding reaction. The same was done for cofactor reconstituted TmrA from

section 2.2.3.3.

2.2.4 Aerobic purification and anaerobic cofactor reconstitution method

This method requires anaerobic manipulation only during the cofactor reconstitution

and refolding steps. Cell lysis and TmrA purification steps were carried out under

ambient conditions in the presence of oxygen.

2.2.4.1 Cell lysis

Cell pellets (pOPIN-TmrA-Chis and pOPIN-TmrA-TF) were resuspended in a suspension

buffer containing 50 mM potassium phosphate pH 8.0, 0.5 M NaCl, 20% glycerol and

EDTA-free Protease Inhibitor Cocktail tablet at a ratio of 1:10. The suspension was

sonicated intermittently (2 sec pulse-on and 2 sec pulse-off) for 8 min at an amplitude

of 50% using BRANSON Digital Sonifier (Emerson Industrial, Australia) and centrifuged

for 15 min at 14,000 g, 4 °C. The supernatant was discarded, and the pellet was

resuspended in denaturation buffer (suspension buffer + 8 M urea). After a second

round of sonication, the suspension was stirred on ice for 1 h to solubilize the pellet. The

remaining insoluble material was removed by centrifugation for 15 min at 14,000 g and

4 °C.

2.2.4.2 Purification of TmrA

The supernatants containing denatured TmrA-CHis and TmrA-TF respectively were

loaded onto the Ni-affinity column (His-Trap FF, GE Healthcare, Buckinghamshire,

United Kingdom), which had been washed with five column volumes of the denaturation

buffer and eluted with four column volumes of 250 mM imidazole in the denaturation

buffer.

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2.2.4.3 Cofactor reconstitution

The reconstitution of TmrA-Chis and TmrA-TF was done inside an anaerobic chamber.

All buffers were purged with N2 gas before use. Denatured TmrA-CHis was reduced by

mixing with a reduction buffer (100 mM Tris-HCl pH 7.5, 200 mM DTT) to 5 mM of DTT.

After incubation at room temperature for 30 min with stirring, iron buffer (100 mM Tris-

HCl pH 7.5, 100 mM FeCl3) and sulfide buffer (100 mM Tris-HCl pH 7.5, 30 mM Na2S)

were added up to 50 mol excess of Fe and S to TmrA-CHis. Following a 90 min incubation

with stirring cyanocobalamin was added to a final concentration of 10 mg/ml. Next,

TmrA-CHis solution was applied to a PD-10 column (GE Healthcare) previously

equilibrated with the refolding buffer containing 50 mM Tris-HCl pH 8.0, 0.5 M NaCl,

20% glycerol, 0.2% CHAPS, and 10 mM DTT. The reconstituted TmrA-Chis and TmrA-TF

containing fractions were collected and stored at -20 ᵒC for future analysis.

2.2.5 Dechlorination activity assay

Standard enzyme assays were performed anoxically in the anaerobic chamber, as

previously described by (Jugder et al., 2017). where titanium (III) citrate and methyl

viologen was used as electron donors. A mastermix was prepared by adding titanium

(III) citrate and methyl viologen to a washing buffer (100 mM Tris-HCl buffer, 150 mM

NaCl, pH 7.5) to reach a final concentration of 2mM and 1 mM, respectively. All reactions

were carried out in 2 mL amber GC vials. The required amount of mastermix, protein

extract and CF added to the crimp sealed GC vials leaving no headspace. Samples were

prepared in triplicate. The vials were incubated at 30 °C for 3 hours inside the anaerobic

chamber. After incubation, the enzymatic reaction was stopped by transferring 1 mL of

each reaction mixture to 10 mL headspace vials containing 0.5 g of anhydrous sodium

sulphate and 1 mL of 1 M sulphuric acid. The headspace of each of the vials was then

analysed with a Focus DSQ II GC/MS equipped with a TrisPlus autosampler (Thermo

Scientific). Each vial was heated at 80 °C for 2.0 min whilst being intermittently agitated

(10s on, 10s off), the autosampler injections were made with a syringe heated to 80 °C.

An injection volume of 0.5 mL was used with a variable split ratio of between 150:1 and

38:1 dependent of analyte concentration, an inlet temperature of 175°C, transfer line

88

and source temperature of 220°C, with helium used as the carrier gas at a constant flow

of 1.9 mL per min vacuum compensated. A RESTEK Rt-Q-BOND column (30 m × 0.32

mm, 10 µm) to separate analytes of interest. Separation was carried out at a

temperature gradient of 100°C for 1 min followed by 4 min ramping with 30°C/min to

reach a final temperature of 220°C. Data was acquired in positive EI using a full mass

scan from 50 to 150 Da. CF and DCM standards (0.025 mM, 0.05 mM, 0.075 mM, 0.1

mM, 0.15 mM, 0.2 mM, 0.5 mM) were prepared fresh on the day of the analysis and

treated the same way as the samples. The total GC/MS analysis time was 8.0 minutes,

with DCM and CF retention times at 4.0 min and 5.0 min (± 0.2 min), respectively. Data

analysis was carried out using the qual browser function of the XCalibur software

(Thermo Scientific). Enzyme specific activity was described as the formation of nmol

DCM per mg of protein per minute. The protein concentration was determined

spectroscopically at 280 nm with a nanodrop spectrophotometer using the molar

extinction coefficient value of 72,140 M-1cm-1 as described in section 2.2.3.3. Heat-

deactivated (incubated at 100°C for 20 min) enzymes were assayed as negative controls

to eliminate the possibility of non-enzymatic CF reductive activity.

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2.3 RESULTS

2.3.1 Large-scale heterologous production of TmrA in E. coli

5 L fermentation of BL21(DE3)pLysS cells containing pOPIN-TmrA-CHis/pOPIN-TmrA-

MBP/pOPIN-TmrA-TF in 2YT media was carried out, cells were harvested after a 3 h

induction with 0.5 mM IPTG (Figure 2.2). Pre- and post-induction samples were collected

and fractionated into soluble and insoluble portions using BugBuster. SDS-PAGE shows

the expression of TmrA-CHis, TmrA-MBP and TmrA-TF in the insoluble fraction (Figure

2.3). Expression of TmrA was further confirmed by tryptic digest and liquid

chromatography tandem mass spectrometry (LC-MS/MS) at the Biomedical Mass

Spectrometry Facility, UNSW (methods and results shown in Appendices, A.6).

Figure 2.2. Fermentation of BL21(DE3)pLysS cells transformed with pOPIN-TmrA-CHis, pOPIN-TmrA-MBP and pOPIN-TmrA-TF. The blue, grey and orange lines represent OD600, pH and dissolved oxygen (%), respectively. The black circle shows the time of induction.

90

Figure 2.3. SDS-PAGE analysis of heterologously expressed TmrA fusion proteins. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Pre-induced insoluble TmrA-CHis; Lane 3. Post-induced insoluble TmrA-CHis; Lane 4. Pre-induced soluble TmrA-CHis; Lane 5. Post-induced soluble TmrA-CHis; Lane 6. Pre-induced insoluble TmrA-MBP; Lane 7. Post-induced insoluble TmrA-MBP; Lane 8. Pre-induced soluble TmrA-MBP; Lane 9. Post-induced soluble TmrA-MBP; Lane 10. Pre-induced insoluble TmrA-TF; Lane 11. Post-induced insoluble TmrA-TF; Lane 12. Pre-induced soluble TmrA-TF; Lane 13. Post-induced soluble TmrA-TF. The framed sections show the protein of interest. The expected sizes of TmrA-Chis, TmrA-MBP and TmrA-TF are approximately 46, 88 and 96 kDa, respectively.

2.3.2 Recovery and refolding of TmrA from inclusion bodies: SDS-MPD based system

The cell lysis, purification and refolding steps were carried out anaerobically. After

separating the soluble and insoluble parts, the inclusion bodies (insoluble portion) were

solubilized with a solubilizing buffer containing 8M urea. The soluble inclusion bodies

(TmrA-CHis and TmrA-MBP) were loaded onto a Ni-NTA and DEAE columns for separate

one-step purifications. SDS-PAGE confirmed the binding of TmrA fusion proteins on both

the columns, albeit quite poorly for the DEAE column (Figures 2.4 and 2.5). In case of Ni-

NTA column, the best amount of elution was observed with the 50 mM imidazole

containing buffer (Figure 2.4).

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Figure 2.4. Purification of TmrA-CHis by affinity chromatography and anion exchange chromatography. Ni-NTA column: Lane. 1. SeeBlue Pre-Stained Protein Standard; Lane. 2. Load; 3. Flowthrough; Lane 4. Elution with buffer containing 10 mM imidazole; Lane 5. Elution with buffer containing 50 mM imidazole; Lane 6. Elution with buffer containing 250 mM imidazole. DEAE column: Lane 7. SeeBlue Pre-Stained Protein Standard; Lane 8. Load; Lane 9. Flowthrough; Lane 10. Eluate with buffer containing 1 M NaCl. The framed sections show the protein of interest.

Figure 2.5. Ni-NTA column purification of TmrA-MBP. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Load; Lane 3. Flowthrough; Lane 4. Wash; 5. Elution with buffer containing 10 mM imidazole; Lane 6. Elution with buffer containing 50 mM imidazole; Lane 7. Elution with buffer containing 250 mM imidazole. The framed sections show the protein of interest.

92

Refolding of the fusion proteins were carried out with and without cofactor

reconstitution. The eluate fractions were buffer-exchanged on a PD-10 column

equilibrated with a buffer containing 120 mM SDS. The SDS-unfolded protein was then

diluted 1:1 in a refolding buffer and incubated at room temperature for 24, 48, 72 and

96 hours. The enzyme mixture was then stored at -20 °C to stop the refolding reaction.

Dechlorination activity assays were then performed with the samples from each

refolding condition. No DCM formation was detected via gas chromatography,

suggesting the refolded proteins were not active. The negative controls with heat-

inactivated enzymes also showed no sign of dechlorination.

2.3.3 Aerobic purification and anaerobic cofactor reconstitution method

In this method, cell lysis and purification of TmrA fusion proteins were carried out

aerobically. Cells were lysed by sonication and inclusion bodies were solubilized with

buffer containing 8 M urea. Solubilized inclusion bodies were then loaded onto a Ni-NTA

column for purification. Binding of TmrA-CHis and TmrA-TF to Ni-NTA column was

confirmed by SDS-PAGE (Figure 2.6).

Figure 2.6. Purification of TmrA-CHis and TmrA-TF by affinity chromatography. TmrA-CHis: Lane 1. SeeBlue Pre-Stained Protein; Lane. 2. Load; Lane 3. Flowthrough; Lane 4. Wash; Lane 5. Eluate with buffer containing 250 mM imidazole; TmrA-TF: Lane 6. SeeBlue Pre-Stained Protein; Lane. 7. Load; Lane 8. Flowthrough; Lane 9. Wash; Lane 10. Eluate with buffer containing 250 mM imidazole. The framed sections show the protein of interest.

93

The reconstitution of the purified denatured TmrA-Chis and TmrA-TF was performed

inside an anaerobic chamber. Denatured TmrA was mixed with DTT, FeCl3, Na2S and

cobalamin in the denaturation buffer. The mixture was then loaded onto a PD-10

desalting column equilibrated with refolding buffer to remove the urea, and

unincorporated excess cofactors.

The cofactor reconstituted TmrA-CHis and TmrA-TF were used in dechlorination activity

assays described in section 2.2.5 where methyl viologen and chloroform were the

electron donor and electron acceptor, respectively. Heat-inactivated (100 ᵒC for 20 min)

TmrA-CHis and TmrA-TF were used as negative controls. The negative controls with

heat-inactivated enzymes also demonstrated some dechlorination of CF possibly

because of the presence of the corrinoid cofactors. However, the dechlorination was

much lower than the samples with the active enzymes in them. The abiotic

dechlorination demonstrated by the negative controls were subtracted from the

samples to calculate their specific activity. The calculated specific activity (average of

replicates) for TmrA-CHis and TmrA-TF was 59±34 (standard deviation) and 2.6±0.3

(range) nmol min-1 mg-1, respectively. The standard curves for CF and DCM used in the

calculations are shown in the appendices (A.4).

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Figure 2.7. CF dechlorination by A. TmrA-CHis, B. heat-inactivated TmrA-CHis, C. TmrA-TF, D. heat-inactivated TmrA-TF. The arrows indicate CF and DCM in the chromatogram. The circles show CF and DCM in the scan. DCM was detected in all four cases, but the amounts were significantly higher with the active enzymes than the heat-inactivated ones.

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2.4 DISCUSSION

The potential use of OHRB in bioremediation endeavours has propelled the study of

these microbes and their respective RDases in great detail. Over the past two decades a

number of efforts have been made to produce recombinant RDases in E. coli, the most

commonly used prokaryotic host for heterologous protein expression. Expression of

soluble and active respiratory RDase in E. coli has not been successful so far. Some

efforts have led to the over-expression of the RDase in interest, but they were

aggregated as inactive and misfolded inclusion bodies in the cytoplasm (Sjuts et al.,

2012, Jugder et al., 2018). E. coli cannot synthesize corrinoids de novo, the insufficient

amounts of cobamide cofactor in the cells presumably result in the formation of inactive

RDases (Suyama et al., 2002). However, two RDases have been recovered from inclusion

bodies expressed in E. coli: the vinyl chloride reducing VcrA from Dehalococcoides

mccartyi strain VS (Parthasarathy et al., 2015) and tetrachloroethene reducing PceA

from Geobacter sp. (Nakamura et al., 2018).

In this study no attempt was made to express soluble TmrA in E. coli; the sole focus was

to recover TmrA in a catalytically active form from the inclusion bodies expressed in E.

coli. The IPTG-inducible pOPIN vector constructs pOPIN-TmrA-CHis, pOPIN-TmrA-MBP

and pOPIN-TmrA-TF described in Jugder et al., (2018) were used for the purpose of

expressing the enzyme. The tmrA gene cloned into the pOPIN vector was devoid of the

TAT signal and predicted membrane spanning regions. The pOPIN-TmrA-CHis was

chosen because of the lack of an additional solubility tag. Maltose Binding Protein (MBP)

is one of the most frequently used fusion tags used to increase the solubility of

expressed recombinant proteins (Ki and Pack, 2020). There are reports where the fusion

of MBP tag to the protein of interest did not hinder the formation of inclusion bodies

but enhanced the recovery of soluble protein after refolding (Sachdev and Chirgwin,

1998a, Sachdev and Chirgwin, 1998b); hence pOPIN-TmrA-MBP construct was also

chosen for this study. A previously reported success in recovering a Trigger Factor (TF)-

tagged PceA from inclusion bodies to a catalytically active form (Nakamura et al., 2018)

prompted the choice of the pOPIN-TmrA-TF construct for this study.

96

Recombinant protein expression in bacterial hosts frequently results in the formation of

non-functional inclusion bodies. Recovery of functional protein typically involves

solubilisation of inclusion bodies with high concentrations of a chaotropic agent such as

urea or guanidine hydrochloride, and subsequent protein refolding by chromatographic

and non-chromatographic methods (Swietnicki, 2006). Another method involves

denaturing protein with the highly effective anionic detergent, sodium dodecyl sulfate

(SDS) and its subsequent removal with an amphipathic diol solvent 2-methyl-2, 4-

pentanediol (MPD) (Michaux et al., 2008a, Michaux et al., 2008b). The presence of MPD

enables the denatured protein to detach from the SDS molecules and refold into the

native state (Roussel et al., 2013). The SDS-MPD system has been successfully employed

in recovering proteins from inclusion bodies for membrane proteins, PagP and Omp2a;

enzymes such as human carbonic anhydrase II and lysozyme; single-peptide proteins,

Tryptophan-Cage (TrpCage) and Tryptophan Zipper 1 (TrpZip); viral capsid protein HBc-

MAGE3 II and Grapevine fanleaf virus (GFLV)-coat protein; voltage-dependent anion

channel protein, VDAC36 from plant cells (Lopes‐Rodrigues et al., 2020, Michaux et al.,

2008a, Michaux et al., 2008b, Michaux et al., 2016, Roussel et al., 2013, Yazdani et al.,

2019, Zhang et al., 2020).

The refolding efficiency of the SDS/MPD system is mainly influenced by temperature,

pH, protein concentration, SDS/MPD ratio and ionic strength (Roussel et al., 2013). In

the current study, refolding of both the pOPIN-TmrA-CHis and pOPIN-TmrA-MBP

constructs using the SDS-MPD system was unsuccessful. It has been established that a

high concentration of MPD is needed to alter the SDS denaturing effect on protein

(Michaux et al., 2016). In this study 3 M MPD was used to counter the denaturing effects

of 120 mM SDS which failed to produce positive outcomes. Roussel et al., (2013)

demonstrated full refolding of Omp2a trimeric membrane porin within four days of

incubation at 20 ᵒC. In the current study incubating TmrA in the refolding buffer at room

temperature up to 96 h yielded no dechlorination activity. Even incorporating a cofactor

reconstitution step with FeCl3, Na2S and cobalamin did not induce functionality. The

possible causes for the lack of proper refolding and/or catalytic activity could be the use

of an improper ratio of SDS/MPD, unsuitable ionic strength and/or improper

97

incorporation of Fe-S cofactors. Hence, further study screening these factors is required

for the MBP and TF- tagged TmrA enzyme.

The second method undertaken in this study involved a chromatographic step in the

presence of oxygen followed by an anaerobic cofactor reconstitution. A similar method

established by Nakamaru et al., (2018) resulted in active PceA from Geobacter sp. In the

current study, both pOPIN-TMrA-CHis and pOPIN-TmrA-TF constructs were successfully

refolded into catalytically active forms. The success of this method over the previously

described attempt could be attributed to the proper incorporation of the iron and sulfur

cofactors. In the previous method, where FeCl3 and Na2S buffers were added in 5- and

10-fold molar excess respectively, the resulting refolded TmrA remained inactive. In this

experiment FeCl3 and Na2s were added in 50-fold molar excess and the refolded TmrA

showed distinct dechlorination activity. The negative controls with the heat-inactivated

TmrA also showed dechlorination, possibly due to the presence of cobalamin in the

sample, which is capable of abiotic dechlorination, but it was significantly lower (p <

0.05) than the non heat-inactivated enzymatic dechlorination samples. An extended

incubation of the dechlorination reaction mixtures for over 12 h showed that both CF

and DCM diminished (data not shown), hinting at further breakdown of DCM.

One of the major limitations of studying RDases is their sensitivity towards oxygen. In

this experiment the cell lysis and purification were carried out under aerobic conditions,

but the enzyme still retained dechlorination activity after cofactor reconstitution inside

an anaerobic chamber. Studies have reported that oxygen sensitivity of Fe-S cluster

containing enzymes is attributed to the destruction of their catalytically active [4Fe-4S]

cluster (Bruska et al., 2013, Flint et al., 1993a, Flint et al., 1993b, Khademian and Imlay,

2020). Oxygen converts exposed Fe‐S clusters to unstable forms which releases iron and

renders the enzyme inactive (Imlay, 2006). In this study, the Fe-S reconstitution took

place under anaerobic conditions which likely contributed to the recovery of active

enzyme despite being purified under aerobic conditions.

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2.5 CONCLUSIONS

For the past two decades, heterologous expression of soluble and functional RDase in E.

coli has proven to be a difficult task. A number of studies have failed to produce soluble

RDase in E. coli although over-expression in the form of inclusion bodies has been a

common outcome. Recovery of active RDase from inclusion bodies have proven to be

difficult because of the improper and insufficient incorporation of cofactors.

Maintaining anaerobic conditions during all steps after cell harvest requires purging all

puffers used with N2 and the use of an anaerobic chamber which adds to the complexity

of the procedure. The current study demonstrates successful recovery of active TmrA

from inclusion bodies expressed in E. coli. Only two other recombinant RDases have

been previously recovered from inclusion bodies expressed in E. coli. The cell lysis and

purification of RDase in presence of oxygen in this study makes the process much less

complicated and more achievable. Future work regarding this system could include

condition optimization for cofactor reconstitution, kinetic studies of the successfully

refolded and active recombinant TmrA, use of additional protein-free controls to

conclusively demonstrate the difference between protein activity and abiotic

dechlorination, incubation for longer periods to determine the fate of DCM, the level of

which was demonstrated to decline over time in the current study, other

characterization of the reconstituted TmrA such as estimation of cofactor content and

oxygen sensitivity. This method can also be extended to other expression hosts such as

B. megaterium and S. blattae in which successful soluble expression of RDases has been

reported but also where the majority of the recombinant enzyme is produced as

insoluble inclusion bodies. This aerobic purification and anaerobic cofactor

reconstitution strategy could also be employed to the recombinant production of similar

proteins which are otherwise arduous to express heterologously.

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CHAPTER THREE: Production of functional recombinant chloroform reductive dehalogenase in Bacillus megaterium: improving existing findings

102

3.1 INTRODUCTION

The failure to express soluble and functional RDases in E. coli (Jugder et al., 2018, Kimoto

et al., 2010, Neumann et al., 1998, Sjuts et al., 2012, Suyama et al., 2002) has been

attributed to the inability of E. coli to synthesize corrinoids de novo and therefore there

were insufficient amounts of cobamide cofactor in the cells to support RDase expression

(Neumann et al., 1998; Suyama et al., 2002). Gram-positive Bacillus megaterium

produces 5,6-dimethylbenzimidazolyl cobamide, the standard-type B12 cofactor,

through an oxygen-independent adenosylcobalamin pathway and has been identified as

a suitable host for heterologous production of functional RDases (Eppinger et al., 2011,

Payne et al., 2015, Wolf and Brey, 1986a).

B. megaterium is a rod-shaped, Gram-positive, mainly aerobic spore-forming bacterium.

It is commonly regarded as a soil bacterium though it can be found in diverse habitats

(Vary et al., 2007). With the dimension of 4 x 1.5 µm, B. megaterium is one of the biggest

known bacteria (Bunk et al., 2010). The ability to grow on different carbon sources and

over a wide range of temperatures has made this non-pathogenic bacterium an

important industrial organism for the production of different vitamins, enzymes and

drugs for decades (Goswami et al., 2018, Grage et al., 2017, Korneli et al., 2012, Vary et

al., 2007). The stable plasmid replication system, absence of endotoxins, lack of alkaline

proteases and protein secretion capacity makes B. megaterium an amenable host for

producing recombinant proteins (Gamer et al., 2009, Korneli et al., 2013).

The first report of soluble and functional expression of reductive dehalogenase in B.

megaterium was described by Payne et al., (2015). NpRdhA is an oxygen-tolerant ortho-

dibromophenol reductase from Nitratireductor pacificus pht-3B. It is a soluble

cytoplasmic enzyme lacking both the twin-arginine signal and the associated

transmembrane rdhB protein. Payne et al., (2015) expressed NpRdhA in B. megaterium

MS941 containing pT7-RNAP plasmid which allows xylose-inducible expression of T7

polymerase.

103

Jugder et al., (2018) successfully expressed TmrA in a soluble and functional form in B.

megaterium MS941. Xylose-inducible expression of TmrA was done aerobically in TB

media at 17 °C. The NHis-tagged TmrA was purified by first loading onto a Ni-NTA drip

column (anaerobic condition) followed by an anion exchange ResQ column (in the

presence of oxygen). The ResQ-purified TmrA had a specific activity of 110 U mg of

protein−1 which is 11-fold lower than the activity of native TmrA (1.27 × 103 U mg of

protein−1) determined previously (Jugder et al., 2017). The lower specific activity of the

recombinant TmrA was attributed to either the incorrect incorporation of corrinoid

cofactors, the absence of a trigger factor to facilitate correct folding of the enzyme or

partial oxidation of the enzyme during purification.

This study aimed to manipulate and screen different culture conditions in order to

improve the soluble expression and specific activity of TmrA in B. megaterium from what

has been described by Jugder et al., (2018). The B. megaterium transformant containing

NHis-TmrA used in this study was received from Jugder et al., (2018).

Figure 3.1. TmrA fusion construct used in B. megaterium expression using pPT7 vector.

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In the study of Jugder et al., (2018) B. megaterium cell transformants were cultured in

Terrific Broth (TB) at 37°C/180 rpm, at an OD578 nm of 0.4-1.0. The culture was

supplemented with 50 μM ammonium iron (II) sulfate, 1 μM B12 and induced with 0.5%

xylose. After growing overnight at 17°C/180 rpm, the culture was harvested by

centrifugation (4°C, 7,000 g for 10 min). In the current study, different culture conditions

were screened to determine if they had any significant influence on the soluble

expression and specific activity of the TmrA expressed in B. megaterium. The screening

conditions were determined after reviewing literature on recombinant protein

expression in B. megaterium. The screenings were carried out in factorial design

experiments. A full factorial design considers all the possible combinations with all levels

of all the factors screened whereas a fractional factorial design consists of a carefully

considered subset of a full factorial design.

3.2.1 Preliminary screening

The different factors and levels considered in this step are described in Table 3.1. A

fractional factorial design was generated using the JMP software shown in the result

section (Table 3.3). The 20 different combinations of culture conditions were each done

in duplicate and the culture volume was 25 mL carried out in 250 mL baffled flasks. The

pre-induction cultures of all the conditions were maintained at 37 °C/180 rpm. At the

end of the culture cell pellets were harvested by centrifugation at 8,000 g for 10 min at

4 ᵒC, washed with 1 x PBS buffer and stored at -20 °C for future use.

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Table 3.1 Culture conditions for primary screening: factors and levels

Factors Levels

Growth media Luria broth (LB)

Terrific broth (TB)

Induction OD600 nm 0.3

0.6

Xylose concentration (%) 0.1

0.3

0.5

Length and temperature of induction Up to OD600 nm 1.5 (37 °C)

3 hrs (37 °C)

6 hrs (30 °C)

Overnight (17 °C)

3.2.2 Second screening: full factorial design

For the second step of screening, different concentrations of xylose and induction

temperature and lengths (Table 3.2) were considered. The different combinations for a

full factorial design were determined using JMP software (Table 3.4). Expression

experiments were carried out in duplicate in 5 mL TB cultures in 50 mL tubes. Bacteria

was grown at 37 °C/180 rpm, when the OD600 reached 0.6, 50 μM ammonium iron (II)

sulfate, 1 μM B12 was added to the culture and induced with xylose. Cell pellets were

harvested by centrifugation (8,000 g for 10 min at 4 ᵒC) and after washing with 1 x PBS

buffer were stored at -20 °C for further use.

Based on the findings, further screening was carried out with 0.03, 0.01, 0.003, 0.001

and 0% xylose induction for 3 h. The other culture conditions remained the same.

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Table 3.2 Culture conditions for second screening: factors and levels

Factors Levels

Xylose Concentration (%) 0.1

0.3

0.5

Length and temperature of induction 3 h (37 °C)

Overnight (17 °C)

3.2.3 SDS-PAGE

Samples collected before and after induction for each screening condition were

normalised according to an OD600 of 1.0 and chemically lysed using BugBuster Protein

Extraction Reagent (Merck Millipore) according to the manufacturer’s instructions.

Proteins were mixed with 4x NuPAGE® gel loading buffer, heated at 95 °C for 5 min and

centrifuged before loading on a Bolt 4-12% Bis-Tris Plus polyacrylamide precast gel

(Thermo Scientific). SeeBlue Plus2 Prestained Standard (Thermo Scientific) was used as

the molecular weight marker and the gel was stained with GelCode Blue Stain Reagent

(Thermo Scientific).

3.2.4 Western blot analysis

Following SDS-PAGE the unstained NHis-tagged proteins were transferred to a

nitrocellulose membrane using the iBlot2 Gel Transfer Device (Life Technologies). The

membrane was blocked overnight with 5% non-fat dry milk in 1 x PBS. After washing in

1 x PBS containing 0.05% TWEEN-20 (Sigma), primary antibody incubation was done for

2 h using the Monoclonal Anti polyHistidine-Peroxidase clone HIS-1 (Sigma, Catalog

Number A7058) at a dilution of 1: 2,000 in PBS containing 0.05% TWEEN-20 and 1%

bovine serum albumin. The membrane was washed in washing buffer and His-tagged

proteins were detected using Amersham™ ECL Select™ Western Blotting Detection

Reagent and the images were visualized by ImageQuant LAS 500.

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3.2.5 Determination of protein concentration

The concentration of the soluble cell fractions was measured using absorbance at 280

nm by a NanoDrop One (Thermo Fisher Scientific). The molar extinction coefficient

ε280nm of 72,140 M-1cm-1 was calculated from the primary amino acid sequence using

ProtParam-ExPaSy.


The activity assay was performed as described in section 2.2.5. TmrA dechlorination of

CF by the soluble protein fractions was measured by GC/MS using titanium (III) citrate

and methyl viologen as electron donors. Heat-inactivated TmrA (100 °C for 20 min) was

used as a negative control.

3.2.7 Large-scale culture and cell lysis

1 L cultures were carried out under the finally decided upon conditions. TB media

containing 10 µg/mL tetracycline and 4.5 µg/mL chloramphenicol was inoculated to a

cell density (OD600) of 0.1 with B. megaterium containing NHis-TmrA. The culture was

maintained at 37 ᵒC and 180 rpm. When the OD600 reached 0.6 the culture was

supplemented with 50 μM ammonium iron (II) sulfate and 1 μM B12 and induced with

0.03% xylose. After 3h, the cells were harvested by centrifugation at 8,000 g for 10 min

at 4 ᵒC. The cells were washed with 1 x PBS and stored at -20 ᵒC until further processing.

The cells were lysed chemically under anaerobic conditions; all the buffers used were

purged with N2, addition of buffer and chemicals to the cell pellet and decanting of

soluble portions were done inside the anaerobic chamber. The cell pellet was

resuspended in a lysis buffer (50mM Tris–HCl pH 8, 17mM NaCl, PMSF tablets, 250

μg/mL lysozyme) purged with N2 at a ratio of 1:5 for 20 minutes at 25 °C. For further cell

breakdown, 1 mg/mL of sodium deoxycholate was added to the cell suspension and

incubated at 37 °C for 60 min with constant shaking, followed by the addition of 25

μg/mL of DNase I (Sigma Aldrich D5025) for 60 minutes at 25 °C. The cell suspension was

then centrifuged at 14,000 g for 20 minutes at 4 °C to collect the soluble fraction.

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3.2.8 Purification of NHis-TmrA

For purification of NHis-TmrA from the soluble fraction of B. megaterium, affinity

chromatography with 1 mL Ni-NTA Superflow column and anion exchange

chromatography with 1 mL HiTrap™ Q FF were carried out. The purification was done

inside an anaerobic chamber (Baker Ruskinn Concept) and all buffers were made anoxic

by purging with N2.

The soluble fraction was loaded onto a 1 mL Ni-NTA Superflow Column using a peristaltic

pump (Gilson Minipuls 2) calibrated to a flowrate of 1 mL/min. The column was

previously equilibrated with 10 CV of buffer A (50 mM NaH2PO4, 300 mM NaCl, 10 mM

imidazole, pH 8.0). Following a wash step with 10 CV of Buffer B (50 mM NaH2PO4, 300

mM NaCl, 20 mM imidazole, pH 8.0), NHis-TmrA was eluted with 5 CV of Buffer C (50

mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8.0).

For anion exchange chromatography, a 1 mL HiTrap™ Q FF column was equilibrated with

10 CV of buffer A (100 mM Tris, pH 8, 10% glycerol). The soluble fraction was then loaded

onto the column, followed by a wash step with 10 CV of the same buffer. Gradient

elution of the protein was performed with buffer containing 0 to 1 M NaCl. The flowrate

was always maintained at 1 mL/min by using a peristaltic pump.

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3.3 RESULTS

3.3.1 Primary screening of culture conditions

To assess the impact of different culture conditions on the soluble expression of TmrA

in B. megaterium four factors were selected: growth media, induction OD600,

concentration of xylose used for induction and length of induction. Two types of culture

media (LB and TB), two induction OD600 (0.3 and 0.6), three xylose concentrations (0.1%,

0.3%, 0.5%) and four induction temperature and lengths (up to OD600 1.5 at 37 °C, 3hrs

at 37 °C, 6hrs at 30 °C and overnight at 17 °C) were considered. The fractional factorial

design consisted of 20 different culture conditions using JMP software shown in Table

3.3. Each condition was done in duplicate in 25 mL cultures in 50 mL tubes.

The cell pellets harvested from each culture condition were fractionated to soluble and

insoluble parts and were analysed by western blotting (shown in Appendices). While no

conclusion could be reached on identifying an optimal culture condition based on the

western blot results, the different culture conditions demonstrated varying levels of

soluble and insoluble expression of recombinant TmrA which confirmed that the soluble

expression could be controlled by differing culture conditions. The conditions producing

higher soluble expression are highlighted in Table 3.3.

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Table 3.3 Primary screening conditions generated by JMP software

Media Type Induction OD600

Xylose Conc.* (%)

Length of Induction

1 LB 0.3 0.3 Up to OD600 1.5 at 37 °C

2 LB 0.6 0.3 Up to OD600 1.5 at 37 °C

3 LB 0.3 0.3 6 h at 30 °C

4 LB 0.3 0.5 6 h at 30 °C

5 LB 0.6 0.3 3 h at 37 °C

6 LB 0.6 0.5 3 h at 37 °C

7 LB 0.3 0.1 3 h at 37 °C

8 LB 0.6 0.1 Overnight at 17 °C



11 TB 0.3 0.1 Up to OD600 1.5 at 37 °C

12 TB 0.6 0.3 Up to OD600 1.5 at 37 °C

13 TB 0.3 0.5 6 h at 30 °C

14 TB 0.6 0.3 6 h at 30 °C

15 TB 0.6 0.3 3 h at 37 °C

16 TB 0.6 0.5 3 h at 37 °C

17 TB 0.3 0.1 3 h at 37 °C

18 TB 0.6 0.5 Overnight at 17 °C



*Due to miscalculation during xylose stock solution preparation, the actual concentrations added were 0.001%, 0.003% and 0.005%. This mistake was identified in later screenings.

111

Miscalculation in preparing the xylose stock solution during the primary screening led to

an unexpected finding. Instead of adding the decided upon xylose concentrations, much

lower concentrations (100 x lower) were added which resulted in recombinant TmrA

with much higher specific activity. This led to a second phase of screening focusing on

xylose concentration and length and temperature of induction.

3.3.2 Second screening of culture conditions

A full factorial design (Table 3.4) with 3 xylose concentrates and 2 induction temperature

and lengths (3 h at 37 ᵒC and overnight at 17 ᵒC) were carried out in duplicate in 5 mL of

TB media in 50 mL tubes.

Table 3.4 Second screening conditions

Xylose Conc. (%) Induction Temperature and Length

1 0.3 17 ᵒC, overnight

2 0.5 37 ᵒC for 3 h



5 0.1 37 ᵒC for 3 h

6 0.3 37 ᵒC for 3 h

SDS-PAGE shows a band consistent with the expected molecular weight of soluble NHis-

TmrA (Figure 3.2), however, a band is also expressed in the native strain at the same

molecular weight. Western blot and activity assays, however, did confirm expression of

functional TmrA in the transformed strain (Appendix, A.5). An example of the

chromatogram for the dechlorination activity assay is shown in Figure 3.3. The specific

activity of the recombinant TmrA increased with decreasing xylose concentration when

induced for 3 h at 37 ᵒC, however that was not the case when induction was done at 17

ᵒC overnight (Figure 3.4). The prediction profiler with the JMP software showed that

among the six conditions tested, the desirability is highest (0.291) when induction is

done with 0.1% xylose for 3 h at 37 ᵒC (Figure 3.5).

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Figure 3.2. SDS-PAGE of the soluble protein fractions from cell pellets obtained from the second screening conditions. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Native B. megaterium; Lane 3. Pre-induction sample; Lane 4. 0.1% xylose, 3 h induction; Lane 5. 0.1% xylose, overnight induction; Lane 6. 0.3% xylose, 3 h induction; Lane 7. 0.3% xylose, overnight induction; Lane 8. 0.5% xylose, 3 h induction; Lane 9. 0.5% xylose, overnight induction. The framed section shows the protein of interest.

Figure 3.3. Dechlorination of CF with B. megaterium cell lysates. All of the 0.5 mM CF added was converted to DCM in this experiment.

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Figure 3. 4. Effect of different culture conditions on specific activity (n=2) of TmrA. The range bars show the lowest and highest value for each condition. o/n refers to overnight.

Figure 3.5. Prediction profiler generated with JMP software.

Since induction at 3 h for 37 ᵒC resulted in more soluble protein the next screenings with

lower xylose concentrations were done at this induction length and temperature. When

induced for 3 h at 37 ᵒC, lowering the xylose concentrations resulted in better specific

activity of the recombinant NHis-TmrA. Also, in the primary screening where

accidentally much lower xylose concentrations were added, specific activity was high,

114

so further screening was done with no xylose and 10 x and 100 x dilutions of the 0.3%

and 0.1% xylose used in the previous step. SDS-PAGE showed that all the xylose

concentrations tested yielded soluble NHis-TmrA (Figure 3.6). The specific activity with

cell lysates from these culture conditions are presented in Figures 3.7. The prediction

profiler generated with JMP showed that the desirability was highest (0.496) when

induction was done with 0.03% xylose (Figure 3.8).

Figure 3.6. SDS-PAGE of the soluble protein fractions from cell pellets obtained from cultures with reduced xylose concentrations. Lane 1. SeeBlue Pre-Stained Protein Standard, Lane 2. Native B. megaterium; Lane 3. Pre-induction sample; Lane 4. No xylose; Lane 5. 0.01% xylose; Lane 6. 0.001% xylose; Lane 7. 0.03% xylose; Lane 8. 0.003% xylose. The framed section shows the protein of interest.

115

Figure 3.7. Effect of culture conditions on specific activity (n=2) of TmrA. The range bars show the lowest and highest value for each condition.

Figure 3.8. Prediction profiler generated with JMP for induction with different xylose concentrations.

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3.3.3 Purification of NHis-TmrA

Purification was carried out with cell lysates derived from 1 L culture induced with 0.03%

xylose for 3 h at 37 ᵒC. In case of Ni-NTA column purification, the protein did not bind

very well to the column and most of it came out with the flowthrough (Figure 3.9). With

Q FF column purification, gradient elution was done with buffer containing 0 to 1 M

NaCl. NHis-TmrA eluted with buffer containing 0.1-0.4 M NaCl (Figure 3.10). The

concentration of NHis-TmrA was calculated using absorbance at 280 nm with a molar

extinction coefficient value of 72,140 M-1 cm-1 (calculated from the primary amino acid

sequence using ProtParam-ExPaSy). The estimated yield of TmrA from a 1L culture was

approximately 50 mg.

CF dechlorination assay was carried out with the partially purified NHis-TmrA. The

calculated specific activity was (3.6 ± 0.3) x 103 nmol/min/mg protein which is 2.8 times

higher than the specific activity for native TmrA calculated by Jugder et al., (2017) and

32.7 times higher than specific activity of the recombinant TmrA expressed in B.

megaterium by Jugder et al., (2018).

Figure 3.9. Ni-NTA column purification. Lane 1. SeeBlue Pre-Stained Protein Standard, Lane 2. Load, Lane 3. Flowthrough, Lane 4. Wash, Lane 5. Elute with buffer containing 250 mM imidazole.

117

Figure 3.10. Q FF column purification. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Load; Lane 3-5. Flowthrough; Land 6 & 7. Wash; Lane 8. Elute with 0.1 M NaCl; Lane 9. Elute with 0.2 M NaCl; Lane 10. Elute with 0.3 M NaCl; Lane 11. Elute with 0.4 M NaCl; Lane 12. Elute with 0.5 M NaCl; Lane 13. SeeBlue Pre-Stained Protein Standard; Lane 14. Elute with 0.6 M NaCl; Lane 15. Elute with 0.7 M NaCl; Lane 16. Elute with 0.8 M NaCl; Lane 17. Elute with 0.9 M NaCl; Lane 18. Elute with 1 M NaCl. The framed section shows the protein of interest.

118

3.4 DISCUSSION

CF reductive dehalogenase, TmrA, has been successfully produced heterologously in a

soluble and active form in B. megaterium by Jugder et al., (2018). He cloned a tmrA gene

devoid of the TAT signal and membrane spanning region, with a His-tag at the N-

terminal, into a pPT7 plasmid and transformed into B. megaterium MS941 already

containing a pT7-RNAP plasmid that allows xylose-inducible expression of T7

polymerase. Their study resulted in the soluble expression of TmrA with the specific

activity of 110 U/mg of protein with was 11-fold lower than the specific activity of the

native TmrA (1.27 x 103 U/mg of protein) measured by Jugder et al. (2017). In the present

study the B. megaterium MS941 transformant produced by Jugder et al., (2018) was

used in multiple screening experiments with an aim to improve the productivity of the

process with respect to protein yield and potentially improve the specific activity of the

expressed recombinant protein.

In the xylose-inducible plasmid system used in B. megaterium, the gene of interest is

under the control of the xylose inducible promoter PxylA with a down-stream multiple

cloning site (Korneli et al., 2013). PxylA is located upstream of the operon consisting of

genes for xylose isomerase XylA, xylulokinase XylB, and xylose permease XylT (Rygus and

Hillen, 1991). XylA and XylB are responsible for the phosphorylation of xylose to form

xylose-5-phosphate, XylT aids in transporting xylose into the cell (Biedendieck et al.,

2011). The gene encoding the repressor protein XylR is divergently oriented upstream

of the operon, resulting in an overlap of the promoter regions of xylR and xylABT operon

(Biedendieck et al., 2010). In the absence of xylose, XylR binds to the operator sequences

located in PxylA and inhibits transcription of the xylABT operon (Gärtner et al., 1988).

When xylose is present, it binds to the repressor XylR causing a conformational change

of XylR which prevents promoter binding (Biedendieck et al., 2011). In the double vector

system used in the present study, the t7 rnap gene is under the control of the strong

PxylA promoter and pPT7 controls the T7 RNAP-dependent expression of the target gene

(Gamer et al., 2009, Korneli et al., 2012). The stable plasmid replication system of B.

megaterium aids in the stable co-existence of the two vectors (Rygus and Hillen, 1991).

119

Although Gamer et al., (2009) demonstrated a tight regulation of xylose-inducible

promoter in the double vector system, in the current study expression of soluble TmrA

without xylose induction was fairly high (the concentration was about 429 mg/mL in cell

lysate), indicating leaky expression. The dechlorination of CF using the cell lysates of

non-induced samples was higher compared to the frequently used xylose concentration

of 5 g/L described in literature (data not shown). Stammen et al. (2010a), (2010b)

reported leaky expression of GFP in B. megaterium before the addition of xylose.

(Marchand and Collins, 2013) described leaky expression from xylose-inducible

promoter controlling the expression of genes agrC (gene encoding transmembrane

histidine kinase receptor in Staphylococcus aureus) and agrA (gene encoding

transcriptional activator in S. aureus) in B. megaterium.

The most frequently described xylose concentration used for the induction of

recombinant protein expression in B. megaterium ranges from 0.1 to 0.5 % (Biedendieck

et al., 2011, Biedendieck et al., 2010, Burger et al., 2003, Gamer et al., 2009, Nehru et

al., 2020, Payne et al., 2015). However, in the current study using much lower

concentrations produced more soluble and active TmrA. The best results were obtained

when induced with 0.03% xylose (much lower than the most commonly used

concentration of 0.5%) for 3 h. However, protein expression under xylose inducible

promotors with very low xylose concentrations have been reported in B. subtilis.

(Bhavsar et al., 2001) developed a xylose-inducible promotor for B. subtilis where the

xyl system-controlled protein expression over a xylose concentration range of 0.0002 to

0.63%. Xylose-inducible expression of protein with very little xylose concentration (0.1

to 10 mM) have been reported in Staphylococcus carnosus (Wieland et al., 1995),

Caulobacter crescentus (Meisenzahl et al., 1997), Hypocrea jecorina (Mach-Aigner et al.,

2010), Aspergillus niger (Mach-Aigner et al., 2012).

The recombinant TmrA expressed in this study had a His-tag at the N-terminal to

facilitate purification via affinity chromatography. However, there was little to no

binding of NHis-TmrA to the Ni-NTA column even though Western Blot analysis

confirmed the presence of the His-tag in the protein prior to loading the cell lysate onto

the column (Figure in Appendices, A.5.5). The NHis-TmrA produced in B. megaterium by

120

Jugder et al., (2018) was purified by affinity chromatography using a Ni-NTA column,

followed by anion exchange chromatography using a ResQ column. The failure of NHis-

TmrA produced in the current study to bind to the Ni-NTA column might be attributed

to a difference in the folding of TmrA after translation due to the shorter induction

period at a higher temperature (3 h at 37 ᵒC in contrast to overnight at 17 ᵒC) which

might have made the His-tag inaccessible to the Ni-NTA column. The effect of

temperature on protein folding has been described by multiple authors (Jagielska and

Scheraga, 2007, Tikhonov et al., 2002, Xu et al., 2012). In this study, TmrA was partially

purified via anion exchange chromatography using a Q FF column. Gradient elution

showed that elution mostly occurred with 100 – 400 mM NaCl. Purification was

performed under anaerobic conditions, at no point of processing, from cell lysis to

dechlorination assay, was TmrA exposed to air.

The calculated specific activity of purified TmrA in this study was (3.6 ± 0.3) x 103

nmol/min/mg of protein which is 2.8 times higher than the specific activity calculated

for native TmrA (1.27 x 103 nmol/min/mg of protein). Similar results were described by

Parthasarathy et al. (2015) where the recombinant VcrA recovered from inclusion

bodies expressed in E. coli, exhibited higher specific activity than the native VcrA from

Dehalococcoides sp. Strain VS. The enhanced specific activity of the recombinant TmrA

might be attributed to the lack of the TAT signal at the N-terminal and the resulting

conformational changes it causes. Enhanced lytic activity upon removal of the binding

domain has been reported for multiple enzymes (Fenton et al., 2010) A truncated

derivative of the phage endolysin LysK exhibited two-fold higher lytic activity than the

native enzyme (Horgan et al., 2009). The specific activity of recombinant TmrA in this

study is 32.7-fold higher the specific activity of recombinant TmrA expressed in B.

megaterium by Jugder et al., (2018). This might be attributed to the oxygen-sensitivity

of the enzyme. In the study by Jugder and co-workers, cell lysis was done using a French

press whereas in the current study the cells were lysed chemically which offered more

control in maintaining anaerobic conditions. More importantly in the previous study the

second purification step was done aerobically; in the current study the single

purification step was carried out inside an anaerobic chamber. The maintenance of

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anaerobic conditions throughout the entire process in this study ensured that activity

loss due to oxygen-sensitivity was minimized.

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3.5 CONCLUSIONS

B. megaterium is proving to be a suitable host for the expression of soluble and

catalytically active RDases. Both respiratory and catabolic RDases have been successfully

expressed in B. megaterium. This study is first to report the screening of culture

conditions for the recombinant expression of RDases in B. megaterium. The findings

clearly demonstrate that varying the culture conditions impact both the level of soluble

expression and the specific activity of the expressed RDase. The recombinant TmrA

expressed in this study had a specific activity higher than the native enzyme which is a

vast improvement than the previous attempts. Further studies might include enzyme

kinetics of the recombinant TmrA, finer tuning of the xylose-inducible promotor to

reduce the non-inducible expression and elucidation of the role of the removal of the

TAT signal on the structure and function of RDases.

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KORNELI, C., BOLTEN, C. J., GODARD, T., FRANCO‐LARA, E. & WITTMANN, C. 2012. Debottlenecking recombinant protein production in Bacillus megaterium under large‐scale conditions—targeted precursor feeding designed from metabolomics. Biotechnology and Bioengineering, 109(6): 1538-1550.

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CHAPTER FOUR: Heterologous expression of chloroform reductive dehalogenase TmrA in Shimwellia blattae: co-expression with chaperone proteins

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4.1 INTRODUCTION

Shimwellia blattae is an enteric bacterium isolated from the hindgut of a cockroach

(Burgess et al., 1973). S. blattae can uptake and synthesize B12 de novo aerobically and

anaerobically and the molecular tools and techniques developed for Escherichia coli can

be used with this organism (Andres et al., 2004). Hence, S. blattae has been identified

as a promising alternative to the standard host E. coli for expressing cobalamin

containing proteins. S. blattae has been successfully employed in expressing

recombinant proteins where otherwise the requirement of adding high-cost vitamin B12

are considered as major drawbacks (Andreeßen et al., 2014, Heinrich et al., 2013, Sato

et al., 2015, Urano et al., 2015).

The failure to express soluble and functional reductive dehalogenases (RDases) in E. coli

(Jugder et al., 2018, Kimoto et al., 2010, Neumann et al., 1998, Sjuts et al., 2012, Suyama

et al., 2002) led to the use of expression hosts capable of producing corrinoids de novo.

The first demonstration of heterologous expression of soluble and active RDase was

PceA from Desulfitobacterium hafniense strain Y51 and RdhA3 of strain DCB-2 produced

in Shimwellia blattae (Mac Nelly et al., 2014). Co-expression of the RDases with their

respective chaperone proteins, PceT and RdhT improved over-expression of the

functional enzymes. Adding the corrinoid hydroxocobalamin and 5,6-

dimethylbenzimidazole to the growth media and using glycerol as the growth substrate

for S. blattae also increased the formation of active RDases. Kunze et al. (2017) produced

functional DcaA from D. dichloroeliminans DCA1 in S. blattae, increased dechlorination

activity was achieved by co-expression of chaperone protein, DcaT.

The minimal rdhAB gene cluster in OHRB is frequently associated with a varied set of

accessory genes, the exact function of most of them still unknown. One of these genes

is the rdhT which encodes a trigger factor-like protein presumed to assist in folding and

maturation of RdhA (Smidt et al., 2000). The PceT from D. hafniense Y51 and TCE1 binds

specifically to the TAT signal sequence of PceA which possibly delays the translocation

of PceA and allows the proper folding and incorporation of cofactors (Maillard et al.,

2011, Morita et al., 2009). RdhT lacks the N-terminal ribosome-binding domain of a

classical trigger factor chaperone protein and thus evolves toward a specific substrate,

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the RdhA (Maillard et al., 2011). In some OHRB the rdhT gene is adjacent to the rdhAB

operon, such as the Desulphitobacterium hafniense strains, but in others it is not found

in any rdhA related operons and is located elsewhere in the genome; the tmrAB operon

of the strain Dehalobacter sp. UNSWDHB (Figure 4.1) is an example (Jugder et al., 2015).

The transcription and function of the rdhT genes located remotely from the rdhAB

operon are mostly unknown.

Figure 4.1. tmrAB operon of Dehalobacter sp. UNSWDHB (adapted from Jugder et al., 2015).

The RDases so far expressed functionally in S. blattae have also been co-expressed with

their respective chaperone RdhT proteins (Mac Nelly et al., 2014; Kunze et al., 2017). In

all three successful cases the RdhA had a chaperone RdhT in the operon structure that

helped in RdhA maturation. To date there have been no reports on RdhA enzymes

encoded in rdh gene clusters lacking genes for a specific folding helper protein expressed

in S. blattae, with or without the co-expression of a chaperone protein.

This chapter describes the expression of TmrA with and without chaperone proteins to

determine their influence on the level of soluble expression of TmrA and hence the

specific activity. Four trigger factors have been identified in the genome of Dehalobacter

sp. UNSWDHB (Accession numbers WP_015043727, WP_034378503, WP_034378507,

WP_015044672). Since there are no trigger factors in the tmrAB operon to which the

function of rdhT could be assigned, we chose two trigger factors for this experiment: the

trigger factors belonging to contig_104 (WP_015043727) and contig_161

(WP_015044672) based on discussions with Dr Haluk Ertan. The trigger factor from

contig_104 (TF104) is found in Dehalobacter sp. UNSWDHB, Dehalobacter sp. CF,

Dehalobacter sp. DCA and in the last two species they have been identified as putative

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PceT. The trigger factor in contig_161 (TF161) is also found in the same three

Dehalobactor species and is identical to the cell division trigger factors in the respective

species. The cell division trigger factor in Dehalobactor sp. UNSWDHB and Dehalobacter

sp. CF is upregulated during chloroform respiration (Jugder et al., 2016, Tang and

Edwards, 2013).

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4.2.1 Plasmid construction

For heterologous expression of TmrA in S. blattae both the tmrA gene including the TAT

signal (NCBI Reference Sequence: WP_034377773.1 GI:736352001), which will be

refered to as tmrA_full throughout the study, and the TAT signal removed tmrA as used

in the previous two chapters, were used. Codon optimized gBlocks of tmrA_full, tf104

and tf161 were ordered from Integrated DNA Technologies. The pD864-TmrA construct

described in (Jugder et al., 2018) was used as the DNA template for the TAT signal

removed tmrA.

Two expression vectors were used in this study: pASK-IBA63c-plus, where the expression

cassette is under the transcriptional control of tetracycline promotor and pBBR1MCS-2

controlled by Lac promoter. Both vectors have been previously described for

heterlogous gene expression in S.blattae. The vector maps for both plasmids are found

in the Appendices.

4.2.1.1 PCR

The vectors used in this experiment were linearized using a polymerase chain reaction

(PCR). The insert genes were amplified to include required overhangs and restriction

sites to facilitate assembly. Each reaction volume of 25 µL contained 12.5 µL Q5 High-

Fidelity 2X Master Mix (New England Biolabs, Ipswich, MA, USA), 0.5 µM of forward and

reverse primers, ~100 ng of template DNA and the required amount of nuclease-free

water to make up the reaction volume. The amplification reactions were carried out in

a BIO-RAD C1000 TouchTM Thermal Cycler using the following cycling conditions: Initial

denaturation at 98 °C for 30 s, 35 cycles of 10 s denaturation at 98 °C, 30 s at respective

annealing temperature (the annealing temperature was determined using the NEB Tm

calculator, https://tmcalculator.neb.com/) and 30 s per kb extension at 72 °C, followed

by a final extension of 3 min at 72 °C. Where the annealing temperature determined by

NEB Tm calculator failed to generate the desired gene product or resulted in unwanted

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non-specific bindings, temperature gradients were used to determine the best

annealing temperature.

The primers used in different PCR reactions used in this study are listed in Table 4.1.

Primers were purchased from Sigma-Aldrich.

4.2.1.2 PCR purification

All PCR products went through a purification step to remove primers and impurities

using a PCR purification Kit (Qiagen) following the manufacturer’s instructions.

4.2.1.3 DpnI digestion

The purified PCR products were treated with DpnI (NEB) to remove the methylated

template DNA. Digestion was carried out in 1X CutSmart® Buffer for 1 h at 37 °C followed

by an enzyme inactivation step at 80 °C for 20 min.

4.2.1.4 Gibson assembly

The tmrA gene without the TAT signal was cloned into vectors pASK-IBA63c-plus and

pBBR1MCS-2 using a Gibson assembly method (Gibson et al., 2010, Gibson et al., 2009).

A Gibson assembly kit from NEB was used for this purpose. A 20 µL reaction mixture

consisting of 0.02–0.5 pmols DNA (2 to 5-fold molar excess of insert than vector was

used), 10 μl of Gibson Assembly Master Mix (2X) and the required amount of deionized

H2O was incubated at 50 °C for 1 h. 2-4 µL of assembled plasmid was used to transfer

competent E . coli JM109 (Promega) cells by heat shock according to the manufacturer’s

instructions.

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4.2.1.5 Golden Gate cloning

For cloning the tmrA_full gene (tmrA with TAT signal) into the pASK-IBA63c-plus vector

and cloning putative trigger factor genes tf104 and tf161 into both pASK-IBA63c-plus

and pBBR1MCS-2 vectors, Golden Gate cloning was performed (Engler et al., 2009,

Engler et al., 2008). The type IIS restriction enzyme BsaI and T4 DNA ligase from NEB was

used for these reactions. A 20 μL reaction volume consisted of 2 μL 10X CutSmart buffer,

2 μL 10 mM ATP, 0.5 μL BsaI enzyme, 0.25 μL T4 DNA ligase, 5:1 ratio of insert to plasmid

and deionized H2O. The reaction mixture was incubated at 37 °C for 20 min. After

incubation 2-4 µL of the reaction mix was used to transfer competent E . coli JM109 cells

by heat shock.

4.2.1.6 Colony PCR

A suitable number of colonies were selected to screen for successful transformation of

recombinant plasmids. Each colony to be screened was boiled in 15 μL of lysis buffer (TE

buffer + 0.1% Triton-X100) for 5 min, then cooled on ice for another 5 min. The lysates

were then centrifuged at 13,000 g for 10 min and 2-4 μL of the supernatant was used as

template for colony PCR. The PCR protocol was the same as described in section 4.2.1.1.

Primers no. 3,4, 19-28 in Table 4.1 were used in the respective PCR reactions.

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Table 4.1 List of primers used in plasmid construction for soluble expression of TmrA with and without chaperone proteins in S. blattae and subsequent transcript analysis studies.

No. Primer name Sequence (5’ → 3’) Reference

1 pASK_fwd AATGAAAAGGCTAAAGCCGATTGGTGGAAGTGGAGCCACCCGCAG This study 2 pASK_rev ACCCGCGTAAACTTGTTTAGGATCAATCATGTCGACGGAGCTCGAATTCG This study 3 tmrA_fwd ATGATTGATCCTAAACAAGTTTACG This study 4 tmrA_rev CTTCCACCAATCGGCTTTA This study 5 BsaI_pASK_fwd AGGTCTCAAGCTCGAATTCGGATCCACAG This study 6 BsaI_pASK_rev AGGTCTCTTGGAGCCACCCGCAGTTC This study 7 BsaI_pBBR_fwd AGGTCTCATCGATATCAAGCTTATCGATACC This study 8 BsaI_pBBR_rev AGGTCTCTAGCTCCAATTCGCCCTATAG This study 9 BsaI_tmrA_full_fwd AGGTCTCAAGCTATGGACAAGGAGAAGTCG This study

10 BsaI_tmrA_full_rev AGGTCTCTTCCATTTCCACCAGTCCGCC This study 11 BsaI_pASK_TF104_fwd AGGTCTCAAGCTATGAAGCAATTCGAGCTTG This study 12 BsaI_pASK_TF104_rev AGGTCTCTTCCAACTAGCTAAATTAATTTCTAAAAGATAATCC This study 13 BsaI_pASK_TF161_fwd AGGTCTCAAGCTATGTCGGTCAAGATCGAG This study 14 BsaI_pASK_TF161_rev AGGTCTCTTCCATTCGTTCTCTTCCGCCAC This study 15 BsaI_pBBR_TF104_fwd AGGTCTCAAGCTATGAAGCAATTCGAGCTTG This study 16 BsaI_pBBR_TF104_rev AGGTCTCTTCGAACTAGCTAAATTAATTTCTAAAAGATAATCC This study 17 BsaI_pBBR_TF161_fwd AGGTCTCAAGCTATGTCGGTCAAGATCGAG This study 18 BsaI_pBBR_TF161_rev AGGTCTCTTCGATTCGTTCTCTTCCGCCAC This study 19 TmrA_full_fwd ATGGACAAGGAGAAGTCG This study 20 TmrA_full_rev TTTCCACCAGTCCGCC This study 21 TF_104_fwd ATGAAGCAATTCGAGCTTGG This study 22 TF_104_rev ACTAGCTAAATTAATTTCTAAAAGATAATCC This study

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23 TF_161_fwd ATGTCGGTCAAGATCGAG This study 24 TF_161_rev TTCGTTCTCTTCCGCCAC This study 25 pASK_seq_fwd GAGTTATTTTACCACTCCCT This study 26 pASK_seq_rev CGCAGTAGCGGTAAACG This study 27 pBBR_seq_fwd ACGTTGTAAAACGACGGCCAG This study 28 pBBR_seq_rev ACTAAAGGGAACAAAAGCTGG This study 29 515F-Y GTGYCAGCMGCCGCGGTAA Parada et al.

(2016) 30 806R GGACTACNVGGGTWTCTAAT Apprill et al.

(2015)

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4.2.1.7 Sequencing

Plasmid DNA was isolated from overnight cultures of E. coli using a QIAprep Spin

Miniprep Kit (Qiagen) using the manufacturer’s instructions. The quality (A260/A280 and

A260/A230) and quantity (ng/ μL) of the extracted DNA were measured using a NanoDrop

One and a Qubit™ 4 Fluorometer from Thermo Fisher Scientific respectively. Samples

were prepared according to the guidelines described in

http://www.ramaciotti.unsw.edu.au and sent to the Ramaciotti Center for Genomics at

UNSW for Sanger sequencing.

4.2.2 Preparing competent S. blattae cells and transformation

4.2.2.1 Chemically competent cells

An overnight culture of S. blattae in LB media was sub-cultured (1:100) in LB media. After

they reached an OD600 of 0.5 - 0.7, cells were collected in sterile 50 mL tubes by

centrifugation at 8,000 g, 4 °C for 10 min. Cells were washed once with ice-cold 30 mM

CaCl2 and then resuspended in the same. Competent cells were then transformed with

1-4 µL of assembly product by heat shock and were plated on LB agar supplemented

with suitable antibiotics. Aliquots of 50 μL were stored at -80 °C for future use.

4.2.2.2 Electrocompetent cells

An overnight culture of S. blattae was sub-cultured in SOB media. When the OD600

reached a value of 0.5 - 0.7 the culture was placed on ice for 15 min. From this point

forward the cells were kept ice cold. Cells were harvested by centrifugation at 5,000 g

for 10 min at 4 °C. Cells were then washed two times with sterile ice-cold 10% glycerol

solution. Cells were resuspended in a suitable volume of sterile 10% glycerol solution

and stored at -80 °C in aliquots of 100 μL.

For electroporation 2-4 μL of plasmid was mixed with 100 μL of electrocompetent cells

and transferred into an ice-cooled 2 mm electroporation cuvette. The cells were

http://www.ramaciotti.unsw.edu.au/

136

electroporated at 2.5 kV, 200 ohms and 25 µF. Cells were then immediately added to

900 μL of pre-warmed SOC media and incubated at 37 °C for an hour with shaking. Cells

were then plated out on LB agar containing suitable antibiotics.

The following S. blattae transformants were produced and and stored in 30% glycerol at

-80 °C for future use.

Table 4.2 List of S. blattae transformants created and used in recombinant protein expression experiments

Competent cell

Method of transformation

Plasmids Name designated for this study

S. blattae Heat shock pASK_tmrA SB_1

Electroporation pBBR_tmrA SB_2

Heat shock pASK_tmrA_full SB_3

Heat shock pASK_tmrA_full and pASK_TF104

SB_4

Heat shock pASK_tmrA_full and pASK_TF161

SB_5

Electroporation pASK_tmrA_full and pBBR_TF104

SB_6

Electroporation pASK_tmrA_full and pBBR_TF161

SB_7

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4.2.3 Heterologous expression of CF reductive dehalogenase in S. blattae

4.2.3.1 Anhydrotetracycline-inducible expression

For the heterologous production of the TmrA, S. blattae strains were cultivated

anaerobically in a defined mineral medium described by Kunze et al., (2017). The recipe

of the media is described in the Appendices (A.3). For each transformant, LB-grown pre-

culture cultivated aerobically overnight at 28 °C was used for inoculation of a second 20

mL LB pre-culture which was grown aerobically at 28 °C for 7–8 h. This second culture

was used as an inoculum (initial OD600 of 0.1) for a third pre-culture grown anaerobically

at 24 °C in the defined mineral medium. When the OD600 of this pre-culture reached a

value of 1.2–2.1, it was used to inoculate the main anaerobic culture of S. blattae

transformant at an initial OD600 of 0.02. The main culture was supplemented with 1 μM

vitamin B12. The cells were cultured at 18 °C up to an OD600 of 0.2–0.4 and were then

induced by the addition of 200 ng/mL anhydrotetracycline. S. blattae strains were

harvested after 6 h by centrifugation at 10,000 g at 4 °C for 10 min under aerobic

conditions.

4.2.3.2 IPTG-inducible expression

A single colony of transformant S. blattae was incubated overnight in 5 mL LB with 50

µg/mL kanamycin. The next day the overnight culture was diluted 1:50 in fresh LB media.

The culture was maintained at 28 °C and agitated at 180 rpm until it reached an OD600

of 0.4. The culture was then inducted with 1 mM IPTG for 3 hours at the same

temperature and agitation speed.

For anaerobic expression, an overnight culture of S. blattae was used to inoculate anoxic

defined media (recipe in the Appendices, A.3) at OD600 of 0.1. The culture was

maintained at 30° C without shaking and induced with 1 mM IPTG when the OD600

reached 0.6. The cells were harvested under aerobic conditions after 6 h of induction.

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4.2.3.3 Co-expression with chaperone proteins

The culture conditions for co-expression of TmrA (including TAT sequence) and

chaperone protein TF104/TF161 was the same as described in section 4.2.3.2. For

induction of the transformants SB_7 and SB_8 200 ng/mL anhydrotetracycline and 1

mM IPTG was added and induction was done for 6 h or 18 h then harvested.

4.2.4 Cell lysis

Cell lysis was performed under anaerobic conditions. Cell pellets were resuspended in

anoxic lysis buffer (50 mM Tris–HCl pH 8, 17 mM NaCl, 250 μg/mL lysozyme, amended

with protease cocktail inhibitor tablet) at a ratio of 2 mL per 1 g wet cells. Cell suspension

was added to 2 ml microcentrifuge tubes containing 0.5 mm ZR BashingBead™ lysis

matrix. Cells were lysed using a TissueLyser LT from Qiagen at 30 Hz for 10 min. The

supernatant obtained after centrifugation at 14,000 g for 15 min was stored at -20 °C

and used in future assays.


The activity assay was performed as described in section 2.2.5. The initial concentration

of chloroform was 0.1 mM and the samples were incubated for 10 h. TmrA

dechlorination activity of CF to DCM by the soluble protein fraction extracted in the

previous section was measured by GC/MS using titanium (III) citrate and methyl viologen

as electron donors.

4.2.6 SDS-PAGE

Pre- and post-induction samples were collected during expression experiments and

were lysed with BugBuster® Plus Lysonase™ Kit (Merck Millipore) according to the

manufacturer’s instructions. The soluble and insoluble portions were mixed with 4x

NuPAGE® gel loading buffer (Thermo Fisher, USA) to reach a concentration of 2x and

denatured at 95°C for 5 min. 20 μL of each sample and 10 μL of SeeBlue Pre-stained

Protein Standard (Life Technologies, USA) were loaded on Bis-Tris gradient gels (4-15%)

139

(Life Technologies, USA) in NuPAGE® MES running buffer (Invitrogen, USA) and

separated at 150 V for 40 min. After washing and staining with GelCode™ Blue Stain

Reagent (Thermo Fisher, USA) gels were imaged using a Gel Doc™ XR+ Imager (BIORAD).

The relevant proteins were identified by LC-MS/MS performed at the Biomedical Mass

Spectrometry Facility, UNSW.

4.2.7 Transcript analysis

Bacterial cell pellets were analysed for the presence of transcripts corresponding to

tmrA, tmrA_full, tf104 and tf161 genes. Cells were lysed for RNA extraction by adding 1

mL of TRI Reagent® (Sigma-Aldrich, St. Louis, MO, USA) to approx. 107 bacterial cells and

pipetting. Lysates were stored at -80°C until needed.

Cell lysates were thawed, and RNA was extracted using Direct-zol™ RNA MiniPrep Kit

(Zymo Research, Irvine, CA, USA) following the manufacturers’ instruction. The RNA

extracts were checked for DNA contamination by 16s rRNA gene PCR amplification using

515F–806R primer pair. 25 µL reactions contained 12.5 µL Q5 High-Fidelity 2X Master

Mix (New England Biolabs, Ipswich, MA, USA), 0.5 µM of respective primers, ~100 ng of

template DNA and required amount of nuclease-free water to make up the reaction

volume. The PCR reactions were performed in a thermocycler using the following cycling

conditions: Initial denaturation at 98 °C for 30 s, 35 cycles of 10 s denaturation at 98 °C,

30 s annealing at 60 °C (the annealing temperature was determined using the NEB Tm

calculator, https://tmcalculator.neb.com/) and 30 s per kb extension at 72 °C, followed

by a final extension of 3 min at 72 °C. PCR products were analysed using E-Gel Precast

Agarose Electrophoresis System (Thermo Fisher Scientific). If the gels showed presence

of DNA contamination in the RNA extracts, the residual DNA was removed using

Ambion® Turbo DNA-free™ Kit (Life Technologies, Carlsbad, CA, USA) as described by

the manufacturer.

DNA-free RNA extracts were used for cDNA synthesis using ProtoScript® II First Strand

cDNA Synthesis Kit (New England Biolabs, Ipswich, MA, USA). Reactions were performed

following the standard protocol supplied by the manufacturer using the Random Primer

Mix. Successful cDNA synthesis was verified via 16S rRNA gene PCR amplification as

140

described above. The presence of transcripts of interest was detected by amplifying the

genes of interest using specific primer pairs (Table 4.1). The PCR reactions and product

analysis were performed as described above.

141

4.3 RESULTS

Several different approaches were tested and compared for heterologous expression of

functional TmrA in cobalamin-synthesizing S. blattae. Anhydrotetracycline-inducible and

IPTG-inducible expression systems were used. Initially tmrA gene devoid of a TAT signal

and predicted membrane spanning regions was expressed. For co-expression with

chaperone proteins tmrA gene including the TAT signal (tmrA_full) was synthesized.

4.3.1 Anhydrotetracycline-inducible expression of TmrA in S. blattae

4.3.1.1 Expression of TmrA without the TAT-signal

The tmrA gene cloned into pD864 vector obtained from Jugder et al., (2018) was excised

by PCR and used as the insert (Figure 4.2). pASK-IBA63c-plus vector was linearized by

PCR using a primer pair that contained the relevant overhangs (Figure 4.2) to facilitate

Gibson assembly as described by Gibson et al., (2009). pASK-IBA63c-plus vector includes

a strep-tag sequence, the tmrA gene was cloned into the vector so that the strep-tag

was fused to the C-terminus. The strep-tag could facilitate in immunological detection

using Strep-tag-specific antibody and purification with strep-tactin columns. The correct

assembly after cloning was confirmed by colony PCR (Figure 4.2) and also Sanger

sequencing. Purified vector was then used to transform competent S. blattae cells.

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Figure 4.2. PCR products in Gibson assembly of tmrA and pASK-IBA63c-plus. Lane 1. PCR product of pD864 vector, tmrA (1,233 bp); Lane 2. Linearized pASK-IBA63c-plus (3,260 bp); Lane 3. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lanes 4-8. tmrA gene after colony PCR of 5 different colonies. The framed sections show the desired bands.

TmrA was expressed in S. blattae cultured anaerobically in a defined mineral media

following a protocol described by Kunze et al., (2017). Cyanocobalamin was added to

the growth medium to support the cobamide supply. Cells were cultured for 6 h for after

induction of gene expression with anhydrotetracycline. Cells were lysed into soluble and

insoluble fractions using BugBuster® Plus Lysonase™ Kit and analysed by SDS-PAGE.

Bands of the expected size (45 kDa) were visible in both soluble and insoluble fractions

of the gel (Figure 4.3). Expression of TmrA was further confirmed by tryptic digest and

liquid chromatography tandem mass spectrometry (LC-MS/MS) at the Biomedical Mass

Spectrometry Facility, UNSW (methods and results shown in Appendices, A.6).

143

Figure 4.3. SDS-PAGE analysis of insoluble and soluble fractions of cell lysate of S. blattae expressing TmrA. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Insoluble fraction; Lane 3. Soluble fraction. The framed portion contains the bands expressing TmrA.

4.3.1.2 Expression of TmrA with the TAT-signal

pASK-IBA63c-plus vector was linearized by PCR using a primer pair that contained BsaI

recognition sites. Codon optimized tmrA_full gBlock purchased from Integrated DNA

Technologies was amplified using a primer pair also containing BsaI recognition sites.

The size of both the vector and insert was checked by gel electrophoresis (Figure 4.4).

tmrA_full was cloned into the pASK-IBA63c-plus vector with strep-tag sequence at the

C-terminus using a Golden Gate cloning method (Engler et al., 2009, Engler et al., 2008).

The correct gene assembly was confirmed by colony PCR (Figure 4.4) followed by Sanger

sequencing. Competent S. blattae cells were then transformed with purified vector.

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Figure 4.4. PCR products for Golden Gate cloning and subsequent colony PCR. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Linearized pASK-IBA63c-plus; Lane 3. tmrA_full; Lanes 4-13. Colony PCR products from colonies 1-10 (Colony 6 in Lane 9 shows the correct band, albeit very faint); Lane 14. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 15. tmrA_full (Plasmid was harvested by a mini-prep protocol from the colony with the correct gene insert. PCR using the plasmid as template showed strong band at correct position). Framed sections show the desired bands.

TmrA including the TAT sequence was expressed in S. blattae cultured anaerobically

following the same protocol described in section 4.3.1.1. Cell lysates were analysed on

SDS-PAGE gel (Figure 4.5.). However, distinct bands of the expected size (~50 kDa) were

not distinguishable. Expression of TmrA was confirmed by dehalogenation of CF to DCM

described further in section 4.3.5.

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Figure 4.5. SDS-PAGE analysis of soluble fraction of cell lysates of S. blattae expressing TmrA. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Native S. blattae, pre-induction; Lane 3. Native S. blattae, post-induction; Lane 4. TmrA without TAT sequence, pre-induction; Lane 5. TmrA without TAT sequence, post-induction; Lane 6. TmrA with TAT sequence, pre-induction; Lane 7. TmrA with TAT sequence, post-induction; Lane 8. Co-expression of TmrA and TF104, pre-induction; Lane 9. Co-expression of TmrA and TF104, post-induction; Lane 10. Co-expression of TmrA and TF161, pre-induction; Lane 11. Co-expression of TmrA and TF161, post-induction; Lane 12. SeeBlue Pre-Stained Protein Standard. The framed sctions represent the area of interest; the yellow frame for TmrA without TAT sequence (45 kDa), the orange frame for TmrA with the TAT sequence and TF161 (both ~50 kDa) and the blue frame for TF104 (~37 kDa).

4.3.2 IPTG-inducible expression of TmrA in S. blattae

For IPTG-inducible expression of TmrA, the tmrA gene was cloned into pBBR1MCS-2

vector by Gibson assembly. tmrA in pD864 vector from Jugder et al., (2018) was

amplified by PCR to use as the insert gene. pBBR1MCS-2 was linearized by PCR with a

primer pair containing suitable overhangs. The fidelity of the plasmid construction was

tested by colony PCR (Figure 4.6) and confirmed by Sanger sequencing.

Electrocompetent S. blattae cells were then transformed with purified vector.

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Figure 4.6. PCR products used in Gibson assembly and tmrA from colony PCR. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Linear pBBR1MCS-2; Lane 3. tmrA; Lane 4. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lanes 5-7. Colony PCR products from 5 different colonies. Framed sections show the desired bands.

S. blattae was cultured aerobically in LB media supplemented with vitamin B12.

Expression of TmrA was induced by 1 mM IPTG for 3 h. Expression of TmrA under

anaerobic conditions were carried out in the defined mineral media with vitamin B12,

with the induction phase being 6 h in duration. Cells were lysed into soluble and

insoluble fractions and analysed on SDS-PAGE gel (Figure 4.7). Faint bands of the

expected size (45 kDa) were visible in both soluble and insoluble fractions. However,

liquid chromatography tandem mass spectrometry (LC-MS/MS) confirmed that TmrA

was only expressed as inclusion bodies (Results shown in Appendices, A.6). This was

further demonstrated by an activity assay where no conversion of CF to DCM was

observed (data not shown).

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Figure 4.7. SDS-PAGE of cell lysate of S. blattae. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Soluble fraction of pre-induction sample; Lane 3. Insoluble fraction of pre-induction sample; Lane 4. Soluble fraction of post-induction sample; Lane 5. Insoluble fraction of post-induction sample. Framed segment shows the location of the expected band.

4.3.3 Co-expression of TmrA with chaperone proteins

The tmrAB operon of strain Dehalobacter sp. UNSWDHB does not contain any genes

encoding chaperone proteins. As mentioned in section 4.1, for this experiment we chose

two trigger factors found in different Dehalobacter sp.: TF104 (WP_015043727) and

TF161 (WP_015044672). The genes were cloned into separate vectors by Golden Gate

assembly, both pASK-IBA63c-plus and pBBR1MCS-2 vectors were used. The success of

the assembly was tested by colony PCR (Figure 4.8 and 4.9) and confirmed by Sanger

sequencing. Competent S. blattae cells were transformed with two vectors

simultaneously: 1. pASK_tmrA_full & pASK_TF104, 2. pASK_tmrA_full & pASK_TF161, 3.

pASK_tmrA_full & pBBR_TF104, 4. pASK_tmrA_full & pBBR_TF1616. Double

transformation with vectors having different antibiotic resistent cassettes was

confirmed by growth on agar plates supplemented with both ampicillin and kanamycin.

The success of the transformation with pASK_tmrA_full & pASK_TF104, pASK_tmrA_full

& pASK_TF161 was confirmed by colony PCR using primer pairs targeting the gene

inserts (Primers 19-24 in Table 4.1.).

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For expression of TmrA (with the TAT-signal) together with a chaperone protein

anaerobic culture in defined mineral media was carried out. All cultures were

supplemented with vitamin B12 to ensure there was enough corrinoid cofactor to

facilitate TmrA expression. Inductions were done for 6 h and 16h. The cultures where

the chaperones were cloned into pBBR1MCS-2 vector showed no soluble expression of

TmrA or the chaperones TF104 and TF161 (Figure 4.10). This was also confirmed by the

absence of any “hits” using LC-MS/MS on extracted gel bands and the absence of

dehalogenating activity in the lysates. Expression tests with the transformants where

TmrA and chaperone proteins were both cloned into pASK-IBA63c-plus vector showed

no distinct bands for soluble expression of TmrA, TF104 and TF161 (Figure 4.5) but

demonstrated dehalogenation of CF to DCM in the activity assay.

Figure 4.8. PCR products in Golden Gate assembly of tf104 into vectors pASK-IBA63c-plus and pBBR1MCS-2. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Linearized pASK-IBA63c-plus; Lane 3. Linearized pBBR1MCS-2; Lanes 4. tf104 insert for pASK-IBA63c-plus; 5. tf104 insert for pBBR1MCS-2; Lane 6. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lanes 7-9. Colony PCR products of cells transformed with pBBR_TF104, Lane 10-11. Colony PCR products of cells transformed with pASK_TF104; Lane 12. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder. Framed sections show the genes of interest.

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Figure 4.9. PCR products in Golden Gate assembly of tf161 into vectors pASK-IBA63c-plus and pBBR1MCS-2. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Linearized pASK-IBA63c-plus; Lane 3. Linearized pBBR1MCS-2; Lanes 4. tf161 insert for pASK-IBA63c-plus; 5. tf161 insert for pBBR1MCS-2; Lane 6. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lanes 7-12. Colony PCR products of cells transformed with pASK_TF161; Lane 13. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lanes 13-17. Colony PCR products of cells transformed with pBBR_TF161. Framed sections show the genes of interest.

Figure 4.10. SDS-PAGE analysis of S. blattae cells containing TmrA and chaperone proteins cloned into pBBR1MCS-2. Transformants with TmrA and pBBR-TF104: Lane 1. Pre-induction soluble fraction; Lane 2. Pre-induction insoluble fraction; Lane 3. Post-induction soluble fraction; Lane 4. Post-induction insoluble fraction; Lane 5. SeeBlue Pre-Stained Protein Standard. Transformants with TmrA and pBBR-TF161: Lane 1. Pre-induction soluble fraction; Lane 2. Pre-induction insoluble fraction; Lane 3. Post-induction soluble fraction; Lane 4. Post-induction insoluble fraction. The framed areas show the region of protein of interest.

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In preliminary expression tests, the co-expression of TmrA including TAT sequence

(TmrA_full) and chaperone proteins TF104 and TF161 was not showing any

distinguishable bands on the SDS-PAGE gel. LC-MS/MS analysis on bands cut out from

gels also could not detect the presence of the desired proteins. It was decided to check

the transcription of the genes tmrA_full, tf104 and tf161. The presence of mRNA for

these genes was analysed (Figures 4.11, 4.12, 4.13, 4.14 and 4.15). Transformed S.

blattae was cultured in defined mineral media, samples were from induced and non-

induced cultures. Results show that tmrA_full was transcribed when expressed alone or

together with the chaperone proteins, in both cases where tf104 and tf161 was cloned

into pASK-IBA63c-plus and pBBR1MCS-2 vectors. Bands were observed in both pre- and

post-induction samples (Figures 4.11, 4.12, 4.13, 4.14 and 4.15). For samples with tf104

cloned into pBBR1MCS-2, very faint bands of tf104 were detected when expressed alone

and with tmrA_full (Figure 4.12). Prominent bands were visible for tf161 cloned into

pBBR1MCS-2 (Figure 4.13). In case of tf104 and tf161 cloned into pASK-IBA63c-plus,

bands were observed both when expressed alone or consequently with tmrA_full

(Figures 4.14 and 4.15).

Figure 4.11. Transcript analysis of tmrA_full gene expressed in S. blattae. Lane 1. Positive control (tmrA_full gBlock purchased from IDT); lane 2. Induced sample; lane 3. Non-induced sample; Lane 4. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder.

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Figure 4.12. Transcript analysis of tf104 and with tmrA_full genes cloned in pBBR1MCS-2 vector and expressed in S. blattae. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Positive control for tf104 (tf104 gBlock purchased from IDT); Lane 3. Induced tf104 expressed alone; Lane 4. Non-induced tf104 expressed alone; Lane 5. Induced tf104 expressed with tmrA_full; Lane 6. Non-induced tf104 expressed with tmrA_full; Lane 7. Negative control for tf104; Lane 8. Positive control for tmrA_full (tmrA_full gBlock purchased from IDT); Lane 9. Induced tmrA_full; lane 10. Non-induced tmrA_full.

Figure 4.13. Transcript analysis of tf161 and with tmrA_full genes cloned in pBBR1MCS-2 vector and expressed in S. blattae. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Positive control for tf161(tf161 gBlock purchased from IDT); Lane 3. Induced tf161 expressed alone; Lane 4. Non-induced tf161 expressed alone; Lane 5. Induced tf161 expressed with tmrA_full; Lane 6. Non-induced tf161 expressed with tmrA_full; Lane 7. Positive control for tmrA_full; Lane 8. Induced tmrA_full; Lane 9. Non-induced tmrA_full.

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Figure 4.14. Transcript analysis of tf104 and with tmrA_full genes cloned in pASK-IBA63c-plus vector and expressed in S. blattae. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Induced tmrA_full; Lane 3. Non-induced tmrA_full; Lane 4. Induced tf104 expressed with tmrA_full; Lane 5. Non-induced tf104 expressed with tmrA_full; Lane 6. Induced tf104 expressed alone; Lane 7. Non-induced tf104 expressed alone.

Figure 4.15. Transcript analysis of tf161 and with tmrA_full genes cloned in pASK-IBA63c-plus vector and expressed in S. blattae. Lane 1. InvitrogenTM E-GelTM 1 Kb Plus DNA Ladder; Lane 2. Positive control for tmrA_full (tmrA_full gBlock purchased from IDT); Lane 3. Induced tmrA_full; Lane 4. Non-induced tmrA_full; Lane 5. Induced tf161 expressed with tmrA_full; Lane 6. Non-induced tf161 expressed with tmrA_full; Lane 7. Positive control for tf161 (tf161 gBlock purchased from IDT); Lane 8. Induced tf161 expressed alone; Lane 9. Non-induced tf161 expressed alone.

153

4.3.5 Dechlorination activity of TmrA expressed in S. blattae

The functionality of all variants of S. blattae were tested by measuring chloroform (CF)

dechlorination in crude extracts with reduced methylviologen as the electron donor. 100

µM CF was added to each reaction and incubated for 10 h. Strains SB_1 (containing

pASK_tmrA), SB_3 (containing pASK_tmrA_full), SB_4 (containing pASK_tmrA_full and

pASK_TF104) and SB_5 (containing pASK_tmrA_full and pASK_TF161) demonstrated

conversion of CF when TmrA expression was induced by the addition of

anhydrotetracycline. SB_1 exhibited the highest rate of conversion of CF to DCM (Figure

4.16). No CF-dechlorinating enzyme activity was detectable in extracts of SB_2

(containing pBBR_tmrA), SB_6 (containing pBBR_tmrA and pBBR_TF104) and SB_7

(containing pBBR_tmrA and pBBR_TF161). Native S. blattae and denatured crude extract

also showed no evidence of CF dechlorination.

Figure 4.16. Dechlorination of CF to DCM by S. blattae strains extracts (n=2). The range bars show the lowest and highest specific activity for each strain.

154

4.4 DISCUSSION

S. blattae was chosen as an expression host for TmrA because of its capability to

synthesize vitamin B12 de novo and its compatibility with the molecular tools developed

for E. coli. The corrinoid cofactor is an intricate part of RDase structure and is buried

inside a substrate binding pocket of the active site of properly folded proteins (Bommer

et al., 2014, Payne et al., 2015). The lack of proper corrinoid cofactor and their improper

incorporation has been attributed as the reason behind the failure of numerous

heterologous expression studies in E. coli. S. blattae has proven to be one solution to

this issue; so far three RDases have been reported to be produced heterologously in a

catalytically active state in S. blattae: the tetrachloroethene RDase of D. hafniense strain

Y51 (PceAY51), the 3,5-dichlorophenol RDase of D. hafniense strain DCB-2 (RdhA3) and

the 1,2-dichloroethane RDase of Desulfitobacterium dichloroeliminans (DcaA) (Mac

Nelly et al., 2014; Kunze et al., 2017).

For soluble and functional expression of TmrA, S. blattae was cultured anaerobically in

a defined mineral media described by Kunze et al., (2017). Glycerol was used as the sole

energy and carbon source. Approximately 1% of the genome of a typical B12-producing

enteric bacterium is involved in B12 biosynthesis and transportation (Roth et al., 1996).

de novo corrinoid synthesis involves more than 30 enzymatic steps (Moore et al., 2013).

Regardless of this large genetic expense only five B12-dependent enzymes are found in

one or more species of enteric bacteria. In S. blattae DSM 4481 metH is the only gene

encoding a functional B12-dependent enzyme, methionine synthase. Unlike S. blattae

strains ATCC 33429 and ATCC 33430, strain DSM 4481 does not harbour glycerol

dehydratase which is essential for the fermentation of glycerol (Daniel et al., 1998,

Toraya et al., 1980). The insertion of a prophage (MuEb) in the dha regulon encoding

glycerol dehydratase in strain DSM 4481 is the reason for the loss of the functional

enzyme (Andres et al., 2004). Roth et al., (1996) suggested that the MuEb insertion

maybe a step towards the loss of the B12 synthetic ability of strain DSM 4481 in the

future. To ensure the continual production of B12 to support the expression of RDases,

Mac Nelly et al., (2014) used glycerol as the carbon source in the culture media.

155

In this study, TmrA was initially expressed without the TAT signal and the membrane

spanning regions as was the case in the previous two chapters. Next, TmrA was co-

expressed with two different chaperone proteins identified from the Dehalobacter

UNSWDHB genome with an aim to increase the level of soluble expression. The N-

terminal TAT signal serves as the recognition site of the precursor by the TAT export

system in the membrane (Palmer and Berks, 2012). Chaperone proteins bind specifically

to the TAT signal sequence and are believed to delay the translocation of the RdhA,

increasing the chance of correct folding and inclusion of cofactors (Maillard et al., 2011,

Morita et al., 2009). Hence, the tmrA gene including the TAT signal was synthesized to

be used in these co-expression tests. The tmrAB operon however, does not include a

gene encoding a chaperone protein so two chaperone proteins, TF104 and TF161, were

chosen to be used in this study. The tmrA, tmrA_full, tf104 and tf161 genes were

encoded into the vectors pASK-IBA63c-plus and pBBR1MCS-2 and competent S. blattae

cells were transformed with the desired vectors (Figure 4.17).

Figure 4.17. The different plasmid constructs and S. blattae transformants used in the expression tests.

The three RDases previously expressed successfully in S. blattae so far all had a trigger

factor-like chaperone in their operon structure and co-expression with the respective

chaperone protein resulted in increased soluble expression of the RDase (Mac Nelly et

al., 2014; Kunze et al., 2017). D. hafniense Y51 possesses at least three putative trigger

156

factor proteins, the PceT-encoding gene is located in the pce operon and recombinant

expression experiments suggest that PceT is a specific trigger factor for PceA (Morita et

al., 2009). Maillard et al., (2011) hypothesized that by lacking the N-terminal ribosome-

binding domain, RdhT distinguishes from generic trigger factor proteins and establishes

itself as the dedicated chaperone for RdhAs. Sjuts et al., (2012) demonstrated soluble,

albeit non-functional expression of D. restrictus PceA when in E. coli when fused with E.

coli trigger factor. This has been the first attempt to co-express a RDase with a

chaperone that does not belong to rdhAB operon or the host organism. However, co-

expression with the chaperones TF104 and TF161 seemingly had no impact on the level

of soluble expression. This suggests the need for defined chaperone proteins for the

increase of soluble expression of TmrA.

In terms of specific activity, the truncation of the TAT-signal seemed to have a significant

effect (p<0.05) on the dechlorinating activity of the enzyme. In the previous two

chapters TmrA devoid of the TAT signal and predicted membrane spanning regions was

expressed. This strategy was attempted to navigate expression of the originally

periplasmic RDases toward the cytoplasm (Jugder et al., 2018, Parthasarathy et al.,

2015). The studies of Parthasarathy et al., (2015) however did not result in a soluble

cytoplasmic expression of recombinant VcrA. Jugder et al., (2018) on the other hand

expressed soluble TmrA in B. megaterium but no comments were made on the role of

the lack of TAT sequence. In this chapter when the expression of TmrA both with and

without the twin-arginine sequence was carried out, a difference in their dechlorination

activity was observed. While in both cases the TmrA was expressed in a soluble and

active form, the dechlorination rate was much higher without the TAT sequence. It can

be postulated that the lack of the twin-arginine sequence effects the folding of TmrA

and in turn modulates the dechlorination activity. Signal peptides not only play roles in

the membrane translocation of the target proteins but also impact their folding kinetics

and stability (Freudl, 2018, Punginelli et al., 2004). Another possible explanation could

be that when the TAT signal is included it makes the active site of TmrA less accessible

to the substrate, CF.

157

4.5 CONCLUSIONS

This is the first report of expression of a functional RDase encoded in a rdh operon

lacking a specific chaperone protein in S. blattae. While the RDase alone (both with and

without the TAT signal) was expressed in a soluble and functional state, co-expression

with two other trigger factor proteins did not have much effect on the level of soluble

expression or the dechlorination activity. The expression of TmrA without the TAT signal

sequence and the membrane spanning region however resulted in soluble expression

exhibiting a significantly higher dechlorination activity (p < 0.05). This strategy can be

utilized in expressing RDases lacking a definite chaperone protein in their rdh gene

cluster. Further studies need to be carried out on the role of the TAT signal when RDases

are expressed recombinantly. The co-expression of chaperon protein with TmrA

excluding the TAT signal would also aid in the clarification of the role of the TAT signal.

158

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CHAPTER FIVE: Cupriavidus necator as a chloroform reducing organism: a molecular biology approach

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5.1 INTRODUCTION

The isolation of Desulfomonile tiedjei, which demonstrated growth by coupling

oxidation of formate or hydrogen to the reduction of 3-chlorobenzoate, shed light on a

previously unknown respiratory process, organohalide respiration, where reductive

dehalogenation of organohalides is coupled to a chemiosmotic mechanism (Dolfing and

Tiedje, 1987, Louie and Mohn, 1999a, Shelton and Tiedje, 1984). Since then, numerous

bacterial strains capable of OHR have been isolated and studied in detail. Though

considerable progress has been made in understanding the phylogenetic, physiological,

genomic, and biochemical features, the molecular composition of the organohalide

respiratory electron transport chains in OHRB is an area that still requires extensive

study. RDases are the enzymes directly catalyzing dechlorination reactions; the redox

partners and mechanism for proton translocation however is yet to be fully determined

for respiratory RDases. This is not the case for non-respiratory RDases. The NpRdhA

gene in the N. pacificus genome is located in a putative operon consisting of a

ferredoxin-NADPH oxidoreductase and [4Fe−4S] and [2Fe−2S] ferredoxins (Lai et al.,

2012). The BhbA in Comamonas sp. 7D-2 has a ferredoxin reductase-like NAD(H)-binding

domain and an iron-binding domain (Chen et al., 2013). These features in the non-

respiratory RDases link substrate reduction to NADPH oxidation. Collins et al. (2018)

demonstrated in vitro and in vivo reconstitution of a NADPH-dependent reducing system

for NpRdhA expressed in B. megaterium. They showed that a non-physiological system

consisting of E. coli flavodoxin reductase (EcFldr) and spinach ferredoxin (SpFd) was able

to support NADPH-dependent 3,5-dibromo-4-hydroxybenzoic acid reduction by

NpRdhA.

Molecular hydrogen is the sole electron donor in Dehalobacter sp. UNSWDHB, as is the

case for other obligate OHRB. H2 is presumably oxidized by membrane associated uptake

hydrogenases. Five hydrogenases showed increased abundance in the genome of

Dehalobacter sp. UNSWDHB in response to CF; one Hup-type (Hya) Ni-Fe hydrogenase,

two membrane associated energy-conserving Ech and Hyc-type Ni-Fe hydrogenases and

two cytoplasmic Fe-Fe hydrogenases; suggesting their roles in electron transfer (Jugder

et al., 2016c). Heterologous reconstitution of the oxidation-reduction of H2/CF by uptake

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hydrogenase and TmrA would require the recombinant expression of both the enzymes.

However, recombinant expression of functional hydrogenases has proven to be difficult

due to the complexity and enzyme specificity of the post-translational maturation

process (English et al., 2009, Jugder, 2013).

C. necator H16 is a Gram-negative lithoautotrophic β-proteobacterium. It is found in soil

and freshwater habitats and can grow at the interface of anoxic and oxic environments

(Yu and Munasinghe, 2018). The organism can acclimate between heterotrophic and

autotrophic lifestyles utilizing organic compounds and hydrogen respectively as energy

source (Pohlmann et al., 2006). C. necator has three O2-tolerant hydrogenases which are

encoded on the mega-plasmid pHG1; a membrane-bound hydrogenase (MBH), a soluble

hydrogenase (SH) and a regulatory hydrogenase (RH)(Burgdorf et al., 2005, Schwartz et

al., 2003). Growth under autotrophic conditions induces the expression of the uptake

hydrogenases MBH and SH in the environment. Culture in heterotrophic media

promotes MBH and SH expression with a higher specific activity (Jugder et al., 2015,

Lauterbach and Lenz, 2013). Culture in minimal medium FGN (fructose-glycerol-

nitrogen) is characterized by initial growth on the preferred carbon source, fructose,

which represses hydrogenase expression; fructose depletion leads to growth on the less-

preferred substrate glycerol, thus stimulating de-repression of hydrogenase expression

(Friedrich et al., 1981, Jugder et al., 2016b).

In this chapter the feasibility of C. necator H16 as an expression host for expression of

soluble active chloroform reductive dehalogenase (TmrA) is assessed. The potential to

utilize the uptake Ni-Fe hydrogenases of C. necator as redox partners and induce in vivo

and in vitro dehalogenation of CF is also evaluated. An important consideration for

recombinant expression of any RDase is the capacity of the host to synthesize/uptake

vitamin B12. C. necator H16 cannot synthesize B12 de novo but has the ability to assimilate

cobalamin and its precursors (Pohlmann et al., 2006).

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5.2.1 C. necator H16 strain confirmation

C. necator H16 strain DSM 428 was retrieved from -80 °C freezer and was plated on LB

agar and incubated at 30 °C for 48 h. The culture was originally obtained from Deutsche

Sammlung von Mikroorganismen und Zellkulturen GmbH (Germany) in 2013 as a freeze-

dried culture, rehydrated according to the manufacturer’s instructions and glycerol

stocks were made.

5.2.1.1 Genomic DNA extraction of C. necator

A single colony was picked from the LB plate and grown in 5 mL LB broth at 30 °C for 24

h. The cells were harvested by centrifugation and genomic DNA (gDNA) was extracted

using Invitrogen™ PureLink™ Genomic DNA Mini Kit (Thermo Fisher Scientific) as

described by the manufacturer.

5.2.1.2 16s rRNA sequencing

Samples were prepared according to the guidelines described in

http://www.ramaciotti.unsw.edu.au and sent to the Ramaciotti Center for Genomics at

UNSW for 16S rRNA gene sequencing. The 515F–806R primer pair was used for the

purpose of validating the identity of the organism.

5.2.2 Chloroform and dichloromethane tolerance test

Prior to contemplating the ultimate goal of engineering C. necator to perform as a

recombinant chloroform dehalogenating organism, it was necessary to determine the

tolerance of C. necator against different concentrations of chloroform and

dichloromethane. An overnight culture of C. necator was used to inoculate LB media

and was maintained at 30 °C, 200 rpm until it reached the log phase of growth. The

tolerance test was carried out in two separate 24-well deep well Deutz microplates for

CF and DCM. The concentrations of CF and DCM tested were 0, 0.83, 1.67 and 2.5 mM.

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The layout of the Deutz microplates with the different concentrations of CF and DCM is

shown in Figure 5.1. The initial OD600 in each cell was 0.16 and the plates were

maintained at 30 °C, 200 rpm throughout the culture period. The OD600 of cells from

each level of CF and DCM concentration was measured in duplicate every 90 min over

7.5 h. At the end, cultures from the cells with the highest concentration of CF and DCM

was diluted in sterile 1x PBS by 10-5 and 100 μL was plated out in triplicate on LB agar to

determine viable cell counts. The plates were incubated at 30 °C for 48 h and then

colonies were counted.

1 2 3 4 5 6

A 0 mM 0 mM 0 mM 0 mM 0 mM 0 mM

B 0.83 mM 0.83 mM 0.83 mM 0.83 mM 0.83 mM 0.83 mM

C 1.67 mM 1.67 mM 1.67 mM 1.67 mM 1.67 mM 1.67 mM

D 2.5 mM 2.5 mM 2.5 mM 2.5 mM 2.5 mM 2.5 mM

Figure 5.1. Layout of the Deutz microplates with different concentrations of CF and DCM for tolerance test.

5.2.3 Vitamin B12 uptake assay

A microbiological assay of vitamin B12 according to the procedures of Vitamin B12 activity

assay described in the United States Pharmacopeial Convention (2008) was carried out

to confirm the capability of C. necator to uptake vitamin B12 from the growth media.

Lactobacillus delbrueckii subsp. lactis ATCC™ 7830 (Lactobacillus leichmannii) was used

as the test organism in this procedure.

An overnight culture of C. necator was refreshed in LB media and grown with and

without vitamin B12 (1 µM) for 24 h at 30 °C, 200 rpm. Cells were then harvested by

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centrifugation at 10,000 g, 4 °C for 10 min, washed three times in sterile 1 x PBS and

resuspended in 0.5 mL of the same buffer. The cells were lysed by a freeze-thaw method

using liquid nitrogen and a water bath maintained at 37 °C. The lysates were centrifuged

at 4 °C, 10,000 g for 10 min and the supernatant was collected to use in the vitamin B12

assay. Sterile conditions were maintained at every step to eliminate the possibility of

contamination.

Lactobacillus leichmannii was cultured overnight at 37 °C, 200 rpm in MRS broth (De

Man, Rogosa, Sharpe) from Thermo Scientific. The cells were harvested by

centrifugation at 4 °C, 10,000 g for 10 min, washed three times in sterile saline and

resuspended in 10 mL of the same buffer.

Difco™ B12 Assay Medium from BD diagnostics was prepared according to the

manufacturer’s instruction. The media was aliquoted to sterile glass tubes, 5 mL each.

Standards and samples were prepared by adding required amounts of vitamin B12 (final

concentrations of 0, 10, 20, 40 and 50 pg/mL for standards) and previously prepared cell

lysates of C. necator grown with and without vitamin B12 to the B12 Assay Medium. Each

standard and sample were done in duplicate. 50 µL of Lactobacillus leichmannii cell

suspension was added to all the standards and samples and cultured at 37 °C, 100 rpm

for 24 h. Cultures were visually observed for turbidity to assess cell growth.

5.2.4 Expression of native Ni-Fe hydrogenase in C. necator

To induce the expression of the native soluble Ni-Fe hydrogenase, C. necator was grown

heterotrophically in minimal medium FGN (Fructose-Glycerol-Nitrogen) which facilitates

substrate shift from fructose to glycerol and thereby resulting in the de-repression of

hox regulon containing the hydrogenase structural and auxiliary genes. The recipe of the

media is illustrated in the Appendices (A.3). Overnight cultures of C. necator were used

to inoculate fresh FGN media at an OD600 of 0.1. The culture was maintained at 30 ᵒC,

250 rpm for 48 h. The culture was sampled periodically to check the OD600. Cells were

harvested by centrifugation at 10,000 g for 15 min at 4 °C and washed with 50 mM KPi

buffer (pH 7.0). The cell pellets were stored at -80 °C for future assays. A control culture

was carried out under the same conditions in FN (Fructose-Nitrogen) media (A.3).

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5.2.5 Cell lysis

The cell pellets were thawed and resuspended in 50 mM KPi buffer containing EDTA-

free Protease inhibitor and DNAse I (20 µg/ml) at a ratio of cell wet weight and

resuspension buffer of 1:5. The cell suspensions were sonicated using a Branson Digital

Sonifier equipped with 1/8” Tapered Microtip (Branson Ultrasonics Corporation, USA)

at 50% amplitude with 0.5 sec pulse on and 0.5 sec pulse off for 8 minute/g of cell pellet.

The supernatant was separated by centrifugation at 20,000 x g for 50 min at 4 ᵒC.

5.2.6 Hydrogenase activity assay

The protocol described by Jugder et al. (2016b) was followed to confirm the activity of

the soluble hydrogenase in the C. necator lysates. The assay was carried out under

anaerobic conditions. In a septum-sealed special optical glass cuvette, 100 µL of cell

lysate from the previous step was added to 2.9 mL of reaction mixture containing 50

mM H2-saturated KPi buffer (pH 7.0) and 1 mM NAD+ (the electron acceptor). The

formation of NADH was monitored using a Cary-100 UV-Visible Spectrophotometer with

continual incubation at 30 ᵒC for 10 min at 0.5 min intervals at 340 nm.

5.2.7 Hydrogenation-dechlorination activity assay

In vitro dehalogenation of CF utilizing the active soluble hydrogenase expressed in native

C. necator and recombinant active TmrA expressed in B. megaterium was tested. C.

necator and B. megaterium pellets were separately resuspended in lysis buffer at a ratio

of 1:5. C. necator was lysed by sonication (section 5.2.5) and B. megaterium was lysed

chemically as described in section 3.2.6.

The activity assay was set up inside the anaerobic chamber. B. megaterium and C.

necator cell lysates containing active TmrA and Ni-Fe hydrogenase was added at 1:1 ratio

to a 10 mL headspace vial containing H2 saturated KPi buffer and 0.1 mM chloroform,

this was done in duplicate. The vials were incubated at 30 °C for 12 h. After incubation,

the enzymatic reaction was quenched by transferring 1 mL of each reaction mixture

(done in triplicate) to a 10 mL headspace flask containing anhydrous sodium sulphate

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(0.5 g) and 1 M sulphuric acid (1 mL). The headspace of the vials was then analysed by

GC/MS as mentioned in section 2.2.5. The different combinations of the cell lysates used

as sample and control for the activity assay is described in Table 5.1.

Table 5.1 Sample and controls used in the Hydrogenation-dechlorination assay

Sample Control

CN in FGN + induced BM (boiled for 15 min)

CN in FGN +non-induced BM

CN in FGN + non-induced BM (boiled for 15 min)

CN in FGN + induced BM CN in FN + induced BM

CN in FN + induced BM (boiled for 15 min)

CN in FN + non-induced BM

CN in FN + non-induced BM (boiled for 15 min)

CN = C. necator; BM = B. megaterium expressing TmrA

5.2.8 Gene cloning

The pBBR-TmrA plasmid constructed in section 4.2.1.4. was used for IPTG-inducible

expression of TmrA in C. necator.

pKRrha plasmid described in (Sydow et al., 2017) was used for rhamnose-inducible

expression of TmrA. pKRrha plasmid encodes for mobilization sequences (mob,

RSF1010), origin of replication (rep, RSF1010), RP4 partitioning region (par),

transcription activator genes (rhaR, rhaS), tetracycline resistance cassette (Tcr), a

rhamnose-inducible promoter (rhaPBAD) and eGFP. The vector map of the plasmid is in

Appendices.

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5.2.8.1 PCR

The insert tmrA gene (TAT signal removed) was amplified from pD864-TmrA plasmid

described in (Jugder et al., 2018) to include BsaI recognition sites to facilitate assembly.

Linearization of pKRrha plasmid and insertion of BsaI recognition sites were carried out

by a touchdown polymerase chain reaction (TD-PCR). 25 µL reaction mixture contained

12.5 µL Q5 High-Fidelity 2X Master Mix (New England Biolabs, Ipswich, MA, USA), 0.5

µM of forward and reverse primers, 3 ng of template DNA and required amount of

nuclease-free water to make up the reaction volume. The amplification reactions were

carried out in a BIO-RAD C1000 TouchTM Thermal Cycler. The cycling conditions for the

insert gene was as following: Initial denaturation at 98 °C for 30 s, 35 cycles of 10 s

denaturation at 98 °C, annealing at 61 °C for 30 s and 30 s per kb extension at 72 °C,

followed by a final extension of 5 min at 72 °C. The cycling conditions for the TD-PCR is

described in Table 5.2.

The primers used in different PCR reactions used in this study are listed in Table 5.3.

Primers were purchased from Sigma-Aldrich.

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Table 5.2 TD-PCR thermal cycling conditions for linearizing pKRrha plasmid

Cycle Number Denaturation Annealing Extension

Initial cycle 30 sec at 98 ᵒC

1 10 sec at 98 ᵒC 30 sec at 72 ᵒC 6 min at 72 ᵒC













14-35 10 sec at 98 ᵒC 30 sec at 59 ᵒC 6 min at 72 ᵒC

Last cycle 10 min at 72 ᵒC

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Table 5.3 List of primers used in plasmid construction for rhamnose-inducible expression of TmrA in C. necator and transcript analysis studies

No. Primer name Sequence (5’ → 3’) Reference

1 pKRrha_fwd AGGTCTCACCTTCTTAAGAATTGTTCATTACGACC This study

2 pKRrha_rev AGGTCTCTCTTGGCTGTTTTGGCG This study

3 pKRrha_tmrA_fwd AGGTCTCAAAGGATGATTGATCCTAAACAAGTTTACG This study

4 pKRrha_tmrA_rev AGGTCTCTCAAGCTTCCACCAATCGGCTTTAG This study

5 tmrA_fwd ATGATTGATCCTAAACAAGTTTACG This study

6 tmrA_rev CTTCCACCAATCGGCTTTA This study

7 515F-Y GTGYCAGCMGCCGCGGTAA Parada et al. (2016)

8 806R GGACTACNVGGGTWTCTAAT Apprill et al. (2015)

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5.2.8.2 Golden Gate cloning

After PCR product purification using a PCR purification kit (Qiagen) and DpnI digestion

(NEB), TAT signal removed tmrA gene was cloned into pKRrha plasmid using a Golden

Gate cloning protocol described in section 4.2.1.5.

5.2.8.3 Colony PCR

A suitable number of colonies were screened for successful transformation with the

assembled plasmid following the protocol of colony PCR described in section 4.2.1.4.

5.2.8.4 Sequencing

QIAprep Spin Miniprep Kit (Qiagen) was used to extract plasmid DNA from overnight

cultures of E. coli. DNA samples were then prepared according to the guidelines

described in http://www.ramaciotti.unsw.edu.au by adding the desired amount of DNA

and primer to each sample and sent to the Ramaciotti Center for Genomics at UNSW for

Sanger sequencing.

5.2.9 Transformation of C. necator

5.2.9.1 Preparation of competent C. necator cells

A pre-culture of C. necator in LB media was grown till late exponential phase (~18-20

hours). This pre-culture was used to inoculate a 50 mL culture in LB media to an OD600

of approximately 0.05 - 0.1. When the growth reached an OD600 of 0.5-0.7 the culture

was transferred to a sterile 50 ml centrifuge tube and incubated on ice for 15 minutes.

The culture was then centrifuged at 5,000 g, 4 °C for 10 minutes using a pre-cooled rotor.

The cell pellet was washed three times with cold and sterile 10% glycerol. After the final

wash, the pellet was resuspended in 500 μL 10% glycerol and aliquoted in 100 μL

volumes in pre-cooled sterile Eppendorf tubes.

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5.2.9.2 Electroporation

For electroporation, 2 µl DNA (50-100 ng/µL) was added to 100 μL of electrocompetent

cells and transferred to a pre-cooled 2mm electroporation cuvette. The cuvette was

pulsed with 2500 V, 25 μF und 400 Ω and 900 µL LB medium was immediately added

and mix well inside the cuvette. The mixture was transferred to a 1.5 mL Eppendorf tube

and was cultured at 30 °C for 3h (with shaking). The cells were then centrifuged at 14,000

g for 2 min, 700 µL of the supernatant was removed and the pellet was resuspended in

the remaining supernatant. Finally, the cells were plated on LB agar plates containing

200 ng/mL kanamycin and incubated at 30 °C for 36-40 hours.

5.2.9.3 Conjugation

E. coli S17-1(ATCC 47055) cells were made chemically competent following the protocol

described in section 4.2.2.1 and were transformed with 1-4 µL of Golden Gate assembly

product by heat shock and were plated on LB agar supplemented with tetracycline.

Plasmids were then transferred to the recipient C. necator H16 by conjugation. E. coli

S17-1 strain (containing pKRrha_tmrA) and C. necator were cultured until exponential

to early stationary phase. For each strain, 25 mL culture was harvested, and cell pellets

were washed once in the same volume of 0.9% NaCl. Cells were then resuspended

together in 0.2 mL of 0.9% NaCl and dropped in the middle of a LB agar plate. Following

24 h incubation at 30 °C for mating, cells were scrapped off the agar plate with 1 mL

0.9% NaCl and plated on selective M9 agar supplemented with tetracycline and

gentamicin for the selection of trans-conjugants. Single colonies were purified by sub-

cultivation on selective LB agar.

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5.2.10 Recombinant expression of TmrA in C. necator

5.2.10.1 IPTG-inducible expression

Different culture conditions were screened to study the IPTG-inducible expression of

TmrA in C. necator (Table 5.4.) Different culture media, induction OD600, concentration

of IPTG and length of induction in various combinations were examined, with all

combinations carried out in duplicate. The cultures were inoculated at OD600 0.1 and

maintained at 30 °C, 250 rpm. 1 μM B12 and 50 μM ammonium iron (II) sulfate was added

to the culture at the time of induction to facilitate the expression of TmrA.

Table 5.4 Different culture conditions tested for the IPTG-inducible expression of TmrA in C. necator

Culture Media* Induction OD600 IPTG concentration (mM)

Length of induction (h)

LB 0.5 0.1 2

2YT 0.6 0.5 4

TB 0.8 1.0 6

Rich media 24

*LB= Luria broth, 2YT = Yeast extract and tryptone media, TB = Terrific broth

5.2.10.2 Rhamnose-inducible expression

For the rhamnose-inducible expression of TmrA in C. necator the culture conditions

described in Table 5.5. were screened. Pre-cultures of C. necator were used to inoculate

the culture media at an OD600 of 0.1. 1 μM B12 was added to the culture media to

facilitate the expression of soluble TmrA. The cultures were maintained at 30 °C, 180

rpm. C. necator in LB media was cultured for 24 h after induction. C. necator in FGN

media was grown for 4 d. Cells were harvested at the end of the culture by centrifugation

at 10,000 g, 4 °C for 10 min, washed with 1 x PBS and stored at -20 °C for future

processing.

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Table 5.5 Culture conditions for screening of rhamnose-inducible expression of TmrA in C. necator

Media Induction time Rhamnose conc. (mM)

LB Inoculation 0

FGN ~OD600 0.6 0.5

1.0

1.5

2.0

2.5

3.0

5.2.11 Cell lysis

Cell pellets from each culture condition were chemically lysed under anoxic conditions

with BugBuster® Plus Lysonase™ Kit (Merck Millipore) according to the manufacturer’s

instructions.

5.2.12 SDS-PAGE

SDS-PAGE was carried out for the pre- and post-induction samples collected from each

culture condition following the process described in section 4.2.7.


The soluble fractions derived from section 5.2.11. were tested for CF dechlorination

activity of TmrA. Activity assays were performed as mentioned in section 2.2.5. Heat-

inactivated (boiled for 15 min) native C. necator samples were used as negative control.

A second in vitro dechlorination assay was performed to assay the capability of soluble

hydrogenase and TmrA in dechlorinating CF to DCM without adding an external electron

donor to the reaction mix, instead H2-saturated Kpi buffer was used.

176

Cell pellets were lysed according to the protocol described in section 5.2.5. Inside the

anaerobic chamber, to a previously crimped 2 mL GC vial, 0.5 mL of cell lysate and 0.1

mM CF was added to 1.4 mL of H2 saturated KPi buffer (saturated by purging with 100%

H2), this was done in triplicates. The vial was incubated at 30 ᵒC for 12 h, after which the

enzymatic reaction was stopped by transferring 1 mL of the reaction mixture to a 10 mL

headspace vial containing anhydrous sodium sulphate (0.5 g) and 1 M sulphuric acid (1

mL). The headspace of the vial was then analysed by GC/MS as mentioned in section

2.2.5.


C. necator cell pellets from each culture condition in the IPTG-inducible expression

studies were analysed for the presence of transcripts corresponding to tmrA gene. RNA

extraction, cDNA synthesis and transcript analysis were performed as described in

section 4.2.8. The primers used in this study are listed in Table 5.3.

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5.3 RESULTS

In this chapter the suitability of C. necator as a novel expression host for TmrA and its

potential to act as a CF dechlorinating organism was initially tested. Three factors were

considered to determine its suitability:

1. Tolerance to the substrate, CF and the dechlorination product, DCM,

2. Capability to uptake B12 from the culture media

3. Active hydrogenase that can facilitate TmrA in CF dechlorination in the presence of

molecular hydrogen.

The C. necator H16 strain used by (Jugder et al., 2015a) was used in this study. The strain

was confirmed by 16S sequencing done at the Ramaciotti Center for Genomics, UNSW .

5.3.1 C. necator chloroform and dichloromethane tolerance test

The tolerance of C. necator to different concentrations of CF and DCM was tested in

cultures grown in 24-well Deutz microplates. The OD600 of the cultures were sampled

every 90 min, there was no obvious difference in the cells grown with CF and DCM up to

a concentration of 2.5 mM (Figures 5.2 and 5.3). After sampling the OD600 for 7.5 h,

cultures from the cells with 2.5 mM CF and DCM respectively were diluted in sterile 1x

PBS at a dilution factor of 10-5, plated in triplicate on LB agar and incubated at 30 ᵒC for

48 h. The CFU for C. necator grown with CF was greater than 5 x 108 CFU per mL, whilst

for DCM, it was ~ 2.5 x 108 CFU per mL. It was concluded that the organism was suitably

tolerant to both CF and DCM.

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Figure 5.2. OD600 of C. necator grown with various concentrations of chloroform (CF) at different time points.

Figure 5.3. OD600 of C. necator grown with different concentrations of dichloromethane (DCM).

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5.3.2 C. necator vitamin B12 uptake assay

Vitamin B12 activity assay described in the United States Pharmacopeial Convention was

carried out to check the capability of C. necator to uptake vitamin B12 from the growth

media. No quantification of vitamin B12 uptake was done, just the visual difference

between the cells grown with and without vitamin B12 was observed. Lactobacillus

leichmannii grew to higher densities with lysates from C. necator grown with B12 than

without B12. It was concluded that C. necator, could successfully uptake vitamin B12 as

expected from the literature.

Figure 5.4. Vitamin B12 uptake assay. A. Standards containing 0, 10, 20, 30, 40 and 50 pg/mL vitamin B12 from left to right; B. Lactobacillus leichmannii grown with C. necator samples: Tubes 1 and 2 were grown with lysates of C. necator without B12 supplementation, tubes 3 and 4 on the right are grown with lysates of C. necator with B12 supplied in the growth media.

5.3.3 Expression of native Ni-Fe hydrogenase in C. necator

NAD+ as used as the electron acceptor in the H2 oxidation assay. NADH formation was

observed at 340 nm using a Cary 100 UV-Visible Spectrophotometer (Figure 5.5). The

enzyme activity was not calculated, the reduction of NAD+ to NADH was monitored with

continual incubation for 10 min at 340 nm using lysates from cells grown in FGN media

and FN media as control. Cell lysates from FN media culture also demonstrated

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reduction of NAD+ to NADH because of the presence of hydrogenases other than SH, but

to a lesser extent than cell lysates grown on FGN media.

Figure 5.5. Reduction of NAD+ to NADH (observed at 340 nm) coupled with H2 oxidation by soluble hydrogenase from C. necator grown in A. FGN media; B. FN media; and standards C. 1 mM NAD+ in KPi buffer; D. 1 mM NADH in KPi buffer.

5.3.4 Hydrogenation-dechlorination activity assay using cell lysates of C. necator and B. megaterium

Native soluble hydrogenase expressed in heterotrophically grown C. necator and active

recombinant TmrA expressed in B. megaterium was used to reduce CF in vitro. Cell lysate

from C. necator and B. megaterium was incubated with H2-saturated KPi buffer and 0.1

mM CF at 30 ᵒC for 12 h. GC/MS results showed that albeit at a very small quantity, CF

was dechlorinated to DCM, the specific activity calculated to be 0.015±0.000 (SD)

nmol/min/mg. The various controls tested (Table 5.1) showed no detectable amounts

of DCM.

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5.3.5 IPTG-inducible expression of TmrA in C. necator

The pBBR_tmrA construct prepared in section 4.2.1.4 was used for IPTG-inducible

expression of TmrA in C. necator. C. necator transformants were cultured in different

media i.e., LB, 2YT, TB and rich media with varying concentrations of IPTG for different

lengths of time. However, no soluble expression of TmrA was detected on SDS-PAGE and

dechlorination assays showed no detectable DCM (data not shown). To check whether

transcription of tmrA was occurring, samples were collected from different induction

conditions grown in rich media and prepared for transcript analysis. The DNA gel showed

no presence of presence of mRNA for tmrA (Figure 5.8).

Figure 5.6. Transcript analysis of tmrA expressed in C. necator grown in rich media. Lane 1. E-Gel™ 1 Kb Plus DNA Ladder; Lane 2. positive control (pBBR_tmrA plasmid); Lane 3. 0.1 mM IPTG for 2 h; Lane 4. 0.1 mM IPTG for 4 h; Lane 5. 0.1 mM IPTG for 6 hr; Lane 6. 0.5 mM IPTG for 2 h; Lane 7. 0.5 mM IPTG for 4 h; Lane 8. 0.5 mM IPTG for 6 h; Lane 9. 1 mM IPTG for 2 h; Lane 10. 1 mM IPTG 4 h; Lane 11. 1mM IPTG for 6 h; Lane 12. Native C. necator. The framed section shows tmrA in the positive control.

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5.3.6 Rhamnose-inducible expression of TmrA in C. necator

tmrA gene without the TAT-signal and membrane spanning region was cloned into the

plasmid pKRrha by Golden Gate cloning (Figure 5.7). After confirming the correct

insertion of the gene by Sanger sequencing, C. necator was transformed by conjugation

with E. coli strain S17 containing pKRrha_tmrA.

Figure 5.7. Gene cloning and subsequent colony PCR. Gel A: Lane 1. E-Gel™ 1 Kb Plus DNA Ladder; Lanes 2-4. Linearized pKRrha; Lanes 5 & 6. tmrA Gel B: Lane 1. E-Gel™ 1 Kb Plus DNA Ladder; Lanes 2-8. Colonies 1-7 picked from plates containing E.coli transformed with tmrA cloned into pKRrha. The framed sections show the DNA of interest.

For expression of TmrA, C. necator was cultured in LB and FGN media. Expression was

induced upon inoculation and when OD600 reached 0.6. Induction with different

concentrations of rhamnose was tested. SDS-PAGE showed no distinguishable

prominent bands for soluble TmrA (Figure 5.8). Attaching a purification tag was not

considered while constructing the plasmid since the focus of the experiments in this

study was the dechlorination capacity of the recombinant protein and the expression

host rather than the purification of the recombinant protein, though in hindsight having

a tag would have aided in the detection of the expressed protein via western blotting.

Cell lysates at all rhamnose concentrations tested, both in LB and FGN media,

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demonstrated dechlorination of CF to DCM when analysed by GC/MS. Dechlorination

rates were higher for TmrA expressed in LB than FGN media (data not shown).

Figure 5.8. SDS-PAGE of rhamnose-inducible expression of TmrA in C. necator. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. LB, no induction; Lane 3. LB, 0.5 mM at inoculation; Lane 4. LB, 1.0 mM at inoculation; Lane 5. LB, 1.5 mM at inoculation; Lane 6. LB, 2.0 mM at inoculation; Lane 7. LB, 2.5 mM at inoculation; Lane 8. LB, 3.0 mM at inoculation; Lane 9. LB, 0.5 mM at OD600 0.6; Lane 10. LB, 1.0 mM at OD600 0.6; Lane 11. LB, 1.5 mM at OD600 0.6; Lane 12. LB, 2.0 mM at OD600 0.6; Lane 13. LB, 2.5 mM at OD600 0.6; Lane 14. LB, 3.0 mM at OD600 0.6; Lane 15. SeeBlue Pre-Stained Protein Standard; Lane 16. FGN, no induction; Lane 17. FGN, 0.5 mM at inoculation; Lane 18. FGN, 1.0 mM at inoculation; Lane 18. FGN, 1.5 mM at inoculation; Lane 19. FGN, 2.0 mM at inoculation; Lane 20. FGN, 2.5 mM at inoculation; Lane 21. FGN, 3.0 mM at inoculation; Lane 22. FGN, 0.5 mM at OD600 0.6; Lane 23. FGN, 1.0 mM at OD600 0.6; Lane 24. FGN, 1.5 mM at OD600 0.6; Lane 25. FGN, 2.0 mM at OD600 0.6; Lane 26. FGN, 2.5 mM at OD600 0.6; Lane 27. FGN, 3.0 mM at OD600 0.6; Lane 28. SeeBlue Pre-Stained Protein Standard. The framed section shows the expected position of TmrA.

C. necator expressing recombinant TmrA was lysed by sonication under anaerobic

conditions and incubated with a mastermix containing titanium (III) citrate and methyl

viologen and 0.1 mM CF at 30 ᵒC for 12 h. GC/MS showed dechlorination of CF to DCM

(Figure 5.9), the calculated specific activity was 44.4 ± 0.3 (n=2, uncertainty is the range)

nmol/min/mg. The same assay with heat-inactivated cell lysate demonstrated no

presence of DCM.

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Figure 5.9. CF dechlorination assay by A. C. necator grown in FGN media, no induction; B. C. necator grown in FGN media, induced with 3.0 mM Rhamnose. The arrows indicate CF and DCM in the chromatogram. The circles show CF and DCM in the scan. DCM was detected only in the rhamnose-induced sample.

The same activity assay when performed with H2-saturated KPi buffer instead of the

artificial electron donors also demonstrated CF dechlorination, the specific activity was

calculated to be 13.9 ± 0.0 (SD) nmol/min/mg. Negative control with heat-inactivated

cell lysates showed no CF dechlorination.

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5.4 DISCUSSION

In this chapter, the potential for C. necator to be a novel host for recombinant expression

of soluble and functional reductive dehalogenases was evaluated. The CF RDase, TmrA,

was successfully expressed heterologously in this host. The feasibility to utilize the

uptake Ni-Fe hydrogenases of C. necator as redox partners and induce in vivo and in vitro

dehalogenation of CF was also tested and yielded promising results.

C. necator has gained increasing attention in many biotechnological applications in the

recent years. It is well known for the production of biodegradable plastic (Atlić et al.,

2011, Reinecke and Steinbuechel, 2009) and has provided a platform for the production

of fuels (Bi et al., 2013), industrial chemicals (Crépin et al., 2016, Fei et al., 2013,

Grousseau et al., 2014, Lee et al., 2016, Li et al., 2012, Müller et al., 2013, Nybo et al.,

2015), polymers (Przybylski et al., 2015) and proteins (Barnard et al., 2005, Srinivasan et

al., 2003). The accessibility of its genome sequence (Pohlmann et al., 2006), flexible

metabolism (Alagesan et al., 2018a, Cramm, 2009, Fukui et al., 2014a, Riedel et al.,

2014), fast growth up to high biomass densities (more than 200 g/liter of biomass) when

cultured under heterotrophic conditions (Reinecke and Steinbuechel, 2009, Srinivasan

et al., 2002) and availability of genetic tools (Aboulnaga et al., 2018, Alagesan et al.,

2018b, Arikawa and Matsumoto, 2016, Bi et al., 2013, Fukui et al., 2011, Gruber et al.,

2016, Hanko et al., 2017, Li and Liao, 2015, Sydow et al., 2017) make it an ideal

expression host.

An important requirement for the recombinant expression of RDases is the availability

of sufficient corrinoid cofactors in the host cell. A number of attempts at expressing

RDases in soluble and active form in E. coli has failed presumably because of its inability

to synthesize corrinoids de novo leading to insufficient levels of corrinoid available in the

cells to support RDase expression (Jugder et al., 2018, Neumann et al., 1998, Suyama et

al., 2002). E. coli utilizes vitamin B12 as coenzyme in enzymes such as ethanolamine

ammonia-lyase (Scarlett and Turner, 1976) and methionine synthetase (Roth et al.,

1996). E. coli uptakes vitamin B12 from the environment to meet its requirements. The

transport of vitamin B12 across the outer membrane of E. coli is facilitated by the

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transmembrane receptor protein BtuB belonging to the family of TonB–dependent

transporters (Pieńko and Trylska, 2020). However, the presence of B12 in the growth

media represses the expression of Btub; the level of intracellular, rather than

extracellular B12 regulates the repression (Kadner, 1978). In this experiment the vitamin

B12 uptake capability of C. necator was demonstrated by undertaking a vitamin B12 assay

using Lactobacillus leichmannii. Soluble and active expression of TmrA in C. necator in

this study also suggests that it has sufficient levels of corrinoid available in the cells to

support expression of RDases. This finding aligns with the results of Black et al. (2018).

They reported the enhanced expression of isobutanol, whose activity is highly

dependent on the presence of vitamin B12 in the cytoplasm, when 1 mM of vitamin B12

was added to the culture media.

To consider C. necator as a dechlorinating organism, it is imperative that the bacterium

is tolerant to both the substrate and product organochloride, at least to some extent.

Chloroform and dichloromethane both are toxic organic compounds. Chloroform is a

widespread groundwater contaminant and is known to inhibit many microbial processes

such as methanogenesis (Weathers and Parkin, 2000), organohalide respiration of

chlorinated ethanes and ethenes (McMurdie et al., 2011), gene transcription in

Desulfitobacterium species (Futagami et al., 2013). Chloroform can inhibit microbial

activity at concentrations as low as 10 mg/L by interfering with membrane function

(Chidthaisong and Conrad, 2000). The genotoxic effects of DCM on bacterial strains are

also well documented (Gisi et al., 1999, Kayser et al., 2000, Kayser and Vuilleumier, 2001,

Vuilleumier, 2002). In this study C. necator grew in media containing up to 2.5 mM

chloroform and dichloromethane without any visible negative impact on growth. At the

end of 48 h incubation, the number of CFU on plates with C. necator growing with 2.5

mM CF and DCM was evaluated. High viable counts were observed in both cases, albeit

lower in the presence of DCM compared to CF.

In this study, the expression of TmrA in C. necator was first attempted with an IPTG-

inducible expression system. The tmrA gene cloned into pBBR1MCS-2 vector,

constructed in chapter four, was used for this purpose. Expression was attempted using

various concentrations of IPTG, over varying lengths of time, in different culture media.

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However, no soluble expression of TmrA was observed. Transcript analysis of cells

cultured in rich media with various concentrations of IPTG over 2 to 6 h showed no

evidence of TmrA mRNA. This was possibly due to poor plasmid stability; plasmid loss

has been previously reported for C. necator cells carrying expression plasmid with a

pBBR1 origin of replication (Fukui et al., 2014b, Gruber et al., 2014, Voss and

Steinbüchel, 2006). After the failure to express TmrA under the IPTG-inducible

expression system, tmrA (devoid of the TAT-signal and the membrane spanning region)

was cloned into the pKRrha plasmid described by Sydow et al. (2017) which had a

tetracycline resistance cassette (Tcr) and a rhamnose-inducible promoter (rhaPBAD).

Soluble and active TmrA was expressed with all the rhamnose concentrations tested (0.5

to 3 mM) in both LB and FGN media. Sydow et al. (2017) observed high tunability of

eGFP expressions with 0.4 to 1 mM rhamnose in different media, the highest level of

protein expression was achieved with 11 mM rhamnose; expression was increased 140-

fold in LB medium and around 60-fold in minimal medium. Similar to the observations

of Sydow and co-workers, the expression of TmrA in this study was tightly regulated and

expression was better in complex media than defined media.

Obligate OHRB such as Dehalobacter strain UNSWDHB are almost always restricted to

H2 as electron donors, however, the redox partners for the terminal RDases are still

unknown. The non-respiratory RDases on the other hand, are linked to genes encoding

putative redox partners or are fused to a redox module which links substrate reduction

to NADPH oxidation (Collins et al., 2018). Genomic, transcriptional, and biochemical

studies have postulated the involvement of membrane-bound hydrogenases in electron

transport in OHRB (Kruse et al., 2017, Kruse et al., 2015, Kube et al., 2005, Mansfeldt et

al., 2014, Morris et al., 2006, Nonaka et al., 2006). In this study, heterologous

reconstitution of oxidation-reduction of H2/CF by uptake hydrogenase from and TmrA

was evaluated.

Recombinant expression of hydrogenases is complex and arduous, so the native O2-

tolerant soluble hydrogenase (SH) of C. necator which is encoded on the mega-plasmid

pHG1 was chosen for this study. Culture of C. necator in minimal medium FGN facilitates

initial growth on the preferred carbon source, fructose and represses hydrogenase

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expression; fructose depletion switches to growth on glycerol and stimulate de-

repression of hydrogenase expression (Jugder et al., 2016b). Growth on heterotrophic

media also promotes expression of membrane-bound hydrogenase (Jugder et al., 2015,

Lauterbach and Lenz, 2013). In this study, C. necator was cultured in FGN media to

promote expression of SH, the functionality of which was tested by the activity assay

described by (Jugder et al., 2016b). When C. necator cell lysate containing functional SH

was mixed with B. megaterium cell lysate containing active recombinant TmrA and

incubated with H2-saturated KPi buffer and CF at 30 ᵒC for 12 h, very little dechlorination

of CF to DCM (specific activity 0.014 ± 0.000 nmol/min/mg) could be detected by GC/MS.

The various negative controls tested showed no sign of dechlorination. This could

suggest that the C. necator hydrogenase oxidized H2 which made electron available to

TmrA for the reduction of CF.

When recombinant TmrA was expressed in C. necator cultured in FGN media, both SH

and recombinant TmrA were active, the activity was confirmed for both by separate

assays. The specific activity for SH was not calculated whereas the specific activity for

recombinant TmrA was 44.4 ± 0.3 nmol/min/mg when titanium (III) citrate reduced

methyl viologen was used as the electron donor. Activity assay with the heat inaed cell

lysate showed no DCM formation. Incubation of cell lysate from C. necator cultured in

FGN media expressing recombinant TmrA with H2-saturated KPi buffer and CF at 30 ᵒC

showed dechlorination of CF to DCM which was detected by GC/MS, the calculated

specific activity was 13.9 ± 0.0 nmol/min/mg. The same assay carried out with heat-

inactivated cell lysate showed no dechlorination. The small amount of dechlorination in

the activity assays might be because of lack of sufficient levels of H2 in the assay mixture.

Due to infrastructural limitations the hydrogenation of KPi buffer was done at a different

location to where the activity assay was set up. H2 loss from the buffer could have

occurred during transportation or over the 12 h incubation period. Despite the various

experimental limitations, this novel study has demonstrated that a non-physiological

hydrogenase can support a H2-dependent organohalide reduction by TmrA. Due to time

constraints in vivo dechlorination by recombinant C. necator could not be tested.

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5.5 CONCLUSIONS

This is the first report of C. necator being used as an expression host for recombinant

RDases after confirming its capacity to uptake exogenous Vitamin B12. TmrA has been

successfully expressed in a soluble and functional form in C. necator. The soluble

expression of TmrA in this host also refutes the notion that RDases cannot be expressed

in hosts incapable of synthesizing vitamin B12 de novo. C. necator expressing

recombinant TmrA has also demonstrated in vitro dechlorination of CF with H2 as the

electron donor, instead of the artificial electron donor methyl viologen which is

commonly used otherwise. The tolerance of C. necator to both substrate (CF) and

dechlorination product (DCM) was also confirmed. Further studies will be needed for

the optimization of H2 oxidation coupled reduction of CF by TmrA and to test the

potential of in vivo CF dechlorination by recombinant C. necator. The findings in this

chapter will provide a platform to better understand the redox partners of RDases and

demonstrate prospects for engineering C. necator into a dechlorinating organism to be

used in bioremediation applications.

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CHAPTER SIX: General discussion and concluding remarks

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Dehalobacter sp. strain UNSWDHB is an obligate OHRB capable of respiring CF to DCM.

The sole electron donor for CF respiration is H2, which is oxidised by a membrane-bound

hydrogenase and the electrons are utilised by the reductive dehalogenase, TmrA for the

reductive dechlorination of CF. In this thesis, TmrA has been expressed in four different

expression hosts, Escherichia coli BL21(DE3)pLysS, Bacillus megaterium MS941,

Shimwellia blattae and Cupriavidus necator H16 strain DSM 428, with an aim to

compare the different expression systems and provide useful guidelines for future

recombinant RDase studies.

6.1 Reductive dehalogenases in organohalide respiration

Since the discovery of OHRB in the 1980s, extensive research has been carried out to

reveal the ecological, genomic, physiological, and biochemical features of this

taxonomically diverse group of bacteria. Research over the past two decades on OHRB

and their RDases has vastly improved our understanding of organohalide respiration and

their potential for application in bioremediation of organohalide-contaminated sites.

The discovery of the crystal structure of the respiratory PceA from Sulfurospirillum

multivorans and the catabolic NpRdhA from Nitratireductor pacificus has shed light on

the catalytic mechanism of RDases (Bommer et al., 2014, Payne et al., 2015).

Breakthroughs such as this opens doors for new paradigms to be explored and

understood. However, there are still many unanswered questions and many knowledge

gaps to be filled, such as 1) what are the biophysical and chemical properties that govern

substrate specificity of different RDases? 2) what factors govern the difference in

reaction mechanisms of RDases? 3) what are the different electron transfer steps in OHR

and why do they vary among OHRB? To answer these important questions, it is

imperative to produce sufficient amounts of RDases to conduct further studies. Isolation

of native RDases has many drawbacks; oxygen sensitivity and slow growth of OHRB, low

biomass and protein yield, the difficulty of working with membrane proteins are among

the major hurdles. Hence, efforts have been made to produce RDases heterologously.

However, the oxygen intolerance of respiratory RDases and their requirement for

cobalamin and Fe-S cluster cofactors complicate the process and not very many success

stories have been reported so far.

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6.2 Escherichia coli: the recombinant protein expression factory

E. coli is the most commonly used prokaryotic host for heterologous expression of

protein (Jia and Jeon, 2016). However, the soluble expression of RDases, particularly

respiratory RDases, in E. coli has not been very successful so far, though recovery from

inclusion bodies by in vitro reconstitution of cofactors has been possible for VcrA from

Dehalococcoides mccartyi strain VS and PceA from Geobacter sp. (Nakamura et al., 2018,

Parthasarathy et al., 2015). The incapability of E. coli to synthesize corrinoids de novo

resulting in insufficient amounts of cobamide cofactor in the cells to support RDase

expression has been suggested as the reason behind the failures (Neumann et al., 1998,

Suyama et al., 2002). The catabolic RDase, NpRdhA, from Nitratireductor pacificus has

been expressed in a soluble and active form in E. coli strains BL21 and HMS174(DE3) by

co-expression with a vitamin-B12 transporter btuB (Collins, 2017, Halliwell et al., 2021).

In this study, TmrA was recovered in a catalytically active form from inclusion bodies

expressed in E. coli BL21(DE3)pLysS. The refolding and cofactor reconstitution methods

used in the experiments reveal that successful refolding depends on the availability of

proper cofactors to assist in the correct refolding. Lack of sufficient cofactors in SDS-

MPD refolding method attempted in this study resulted in inactive TmrA. The most

promising aspect of this study was the revelation that the TmrA inclusion bodies could

be purified in the presence of air prior to the cofactor reconstitution and resume activity

when the reconstitution is done under anaerobic conditions. Protein purification inside

an anaerobic chamber is complicated and laborious. Long hours of working inside an

anaerobic chamber at a stretch is also not very ergonomic for the technician. The

method of aerobic purification and anaerobic cofactor reconstitution described in this

study offers a sustainable production pipeline for RDases and other similar proteins.

6.3 Bacillus megaterium: the big beast

The corrinoid producing Gram-positive Bacillus megaterium has been identified as a

suitable host for the expression of soluble and functional RDases. Both respiratory and

catabolic RDases have been successfully expressed in B. megaterium (Jugder et al., 2018,

Payne et al., 2015). Jugder and co-workers (2018) successfully expressed TmrA, the

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RDase of interest in this thesis, in B. megaterium. However, the recombinant TmrA had

an 11-fold lower specific activity than that calculated for the native enzyme (Jugder et

al., 2017). In this experiment various culture and induction conditions were screened to

improve the soluble expression and specific activity of recombinant TmrA in B.

megaterium. A serendipitous find was that adding much lower concentrations of xylose

than what is conventionally used, particularly when induction is done at 37 °C for 3 h,

leads to higher specific activity of the recombinant TmrA. Screening experiments done

by full factorial design supported this assertion. The best induction conditions revealed

by the screening experiments were induction with 0.03% xylose for 3 h at 37 °C.

Statistical analysis by JMP of the soluble expression and specific activity of recombinant

TmrA supports this finding, the desirability function (0.5) was the highest for this

condition. The desirability function endeavours to determine the best compromise

among all the responses involved in a process (Candioti et al., 2014). The overall

desirability for all responses (soluble expression and specific activity in this study) is

interpreted as the geometric mean of the desirability functions for the individual

responses. Recombinant TmrA purified from B. megaterium induced with 0.03% xylose

for 3 h at 37 °C had a specific activity of (3.6 ± 0.3) x 103 nmol/min/mg protein which is

32.7 times higher than specific activity of the recombinant TmrA expressed in B.

megaterium by Jugder et al., (2018) and 2.8 times higher than the specific activity for

native TmrA calculated by Jugder et al., (2017).

6.4 Shimwellia blattae: the vitamin B12-producing gut bacteria from cockroach

The corrinoid-producing Gram-negative gammaproteobacterium Shimwellia blattae has

been used as an expression host for the soluble expression of three respiratory RDases;

PceA from Desulfitobacterium hafniense strain Y51, RdhA3 from Desulfitobacterium

hafniense strain DCB-2, and DceA from Desulfitobacterium dichloroeliminans DCA1 and

also the catabolic RDase NpRdhA from Nitratireductor pacificus (Halliwell et al., 2021,

Kunze et al., 2017, Mac Nelly et al., 2014). In cases of all three respiratory RDases, co-

expression of the respective chaperone protein resulted in higher soluble expression

and specific activity of the recombinant RDase. TmrA does not have a specific chaperone

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in the tmrAB operon. To co-express TmrA with a chaperone protein, two chaperones

(WP_015043727 and WP_015044672) were chosen from the genome of Dehalobacter

sp. UNSWDHB, they were referred to as TF104 and TF161 in this study. TmrA was

expressed in S. blattae in different forms and combinations: TmrA without the TAT

signal, TmrA including the TAT signal, co-expression of TAT signal included TmrA with

TF104 and co-expression of TAT signal included TmrA with TF161. All these experiments

resulted in the soluble and functional expression of TmrA in S. blattae. However, the

results were not what was expected. Co-expression with the chaperone proteins did not

seem to have any apparent effect on the soluble expression level, however, the

TmrA devoid of the TAT signal displayed the highest specific activity amongst them all.

These findings give rise to an interesting puzzle; the three respiratory RDases previously

expressed in S. blattae were all expressed with their TAT signal intact, and all of them

exhibited soluble expression alone while co-expression with their respective chaperone

proteins increased their soluble expression, they, however, were not expressed without

their TAT signals. The questions that arise here are: 1. Would eliminating the TAT signal

from the other three respiratory RDases increase their specific activity? 2. Do the TF104

and TF161 chaperones co-expressed in this study compete/interact/antagonise with the

native S. blattae chaperones? The answer to these questions would assist in developing

an ideal recombinant RDase expression system in S. blattae and other expression hosts

alike.

6.5 Cupriavidus necator: the “knallgas” bacteria

In this study C. necator was introduced as a novel host for recombinant RDase

expression. The broader and longer-term aim was to engineer C. necator into a

dechlorinating organism. The preliminary studies done to verify its suitability as a

dechlorinating organism were: 1. The capability to uptake vitamin B12 from the media;

2. Tolerance to substrate, CF and breakdown product, DCM; 3. The suitability of the C.

necator soluble hydrogenase to oxidize the electron donor H2 to support CF reduction

by TmrA. C. necator demonstrated favourable results in all three experiments. TmrA was

expressed in a soluble and active form in C. necator. The specific activity was higher

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when expressed in LB media than in FGN media but culture in the heterotrophic FGN

media promotes expression of the soluble hydrogen which is required for the cellular

dechlorination of CF in a reductive dehalogenation process. In vitro activity assay with

cell lysates of the recombinant C. necator using H2 as the electron donor demonstrated

dechlorination of CF albeit at very small quantities and high amount of background

noise. Due to time constraints this dechlorination system could not be optimized and in

vivo dechlorination assays could not be tested. However, the results obtained

demonstrate C. necator as a potential expression host for recombinant RDases.

6.6 Comparing the four expression systems

To-date no single respiratory RDase has been successfully expressed in multiple

expression hosts, so the comparison of the available expression systems for RDases

could not be compared. Very recently Halliwell et al., (2021) published a comparative

study of the heterologous expression of a catabolic reductive dehalogenase, NpRdhA,

from Nitratireductor pacificus strain pht-3B in Bacillus megaterium, Escherichia coli

HMS174(DE3), Shimwellia blattae and a commercial strain of Vibrio natrigenes. In this

study CF RDase from Dehalobacter sp. UNSWDHB was expressed in a soluble and active

form in four expression hosts Escherichia coli BL21(DE3)pLysS, Bacillus megaterium

MS941, Shimwellia blattae and Cupriavidus necator H16 strain DSM 428; albeit to a

varying degree of success. Halliwell et al., (2021) reported that recombinant NpRdhA

expressed in B. megaterium had the highest specific activity while the recombinant

NpRdhA in S. blattae had the lowest specific activity. In this study, the highest specific

activity of recombinant TmrA was achieved by B. megaterium, matching the findings of

Halliwell et al., (2021).

Halliwell and co-workers attempted to optimize the culture conditions for all the

expression hosts tested, however, that was always not possible since different host

organisms have different growth requirements, e.g., V. natrigenes gave better results

when cultured in 2YT media instead of TB media; S. blattae was induced under anaerobic

conditions. In the current study, the best growth and expression conditions

recommended in literature for each expression host was used. In terms of complexity of

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the expression system, culture and protein expression in S. blattae and C. necator was

much more complicated that the other two hosts. Expression in S. blattae required three

pre-culture steps and the main culture was anaerobic which was difficult to maintain

compared to the other systems and also resulted in low yield of cells. Due to time

limitations, purification of recombinant TmrA expressed in S. blattae could not be done

but the previous studies with the respiratory RDases by Mac Nelly et al., (2014) and

Kunze et al., (2017) were not very successful in terms of protein purification. Expression

in C. necator requires a long induction period whereas TmrA expression in E. coli and B.

megaterium can be accomplished in one day. Based on the findings of this study, B.

megaterium demonstrated the best results in terms of production process, soluble

expression, and specific activity. However, a major limitation of the B. megaterium

expression system is the 3-day transformation protocol (Collins, 2017).

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6.7 Standout findings of this study

The expression of recombinant TmrA in a soluble and catalytically active form was

successful in all four expression hosts tested in this study. The most important findings

of this study are summarised below:

Purification of inclusion bodies expressed in E. coli in presence of air does not render

them inactive after cofactor reconstitution. This is a very important finding in terms of

optimizing the recombinant protein production pipeline. The functional and structural

studies of RDases requires the production of a suitable biomass of purified protein.

Purification of protein under anaerobic conditions is complicated, expensive, and not to

mention not very ergonomic. Additionally, purification using an ÄKTA or similar system

would provide a much better control over the purification step than manual injection or

using a peristaltic pump.

Xylose concentrations much lower than the conventionally used amounts increase

specific activity of recombinant TmrA. The xylose concentration used to induce

recombinant RDase production in B. megaterium was 0.1% for NpRdhA and 0.5% for

TmrA. Literature review revealed that similar concentrations have been used for the

expression of other recombinant proteins in B. megaterium. Miscalculations while

preparing the xylose stock solution prompted expression screening with 10 x and 100 x

lower than the conventionally used xylose concentrations. Induction with 0.03% xylose

for 3 h at 37 °C produced recombinant TmrA with a specific activity which is more than

30% higher than that measured in the previous study.

Excluding the TAT signal increases specific activity. In this study recombinant TmrA was

produced in all four expression hosts without the TAT signal and the membrane

spanning region. In S. blattae recombinant TmrA was expressed both with and without

the TAT signal and the TmrA without the TAT signal demonstrated significantly higher

(p<0.05) specific activity than when expressed with the TAT signal. These results brought

to light that removing the TAT signal of the RDase of interest for recombinant expression

could potentially improve their specific activity. The recombinant TmrA expressed in B.

megaterium without the TAT signal showed approximately 3 times higher specific

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activity than that calculated for the native enzyme. Recombinant expression of VcrA

from Dehalococcoides mccartyi strain VS in E. coli was also done devoid of the TAT signal

and demonstrated high specific activity (Parthasarathy et al., 2015).

Non-native hydrogenases can support CF dechlorination by TmrA. The activity assays

done for all the native and recombinant RDases are done using artificial electron donors

such a methyl viologen. The only exceptions are the 3-Chlorobenzoate-RDase from

Desulfomonile tiedjei DCB-1 in which case a partially purified native hydrogenase was

used in the assay mixture alongside methyl viologen (Ni et al., 1995) and the

recombinant catabolic NpRdhA where NADPH, spinach ferredoxin and E. coli flavodoxin

reductase was used. In this study the recombinant TmrA expressed in C. necator

demonstrated dechlorination of CF using H2 as the electron donor which was oxidized

by the hydrogenases of the expression host. These results will provide a platform to

study or even replicate the electron transport chain of RDases in non-native hosts.

6.8 Future research

The structural and functional studies of RDases require their production in sufficient

amounts to support comprehensive biochemical and biophysical studies. This

dissertation provides a comprehensive study on various recombinant expression

systems for one RDase, TmrA from Dehalobacter sp. UNSWDHB. Future studies based

on the findings presented in this document could be

• Optimizing the cofactor reconstitution and refolding protocol described for E.

coli expression

• Determining the oxygen-sensitivity level for TmrA and other RDases which will

aid in designing optimum protein purification protocols

• Studying the kinetics of the recombinant TmrA produced in different hosts and

see how it compares to the native enzyme

• Carry out further investigation into the impact of the TAT signal on the

expression and specific activity of recombinant RDases

• Optimizing the use of non-native redox partners in activity assays for RDases

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• Optimizing the recombinant RDase expression system in C. necator

• In vivo CF dechlorination assay with recombinant C. necator grown under

autotrophic and heterotrophic conditions

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6.9 References


CANDIOTI, L. V., DE ZAN, M. M., CÁMARA, M. S. & GOICOECHEA, H. C. 2014. Experimental design and multiple response optimization. Using the desirability function in analytical methods development. Talanta, 124, 123-138.

COLLINS, F. A. 2017. A biochemical study of a catabolic reductive dehalogenase. PhD, Manchester University.

HALLIWELL, T., FISHER, K., PAYNE, K. A. P., RIGBY, S. E. J. & LEYS, D. 2021. Heterologous expression of cobalamin dependent class-III enzymes. Protein Expression and Purification, 177, 105743.

JIA, B. & JEON, C. O. 2016. High-throughput recombinant protein expression in Escherichia coli: current status and future perspectives. Open Biology, 6(8): 160196.


JUGDER, B. E., BOHL, S., LEBHAR, H., HEALEY, R. D., MANEFIELD, M., MARQUIS, C. P. & LEE, M. 2017. A bacterial chloroform reductive dehalogenase: purification and biochemical characterization. Microbial Biotechnology, 10(6): 1640-1648.

KUNZE, C., DIEKERT, G. & SCHUBERT, T. 2017. Subtle changes in the active site architecture untangled overlapping substrate ranges and mechanistic differences of two reductive dehalogenases. FEBS, 284(20): 3520-3535.

MAC NELLY, A., KAI, M., SVATOŠ, A., DIEKERT, G. & SCHUBERT, T. 2014. functional heterologous production of reductive dehalogenases from Desulfitobacterium hafniense strains. Applied and Environmental Microbiology, 80(14): 4313.



NI, S., FREDRICKSON, J. K. & XUN, L. 1995. Purification and characterization of a novel 3-chlorobenzoate-reductive dehalogenase from the cytoplasmic membrane of Desulfomonile tiedjei DCB-1. Journal of bacteriology, 177(17): 5135-5139.


PAYNE, K. A. P., QUEZADA, C. P., FISHER, K., DUNSTAN, M. S., COLLINS, F. A., SJUTS, H., LEVY, C., HAY, S., RIGBY, S. E. J. & LEYS, D. 2015. Reductive dehalogenase

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structure suggests a mechanism for B12-dependent dehalogenation. Nature, 517(7535): 513-516.

SUYAMA, A., YAMASHITA, M., YOSHINO, S. & FURUKAWA, K. 2002. Molecular characterization of the PceA reductive dehalogenase of Desulfitobacterium sp. strain Y51. Journal of Bacteriology, 184(13): 3419.

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Appendices

209

A.1 Vector Maps

Figure A1.1. Vector map of plasmid used in E. coli experiments.

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Figure A1.2.Vector map of plasmid used in E. coli experiments.

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Figure A1.3. Vector map of plasmid used in E. coli experiments.

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Figure A1.4. Vector maps of plasmids used for recombinant expression of TmrA in B. megaterium (Gamer et al., 2009).

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Figure A1.5. Vector maps of plasmids used in S. blattae experiments.

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Figure A1.6. Vector map of the plasmid used in recombinant TmrA expression in C. necator.

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A.2 Sequence of genes used in the experiments A.2.1 tmrA without TAT-signal ATGATTGATCCTAAACAAGTTTACGCGGGTACCGTTAAAGAACTGGATGAACTGCCGTTCAATATTCCGGCGGATTACAAGCCGTTTACGAACCAGCGTAACATCTTCGGCCAAGCTGTCCTGGGTGTTCCGGAGCCGCTGGCACTGGTCGAGCGTTTCGACGAAGTTCGCTGGAATGGCTGGCAGACCGACGGCAGCCCGGGTCTGACCGTCTTGGATGGCGCCGCGGCACGTGCGTCTTTTGCGGTTGACTATTACTTTAACGGTGAGAACAGCGCCTGCCGTGCGAATAAAGGTTTCTTTGAGTGGCATCCTAAAGTGCCGGAACTGAACTTCAAGTGGGGTGACCCGGAGCGCAATATTCACAGCCCGGGTGTCAAGAGCGCAGAAGAAGGCACCATGGCAGTGAAGCGCATGGCCCGTTTCTTCGGCGCAGCGAAGGCTGGCATTGCGCCGTTCGATAAACGTTGGGTGTTTACGGAAACCGCTGCGTTTGTGAAAACTCCTGAGGGCGAGGACCTGAAGTTCATTCCGCCGGACTTTGGCTTTGAGCCGAAACACGTGATCTCCATGATCATCCCGCAGAGCCTTGAGGGTGTGAAGTGTGCGCCGTCGTTCTTGGGTAGCGCGGAGTATGGTCTGAGCTTTGCACAAATCGGTTACGCAGCATTCGGCCTGTCTATGTTCATTAAAGATCTGGGTTATCACGCAGTTCCAATCGGCAGCGATAGCGCGCTGAGCATTCCAATCGCCATTCAGGCTGGTCTGGGCGAGTACAGCCGTAGCGGCCAAATGATTACCCCGGAATTCGGTCCGAATGTGCGTCTGTGCGAAGTCTTTACGGATATGCCGCTGAATCACGACAAGCCGATTAGCTTCGGTGTTACTGAGTTTTGTAAGACCTGTAAGAAATGCGCAGAGGCGTGTCCGCCGCAGGCCATCAGCTACGAAGATCCGACCATTGATGGTCCGCGCGGCCAGATGCATAACTCCGGCATCAAGCGTTGGTATGTTGACCCGGTAAAATGTTTTGAGTTTTGGAGCCGTGACAACGTCCGCAACTGCTGCGGTGCGTGTATCGCGGCGTGCCCGTTCACCAAACCAGAGGCCTGGCATCACACGTTGATTCGCTCTCTGGTGGGTGCACCGGTGATTACGCCGTTCATGAAAGATATGGACGACATCTTTGGTTACGGTAAGCCGAATGAAAAGGCTAAAGCCGATTGGTGGAAG

216

A.2.2 tmrA with TAT-signal ATGGACAAGGAGAAGTCGAATAATGATAAGCCGGCAACAAAGATCAACCGTCGCCAGTTCCTGAAATTTGGGGCCGGGGCTTCTTCCGGTATTGCGATTGCGACGGCGGCCACGGCCCTGGGAGGAAAGAGCCTTATCGATCCGAAACAGGTGTACGCGGGAACAGTTAAGGAGCTGGACGAATTGCCCTTCAATATTCCTGCAGATTACAAACCTTTCACCAATCAGCGTAACATCTTCGGACAAGCAGTATTAGGCGTACCAGAACCGTTAGCCCTGGTCGAGCGTTTCGACGAAGTACGTTGGAATGGATGGCAAACAGACGGAAGTCCCGGTCTGACTGTGTTAGATGGGGCAGCCGCGCGTGCCTCGTTTGCAGTGGATTATTATTTTAATGGGGAGAACTCTGCGTGTCGTGCCAACAAAGGGTTCTTTGAATGGCACCCAAAAGTCCCGGAACTTAATTTCAAATGGGGAGATCCAGAGCGCAATATTCACTCACCGGGAGTTAAATCAGCGGAAGAAGGCACCATGGCGGTAAAACGTATGGCTCGCTTTTTCGGGGCGGCCAAAGCTGGCATCGCTCCATTCGATAAACGCTGGGTGTTTACCGAAACGGCTGCGTTCGTCAAGACGCCTGAGGGGGAAGACTTGAAATTCATTCCACCCGATTTTGGTTTTGAGCCTAAACACGTAATCTCTATGATTATTCCACAAAGCTTAGAAGGCGTCAAGTGTGCCCCTAGTTTCCTTGGCTCTGCCGAGTATGGTCTTTCCTTCGCGCAGATCGGATATGCAGCTTTCGGTCTTTCTATGTTCATTAAAGATTTAGGCTATCATGCGGTACCGATCGGTAGTGATTCGGCTTTATCCATCCCCATTGCTATTCAGGCCGGACTGGGCGAGTATTCTCGTTCTGGTCAGATGATCACTCCGGAGTTCGGACCTAACGTGCGCTTATGTGAGGTATTTACAGATATGCCATTAAACCACGATAAGCCGATTTCATTCGGTGTAACAGAATTCTGCAAAACTTGTAAAAAATGCGCAGAGGCGTGCCCGCCACAGGCTATTTCCTACGAAGACCCCACGATCGACGGTCCTCGTGGCCAAATGCATAACTCGGGGATTAAGCGCTGGTATGTTGATCCCGTCAAGTGTTTTGAGTTCTGGAGTCGCGATAACGTCCGTAATTGCTGCGGTGCTTGCATTGCGGCATGTCCCTTCACAAAACCTGAGGCTTGGCACCATACCCTTATTCGTTCCTTGGTAGGAGCGCCAGTGATCACTCCGTTTATGAAAGACATGGACGACATCTTTGGGTACGGTAAGCCTAACGAGAAGGCAAAGGCGGACTGGTGGAAA

217

A.2.3 tf104 ATGAAGCAATTCGAGCTTGGACAATACAAGGGGCTTAATGTCAAACGCTTTGATACTACGGTTCAAGAGGAAGAAATTCAGCAGGCGCTTGATTACATTATTGGTAGTTTTGACGAAATCGAAGAAGAAAAACGTAATGAGCCCATTAAGACAAATGACTATGTTATTGTCGACATTGACGGTTTTGAGAAGGACGCGACGGTTCCTGTCATTCGCTCGCTGGACACCAAGCTTCGCGTCGGTTCAGAGGGCGTGTTTCAGGAAGTAAGCGCTAACTTATTAGGCAAAAAAATGGGAGACACTGTTACTTTCGAGGCCGTCATCCAGCCTGACGCTTTAGAGTTCCAGCGCTGGTGGGGTTCCGAAATCCGCTTCACAGTGAAGATCAAGTCTGTGTTTGTAGTGAAGAAGTCGGAACTTACAGAGGAGTTAATTCGCAAGATCGAGCCGGACCTTAAGAACTTGGAAGACTTAAAAAATATGTTGGCATTGAAGATCACGCACGAGAAGGAAGGCAAAGAGCGCGAAGCCAACATCTTGCTGATCTTCCAAGCTTTGGTGAAACAGTGTAAATATGAATTTGACGAAGAGGAACTTGACTCTGCGGCGGAAGACCTTTACAAAAAGTTTACCGAGGAGTTAAAGATCGTCGACGACATGGAGTTAATGGAGTACCTGATCCACCGTAAAATTACAGCCGATCAACTTTTAGTAGAATGCAAGGAGGAAGCTAGTCGCCGTATCTTGTGGGAACTTATGATCAACTCAGTCATTGAGAAAGAGGAAATCAATCTTACCCCTGATGAGATCAAATACTTGGAGAAACGTATTAATGAATCGCGTCAGAATGGTCAATTACCGGAAGAATTCATGGATATTAATTTTCTTCAAGCACCCTATCTGCGCAAAAAGACCATGGATTATCTTTTAGAAATTAATTTAGCTAGT

218

A.2.4 tf161 ATGTCGGTCAAGATCGAGAAGAAGAACAATAACATTTATGAGATGGAGATCATGGTAGGCGTAGAAGAGGTCTCAGCAGCATTTGACCGCGTGATGAAGCGCGCGGGCCAGGGTTTGAGTATTCCAGGGTTTCGTAAAGGAAAGGCACCAAAGCATATTATTGAACGTTATGTGGATAAAGACGCAGTGAATAATGAGGTCATGGAGCAAGTATCATATCCTGCTTTATTTGAAGCTTATAAAGAACACAGCATTACACCTGTTTCCCGCCCAGCTATGCAGGTCGTCCAATTTGAGGCAGACAAGGACTTGATCTTCAAAGTAACTGTGGAGACAAAACCTGATGTCGAACTTGGCCAATACAAAGGCTTAGGCATTGAAAAGCAAGCGGTAGAGGTCACGGATGAACAGGTAGCCGAGGAGTTGGAGCGTCGTCAAAATCGTCACGCTAAATTGATTCCCGTAGAAGACGGAGAAATCATTGATCAGGATATTGTCACCATCGACTTCGAGGGTTTCCTGGGCGATGTCCCGTTTGAAGGTGGTAAGGGAGAGAACCACGAGTTGACCATCGGATCAGGAACTTTTATTCCGGGATTTGAGGAGCAGATTAAAGGCTCCAAGAATGGACAAGAACTGGAAATTAACGTTAAGTTCCCGGAGGGTTATCACAGCGAGGAACTGTCTGGGAAGGATGCAATTTTCAAGGTTAAGATTAATGGCATCAAACGTAAGGAATTAGCCGCGTTGGACGACGAGTTTGCAAAAGACGTCTCCGAATTTGACACCCTTGAGGAACTTAAGCAGGACATCCGTGAGAAGATGATCTCAGCCGCTGAGACTCGTCGTGATAATGAATACAAAACTGAAGTAATCAAGAAGGTAGCCGAGAACGCGGTTTGCGAAATCCCTGAGGGTATGAAGCAAGAGCGTATTGACGCGCTTATGGAGGATCTTCAACAGAATATGTCCTATCAAGGAATCTCGATGGAGCAGTATTGCCAGTATGTCAATACGAGTCGCGAAGCACTTCGTGAATCGTTCTCTCAGCAAGCTACGGATGGCTTGAAAACCGAACTTGTATTAGAAGCGATCGCAGAAAAAGAGGGGATCACGGTCACGGACGAGGAAATTGAAAGTGAGATCAACCGTATGGCGATGCAATATGGACGTACTGCGGAAGAGTTGAAGAGTGCTCTTGAAGCGCGTGGCGAGATGCAGATGTTCAAAATTTCTCTTACAAGCGAGCATACTGTGGACTTCTTAGTAAAAAATAATGGCGCGGGTCAAGAATCTGCAGCCGCCCAAAATACCGACTCCGAGTCTAATGTGGCGGAAGAGAACGAA

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A.3 Composition of culture media and buffers Media Composition

Defined media for S. blattae 14.0 g/L K2HPO4 6.0 g/L KH2PO4 3.0 g/L (NH4)2SO4 0.2 g/L MgSO4 x 7 H2O 8.66 µM CoCl2 300 mM Glycerol 0.02% Yeast Extract 0.2 mL SL10 trace element solution Milli-Q Water up to 1 L

Fructose-Glycerol-Nitrogen (FGN) medium 100 mL 10X H16 buffer 10 mL 20% NH4Cl 1 mL 20% MgSO4 x 7 H2O 1 mL 1% CaCl2 x 2 H2O 1 mL 0.5% FeCl3 x 6 H2O 1 ml 0.02% NiCl2 5 mL 40% Fructose 5 mL 40% Glycerol 876 mL Milli-Q water

Fructose-Nitrogen (FN) medium 100 mL 10X H16 buffer 10 mL 20% NH4Cl 1 mL 20% MgSO4 x 7 H2O 1 mL 1% CaCl2 x 2 H2O 1 mL 0.5% FeCl3 x 6 H2O 1 ml 0.02% NiCl2 10 mL 40% Fructose 876 mL Milli-Q water

Buffers Composition

10X H16 buffer pH 7.0 90 g Na2HPO4 x 12 H2O 15 g KH2PO4 Milli-Q water up to 1 L

50 mM KPi (potassium phosphate) buffer pH 21.1 mL 1 M KH2PO4 7.0 28.9 mL 1 M K2HPO4

Milli-Q water up to 1 L

SL10 Trace Element solution 10 mL 25% HCl 1.5 g FeCl2 x 4 H2O 190 mg CoCl2 x 6 H2O 100 mg MnCl2 x 4 H2O 70 mg ZnCl2 6 mg H3BO3 36 mg Na2MoO4 x 2 H2O 24 mg NiCl2 x 6 H2O 2 mg CuCl2 x 2 H2O Milli-Q water up to 1 L

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A.4 Dechlorination activity assay calculation data

Table A.4.1 Raw data for standard curve construction

Concentration (mM) Relative Abundance

DCM 0.025 8107993 0.05 15518906

0.075 24144500 0.1 35268650

0.15 47139389 0.2 61668525 0.5 202290523 CF

0.025 3204185 0.05 6177159 0.75 10591086 0.1 14847746

0.15 23596729 0.2 29819616 0.5 81328068

Figure A.4.1. DCM standard curve for dehalogenation activity calculation.

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Figure A.4.2. CF standard curve for dehalogenation activity calculation.

Table A.4.2 Raw data for dechlorination activity assay for E. coli samples

Construct DCM produced (nmol)

Amount of protein added to assay

(mg)

Specific activity (nmol/mg/min)

Corrected specific activity (nmol/mg/min)

TmrA-TF_1 10510.67 0.04 1460 30 TmrA-TF_2 10995.37 0.04 1527 97 TmrA-TF_3 10674.415 0.04 1483 53 Heat-inactivated TmrA-TF_1

10339.08 0.04 1436 -

Heat-inactivated TmrA-TF_1

10257.885 0.04 1425 -

TmrA-Chis_1 10865.37 0.3 3018 34 TmrA-Chis_2 10899.405 0.3 3028 44 Heat-inactivated TmrA-CHis

10742.795 0.3 2984 -

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Table A.4.3 Raw data for dechlorination activity assay for B. megaterium samples

Culture Condition DCM produced (nmol)


(mg)


TB media, 0.1% xylose, 3h_1 19985.005 273 0.41 TB media, 0.1% xylose, 3h_2 23254.015 247 0.52 TB media, 0.1% xylose, overnight_1 12382.895 7 9.83 TB media, 0.1% xylose, overnight_2 12397.895 6.5 10.6 TB media, 0.3% xylose, 3h_1 11262.17 266 0.24 TB media, 0.3% xylose, 3h_2 11757.91 261 0.25 TB media, 0.3% xylose, overnight_1 10197.435 5 11.33 TB media, 0.3% xylose, overnight_2 13851.935 6.5 11.84 TB media, 0.5% xylose, 3h_1 11045.54 271.5 0.23 TB media, 0.5% xylose, 3h_2 10313.23 254.5 0.001 TB media, 0.5% xylose, overnight_1 10200.775 3 18.89 TB media, 0.5% xylose, overnight_1 10549.935 5.5 10.66 TB media, 0.01% xylose, 3h_1 142558.3272 193 4.1 TB media, 0.01% xylose, 3h_2 158953.9982 170 5.19 TB media, 0.001% xylose, 3h_1 99437.80481 156 3.54 TB media, 0.001% xylose, 3h_1 121439.113 157.5 4.28 TB media, 0.03% xylose, 3h_1 191791.6906 171 6.23 TB media, 0.03% xylose, 3h_2 151955.6192 171 4.93 TB media, 0.003% xylose, 3h_1 122957.7172 162.5 4.2 TB media, 0.003% xylose, 3h_2 108763.5867 174 3.47 TB media, 0% xylose, 3h_1 92139.70159 216.5 2.36 TB media, 0% xylose, 3h_2 99622.84708 212 2.61 Purified protein_1 12599.08 0.02 3499.743 Purified protein_2 13987.79 0.02 3885.497 Purified protein_3 11940.39 0.02 3316.775

Table A.4.4 Raw data for dechlorination activity assay for S. blattae samples

Construct DCM produced (nmol)


(mg)


TmrA_1 43496.06 66 1.098 TmrA_2 48009.97 66 1.212 TmrA_full_1 10142.75 100 0.169 TmrA_full_2 10241.72 100 0.171 TmrA_full co-expressed with TF104_1 10803.6 66 0.273 TmrA_full co-expressed with TF104_2 10712.83 66 0.271 TmrA_full co-expressed with TF161_1 10626.54 76 0.233 TmrA_full co-expressed with TF161_1 10681.25 76 0.234

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Table A.4.5 Raw data for dechlorination activity assay for C. necator samples

Sample DCM produced (nmol)

Amount of protein added to assay (mg)


CN+BN_1.1 10041.52 992.5 0.014 CN+BN_1.2 10041.97 992.5 0.014 CN+BN_1.3 10020.37 992.5 0.014 CN+BN_2.1 10035.84 992.5 0.014 CN+BN_2.2 10051.1 992.5 0.014 CN+BN_2.3 10042.92 992.5 0.014 Rec. CN_1 10304.17 0.32 44.722 Rec. CN_2 10164.05 0.32 44.115 Rec. CN+BN_1.1 10005.68 1 13.897 Rec. CN+BN_1.2 10014.09 1 13.908 Rec. CN+BN_1.3 - 1 - Rec. CN+BN_2.1 10015.48 1 13.910 Rec. CN+BN_2.2 10010.45 1 13.903 Rec. CN+BN_2.3 10018.18 1 13.914

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A.5 Screening of recombinant B. megaterium for expression of TmrA using an anti-His monoclonal antibody in a Western blot assay

Figure A.5.1. Western blot images of screening conditions A. LB, OD600 0.3, xylose conc. 0.3%, up to OD600 1.5; B. LB, OD600 0.6, xylose conc. 0.3%, up to OD600 1.5; C. TB, OD600 0.3, xylose conc. 0.1%, up to OD600 1.5; D. TB, OD600 0.6, xylose conc. 0.3%, up to OD600 1.5. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Pre-induction soluble fraction (R1); Lane 3. Pre-induction insoluble fraction (R1); Lane 4. Post-induction soluble fraction (R1); Lane 5. Post-induction insoluble fraction (R1); Lane 6. Pre-induction soluble fraction (R2); Lane 7. Pre-induction insoluble fraction (R2); Lane 8. Post-induction soluble fraction (R2); Lane 9. Post-induction insoluble fraction (R2).

R1 = Replicate 1; R2 = Replicate 2

225

Figure A.5.2. Western blot images of screening conditions. A. LB, OD600 0.3, xylose conc. 0.3%, 6 h; B. LB, OD600 0.3, xylose conc. 0.5%, 6 h; C. TB, OD600 0.3, xylose conc. 0.5%, 6 h; D. TB, OD600 0.6, xylose conc. 0.3%, 6 h. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Pre-induction soluble fraction (R1); Lane 3. Pre-induction insoluble fraction (R1); Lane 4. Post-induction soluble fraction (R1); Lane 5. Post-induction insoluble fraction (R1); Lane 6. Pre-induction soluble fraction (R2); Lane 7. Pre-induction insoluble fraction (R2); Lane 8. Post-induction soluble fraction (R2); Lane 9. Post-induction insoluble fraction (R2).


226

Figure A.5.3. Western blot images of screening conditions. A. LB, OD600 0.6, xylose conc. 0.3%, 3 h; B. LB, OD600 0.6, xylose conc. 0.5%, 3 h; C. LB, OD600 0.3, xylose conc. 0.1%, 3 h; D. TB, OD600 0.6, xylose conc. 0.3%, 3 h; E. TB, OD600 0.6, xylose conc. 0.5%, 3 h; F. TB, OD600 0.3, xylose conc. 0.1%, 3 h. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Pre-induction soluble fraction (R1); Lane 3. Pre-induction insoluble fraction (R1); Lane 4. Post-induction soluble fraction (R1); Lane 5. Post-induction insoluble fraction (R1); Lane 6. Pre-induction soluble fraction (R2); Lane 7. Pre-induction insoluble fraction (R2); Lane 8. Post-induction soluble fraction (R2); Lane 9. Post-induction insoluble fraction (R2).


227

Figure A.5.4. Western blot images of screening conditions. A. LB, OD600 0.6, xylose conc. 0.1%, overnight; B. LB, OD600 0.6, xylose conc. 0.5%, overnight; C. LB, OD600 0.3, xylose conc. 0.1%, overnight; D. TB, OD600 0.6, xylose conc. 0.5%, overnight; E. TB, OD600 0.3, xylose conc. 0.1%, overnight; F. TB, OD600. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. Pre-induction soluble fraction (R1); Lane 3. Pre-induction insoluble fraction (R1); Lane 4. Post-induction soluble fraction (R1); Lane 5. Post-induction insoluble fraction (R1); Lane 6. Pre-induction soluble fraction (R2); Lane 7. Pre-induction insoluble fraction (R2); Lane 8. Post-induction soluble fraction (R2); Lane 9. Post-induction insoluble fraction (R2).


228

Figure A.5.5. Western blot image of pre- and post-induction samples from 1 L culture of B. megaterium. Lane 1. SeeBlue Pre-Stained Protein Standard; Lane 2. TmrA in soluble cell fraction; Lane 3. TmrA in insoluble cell fraction.

229

A.6 LC-MS/MS Results to confirm identity of TmrA in excised gel bands

Mass spectrometric analysis for this work was carried out at the Bioanalytical Mass

Spectrometry Facility, UNSW and was supported in part by infrastructure funding from

the New South Wales Government as part of its coinvestment in the National

Collaborative Research Infrastructure Strategy”.

Digest peptides were separated by nano-LC using an Ultimate 3000 HPLC and

autosampler system (Dionex, Amsterdam, Netherlands). Samples (2.5 µl) were

concentrated and desalted onto a micro C18 precolumn (300 µm x 5 mm, Dionex) with

H2O:CH3CN (98:2, 0.05 % TFA) at 15 µl/min. After a 4 min wash the pre-column was

switched (Valco 10 port valve, Dionex) into line with a fritless nano column (75µ x

~10cm) containing C18 media (1.9 µ, 120Å, Dr Maisch, Ammerbuch-Entringen Germany)

manufactured according to Gatlin et al. (1998). Peptides were eluted using a linear

gradient of H2O:CH3CN (98:2, 0.1 % formic acid) to H2O:CH3CN (64:36, 0.1 % formic

acid) at 200 nl/min over 30 min. High voltage 2000 V) was applied to low volume tee

(Upchurch Scientific) and the column tip positioned ~ 0.5 cm from the heated capillary

(T=275°C) of an Orbitrap Velos (Thermo Electron, Bremen, Germany) mass

spectrometer. Positive ions were generated by electrospray and the Orbitrap operated

in data dependent acquisition mode (DDA).

A survey scan m/z 350-1750 was acquired in the Orbitrap (Resolution = 30,000 at m/z

400, with an accumulation target value of 1,000,000 ions) with lockmass enabled. Up to

the 10 most abundant ions (>4,000 counts) with charge states > +2 were sequentially

isolated and fragmented within the linear ion trap using collisionally induced

dissociation with an activation q = 0.25 and activation time of 30 ms at a target value of

30,000 ions. M/z ratios selected for MS/ MS were dynamically excluded for 30 seconds.

All MS/MS spectra were searched against NCBI database using MASCOT (version 2.3)

with the following search criteria: enzyme specificity was trypsin; precursor and product

ion tolerances were at 4 ppm and ± 0.4 Da, respectively; variable modification of

methionine oxidation; and one missed cleavage was allowed. The ions score significance

230

threshold was set to 0.5 and each protein was provided with a probability based Mowse

(Molecular Weight Search) score (Pappin et al., 1993). Mass spectrometric analysis was

carried out at the Bioanalytical Mass Spectrometry Facility, University of New South

Wales, Australia.

PAPPIN, D.J.C, HOJRUP, P., BLEASBY, A.J. 1993. Rapid identification of proteins by

peptide-mass fingerprinting. Current Biology, 3(6): 327-332.

GATLIN, C., KLEEMANN, G., HAYS, L., LINK, A., YATES, JR. 1998. Protein identification at

the low femtomole level from silver-stained gels using a new fritless electrospray

interface for liquid chromatrography-microspray and nanospray mass

spectrometry. Analytical Biochemistry, 263(1): 93–101.

231

Figure A.1. LC-MS/MS result for insoluble TmrA-Chis expressed in E. coli.

232

Figure A.2. LC-MS/MS result for insoluble TmrA-Chis expressed in E. coli (continued).

233

Figure A.3. LC-MS/MS result for insoluble TmrA-MBP expression in E. coli.

234

Figure A.4. LC-MS/MS result for insoluble TmrA-MBP expression in E. coli (continued).

235

Figure A.5. LC-MS/MS data for pASK_TmrA expressed in S. blattae (soluble cell fraction).

Figure A.6. LC-MS/MS data for pASK_TmrA expressed in S. blattae (insoluble cell

fraction).

236

Figure A.7 LC-/MS/MS data for pBBR_TmrA expressed in S. blattae (insoluble cell fraction).

production of functional recombinant chloroform reductive

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