Siderophore Production by Marine Alpha-Proteobacterium

UNIVERSITY OF CALIFORNIA

Santa Barbara

Siderophore production by marine α-proteobacterium Ochrobactrum sp. SP18

A Dissertation submitted in partial satisfaction of the

requirements for the degree Doctor of Philosophy

in Chemistry

Jessica Elanor Martin

Committee in charge:

Professor Alison Butler, Chair

Professor Peter Ford

Professor Richard Watts

Professor Stanley Parsons

June 2006 UMI Number: 3218830

UMI Microform 3218830 Copyright 2006 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.

ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, MI 48106-1346 The dissertation of Jessica Elanor Martin is approved.

______Alison Butler

______Peter Ford

______Richard Watts

______Stanley Parsons

May 2006

Siderophore production by marine α-proteobacterium Ochrobactrum sp. SP18

Jessica Elanor Martin

iii

ACKNOWLEDGMENTS

I would like to thank Prof. Alison Butler, Department of Chemistry and

Biochemistry, University of California, Santa Barbara for her assistance, patience, and support throughout my graduate education. Prof. Butler’s ideas and suggestions were the foundation for the work in this dissertation. I thank my committee,

Professors Peter Ford, Richard Watts, and Stanley Parsons for their time, commentary, and helpful advice.

Sincere thanks go to Dr. James Pavlovich for his invaluable assistance with mass spectrometry and for many interesting conversations. I would also like to thank the

Butler lab members, current and past, for their friendship, assistance, and support.

I would especially like to thank my husband Chris Martin for his patience, care, and support. Without his help, this work would not have been possible. I would also like to thank my parents, Charles and Marilyn Dryden, and my sister, Amy

Dryden, for their love and support throughout my education. It was because of their care and faith that I have been able to accomplish my goals.

VITA OF JESSICA ELANOR MARTIN May 2006

Education May 1998 M.S. in Chemistry Thesis: Relationships between nutrient concentrations and fecal coliform levels at Pine Flats in Oak , Arizona Thesis advisor: Professor Richard Foust Northern Arizona University, Flagstaff, Arizona

May 1996 B.S. in Chemistry Northern Arizona University, Flagstaff, Arizona

Research Experience 2000 – 2006 Graduate Research Assistant Advisor: Professor Alison Butler Topic: Marine bacterial siderophores University of California, Santa Barbara, California

2004 – 2005 California Sea Grant Graduate Trainee Advisor: Professor Alison Butler Topic: Marine red algal vanadium-dependent bromoperoxidases University of California, Santa Barbara, California

1996 – 1998 Graduate Research Assistant/Associate Advisor: Professor Richard Foust Topic: Oak Creek Water Quality Monitoring Project, EPA Section 319(h) National Monitoring Program Northern Arizona University, Flagstaff, Arizona

Publications Martin, J.D., Homann, V.V., Haygood, M.G., and Butler, A. Structure and membrane affinity of novel lipophilic siderophores produced by Ochrobactrum sp. SP18. Journal of Biological Inorganic Chemistry in press.

Butler, A. and Martin, J.D. The marine biogeochemistry of iron. In Metal Ions in Biological Systems (A. Sigel, H. Sigel, and R.K.O. Sigel, Eds.). 2005, Vol. 44, 21 – 46, Marcel Dekker, New York.

Martinez, J.S., Carter-Franklin, J.N., Mann, E.L., Martin, J.D., Haygood, M.G., and Butler, A. Structure and membrane affinity of a new suite of amhiphilic siderophores produced by a marine bacterium. Proceedings of the National Academy of Sciences USA 2003, 100, 3754 – 3759.

ABSTRACT

Siderophore production by marine α-proteobacterium Ochrobactrum sp. SP18

Jessica Elanor Martin

Iron is required for growth of nearly all microorganisms. While iron is the fourth most abundant element on the earth’s surface, iron is only sparingly soluble in the aerobic, near neutral conditions under which most microorganisms grow.

Therefore, microorganisms experience iron limitation in nearly every environment where they grow, from infection of a mammalian host (where iron is highly controlled by protein complexation) to aquatic and marine environments (where iron is relatively insoluble or is complexed by organic ligands). Bacteria, fungi, and other microbes have developed complex strategies to compete for iron under these conditions. Specifically, many microbes produce low molecular weight, iron binding compounds called siderophores to acquire iron from the environment.

Siderophores are secreted by microorganisms and then are taken back into the cells as the ferric complex to promote microbial growth.

The siderophores produced by marine α-proteobacterium Ochrobactrum sp.

SP18 were structurally characterized. This marine bacterium produces a suite of three aerobactin-derived, amphiphilic siderophores composed of citrate,

symmetrically derivatized with L-lysine which is N-hydroxylated and N-acylated to form two hydroxamate binding groups. Each siderophore contains one (E)-2- decenoic acid moiety and an (E)-2-octenoic, octanoic, or (E)-2-decenoic acid moiety. The photoreactivity and membrane affinity of these siderophores were investigated.

In addition to isolation of the cell-associated ochrobactin siderophores from the marine α-proteobacterium Ochrobactrum sp. SP18, these amphiphilic siderophores were isolated from the marine γ-proteobacterium Vibrio sp. DS40M5.

The isolation of these siderophores from two widely different strains, that are in distinct clades of bacteria suggests that these siderophores may be widely utilized in marine environments.

vii

TABLE OF CONTENTS

I. Introduction...... 1

A. Microbial iron acquisition...... 1

1. Ferric-ion-specific chelators: siderophores...... 1

2. Siderophore structures...... 2

3. Ferric ion affinity ...... 4

4. Outer membrane transport proteins...... 5

5. Siderophore transfer to the periplasm ...... 7

6. Iron transport across the cytoplasmic membrane...... 8

7. Outer membrane transport proteins (OMPs) versus OMPNs ...... 9

8. Apo- versus ferric-siderophore detection...... 11

B. Iron in the oceans...... 14

1. The iron hypothesis...... 14

2. Iron fertilization experiments...... 15

C. Iron speciation in the ocean...... 19

1. Iron binding ligands ...... 19

2. Photochemical iron cycling in marine waters ...... 20

3. Dissolution of iron minerals...... 23

D. Marine siderophores...... 24

1. Amphiphilic character of marine siderophores...... 30

2. Photoreactivity of marine siderophores ...... 33

E. Siderophores in natural seawater environments ...... 34

viii

F. Organization of this thesis ...... 37

G. References ...... 39

II. Isolation and structural characterization of siderophores produced by

Ochrobactrum sp. SP18 ...... 49

A. Introduction ...... 49

1. Marine α-proteobacteria...... 49

2. Citrate-derived siderophores ...... 51

B. Materials and methods...... 53

1. Maintenance of Ochrobactrum sp. SP18 ...... 53

2. Siderophore production...... 53

3. Biofilm formation...... 55

4. Siderophore isolation ...... 55

5. Structure determination...... 57

a. Amino acid analysis ...... 57

b. Fatty acid analysis...... 57

c. NMR...... 58

d. Mass spectrometry ...... 58

C. Results ...... 59

1. Ochrobactrum sp. SP18 siderophores...... 59

a. Biofilm determination ...... 59

b. Sodium chloride content of medium...... 59

c. Tandem mass spectrometry ...... 62

d. Mass spectrometry ...... 62

a. Marfey's amino acid analysis ...... 62

b. Fatty acid analysis...... 69

c. NMR...... 71

D. Discussion ...... 90

E. References...... 93

III. Siderophore production by Vibrio sp. DS40M5, Citrobacter koseri ATCC 27156,

and Klebsiella pneumoniae ATCC 25306...... 97

A. Introduction ...... 97

1. Structrual similarities between aerobactin and the ochrobactins...97

2. Diversity of aerobactin production ...... 97

3. Citrobacteria...... 100

4. Siderophore isolation: cell-associated versus secreted siderophores

...... 100

B. Materials and methods...... 103

1. Siderophore production by Vibrio sp. DS40M5 ...... 103

a. Growth and maintenance of Vibrio sp. DS40M5...... 103

b. Isolation of siderophores from Vibrio sp. DS40M5...... 103

c. Analysis of siderophores produced by Vibrio sp. DS40M5...

...... 105

2. Siderophore production by Citrobacter koseri ATCC 27156...... 106

a. Growth and maintenance of Citrobacter koseri ATCC

27156...... 106

b. Isolation of siderophores from Citrobacter koseri ATCC

27156...... 106

c. Analysis of siderophores produced by Citrobacter koseri

ATCC 27156 ...... 109

3. Siderophore production by Klebsiella pneumoniae ATCC 25306

...... 109

a. Growth and maintenance of Klebsiella pneumoniae ATCC

25306...... 109

b. Isolation of siderophores from Klebsiella pneumoniae

ATCC 25306 ...... 110

4. Biosynthesis of 15N aerobactin and feeding experiments ...... 110

a. Biosynthesis and purification of 15N labeled aerobactin..110

b. Aerobactin feeding experiment with Ochrobactrum sp.

SP18 ...... 111

c. Attempts to produce 15N labeled ochrobactins...... 112

C. Results ...... 114

1. Isolation of siderophores from Vibrio sp. DS40M5...... 114

2. Siderophore production by Citrobacter koseri ATCC 27156...... 126

3. Siderophore production by Klebsiella pneumoniae ATCC 23056

...... 133

4. Biosynthesis of 15N labeled aerobactin and feeding experiments 133

5. Attempts to produce 15N labeled ochrobactins ...... 144

F. Discussion...... 145

1. Ochrobactins: lipophilic aerobactin-derived marine siderophores

...... 145

2. Siderophore production by Citrobacter koseri ATCC 27156...... 148

G. References ...... 149

IV. Photochemical reactions of the ochrobactin siderophores...... 152

A. Introduction ...... 152

1. Photoreactivity of citrate siderophores (aerobactin, petrobactin) 152

B. Materials and methods...... 159

1. Production and isolation of ochrobactin siderophores...... 159

2. Extinction coefficient determination...... 160

3. Natural sunlight irradiation of Fe(III)-ochrobactin C ...... 161

4. Characterization of the photoreaction products ...... 162

a. Ferrous iron release ...... 162

b. Photoproduct ligand structure ...... 162

c. Carbon dioxide evolution ...... 162

5. Determination of quantum yield ...... 163

C. Results ...... 164

1. Isolation of ochrobactins...... 164

2. Extinction coefficient determination...... 170

3. Natural sunlight irradiation of Fe(III)-ochrobactin C ...... 173

xii

4. Characterization of the photoreaction products ...... 176

a. Ferrous iron release ...... 176

b. Photoproduct ligand structure ...... 176

c. Carbon dioxide evolution ...... 182

5. Quantum yield...... 186

F. Discussion...... 197

G. References ...... 200

V. Membrane partition coefficient determination with the ochrobactin siderophores

...... 202

A. Introduction ...... 202

1. Amphiphilicity of marine siderophores ...... 202

a. Siderophore structures...... 202

b. Cell-associated siderophores...... 202

c. Membrane partitioning studies...... 205

B. Materials and methods...... 208

1. Purification and preparation of siderophores ...... 208

2. Partition coefficient determination...... 211

C. Results ...... 213

1. Isolation and purification of ochrobactins and ochrobactin

photoproducts...... 213

2. Partition coefficient determination...... 213

F. Discussion...... 226

xiii

G. References ...... 229

VI. Concluding Remarks and Future Directions...... 231

A. The ochrobactins ...... 231

B. Biosynthesis of citrate-derived siderophores ...... 232

C. Putative outer membrane siderophore transport protein ...... 236

D. Photoreactivity of the ochrobactins...... 237

E. References...... 238

xiv

LIST OF FIGURES

Figure 1.1 Structures of certain siderophores ...... 3

Figure 1.2 Diagram of outer membrane transport proteins and their cognate

siderophores ...... 6

Figure 1.3 Diagram of the diferric-dicitrate outer membrane sensor and transport

system...... 10

Figure 1.4 Hydroxycarboxylic acids and known conditional stability constants with

iron(III)...... 22

Figure 1.5 Amphiphilic marine siderophores...... 26

Figure 1.6a Hydrophilic bacterial marine siderophores...... 27

Figure 1.6b Hydrophilic bacterial marine siderophores continued...... 28

Figure 1.7 Schematic diagram of competing reactions controlling iron(II)

concentrations ...... 29

Figure 1.8 Photoreaction of aquachelin D in natural sunlight ...... 35

Figure 1.9 Photoreaction of petrobactin in natural sunlight...... 36

Figure 2.1 Citrate-derived siderophores...... 50

Figure 2.2 RP-HPLC of cell-extract from Ochrobactrum sp. SP18 grown in normal

ASW-Fe (0.31 M sodium chloride) ...... 60

Figure 2.3 RP-HPLC of cell-extract from Ochrobactrum sp. SP18 grown in low

NaCl ASW-Fe (0.01 M sodium chloride) ...... 61

Figure 2.4 Tandem mass spectrum from ochrobactin A, m/z 757.5 ...... 64

Figure 2.5 Tandem mass spectrum of ochrobactin B, m/z 759.4...... 65

Figure 2.6 Tandem mass spectrum of ochrobactin C, m/z 785.43...... 66

Figure 2.7 Marfey’s amino acid analysis ...... 67

Figure 2.8 The three possible FDAA-L-lysine derivatives...... 68

Figure 2.9 GC-MS fragmentation patterns of methyl esters generated from the

ochrobactin siderophores ...... 70

Figure 2.10 1H NMR of ochrobactin C ...... 74

Figure 2.11 13C NMR of ochrobactin C...... 75

Figure 2.12 gCOSY of ochrobactin C...... 76

Figure 2.13 Expanded region of gCOSY of ochrobactin C ...... 77

Figure 2.14 gDQCOSY of ochrobactin C...... 78

Figure 2.15 Expanded region of gDQCOSY of ochrobactin C ...... 79

Figure 2.16 gHMQC of ochrobactin C ...... 80

Figure 2.17 Expanded region of gHMQC of ochrobactin C...... 81

Figure 2.18 CIGAR of ochrobactin C...... 82

Figure 2.19 APT of ochrobactin C...... 83

Figure 2.20 MSMS fragmentation of the ochrobactin siderophores...... 87

Figure 2.21 Rationalization of fragments in ochrobactin B tandem mass spectrum .88

Figure 2.22 Structures of the three siderophores produced by Ochrobactrum sp.

SP18 ...... 89

Figure 3.1 Structures of aerobactin and the ochrobactins...... 98

Figure 3.2 RP-HPLC purification of aerobactin from Vibrio sp. DS40M5...... 118

Figure 3.3 MSMS of aerobactin from Vibrio sp DS40M5, m/z 565...... 119

Figure 3.4 Fragmentation pattern of aerobactin...... 120

xvi

Figure 3.5 RP-HPLC of cell-associated siderophores from Vibrio sp. DS40M5 ....122

Figure 3.6 MSMS of m/z 759, Vibrio sp. DS40M5...... 123

Figure 3.7 MSMS of m/z 785, Vibrio sp. DS40M5...... 124

Figure 3.8 Fragmentation patterns for ochrobactins B and C ...... 125

Figure 3.9 RP-HPLC purification of aerobactin from C. koseri ATCC 27156 ...... 128

Figure 3.10 MSMS of aerobactin from C. koseri ATCC 27156, m/z 565...... 129

Figure 3.11 RP-HPLC of DFOG from C. koseri ATCC 27156...... 130

Figure 3.12 MSMS of m/z 619, C. koseri ATCC 27156 ...... 131

Figure 3.13 Rationalization of fragmentation pattern from desferrioxamine G ...... 132

Figure 3.14 RP-HPLC isolation of 15N labeled aerobactin...... 136

Figure 3.15 ESI-MS spectrum of 15N labeled aerobactin, m/z 569 ...... 137

Figure 3.16 MSMS of 15N labeled aerobactin, m/z 569...... 138

Figure 3.17 Fragmentation pattern for 15N labeled aerobactin ...... 139

Figure 3.18 RP-HPLC resolution of ochrobactins from 15N aerobactin supplemented

cultures of Ochrobactrum sp. SP18 ...... 141

Figure 3.19 ESI-MS spectrum of ochrobactin C...... 142

Figure 3.20 Expanded ESI-MS spectrum of ochrobactin C...... 143

Figure 3.21 Phylogenetic tree indicating characterized marine siderophores ...... 146

Figure 4.1 Photoreactive siderophores isolated from marine bacteria...... 153

Figure 4.2 Fe(III)-petrobactin photoreaction ...... 154

Figure 4.3 Photolysis of Fe(III)-aerobactin...... 156

Figure 4.4 Proposed coordination of Fe(III) by the petrobactin photoproduct...... 157

Figure 4.5 ESI-MS spectrum of methylated ochrobactin C...... 166

xvii

Figure 4.6 MSMS fragmentation of methylated ochrobactin C, m/z 799...... 167

Figure 4.7 Fragmentation pattern of methylated ochrobactin C ...... 168

Figure 4.8 UV-visible spectra of Fe(III) addition to apo-ochrobactin C ...... 171

Figure 4.9 Plot of titration of ochrobactin C solution with standard Fe(III)...... 172

Figure 4.10 Irradiation of Fe(III)-ochrobactin C in natural sunlight ...... 174

Figure 4.11 ESI-MS of Fe(III)-ochrobactin C during photolysis ...... 175

Figure 4.12 Fe(II) trapping with 1,10-phenanthroline ...... 178

Figure 4.13 MSMS of apo-ochrobactin C photoproduct, m/z 739 ...... 179

Figure 4.14 Rationalization of fragments from apo-ochrobactin C photoproduct...181

Figure 4.15 Carbon dioxide detection...... 184

Figure 4.16 Photoreaction scheme of Fe(III)-ochrobactin C ...... 185

Figure 4.17 RP-HPLC resolution of photoreaction...... 188

Figure 4.18 Plot of Ф determined from consumption of Fe(III)-ochrobactin C ...... 195

Figure 4.19 Plot of Ф determined from production of Fe(III)-ochrobactin C

photoproduct ...... 196

Figure 5.1 Amphiphilic marine siderophores...... 203

Figure 5.2 RP-HPLC resolution of apo-ochrobactin C...... 216

Figure 5.3 Partitioning traces for ochrobactin C...... 217

Figure 5.4 Partitioning traces for ochrobactin B...... 218

Figure 6.1 Structures of the ochrobactin siderophores ...... 231

Figure 6.2 Biosynthetic pathway for rhizobactin 1021...... 233

Figure 6.3 Biosynthetic pathway for aerobactin ...... 234

xviii

LIST OF TABLES

Table 1.1 Summary of membrane partitioning studies of siderophores into artificial

DMPC vesicles (200 nm)...... 31

Table 2.1 1H and 13C assignments for ochrobactin C...... 84

Table 2.2 Coupling and 2D NMR data for ochrobactin C ...... 85

Table 3.1 Aerobactin producing bacterial strains ...... 99

Table 3.2 Artificial seawater (ASW) media utilized for growth of 15N-labeled

ochrobactins ...... 113

Table 3.3 Growth media tested with each strain...... 114

Table 3.4 MSMS fragmentation of aerobactin...... 121

Table 3.5 MSMS fragmentation of 15N labeled aerobactin...... 140

Table 4.1 Fragmentation of methylated ochrobactin C, m/z 799...... 169

Table 4.2 Fragmentation of apo-ochrobactin C photoproduct, m/z 739...... 180

Table 4.3 Data from photoreaction, consumption of Fe(III)-ochrobactin C trial 1 .189

Table 4.4 Data from photoreaction, consumption of Fe(III)-ochrobactin C trial 2 .190

Table 4.5 Data from photoreaction, consumption of Fe(III)-ochrobactin C trial 3 .191

Table 4.6 Data from photoreaction, production of Fe(III)-ochrobactin C

photoproduct trial 1...... 192

Table 4.7 Data from photoreaction, production of Fe(III)-ochrobactin C

photoproduct trial 2...... 193

Table 4.8 Data from photoreaction, production of Fe(III)-ochrobactin C

photoproduct trial 3...... 194

xix

Table 5.1 Data from apo-ochrobactin C partitioning experiments ...... 219

Table 5.2 Data from Fe(III)-ochrobactin C partitioning experiments ...... 220

Table 5.3 Data from Fe(III)-ochrobactin C photoproduct partitioning experiments

...... 221

Table 5.4 Data from apo-ochrobactin B partitioning experiments ...... 222

Table 5.5 Data from Fe(III)-ochrobactin B partitioning experiments ...... 223

Table 5.6 Data from Fe(III)-ochrobactin B photoproduct partitioning experiments

...... 224

LIST OF ABBREVIATIONS

ABC: ATP Binding Cassette

APT: Attached proton test

ASW: Artificial seawater medium

ATCC: American Type Culture Collection

CAS: Chrome azurol sulfonate

CIGAR: Constant time inverse detected gradient accordion rescaled long range

CLE/ACSV: Competitive ligand equilibration/adsorptive cathodic stripping

voltammetry

CMC: Critical micelle concentration

DL: Concentration of siderophore in the lipid phase

DT: Total concentration of siderophore

DW: Concentration of siderophore remaining in solution

DFOG: Desferrioxamine G

DMPC: 1,2-dimyristoyl-sn-glycero-3-phosphocholine

DMSO: Dimethylsulfoxide

EDTA: Ethylenediaminetetraacetic acid

Eifex: European Iron Fertilization Experiment

ESI-MS: Electrospray ionization mass spectrometry

FDAA: (1-fluoro-2,4-dinitrophenyl)-5-L-alaninamide

FRET: Fluorescence resonance energy transfer gCOSY: Gradient selected correlation spectroscopy

xxi

gDQCOSY: Gradient selected double-quantum filtered correlation spectroscopy gHMQC: Gradient selected heteronuclear multiple quantum correlation

HCAs: Hydroxycarboxylic acids

HDPE: High-density polyethylene

HEPES: 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid

HNLC: High-nitrate-low-chlorophyll

GC-MS: Gas chromatography mass spectrometry

I0: Incident light intensity

ID: Internal diameter

IronEx: Iron Enrichment Experiment

L: Length

L1 and L2: Iron binding ligands

LB: Luria-Bertani

MSMS: Tandem mass spectrometry

NMR: Nuclear magnetic resonance

O: Ochrobactin

OMPs: Outer membrane transport proteins

OMPNs: Outer membrane signaling and transport proteins

PDB: Protein Data Bank phen: 1,10-phenanthroline

PP: Photoproduct

PVDF: Polyvinylidiene fluoride rRNA: Ribosomal ribonucleic acid

xxii

RP-HPLC: Reversed-phase high performance liquid chromatography

SDS-PAGE: Sodium-dodecyl-sulfate polyacrylamide gel electrophoresis

SEEDS: Subarctic Pacific Iron Experiment for Ecosystems Dynamics Study

SERIES: Subarctic Ecosystem Response to Iron Enrichment Study

SOFeX: Southern Ocean Iron Experiment

SOIREE: Southern Ocean Iron Release Experiment

Sid: Siderophore

TEAP: Triethylamine phosphate

TFA: Trifluoroacetic acid

Tris: Tris(hydroxymethyl)aminomethane

UVA: Ultraviolet A (315 – 400 nm)

UVB: Ultraviolet B (280 – 315 nm)

xxiii Chapter One

Introduction

Microbial Iron Acquisition

Ferric-Ion-Specific Chelators: Siderophores

Iron is required for growth of the vast majority of all bacteria (Sigel 1998,

Templeton 2002). While iron is the fourth most abundant element by weight in the

Earth’s surface, the insolubility of iron in the aerobic, near neutral pH conditions in

which most bacteria grow severely limits the availability of this essential nutrient

-39 (KSP Fe(OH)3 = 10 ). To overcome this apparent lack of iron, microorganisms have

evolved an elaborate mechanism to acquire iron. Under aerobic growth conditions, bacteria and other microorganisms produce siderophores, which are low molecular weight, ferric-ion-specific compounds for the solubilization and sequestration of iron(III). Siderophores are generally produced under conditions of low iron availability and secreted into the surrounding environment where they can complex ferric ion. Iron(III)-bound siderophores are returned to the cell through an active transport system, based on siderophore-specific outer membrane transport proteins.

The synthesis of both the siderophores and the outer membrane transport proteins is tightly regulated by iron concentrations within the cell. Once the iron concentration inside the cell is sufficient, the biosynthesis of siderophores and transport proteins is repressed.

1 Siderophore Structures

The structures of more than two hundred siderophores produced by terrestrial

and freshwater bacteria are known. Two common iron(III)-binding motifs found

among the known structures are catechols and hydroxamic acids but other iron-

binding functional groups are also present. Several species of Enterobacteriaceae

such as Escherichia coli produce enterobactin (Figure 1.1a), an example of a tris-

catecholate-containing siderophore (O’Brien and Gibson 1970). Desferrioxamine B

(Figure 1.1b), produced by Streptomyces species, is an example of a tris-hydroxamate siderophore (Bickel et al. 1960). While these two examples each only contain a single type of iron-binding group, many siderophores coordinate iron(III) with a

combination of ligand types.

Hexadentate siderophores such as enterobactin and desferrioxamine B generally bind iron with one-to-one stoichiometry. Enterobactin is the cyclic triester of 2,3-dihydroxy-N-benzoyl-S-serine and forms an octahedral complex with ferric ion

(Isied et al. 1976, McArdle et al. 1978). While many hexadentate siderophores are known, others are only tetradentate, such as rhodotorulic acid (Figure 1.1c) (Atkin and Neilands 1968), alcaligin (Figure 1.1d) (Nishio et al. 1988), bisucaberin (Figure

1.1e) (Takahashi et al. 1987), and putrebactin (Figure 1.1f) (Ledyard and Butler

1997). Tetradentate siderophores often form Fe2(siderophore)3 complexes to achieve

octahedral coordination of iron(III). The crystal structure of the Fe2(alcaligin)3

complex revealed that only one alcaligin acts to bridge the iron(III) ions as opposed to

a triple bridged helicate structure (Hou et al. 1998).

OH O N O OH a NH OH b CH3 O H N OH 2 O O O OH O NH O HN NH O O O O HO N O N N OH OH H O HO OH H HO N O c H d N O N OH N O O O N N O OH O H N O N OH H OH e O f H HO HO N O N N N H O O O H N N O OH N O O N H OH

Figure 1.1: Structures of certain siderophores. a) enterobactin (O’Brien and Gibson 1970), b) desferrioxamine B (Bickel et al. 1960), c) rhodotorulic acid (Atkin and Neilands 1968), d) alcaligin (Nishio et al. 1988), e) bisucaberin (Takahashi et al. 1987), and f) putrebactin (Ledyard and Butler 1997). Enterobactin and desferrioxamine B are hexadentate siderophores, the others are tetradentate structures.

3 Ferric Ion Affinity

Siderophores complex ferric ion with particularly high affinity constants. The stability constant is often reported as the complex formation constant, βFeLH between

3+ n- hydrated ferric ion, Feaq , and the fully deprotonated siderophore ligand (Sid ):

3+ n− (3-n) Feaq + Sid ' FeSid

[FeSID(3-n) ] β siderophore = 110 3+ n− [Feaq ][Sid ]

The value of the formation constants span a wide range, from 1022.5 for ferric aerobactin to 1049 for the ferric enterobactin complex. Determination of these proton- independent formation constants requires knowledge of the pKa value of each coordinating group.

Because little fully deprotonated siderophore or hexaquo iron(III) species would be present in solution at physiological pH, as well as to avoid problems associated with hydrolysis of ferric ion, the stability constant is often determined by competing the siderophore against another ligand with well defined thermodynamics of interaction with ferric ion, such as EDTA:

Fe(EDTA)1- + Sid n- ' FeSid (3-n) + EDTA 4-

4- 3+ [FeSid(3-n) ][EDTA 4- ] ⎛ [FeSid(3-n) ] ⎞⎛[EDTA ][Fe ]⎞ β siderophore K = = ⎜ ⎟⎜ aq ⎟ = 110 overall - n- ⎜ 3+ n- ⎟⎜ - ⎟ EDTA [FeEDTA ][Sid ] ⎝[Feaq ][Sid ]⎠⎝ [FeEDTA ] ⎠ β110

β values are comparable only for ligands of the same βFeLH, thus, β values are only comparable for ligands of the same denticity.

4 Outer Membrane Transport Proteins

Bacteria have evolved a class of high affinity outer membrane transport proteins which recognize specific iron(III)-siderophore complexes and are involved in energy-dependent active transport of the ferric siderophore complex across the outer membrane. Three transporters from E. coli whose crystal structures have been determined are FhuA (ferric hydroxamate uptake) (Ferguson et al. 2000) which recognizes ferric ferrichrome siderophores, FepA (ferric enterobactin permease)

(Buchanan et al. 1999) which recognizes ferric enterobactin, and FecA (ferric citrate protein) (Ferguson et al. 2002) which recognizes diferric-dicitrate (Figure 1.2).

Recently, the structures of FpvA the ferric pyoverdin outer membrane transport protein (Cobessi et al. 2005a) and FptA the ferric pyochelin outer membrane transport protein from Pseudomonas aeruginosa were determined (Cobessi et al.

2005b). The structural core for these transporters is a membrane-spanning, 22-strand, anti-parallel β-barrel similar in overall structure to other membrane porins, although each of the siderophore transporters has a segment of residues at the N-terminus which folds inside the β-barrel, effectively plugging the barrel from the periplasmic side of the outer membrane. When the ferric siderophore complex binds to the transporter, FhuA, FepA, FecA, and FpvA undergo conformational changes, causing a five amino acid region near the N-terminus (the TonB Box) to interact with the

TonB protein, initiating the transport and release of the ferric siderophore complex into the periplasmic space.

OH N H R O O H O R N O NH O H HO N N N O OH O H HN O HO III N O O NH H O OH + Fe HN HO HO N N N N O OH N O OH OHO O O H O O H HN N NH HO N N III OH O H O H + Fe O N O HO NH O O OH O III OH + Fe 2 O NH2 O O O HO III H + 2 Fe OH O N O HO O N N OH HO O HN S OH O OH S III OH +Fe

Outer FhuA FecA FpvA membrane FepA FptA

Cytoplasmic ExbD TonB membrane ExbB

FepA FhuA FecA FpvA FptA

Figure 1.2: Diagram of outer membrane transport proteins and their cognate siderophores. Shown along the bottom are the crystal structures for FepA (PDB ID 1FEP) (Berman et al. 2000, Buchanan et al. 1999), FhuA (PDB ID 1QFG) (Ferguson et al. 2000, Berman et al. 2000), FecA (PDB ID 1KMO) (Ferguson et al. 2002, Berman et al. 2000), FpvA (PDB ID 1XKH) (Cobessi et al. 2005a, Berman et al. 2000), and FptA (PDB ID 1XKW) (Cobessi et al. 2005b, Berman et al. 2000). Protein images drawn with MBT Protein Workshop 1.0 (Moreland et al. 2005).

6 Siderophore Transfer to the Periplasm

TonB is a cytoplasmic membrane protein which spans the periplasmic space binding the siderophore receptor proteins and also binding the cytoplasmic membrane proteins, ExbB and ExbD (Braun and Braun 2002). The TonB/ExbB/ExbD ternary complex mediates the signal transduction of the electrochemical potential of the cytoplasmic membrane to the outer membrane, allowing transport and release of the ferric siderophore complex. Once the ferric siderophore is released to the periplasmic space, it is bound by a high affinity periplasmic binding protein (e.g., FhuD Kd 0.1

µM for ferric ferrichrome siderophore complex), preventing the reverse transport of the iron-siderophore complex across the outer membrane (Braun and Braun 2002).

The binding and transport of a ferric siderophore complex is usually highly specific. The kinetics of ferric siderophore uptake show saturation behavior with an apparent Kd value in the range of less than 1 to 100 nM. For example FepA binds

Fe(III)-enterobactin with a Kd of less than 0.1 nM (Cao et al. 2000). Similarly, the apparent Kd value for Fe(III)-pyoverdin with FpvA was determined to be 0.37 nM, for Fe(III)-pyochelin with FptA (ferric pyochelin transport) was found to be 0.54 nM, and for FhuA with Fe(III)-ferrichrome was identified as 0.65 nM (Hoegy et al. 2005).

3- For Fe(III)-enterobactin, other complexes such as Rh(III)-(catecholate)3 (a kinetically inert complex) do not affect the rate of uptake of Fe(III)-enterobactin, however, Rh(III)-(N,N-dimethyl-2,3-dihydroxybenzamide)3 (a complex containing the catecholamide but not the trilactone moiety of enterobactin) does block uptake of

Fe(III)-enterobactin at about 100 µM (Ecker et al. 1988). Thus, while the trilactone

7 backbone of enterobactin is not required for recognition and uptake, the catecholamide moiety is an essential feature of enterobactin for iron uptake to occur.

Investigation of uptake of Fe(III)-enterobactin using a fluorescent FepA derivative revealed biphasic binding kinetics of the siderophore complex characterized by an initial fast step followed by a slower step (Payne et al. 1997). This biphasic binding was proposed to arise from an initial binding of the Fe(III)-enterobactin complex to a site on or near the outside of FepA followed by binding within the protein cavity

(Payne et al. 1997). Site directed mutagenesis of FepA supports this hypothesis, suggesting the presence of two distinct binding sites within the protein, one consisting primarily of hydrophobic bonds with residues in the extracellular loops, notably with

F329 and Y272, and a second site deeper within the protein with hydrophobic and electrostatic interactions combining to produce an extremely selective binding pocket

(Cao et al. 2000). Other outer membrane siderophore transport proteins have similar motifs of aromatic residues in the extracellular loops which may contribute to similar biphasic binding events.

Iron Transport across the Cytoplasmic Membrane

Several types of systems exist for the transport of iron across the cytoplasmic membrane. These include the translocation of ionic ferric ion, ferrous iron and the ferric siderophore complexes. Transport of the ferric-ion and ferric-siderophore complexes typically requires use of an ABC (ATP Binding Cassette) transporter system in an energy-dependent process. One example from the diferric-dicitrate

8 transporter system involves FecBCDE (Figure 1.3); analogous systems have been identified for other known siderophore transporters. Four proteins are involved in transport of iron from the periplasmic space across the cytoplasmic membrane: FecB

(a periplasmic iron-binding protein), FecC and FecD (cytoplasmic membrane proteins), and FecE (ATP-binding protein) (Braun and Mahren 2005). In the E. coli diferric-dicitrate transport pathway, iron is reduced and removed from the citrate within the periplasmic space and iron(II) is transported across the cytoplasmic membrane (Braun and Mahren 2005). Isotopic labeling studies demonstrate that citrate does not cross the cytoplasmic membrane (Hussein et al. 1981).

Outer Membrane Transport Proteins (OMPs) versus OMPNs

In addition to acting as a diferric-dicitrate transport protein, FecA has an additional function as an outer membrane diferric-dicitrate sensor (Figure 1.3).

Under iron limitation, derepression of fecA and fecIR occurs and small amounts of

FecA, FecI, and FecR are produced (Braun and Mahren 2005). When diferric- dicitrate binds in the extracellular cavity of FecA, a conformational change is induced such that the N-terminal region of FecA contacts FecR, which is a cytoplasmic membrane spanning protein. FecR, in turn, contacts the sigma factor FecI, which directs RNA polymerase to the promoter region for FecA, initiating production of the

FecABCDE proteins (Braun and Mahren 2005). Thus, detection of the diferric- dicitrate complex by FecA induces synthesis of the proteins required for transport of diferric-dicitrate into the periplasmic space and transport of iron(II) across the

Outer Membrane

FecB TonB Box N

Cytoplasmic Membrane FecD FecC ExbD ExbB TonB FecR FecE FecI Fe2+ RNAP

P P fecIR P P fecABCDE Fur fecI Fur fecA

Figure 1.3: Diagram of the diferric-dicitrate outer membrane sensor and transport system. FecA, FecI, and FecR are involved in sensing extracellular diferric-dicitrate. FecABCDE are involved in transport of diferric-dicitrate into the periplasmic space and transport of Fe(II) across the cytoplasmic space.

10 cytoplasmic membrane (Braun and Mahren 2005). Some outer membrane siderophore transport proteins, such as FhuA (Ferguson et al. 2000) and FptA

(Cobessi et al. 2005b), have a short N-terminal sequence containing the TonB Box sequence and sufficient linking amino acids to contact TonB. In the case of FecA

(Enz et al. 2000), FpvA (James et al. 2005), and PupB (pseudobactin uptake protein from Pseudomonas putida) (Koster et al. 1994), the N-terminal chain contains approximately 100 additional amino acids past the TonB Box which contain the signaling sequence for interaction with FecR. The outer membrane transport proteins

(OMPs) and the outer membrane signaling and transport proteins (OMPNs) are thus distinguishable by their N-terminal sequences.

Apo- versus Ferric-Siderophore Detection

Recent evidence suggests that under physiological conditions, the outer membrane transport proteins are pre-loaded with apo-siderophore. The first indication of this phenomenon was the copurification of apo-pyoverdin and FpvA from Pseudomonas aeruginosa (Schalk et al. 1999). Fluorescence and fluorescence resonance energy transfer (FRET) experiments indicated that in vivo FpvA proteins were normally loaded with apo-pyoverdin which was rapidly displaced in the presence of Fe(III)-pyoverdin (Schalk et al. 1999, Schalk et al. 2002). Release of apo-pyoverdin was found to be TonB dependent (Clément et al. 2004). The crystal structure of FpvA loaded with apo-pyoverdin was recently reported at 3.6 Å resolution and further elucidates the interaction (Cobessi et al. 2005a). Apo-

11 pyoverdin is bound in a receptor pocket near the protein surface which is lined with aromatic residues (Cobessi et al. 2005). The extracellular loops do not fully cover the apo-siderophore, leaving the apo-siderophore partially exposed to the solvent

(Cobessi et al. 2005). The chromophore portion of pyoverdin, however, is buried within the protein and is not solvent accessible (Cobessi et al. 2005). In the presence of TonB and Fe(III)-pyoverdin, the apo-siderophore may be expelled without significant conformational changes in exchange for Fe(III)-pyoverdin (Cobessi et al.

2005). Tantalizing new data even suggest that Fe(III) is removed from pyoverdin while bound within the FpvA outer membrane transport protein (Schons et al. 2005).

In 2003, Yue et al. demonstrated that the FecA outer membrane transport protein was also able to bind apo-citrate although with significantly lower affinity than for the diferric complex (Yue et al. 2003). The crystal structures of the unliganded, apo-citrate, and diferric-dicitrate FecA complexes were determined and suggested a sensing mechanism for discrimination between apo-citrate and diferric- dicitrate (Yue et al. 2003). The unliganded and apo-citrate loaded proteins have very similar conformations (Yue et al. 2003). Apo-citrate is accommodated within a binding pocket located near the protein surface, with two citrate ions arranged orthogonally to each other (Yue et al. 2003). The citrate ions accept 13 hydrogen bonds from R155 (in the plug domain), R365, R380, and R438 (each within the β- barrel), and through five bridging water molecules (Yue et al. 2003). Upon coordination of iron (or through displacement of apo-citrate by diferric-dicitrate), the conformation dramatically changes (Yue et al. 2003). Diferric-dicitrate resides in the

12 same binding pocket but the two citrate ions are now positioned in a parallel configuration with two ferric ions between them (Yue et al. 2003). Each iron is coordinated with three carbonyl and two hydroxyl bonds from citrate and one water molecule (Yue et al. 2003). Extracellular loops seven and eight fold in to form a lid, trapping diferric-dicitrate within the binding pocket (Yue et al. 2003). Residues S568,

Q570, and D573 form hydrogen bonds and van der Waals interactions with diferric- dicitrate (Yue et al. 2003). Importantly, residues Q178 (from loop eight) and N721 move to within hydrogen bonding distance of the diferric-dicitrate, indicating significant changes in the protein configuration (Yue et al. 2003). The plug domain shifts downward and the TonB-box domain becomes quite flexible and is no longer resolved in the crystal structure (Yue et al. 2003).

In vitro and in vivo experiments with FhuA and apo-ferrichrome and with

FptA and its cognate siderophore pyochelin demonstrated that both of these outer membrane transport proteins also bind the apo-siderophore with high affinity (Hoegy et al. 2005). Interestingly, the measured in vivo affinity is much higher than the in vitro affinity. It is proposed that the outer membrane transport proteins are normally loaded with apo-siderophore under physiological conditions (Hoegy et al. 2005).

Thus the transport of iron from the extracellular milieu into the cell depends on the secretion of ferric-ion chelators which are taken back into the cell through a highly specific system of proteins which recognize the ferric-siderophore complex.

13 Iron in the Oceans

One third to one half of the world’s fixation of carbon dioxide is estimated to occur in the oceans as a result of photosynthetic activity by phytoplankton (Field et al. 1998). Specifically, the majority of this marine carbon dioxide fixation has been shown to occur in coastal environments. In vast regions of the world’s oceans however, chlorophyll levels from photosynthetic microorganisms are unusually low, leading to low levels of primary production, despite the fact that these waters are replete in major nutrients like nitrate, phosphate, and silicate. These high-nitrate-low chlorophyll (HNLC) regions also coincide with very low iron levels, ranging from 20 pM to 1 nM in surface seawater (Morel et al. 2003, Morel and Price 2003, Moore et al. 2002, Johnson et al. 1997, Johnson et al. 1994, Martin et al. 1994, O’Sullivan et al. 1991, Martin 1990, Martin and Fitzwater 1988).

The Iron Hypothesis

The recognition that the HNLC regions correlated with low iron concentration led to the development of the “Iron Hypothesis” by John Martin (Martin 1990, Martin et al. 1991), which states that primary productivity in large areas of the world’s oceans is limited by low iron concentrations. The assertion is that photosynthetic microorganisms are unable to utilize the available nitrate, phosphate, and silicate due to a lack of iron, and that addition of iron to the HNLC regions would result in an increase in growth of photosynthetic microorganisms and primary productivity. This increase in growth could efficiently consume carbon dioxide from the atmosphere,

14 leading to removal of carbon dioxide to the deep oceans as the remains of the phytoplankton sink away from the ocean surface. Consequently, as suggested by the

Iron Hypothesis, atmospheric carbon dioxide levels could be significantly decreased if iron addition to a large area of the ocean could stimulate sufficient primary production and if this carbon could be exported to the deep oceans.

The region of high primary productivity to the west of the Galapagos Islands supported the Iron Hypothesis. The conjecture was that an influx of iron in the form of volcanic ash from the Galapagos Islands which was carried by the prevailing winds and currents was fertilizing these waters. The hypothesis that increasing iron concentrations could allow photosynthetic microorganisms to proliferate was tantalizing and quickly led to early in vitro supplementation studies.

Iron Fertilization Experiments

The Iron Hypothesis has now been tested at least eight times on large (~70-

100 km2) patches of ocean water in the equatorial Pacific, the eastern subarctic

Pacific, and the Southern Oceans. The first open-ocean iron enrichment experiment

(IronEx I) was completed in 1993 in the equatorial Pacific Ocean (Martin et al. 1994,

Boyd et al. 1999, Coale et al. 1998, Stanton et al. 1998, Gordon et al. 1998). In this experiment, a single 4 nM iron enrichment was made to a patch of open surface water, resulting in a three-fold increase in chlorophyll. Within four days, the added iron was no longer detectable. The magnitude and longevity of the biological and geochemical responses were much smaller than predicted, most likely due to the rapid

15 loss of iron from the system (Martin et al. 1994, Cullen 1995). A second series of experiments (IronEx II, 1996) tested the idea that iron loss decreased efficacy in

IronEx I and was completed in the equatorial Pacific Ocean, near the Galapagos

Islands (Wells 2003, Cavender-Bares et al. 2001, Landry et al. 2000, Bollens and

Landry 2000, Mann and Chisholm 2000, Erdner and Anderson 1999, Steinberg et al.

1998, Behrenfeld et al. 1996, Rue and Bruland 1997, Coale et al. 1996). Multiple additions of iron over several days stimulated massive phytoplankton blooms reaching a maximum two days after the last addition of iron, resulting in a 90 µatm drawdown of CO2 and a 5 µM drawdown of nitrate. The iron(III) present in surface ocean waters has been shown to be essentially fully complexed by an organic ligand or class of ligands, “L” (Rue and Bruland 1997, Debarr et al. 1995, Wu and Luther

1995, Rue and Bruland 1995, Gledhill and Vandenberg 1994). An intriguing result of IronEx II was that the concentration of the organic ligand, “L”, increased in a short time-span, meeting the concentration of added iron; thus it has been proposed that

“L” is biologically derived (Rue and Bruland 1997). The results from IronEx II unequivocally support the iron hypothesis for the limitation of primary productivity by iron concentration.

In addition to IronEx I and IronEx II, mesoscale additions of ferrous sulfate have been made in the subarctic Pacific Ocean, during the Subarctic Pacific Iron

Experiment for Ecosystems Dynamics Study (SEEDS, 2001) (Tsuda et al. 2003) and the Subarctic Ecosystem Response to Iron Enrichment Study (SERIES, 2002) (Boyd et al. 2004). SEEDS resulted in a 40-fold increase of chlorophyll and a 13 µM

16 drawdown of nitrate; SERIES resulted in a greater than ten-fold increase in chlorophyll and more than 5 µM drawdown of nitrate.

Three large-scale experiments in the Southern Ocean have been completed and a fourth is in progress. These include the Southern Ocean Iron Release

Experiment (SOIREE, 1999) (Boyd et al. 2000, Bakker et al. 2001, Bakker et al.

2005, Bowie et al. 2001, Boyd and Law 2001, Boyd and Abraham 2001, Croot et al.

2001, Frew et al. 2001, Gall et al. 2001a, Gall et al. 2001b, Hall and Safi 2001,

Karsh et al. 2003, Law and Ling 2001, Law et al. 2003, Maldonado et al. 2001,

Nodder et al. 2001, Nodder and Waite 2001, Trull et al. 2001, Trull and Armand

2001, Waite and Nodder 2001, Watson et al. 2000, Zeldis 2001), the Southern Ocean

Iron Experiment (SOFeX, 2002) (Buessler et al. 2004, Cassar et al. 2004, Coale et al.

2004), an experiment in the Atlantic sector of the Southern Ocean called EisenEx

(named for the German word for iron, Eisen; 2000) (Bakker et al. 2005, Gervais et al.

2002, Croot and Laan 2002), and the European Iron Fertilization Experiment (Eifex) which took place in early 2004; the results from Eifex have not yet been fully reported (Walter et al. 2005). The completed experiments have all demonstrated significant increases in primary productivity (e.g., 40-fold increase in chlorophyll) and biomass of phytoplankton in response to iron addition. Corresponding reductions in atmospheric carbon dioxide and nitrate concentrations also occurred. The bloom stimulated by SOIREE lasted for over 50 days. The unexpected persistence of this bloom led to the proposal that phytoplankton release compounds to keep iron in a useable form, although the nature of this compound or compounds is not yet known.

17 During SOIREE, the fugacity of CO2 in surface waters decreased by 3.8 µatm/day after four to five days to a total reduction of 35 µatm after 13 days (Bakker et al.

2005). Results during EisenEx were similar although storm conditions resulted in greater deep water mixing (Bakker et al. 2005). Total biological uptake of carbon across the patch was 1389 ton (SOIREE) and 1433 ton (EisenEx) (Bakker et at.

2005). It is likely that the amount of carbon removed was similar for the two experiments because the amount of iron added was similar, despite differences in weather conditions (Bakker et al. 2005). Results of SOFeX, which were recently reported (Coale et al. 2004), demonstrated that carbon could be exported to the deep oceans (below 100 m). However, instead of the predicted sequestration of 100,000 tons carbon per 1000 tons Fe added, only 1000 tons of carbon was exported (Coale et al. 2004). Thus the results of these studies support the Iron Hypothesis and also provide further evidence for the presence of biologically derived iron binding ligands which play an integral role in iron cycling in the oceans.

In addition to phytoplankton such as diatoms and cyanobacteria, heterotrophic bacteria constitute an important class of microorganisms in the ocean that are also limited by the low iron levels in HNLC regions. Heterotrophic bacteria comprise up to half the total particulate organic carbon in ocean waters (Pakulski et al. 1996) and in some regions, such as the subarctic Pacific, heterotrophic bacteria can even contain higher cellular concentrations of iron than phytoplankton (Tortell et al. 1996).

During iron supplementation-induced phytoplankton blooms, heterotrophic bacteria also increase in numbers. Thus heterotrophic bacteria compete successfully for iron

18 against phytoplankton and cyanobacteria.

Iron Speciation in the Ocean

Iron Binding Ligands

The speciation of iron in the oceans is of utmost importance for understanding the functionality and purpose of siderophores in the natural environment. The very low concentrations of “free” iron in seawater make accurate concentration determinations very difficult. A competitive ligand equilibration/adsorptive cathodic stripping voltammetry (CLE/ACSV) technique was developed to determine the speciation of iron in seawater (Rue and Bruland 1995). In the central North Pacific,

CLE/ACSV titration data revealed the existence of two classes of iron binding ligands. These ligands were designated as L1 and L2 based on conditional stability

cond 13 -1 cond 11 -1 constants: K L1/Fe' = 1.2 × 10 M versus K L2/Fe' = 3.0 × 10 M (Rue and Bruland

1995). The total concentration of iron-binding ligands was estimated to be approximately 2 nM with 0.44 nM identified as L1 while the remaining 1.5 nM was identified as L2; the concentration of iron binding ligands far exceeded the concentration of dissolved iron (Rue and Bruland 1995). Two classes of ligand, L1 and L2, were also detected in the equatorial Pacific Ocean and the conditional stability constants were very similar to those determined in the previous experiments.

cond Specifically, L1 was identified at a concentration of 310 ± 80 pM with K L1/Fe' = 4.7 ±

12 -1 cond 11 -1 (2.7) × 10 M and L2 at 190 ± 90 pM with K L2/Fe' = 6.5 ± (3.7) × 10 M (Rue and

Bruland 1997).

19 Iron-binding compounds from more than 200 L of natural seawater were concentrated using an adsorption resin, were assayed for hydroxamate and catecholate binding groups, and the conditional stability constants with Fe(III) were determined using CLE/ACSV (Macrellis et al. 2001). Each compound with iron- binding activity tested positive for hydroxamate or catecholate moieties using qualitative colorimetric assays (Macrellis et al. 2001). The conditional stability

cond 11.5 11.9 -1 constants determined for the iron-binding compounds, K FeL,Fe' = 10 – 10 M , were very similar to the conditional stability constants of isolated siderophores and of

L1 and L2 (Macrellis et al. 2001).

Photochemical Iron Cycling in Marine Waters

The presence of oxalic, citric, malic, glyceric, salicylic, tartaric, glucaric, gluconic, or p-hydroxybenzoic acids in solution all have been shown to increase photoproduction of iron(II) from iron(III) (Figure 1.4) (Cunningham et al. 1988,

Kuma et al. 1992). Such organic complexes and, possibly, siderophores containing similar ferric-ion binding moieties, potentially undergo photooxidative reactions resulting in release of iron(II). Iron(II) is widely accepted as a bioavailable iron species.

Iron concentrations in oceanic surface waters have significant diurnal patterns with increasing concentrations of iron(II) during daylight hours (Johnson et al. 1994,

Rijkenberg et al. 2005). Generally, the highest total dissolved iron concentrations occur at midday (Waite and Nodder 2001). The impact of ultraviolet irradiation on

20 photoreduction of iron(III) in the Southern Ocean was assessed during the EisenEx iron fertilization experiment (Rijkenberg et al. 2005). Ultraviolet B wavelengths

(UVB; 280 – 315 nm) produced the highest concentration of iron(II) followed by ultraviolet A (UVA; 315 – 400 nm) and visible light (400 – 700 nm) during shipboard irradiation experiments (Rijkenberg et al. 2005). As in the equatorial Pacific, a distinct diurnal iron cycle was evident as well (Rijkenberg et al. 2005). It is evident that the photoreductive cycling of iron is important for the biogeochemistry of dissolved iron in the surface waters of the oceans.

Laboratory investigations have helped to clarify the photoreductive processes of iron in the ocean. It has been noted that more than 99.9% of dissolved iron in the surface waters of the oceans is complexed by organic ligands (Debarr et al. 1995,

Rue and Bruland 1995, Wu and Luther 1995, Rue and Bruland 1997). Hydroxamate and catecholate binding groups have been detected in the compounds identified as L1 and L2 (Macrellis et al. 2001). Hydroxycarboxylic acids (HCAs; Figure 1.2) are present in dissolved organic matter in the oceans (Creach 1955, Degens et al. 1964,

Handa and Tominaga 1969) and are also common functional groups in marine siderophores (as α-hydroxy carboxylic acids) such as the marinobactins, aquachelins, and alterobactins (Martinez et al. 2000, Reid et al. 1993).

In the dark at pH 8, HCAs such as glucaric acid-1,4-lactone, glucaric, tartaric, gluconic, citric, glyceric, malic, and glucuronic acids increase dissolved iron concentrations by stabilizing ferric species and therefore decreasing hydrolytic precipitation of iron(III) (Kuma et al. 1995). With ultraviolet or natural sunlight

HO OH OH OH O O OH OH OH O HO O HO HO OH HO OH O O O OH OH OH OH OH OH O OH glucaric acid- glucaric acid tartaric acid gluconic acid 1,4-lactone KFeL = 6.49

O O OH O OH O OH O OH HO OH OH OH HO HO O OH HO O O HO OH OH citric acid glyceric acid malic acid glucuronic acid KFeL = 11.5 KFeL = 7.1

O O O OH HO OH OH O OH HO oxalic acid salicylic acid p-hydroxybenzoic acid

Figure 1.4: Hydroxycarboxylic acids and known conditional stability constants with iron(III) (Kuma et al. 1995).

22 irradiation in the presence of HCAs (50 µM), photoreduction rates of iron(III) (5 µM) are significantly increased; rates increased linearly with sunlight intensity supporting the conclusion that this is genuinely a photochemical reaction and not a purely chemical process (Cunningham et al. 1988, Kuma et al. 1992, Kuma et al. 1995).

HCAs also increased rates of photoreduction of solid ferric HCA complexes, but rates depend on surface concentrations of HCAs (Kuma et al. 1995). In the presence of

HCAs, fast reduction of iron(III) relative to rates of oxidation of iron(II) results in significant concentrations of iron(II) in surface waters exposed to sunlight (Kuma et al. 1995). In the Southern Ocean, simultaneous addition of iron(III) and glucaric acid to surface ocean water produced higher levels of iron(II) than addition of iron(III) without an organic acid (Öztürk et al. 2004).

Dissolution of Iron Minerals

In organic-ligand free waters (or artificial seawater), irradiation with natural sunlight or ultraviolet light has been shown to initiate reduction of iron containing mineral surfaces, resulting in release of small amounts of iron(II). These reactions have been predicted to increase both the concentration of iron(II) and the dissolution of iron-containing minerals such as goethite. In the equatorial Pacific, Johnson et al. proposed a model for iron cycling in the surface oceans involving photoreductive dissolution of colloidal iron as iron(II), oxidation, and biological uptake of iron(III)

(Johnson et al. 1994).

In the dark, siderophores accelerate the rate of dissolution of iron(III)

23 containing minerals such as goethite, lepidocrocite, and amorphous iron oxides

(Hersman et al. 1995, Yoshida et al. 2002, Cheah et al. 2003, Kraemer 2004).

Mineral dissolution in the dark is a ligand-controlled mechanism, dependent on the surface concentration of siderophore or other ligand. Light-induced dissolution of iron(III) containing minerals such as lepidocrocite or goethite in the presence of iron- binding compounds increases the release of iron during irradiation (Borer et al. 2005,

Kraemer et al. 2005). Surprisingly, both photoreactive siderophores (such as aerobactin) and photostable siderophores (such as desferrioxamine B) increase the amount of iron released during irradiation (Borer et al. 2005). It has been proposed that in this light-initiated reaction, surface sites of iron(III) are reduced and released from the mineral surface. Siderophores then act to shuttle iron(II) away from the surface, preventing reoxidation and precipitation on the mineral surface (Borer et al.

2005). Once away from the mineral surface, reoxidation to the ferric species may occur, and formation of the ferri-siderophore complex is likely to increase the overall solubility of iron (and thus the total concentration of dissolved iron).

Marine Siderophores

While the study of siderophores produced by bacteria of marine origin is relatively new, a modest number of structures from many marine bacteria have been elucidated (Figures 1.5 and 1.6). Two interesting themes have become evident from the known siderophore structures from marine bacteria in contrast to known terrestrial bacteria: (1) the production of suites of amphiphilic siderophores composed of a

24 peptidic headgroup and one of a series of fatty acid appendages (Martinez et al. 2000,

Martinez et al. 2003), and (2) the presence of photolabile moieties that are photoreactive when bound to Fe(III), producing an oxidized siderophore ligand and

Fe(II) under irradiation with natural sunlight or ultra-violet light (Barbeau et al. 2001,

Barbeau et al. 2002).

Suites of marine amphiphilic siderophores include the aquachelins produced by Halomonas aquamarina DS40M3 (Holt 1998, Martinez et al. 2000), the marinobactins produced by Marinobacter sp. DS40M6 and Marinobacter sp.

DS40M8 (Martinez et al. 2000), the amphibactins produced by Vibrio sp. R-10

(Martinez et al. 2003), and the synechobactins produced by Synechococcus sp.

PCC7002 (Figure 1.5) (Ito and Butler 2005). H. aquamarina DS40M3,

Marinobacter sp. DS40M6, and Marinobacter sp. DS40M8 are open ocean isolates, collected in the same water sample at a depth of 40 m over the continental slope in the eastern equatorial Atlantic (Haygood et al. 1993, Martinez et al. 2000). Vibrio sp.

R-10 is a coastal isolate, collected near Roatan, Honduras (Martinez et al. 2003).

Synechococcus sp. PCC7002 is a coastal marine cyanobacterium; the synechobactins are the only structurally characterized marine cyanobacterial siderophores (Ito and

Butler 2005).

In the ocean, siderophores participate in iron cycling and increase the total concentration of dissolved iron available to microorganisms. Competing processes controlling the concentrations of dissolved iron in oceanic surface waters are shown in Figure 1.7. Iron(II) oxidation in the surface ocean is rapid. The process is pH

a O O c O O O H N N N N N HO HO HO HO HO H O H O H O H O R N N N OH R N N OH H N N N N N O H O H O H O H O H O OH OH OH OH N O R = O R = O E O I D O O 2 H OH O D1 O G C OH O O F O B O E A O D b O O OH O OH N N C HO HO HO OH OH O O B H O H O H O N N N OH R N N N N O H O H O H O H O OH O C OH d H2NO O B O O R = N N A D H O OH OH C O OH OH B H O O N N O A O

Figure 1.5: Amphiphilic marine siderophores. A) marinobactins produced by Marinobacter sp. DS40M6 and DS40M8 (Martinez et al. 2000), B) aquachelins produced by Halomonas aquamarina DS40M3 (Martinez et al. 2000), C) amphibactins produced by Vibrio sp. R-10 (Martinez et al. 2003), and D) synechobactins produced by the cyanobacterium Synechococcus sp. PCC7002 (Ito and Butler 2005).

O OH O O a O b H N N HN N NH2 H OH HO OH NH OH O OH O N COOH OH H O NH2 O NH H N N HN OH O HO NH O O OH O H HN O O N O O c H HN NH2 HO COOH N OH HN O OH OH OH O H O H O H N N N COOH N N N NH2 H O H O H O HO COOH HO COOH O d H2NOC N HO3S H HN O H OH O H O H O N N N COOH HO N N N OH O NH2 H O H O H OH A: R=CH NH HO COOH 2 2 R B: R=NHC(NH)NH2 OH O HO N H H e OH N N O HO N O H O HO H2N f HO O NH R O O H HO N O N N O N H H O N N OH HO OH H O R = H or SO3

Figure 1.6a: Hydrophilic bacterial marine siderophores. a) aerobactin produced by Vibrio sp. DS40M5 (Haygood et al. 1993), b) and c) alterobactins produced by Pseudoalteromonas luteoviolacea (Reid et al. 1993), d) pseudoalterobactins produced by Pseudoalteromonas sp. KP20-4 (Kanoh et al. 2003), e) desferrioxamine G produced by Vibrio sp. BLI-41 (Martinez et al. 2001), and f) petrobactin and petrobactin sulfonate produced by Marinobacter hydrocarbonoclasticus (Barbeau et al. 2002, Bergeron et al. 2003, Hickford et al. 2004).

a H b HO N O HO N N NH O O O O O O N O N OH HN N H OH c d H OH O H OH O O N N O N N N N O H O H O H O OH N N O N N O N N OH H O OH H O

e H HO O N COOH O N OH O O COOH OH O

Figure 1.6b: Hydrophilic bacterial marine siderophores continued. a) putrebactin produced by Shewanella oneidensis (Ledyard and Butler 1997), b) bisucaberin produced by Vibrio salmonicida (Winkelmann et al. 2002), c) dehydronocardamine produced by Streptomyces sp. (Lee et al. 2005), d) desmethylenylnocardamine produced by Streptomyces sp. (Lee et al. 2005), e) vibrioferrin produced by Vibrio parahaemolyticus (Yamamoto et al. 1994).

atmospheric dust upwelling - sid reduction biological hν biological FeIII-siderophore FeIII FeII uptake (aq) DOC (aq) uptake

+ sid -nH+ oxidation red III Fe (OH)n (Fe minerals)

hν

Figure 1.7: Schematic diagram of competing reactions controlling iron(II) concentrations. It is thought that FeII is a bioavailable species, available for direct uptake as the “free” ion.

29 dependent and is likely the result of oxidation by oxygen, hydrogen peroxide, organic peroxides, superoxide, or other reactive oxygen species (Miller et al. 1995).

Amphiphilic Character of Marine Siderophores

Compared to known terrestrial siderophores, a very high percentage of marine siderophores are amphiphilic and lipophilic. The high amphiphilicity or lipophilicity of these molecules suggest significant surface reactivity in aqueous solution. The apo-marinobactins have low critical micelle concentrations (CMCs) ranging from 25

µM for marinobactin E (ME) to 150 µM for MA indicating significant amphiphilic interactions in aqueous solution (Martinez et al. 2000). The ferri-marinobactins have slightly higher CMCs than their apo counterparts (Martinez et al. 2000).

Despite their amphiphilic characteristics, siderophores such as the marinobactins and aquachelins are generally secreted into the culture medium by the bacteria. To assess the potential interactions of the amphiphilic siderophores with bacterial cells, the membrane partition coefficients for the amphibactins (Martinez et al. 2003) and the marinobactins (Martinez et al. 2000, Xu et al. 2002) have been determined with artificial 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) vesicles (Table 1.1). Briefly, solutions of physiological mixtures of amphibactins or marinobactins or purified individual siderophores were incubated with varying concentrations of DMPC ranging from 0 to 30 mM. The vesicles were removed by centrifugation and the siderophore remaining in the supernatant was determined by

RP-HPLC. Partition coefficients were calculated from a plot of the concentration of

Siderophore Acyl Appendage Partition Coefficient, M-1 Apo FeIII AG C18:1, 3-OH 833 ± 108 AI C18:1 1018 ± 194 AB C14:0, 3-OH 1338 ± 161 AD C14:0 1915 ± 140 AH C16:0 3784 ± 225 MA C12:0 36 ± 7 MB C14:1 25 ± 4 MC C14:0 195 ± 28 MD (mixture) C16:1 209 ± 28 ME C16:0 5818 ± 694 174 ± 14

Table 1.1: Summary of membrane partitioning studies of siderophores into artificial DMPC vesicles (200 nm). A denotes Amphibactin (Martinez et al. 2003), M denotes Marinobactin (Martinez et al. 2000). Adapted from Martinez et al. 2003.

31 DMPC versus the ratio of siderophore remaining in solution to the total siderophore

(see Chapter 5).

Partition coefficients measured for the individual siderophores agreed well with partition coefficients measured for each siderophore as a part of the physiological mixture. In general, the longer acyl appendages produced the largest partition coefficients. Reduction of the marinobactin acyl appendage by two methylene carbons (such as from ME to MC) or the introduction of a double bond

(such as from ME to MD) results in a decrease in partition coefficient of approximately one order of magnitude (Xu et al. 2002). Membrane partition coefficients for incubation of the marinobactins with vesicles formed from E. coli lipopolysaccharide demonstrated very similar results (Xu et al. 2002). The amphibactins produced a similar trend, although the differences in partition coefficient were less (Martinez et al. 2003). The disparity between the partition coefficients of the marinobactins and the amphibactins may derive from the significant differences in the peptidic headgroups. The marinobactin headgroup is composed of six amino acid residues while the amphibactin headgroup is composed of only four amino acid residues. The smaller headgroup of the amphibactins may interfere less with the vesicle or cell surface, therefore contributing less to the overall partition coefficient measurement than the larger headgroup of the marinobactins

(Martinez et al. 2003).

It has been proposed that the amphiphilic or lipophilic character of known marine siderophores could contribute to the formation of a concentration gradient

32 radiating away from bacterial cells in the open ocean, prohibiting diffusion of the siderophores away from the cells, potentially increasing iron availability for these cells (Martinez et al. 2000). It is also conceivable that the amphiphilic character could lead to useful interactions between these siderophores and iron-containing particles or extracellular material such as biofilms or marine snow, thus contributing to increased bioavailability of iron.

Photoreactivity of Marine Siderophores

The prevalence of photoreactive moieties in marine siderophores is evident in a wide variety of known structures including the citrate-derived siderophores aerobactin (Vibrio sp. DS40M5) (Haygood et al. 1993) and petrobactin

(Marinobacter hydrocarbonoclasticus) (Barbeau et al. 2002, Bergeron et al. 2003,

Hickford et al. 2004), the amphiphilic aquachelins (Halomonas aquamarina

DS40M3) (Martinez et al. 2000) and marinobactins (Marinobacter sp. DS40M6 and

DS40M8) (Martinez et al. 2000), and the hydrophilic alterobactins

(Pseudoalteromonas luteoviolacea) (Reid et al. 1993). The photoreactions of ferri- siderophores in the upper ocean could contribute to the bioavailability of iron

(Barbeau et al. 2001, Barbeau et al. 2002). The photochemical reaction of ferri- siderophores generates an oxidized ligand and Fe(II), which could be directly taken up by microorganisms or, in aerobic environments, could be oxidized to Fe(III) and rechelated by another siderophore or by the photoproduct itself (Barbeau et al. 2001,

Barbeau et al. 2002). Thus, the cycling of iron in the upper ocean could be

33 moderated by the photolysis of photoreactive siderophore ligands.

Beta-hydroxyaspartate containing siderophores such as the aquachelins, marinobactins, and alterobactins are photoreactive when coordinated to Fe(III).

Natural sunlight is sufficient to induce photolysis of the ferri-aquachelins, producing an oxidized, truncated siderophore photoproduct, a lipidic fragment, and Fe(II)

(Figure 1.8) (Barbeau et al. 2001). The aquachelin photoproduct retains the ability to coordinate Fe(III) through the two remaining hydroxamate binding groups although

cond 12.2 the conditional stability constant is significantly reduced ( K FeL,Fe' = 10 for aquachelin B versus 1011.5 for the photoproduct) (Barbeau et al. 2001). It is unknown whether the photoproduct is available for direct uptake into the bacterium.

Citrate-derived siderophores such as the petrobactins are also photoreactive when coordinated to Fe(III) (Figure 1.9) (Barbeau et al. 2002). Irradiation with natural sunlight or ultraviolet light initiates a photoreaction resulting in an oxidized siderophore photoproduct and Fe(II). As with the aquachelin photoproduct, the petrobactin photoproduct retains the ability to coordinate Fe(III) through the two catechol moieties and possibly through the truncated citrate backbone.

Siderophores in Natural Seawater Environments

Although siderophores have been isolated and structurally characterized from pure cultures of marine bacteria and the ligands L1 and L2 have been detected in seawater, the presence of siderophores in natural seawaters had not been directly detected until recently. Development of HPLC-ESI-MS techniques has allowed the

FeIII

O O O N N O O O O OH O H O H O H O N N N N N N N OH H O H O H O H O OH OH H2NO

hν Fe(II)

O O N N HO HO OH H O H O H O H N N N N N N OH O H O H O H O OH OH H2NO

Figure 1.8: Photoreaction of aquachelin D in natural sunlight. Irradiation of the ferri- siderophore produces a truncated siderophore photoproduct and Fe(II) (Barbeau et al. 2001).

O O HN HN HN HN

HN O O HN HO OH O O O hν O FeIII OH O O O HN O O Fe(II) HN HO OH

HN HN HN HN O O

Figure 1.9: Photoreaction of petrobactin in natural sunlight (Barbeau et al. 2002).

36 isolation and tentative structural characterization of siderophores from enriched seawater incubations containing natural assemblages of heterotrophic bacteria

(McCormack et al. 2003). In one such experiment, seawater collected from

Wembury Bay in the English Channel was supplemented with glucose, ammonia, and phosphate and incubated; typical coastal iron concentrations for this area are approximately 2 to 3 nM (Gledhill et al. 2004). These natural bacterial populations produced detectable levels of siderophores in culture (Gledhill et al. 2004). Several amphibactins (D, E and two previously uncharacterized amphibactins) and desferrioxamines B and G (and an uncharacterized desferrioxamine) were detected from enriched seawater collected from Wembury Bay in the English Channel

(Gledhill et al. 2004).

Organization of this Thesis

The goal of this project was to investigate the nature of siderophores produced by marine α-proteobacterial species. The siderophores produced by a coastal marine

α-Proteobacterium, Ochrobactrum sp. SP18, were extracted from the cell membranes. The structures of these novel siderophores were determined using a combination of chiral amino acid analysis, mass spectrometry, tandem mass spectrometry, fatty acid analysis, and both one and two dimensional NMR. These siderophores, named the ochrobactins, were found to consist of citrate, symmetrically derivatized with L-lysine at each end, which is then Nε-hydroxylated and Nε-acylated

37 to form two hydroxamates per molecule. The ochrobactins coordinate iron through the two hydroxamates and also through the central citrate moiety. The ochrobactins are the first example of an acylated aerobactin structure and are also the first membrane associated citrate-derived siderophores. The photoreactivity and lipid partition properties were investigated.

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48 Chapter Two

Isolation and Structural Characterization of Siderophores Produced by Ochrobactrum sp. SP18

Introduction

Marine α-Proteobacteria

Standard culture-dependent techniques have been utilized for decades for the isolation of bacteria from many environments. The vast majority of bacteria isolated in this way from marine environments are gram-negative chemoorganotrophs, principally γ-proteobacteria (Munn 2004). Recent advances in community analysis techniques have significantly changed our understanding of marine microbial ecology. Culture-independent techniques for analysis of the diversity of microbial communities, specifically analysis of 16S rRNA sequences from environmental samples, have identified numerous previously unculturable species of bacteria

(Schmidt et al. 1991, Giovannoni and Rappe 2000, Rappe et al. 2000, Munn 2004,

Buchan et al. 2005, Giovannoni and Stingl 2005). In fact, α-proteobacteria represent a significant portion of bacterial species present in the oceans and, in contrast to culture-based studies, culture-independent studies have revealed that α- proteobacteria constitute a much larger fraction of microbial species in marine environments than γ-proteobacteria (Giovannoni and Rappe 2000, Munn 2004).

To date, siderophores isolated from marine bacteria have all been isolated from species of γ-proteobacteria. Alpha-proteobacteria have historically been difficult to cultivate and no siderophores have previously been isolated from marine

49 O O O H HO O N N N O N OH H O OH OH OH O OH D Schizokinen O O OH H OH Vibrioferrin O OH N N H O O H OH N N OH OH O O A OH Aerobactin O O OH H O OH O N N O HO O N N O N O H O O OH OH OH O Acinetoferrin OH O OR E OH Achromobactin O OH I OH O H O O N N N N H O H OH O N N OH O OH Nannochelins O HO B OH R = H or CH O O 3 O HO O H O OH N N O N N H O O OH OH N OH O Rhizobactin 1021 H NH2 O OR F OH OH OH OH Staphyloferrin B H H H O J OH O N N N N N OH O O H O O O H O O N OH O O N OH C Petrobactins H O OH R = H or SO3 O O R O H O O R O H N OH N N N N N H N O OH OH H HO H H OH OH G OH Synechobactins K,L HO OH OH H O O N N O HO O OH O O O Rhizoferrin: R=H Staphyloferrin A: R=CO2H

Figure 2.1: Citrate-derived siderophores. (A) Aerobactin, produced by Klebsiella pneumoniae (Aerobacter aerogenes 62-1) (Gibson and Magrath 1969) and others; (B) Nannochelins, produced by Nannocystis exedens (Kunze et al. 1992); (C) Petrobactins, produced by Marinobacter hydrocarbonoclasticus (Barbeau et al. 2002, Bergeron et al. 2003, Hickford et al. 2004); (D) Schizokinen, produced by Bacillus megaterium (Byers et al. 1967), Anabaena (Simpson and Neilands 1976), and others; (E) Acinetoferrin, produced by Acinetobacter haemolyticus (Okujo et al. 1994); (F) Rhizobactin 1021, produced by Rhizobium meliloti 1021 (Persmark et al. 1993); (G) Synechobactins, produced by Synechococcus PCC7002 (Ito and Butler 2005); (H) Vibrioferrin, produced by Vibrio parahaemolyticus (Yamamoto et al. 1994); (I) Achromobactin, produced by Erwinia chrysanthemi (Munzinger et al. 2000); (J) Staphyloferrin B, produced by Staphylococcus hyicus (Haag et al. 1994); (K) Rhizoferrin, produced by Rhizopus microsporus var. rhizopodiformis (Drechsel et al. 1991); and (L) Staphyloferrin A, produced by Staphylococcus hyicus DSM 20459 (Meiwes et al. 1990).

50 α-proteobacterial species. One terrestrial α-proteobacterial siderophore has been

characterized: rhizobactin 1021 (Persmark et al. 1993). The structure of rhizobactin

1021 is similar to that of schizokinen, with a central citrate moiety, modified at each

end with diamino propane which is N-hydroxylated and acetylated to form two

hydroxamate binding groups (Mullis et al. 1971). In the case of rhizobactin 1021,

one acetyl group is replaced with an (E)-2-decenoic acid moiety to form one of the

few known amphiphilic terrestrial siderophores (Mullis et al. 1971).

Citrate-Derived Siderophores

Citrate is a known microbial siderophore (Guerinot et al. 1990, Lesueur et al.

1993). Citrate is secreted by bacteria such as Bradyrhizobium japonicum under low-

iron conditions or is pirated from other sources (such as citrate secreted by or

contained in plants) and is then taken up as the diferric-dicitrate complex through the

FecA outer membrane siderophore transport protein or similar proteins (see summary

in Chapter 1) (Wagegg and Braun 1981, Yue et al. 2003). Citrate-derived

siderophores have been isolated from a variety of bacteria including Gram-negative γ- proteobacteria, Gram-positive bacteria, and even cyanobacteria (Figure 2.1). Three principal classes of citrate-derived siderophores have been isolated: (1) citrate modified at each end with diamino alkane subunits, (2) citrate modified at each end with lysine residues, and (3) asymmetric structures composed of one or more citrate moieties. Citrate-derived siderophores are a varied and interesting class of siderophores isolated from a variety of bacterial species.

Described in this chapter are the isolation and structural characterization of a

new suite of cell-associated, amphiphilic, photoreactive siderophores from a near- shore marine α-proteobacterium, Ochrobactrum sp. SP18.

52 Materials and Methods

Maintenance of Ochrobactrum sp. SP18

Ochrobactrum sp. SP18 was isolated from a water sample collected from the

pier at Scripps Institution of Oceanography, La Jolla, California during a

phytoplankton bloom by Prof. Margo Haygood. Ochrobactrum sp. SP18 was

maintained on low-iron natural seawater agar plates containing 0.5 g yeast extract

(Difco), 15 g BactoAgar (Difco), and 5 g BactoPeptone (Difco) per liter of natural

seawater at room temperature. Natural seawater was obtained from the Biology II

pipeline, UCSB. Seawater was pumped through an 18 inch diameter polyvinyl chloride pipe located off the coast of UCSB and aged for one to five months at 4 °C in 20 L high-density polyethylene (HDPE) or polycarbonate carboys in the dark.

Glycerol stocks of SP18 were prepared from 800 µL of overnight culture in Marine

Broth 2216 (Difco) and 200 µL of sterile glycerol, frozen in liquid nitrogen, and

stored at -80 °C.

Siderophore Production

For siderophore production, Ochrobactrum sp. SP18 was grown in low-iron artificial seawater medium (ASW-Fe) containing 10 g casamino acids, 1 g ammonium chloride, 1 g disodium glycerophosphate hydrate, 12.35 g magnesium sulfate

heptahydrate, 1.45 g calcium chloride dihydrate, 17.55 g sodium chloride

(SigmaUltra, Sigma), 0.75 g potassium chloride, and 3 mL glycerol per liter of

doubly deionized water (Barnstead Nanopure II); following autoclaving, 10 mL of

53 filter-sterilized 1 M HEPES pH 7.4, 2 mL of filter-sterilized 1 M sodium bicarbonate, and 5 mL of filter-sterilized vitamin stock were added per liter of medium. The vitamin stock solution contained 40 mg biotin, 4 mg niacin, 2 mg thiamin, 4 mg para- aminobenzoic acid, 2 mg calcium pantothenic acid, 20 mg pyridoxine hydrochloride,

2 mg cyanocobalamin, 4 mg riboflavin, and 4 mg folic acid in 200 mL of doubly deionized water (Barnstead Nanopure II).

For siderophore production, a single colony of Ochrobactrum sp. SP18 was grown for 12 to 24 hours at room temperature on a fresh maintenance medium plate.

Cells were resuspended in 5 mL ASW-Fe and 2.5 mL of this suspension transferred to a 4 L acid-washed Erlenmeyer flask containing 2 L ASW-Fe. Cultures were grown at room temperature with shaking at 170 rpm on an orbital shaker for 48 to 72 hours.

Siderophore production was monitored by chrome-azurol sulfonate (CAS) assay using 1 part CAS solution containing CAS/Fe(III)/hexadecyl-trimethyl ammonium bromide and 1 part culture (Schwyn and Neilands 1987). On some occasions, 1 mL of culture was centrifuged at 16,000 × g for 30 seconds to pellet cells, the supernatant removed, the cells resuspended in 1 mL of doubly deionized water (Barnstead

Nanopure II), and then the supernatant and cell suspension each tested as described above. A color change to pink within 1 minute was interpreted as a positive CAS response.

To evaluate the affect of sodium chloride concentration, ASW-Fe media was prepared without added sodium chloride (0.01 M NaCl). These cultures were inoculated as described above for normal ASW-Fe media (0.31 M NaCl) and the

54 siderophores produced were compared.

Biofilm Formation

The ability of Ochrobactrum sp. SP18 to form biofilms was assessed using a modified version of the method described by Djordjevic et al. (2002). Briefly, a single colony of Ochrobactrum sp. SP18 was used to inoculate 10 mL each of Marine

Broth 2216 (Difco) and ASW-Fe and this suspension was grown for 12 hours at room temperature with shaking on an orbital shaker at 170 rpm. After 12 hours of growth,

100 µL of each culture was transferred to fresh tubes containing 10 mL of ASW-Fe and gently mixed. Six 100 µL aliquots of each inoculum and of sterile Marine Broth

2216 and ASW-Fe were transferred to a sterile microtiter plate and grown undisturbed for 48 hours at room temperature. The liquid was removed by aspiration, rinsed with doubly deionized water (Barnstead Nanopure II), and air dried for 45 minutes. Each well was then stained with 150 µL 1% (w/v) crystal violet for 20 minutes. The stain was removed by aspiration and the wells rinsed five times with doubly deionized water (Barnstead Nanopure II). A purple ring remaining in the well corresponds to the presence of a biofilm.

Siderophore Isolation

Cells from CAS positive cultures were harvested by centrifugation at 4800 × g

for 25 minutes at 4 °C in 6 acid-washed polycarbonate centrifuge tubes (Sorvall).

Cell pellets from individual tubes (i.e. from approximately 330 mL of culture) were

55 transferred to sterile 50 mL polyethylene conical tubes (Fisher) with 5 mL doubly deionized water (Barnstead Nanopure II) and extracted with 45 mL 200 proof ethanol

overnight at room temperature with shaking at 120 rpm on an orbital shaker. Extracts were filtered through 0.22 µm polyvinylidiene fluoride filters (Millipore) and concentrated under vacuum to roughly 10% of the initial volume. The concentrate was then applied to a Sep-Pak® Vac C18 cartridge (1 g sorbent, Waters Corp.), rinsed sequentially with 25 mL each of water and 50% methanol and eluted with 100%

methanol. Fractions were hand collected and tested for the presence of siderophore

with 1:1 CAS:eluent. Positive fractions were pooled and concentrated under vacuum

to approximately 10% of the original volume. Siderophores were purified by

reversed-phase HPLC (RP-HPLC) on a preparative C4 column (22 mm internal

diameter (ID) × 250 mm length (L), Vydac) with a gradient from 49.95% doubly

deionized water (Barnstead Nanopure II), 50% methanol, 0.05% trifluoroacetic acid

(TFA) to 99.95% methanol, 0.05% TFA over 37 minutes. Eluent was continuously

monitored at 215 nm and 415 nm with a Waters 2487 dual wavelength detector.

Peaks were hand collected, concentrated under vacuum in a Jouan concentrator

(Jouan, model 10.10) or by rotary evaporation, and then ultra-purified by application

to a preparative or semipreparative C4 column (10 mm ID × 250 mm L, Vydac) using the same gradient. Samples were concentrated as above and lyophilized. Purified siderophore samples were stored at -80 °C.

56 Structure Determination

Amino Acid Analysis

The amino acid composition of the siderophores produced by Ochrobactrum

sp. SP18 was determined by Marfey’s technique for amino acid analysis (Marfey

1984). Lyophilized samples of 1-10 mg of purified siderophore were hydrolyzed for

48 hours with 57% hydriodic acid (Inglis et al. 1971), repeatedly evaporated and washed with doubly deionized water (Barnstead Nanopure II) under vacuum, the resulting amino acid fragments derivatized with (1-fluoro-2,4-dinitrophenyl)-5-L- alaninamide (Marfey’s reagent, FDAA), and resolved by RP-HPLC on an analytical

YMC ODS-AQ C18 column (4.6 mm ID × 250 mm L, Waters Corp.) using a linear

gradient from 90% 50 mM triethylamine phosphate (TEAP) pH 3.0, 10% acetonitrile

to 60% 50 mM TEAP pH 3.0, 40% acetonitrile over 45 minutes (Marfey 1984). The eluent was continuously monitored at 340 nm on a Waters 2487 UV-visible detector.

Samples were compared to amino acid standards prepared in the same way.

Assignments were verified by coinjection of the siderophore sample and amino acid standard.

Fatty Acid Analysis

Fatty acid composition was determined by digestion of purified, lyophilized siderophores (roughly 1 to 10 mg) with 1 mL 3 N methanolic hydrochloride (Sigma) for 3 hours at 110 °C to generate methyl esters, extraction into 1 mL hexanes, and resolution by GC-MS (Hewlett-Packard 5800 with EI). Commercially available fatty

57 acids and fatty acid methyl esters were used as standards. NMR spectra (see below) confirmed the assignments for ochrobactin C.

NMR

Three independent 7 to 10 mg samples of ochrobactin C in 700 µL d6-

dimethylsulfoxide (d6-DMSO, 99.9%, Cambridge Isotopes, Inc.) were analyzed by

NMR. 1H, 13C, APT, gCOSY, gDQCOSY, gHMQC, and CIGAR were completed at

25 °C on a 500 MHz Varian UNITY INOVA instrument with the standard pulse

sequences and the data were analyzed with the provided software.

Mass Spectrometry

The masses of the siderophores were determined using electrospray ionization

mass spectrometry (ESI-MS) on a Micromass Q-TOF2 (Waters Corp.). Exact mass

determinations were made on the same instrument using a peptide standard for

calibration. Tandem mass spectrometry of the ochrobactins with the Q-TOF2 using

argon collision gas in concert with fatty acid analysis and Marfey’s amino acid

analysis were used to deduce the structures of ochrobactins A and B.

58 Results

Ochrobactrum sp. SP18 Siderophores

Ochrobactrum sp. SP18 produces a suite of at least three siderophores. The

siderophores were found associated with the cells under culture conditions, rather than in the culture supernatant as with most other known siderophores. The siderophores were easily extracted from the cell pellets with ethanol.

The rapid biofilm assay indicated that Ochrobactrum sp. SP18 does produce

biofilm material during growth in both Marine Broth 2216 and ASW-Fe media.

Assay results suggest that Ochrobactrum sp. SP18 produced more biofilm material

under low iron conditions (i.e. growth in ASW-Fe media). Growth in large ASW-Fe

cultures with shaking produces a dense foam plug on the surface of the culture,

ascribed to biofilm production during growth. No significant amount of siderophore

was found associated with this material and it was not studied further.

Alteration of the sodium chloride content of the culture significantly altered the relative amounts of the siderophores (Figures 2.2 and 2.3). In normal ASW-Fe

media (0.31 M sodium chloride), ochrobactin C was the principal product with small

amounts (roughly 10% of the total) of ochrobactin B produced (Figure 2.2). When

the sodium chloride content of the medium was decreased (0.01 M sodium chloride),

an additional siderophore was produced in quantities large enough to analyze

(ochrobactin A) and the ratio of ochrobactin B to C increased (Figure 2.3). All three

known ochrobactin siderophores were found associated with the cell pellet after

centrifugation, indicating the lipophilic nature of these siderophores.

1 C Abs (215 nm)

0 0 Time (min) 40

Figure 2.2: RP-HPLC of cell-extract from Ochrobactrum sp. SP18 grown in normal ASW-Fe (0.31 M sodium chloride). B, C = ochrobactins B and C, respectively.

B Abs (215 nm) A

0 0 Time (min) 40

Figure 2.3: RP-HPLC trace of cell-extract from Ochrobactrum sp. SP18 grown in reduced NaCl ASW-Fe media (0.01 M NaCl), showing the separation of ochrobactins A, B, and C.

61 Tandem mass spectrometry demonstrated similar fragmentation patterns for the individual siderophores, suggesting common structural components between the three siderophores (Figures 2.4, 2.5, and 2.6). Specifically, each fragmentation pattern demonstrated a loss of 46 mass units from the parent ion, similar to the petrobactins and aerobactin, indicating the presence of a citrate moiety (Barbeau et al. 2002). In addition, each fragmentation pattern included peaks at m/z 633, m/z

425, and m/z 315, suggesting the presence of common structural features in each siderophore. Each fragmentation pattern included the m/z 163, m/z 145, and m/z 128 pattern corresponding to fragmentation of lysine.

A small peak is evident in each fragmentation pattern corresponding to the acyl appendages; all patterns include a peak at m/z 153 and ochrobactins A and B also include a peak at m/z 125 or m/z 127, respectively, corresponding to the other fatty acid. Ochrobactin C demonstrates a single acyl appendage peak at m/z 153.

Exact mass experiments determined the mass of ochrobactin A to be m/z 757.4222

+ (M + H) , consistent with a molecular formula of C36O13N4H61 (∆ 1.7 ppm), ochrobactin B to be m/z 759.4396 (M + H)+, consistent with a molecular formula of

+ C36O13N4H63 (∆ 0.5 ppm), and ochrobactin C to be m/z 785.4513 (M + H) , consistent with a molecular formula of C38O13N4H65 (∆ 3.8 ppm). The exact mass of

Fe(III)-ochrobactin C was m/z 838.3655 (M-3H + FeIII + H)+, consistent with a molecular formula of C38O13N4H62Fe (∆ -0.27 ppm).

Marfey’s amino acid analysis indicated the presence of only L-lysine in each siderophore, as demonstrated by the presence of three peaks when resolved by RP-

62 HPLC (Figure 2.7). Three peaks would be anticipated since lysine can be derivatized at either or at both ends (Figure 2.8).

Mass spectrometry of purified samples of the siderophores determined the masses of the siderophores to be, in order of increasing quantity, ochrobactin A m/z

757, ochrobactin B m/z 759, and ochrobactin C m/z 785. Iron(III)-bound adducts are also apparent at 53 mass units higher in the mass spectrum for each siderophore, particularly when a small amount of formic acid is added to the sample. Fe(III) binding by citrate-derived dihydroxamate siderophores (see below) would be predicted to involve both hydroxamate functionalities and the citrate hydroxyl and carboxylic acid moieties. In order to observe the Fe(III)-ochrobactin complex in positive mode, the negatively charged ion must be neutralized (with one proton or sodium) and then must be protonated to be positively charged. The observed complex is therefore ([M-4H+Fe]+2H)+.

Figure 2.4: Tandem mass spectrum from ochrobactin A, m/z 757.5.

Figure 2.5: Tandem mass spectrum of ochrobactin B, m/z 759.4.

Figure 2.6: Tandem mass spectrum for ochrobactin C, m/z 785.43.

66 2.0 a Marfey’s reagent Marfey’s -derivative -derivative -derivative -derivative =340 nm) ε

1.0 α bis-derivative λ AU ( AU mono- mono-

0 0 30 60 Time (minutes)

2.0 b

=340 nm) 1.0 λ AU ( AU

0 0 30 60 Time (minutes) Figure 2.7: Marfey’s amino acid analysis. a) RP-HPLC resolution of derivatized hydrolysis fragments generated from ochrobactin C. b) Coinjection of the sample from a) with derivatized L-lysine. The peaks at 30 minutes are a degradation product of Marfey’s reagent.

NO2 O NO2 O NO2 O H H H N N N NH2 NH2 NH2

H3C H H3C H H3C H O2N O O2N O2N HN NH NH OH NH2 H O H CH H2N H HN 3 H OH OH H2N O2N NH O O

NO2

mono-α-derivative mono-ε-derivative bis-derivative

Figure 2.8: The three possible FDAA-L-lysine derivatives.

68 Qualitative observations and tandem mass spectrometry suggested the

presence of fatty acid appendages. The identities of the acyl appendages were

determined by digestion of purified siderophore samples with methanolic hydrochloride to generate fatty acid methyl esters which were resolved by GC-MS

(Figure 2.9). Ochrobactin C, the principal siderophore produced by Ochrobactrum sp. SP18, was found to contain only (E)-2-decenoic acid moieties. The secondary product, ochrobactin B, was found to contain both octanoic and (E)-2-decenoic acid moieties and the tertiary product, ochrobactin A, was found to contain both (E)-2-

octenoic acid and (E)-2-decenoic acid appendages. The identity of each fatty acid methyl ester was verified by comparison to authentic standards and published values

(Persmark et al. 1993, Okujo et al. 1994).

Figure 2.9: GC-MS fragmentation patterns of methyl esters generated from the ochrobactin siderophores, arrows indicate the parent ion in each case: a) ochrobactin A, methyl ester m/z 156; b) ochrobactin B, methyl ester m/z 158; c) ochrobactin C, methyl ester m/z 184. Ochrobactins A and B produce two peaks in the chromatogram (not shown), indicating the presence of an octanoic or (E)-2-octenoic acid moiety (respectively) and an (E)-2-decenoic acid moiety in each siderophore.

70 Spectral values from 1H and 13C NMR experiments with ochrobactin C are summarized in Tables 2.1 and 2.2. The 1H NMR spectrum (Figure 2.10) revealed the

α presence of two amide protons at δH 8.19 and δH 8.12, two lysine C protons at δH

4.14, an intense peak at δH 1.25 consistent with methylene protons, as well as an

13 intense triplet at δH 0.86. The C NMR spectrum (Figure 2.11) revealed the presence

of a quaternary carbon signal at δC 73.3 and five carbonyl carbon signals at δC 174.8,

δC 173.5, δC 169.7, δC 169.3, δC 165.2; the signals at δC 173.5 and δC 165.2 each

likely result from the overlap of two nearly identical carbonyl resonances (see below).

The 1H-1H gCOSY (Figures 2.12 and 2.13) revealed an alkyl group starting with the

triplet methyl signal, H1 (H1'), through the methylene resonance, H5 (H5'), via the

large peak at δH 1.25. Furthermore, the gCOSY and gHMQC (Figure 2.14) spectra

showed the connection of the methylene to one vinyl system, i.e., H8 (H8’) and H9

(H9’), which was also indicated by CIGAR-HMBC correlations (Figure 2.15) from

H5 (H5') at δH 2.17 to both C8 (C8’) at δC 145.2 and C9 (C9’) at δC 119.9. The large

coupling constants (15.5 Hz) between the vinyl protons are indicative of a trans configuration of the double bond. In the CIGAR spectrum, the vinyl protons showed correlations to a carbonyl carbon, C10 (C10') at δC 165.2. In addition to these data, the number of carbons corresponding to the alkyl chain at δH 1.25 seen in gHMQC

and 13C NMR spectra indicated that this monounsaturated alkyl group is (E)-2- decenoic acid. Comparison to previously published NMR data for (E)-2-decenoic acid and rhizobactin 1021 (containing the same fatty acid moiety) further supports this assignment (Persmark et al. 1993).

71 The APT spectrum (Figure 2.16) identified the peak at δC 51.6 as the α-carbon of the previously identified L-lysine residues and gHMQC correlations identified the

α-proton signals as δH 4.14. 2D NMR data and comparison to published values for aerobactin and L-lysine confirm the presence of lysine in ochrobactin C. The

presence of a citrate moiety was confirmed by gHMQC and CIGAR-HMBC

correlations from the methylene signals H18 (H18') to the quaternary carbon signal

(C19), two carbonyl signals C17 (C17'), and C20.

CIGAR-HMBC correlation from H11 (H11') in lysine to the carbonyl carbon

C10 (C10') indicated that each lysine is Nε-acylated by (E)-2-decenoic acid and the

absence of the Nε-amide proton of each lysine residue showed the presence of

hydroxamate groups at this position. Consequently, ochrobactin C was found to have

an acylated structure similar to aerobactin which is a well known siderophore produced by a wide variety of bacteria, including the lung pathogen Klebsiella pneumoniae (Gibson and Magrath 1969), many enteric strains (Crosa et al. 1988,

Lawlor and Payne 1984, Stuart et al. 1986), and has also been isolated from several species of marine bacteria (Haygood et al. 1993, Okujo and Yamamoto 1994,

Murakami et al. 1995, Murakami et al. 1998, Moon et al. 2004).

Interestingly, two distinct α-amide protons (δH 8.12 and δH 8.19) from the lysines are observed and the chemical shifts of citrate are unsymmetric, although the structure of ochrobactin C is symmetric. The distinct resonances of the two amide protons of lysine suggest a hydrogen bonding effect between one of the amide protons and the free carboxylic acid group (C20). This same asymmetry is apparent

72 1 in the H NMR of nannochelin A in 1:1 d6-DMSO:CHCl3 where the amide protons appear as two doublets at δH 8.19 and δH 8.13 (Bergeron and Phanstiel 1992).

1 Figure 2.10: H NMR of 7 mg ochrobactin C in 700 µL d6-DMSO.

13 Figure 2.11: C NMR of 7 mg ochrobactin C in 700 µL d6-DMSO.

Figure 2.12: gCOSY from 7 mg ochrobactin C in 700 µL d6-DMSO.

Figure 2.13: Expanded region of gCOSY from 7 mg ochrobactin C in 700 µL d6- DMSO.

Figure 2.14: gDQCOSY of 7 mg ochrobactin C in 700 µL d6-DMSO.

Figure 2.15: Expanded region of gDQCOSY of 7 mg ochrobactin C in 700 µL d6- DMSO.

Figure 2.16: gHMQC of 7 mg ochrobactin C in 700 µL d6-DMSO.

Figure 2.17: Expanded region of gHMQC of 7 mg ochrobactin C in 700 µL d6- DMSO.

Figure 2.18: CIGAR for 7 mg ochrobactin C in 700 µL d6-DMSO.

Figure 2.19: APT from 7 mg ochrobactin C in 700 µL d6-DMSO. The region from 60 to 180 ppm is expanded in the inset.

O OH 16' O O ii'13' 11' i' 8' 6' 4' 2' N15' N 10' 1' 17' H 14' 12' 9' 7' 5' 3' 18' OH OH 19 OH 20 18 O OH H 14 12 9 7 5 3 17 N 15 N 10 1 O ii 13 11 i 8 6 4 2 16 O O OH

Position 13C 1H C1, C1’ 13.9 0.86 (t, 6H, J = 6.5) C2, C2’ 25.9 (1.25m) C3, C3’ 30.7 (1.25m) C4, C4’ 28.6 (1.25m) C5, C5’ 28.5 (1.25m) C6, C6’ 27.9 1.40 (m, 4H) C7, C7’ 31.6 2.17 (q, 4H, J = 7.0) C8, C8’ 145.2 6.68 (dt, 2H, J = 15.5, 7.0) C9, C9’ 119.9 6.51 (d, 2H, J = 15.5) C10, C10’ 165.2 Ni,i’OH 9.8 (s, 2H) C11, C11’ 69.8 3.51 (m, 4H) C12, C12’ 22.4 1.52 (m, 4H) C13, C13’ 22.1 1.26 (m, 4H) C14, C14’ 31.2 1.68 (m, 2H) 1.56 (m, 2H) C15, C15’ 51.6 4.14 (m, 2H) NiiH 8.19 (d, 1H, J = 7.5) Nii’H 8.12 (d, 1H, J = 7.5) C16, C16’ 173.5 OH 12.5 (br) C17 169.7 C17’ 169.3 C18 43.0 2.55 (d, 1H, J = 12.5) 2.65 (d, 1H, J = 12.5) C18’ 42.8 2.61 (s, 2H) C19 73.3 C20 174.8 OH 12.5 (br)

Table 2.1: 1H and 13C assignments for ochrobactin C.

O OH O O N N H OH OH OH OH HO N N O O O OH = gCOSY = CIGAR

Position gCOSY gHMQC CIGAR C1, C1’ H2/H2’ H1/H1’ H2/H2’, H3/H3’ C2, C2’ H1/H1’ C3, C3’ H1/H1’, H2/H2’, H4/H4’ C4, C4’ C5, C5’ C6, C6’ H7/H7’ C7, C7’ H6/H6’, H8/H8’, H9/H9’ H7/H7’ H5/H5’, H6/H6’, H8/H8’, H9/H9’ C8, C8’ H7/H7’, H9/H9’ H8/H8’ H7/H7’ C9, C9’ H7/H7’, H8/H8’ H9/H9’ H7/H7’, H8/H8’ (weak) C10, C10’ H8/H8’ C11, C11’ H12/H12’ H11/H11’ C12, C12’ H11/H11’, H13/H13’ H12/H12’ C13, C13’ H12/H12’, H14/H14’ H13/H13’ H12/H12’ (weak) C14, C14’ H13/H13’, H15/H15’ H14/H14’ H13/H13’, H15/H15’ C15, C15’ H14/H14’, NiiH/Nii’H H15/H15’ H13/H13’, H14/H14’ C16, C16’ H14/H14’, H15/H15’ C17 NiiH, H15 (weak), H18 C17’ Nii’H, H15’ (weak), H18’ C18 H18 C18’ H18’ C19 H18/H18’ C20 H18/H18’

Table 2.2: Coupling and 2D NMR data for ochrobactin C.

85 The structures of ochrobactins A and B were deduced from amino acid

analysis, fatty acid analysis, and tandem mass spectrometry (Figure 2.17).

Ochrobactins A and B were found to be structurally related to ochrobactin C. The

difference between the siderophores was the composition of fatty acid appendages

attached to the Nε position of the lysine residues. Each siderophore contains one (E)-

2-decenoic acid moiety and ochrobactin A contains one (E)-2-octenoic acid moiety while ochrobactin B contains one octanoic acid moiety. Thus, the ochrobactins were found to be composed of citrate, symmetrically derivatized with L-lysine which is Nε- acylated and Nε-hydroxylated to form two hydroxamate binding groups. The three

siderophores differ only in the identities of the acyl appendages. Unlike most other

known siderophores, the ochrobactins are found associated with the cell membrane in

culture and are the first example of a citrate-type, cell-associated siderophore.

A common feature of mass spectra from citrate-derived siderophores is the

repetitive loss of 46 mass units; this feature is evident in the tandem mass spectra for

the ochrobactins and is the result of loss of the citryl carboxylic acid and two H+.

This loss of 46 mass units occurs in each daughter ion which contains the citrate moiety. Rationalization of the fragmentation for ochrobactin B is shown in Figure

2.18. For ochrobactin A, the peaks corresponding to loss of the citryl carboxylic acid moiety are m/z 711, m/z 587, m/z 559, m/z 425, m/z 397, and m/z 273. For ochrobactin C, the corresponding peaks are m/z 739, m/z 587, m/z 425, and m/z 273.

86 605 443 287 125

OH H H HO A N N N N O O OH O O O OH O OH O OH

153 315 471 633

607 445 289 127

OH H H HO B N N N N O O OH O O O O OH O OH OH

153 315 471 633

633 471 315 153

OH H H HO C N N N N O O OH O O O OH O OH O OH

153 315 471 633

Figure 2.20: Fragmentation of the ochrobactin siderophores by ESI-MSMS, using argon collision gas to generate daughter ions; deduced from the spectra in Figures 2.4, 2.5, and 2.6.

OH H H HO N N N N m/z 759 O O OH O O O O OH O OH OH

-46 (C, 2O, 2H)

OH H H HO N N N N O O O O O O O OH OH m/z 713 OH H H HO N N N N O O OH O O O O OH OH

Figure 2.21: Rationalization of fragments in ochrobactin B tandem mass spectrum. Fragmentation of the central citrate moiety results in loss of the citryl carboxylic acid functionality producing a 46 mass unit lighter stable fragment. Each fragment in the ochrobactin B tandem MS spectrum containing the citrate moiety produced corresponding fragments. The peaks at m/z 713, m/z 587, m/z 425, m/z 399, and m/z 273 are produced from ochrobactin B. Ochrobactins A and C also produced similar fragmentation patterns (see text).

O OH O O N N H OH OH OH A OH H O N N O O O OH O OH O O N N H OH OH OH B OH H O N N O O O OH O OH O O N N H OH OH OH C OH H O N N O O O OH

Figure 2.22: Structures of the three siderophores produced by Ochrobactrum sp. SP18.

89 Discussion

The ochrobactins are the first siderophores isolated from a marine α-

proteobacterium. The ochrobactins were determined to be citrate-derived

amphiphilic siderophores and are the first example of a membrane-associated citrate-

type siderophore, more specifically, the first amphiphilic derivatives of aerobactin.

These structures contain both of the common features identified from marine γ-

proteobacterial species: moieties that are photoreactive when coordinated to ferric ion and lipidic portions conferring amphiphilicity. Thus, the ochrobactin structures

support the notion that marine siderophores have distinct properties that are adaptive

in marine environments. The ochrobactins also share characteristics with rhizobactin

1021, the only structurally characterized terrestrial α-proteobacterial siderophore, as

well as the synechobactins, the only structurally characterized marine cyanobacterial

siderophores. Other known cell-associated siderophores include the mycobactins

produced by Mycobacteria (Ratledge and Dale 1999, Gobin and Horwitz 1996), the

structurally related formobactins (Murakami et al. 1996), nocobactins (Ratledge and

Patel 1976), and amamistatins (Kokubo et al. 2000, Suenaga et al. 1999), and the recently characterized amphibactins produced by marine Vibrio sp. R-10 (Martinez et

al. 2003). While amphiphilic and lipophilic siderophores are relatively uncommon

among known terrestrial bacteria, more than half of the known marine siderophores

are amphiphilic, predominantly produced as suites of compounds differing only

within a family of siderophores by the nature of the fatty acid appendage.

Important questions remain regarding the functional importance of these

90 amphiphilic moieties. It has been proposed that amphiphilic siderophores would be

retained near the bacterial cell, limiting diffusion and potentially increase iron

availability to the cell (Martinez et al. 2000). However, evidence is accumulating that bacterial cells in the water column primarily exist in consortia attached to particles or surfaces. These cells are often surrounded or encased by exopolysaccharides such as

in biofilms or carbohydrates such as in marine snow. The potential interactions

between these materials and amphiphilic siderophores have not been investigated.

These interactions could suggest other functions for the amphiphilic moieties of the

ochrobactins and other amphiphilic siderophores such as the marinobactins,

aquachelins, amphibactins, and synechobactins.

The presence of suites of siderophores varying only by the length of the fatty

acid appendage is also intriguing. Although other citrate-derived amphiphilic

siderophores have been isolated (namely rhizobactin 1021 (Lynch et al. 2001,

Persmark et al. 1993), acinetoferrin (Okujo et al. 1994), and the synechobactins (Ito

and Butler 2005) the ochrobactins are unique in that they possess two fatty acid

appendages that are not always identical. Variation of the length of appendage or the

ratio of siderophores in response to changes in environmental conditions is an

intriguing response, suggesting exquisite sensitivity to environmental conditions and

functionality dependent on the fatty acid composition of the siderophore complexes.

The first family of marine α-proteobacterial siderophores thus expand our

knowledge of iron acquisition strategies of marine bacteria and lend support to the

idea that amphiphilicity and photoreactivity contribute to the success of bacteria in

91 marine environments.

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96 Chapter Three

Siderophore production by Vibrio sp. DS40M5, Citrobacter koseri ATCC 27156, and Klebsiella pneumoniae ATCC 25306

Introduction

Structural Similarities between Aerobactin and the Ochrobactins

The structural similarities between the ochrobactins and aerobactin are evident

(Figure 3.1). Both are constructed of citrate symmetrically derivatized with L-lysine

which is Nε-acylated and Nε-hydroxylated to form two hydroxamate moieties. In the

case of aerobactin, each lysine is Nε-acetylated (Gibson and Magrath 1969); in the

ochrobactins, one lysine is Nε-acylated with (E)-2-decenoic acid while the other is Nε- acylated with (E)-2-octenoic, octanoic, or (E)-2-decenoic acid (Chapter 2).

Diversity of Aerobactin Production

Aerobactin is produced by a wide variety of bacteria (Table 3.1), including the

lung pathogen Klebsiella pneumoniae (Gibson and Magrath 1969), many enteric

strains (Lawlor and Payne 1984, Stuart et al. 1986, Crosa et al. 1988), and numerous

species of marine bacteria (Haygood et al. 1993, Okujo and Yamamoto 1994,

Murakami et al. 1995, Murakami et al. 1998, Moon et al. 2004). In addition to

bacterial strains that synthesize aerobactin, bacteria such as Vibrio vulnificus M2799

and Vibrio parahaemolyticus WP1 produce the outer membrane transport proteins to

utilize exogenous ferric-aerobactin as an iron source, yet do not synthesize the

siderophore themselves (Funahashi et al. 2003, Tanabe et al. 2005). Aerobactin has

O OH O O N N H OH OH OH A OH H O N N O O O OH O OH O OH O O O O N N N N H H OH OH OH OH OH OH B OH OH H O H O N N N N O O O O O OH O OH O OH O O N N H OH OH OH C OH H O N N O O O OH

Figure 3.1: Structures of aerobactin (left (Gibson and Magrath 1969)) and the ochrobactins (right, Chapter 2 (Martin et al. in press)).

Bacterial species Techniques References Citrobacter diversus B,D Chen et al. 2000 Citrobacter freundii (1/15 strains) F Mokracka et al. 2004 Citrobacter koseri (3/3 strains) F1 Mokracka et al. 2004 Citrobacter koseri (5/5 strains) F1 Khashe and Janda 1996 Enterobacter cloaceae (42/85 strains) F1 Mokracka et al. 2004 Enterobacter cloacae EK33 B,D Crosa et al. 1988 Escherichia coli: A Ewers et al. 2005 14/14 avian pathogenic 3/9 non pathogenic 1/6 enterotoxic 1/5 uropathogenic Escherichia fergusonii (3/50 strains) F2 Šmajs et al. 2003 Klebsiella pneumoniae B,M,N Gibson and Magrath 1969 (Aerobacter aerogenes 62-1) Original Structure Klebsiella pneumoniae (17/34 strains) F1 Koczura and Kaznowski 2003 Pseudomonas sp. X40 B,M,N Buyer et al. 1991 Shigella boydii 1392 G Lawlor and Payne 1984 Shigella flexneri 2457 B,G,H Payne 1980 Shigella flexneri NTCC4839 Shigella flexneri UP4204 Shigella flexneri SA100 G Lawlor and Payne 1984 Shigella flexneri (9 strains) I Payne et al. 1983 Shigella sonnei E Payne 1988 Vibrio hollisae ATCC 33564 B,C,D Okujo and Yamamoto 1994 Vibrio mimicus ATCC 33653 B,C,D Okujo and Yamamoto 1994 E Moon et al. 2004 Vibrio sp. B,C,O Murakami et al. 1995 Vibrio orientalis sp. SD 004 G,J Murakami et al. 1998 Vibrio nereis sp. SD 248 G,J Murakami et al. 1998 Vibrio splendidus sp. SD 101 G,J Murakami et al. 1998 Vibrio tubiashi sp. SD 102 G,J Murakami et al. 1998 Vibrio sp. DS40M5 B,H,K,L Haygood et al. 1993 Yersinia frederiksenii (7/12 strains) D,N Stuart et al. 1986 Yersinia intermedia (5/5 strains) D,N Stuart et al. 1986 Yersinia kristensenii (1/5 strains) D,N Stuart et al. 1986 Table 3.1: Aerobactin producing strains, indicating method of characterization. A, PCR amplification of iucD; B, 1H NMR; C, 13C NMR; D, FABMS; E, genetic analysis; F1, biological assay involving cross feeding to siderophore-deficient receptor-proficient E. coli LG1522; F2, biological assay involving cross feeding to siderophore-deficient receptor-proficient E. coli H1887; G, TLC using authentic aerobactin as a standard; H, UV-visible spectroscopy; I, biological assay; J, HPLC with authentic aerobactin as a standard; K, IR spectroscopy; L, 2D 1H NMR; M, amino acid analysis; N, paper electrophoresis; O, secondary ion mass spectrometry.

99 been isolated from numerous pathogenic species and it has been proposed that

aerobactin may be a virulence factor. However, it has also been isolated from many

marine and soil bacteria suggesting that this siderophore is widely utilized, and not

just by pathogenic bacteria.

Citrobacteria

Citrobacteria are important environmental strains and many strains are

pathogenic. Investigation of the iron acquisition strategies by Citrobacter species has

revealed that the majority are able to produce enterobactin (70 of 71 species in one

study) (Khashe and Janda 1996). In addition, roughly 25% carry the genes to

synthesize yersiniabactin (Schubert et al. 2000, Mokraka et al. 2004). Aerobactin has been detected in the supernatants of many Citrobacter species, particularly from uropathogenic species and from environmental samples. One recent study also detected the presence of an undetermined hydroxamate siderophore, distinct from aerobactin, from cultures of a Citrobacter koseri species using a biological assay

(Mokraka et al. 2004).

Siderophore Isolation: Cell-Associated versus Secreted Siderophores

The vast majority of structurally characterized siderophores from terrestrial

and even marine bacteria have been detected in and isolated from the culture

supernatant. Notably, the mycobactins produced by Mycobacteria and the

structurally related formobactins (Murakami et al. 1996), nocobactins (Ratledge and

100 Patel 1976), and amamistatins (Kokubo et al. 2000, Suenaga et al. 1999) are cell- associated siderophores; these siderophores have been extracted from the cell pellets with ethanol for structural elucidation. The discovery of the amphibactins from marine Vibrio sp. R-10 (Martinez et al. 2003) and the ochrobactins from marine

Ochrobactrum sp. SP18 (Chapter 2), both of which were easily extracted from the cell pellet with ethanol, suggested that other bacteria might also contain cell- associated siderophores, including in particular strains producing aerobactin.

Following the discovery of the ochrobactins, three aerobactin-producing bacterial strains were evaluated for production of aerobactin-based cell-associated siderophores. Vibrio sp. DS40M5, a marine γ-proteobacterium has previously been shown to produce aerobactin (Haygood et al. 1993). This isolate was collected from a depth of 40 m over the continental slope off West Africa (Haygood et al. 1993).

Three other isolates from the same water sample produce suites of amphiphilic siderophores: the marinobactins produced by Marinobacter sp. DS40M6 and

DS40M8 and the aquachelins produced by Halomonas aquamarina sp. DS40M3

(Martinez et al. 2000). Vibrio sp. DS40M5 was therefore selected for evaluation for production of cell-associated siderophores. All nine of the reported strains of

Citrobacter koseri (previously C. diversus) have been found to produce aerobactin

(Khashe and Janda 1994, Chen et al. 2000, Mokracka et al. 2004). It was thus reasonable to expect that C. koseri ATCC 27156, a strain not previously studied,

101 would also synthesize aerobactin. This soil bacterium was therefore selected for

analysis as an example of a terrestrial aerobactin-producing strain. Klebsiella

pneumoniae ATCC 25306 is the original source of aerobactin (Gibson and Magrath

1969). This pathogenic strain was also evaluated for the presence of cell-associated siderophores.

102 Materials and Methods

Siderophore Production by Vibrio sp. DS40M5

Growth and Maintenance of Vibrio sp. DS40M5

Vibrio sp. DS40M5 was maintained on natural seawater agar plates containing

0.5 g yeast extract (Difco), 5 g BactoPeptone (Difco), and 15 g BactoAgar (Difco) per liter of natural seawater. Natural seawater was obtained from the Biology II pipeline,

UCSB. Seawater was pumped through an 18 inch diameter polyvinyl chloride pipe located off the coast of UCSB and aged for one to five months at 4 °C in 20 L high- density polyethylene (HDPE) or polycarbonate carboys in the dark. Freezer stocks of

Vibrio sp. DS40M5 were prepared from 800 µL overnight culture in Marine Broth

2216 (Difco) and 200 µL sterile glycerol, frozen in liquid nitrogen, and stored at -80

°C.

Isolation of Siderophores from Vibrio sp. DS40M5

For siderophore production, Vibrio sp. DS40M5 was grown in artificial seawater medium M5 (ASW-M5) containing 15.5 g sodium chloride (SigmaUltra),

0.75 g potassium chloride, 12.35 g magnesium sulfate heptahydrate, 2.9 g calcium chloride dihydrate, 2 g casamino acids, 1 g ammonium chloride, and 0.1 g disodium glycerophosphate hydrate per liter of doubly deionized water (Barnstead Nanopure

II); the pH after autoclaving was 7.4. A single colony of Vibrio sp. DS40M5 was streaked onto a natural seawater agar plate to form a bacterial lawn and grown overnight at room temperature. The bacterial lawn was gently resuspended in 5 mL

103 ASW-M5 and 2.5 mL of this suspension was used to inoculate 2 L ASW-M5 in a 4 L acid-washed Erlenmeyer flask. Cultures were grown at room temperature with shaking at 120 rpm on an orbital shaker for four days.

Cultures were harvested by centrifugation at 4800 × g for 30 minutes at 4 °C.

For isolation of aerobactin, the culture supernatant was incubated with 200 mL of

Amberlite XAD-2 resin (Supelco Inc.) with shaking at 120 rpm for one hour or until a

1:1 supernatant:Fe(III)-CAS solution assay was negative (Schwyn and Neilands

1987). The resin was washed with 200 mL doubly deionized water (Barnstead

Nanopure II) to remove salts and aerobactin was eluted with methanol. Positive fractions were concentrated by rotary evaporation and then purified by RP-HPLC on a preparative C4 column (22 mm internal diameter (ID) × 250 mm length (L), Vydac) using a linear gradient from 0.05% trifluoroacetic acid (TFA) in doubly deionized water (Barnstead Nanopure II) to 0.05% TFA, 74.95% doubly deionized water

(Barnstead Nanopure II), 25% acetonitrile over 30 minutes. The eluent was continuously monitored at 215 nm with a Waters 2487 detector. Fractions were hand collected and immediately concentrated under vacuum in a Jouan concentrator

(Jouan, model 10.10). Aerobactin was ultra-purified by application to a semipreparative C4 column (10 mm ID × 250 mm L, Vydac) using the same gradient, fractions hand collected, concentrated under vacuum, lyophilized, and stored as a powder at -80 °C.

For isolation of cell-associated siderophores, the cell pellets from approximately 330 mL of culture were transferred to sterile 50 mL polyethylene

104 conical tubes and extracted overnight with 90% ethanol at room temperature with

shaking at 60 – 120 rpm on an orbital shaker. The extracts were filtered through a

0.22 µm polyvinylidiene fluoride filter (PVDF, Millipore) to remove cell debris and concentrated under vacuum to roughly 10% of the initial volume. The concentrate was then applied to a Sep-Pak® Vac C18 cartridge (1 g sorbent, Waters, Corp.), rinsed sequentially with 25 mL each of doubly deionized water (Barnstead Nanopure II),

50% methanol, and 100% methanol. Fractions were hand collected and tested for the presence of siderophores with 1:1 Fe(III)-CAS solution assay:eluent. Positive fractions from 50% methanol were pooled and concentrated under vacuum to approximately 10% of the initial volume. Positive fractions from 100% methanol

were pooled separately and concentrated as above. Siderophores were purified by

RP-HPLC on a preparative C4 column (22 mm ID × 250 mm L, Vydac) with a gradient from 49.95% doubly deionized water (Barnstead Nanopure II), 50% methanol, 0.05% TFA to 99.95% methanol, 0.05% TFA over 37 minutes. The eluent was continuously monitored at 215 nm and 415 nm with a Waters 2487 dual wavelength detector. Fractions were collected by hand and concentrated as described above.

Analysis of Siderophores Produced by Vibrio sp. DS40M5

Concentrated, purified siderophore solutions were analyzed by electrospray ionization mass spectrometry (ESI-MS), tandem mass spectrometry (MSMS), and exact mass determination on a Micromass Q-TOF2 mass spectrometer (Waters,

105 Corp.). For MSMS, argon collision gas was used to generate daughter ions. Exact

mass determinations were made with coinjection of the sample and a peptide standard

for calibration. Samples were dissolved in methanol with a small amount of formic acid.

Siderophore Production by Citrobacter koseri ATCC 27156

Growth and Maintenance of Citrobacter koseri ATCC 27156

Citrobacter koseri ATCC 27156 was purchased from the American Type

Culture Collection (ATCC). The lyophilized pellet was resuspended in Luria-Bertani

(LB) broth and 20 µL spread on a fresh Nutrient Agar (Difco) plate. Plates were

maintained at room temperature. Freezer stocks were prepared from 800 µL

overnight culture in LB and 200 µL sterile glycerol, frozen in liquid nitrogen, and

stored at -80 °C.

Isolation of Siderophores from Citrobacter koseri ATCC 27156

For siderophore production, a single colony of C. koseri ATCC 27156 was

streaked onto a fresh Nutrient Agar or natural seawater agar plate to form a bacterial

lawn and grown overnight at room temperature. C. koseri ATCC 27156 was grown

in four different media to test for siderophore production: HEPES-succinate

(modified from (Simon 1963, Khashe and Janda 1996)), Medium 56 (modified from

(Gibson and Magrath 1969, Monod et al. 1951)), low-iron artificial seawater (ASW-

Fe; 0.31 M NaCl), and low NaCl ASW-Fe (0.01 M NaCl). In each case, 5 mL sterile

106 medium was used to gently resuspend overnight bacterial lawns and 2.5 mL of the

cell suspension was transferred to 2 L of sterile medium in 4 L acid-washed

Erlenmeyer flasks. Cultures were grown for 0.5 to 5 days at room temperature with shaking at 170 rpm on an orbital shaker. HEPES-succinate medium contained 5.8 g sodium chloride (SigmaUltra), 3.7 g potassium chloride, 0.15 g calcium chloride dihydrate, 0.1 g magnesium chloride hexahydrate, 1.1 g ammonium chloride, 0.15 g sodium sulfate, and 0.27 g monobasic potassium phosphate per liter of doubly deionized water (Barnstead Nanopure II). After autoclaving, 10 mL filter-sterilized

50% disodium succinate (w/v) and 10 mL filter-sterilized 1.0 M HEPES pH 7.4 were added. Medium 56 contained 13.6 g monobasic potassium phosphate, 2 g ammonium sulfate, 0.2 g magnesium sulfate heptahydrate, and 0.01 g calcium chloride dihydrate per liter of doubly deionized water (Barnstead Nanopure II); 10 mL filter-sterilized

50% glucose (w/v) was added after autoclaving. ASW-Fe contained 10 g casamino acids, 1 g ammonium chloride, 1 g disodium glycerophosphate hydrate, 12.35 g magnesium sulfate heptahydrate, 1.45 g calcium chloride dihydrate, 17.55 g sodium chloride (SigmaUltra), 0.75 g potassium chloride, and 3 mL glycerol per liter of doubly deionized water (Barnstead Nanopure II); following autoclaving, 10 mL filter- sterilized 1 M HEPES pH 7.4, 2 mL filter-sterilized 1 M sodium bicarbonate, and 5 mL of filter-sterilized vitamin stock solution were added per liter of medium. The vitamin stock solution contained 40 mg biotin, 4 mg niacin, 2 mg thiamin, 4 mg para- aminobenzoic acid, 2 mg calcium pantothenic acid, 20 mg pyridoxine hydrochloride,

2 mg cyanocobalamin, 4 mg riboflavin, and 4 mg folic acid in 200 mL doubly

107 deionized water (Barnstead Nanopure II). Low-NaCl ASW-Fe is as above, but does

not contain added sodium chloride.

Cells were pelleted by centrifugation at 4800 × g for 30 minutes at 4 °C. To

isolate secreted siderophores, the supernatant was incubated with 200 mL of

Amberlite XAD-2 resin (Supelco) at room temperature with gentle shaking for 1 hour

or until a 1:1 Fe(III)-CAS:supernatant assay was negative. The resin was rinsed with

200 mL doubly deionized water (Barnstead Nanopure II) to remove salts and iron- binding compounds were eluted with 200 mL 50% methanol and 200 mL 100% methanol. Fractions were hand collected and assayed for siderophore with 1:1

Fe(III)-CAS:eluent. Positive fractions from 50 or 100% methanol were independently pooled and concentrated by rotary evaporation. Siderophores from the

50% methanol fractions were purified by RP-HPLC using a preparative C4 column

(22 mm ID × 250 mm L, Vydac) using a linear gradient from 0.05% TFA in doubly

deionized water (Barnstead Nanopure II) to 0.05% TFA, 74.95% doubly deionized

water (Barnstead Nanopure II), 25% acetonitrile over 30 minutes. Siderophores from

100% methanol fractions were purified by RP-HPLC on a preparative C4 column

with a linear gradient from 49.95% doubly deionized water (Barnstead Nanopure II),

50% methanol, 0.05% TFA to 0.05% TFA in methanol over 37 minutes. In each case, the eluent was continuously monitored at 215 nm with a Waters 2487 detector,

fractions were hand collected, and immediately concentrated under vacuum in a

Jouan concentrator (Jouan, model 10.10).

The cell pellets were transferred to sterile 50 mL polyethylene conical tubes

108 and extracted with 90% ethanol overnight at room temperature with shaking at 120 rpm on an orbital shaker. The ethanol extracts were filtered through a 0.22 µm PVDF filter (Millipore) and concentrated under vacuum to roughly 10% of the initial volume. Concentrated extracts were applied to a Sep-Pak® Classic C18 cartridge (360 mg sorbent, Waters, Corp.), sequentially rinsed with 20 mL each of doubly deionized water (Barnstead Nanopure II) and 50% methanol and eluted with 20 mL of 100% methanol. Fractions were hand collected, concentrated under vacuum, and each fraction was assayed for iron-binding compounds by 1:1 Fe(III)-CAS:concentrated eluent assay.

Analysis of Siderophores Produced by Citrobacter koseri ATCC 27156

Siderophores produced by C. koseri ATCC 27156 were analyzed by ESI-MS and MSMS on a Micromass Q-TOF2 mass spectrometer (Waters, Corp.) and compared to published data (Martinez et al. 2001, Martinez 2002, Küpper et al. submitted 2006).

Siderophore Production by Klebsiella pneumoniae ATCC 25306

Growth and Maintenance of Klebsiella pneumoniae ATCC 25306

K. pneumoniae ATCC 25306 was obtained from the ATCC as a lyophilized pellet. The pellet was resuspended in LB broth and 20 µL spread on a fresh Nutrient

Agar (Difco) plate. Plates were incubated at room temperature. Freezer stocks were prepared from 800 µL overnight culture in LB and 200 µL sterile glycerol, frozen in

109 liquid nitrogen, and stored at -80 °C.

Isolation of Siderophores from Klebsiella pneumoniae ATCC 25306

For siderophore studies, K. pneumoniae ATCC 25306 was grown in ASW-Fe,

low-NaCl ASW-Fe, Medium 56, and HEPES-succinate media. In each case, a single

colony was transferred to a fresh Nutrient Agar or natural seawater agar plate to form

a bacterial lawn and grown overnight at room temperature. Cells were resuspended in

5 mL sterile medium and 2.5 mL used to inoculate 2 L of medium in a 4 L acid-

washed Erlenmeyer flask. Cultures were grown at room temperature with shaking on

an orbital shaker at 120 rpm for up to 5 days.

Cultures were harvested by centrifugation at 4800 × g for 30 minutes at 4 °C.

Aerobactin was isolated from the supernatant as described above for Vibrio sp.

DS40M5. Cell pellets from K. pneumoniae ATCC 25306 were extracted and assayed

as described above.

Biosynthesis of 15N-labeled Aerobactin and Feeding Experiments

Biosynthesis and Purification of 15N-labeled Aerobactin

15N-labeled aerobactin was produced by growing C. koseri ATCC 27156 in

15 HEPES-succinate medium prepared with NH4Cl (Cambridge Isotopes). A single colony of C. koseri ATCC 27156 was transferred to a fresh natural seawater agar plate to form a bacterial lawn and grown at room temperature overnight. Cells were gently resuspended in 5 mL 15N supplemented medium and this inoculum transferred

110 to 1 L of 15N enriched medium in a 2 L acid-washed Erlenmeyer flask. Cultures were grown with shaking on an orbital shaker at 170 rpm for 4 days at room temperature.

Cultures were harvested and purified as described above. 15N-labeled aerobactin production was verified by ESI-MS and MSMS on a Micromass Q-TOF2 mass spectrometer (Waters, Corp.).

Aerobactin Feeding Experiment with Ochrobactrum sp. SP18

A single colony of Ochrobactrum sp. SP18 was transferred to a fresh natural seawater agar plate and grown overnight at room temperature to form a bacterial lawn. Cells were gently resuspended in 5 mL ASW-Fe and this inoculum was added to 1 L ASW-Fe. Cultures were grown at room temperature with shaking on an orbital shaker at 170 rpm for 72 hours, early in stationary phase. Cells were pelleted by centrifugation at 4800 × g for 30 minutes at 4 °C. Pellets were gently rinsed and resuspended in fresh ASW-Fe, diluted to a final volume of 1 L with ASW-Fe and supplemented with approximately 2.5 mg 15N-labeled aerobactin. Supplemented cultures were incubated at room temperature with shaking on an orbital shaker at 170 rpm for 24 hours and then one-half of the culture was harvested. The remaining one- half was grown for an additional 24 hours and then harvested at 48 hours. At each timepoint, cells were pelleted by centrifugation, cell pellets transferred to sterile 50 mL polyethylene conical tubes, and extracted overnight with shaking at 60 – 120 rpm on an orbital shaker in 90% ethanol. Ethanol extracts were filtered with a 0.22 µm

111 PVDF SteriFlip® filter unit (Millipore) and concentrated under vacuum to approximately 10% of the initial volume.

Concentrated cell extracts were applied to a Sep-Pak® Classic C18 cartridge

(360 mg sorbent; Waters, Corp.), washed with 20 mL doubly deionized water

(Barnstead Nanopure II), and eluted with 100% methanol. Fractions were hand collected and siderophores detected with 1:1 Fe(III)-CAS:eluent assay. Positive fractions were pooled and concentrated under vacuum. Siderophores were purified by RP-HPLC on a preparative C4 column (22 mm ID × 250 mm L, Vydac) with a gradient from 49.95% doubly deionized water (Barnstead Nanopure II), 50% methanol, 0.05% TFA to 99.95% methanol, 0.05% TFA over 37 minutes. Eluent was continuously monitored at 215 nm and 415 nm with a Waters 2487 dual wavelength detector. Peaks were hand collected and concentrated under vacuum. Purified siderophores were analyzed by ESI-MS and MSMS on a Micromass Q-TOF2 mass spectrometer (Waters, Corp.) in methanol with a small amount of formic acid.

Attempts to Produce 15N-labeled Ochrobactins

In order to produce 15N-labeled ochrobactins, several media containing only

NH4Cl as a nitrogen source were prepared. The media are summarized in Table 3.2.

112

ASW-Fe (usual medium) ASW+Succinate 5.00 g casamino acids 7.75 g NaCl 0.50 g NH4Cl 0.40 g KCl 0.05 g glycerophosphate 0.10 g MgSO4·7H2O 6.20 g MgSO4·7H2O 0.05 g CaCl2·2H2O 0.75 g CaCl2·2H2O 0.50 g NH4Cl 8.80 g NaCl 2.50 g disodium succinate 0.40 g KCl 1.50 g Na2HPO4 1.5 mL glycerol 500 mL doubly deionized 500 mL doubly deionized water (Barnstead water (Barnstead Nanopure II) Nanopure II) after autoclaving, add: 5.0 mL 1M HEPES pH 7.4 1.0 mL 1M NaHCO3 5.0 mL vitamin stock (see p. 107) Modified ASW-M5 ASW+Pyruvate 7.75 g NaCl 7.75 g NaCl 0.38 g KCl 0.40 g KCl 6.18 g MgSO4·7H2O 0.10 g MgSO4·7H2O 1.45 g CaCl2·2H2O 0.05 g CaCl2·2H2O 1.00 g NH4Cl 0.50 g NH4Cl 0.13 g glycerophosphate 2.50 g pyruvate 500 mL doubly deionized 1.50 g Na2HPO4 water (Barnstead 500 mL doubly deionized Nanopure II) water (Barnstead Nanopure II) Table 3.2: Artificial seawater (ASW) media utilized for growth of 15N-labeled ochrobactins.

113 Results

Vibrio sp. DS40M5, Citrobacter koseri ATCC 27156, and Klebsiella pneumoniae ATCC 25306 were evaluated for production of cell-associated siderophores. Media utilized for these experiments are summarized in Table 3.3.

Strain Medium Vibrio sp. DS40M5 ASW-M5 ASW-Fe (0.31 M NaCl) C. koseri ATCC 27156 ASW-Fe (0.31 M NaCl) ASW-Fe (0.01 M NaCl) HEPES-succinate Medium 56 K. pneumoniae ATCC 25306 ASW-Fe (0.31 M NaCl) ASW-Fe (0.01 M NaCl) HEPES-succinate Medium 56 Ochrobactrum sp. SP18 ASW-Fe (0.31 M NaCl) ASW-Fe (0.01 M NaCl) Table 3.3: Growth media tested with each strain. Media recipes appear on page 103 (ASW-M5) and 106-107 (HEPES-succinate, Medium 56, and ASW-Fe).

Isolation of Siderophores from Vibrio sp. DS40M5

Vibrio sp. DS40M5 has previously been shown to produce aerobactin as a primary siderophore (Haygood et al. 1993). As expected (Haygood et al. 1993), roughly 2 mg/L of aerobactin was isolated from Vibrio sp. DS40M5 cultures (Figure

3.2, 3.3, and 3.4). Under acidic conditions such as ESI-MS, the presence of aerobactin is evident in the ESI-mass spectrum aerobactin (m/z 565 (M+H)+).

Cleavage at the amide bonds to generate y- and b-type fragments (Roepstorff and

Fohlman 1984) produces fragments at m/z 361 ((M)+, composed of N-Ac, N-OH- lysine and citrate) and m/z 205 ((M+2H)+, consisting of N-Ac, N-OH-lysine) (Table

114 3.4). Characteristic fragmentation of the citryl α-hydroxy carboxylic acid moiety produces m/z 519 and m/z 315, corresponding to loss of 46 mass units (Bandu et al.

2006). Loss of one terminal acetyl group produces the peaks at m/z 523, m/z 477, m/z 273, and m/z 163. Peaks corresponding to loss of the second acetyl group are also present but are of very low intensity. Fragmentation of N-Ac, N-OH lysine produces fragments at m/z 205, m/z 163, m/z 142, and m/z 128. A citryl fragment is visible at m/z 159.

Extraction of the cell pellets of Vibrio sp. DS40M5 revealed the presence of cell-associated iron binding compounds, in addition to the previously identified hydrophilic siderophore aerobactin. Resolution by RP-HPLC revealed the presence of at least two Fe(III)-CAS positive compounds (Figure 3.5). Analysis of the purified siderophores by ESI-MS and MSMS (Figures 3.6, 3.7, and 3.8) indicated that the extracted iron-chelating fractions collected at 27.5 and 29.0 minutes (Figure 3.5) were ochrobactins B and C, respectively. The peak collected at 27.5 minutes, identified as ochrobactin B (Figures 3.6 and 3.8), exhibited virtually a identical fragmentation pattern to ochrobactin B isolated from Ochrobactrum sp. SP18 (Figure 2.5).

Fragmentation of the parent ion at m/z 759 generates the primary fragments noted previously, specifically, m/z 633, m/z 607, m/z 471, m/z 445, m/z 315, m/z 289, and m/z 153. The peak at m/z 127 corresponding the octanoate acyl appendage of ochrobactin B is either of low intensity or is not present in the fragmentation pattern.

The acyl appendage peaks are normally of low intensity in MSMS spectra, particularly when the concentration of the parent ion is low. As has been noted for

115 other citrate-derived siderophores, the loss of 46 mass units is also evident in the

fragmentation pattern. Peaks at m/z 713, m/z 587, m/z 425, m/z 399, and m/z 273

each derive from fragmentation of the citrate moiety (i.e. loss of 46 amu) (Bandu et

al. 2006). The remaining peaks derive from dehydration as noted for MSMS of

peptide-like compounds. Similarly, the fragmentation pattern for the peak collected

at 29 minutes (Figures 3.7 and 3.8) aligns well with the pattern generated from

ochrobactin C isolated from Ochrobactrum sp. SP18 (Figure 2.6). The primary

fragments at m/z 633, m/z 471, m/z 315, m/z 153 were generated from fragmentation

of the parent ion at m/z 785. The characteristic pattern deriving from citrate is also

evident in the fragmentation pattern; peaks at m/z 739, m/z 587, m/z 425, and m/z

273 are all attributed to fragmentation of the citryl carboxyl moiety (Bandu et al.

2006). As above, the remaining peaks derive from cyclization or dehydration as seen

in peptide-like compounds.

The exact mass for ochrobactin B isolated from Vibrio sp. DS40M5 was m/z

+ 759.4394 for (M+H) , which is consistent with C36H63N4O13 (∆1.03 ppm). The exact mass for ochrobactin C from Vibrio sp. DS40M5 was m/z 785.4555 for (M+H)+, which is consistent with C38H65N4O13 (∆1.57 ppm). These numbers are very similar

to those determined for ochrobactins B and C isolated from Ochrobactrum sp. SP18;

+ specifically, m/z 759.4296 for (M+H) , consistent with C36H63N4O13 (∆0.5 ppm) for

+ ochrobactin B and m/z 785.4513 for (M+H) , consistent with C38H65N4O13 (∆-3.77

ppm) for ochrobactin C.

To evaluate the affect of growth media on ochrobactin production, Vibrio sp.

116 DS40M5 was also grown in ASW-Fe medium. As seen with this strain in ASW-M5 medium, aerobactin was produced as the principal siderophore with small amounts of ochrobactins B and C extractible from the cell pellet. The total amount of siderophore was slightly reduced (likely due to higher iron(III) content in the ASW-

Fe medium). Ochrobactrum sp. SP18 was not found to produce aerobactin in ASW-

Fe medium but would not grow in ASW-M5 medium. It is likely that Ochrobactrum sp. SP18 has an absolute requirement for one or more of the vitamins in the vitamin stock solution that is added to ASW-Fe medium; further attempts to alter the growth medium for Ochrobactrum sp. SP18 may be successful if vitamin stock is added.

Thus, based on MS, MSMS, and exact mass determinations aerobactin and ochrobactins B and C were produced in the same culture by Vibrio sp. DS40M5.

117

Figure 3.2: RP-HPLC purification of aerobactin from 50% methanol XAD-2 fractions from cultures of Vibrio sp. DS40M5. The large peak at 38.2 minutes was determined to be aerobactin.

118

Figure 3.3: MSMS spectrum from fragmentation of aerobactin purified from Vibrio sp. DS40M5, m/z 565 (M+H)+.

119

523 361 205 (43)

OH H H HO N N N N O O OH O O O OH O OH O OH

(43) 205 361 523

Figure 3.4: Fragmentation pattern of aerobactin; parent ion m/z 565. The fragmentation pattern also includes loss of 46 mass units from each fragment containing the central citryl moiety, i.e. the peaks at m/z 519, m/z 501, m/z 315.

120

OH H H HO N N N N 565 O O OH O O O OH O OH O OH

OH H H HO N N N N H 523 O OH O O O OH O OH O OH

OH H H HO N N N N 519 O O OH O O O OH O OH

OH H H HO N N N N H 477 O OH O O O OH O OH H HO N N 361 O OH O O O OH O OH H HO N N 315 O OH O O O OH

H HO N N 273 O H OH O O OH H HO N N H 205 O O OH H HO N N H H 163 O OH

Table 3.4: Fragments arising from aerobactin in MSMS; fragment shown at left, m/z indicated at right. Remaining peaks in MSMS spectrum arise from internal dehydration. Peaks at m/z 142 and 128 are fragments of lysine.

121

Figure 3.5: RP-HPLC purification of cell-associated siderophores from cultures of Vibrio sp. DS40M5. The peaks at 27.5 and 29.0 minutes were collected by hand and further analyzed.

122

Figure 3.6: MSMS of the peak collected at 27.5 min from the RP-HPLC purification. Fragmentation of the principal component, m/z 759 (M+H)+, using Ar collision gas to generate daughter ions.

123

Figure 3.7: MSMS of the peak collected at 29.0 min from the RP-HPLC purification. Fragmentation of the principal component, m/z 785 (M+H)+, using Ar collision gas to generate daughter ions.

124

607 445 289 127

OH H H HO B N N N N O O OH O O O O OH O OH OH

153 315 471 633

633 471 315 153

OH H H HO C N N N N O O OH O O O OH O OH O OH

153 315 471 633

Figure 3.8: Fragmentation patterns for ochrobactins B and C. Ochrobactin B contains octanoic and (E)-2-decenoic acid moieties while ochrobactin C contains two (E)-2-decenoic acid moieties.

125 Siderophore Production by Citrobacter koseri ATCC 27156

Several strains of the soil bacterium Citrobacter koseri have been shown to produce aerobactin using NMR and mass spectrometric analysis of purified siderophores or by biological assay (Chen et al. 2000, Mokracka et al. 2004). C. koseri ATCC 27156 was selected for analysis; this strain had not previously been studied, but it was reasonable to expect that this strain would produce aerobactin as a principal siderophore since all strains previously studied produced aerobactin

(Khashe and Janda 1996, Chen et al. 2000, Mokracka et al. 2004). Aerobactin was detected in the culture supernatant of this strain in all media examined (Figure 3.9).

Aerobactin was positively identified from the 50% methanol eluent from XAD-2 by

ESI-MS and MSMS of the purified siderophore (Figure 3.10) and through comparison to published values for aerobactin (Küpper et al. submitted 2006). As was seen for Vibrio sp. DS40M5 (Figures 3.3 and 3.4, Table 3.3), MSMS analysis of aerobactin principally generates y- and b-type fragments (Roepstorff and Fohlman

1984). Fragments with m/z 361 ((M)+, composed of N-Ac, N-OH-lysine and citrate) and m/z 205 ((M+2H)+, consisting of N-Ac, N-OH-lysine) are evident in the fragmentation pattern as are the characteristic citrate-derived fragments (m/z 519 and m/z 315) (Bandu et al. 2006), loss of the acetyl moiety (m/z 523, m/z 477, m/z 273, and m/z 163), fragmentation of N-Ac, N-OH lysine (m/z 205, m/z 163, m/z 142, and m/z 128), and the citrate fragment at m/z 159.

Desferrioxamine G (DFOG) was also isolated from the supernatant of C. koseri cultures in all media tested, as established by ESI-MS and MSMS. DFOG

126 eluted from XAD-2 resin in 100% methanol. A large peak with a poorly-resolved leading peak and a trailing shoulder were evident by RP-HPLC (Figure 3.11).

Fragmentation of the primary component at m/z 619 by MSMS indicated that this was desferrioxamine G (Figure 3.12 and 3.13). Fragmentation of desferrioxamine G along the amide bonds produces the typical y- and b-type fragments according to the nomenclature of Roepstorff and Fohlman (1984). Y-type fragments are evident at m/z 519, m/z 419, m/z 319, m/z 219, and m/z 119 while m/z 501, m/z 401, m/z 301, m/z 201, and m/z 101 are b-type fragments.

No extractible cell-associated iron-binding compounds were detected in any media examined. Thus, this strain of Citrobacter was found to produce aerobactin and desferrioxamine G but not ochrobactins under the culture conditions examined.

127

Figure 3.9: RP-HPLC of 50% methanol XAD-2 fractions from purification of aerobactin from C. koseri ATCC 27156. The peak at approximately 31.5 minutes was identified as aerobactin.

128

Figure 3.10: MSMS fragmentation pattern for aerobactin produced by C. koseri ATCC 27156. See Table 3.3 for assignment of fragments.

129

Figure 3.11: RP-HPLC resolution of siderophores from 100% methanol XAD-2 fractions. The peak at approximately 31 minutes was identified as DFOG.

130

Figure 3.12: MSMS fragmentation pattern of m/z 619 collected from the RP-HPLC purification at 31 minutes using argon collision gas to generate fragments.

131

519 401 319 201 119

O OH H O O OH N N N NH HO N N 2 O O OH H O 101 219 301 419 501

Figure 3.13: Rationalization of fragmentation pattern from desferrioxamine G isolated from Citrobacter koseri ATCC 27156. Additional fragments at m/z 283 and m/z 183 arise from internal dehydrations.

132 Siderophore Production by Klebsiella pneumoniae ATCC 25306

The original aerobactin producing strain, Klebsiella pneumoniae ATCC 25306

(Aerobacter aerogenes 62-1) (Gibson and Magrath 1969) was also analyzed for production of ochrobactin siderophores. Although aerobactin was detected in each culture, no extractable cell-associated iron-binding compounds were detected. Thus,

K. pneumoniae ATCC 25306 was not found to produce ochrobactins under the culture conditions examined.

Biosynthesis of 15N-labeled Aerobactin and Feeding Experiments

Growth of C. koseri ATCC 27156 in 15N supplemented HEPES-succinate

media resulted in production of 15N-labeled aerobactin, as identified by ESI-MS and

MSMS (Figures 3.14, 3.15, 3.16, and 3.17 and Table 3.5). ESI-MS data revealed that

the principal component of the peak at 32 minutes had a mass of m/z 569, the mass

predicted for aerobactin fully labeled with 15N. The fragmentation pattern of 15N-

labeled aerobactin follows the same pattern as that of natural isotopic aerobactin but

the fragments containing the α or ε nitrogen of lysine are mass enhanced by the 15N isotope. Specifically, m/z 569 (the parent ion) and fragments at m/z 527, m/z 523, and m/z 481 are 4 mass units higher than their natural isotope counterparts.

Fragments containing only one lysine residue are enhanced by only two mass units and include m/z 363, m/z 317, m/z 275, m/z 207, and m/z 165.

15N-labeled aerobactin (2.5 mg) was added to 1 L cultures of early stationary phase Ochrobactrum sp. SP18 and incubated at room temperature for 24 to 48 hours.

133 Early stationary phase cells were used for feeding experiments because this is the

time period when a rapid increase in the amount of extractible ochrobactins normally

occurs. Exogenous 15N-labeled aerobactin added to Ochrobactrum sp. SP18 cultures

was not converted to labeled ochrobactins. Although these cultures produced

significant quantities of ochrobactin, no evidence could be found of conversion of the

labeled aerobactin (Figures 3.21 and 3.22). Fully 15N-labeled ochrobactins would be predicted to have masses of m/z 761, m/z 763, and m/z 789 for A, B, and C,

respectively. Purified ochrobactins were found to have the normal isotopic masses

m/z 757, m/z 759, and m/z 785, for ochrobactins A, B, and C, respectively. 15N

aerobactin was recovered from the cultures. No significant change in cell density was

noted during the experiment; Ochrobactrum sp. SP18 cultures generally reach

stationary phase in 24 to 48 hours with no significant changes in cell density after this

point. It is likely that the cell density of the culture, even after transfer to fresh

medium, was too dense to support cell growth.

Purified ochrobactin C, for example, demonstrated major peaks at m/z 785

(M+H)+, m/z 807 (M+Na)+, and m/z 633 (a fragment resulting from loss of one acyl

appendage; Figure 3.21). The isotopic pattern observed for these peaks matches the

natural abundance patterns seen previously for ochrobactins isolated from

Ochrobactrum sp. SP18 (Chapter 2).

There are many possible explanations for this negative result. Early stationary

phase cultures of Ochrobactrum sp. SP18, while secreting significant quantities of

ochrobactins, may not be actively synthesizing the siderophores. It is also possible

134 that Ochrobactrum sp. SP18 is not able to take up aerobactin or that sufficient quantities of ochrobactin siderophores had already been produced.

135

Figure 3.14: RP-HPLC spectrum from isolation of 15N-aerobactin produced by C. koseri ATCC 27156. The large peak at approximately 32 minutes was identified as 15N-aerobactin.

136

Figure 3.15: Mass spectrum of 15N-labeled aerobactin m/z 569. The peak at m/z 591 is the sodium adduct, m/z 551 is the loss of water, m/z 523 is fragmentation of the citryl moiety (loss of 46 mass units), and m/z 207 is the fragment consisting of N-Ac, N-OH lysine.

137

Figure 3.16: MSMS fragmentation pattern of 15N-labeled aerobactin (m/z 569).

138

527 363 207 (43)

OH H H HO N N N N O O OH O O O OH O OH O OH

(43) 207 363 527

Figure 3.17: Fragmentation pattern for 15N-labeled aerobactin.

139 OH H H HO N N N N 569 O O OH O O O OH O OH O OH

OH H H HO N N N N H 527 O OH O O O OH O OH O OH

OH H H HO N N N N 523 O O OH O O O OH O OH

OH H H HO N N N N H 481 O OH O O O OH O OH

H HO N N O 363 OH O O O OH O OH

H HO N N O 317 OH O O O OH

H HO N N O H 275 OH O O OH

H HO N N H 207 O O OH

H HO N N H H 165 O OH

Table 3.5: Fragments arising from 15N aerobactin in MSMS; fragment shown at left, m/z indicated at right. Peaks at m/z 142 and 128 are fragments of lysine. Remaining peaks in MSMS spectrum arise from internal dehydration. The fragment at m/z 159 seen is MSMS for aerobactin is not present in this spectrum or is of very low intensity. Compare to Table 3.4.

140

Figure 3.18: RP-HPLC resolution of ochrobactins isolated from 15N-aerobactin supplemented cultures of Ochrobactrum sp. SP18.

141

Figure 3.19: ESI-MS spectrum of partially purified ochrobactin C (m/z 785) from 15N aerobactin supplemented cultures. Peaks at m/z 807 and m/z 829 are sodium adducts of m/z 785. The peak at m/z 767 results from dehydration of m/z 785. Remaining peaks are fragments of ochrobactin C.

142

Figure 3.20: Expanded ESI-MS spectrum from purified ochrobactin C from 15N aerobactin supplemented cultures.

143 Attempts to Produce 15N-labeled Ochrobactins

Ochrobactrum sp. SP18 is normally grown in ASW-Fe medium containing

casamino acids. This medium cannot be used to produce 15N-labeled siderophores

15 using NH4Cl, since the large amount of casamino acids (10 g/L) provides a significant source of natural isotopic nitrogen. However, Ochrobactrum sp. SP18 was unable to grow in any of the other media reported here, including ASW-M5,

ASW+succinate, and ASW+pyruvate. There are two primary differences between these media and ASW-Fe, the lack of casamino acids and the lack of added vitamins.

It is suggested that Ochrobactrum sp. SP18 has an absolute requirement for at least one exogenous vitamin (biotin, niacin, thiamin, para-aminobenzoic acid, calcium pantothenic acid, pyridoxine hydrochloride, cyanocobalamin, riboflavin, or 4 mg folic acid). Addition of the vitamin stock, or specific vitamins, to 15N supplemented media

could be used to develop media for the production of 15N-labeled ochrobactins.

144 Discussion

Ochrobactins: Lipophilic Aerobactin-Derived Marine Siderophores

Vibrio sp. DS40M5, a marine γ-proteobacterium, was previously found to produce the hydrophilic siderophore aerobactin (Haygood et al. 1993). Two bacterial

isolates from the same water sample, Marinobacter sp. DS40M6 and Halomonas

aquamarina sp. DS40M3, have been shown to produce the amphiphilic marinobactins

and aquachelins, respectively (Martinez et al. 2000). While hydrophilic siderophores

from marine strains have been isolated, many marine siderophores are produced as

suites of amphiphiles, differing only in the nature of the acyl appendages (Figure

3.21). Following the discovery of the cell-associated ochrobactins, Vibrio sp.

DS40M5 was screened for production of cell-associated siderophores and found to

produce ochrobactins B and C. It is interesting that two marine bacterial species from

widely different bacterial clades produce the lipophilic aerobactin-derived cell- associated ochrobactins. Additionally, this result suggests that either convergent evolution or lateral gene transfer underlie production of the ochrobactin siderophores

by such widely different bacteria. It is therefore of great interest to investigate the

genes responsible for the biosynthesis of the ochrobactins in Ochrobactrum sp. SP18

and Vibrio sp. DS40M5. The sequences of these genes may elucidate the origin of

the production of the lipophilic ochrobactin siderophores in these marine isolates.

Citrobacter koseri ATCC 27156 and Klebsiella pneumoniae ATCC 25306

were selected for analysis for aerobactin-derived cell-associated siderophores as it

was logical to expect the first to produce aerobactin based on experiments with

145 C (Martinez (Martinez . Alterobactin A 40 changes 40 (Reid etal Aquachelins Alterobactin B , desferrioxamine G , alterobactins MycobacVaccae MycobactSmegATC Mycobacterium vaccae Mycobacterium M. smegmatis (Martinez et(Martinez al. 2003) Pseudoalterobactins MycobacTuber M. tuberculosis Putrebactin Norcardiaaster Nocardia asteroides Ochrobactins 97 . Modified from from Modified . (Kanoh et al. 2003) (Martinez et al.2003) et (Martinez (this work) (this , amphibactins , pseudoalterobactins AMB Alut SP25 MBIC3993 Sputrefaciens Alteromonas luteoviolacea NCIMB1893 luteoviolaceaA. Shewanella putrefaciens Halomonas H. aquamarina HTB111 , and ochrobactins Halomonas halodurans HTB111 93 S25 α SP18 100 HA Ochrobactrum S1 100 Vc Ochrobactrum sp. SP18 100 100 Vibrio cholerae Gram + (High GC) + (High Gram 98 O Vibrioferrin 100 SinoRhfredii 85 100 100 Listonella 84 (Yamamoto et al.1994)(Yamamoto et Listonella anguillarum Bisucaberin 98 γ 100 (Ito and Butler(Ito and 2005) 83 Vv 96 87 R 77 S. fredii Vt V. tubiashii , vibrioferrin β 95 D5 100 B PCOB-2 94 100 DS40M5 Sinorhizobium meliloti 1021 Sinorhizobium P. aeruginosa BLI-41 100 100 , synechobactins Vibrio R10 93 Ochrobactins 100 100 PCOB2 (Barbeau et al.2002, Bergeron et al.2003, Hickford et al.2004) Pseudoaerug DS4OM8 P. fluorescens (Winkelmann et al. 2002) Pseudofluor BurkcepCF B. cepacia LS2 B. graminis B. Aerobactin sp. PCC7002 ferm obacterarticus carbonoclasticus Pseudcorrugata Synechobactins rkholderia eto590 netohaem BurkcepPlant o n o (Martinez et al.2000) (Martinez u n A. haemolyticus Pseudomonas corrugata Marinobacter articus Burkholderia cepacia M. hydrocarbonoclasticus , petrobactins , bisucaberin Marinobacter DS40M6 sp. Acinetobacter DSM590 Cyanobacteria: Synechococcus , aquachelins Desferrioxamine G Rhodoferax fermentans Marinobactins (Martinez et al. 2000) (Martinez (Haygood et al. 1993) (Ledyard and Butler) and (Ledyard Amphibactins Marinobactins , aerobactin , putrebactin Petrobactins 1993) Figure 3.21: et al. 2001)

146 related strains and the latter was the original source of aerobactin (Gibson and

Magrath 1969). Neither the soil bacterium C. koseri ATCC 27156 nor the lung

pathogen K. pneumoniae ATCC 25306 were found to produce the cell-associated ochrobactins. It is perhaps telling that the two studied marine bacterial species produce the cell-associated ochrobactin siderophores while the two terrestrial species

produced only hydrophilic siderophores. This result suggests that, as suggested

previously (Martinez et al. 2000), the production of lipophilic siderophores such as

the ochrobactins or the amphibactins (Martinez et al. 2003) may indeed provide some

advantage to marine species for iron sequestration. Specifically, cell association

could serve to localize the siderophores near the cell, potentially increasing the likelihood of iron uptake, increasing iron availability while decreasing the cost of producing siderophores for the cell.

Do apo marine siderophores reside within the outer membrane transport proteins like the pyoverdin (FpvA), pyochelin (FptA), citrate (FecA), and ferrichrome

(FhuA) siderophores? Little is known about the putative outer membrane siderophore transport proteins from marine bacteria. Isolation of the outer membrane proteins of

Alteromonas luteoviolacea with or without added iron revealed the presence of additional outer membrane proteins during iron limited growth (Reid and Butler

1991, Martinez 2002). Resolution of these proteins by SDS-PAGE indicated that they were of approximately the same size as known outer membrane siderophore transport proteins (Reid and Butler 1991, Reid 1994, Martinez 2002). However, the reactivity of these proteins has not been investigated. It is likely that apo marine

147 siderophores will also interact with their cognate outer membrane transport proteins.

Apo loading of outer membrane transport proteins and cell-association could thus play similar roles, both promoting the uptake of ferric-siderophore complexes by increasing the concentration of siderophores near the cell.

Siderophore Production by C. koseri ATCC 27156

C. koseri ATCC 27156 was found to produce aerobactin and desferrioxamine

G, but not cell-associated siderophores. Desferrioxamine G has not previously been

identified in the culture supernatant of a Citrobacter species, although an

“unidentified hydroxamate siderophore” was reported by Mokraka et al. (2004); this

uncharacterized hydroxamate siderophore could be DFOG. The mechanisms of iron

uptake by environmental and pathogenic Citrobacterial strains is important for

controlling infection and for understanding the role of siderophores in environmental

iron cycling.

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151 Chapter Four

Photochemical Reactions of the Ochrobactin Siderophores

Introduction

Photoreactivity of Citrate Siderophores (Aerobactin, Petrobactin)

Many of the known marine siderophores contain binding groups that are

photoreactive when coordinated to iron(III) (Figure 4.1). Specifically these groups include α-hydroxy carboxylic acids such as citrate found in aerobactin (Haygood et

al. 1993), the petrobactins (Barbeau et al. 2002, Bergeron et al. 2003, Hickford et al.

2005), and the synechobactins (Ito and Butler 2005) and β-hydroxy aspartic acid

which is found in the marinobactins (Martinez et al. 2000), aquachelins (Martinez et

al. 2000), alterobactins (Reid et al. 1993), and pseudoalterobactins (Kanoh et al.

2003). Siderophores produced by organisms in the surface waters will be exposed to significant levels of sunlight irradiation which may induce photoreactions. Photolysis of Fe(III)-siderophore complexes has been shown to produce a modified, oxidized ligand and Fe(II) (Barbeau et al. 2001, Barbeau et al. 2002, Küpper et al. submitted

2006). The physiological implications of these photoreactions are not yet known, but the prevalence of photoreactive moieites in known marine bacterial siderophore structures suggests that they confer some advantage. Photoreactive iron(III) chelating groups such as the α-hydroxy carboxylic acids are an intriguing feature of marine siderophores, the biological functions of which are not yet known.

The petrobactins are citrate-derived siderophores produced by the marine γ-

proteobacterium Marinobacter hydrocarbonoclasticus (Barbeau et al. 2002,

152 abc O O O O O O O NH HN O NH OHHN NH HN OH O HO HO O OH OH OH HO O N OH HO N O O HN NH N OH HO N O O O O O O HN NH d O O O NH HN O OH R HO O OH OH HO OH HO OH

R = H or SO3 N OH HO N e O O O O H HN NH2 N OH HN O OH OH OH O H O H O H N N N COOH N N N NH2 H O H O H O HO COOH HO COOH SO H g H 3 CBA N O OH O N H O OH CONH2 f H HN O HN N NH2 OH O H O H O N N COOH HO OH NH N N N O H H H O NH2 O O OH N COOH HO COOH NH O A: R=CH NH 2 NH H HN OH 2 2 R HO NH B: R=NHC(NH)NH2 O H HN O O N O O O O O H N N N HO HO HO COOH HO O H O H O N N N OH h H N N N O H O H O H O OH OH O O OH N N HO HO HO O OH O O O O H H H N N N OH i N N N N H O H O H O H O OH OH H2NO Figure 4.1: Photoreactive siderophores isolated from marine bacteria. a) aerobactin, Vibrio sp. DS40M5 (Haygood et al. 1993) and others; b) synechobactins, Synechococcus sp. PCC7002 (Ito and Butler 2005); c) petrobactin, Marinobacter hydrocarbonoclasticus (Barbeau et al. 2002, Bergeron et al. 2003, Hickford et al. 2005); d) ochrobactins, Ochrobactrum sp. SP18 (Chapter 2); e) alterobactin B, Pseudoalteromonas luteoviolacea (Reid et al. 1993); f) alterobactin A, Pseudoalteromonas luteoviolacea (Reid et al. 1993); g) pseudoalterobactins, Pseudoalteromonas sp. KP20-4 (Kanoh et al. 2003); h) marinobactins (shown is marinobactin E), Marinobacter sp. DS40M6 (Martinez et al. 2000); and i) aquachelins (shown is aquachelin D), Halomonas aquamarina (Martinez et al. 2000).

153

O O HN HN HN HN

HN O O HN HO OH O O O hν O FeIII OH O O O HN O O Fe(II) HN HO OH

HN HN HN HN O O

Figure 4.2: Fe(III)-petrobactin photoreaction. Photoreaction of ferri-petrobactin under natural sunlight results in production of an oxidized siderophore ligand and Fe(II) (Barbeau et al. 2002, Bergeron et al. 2003). The oxidized siderophore ligand differs from the siderophore by 46 mass units as assessed by electrospray ionization mass spectrometry (ESI-MS), presumably due to the loss of CO2 and two protons.

154 Bergeron et al. 2003, Hickford et al. 2005). Irradiation of ferri-petrobactin with

ultraviolet light or natural sunlight into the ligand-to-metal charge transfer band

induces oxidation of the siderophore and formation of iron(II) (Figure 4.2) (Barbeau et al. 2002).

Aerobactin is another citrate-derived siderophore produced by several marine bacteria, including numerous Vibrio species (Haygood et al. 1993, Okujo and

Yamamoto 1994, Murakami et al. 1995, Murakami et al. 1998, Moon et al. 2004) and a halotolerant Pseudomonas species (Buyer et al 1991) as well as many terrestrial and pathogenic strains (including Enterobacter strains (Ewers et al. 2005), K. pneumoniae

(Gibson and Magrath 1969, Koczura and Kaznowski 2003), Citrobacter strains

(Khashe and Janda 1996, Chen et al. 2000, Mokracka et al. 2004), and Yersinia strains (Stuart et al. 1996)). Ferri-aerobactin undergoes a photoreaction similar to that of petrobactin when exposed to natural sunlight or to ultraviolet light from a mercury arc lamp (Figure 4.3). Photooxidation of the iron(III)-bound aerobactin ligand likely begins with a ligand-to-metal charge transfer, generating an oxidized ligand, carbon dioxide, and iron(II) (Küpper et al. 2006 submitted).

Both the petrobactin and aerobactin photoproducts are able to coordinate

Fe(III). In the case of aerobactin, the iron(III) stability constants were measured for both the natural siderophore and the photoproduct. The proton independent formation constant, log KML, for Fe(III)-aerobactin photoproduct was determined to be 28.6

while for Fe(III)-aerobactin, log KML was determined to be 27.6 (Küpper et al. 2006

submitted). The photoproduct retains the ability to coordinate ferric ion and this

155

HOOC HOOC HOOC N N O N HN HO O HN O O Fe(III) NH O O O O O hv FeIII FeIII O OH O O Fe(II) NH O O NH HO O NH O O CO2 O N O N O N HOOC HOOC HOOC

Figure 4.3: Photolysis of Fe(III)-aerobactin. Irradiation of Fe(III)-aerobactin generates an oxidized photoproduct ligand, carbon dioxide, and Fe(II). The photoproduct retains the ability to coordinate Fe(III) and the proposed structure of this complex is shown at the right (Küpper et al. 2006 submitted).

156

O HN HN

HN O O O

O FeIII

O HN O O

HN HN O

Figure 4.4: Proposed coordination of Fe(III) by the petrobactin photoproduct.

157 ligand has a very similar affinity for iron(III) relative to the natural siderophore

(Küpper et al. 2006 submitted). A coordination mode similar to the enolate tautomer of acetoacetone was proposed to account for the high binding affinity of the aerobactin photoproduct (Figure 4.3) (Küpper et al. 2006 submitted) and may be present in petrobactin as well (Figure 4.4).

Similar to other citrate-derived siderophores, ferric complexes of the aerobactin-derived ochrobactin siderophores readily undergo a photochemical reaction. The products of this reaction were evaluated by electrospray ionization mass spectrometry, tandem mass spectrometry, and continuous-flow isotope ratio mass spectrometric analysis for carbon dioxide and the quantum yield for the photoreaction was determined.

158 Materials and Methods

Production and Isolation of Ochrobactin Siderophores

Ochrobactrum sp. SP18 was grown in low-iron artificial seawater medium

(ASW-Fe) containing 10 g casamino acids, 1 g ammonium chloride, 1 g disodium glycerophosphate hydrate, 12.35 g magnesium sulfate heptahydrate, 1.45 g calcium chloride dihydrate, 0.75 g potassium chloride, and 3 mL glycerol per liter of doubly deionized water (Barnstead Nanopure II); after autoclaving, 10 mL of filter-sterilized

1 M HEPES pH 7.4, 2 mL of filter-sterilized 1 M sodium bicarbonate, and 5 mL of filter-sterilized vitamin stock were added per liter of medium. The vitamin stock solution contained 40 mg biotin, 4 mg niacin, 2 mg thiamin, 4 mg para-aminobenzoic acid, 2 mg calcium pantothenic acid, 20 mg pyridoxine hydrochloride, 2 mg cyanocobalamin, 4 mg riboflavin, and 4 mg folic acid in 200 mL of doubly deionized

water (Barnstead Nanopure II). A single colony of Ochrobactrum sp. SP18 was

grown for 12 to 24 hours at room temperature on a fresh maintenance medium plate.

Cells were resuspended in 5 mL of ASW-Fe and 2.5 mL of this suspension was transferred to a 4 L acid-washed Erlenmeyer flask containing 2 L of ASW-Fe.

Cultures were grown at room temperature with shaking at 170 rpm on an orbital

shaker for 48 to 72 hours.

Cells were pelleted by centrifugation at 4800 × g for 30 minutes at 4 °C and

cell pellets were transferred to sterile 50 mL conical tubes. The cells were extracted overnight with 90% ethanol with shaking at 120 rpm on an orbital shaker in order to

release siderophores from the cell membranes. Ethanol extracts were concentrated

159 under vacuum, applied to a C18 SepPak® cartridge (1 g sorbent, Waters Corp.), washed with 50 mL of doubly deionized water (Barnstead Nanopure II) and eluted with methanol. Fractions were hand collected and assayed for the presence of siderophores with 1:1 Fe(III)-CAS:eluent (Schwyn and Neilands 1983). Positive fractions were pooled and concentrated under vacuum to approximately 10% of the initial volume.

Ochrobactins were purified by RP-HPLC using a C4 preparative (22 mm

internal diameter (ID) × 250 mm length (L), Vydac) or a semipreparative (10 mm ID

× 250 mm L, Vydac) column using a gradient from 0.05% trifluoroacetic acid (TFA),

50% methanol, 49.95% doubly deionized water (Barnstead Nanopure II) to 99.95%

methanol, 0.05% TFA over 45 minutes. The eluent was monitored continuously at

215 nm with a Waters 2487 UV-visible detector. Fractions were hand collected and

immediately concentrated under vacuum and repeatedly rinsed with methanol to

remove excess TFA. Samples were evaluated by electrospray ionization-mass

spectrometry (ESI-MS) for purity and ultrapurified as above if necessary. Purified

samples were lyophilized and stored at -80 °C.

Extinction Coefficient Determination

Iron(III) titration experiments were completed with apo-ochrobactin C to

determine the extinction coefficient of the ferri-siderophore complex assuming one to

one iron to ligand complex formation as indicated by ESI-MS. A lyophilized sample

of apo-ochrobactin C was dissolved in 500 µL of HPLC-grade methanol and kept on

160 ice in the dark. Ferri-siderophore solutions were prepared as follows: 20 µL of apo- ochrobactin C solution (approximately 10 to 80 mM by weight) was mixed with 0 to

352 µL of 1.817 mM FeCl3 in 0.1 M HNO3 solution (0 to 0.64 mM final

concentration) in acid-washed glass vials with Teflon lid liners and incubated for 20

minutes at room temperature in the dark. Five-hundred microliters of 200 mM phosphate buffer pH 8.0 was added and the ferri-siderophore solutions were diluted to a final volume of 1 mL with 0.22 µm filtered doubly deionized water (Barnstead

Nanopure II). Solutions were then incubated at room temperature in the dark for an additional 10 minutes. Formation of the ferric-ochrobactin complex was monitored on a Cary 3E UV-visible spectrophotometer, measuring the hydroxamate ligand-to- metal charge transfer band at 407 nm. The extinction coefficient (ε) was determined from a plot of the concentration of Fe(III)-ochrobactin C versus the absorbance at 407 nm; the slope of this plot is the extinction coefficient.

Natural Sunlight Irradiation of Fe(III)-Ochrobactin C

A 60.6 µM Fe(III)-ochrobactin C solution in 50% methanol, 5 mM Tris·HCl

pH 5.0, 50 mM KCl was exposed to natural sunlight in an acid-washed quartz flask at

room temperature. One milliliter aliquots were monitored by UV-visible

spectroscopy from 600 to 200 nm every 0.5 hours with a Cary 3E UV-visible

spectrophotometer. After 3 hours, the sample was evaluated by ESI-MS to determine

the extent of reaction progress.

161 Characterization of the Photoreaction Products

Ferrous Iron Release

Production of Fe(II) during photolysis of Fe(III)-ochrobactin C was verified

by trapping of Fe(II) with 1,10-phenanthroline monohydrate. Fe(III)-ochrobactin

solution (37.5 µM) was irradiated with natural sunlight in the presence of 10 mM

1,10-phenanthroline in 5 mM phosphate buffer (pH 6.0) at room temperature.

Production of Fe(II) was assessed by monitoring the new absorbance at 510 nm,

2+ consistent with the λmax of Fe(phen)3 (Parker 1953).

Photoproduct Ligand Structure

The Fe(III)-ochrobactin C photoproduct was evaluated by ESI-MS and the

exact mass was determined with coinjection of a peptide standard for calibration.

Tandem mass spectrometry (MSMS) was used to evaluate the composition of the

apo-ochrobactin C photoproduct.

Carbon Dioxide Evolution

Fifteen hundred to sixteen hundred microliter aliquots of 60 µM Fe(III)-

ochrobactin C solutions in 1:1 water:methanol (pH 5.4) were exhaustively photolyzed

in 10 mL quartz vials sealed with silicon septa with Teflon linings under air. The

headspace was immediately assayed using a Finnigan MAT Deltaplus XL continuous- flow isotope ratio mass spectrometer interfaced to a GasBench II (Thermo Electron;

Marine Science Institute Analytical Laboratory, UCSB) (Tu et al. 2001). Production

162 of carbon dioxide was verified and quantified by comparison to a gas phase standard

and from mass and isotope ratio data.

Determination of Quantum Yield

The quantum yield for the photolysis reaction of Fe(III)-ochrobactin C was

determined for production of Fe(III)-ochrobactin C photoproduct and the

consumption of Fe(III)-ochrobactin C. Continuous wave photolysis of 1 mL aliquots

of 36 µM Fe(III)-ochrobactin C in 50% methanol, 10 mM phosphate buffer pH 6.0

was accomplished with a 200 W mercury arc lamp using a 314.5 nm interference

filter for 0 to 180 s irradiations in 1.5 mL quartz cuvettes. Samples were stirred

continuously under aerobic conditions and were protected from extraneous light. I0,

the intensity of the incident light reaching the samples, was determined with the

chemical actinometer potassium ferrioxalate according to established methods

(Hatchard and Parker 1956). Following irradiation, samples were incubated at room

temperature for a minimum of three hours to insure that all of the Fe(II) produced

would be reoxidized to Fe(III) and rechelated by the native ligand or the

photoproduct. Fe(III)-ochrobactin C photoproduct and Fe(III)-ochrobactin C were quantified by resolution on an analytical C4 RP-HPLC column (4.6 mm ID × 250 mm

L, Vydac) with a gradient from 50% methanol, 49.95% doubly deionized water

(Barnstead Nanopure II), 0.05% TFA to 99.95% methanol, 0.05% TFA over 37 minutes. Concentrations were determined by comparison to a standard curve prepared for each ligand.

163 Results

Isolation of Ochrobactins

Ochrobactins were isolated from cultures of Ochrobactrum sp. SP18.

Roughly 0.5 to 5 mg of total siderophores were isolated per liter of culture. Purified

samples were stored as a lyophilized white powder. Storage in methanol, even at -80

°C, resulted in methylation of the siderophores (Figures 4.5 to 4.7, Table 4.1).

Fragmentation of the methylated ochrobactin C derivative (m/z 799) indicates

methylation of one lysine carboxylic acid moiety. Many peaks in this fragmentation

pattern also appear in the fragmentation pattern of ochrobactin C (Figure 2.6);

specifically, the major fragments at m/z 471 (citryl N-OH, N-acyl lysine), m/z 315 (N-

OH, N-acyl lysine), and m/z 153 ((E)-2-decenoic acid) appear in both spectra. Peaks

at 14 mass units higher correspond to the methylated lysine moiety, specifically

including m/z 647, m/z 485, and m/z 329. As seen previously, fragmentation of the

citrate moiety (loss of 46 mass units) produces fragments at m/z 753, m/z 601, m/z

439, and m/z 425 corresponding to loss of CO2 and 2 protons, indicating that the site

of methylation is not on the citrate. The peaks at m/z 177 and 142 arise from

fragmentation of methylated N-OH lysine while m/z 163 and m/z 128 correspond to

the unmethylated N-OH lysine fragmentation; peaks at m/z 100 and m/z 82 are from

the fragmentation of lysine. The remaining peaks are the result of internal

dehydrations. The iron(III)-methylated ochrobactin C complex appears by ESI-MS at m/z 852, 53 mass units higher than the apo form, providing evidence that the methylation is not on the hydroxamate, but rather is most likely on the carboxylic

164 acid (Figure 4.5). A doubly methylated ochrobactin derivative is also evident at m/z

813, but is found is lower quantities than the singly methylated derivative.

165

Figure 4.5: ESI-MS spectrum of methylated ochrobactin C (m/z 799).

166

Figure 4.6: MSMS fragmentation of methylated ochrobactin C, m/z 799.

167

647 485 329 153

OH H H HO N N N N O O OH O O O O OH O OH OCH3

153 315 471 633

Figure 4.7: Fragmentation pattern of methylated ochrobactin C, m/z 799.

168 OH H H HO N N N N m/z 799 O O OH O O O O OH O OH OCH3 OH H H HO N N N N m/z 753 O O O O O OH O OH OCH3

OH H H HO N N N N m/z 647 H O OH O O O O O OH OH OCH3 OH H H HO N N N N m/z 601 H O O O O OH O OH OCH3

H HO N N m/z 485 O OH O O O O OH OCH3

OH H N N m/z 471 O O O OH O OH O OH

H HO N N m/z 439 O OH O O O OCH3

OH H N N m/z 425 O O O OH O OH H HO N N m/z 329 H O O OCH3

OH H N N m/z 315 H O O OH

Table 4.1: Fragments generated from methylated ochrobactin C, m/z 799.

169 Extinction Coefficient Determination

In solution at pH 8.0, ferri-ochrobactin C has a distinct absorption band near

407 nm corresponding to the hydroxamate to Fe(III) charge transfer (Figure 4.8). The band at 407 nm is particularly sensitive to changes in solution pH, shifting to longer

wavelength as the pH is decreased. Two significant difficulties impede the

determination of the extinction coefficients for ochrobactin siderophores: 1) the

relative insolubility of iron at near neutral pH, and 2) the insolubility of the ferri-

ochrobactins in aqueous solution. Extinction coefficients were determined in four

independent trials (Figure 4.9). Solutions with excess iron or with Fe(III)- ochrobactin concentrations above about 0.3 mM become cloudy and Fe(III)- ochrobactin begins to precipitate; if left overnight, nearly all the siderophore and any excess iron will precipitate to the bottom, forming an deeply red, oily layer.

Although solubility problems complicate the determination of the extinction coefficient for ochrobactin C, the results of the four trials agree well with each other.

The average extinction coefficient determined from the trials was 2537 ± 83 M-1cm-1.

To prevent iron precipitation and subsequent siderophore precipitation, serial dilutions of the ochrobactin C stock solution were made and then the iron titration experiment was repeated. The extinction coefficient was determined from the slope of the plot of iron(III)-ochrobactin C concentration versus the absorbance at 407 nm.

170 4

0 mM Fe(III) 0.04 mM Fe(III) 0.08 mM Fe(III) 3 0.16 mM Fe(III) 0.20 mM Fe(III)

2 Abs

0 200 300 400 500 600 700

Wavelength (nm) Figure 4.8a: UV-visible spectra of titration of apo-ochrobactin C with Fe(III).

0.30

0 mM Fe(III) 0.25 0.04 mM Fe(III) 0.08 mM Fe(III) 0.16 mM Fe(III) 0.20 0.20 mM Fe(III)

0.15 Abs

0.10

0.05

0.00 300 350 400 450 500 550 600 Wavelength (nm) Figure 4.8b: Expanded view of UV-visible spectra of titration of apo-ochrobactin C with Fe(III).

171

1.0

0.8

0.6

0.4 Abs (407 nm)Abs (407

0.2

0.0 0.00000 0.00005 0.00010 0.00015 0.00020 0.00025 0.00030 0.00035 [Fe(III)] (M)

Figure 4.9: Plot of titration of apo-ochrobactin C (solutions from four trials were approximately 0.2 mM, 0.4 mM, 0.8 mM, and 1.6 mM by weight) with standard Fe(III) solution (1.817 mM) in 100 mM phosphate buffer pH 8.0.

172 Natural Sunlight Irradiation of Fe(III)-Ochrobactin C

In natural sunlight Fe(III)-ochrobactin C undergoes rapid photooxidation of the ligand (Figure 4.10). After only 30 minutes approximately 50% of the ferrated siderophore ligand was photooxidized as evinced by the UV-visible spectrum. After

90 minutes, the reaction had reached a maximum and no further changes were evident by UV-visible spectroscopy. Mass spectrometry was used to monitor the progress of the reaction (Figure 4.11). Fe(III)-ochrobactin C is evident at m/z 838. The majority of the photoproduct was iron bound, corresponding to m/z 792 and the sodium adduct at m/z 814. A small amount of the siderophore photoproduct was in the apo form corresponding to m/z 739 and the sodium adduct at m/z 761. In this case, a small amount of the photoproduct was also found as the methylated form (m/z 806 and m/z

852).

173 3.0

2.5

2.0

1.5 AU

1.0

0.5

0.0 200 300 400 500 600 Wavelength (nm)

0.30

0.25

0.20

0.15 AU

0.10

0.05

0.00 300 350 400 450 500 550 600 Wavelength (nm) Figure 4.10: Irradiation of Fe(III)-ochrobactin C in natural sunlight. UV-visible scans were obtained at 30 minute intervals. After 90 minutes, no further photoreaction was detectable. Progress of the reaction was also monitored by ESI- MS (Figure 4.12).

174

Figure 4.11: Mass spectrum of Fe(III)-ochrobactin C during photolysis. Fe(III)-ochrobactin C (m/z 838), Fe(III)-ochrobactin C photoproduct (m/z 792), see text.

175 Characterization of the Photoreaction Products

Ferrous Iron Release

Irradiation of Fe(III)-ochrobactin C in the presence of 1,10-phenanthroline

under natural sunlight immediately resulted in an increase in absorption at 510 nm,

2+ indicating formation of the Fe(II)(phen)3 complex (Figure 4.12). Continued

irradiation led to increasing absorption at 510 nm with a maximum after 45 minutes,

-8 2+ corresponding to production of 3.79 × 10 moles of Fe(II)(phen)3 indicating

complete photoreaction of Fe(III)-ochrobactin C (3.75 × 10-8 moles before

irradiation). A dark control did not show any increase in absorption at 510 nm over

this time period.

Photoproduct Ligand Structure

The exact mass of the ferri-ochrobactin C photoproduct was determined to be

+ m/z 792.3583 for (M-3H+Fe+H) , consistent with C37H60N4O11Fe (∆-2.46). The

MSMS fragmentation of the apo-ochrobactin C photoproduct (m/z 739) indicates that

the 46 mass unit fragment is lost from the citrate moiety, likely corresponding to the

loss of CO2 and two protons (Figure 4.13, 4.14, and Table 4.2) (Bandu et al. 2006).

The fragmentation pattern of the photoproduct is much less complicated than the fragmentation pattern of the native siderophore (Figure 2.6), because the fragments resulting from the loss of 46 mass units do not appear. The fragmentation of m/z 739 results in primary y- and b-type fragments, specifically, m/z 587, m/z 425, m/z 315 and m/z 153. The fragments at m/z 436, m/z 273, and m/z 163 are internal fragments.

176 Peaks at m/z 145 and m/z 128 are fragments of N-OH lysine. The remaining peaks result from internal dehydrations.

177

0.5

t=0 min t=15 min 0.4 t=30 min t=45 min

0.3 Abs

0.2

0.1

0.0 350 400 450 500 550 600 650

Wavelength (nm)

Figure 4.12: Fe(II) trapping with 1,10-phenanthroline. UV-visible spectra of Fe(III)- ochrobactin C during photolysis in the presence of 1,10-phenanthroline, demonstrating the trapping of Fe(II) by 1,10-phenanthroline by the appearance of the 2+ peak at 510 (λmax for Fe(II)(phen)3 ). The shoulder at 417 nm corresponds to the Fe(III)-ochrobactin C complex at pH 6.0.

178

Figure 4.13: MSMS fragmentation pattern for apo-ochrobactin C photoproduct, m/z 739.

179

OH H H HO N N N N 739 O O O O O O OH O OH

OH H H HO N N N N 587 H O O O O O OH O OH

OH H H HO N N N N 435 H H O O O O OH O OH H HO N N 425 O O O O O OH

H HO N N 315 H O O OH H HO N N 273 O H O O O OH H HO N N H H 163 O OH 153 O

Table 4.2: MSMS fragmentation pattern for apo-ochrobactin C photoproduct, m/z 799.

180

587 425 315 153

OH H H HO N N N N O O O O O O OH O OH

153 315 425 587

Figure 4.14: Fragmentation pattern for apo-ochrobactin C photoproduct.

181 Carbon Dioxide Evolution

Carbon dioxide evolution during photoreaction of Fe(III)-ochrobactin C was

quantified on a Finnigan MAT Deltaplus XL continuous-flow isotope ratio mass

spectrometer interfaced to a GasBench II (Thermo Electron) (Figure 4.15). As

controls, the quantity of carbon dioxide in the helium feed line and in the headspace

above an unirradiated dark control of Fe(III)-ochrobactin C were also determined.

For each sample, the number of moles of carbon dioxide produced was calculated

from the percent carbon dioxide in the headspace, which was determined from the

area under the curve and comparison to a gas phase carbon dioxide standard (0.3%

carbon dioxide in helium, certified gas blend, Air Liquide, Inc.). The carbon dioxide

content in the helium feed line was well below the detection limit for the instrument.

The headspace of the dark control, which was prepared under air, contained

6.5 × 10-8 mol carbon dioxide. The headspace of the irradiated Fe(III)-ochrobactin C

samples, also prepared under air, contained 1.6 × 10-7 mol carbon dioxide. In this

case, 9.5 × 10-8 moles of carbon dioxide were released, equivalent to complete

photolysis of the ferri-siderophore complex (compared to 1.6 mL 60 µM Fe(III)- ochrobactin C, equivalent to 9.6 × 10-8 moles). Detection of carbon dioxide in the

headspace following irradiation of ferric-ochrobactin solutions confirms the evolution

of carbon dioxide during photolysis. It was not possible to calculate a quantum yield

from this experiment because the incident light intensity (I0) is not known for the bulk

irradiation setup.

Thus, photoreactions of the Fe(III)-ochrobactins produce the oxidized ligand,

182 Fe(II), and carbon dioxide (Figure 4.16). In aerobic solutions, Fe(II) is rapidly reoxidized to Fe(III) and can be chelated by the photoproduct or by the native siderophore.

183

Time (seconds)

a) Dark Control Peak # Time (s) Area (Vs) 4 214 1.740 5 284 1.528 6 354 1.337 7 423 1.168 8 493 1.022

b) Irradiated Fe(III)-ochrobactin C Peak # Time (s) Area (Vs) 4 214 4.237 5 284 3.670 6 354 3.184 7 423 2.777 8 493 2.416

Figure 4.15: CO2 detection. a) CO2 detection in the headspace above the dark control; b) CO2 detection in the headspace of photolyzed Fe(III)-ochrobactin C. Arrows indicate the peaks for CO2 from sample injections. Sequential numbers indicate the injection number; other numbers indicate time of injection. Peaks 1, 2, and 3 are assessment injections of 100% CO2 for validation (Air Liquide).

184

HOOC R1 HOOC R1 HOOC R1 N N O N HN HO O Fe(III) HN O O NH O O O O O hv FeIII FeIII O OH O O Fe(II) NH O O NH HO O NH O O CO2 O N O N O N HOOC R2 HOOC R2 HOOC R2

Figure 4.16: Photoreaction scheme of Fe(III)-ochrobactin C. Photolysis into the ligand-to-metal charge transfer band (314 nm) results in reduction of Fe(III) to Fe(II) and oxidation of the ligand. In aerobic solutions, Fe(II) is rapidly re-oxidized to Fe(III) and can then be bound by the photoproduct or by the native siderophore. R1=- CH=CH(CH2)6CH3; A: R2=-CH=CH(CH2)4CH3, B: R2=-(CH2)6CH3, or C: R2=- CH=CH(CH2)6CH3.

185 Quantum Yield

The quantum yield (Ф) for the formation of photoproduct during the

photolysis of Fe(III)-ochrobactin C was determined with solutions of 36 µM ferri-

siderophore in 50% methanol at pH 6.0 by measuring the change in concentration of

the Fe(III)-ochrobactin C complex (Fe(Oc)) and the appearance of the ferri- photoproduct (Fe(PPC)) by RP-HPLC (Figure 4.17). Samples were irradiated at 314 nm using a mercury arc lamp with a 314 nm interference filter. Vigorous stirring under air ensured that samples were well mixed during photolysis as well as allowing for the oxidation of produced Fe(II) to Fe(III), to insure that the ochrobactin C and photoproduct C were Fe(III) bound. The incident light intensity (I0) was

independently determined for each experiment using potassium ferrioxalate. I0 was

calculated to be 1.0215 × 1015 ± 0.004 × 1015 photons/s based on three experiments

with three replicates each.

The quantum yield (Ф) was calculated from Eq. 1 (Tables 4.3 to 4.8):

()∆ [Fe(Oc) or Fe(PPC)] (volume)(6.023×1023 photons/mole) Φ = −A314 Eq. 1 ()1−10 ()(I0 irradiation time ) where ∆[Fe(Oc) or Fe(PPC)] is the change in concentration of Fe(III)-ochrobactin C or Fe(III)-ochrobactin C photoproduct (M), volume is the volume irradiated (L), (1-

10-A314) accounts for the amount of light absorbed by the sample, A314 is the average absorbance of each sample at 314 nm, and I0 is the incident light intensity (1.0215 ×

1015 photons/second.

The initial quantum yield for 30 second irradiation was determined to be

186 0.038 ± 0.001 from production of Fe(III)-ochrobactin C photoproduct (Figure 4.19) and 0.036 ± 0.0003 from the consumption of Fe(III)-ochrobactin C (Figure 4.18).

187 t=0s

t=30s

t=60s

t=90s

t=120s

t=150s

t=180s

Figure 4.17: RP-HPLC resolution of photoreaction, illustrating method of concentration determination. The peak at approximately 26 minutes is Fe(III)- ochrobactin C photoproduct; the peak at approximately 28 minutes is Fe(III)- ochrobactin C.

188 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 1, consumption of Fe(III)-ochrobactin C) Trial 1 Fe(III)-Ochro C Change Sample Peak Area [FeOc] (M) [FeOc] (M) Ave A314 Ф (FeOc) 0 10485455 3.495 x10-5 9 x10-8 0.217105 0 10511155 3.504 x10-5 4 x10-9 0.217105 0 10540057 3.513 x10-5 9 x10-8 0.217105

30 10292763 3.431 x10-5 7.3 x10-7 0.216713 0.0366 30 10290180 3.430 x10-5 7.4 x10-7 0.217613 0.0369 30 10300317 3.433 x10-5 7.1 x10-7 0.216988 0.0353

60 10120055 3.373 x10-5 1.31 x10-6 0.221405 0.0322 60 10120867 3.374 x10-5 1.30 x10-6 0.224549 0.0318 60 10121032 3.374 x10-5 1.30 x10-6 0.220200 0.0322

90 9967307 3.322 x10-5 1.82 x10-6 0.218720 0.0301 90 9963784 3.321 x10-5 1.83 x10-6 0.219742 0.0302 90 9930588 3.310 x10-5 1.94 x10-6 0.222100 0.0317

120 9859077 3.286 x10-5 2.18 x10-6 0.220848 0.0268 120 9843889 3.281 x10-5 2.23 x10-6 0.218804 0.0277 120 9803158 3.268 x10-5 2.36 x10-6 0.247940 0.0267

150 9752171 3.251 x10-5 2.53 x10-6 0.225662 0.0246 150 9732827 3.244 x10-5 2.60 x10-6 0.222466 0.0255 150 9754723 3.252 x10-5 2.52 x10-6 0.231842 0.0240

180 9601674 3.201 x10-5 3.04 x10-6 0.253220 0.0225 180 9616465 3.205 x10-5 2.99 x10-6 0.221748 0.0245 180 9603156 3.201 x10-5 3.03 x10-6 0.221474 0.0248 Table 4.3: Ф from trial 1, calculated from consumption of Fe(III)-ochrobactin C, peak at approximately 28 minutes (Figure 4.17). 36 µM Fe(III)-ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

189 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 2, consumption of Fe(III)-ochrobactin C) Trial 2 Fe(III)-Ochro C Change Sample Peak Area [FeOc] (M) [FeOc] (M) Ave A314 Ф (FeOc) 0 10648405 3.549 x10-5 8 x10-8 0.219055 0 10618912 3.540 x10-5 2 x10-8 0.217377 0 10606692 3.536 x10-5 6 x10-8 0.219635

30 10391291 3.464 x10-5 7.8 x10-7 0.210671 0.0398 30 10397194 3.466 x10-5 7.6 x10-7 0.261795 0.0329 30 10423249 3.474 x10-5 6.7 x10-7 0.210381 0.0344

60 10259056 3.420 x10-5 1.22 x10-6 0.214112 0.0308 60 10252007 3.417 x10-5 1.24 x10-6 0.213738 0.0314 60 10239326 3.413 x10-5 1.28 x10-6 0.245751 0.0292

90 10106177 3.369 x10-5 1.73 x10-6 0.209801 0.0296 90 10107477 3.369 x10-5 1.72 x10-6 0.213379 0.0291 90 10110488 3.370 x10-5 1.71 x10-6 0.216156 0.0286

120 10035959 3.345 x10-5 1.96 x10-6 0.216355 0.0246 120 10039471 3.346 x10-5 1.95 x10-6 0.218582 0.0242 120 10028215 3.343 x10-5 1.99 x10-6 0.215981 0.0249

150 9915606 3.305 x10-5 2.36 x10-6 0.21492 0.0238 150 9877914 3.293 x10-5 2.49 x10-6 0.216896 0.0249 150 9863799 3.288 x10-5 2.54 x10-6 0.21975 0.0251

180 9702894 3.234 x10-5 3.07 x10-6 0.214715 0.0258 180 9703737 3.235 x10-5 3.07 x10-6 0.226609 0.0247 180 9777963 3.259 x10-5 2.82 x10-6 0.216866 0.0235 Table 4.4: Ф from trial 2, calculated from consumption of Fe(III)-ochrobactin C, peak at approximately 28 minutes (Figure 4.17). 36 µM Fe(III)-ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

190 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 3, consumption of Fe(III)-ochrobactin C) Trial 3 Fe(III)-Ochro C Change Ave Sample Peak Area [FeOc] (M) [FeOc] (M) A314 Ф (FeOc) 0 10637190 3.546 x10-5 9 x10-8 0.210671 0 10663261 3.554 x10-5 4 x10-9 0.210381 0 10692582 3.564 x10-5 9 x10-8 0.214111

30 10441709 3.481 x10-5 7.4 x10-7 0.213737 0.0375 30 10439088 3.480 x10-5 7.5 x10-7 0.245751 0.0341 30 10449372 3.483 x10-5 7.2 x10-7 0.209801 0.0368

60 10266502 3.422 x10-5 1.33 x10-6 0.230903 0.0316 60 10267326 3.422 x10-5 1.32 x10-6 0.214615 0.0334 60 10267493 3.423 x10-5 1.32 x10-6 0.215759 0.0332

90 10111543 3.371 x10-5 1.84 x10-6 0.217376 0.0307 90 10107969 3.369 x10-5 1.85 x10-6 0.217072 0.0309 90 10074293 3.358 x10-5 1.97 x10-6 0.215881 0.0329

120 10001747 3.334 x10-5 2.21 x10-6 0.216438 0.0277 120 9986340 3.329 x10-5 2.26 x10-6 0.216972 0.0282 120 9945019 3.315 x10-5 2.40 x10-6 0.231888 0.0285

150 9893294 3.298 x10-5 2.57 x10-6 0.233521 0.0243 150 9873670 3.291 x10-5 2.64 x10-6 0.222435 0.0258 150 9895883 3.299 x10-5 2.56 x10-6 0.217636 0.0255

180 9740620 3.247 x10-5 3.08 x10-6 0.218506 0.0255 180 9755624 3.252 x10-5 3.03 x10-6 0.219162 0.0250 180 9742123 3.247 x10-5 3.07 x10-6 0.214981 0.0258 Table 4.5: Ф from trial 3, calculated from consumption of Fe(III)-ochrobactin C, peak at approximately 28 minutes (Figure 4.17). 36 µM Fe(III)-ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

191 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 1, production of Fe(III)-ochrobactin C photoproduct) Trial 1 Fe(III)-PPC Change Ave Sample Peak Area [FePPC] (M) A314 Ф (FePPC) 0 0 0 0.217105 0 0 0 0.217105 0 0 0 0.2171050

30 219459 7.315 x10-7 0.2167130 0.0370 30 222042 7.401 x10-7 0.2176130 0.0372 30 211905 7.064 x10-7 0.2169880 0.0350

60 392167 1.307 x10-6 0.2214050 0.0321 60 391355 1.305 x10-6 0.2245485 0.0317 60 391190 1.304 x10-6 0.2201995 0.0322

90 544915 1.816 x10-6 0.2187195 0.0305 90 548438 1.828 x10-6 0.2197415 0.0301 90 581634 1.939 x10-6 0.2220995 0.0317

120 653145 2.177 x10-6 0.2208480 0.0271 120 668333 2.228 x10-6 0.2188035 0.0276 120 709064 2.364 x10-6 0.2479400 0.0286

150 760051 2.534 x10-6 0.2256620 0.0245 150 779395 2.598 x10-6 0.2224655 0.0254 150 757499 2.525 x10-6 0.2318420 0.0239

180 910548 3.035 x10-6 0.2532195 0.0230 180 895757 2.986 x10-6 0.2217480 0.0244 180 909066 3.030 x10-6 0.2214735 0.0251 Table 4.6: Ф from trial 1, determined from production of Fe(III)-ochrobactin C photoproduct, peak at approximately 26 minutes (Figure 4.17). 36 µM Fe(III)- ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

192 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 2, production of Fe(III)-ochrobactin C photoproduct) Trial 2 Fe(III)-PPC Change Ave Sample Peak Area [FePPC] (M) A314 Ф (FePPC) 0 0 0 0.2190550 0 0 0 0.2173770 0 0 0 0.2196350

30 233379 8.335 x10-7 0.2106705 0.0426 30 227476 8.124 x10-7 0.2617950 0.0353 30 201421 7.194 x10-7 0.2103805 0.0368

60 365614 1.306 x10-6 0.2141115 0.0330 60 372663 1.331 x10-6 0.2137375 0.0336 60 385344 1.376 x10-6 0.2457505 0.0313

90 518493 1.852 x10-6 0.2098005 0.0317 90 517193 1.847 x10-6 0.2133790 0.0312 90 514182 1.836 x10-6 0.2161560 0.0307

120 588711 2.103 x10-6 0.2163545 0.0263 120 585199 2.090 x10-6 0.2185820 0.0260 120 596455 2.130 x10-6 0.2159805 0.0267

150 709064 2.532 x10-6 0.2149200 0.0255 150 746756 2.667 x10-6 0.2168960 0.0267 150 760871 2.717 x10-6 0.2197495 0.0269

180 921776 3.292 x10-6 0.2147145 0.0276 180 920933 3.289 x10-6 0.2266085 0.0265 180 846707 3.024 x10-6 0.2168655 0.0252 Table 4.7: Ф from trial 2, determined from production of Fe(III)-ochrobactin C photoproduct, peak at approximately 26 minutes (Figure 4.17). 36 µM Fe(III)- ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

193 Calculation of Quantum Yield for Fe(III)-ochrobactin C photolysis at 314 nm (Trial 3, production of Fe(III)-ochrobactin C photoproduct) Trial 3 Fe(III)-PPC Change Ave Sample Peak Area [FePPC] (M) A314 Ф (FePPC) 0 0 0 0.2106705 0 0 0 0.210381 0 0 0 0.214111

30 222635 7.951 x10-7 0.213737 0.0402 30 225256 8.045 x10-7 0.2457505 0.0366 30 214972 7.678 x10-7 0.209801 0.0394

60 397842 1.421 x10-6 0.230903 0.0339 60 397018 1.418 x10-6 0.214615 0.0357 60 396851 1.417 x10-6 0.215759 0.0356

90 552801 1.974 x10-6 0.217376 0.0328 90 556375 1.987 x10-6 0.217072 0.0331 90 590051 2.107 x10-6 0.215881 0.0352

120 662597 2.366 x10-6 0.216438 0.0296 120 678004 2.421 x10-6 0.216972 0.0303 120 719325 2.569 x10-6 0.231888 0.0305

150 771050 2.754 x10-6 0.233521 0.0260 150 790674 2.824 x10-6 0.222435 0.0277 150 768461 2.745 x10-6 0.217636 0.0274

180 923724 3.299 x10-6 0.218506 0.0273 180 908720 3.245 x10-6 0.219162 0.0268 180 922221 3.294 x10-6 0.214981 0.0276 Table 4.8: Ф from trial 3, determined by production of Fe(III)-ochrobactin C photoproduct, peak at approximately 26 minutes (Figure 4.17). 36 µM Fe(III)- ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0.

194

0.045

0.040

0.035 Φ

0.030

0.025

0.020 0 50 100 150 200 Irradiation time (s)

Figure 4.18: Plot of Φ determined for consumption of 36 µM Fe(III)-ochrobactin C in 50% methanol at pH 6.0; irradiation at 314 nm. Symbols: ● trial 1, ○ trial 2, ▼ trial 3. Trial 1 was 0.036 ± 0.001; trial 2 was 0.036 ± 0.004; trial 3 was 0.036 ± 0.001.

195

0.045

0.040

0.035 Φ

0.030

0.025

0.020 0 50 100 150 200 Irradiation time (s)

Figure 4.19: Plot of Φ determined for production of Fe(III)-ochrobactin C photoproduct in 50% methanol at pH 6.0; irradiation at 314.5 nm. Symbols: ● trial 1, ○ trial 2, ▼ trial 3. Trial 1 was 0.036 ± 0.001; trial 2 was 0.038 ± 0.004; trial 3 was 0.039 ± 0.002.

196 Discussion

The photoreaction of Fe(III)-ochrobactin siderophores in natural sunlight or under ultraviolet irradiation with a mercury arc lamp at 314 nm produces an oxidized ligand, Fe(II), and carbon dioxide. The initial quantum yields for 30 second irradiation were determined for the photoreaction of Fe(III)-ochrobactin C (under conditions of 36 µM Fe(III)-ochrobactin C, 50% methanol, 10 mM phosphate buffer pH 6.0; Ф = 0.038 ± 0.001 as measured by production of Fe(III)-ochrobactin C photoproduct; Ф = 0.036 ± 0.0003 as measured by consumption of Fe(III)- ochrobactin C). These are the first quantum yields reported for a photoreactive marine siderophore.

Quantum yields have been determined previously for ferric citrate complexes in solution; for irradiation of 0.1 mM citrate, 0.1 mM Fe(III) pH 6.0 at 436 nm the quantum yield ranged from 0.28 (pH 4.0) to 0.21 (pH 6.0) as measured by Fe(II) production (Faust and Zepp 1993). The Fe(III) speciation was predicted to be 93%

Fe(OH)(citrate)-, 6.6% Fe(citrate)0 in the pH range from 4.0 to 6.0 (Faust and Zepp

1993). Further investigation of the photoreactivity of ferric citrate complexes suggests that the formation of a µ-oxo bridged diferric dicitrate complex is the most likely species to account for pH dependence of the photolysis reactions at low pH (2.7 to 4.0) (Abrahamson et al. 1994). Abrahamson et al. (1994) determined the quantum yield for irradiation of ferric citrate solutions at 366 nm to range from 0.28 (pH 2.7) to 0.45 (pH 4.0) for 1.5 mM citrate, 0.3 mM Fe(III). A µ-oxo bridged diferric dicitrate dimer was proposed to account for the observed pH dependence of the

197 photoreaction of iron(III) citrate complexes (Abrahamson et al. 1994).

The quantum yields determined for ferric citrate complexes were nearly an order of magnitude higher than that determined for Fe(III)-ochrobactin C (Faust and

Zepp 1993, Abrahamson et al. 1994). One significant difference may be the speciation of Fe(III)-ochrobactin compared to the ferric citrate complexes. The chelation of iron(III) by the ochrobactin siderophores involves the photostable hydroxamate moieties and the α-hydroxy carboxylic acid moiety of the citrate backbone. The photostable hydroxamate binding groups may serve to stabilize the ferric ion in the complex, thus decreasing the photolability of the complex relative to the ferric citrate complexes. Thus, while the quantum yield determined for the photoreaction of Fe(III)-ochrobactin C is less than that for ferric citrate, the reaction mechanism for the ochrobactins may be significantly different than that of ferric citrate.

Photoreactivity of marine siderophores is an intriguing feature of these ligands. Given that marine microorganisms producing known marine siderophores live in the surface waters of the oceans and will be exposed to significant amounts of sunlight, the purpose of these photoreactive binding groups (such as citrate and β- hydroxy aspartate) is brought into question. The α-hydroxy carboxylate binding moieties are very common in the marine siderophores that have been characterized

(Haygood et al. 1993, Reid et al. 1993, Martinez et al 2000, Barbeau et al. 2002,

Bergeron et al. 2003, Kanoh et al. 2003, Ito and Butler 2005). Is it merely a coincidence, or does the photoreactivity serve a specific function? It has been

198 proposed that production of Fe(II) by photolysis of these ligands could increase the bioavailability of iron (Barbeau et al. 2001, Barbeau et al. 2002). However, perhaps a more direct effect underlies the presence of these moieties. Could the production of carbon dioxide during photolysis or the structure of the photooxidized ligand serve as a signaling molecule? Such a mechanism could communicate the presence of light

(i.e. open water growth conditions). The recent identification of outer membrane siderophore transport proteins as signaling molecules suggests exciting possibilities for the functions of photoreactive siderophores.

199 References

Barbeau, K., Rue, E.L., Bruland, K.W., and Butler, A. (2001) Nature 413, 409 – 413.

Barbeau, K., Zhang, G., Live, D.H., and Butler, A. (2002) Journal of the American Chemical Society 124, 378 – 379.

Bergeron, R.J., Huang, G., Smith, R.E., Bharti, N., McManis, J.S., and Butler, A. (2003) Tetrahedron 59, 2007 – 2014.

Buyer, J.S., de Lorenzo, V., and Neilands, J.B. (1991) Applied and Environmental Microbiology 57, 2246 – 2250.

Faust, B.C. and Zepp, R.G. (1993) Environmental Science and Technology 27, 2517 – 2522.

Hatchard, C.G. and Parker, C.A. (1956) Proceedings of the Royal Society, Series A 235, 518 – 536.

Haygood, M.G., Holt, P.D., and Butler, A. (1993) Limnology and Oceanography 38, 1091 – 1097.

Hickford, S.J.H., Küpper, F.C., Zhang, G.P., Carrano, C.J., Blunt, J.W., and Butler, A. (2004) Journal of Natural Products 67, 1897 – 1899.

Ito, Y. and Butler, A. (2005) Limnology and Oceanography 50, 1918 – 1923.

Kanoh, K., Kamino, K., Leleo, G., Adachi, K., and Shizuri, Y. (2003) Journal of Antibiotics 56, 871 – 875.

Küpper, F.E., Carrano, C.J., Kuhn, J.-U., and Butler, A. (submitted) Inorganic Chemistry.

Martinez, J.S., Zhang, G.P. Holt, P.D., Jung, H., Carrano, C.J., Haygood, M.G., and Butler, A. (2000) Science 287, 1245 – 1247.

Moon, Y.-H., Tanabe, T., Funahashi, T., Shiuchi, K., Nakao, H., and Yamamoto, S. (2004) Microbiology and Immunology 48, 389 – 398.

Murakami, K., Ohta, S., Fuse, H., Takimura, O., and Kamimura, K. (1995) Microbios 84, 231 – 238.

Murakami, K., Fuse, H., Takimura, O., Kamimura, K., and Yamaoka, Y. (1998) Journal of Marine Biotechnology 6, 76 – 79.

200

Okujo, N. and Yamamoto, S. (1994) FEMS Microbiology Letters 118, 187 – 192.

Parker, C (1953) Proceedings of the Royal Society of London, Series A 220, 104 – 116.

Reid, R.T., Live, D.H., Faulkner, D.J., and Butler, A. (1993) Nature 366, 455 – 458.

Tu, K.P., Brooks, P.D., and Dawson, T.E. (2001) Rapid Communications in Mass Spectrometry 15, 952 – 956.

201 Chapter Five

Membrane Partition Coefficient Determination with the Ochrobactin Siderophores

Introduction

Amphiphilicity of Marine Siderophores

Siderophore Structures

While the study of siderophores produced by marine bacteria is relatively

new, the structures of a modest number of marine siderophores have been determined.

Two structural themes have emerged from the known marine siderophore structures:

(1) the presence of photoreactive moieties (see Chapter 4), and (2) the production of suites of amphiphilic siderophores composed of a peptidic (or peptide-like) headgroup and one of a series of acyl appendages. The appendages vary by length, degree of unsaturation, and substitution (Figure 5.1). Amphiphilic siderophores

constitute a much higher percentage of known marine siderophores than known

terrestrial siderophores.

Cell-Associated Siderophores

The marinobactins (Martinez et al. 2000), aquachelins (Martinez et al. 2000),

and synechobactins (Ito and Butler 2005) are generally found in the culture

supernatants under laboratory conditions. A small subset of amphiphilic siderophores

is cell-associated. The amphibactins (Martinez et al. 2003) and ochrobactins

(Chapter 2) are cell-associated siderophores from marine bacteria which must be

202

O O O O a b OH O H N N N N N HO HO HO O HO HO HO OH H O H O H O H O H O R N N N OH N N N OH H N N N R N N N N O H O H O H O H O H O H O H O OH OH OH OH O R = R = H2NO O E D O O O D2 O C O D1 O B C O A O B O A d A O O O c O OH B N N O HO HO O N N C H O H O H R N N OH N N OH OH H O H O OH OH OH OH H O N N N O R = O O O I O OH O H OHO G e O OH O O F N N O C O H OH E OH O B O OH D A OH O H O OH C N N O OH O O B

Figure 5.1: Amphiphilic marine siderophores. a) marinobactins produced by Marinobacter sp. DS40M6 (Martinez et al. 2000); b) aquachelins produced by Halomonas aquamarina (Martinez et al. 2000); c) amphibactins produced by Vibrio sp. R-10 (Martinez et al. 2003); d) ochrobactins produced by Ochrobactrum sp. SP18 (Chapter 2); and e) synechobactins produced by Synechococcus sp. PCC7002 (Ito and Butler 2005).

203 extracted from the cell pellets during isolation. A few cell-associated siderophores

have been structurally characterized from terrestrial and pathogenic bacterial isolates,

namely, the mycobactins (Gobin et al. 1996, Ratledge and Dale 1999) and the

structurally related formobactins (Murakami et al. 1996), nocobactins (Ratledge and

Patel 1976), and amamistatins (Suenaga et al. 1999, Kokubo et al. 2000).

Mycobacteria produce both secreted siderophores, the carboxymycobactins (in

pathogenic strains) or the exochelins (in saprophytic strains), and both pathogenic and

saprophytic strains produce the cell-associated, lipophilic mycobactins (Ratledge and

Dale 1999). Recent evidence indicates that the function of the extremely lipophilic

mycobactins and related structures is short-term iron storage in the cellular envelope

when the iron uptake pathways are saturated (Ratledge 2004). The secreted

exochelins chelate iron(III) and bring it back to the cell; the ferric-exochelin complex

is recognized at the cell surface by a receptor protein (Rec), transferred across the

envelope, possibly by FxuD, and then transported into the cell by FxuA, FxuB, and

FxuC (Ratledge and Dale 1999, Ratledge 2004). The carboxymycobactins also

chelate iron(III) from the environment and bring it back to the cell where iron(III) is

reduced to iron(II) and shuttled across the membrane, possibly complexed to salicylic

acid (Ratledge 2004). Excess iron provided by either the exochelins or the carboxymycobactins is transferred to the mycobactins for temporary storage

(Ratledge 2004).

The functions of the lipophilic mycobacterial siderophores seem to be different from the amphibactins and the ochrobactins. On some occasions, the

204 amphibactins were found to be secreted, and not cell-associated, likely due to slight alterations in the culture medium (Martinez et al. 2003). The ochrobactins, although lipophilic, are structurally very similar to aerobactin which is transported across the outer membrane through a specific outer membrane protein, IutA (Wooldridge et al.

1992, Murakami et al. 2000), which might suggest that the ochrobactins could be taken up in a similar manner. Thus, the amphibactins and ochrobactins, while found attached to the cell surface like the mycobactins, seem to play a more traditional role as siderophores, scavenging iron from the environment and directly transporting it into the cell.

Membrane Partitioning Studies

Suites of amphiphilic siderophores composed of a peptidic head group and one of a series of acyl appendages have been isolated from a variety of marine bacteria. Because of their surface reactivity and amphiphilicity, the membrane affinities of the marinobactin and amphibactin siderophores were investigated using large, unilamellar 1,2-dimyristoyl-sn-glycero-3-phosphocholine vesicles (DMPC) (Xu et al. 2002, Martinez et al. 2003) (see also Chapter One). Membrane affinity measurements of these amphiphiles permitted comparison of these molecules to other amphiphilic or lipophilic compounds and also provided an estimate of the interaction of these molecules with bacterial cell membranes. Overall, these siderophores partitioned to an extent similar to membrane-active detergents such as octyl glucoside and Triton X-100 (Inoue 1996). For the marinobactins, a reduction by two methylene

205 carbons or the introduction of a cis double bond in the fatty acid appendage resulted in a one order of magnitude reduction in partition coefficient (Xu et al. 2002). The amphibactins have longer acyl appendages and a smaller headgroup than the marinobactins (4 amino acid residues for amphibactins versus 6 amino acid residues in the marinobactins) and generally partitioned more than the marinobactins. For example, amphibactin D and marinobactin C each have a saturated C14 acyl

appendage. The membrane partition coefficient for amphibactin D (1915 M-1

(Martinez et al. 2003)) was significantly higher than for marinobactin C (195 M-1 (Xu

et al. 2002)). This trend was not extended to amphibactin H (3784 M-1 (Martinez et

al. 2003)) and marinobactin E (5818 M-1 (Xu et al. 2002)) each of which have a

saturated C16 acyl appendage, suggesting that a balance between the length of the acyl

appendage and the size of the headgroup collectively determine the extent of

interaction with the membrane.

It has been predicted that the amphiphilicity of marine siderophores serves to

localize the siderophores near the cells, such that a concentration gradient of siderophores radiating out from the cell is created (Martinez et al. 2000). Limiting diffusion of the siderophores away from the cell could decrease loss to the environment. Constraining siderophores near the cell could also help to prevent other cells from “stealing” them, a phenomenon known as siderophore piracy. Formation

of a concentration gradient could therefore serve to increase the availability of iron to

the cells.

206 The membrane partition coefficients for apo-, Fe(III)-, and Fe(III)- photoproduct of ochrobactins B and C, which each have two acyl appendages, were determined using large unilamellar DMPC vesicles.

207 Materials and Methods

Purification and Preparation of Siderophores

Ochrobactins B and C were isolated from cultures of Ochrobactrum sp. SP18.

For siderophore production, Ochrobactrum sp. SP18 was grown in low-iron artificial seawater medium (ASG-Fe) containing 10 g casamino acids, 1 g ammonium chloride,

1 g disodium glycerophosphate hydrate, 12.35 g magnesium sulfate heptahydrate,

1.45 g calcium chloride dihydrate, 0.75 g potassium chloride, and 3 mL glycerol per liter of doubly deionized water (Barnstead Nanopure II) ; after autoclaving, 10 mL of filter-sterilized 1 M HEPES pH 7.4, 2 mL of filter-sterilized 1 M sodium bicarbonate, and 5 mL of filter-sterilized vitamin stock were added per liter of medium. The vitamin stock solution contained 40 mg biotin, 4 mg niacin, 2 mg thiamin, 4 mg para- aminobenzoic acid, 2 mg calcium pantothenic acid, 20 mg pyridoxine hydrochloride,

2 mg cyanocobalamin, 4 mg riboflavin, and 4 mg folic acid in 200 mL of doubly deionized water (Barnstead Nanopure II) . A single colony of Ochrobactrum sp.

SP18 was grown for 12 to 24 hours at room temperature on a fresh maintenance medium plate. Cells were resuspended in 5 mL of ASG-Fe and 2.5 mL of this suspension transferred to a 4 L acid-washed Erlenmeyer flask containing 2 L ASG-

Fe. Cultures were grown at room temperature with shaking at 170 rpm on an orbital shaker for 48 to 72 hours.

Cells were pelleted by centrifugation at 4800 × g for 30 minutes at 4 °C. The cell pellets from approximately 330 mL of culture supernatant (from a 2 L culture) were transferred to sterile 50 mL conical tubes. The cells were extracted overnight

208 with 90% ethanol to remove siderophores from the cell membranes with shaking at

120 rpm on an orbital shaker. Ethanol extracts were concentrated under vacuum,

applied to a C18 SepPak® cartridge (1 g sorbent, Waters, Corp.), washed with 50 mL

of doubly deionized water (Barnstead Nanopure II) and eluted with methanol.

Fractions were hand collected and assayed for the presence of siderophores with 1:1

Fe(III)-CAS solution assay:eluent (Schwyn and Neilands 1983). Positive fractions

were pooled and concentrated under vacuum to approximately 10% of the initial

volume.

Ochrobactins B and C were purified by RP-HPLC using a C4 preparative (22 mm internal diameter (ID) × 250 mm length (L), Vydac) or a semipreparative (10 mm

ID ×250 mm L, Vydac) column using a gradient from 0.05% trifluoroacetic acid

(TFA), 50% methanol, 49.95% doubly deionized water (Barnstead Nanopure II) to

0.05% TFA in methanol over 45 minutes. The eluent was monitored continuously at

215 nm with a Waters 2487 UV-visible detector. Fractions were hand collected and immediately concentrated under vacuum and repeatedly rinsed with methanol to remove excess TFA. Samples were evaluated by electrospray ionization-mass spectrometry (ESI-MS) for purity and ultrapurified as above if necessary. Purified samples were lyophilized and stored at -80 °C. Very little ochrobactin A is produced

by the bacterium and was not purified for membrane partitioning experiments.

Concentrations of ochrobactin solutions were determined by titration with

standardized iron(III) solutions. Purified siderophores were dissolved in methanol

and stored on ice in the dark. Twenty microliters of siderophore solution was

209 incubated with 0 to 0.64 mM Fe(III) from 1.817 mM FeCl3 in 10% HNO3 (Aldrich)

in acid-washed 1 dram glass vials with teflon lid liners for 15 minutes at room

temperature in the dark. Five-hundred microliters of 200 mM phosphate buffer pH

8.0 was added and the final solutions diluted to 1 mL with doubly deionized water

(Barnstead Nanopure II). Solutions were equilibrated for an additional 10 minutes in

the dark at room temperature and were then assayed on a Cary 3E UV-vis spectrophotometer. Formation of the iron(III)-ochrobactin complex was monitored by measuring the absorbance at 407 nm, corresponding to the hydroxamate to

iron(III) charge transfer.

To prepare ferri-ochrobactins, lyophilized samples were dissolved in a small

amount of methanol and then diluted with 50% methanol/100 mM phosphate buffer

pH 8.0. FeCl3 in 10% HNO3 (1.817 mM) was added dropwise with stirring in the

dark to the siderophore solution, forming a deeply red complex. Solutions were

stirred at room temperature overnight and then assayed by ESI-MS. Ferri-

ochrobactins were purified by C18 SepPak® cartridge (1g sorbent, Waters, Corp.) as

above and ultrapurified by RP-HPLC as above to insure that samples contained only

the ferric complexes. Samples were immediately concentrated under vacuum and

rinsed repeatedly with methanol to remove residual TFA and then lyophilized, taking

care not to expose the solutions to light. Ferri-ochrobactins were stored at -80 °C in

the dark.

Ochrobactin photoproducts were prepared from solutions of iron(III)-

ochrobactins in 50% methanol/100 mM phosphate buffer pH 8.0. Ferri-ochrobactins

210 were photolyzed for 1 to 3 hours in natural sunlight in 50 mL quartz flasks at 24 °C.

Reaction progress was monitored by UV-visible spectroscopy, monitoring the

appearance of a new peak at 257 nm. When no further changes were evident by UV-

visible spectroscopy, samples were evaluated by ESI-MS. Ferri-ochrobactin photoproducts were repurified as described for the ferri-ochrobactins, lyophilized, and stored at -80 °C.

Partition Coefficient Determination

Membrane affinity coefficients were determined for the partitioning of apo-

ochrobactin C, Fe(III)-ochrobactin C, and Fe(III)-ochrobactin C photoproduct into

200 nm 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) vesicles using the

method described previously (Xu et al. 2002, Martinez et al. 2003). DMPC vesicles

were prepared by extrusion with a Mini-Extruder (Avanti Polar Lipids) using 200 nm

polycarbonate membranes. Briefly, thin-films of DMPC were prepared by

evaporating a solution of DMPC in chloroform (Avanti Polar Lipids or Sigma) under

vacuum in a 100 mL acid-washed round bottom flask. Residual chloroform was

removed under vacuum overnight. The lipid thin-films were rehydrated for 30

minutes at room temperature with 5 mL of 35 °C 10 mM Tris-HCl/100 mM KCl pH

8.0 followed by 8 freeze (liquid nitrogen)/thaw (35 °C waterbath)/vortex cycles.

Large unilamellar vesicles (LUVs) were formed by extrusion through a 200 nm

polycarbonate membrane (31 passes) (J.S. Martinez and J.N. Carter-Franklin,

personal communication). The vesicle solution was diluted to a final concentration of

211 30 mM and stored in the dark at 4 °C or on ice and used within 48 hours.

For partition coefficient determination, approximately 20 µM siderophore

solutions were incubated in acid-washed polycarbonate centrifuge tubes (Beckman)

with 0 to 30 mM DMPC for 2.5 hours at room temperature in the dark with shaking at

60 rpm on an orbital shaker. Samples were centrifuged at 200,000 × g for 3.5 hours.

The siderophore remaining in solution (DW) was assayed by RP-HPLC with an

analytical C4 column (4.6 mm ID × 250 mm L, Vydac) with a gradient from 0.05%

TFA in 50% methanol, 49.95% doubly deionized water (Barnstead Nanopure II) to

0.05% TFA in methanol over 45 minutes. The eluent was continuously monitored at

215 nm for concentration determination and at 415 nm to monitor for the presence of ferri-siderophore complexes on a Waters 2487 dual wavelength detector. The concentration of siderophore remaining in solution was determined by comparison of the peak area to a standard curve prepared for each siderophore. Partition coefficients were determined from a plot of the concentration of DMPC versus the ratio of siderophore remaining in solution (DW) to the total siderophore (DT) (Martinez et al.

2000, Martinez 2002).

212 Results

Isolation and Purification of Ochrobactins and Ochrobactin Photoproducts

As expected, approximately 0.5 to 5 mg of ochrobactin siderophores were

isolated per liter of culture. Roughly ten percent of the total material was ochrobactin

B while the remainder was predominantly ochrobactin C; small amounts of

ochrobactin A were produced, but not utilized in these experiments. The lipophilicity

of the ochrobactins makes these siderophores adhere to the walls of all containers

during purification and, especially, during concentration. Recovery of purified

siderophores is significantly increased by repeated rinsing of all containers with

methanol and by limitation of the number of transfers made.

Partition Coefficient Determination

The cell association of the ochrobactin siderophores and their obvious

lipophilicity in solution suggested significant surface activity and membrane affinity.

The partition coefficients for apo-ochrobactin B, Fe(III)-ochrobactin B, Fe(III)-

ochrobactin B photoproduct, apo-ochrobactin C, Fe(III)-ochrobactin C, and Fe(III)-

ochrobactin C photoproduct with 200 nm DMPC vesicles were determined using a

previously described technique combining ultracentrifugation and RP-HPLC to analyze the concentration of siderophore or photoproduct remaining in solution (DW).

The concentration of siderophore or photoproduct was determined from the area under the peak by RP-HPLC and comparison to a prepared standard curve for each siderophore (Figure 5.2). A plot of the millimolar concentration of DMPC versus the

213 ratio of DW to DT (the total siderophore concentration; i.e. DW + DL, where DL isthe siderophore in the lipid portion) was prepared; the ratio of siderophore remaining in solution versus the total siderophore concentration, DW/DT, was found to be inversely proportional to the concentration of DMPC. Assuming a dilute solution, the partition coefficient (K) may be expressed as

D K = L Eq. 1 [L]DW where [L] is the millimolar concentration of DMPC. Each curve was therefore fit to the equation

D 1 W = Eq. 2 DT K[L] +1 to determine the partition coefficients (K).

Experiments to investigate the partitioning of the ochrobactins into large unilamellar DMPC showed the partition coefficient of apo-ochrobactin C was 317.9 ±

33.8 M-1 whereas the partition coefficient for Fe(III)-ochrobactin C was only 49.2 ±

3.9 M-1, indicating a significant decrease in partitioning upon iron chelation (Figure

5.3, Table 5.1). The partition coefficient for Fe(III)-ochrobactin C photoproduct, however, was found to be 194.7 ± 12.1 M-1 (Figure 5.3, Table 5.1). For Fe(III)- ochrobactin C it was noted that the curve fitting did not align well with the data.

Attempts were made to improve the fit by removing high DMPC concentration data, shifting emphasis to the lower concentration points. Such attempts slightly improve the fit, but did not significantly change the calculated partition coefficient. Thus the existing fit, with the same concentration range as for apo-ochrobactin C and Fe(III)-

214 ochrobactin C photoproduct, was retained.

Partitioning experiments with apo-ochrobactin B, Fe(III)-ochrobactin B, and

Fe(III)-ochrobactin B photoproduct with DMPC vesicles demonstrated a similar pattern of partition coefficients (Figure 5.4, Table 5.2). The partition coefficient for apo-ochrobactin B was 20.5 ± 1.2 M-1, for Fe(III)-ochrobactin B was 4.4 ± 0.6 M-1, and for Fe(III)-ochrobactin B photoproduct was 25.8 ± 0.9 M-1. In this case, iron coordination also produced a significant decrease in partition coefficient relative to the apo-siderophore. Fe(III)-ochrobactin B photoproduct also demonstrated greater partitioning affinity than Fe(III)-ochrobactin B, following the same trend as was seen with ochrobactin C. Overall, the partitioning coefficients of ochrobactin B are less than ochrobactin C, consistent with the decrease in the acyl chain length of one appendage in ochrobactin B. The overall trend of reduction of partitioning with

Fe(III)-ochrobactin versus apo-ochrobactin is also consistent with previous results with Fe(III)- (174 M-1) versus apo-marinobactin (5818 M-1) (Xu et al. 2002, Martinez et al. 2003).

215 20 mM DMPC

0 mM DMPC

Figure 5.2: RP-HPLC resolution of apo-ochrobactin C. The peak at approximately 24.5 minutes, indicated by the arrow is apo-ochrobactin C. The small peak at approximately 25 minutes is Fe(III)-ochrobactin C; following incubation and ultracentrifugation, a small amount of ferrisiderophore was evident.

216

Figure 5.3: Partitioning traces for apo-ochrobactin C (●), Fe(III)-ochrobactin C (∇), and Fe(III)-photoproduct C (■).

217

Figure 5.4: Partitioning traces for apo-ochrobactin B (●), Fe(III)-ochrobactin B (∇), and Fe(III)-photoproduct B (■).

218

Sample Injection Peak [DMPC] (mM) Volume (µL) Area OC (nmoles) Dw (M) Dw/Dt 0 mM 1b 250 6508275 6.020 2.408 x10-5 1 0 mM 1a 250 5683701 5.278 2.111 x10-5 0.877 0 mM 1b 250 5560125 5.167 2.067 x10-5 0.858 0 mM 1a 250 5806126 5.388 2.155 x10-5 0.895

1 mM 1b 500 7813878 7.195 1.439 x10-5 0.598 1 mM1a 500 8826144 8.106 1.621 x10-5 0.673

2.5 mM 1b 500 7077215 6.532 1.306 x10-5 0.542 2.5 mM 1a 500 6668511 6.164 1.233 x10-5 0.512

5 mM 1b 500 4544693 4.253 8.506 x10-6 0.353 5 mM 1a 500 4326899 4.057 8.114 x10-6 0.337

7.5 mM 2a 500 4399265 4.122 8.244 x10-6 0.342 7.5 mM 1b 500 3611450 3.413 6.826 x10-6 0.283 7.5 mM 1a 500 4244448 3.983 7.965 x10-6 0.331

10 mM 1a 1000 7624054 7.024 7.024 x10-6 0.292

15 mM 1b 1000 4850515 4.528 4.528 x10-6 0.188 15 mM 1a 1000 5839135 5.418 5.418x10-6 0.225

20 mM 1b 1000 4684080 4.378 4.378 x10-6 0.182 20 mM 1a 1000 5103702 4.756 4.756 x10-6 0.198

25 mM 1b 1000 6454662 5.972 5.972 x10-6 0.248 25 mM 1a 1000 4313113 4.044 4.044 x10-6 0.168

30 mM 1b 1000 4533202 4.242 4.242 x10-6 0.176 30 mM 1a 1000 4544331 4.252 4.253 x10-6 0.177

Table 5.1: Data from apo-ochrobactin C partitioning experiments.

219

Sample Injection Peak Fe(III)-OC [DMPC] (mM) Volume (µL) Area (nmoles) Dw (M) Dw/Dt 0 mM 1a 250 4140548 4.339 1.736 x10-5 0.985 0 mM 1b 250 4205788 4.405 1.762 x10-5 1

1 mM 1a 500 7771469 7.970 1.594 x10-5 0.905 1 mM 1b 500 7652146 7.851 1.570 x10-5 0.891

2.5 mM 1a 500 6778155 6.977 1.395 x10-5 0.792 2.5 mM 1b 500 6984747 7.184 1.437 x10-5 0.815

5 mM 1a 500 6638238 6.837 1.367 x10-5 0.776 5 mM 1b 500 6382817 6.582 1.316 x10-5 0.747

7.5 mM 1a 500 5913897 6.113 1.223 x10-5 0.694 7.5 mM 1b 500 5611531 5.810 1.162 x10-5 0.660

10 mM 1a 500 5288911 5.488 1.098 x10-5 0.623 10 mM 1b 500 5334715 5.534 1.107 x10-5 0.628

15 mM 1a 500 5079499 5.278 1.056 x10-5 0.599 15 mM 1b 500 4961724 5.161 1.032 x10-5 0.586

20 mM 1 a 500 4371861 4.571 9.142 x10-6 0.519 20 mM 1b 500 4414668 4.614 9.227 x10-6 0.524

30 mM 1a 500 4254418 4.453 8.907 x10-6 0.506 30 mM 1b 500 4269061 4.468 8.936 x10-6 0.507

Table 5.2: Data from Fe(III)-ochrobactin C partitioning experiments.

220

Sample Injection Peak Fe(III)-OC PP [DMPC] (mM) Volume (µL) Area (nmoles) Dw (M) Dw/Dt 0 mM 1a 250 3242488 6.575 2.630x10-5 1 0 mM 1b 250 3164209 6.419 2.568 x10-5 0.976

1 mM 1a 500 5030642 10.152 2.030 x10-5 0.772 1 mM 1b 500 4976414 10.043 2.009 x10-5 0.764

2.5 mM 1a 500 3960425 8.011 1.602 x10-5 0.609 2.5 mM 1b 500 4656553 9.404 1.881 x10-5 0.715 2.5 mM 1c 500 3702036 7.495 1.499 x10-5 0.570

5 mM 1a 500 3404917 6.900 1.380 x10-5 0.525 5 mM 1b 500 3094533 6.280 1.256 x10-5 0.477

10 mM 1a 1000 3505296 7.101 7.101 x10-6 0.270 10 mM 1b 1000 3590952 7.272 7.272 x10-6 0.276

15 mM 1a 1000 3861938 7.814 7.814 x10-6 0.297 15 mM 1b 1000 3461763 7.014 7.014 x10-6 0.267

20 mM 1a 1000 2126615 4.344 4.344 x10-6 0.165 20 mM 1b 1000 2060439 4.211 4.211 x10-6 0.160

25 mM 1a 1000 2375820 4.842 4.842 x10-6 0.184 25 mM 1b 1000 2575497 5.241 5.241 x10-6 0.199

30 mM 1a 500 977862 2.046 4.092 x10-6 0.156 30 mM 1b 1000 1644344 3.379 3.379 x10-6 0.128

Table 5.3: Data from Fe(III)-ochrobactin C photoproduct partitioning experiments.

221

Sample Injection Peak [DMPC] (mM) Volume (µL) Area OB (nmoles) Dw Dw/Dt 0 mM 1a 500 8439485 8.333 1.67 x10-5 0.993 0 mM 1b 500 8497135 8.391 1.68 x10-5 1

2.5 mM 1a 500 8263809 8.157 1.63 x10-5 0.972 2.5 mM 1b 500 8343179 8.237 1.65 x10-5 0.982

5 mM 1a 500 7554324 7.448 1.49 x10-5 0.888 5 mM 1b 500 7624265 7.518 1.50 x10-5 0.896

7.5 mM 1a 500 7018929 6.912 1.38 x10-5 0.824 7.5 mM 1b 500 7086352 6.980 1.40 x10-5 0.832

10 mM 1a 500 6717109 6.611 1.32 x10-5 0.788 10 mM 1b 500 6675976 6.569 1.31 x10-5 0.783

15 mM 1a 500 6037778 5.931 1.19 x10-5 0.707 15 mM 1b 500 6047484 5.941 1.19 x10-5 0.708

20 mM 1a 500 5960092 5.853 1.17 x10-5 0.697 20 mM 1b 500 5960994 5.854 1.17 x10-5 0.698

25 mM 1a 500 5821483 5.715 1.14 x10-5 0.681 25 mM 1b 500 5920559 5.814 1.16 x10-5 0.693

30 mM 1a 500 5898833 5.792 1.16 x10-5 0.690 30 mM 1b 500 5916123 5.810 1.16 x10-5 0.692

Table 5.4: Data from apo-ochrobactin B partitioning experiments.

222

Sample Injection Peak Fe(III)-OB [DMPC] (mM) Volume (µL) Area (nmoles) Dw Dw/Dt 0 mM 1a 250 4986258 4.374758 1.75 x10-5 0.970

1 mM 1a 500 9633163 9.021663 1.80 x10-5 1 1 mM 1b 500 9434358 8.822858 1.76 x10-5 0.978

2.5 mM 1a 500 9203253 8.591753 1.72 x10-5 0.952 2.5 mM 1b 500 9109156 8.497656 1.70 x10-5 0.942

5 mM 1a 500 9069559 8.458059 1.69 x10-5 0.938 5 mM 1b 500 8957310 8.34581 1.67 x10-5 0.925

7.5 mM 1a 500 8957482 8.345982 1.67 x10-5 0.925

10 mM 1a 500 8940071 8.328571 1.67 x10-5 0.923

15 mM 1a 500 9214024 8.602524 1.72 x10-5 0.954 15 mM 1b 500 8828189 8.216689 1.64 x10-5 0.911

20 mM 1a 500 8874512 8.263012 1.65 x10-5 0.916 20 mM 1b 500 9211138 8.599638 1.72 x10-5 0.953

25 mM 1a 500 8780015 8.168515 1.63 x10-5 0.905 25 mM 1b 500 9001506 8.390006 1.68 x10-5 0.930

30 mM 1a 500 8715388 8.103888 1.62 x10-5 0.898 30 mM 1b 500 8531154 7.919654 1.58 x10-5 0.878

Table 5.5: Data from Fe(III)-ochrobactin B partitioning experiments.

223

Sample Injection Peak Fe(III)-OB PP [DMPC] (mM) Volume (µL) Area (nmoles) Dw Dw/Dt 0 mM 1a 250 3341404 3.240 1.30 x10-5 1 0 mM 1b 500 7017901 6.916 1.38 x10-5 1

2.5 mM 1a 500 5994730 5.893 1.18 x10-5 0.909 2.5 mM 1b 500 6030710 5.929 1.19 x10-5 0.915

10 mM 1a 500 5155712 5.054 1.01 x10-5 0.780 10 mM 1b 500 5297962 5.196 1.04 x10-5 0.802

20 mM 1a 500 4201382 4.100 8.20 x10-5 0.633 20 mM 1b 500 4490645 4.389 8.78 x10-6 0.677

30 mM 1a 500 3835694 3.734 7.47 x10-6 0.576 30 mM 1b 500 3788122 3.687 7.37 x10-6 0.569

1 mM 1a 500 6399052 6.298 1.26 x10-5 0.972 1 mM 1b 500 6204573 6.103 1.22 x10-5 0.942

5 mM 1a 500 5900381 5.799 1.16 x10-5 0.895 5 mM 1b 500 5704376 5.603 1.12 x10-5 0.865

7.5 mM 1a 500 5350694 5.249 1.05 x10-5 0.810 7.5 mM 1b 500 5449163 5.348 1.07 x10-5 0.825

15 mM 1a 500 4611007 4.510 9.02 x10-6 0.696 15 mM 1b 500 4712290 4.610 9.22 x10-6 0.712

25 mM 1a 500 4363874 4.262 8.52 x10-6 0.658 25 mM 1b 500 4169321 4.068 8.14 x10-6 0.628

Table 5.6: Data from Fe(III)-ochrobactin B photoproduct partitioning experiments.

224 Both the RP-HPLC results (A415nm) and the mass spectrometry data support the assertion that the Fe(III)-ochrobactins and the Fe(III)-ochrobactin photoproducts are indeed iron bound. Therefore, these results suggest significant differences in the lipophilicity between the Fe(III)-ochrobactin complex and the Fe(III)-ochrobactin photoproduct, possibly deriving from significant differences in the three-dimensional configuration of the molecules.

225 Discussion

The differences in partition coefficient between apo-ochrobactin, Fe(III)- ochrobactin, and Fe(III)-photoproduct are intriguing. The apo-ochrobactins partitioned much more extensively (apo-ochrobactin B, 21 ± 1 M-1; apo-ochrobactin

C, 318 ± 34 M-1) than the iron(III) siderophore complexes (Fe(III)-ochrobactin B, 4 ±

1 M-1; Fe(III)-ochrobactin C, 49 ± 4 M-1). However, photolysis of the ferric ochrobactin complexes significantly increases the lipid partitioning coefficients (i.e.,

Fe(III)-photoproduct B, 26 ± 1 M-1; Fe(III)-photoproduct C, 195 ± 12 M-1) relative to the iron(III) ochrobactin B and C complexes, respectively.

For the marinobactins, decreasing the length of the acyl appendage by two methylene carbons resulted in a significant reduction of partition coefficient (i.e.

-1 -1 marinobactin E (C16:0) 5818 M to marinobactin C (C14:0) 195 M ) (Xu et al. 2002).

The amphibactins also exhibited a reduction in partition coefficient with decreased

-1 acyl appendage length (i.e. amphibactin H (C16:0) 3784 M to amphibactin D (C14:0)

1915 M-1) (Martinez et al. 2003). As was seen previously with the marinobactins and amphibactins, decreasing the length of one fatty acid appendage in ochrobactin by two methylene carbons results in a significant reduction in partition coefficient (Xu et al. 2002, Martinez et al. 2003).

It is interesting to note that the partition coefficient determined for secreted marinobactin E is higher than the partition coefficients determined for the cell- associated ochrobactins. The partition coefficients for the marinobactins range from

36 for marinobactin A (C12:0) to 5818 for marinobactin E (C16:0) for 200 nm DMPC

226 vesicles, yet these siderophores are found in the supernatant (Xu et al. 2002). The partition coefficients for the amphibactins are generally higher, ranging from 833 for amphibactin G (C18:1 3-OH) to 3784 for amphibactin H (C16:0) and these siderophores are generally isolated from the cell pellet (Martinez et al. 2003). The amphibactins have a smaller hydrophilic headgroup than the marinobactins (four amino acids versus six amino acids) which may contribute to the observed differences in partition coefficients (Martinez et al. 2000, Xu et al. 2002, Martinez et al. 2003). The ochrobactins have lower partition coefficients than either the marinobactins or the amphibactins yet are found in the cell pellet under laboratory conditions. While the ochrobactins have a smaller “headgroup” consisting of two lysine residues and citrate, they also each have two acyl appendages. Both the headgroup size and the acyl appendage length (or number) contribute to the membrane partitioning of amphiphilic molecules. Differences in the outer membrane or perhaps the culture medium may also have important effects on the membrane association of amphiphilic marine siderophores.

The observed differences in partition coefficients could contribute a physiological benefit. Constraining apo-siderophore to the vicinity of the cell could prevent loss by diffusion, thereby potentially increasing iron availability for the cell.

At the same time, releasing Fe(III)-siderophore upon iron binding could promote active uptake, both by keeping the siderophore complex near the cell but also by freeing the ferri-siderophore complex for interaction with the putative outer membrane transport protein. The interaction of ferri-ochrobactin photoproduct

227 complexes with DMPC vesicles is even more intriguing. One could reason that the ferri-ochrobactin photoproducts would only be present if the cell was in open water conditions. Under these conditions it might be beneficial to keep a tight hold on siderophores to prevent loss or piracy and to promote iron uptake. It is also possible that the lipophilicity of the ochrobactin siderophores may permit these siderophores to interact with particles such as iron containing minerals.

Recent 3D modeling of apo- and Fe(III)-acinetoferrin suggest a possible structural explanation for differences in partition coefficients (Luo et al. 2005).

Specifically, the acyl appendages of apo-acinetoferrin are predicted to be in a nearly parallel configuration. Thus, both fatty acid groups could interact with the membrane simultaneously, increasing partitioning. The Fe(III) complex, however, is predicted to have an angle of approximately 130° separating the tails. Therefore, only one tail could interact with the membrane at any one time, reducing the partition coefficient

(Luo et al. 2005, Fadeev et al. 2004). If such a mechanism is operative with the

Fe(III)-ochrobactins, one might expect all the Fe(III)-ochrobactins to partition similarly because each contain the C10:1 fatty acid, yet actually Fe(III)-ochrobactin C with two C10:1 fatty acid appendages partitions more than Fe(III)-ochrobactin B with

C10:1 and C8:0 fatty acids.

The cell association and lipophilicity of the ochrobactins and amphibactins are intriguing features of these siderophores. The mechanism of iron uptake by these siderophores, the specific physiological functions of cell-association, and the factors contributing to cell-association remain to be elucidated.

228 References

Fadeev, E.V., Luo, M.K., and Groves, J.T. (2004) Journal of the American Chemical Society 126, 12065 – 12075.

Gobin, J. and Horwitz, M. (1996) Journal of Experimental Medicine 183, 1527 – 1532.

Inoue, T. (1996) in Vesicles 62, 151 – 193. Rosoff, M., Ed. Marcel Dekker, New York.

Luo, M.K., Fadeev, E.V., and Groves, J.T. (2005) Journal of the American Chemical Society 127, 1726 – 1736.

Kokubo, S., Suenaga, K., Shinohara, C., Tsuji, T., and Uemura, D. (2000) Tetrahedron 56, 6435 – 6440.

Martinez, J.S. (2002) Mechanisms of siderophore-mediated iron sequestration by marine bacteria. Dissertation University of California, Santa Barbara.

Martinez, J.S., Carter-Franklin, J.N., Mann, E.L., Martin, J.D., Haygood, M.G., and Butler, A. (2003) Proceedings of the National Academy of Sciences, USA 100, 3754 – 3759.

Murakami, K., Fuse, H., Takimura, O., Inoue, H., and Yamaoka, Y. (2000) Microbios 101, 137 – 146.

Murakami, Y., Kato, S., Nakajima, M., Matsuoka, M., Kawai, H., ShinYa, K., and Seto, H. (1996) Journal of Antibiotics 49, 839 – 845.

Ratledge, C. (2004) Tuberculosis 84, 110 – 130.

Ratledge, C. and Dale, J. (1999) Mycobacteria: molecular biology and virulence. Blackwell Science, Ltd., Oxford, UK

Ratledge, C. and Patel, P. (1976) Journal of General Microbiology 93, 141 – 152.

Suenaga, K., Kokubo, S., Shinohara, C., Tsuji, T., and Uemura, D. (1999) Tetrahedron Lett 40, 1945 – 1948.

Wooldridge, K.G., Morrisey, J.A., and Williams, P.H. (1992) Journal of General Microbiology 138, 597 – 603.

Xu, G., Martinez, J.S., Groves, J.T., and Butler, A. (2002) Journal of the American

229 Chemical Society 124, 13408 – 10415.

230 Chapter Six

Concluding Remarks and Future Directions

The Ochrobactins

Two species of marine bacteria, Ochrobactrum sp. SP18 and Vibrio sp.

DS40M5, were found to produce the ochrobactins, a new suite of aerobactin-derived

cell-associated siderophores. Each ochrobactin siderophore is composed of citrate symmetrically derivatized with L-lysine; each lysine is Nε-acylated and Nε-

hydroxylated to form two hydroxamate binding moieties. Each ochrobactin

siderophore contains one (E)-2-decenoic acid and a second acyl appendage,

ochrobactin A contains (E)-2-octenoic acid, ochrobactin B contains octanoic acid, and ochrobactin C contains a second (E)-2-decenoic acid moiety (Figure 6.1, see also

Chapter 2 (Martin et al. 2006)).

O A O O OH O B O N N C H OH OH OH OH H O N N O O O OH

Figure 6.1: Structures of the ochrobactin siderophores.

Vibrio sp. DS40M5 is a marine γ-proteobacterium while Ochrobactrum sp. SP18 is a marine α-proteobacterium. It is interesting to note that two strains of bacteria from

231 widely different clades both produce a suite of ochrobactin siderophores. Evaluation of other aerobactin producing marine and terrestrial bacterial strains will help to

establish the validity of the assertion that the ochrobactins confer some special

advantage in marine environments.

Biosynthesis of Citrate-Derived Siderophores

Rhizobactin 1021 is a citrate-derived, amphiphilic siderophore produced by

the alfalfa symbiont Sinorhizobium meliloti 1021 (Figure 6.2) (Persmark et al. 1993).

The biosynthetic pathway of rhizobactin 1021 has been partially elucidated (Lynch et

al. 2001). Six genes were identified and found to be involved in the biosynthesis of

rhizobactin 1021 designated rhbABCDEF (Lynch et al. 2001). Transposon

mutagenesis and sequence analysis were utilized to predict the functions of RhbA,

RhbB, RhbC, RhbD, RhbE, and RhbF. The biosynthesis of rhizobactin is predicted to

begin with the decarboxylation and amination of L-glutamic acid by RhbA to form L-

2,4-diaminobutryic acid, which is then decarboxylated by RhbB to form 1,3-

diaminopropane (Figure 6.2) (Lynch et al. 2001). 1,3-diaminopropane is N4-

hydroxylated by RhbE and N4-acetylated by RhbD (Lynch et al. 2001). Citrate is

derivatized at one end with the resulting N4-acetyl-N4-hydroxy-1-aminopropane by

RhbC followed by derivatization at the other end by RhbF to form schizokinen

(Lynch et al. 2001). Finally, an unidentified gene product replaces one acetyl group

with (E)-2-decenoic acid to form rhizobactin 1021 (Lynch et al. 2001). It is likely that this unidentified gene is not located near the rhbABCDEF operon.

232 O OH OH O RhbA H2N RhbB H2N RhbE HN RhbD HO N O O CO 2 1/2 O2 Acetyl CoA H2N OH H2N OH H2N H2N CoA H2N L-glutamic acid L-2,4-diamino- 1,3-diamino- N4-hydroxy- N4-acetyl-N4-hydroxy- butyric acid propane 1-aminopropane 1-aminopropane Citrate O

HO N RhbC

HO O HO HN O O OH

RhbF

O O

N OH HO N

HO O NH HN O O HO Schizokinen

? O O

N OH HO N

HO O NH HN O O HO Rhizobactin 1021

Figure 6.2: Biosynthetic pathway for rhizobactin 1021. RhbABCDEF produce schizokinen and an unidentified additional enzyme replaces one terminal acetyl group with one (E)-2-decenoic acid moiety (Lynch et al. 2001).

233 OH O H2N HN HO N IucD IucB O O O 1/2 O2 Acetyl CoA H2N OH H2N OH CoA H2N OH L-lysine Nε-OH-lysine Nε-acetyl- Nε-OH-lysine

O Citrate HO N IucA

O HO O HO HN OH O O OH

IucC

O O N OH HO N

O O HO O HO NH HN OH O O OH

Aerobactin

Figure 6.3: Biosynthetic pathway for aerobactin (deLorenzo and Neilands 1986, deLorenzo et al. 1986).

234 The relationship between rhizobactin 1021 and schizokinen is similar to the structural relationship between the ochrobactins and aerobactin. The biosynthetic pathway of aerobactin is known and resembles that of schizokinen and rhizobactin

1021. In the biosynthesis of aerobactin, L-lysine is sequentially Nε-hydroxylated and

Nε-acetylated by IucD and IucB, respectively (deLorenzo et al. 1986, deLorenzo and

Neilands 1986). Citrate is then derivatized with Nε-acetyl, Nε-hydroxy-lysine at both ends by IucA and then IucC to form the siderophore aerobactin (deLorenzo and

Neilands 1986). The discovery of the ochrobactins and the similarity to rhizobactin

1021 suggests interesting parallels between the biosynthetic pathways, specifically in the attachment of the acyl appendages.

The gene sequences for a number of aerobactin biosynthesis operons have been submitted to the National Center for Biotechnology Information. Interestingly, the gene sequences for aerobactin biosynthesis from terrestrial and pathogenic bacteria and from marine Vibrio species are quite diverse. Sequence alignment between distantly related species has fairly low sequence identity or homology. It would be very interesting to pursue the biosynthetic pathways for aerobactin from

Vibrio sp. DS40M5 and for the ochrobactins both from Ochrobactrum sp. SP18 and from Vibrio sp. DS40M5. Sequence similarity between the siderophore biosynthetic pathways might suggest lateral gene transfer; significant differences in the sequences might suggest convergent evolution. With the sequences for aerobactin biosynthesis genes from several marine Vibrio species published, it may be possible to amplify the aerobactin biosynthetic genes using a PCR technique. If the sequences are too

235 dissimilar, it may be possible to generate a genomic library for screening. As was seen for the biosynthesis of rhizobactin 1021 and schizokinen, the biosynthetic pathway for the production of the ochrobactins may be similar to the pathway for biosynthesis of aerobactin. Of additional interest in the biosynthesis of these siderophores is the protein involved in attachment of the acyl appendage. The specific protein responsible for addition of the acyl appendage of rhizobactin 1021 has not yet been identified. If the acyl appendages of the ochrobactins are attached in a similar fashion, then isolation of the ochrobactin biosynthetic genes could potentially help to locate the gene for the final biosynthetic step of rhizobactin 1021.

Putative Outer Membrane Siderophore Transport Protein

While it has been suggested that marine siderophores are transported into the cells by an outer membrane siderophore transport protein, such a protein has not been characterized. Preliminary experiments with Pseudoalteromonas luteoviolacea indicate that an iron repressible outer membrane protein is produced under conditions of iron limitation (Reid et al. 1993, Reid 1994). This protein is suggested to be the outer membrane transport protein for the alterobactins (Reid et al. 1993, Reid 1994).

Aerobactin is taken into the cell through an outer membrane siderophore transport protein, IutA (Wooldridge et al. 1992). Given the structural similarities between the ochrobactins and aerobactin, it seems reasonable that the ochrobactins may be taken up through a similar means. The nature of the putative outer membrane protein and the differences between this putative protein and the aerobactin outer

236 membrane transport protein may clarify siderophore transport in this marine

bacterium.

Photoreactivity of the Ochrobactins

Expansion of the photochemical studies of the ochrobactins may prove

rewarding. Investigation of the concentration dependence and pH dependence of the photoreaction will contribute to understanding the photochemical reaction. While

there are significant limitations on the range of siderophore concentrations that can be

studied in aqueous solution, comparison of the quantum efficiencies at varying

concentrations could be interesting.

The ochrobactins have all of the hallmarks of structurally characterized

marine siderophores: (1) binding moieties which are photoreactive when coordinated

to iron(III), and (2) suites of amphiphilic siderophores which differ by the length,

saturation, or substitution of the acyl appendages. In addition, the structural

similarity of the ochrobactins to aerobactin provides an interesting opportunity to

investigate the biosynthesis of structurally related siderophores. The suite of

ochrobactin siderophores provides a useful system for the study of marine

siderophores, their biosynthesis, and their reactivities.

237 References deLorenzo, V., Bindereif, A., Paw, B.H., and Neilands, J.B. (1986) Journal of Bacteriology 165, 570 – 578. deLorenzo, V. and Neilands, J.B. (1986) Journal of Bacteriology 167, 350 – 355.

Lynch, D., O’Brien, J., Welch, T., Clarke, P., Cuív, P.Ó., Crosa, J.H., and O’Connell, M. (2001) Journal of Bacteriology 183, 2576 – 2585.

Martin, J.D., Ito, Y., Homann, V.V., Haygood, M.G., and Butler, A. (2006) Journal of Biological Inorganic Chemistry in press.

Persmark, M., Pittman, P., Buyer, J.S., Schwyn, B., Gill, P.R.J., and Neilands, J.B. (1993) Journal of the American Chemical Society 115, 3950 – 3956.

Wooldridge, K.G., Morrissey, J.A., and Williams, P.H. (1992) Journal of General Microbiology 138, 597 – 603.

238