RECOMBINANT PRODUCTION OF OMEGA-3 FATTY ACIDS IN E. COL1
USING A GENE CLUSTER ISOLATED FROM SHEWANELLA BALTICA
A Thesis
Presented to
The Faculty of Graduate Studies
of
The University of Guelph
By
MITRA AMIRI-JAMI
In partial fulfillment of requirements
For the degree of
Doctor of Philosophy
January, 2009
© Mitra Amiri-Jami, 2009 Library and Archives Bibliotheque et 1*1 Canada Archives Canada Published Heritage Direction du Branch Patrimoine de I'edition
395 Wellington Street 395, rue Wellington Ottawa ON K1A 0N4 OttawaONK1A0N4 Canada Canada
Your file Votre reference ISBN: 978-0-494-58261 -9 Our file Notre reference ISBN: 978-0-494-58261-9
NOTICE: AVIS:
The author has granted a non L'auteur a accorde une licence non exclusive exclusive license allowing Library and permettant a la Bibliotheque et Archives Archives Canada to reproduce, Canada de reproduire, publier, archiver, publish, archive, preserve, conserve, sauvegarder, conserver, transmettre au public communicate to the public by par telecommunication ou par I'lnternet, prefer, telecommunication or on the Internet, distribuer et vendre des theses partout dans le loan, distribute and sell theses monde, a des fins commerciales ou autres, sur worldwide, for commercial or non support microforme, papier, electronique et/ou commercial purposes, in microform, autres formats. paper, electronic and/or any other formats.
The author retains copyright L'auteur conserve la propriete du droit d'auteur ownership and moral rights in this et des droits moraux qui protege cette these. Ni thesis. Neither the thesis nor la these ni des extraits substantiels de celle-ci substantial extracts from it may be ne doivent etre imprimes ou autrement printed or otherwise reproduced reproduits sans son autorisation. without the author's permission.
In compliance with the Canadian Conformement a la loi canadienne sur la Privacy Act some supporting forms protection de la vie privee, quelques may have been removed from this formulaires secondaires ont ete enleves de thesis. cette these.
While these forms may be included Bien que ces formulaires aient inclus dans in the document page count, their la pagination, il n'y aura aucun contenu removal does not represent any loss manquant. of content from the thesis.
1+1 Canada ABSTRACT
RECOMBINANT PRODUCTION OF OMEGA-3 FATTY ACIDS IN E. COLI
USING A GENE CLUSTER ISOLATED FROM SHEWANELLA BALTICA
Mitra Amiri-Jami Advisor: University of Guelph, 2009 Professor M. W. Griffiths
Omega-3 fatty acids such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) play an important role in many aspects of human health. The main dietary source of EPA and DHA is cold-water fish, the populations of which are in decline.
There has been extensive research to find alternative sources for EPA/DHA. Production of EPA and DHA by some bacterial strains such as Shewanella has been of interest for the purpose of cloning omega-3 fatty acid genes and recombinant production of
EPA/DHA in other microorganism or higher organism such as plants. The first study presented in this thesis demonstrates the production of EPA/DHA in Shewanella baltica strains, which have been previously reported as non-omega-3 fatty acid-producing bacteria. All Shewanella baltica strains produced EPA and Shewanella baltica MAC1 synthesized both EPA and DHA. Next, we cloned EPA/DHA genes from Shewanella baltica MAC1 by constructing a bacterial artificial chromosome (BAC) library.
Recombinant production of EPA/DHA was detected in one clone with a DNA insert size of 200 kb. Attempts to subclone the EPA/DHA gene cluster into a food grade, broad host range vector were not successful due to an inability to digest and to isolate the insert from the BAC vector. In chapter four, first each pfa (pfaA, pfaB, pfaC, pfaD, pfaE) gene of Shewanella baltica MAC1 responsible for EPA/DHA production was partially amplified in order to make probes for rapid detection of positive clones in a fosmid library. Second, to clone the EPA/DHA gene cluster, we prepared a fosmid library from
Shewanella baltica MAC1 with insert size of -40 kb. Fosmid clones were screened by colony hybridization for pfaA and pfaD genes. Recombinant production of EPA was detected in six fosmid clones and one clone produced both EPA and DHA. The insert carrying EPA/DHA genes was isolated from the fosmid DNA using Noil. The research presented in this thesis demonstrated recombinant production of omega-3 fatty acid in E. coli and provided a DNA fragment (35 kb) containing all EPA/DHA genes which can be ligated to a food grade, broad host range vector in order to provide alternative sources of
EPA and DHA. ACKNOWLEDGEMENTS
I would like to thank my supervisor Dr. Mansel Griffiths who gave me the opportunity to work in his lab and for his trust that I was the right person for this project. He has provided advice, support, patience and respect while allowing me to maintain my independence, and for that, I am indebted and grateful.
I would like to thank the members of my advisory committee. Dr. Yukio Kakuda who has always been available for long scientific discussions, support, encouragements and his expert knowledge. Dr. George Van der Merwe for his insightful comments and attention to detail.
I would like to thank the many friends that I have made over the years for their support and memories that I will cherish forever.
I would like to thank my parents who have always encouraged me to pursue my dreams. I would also like to thank my sisters and my brother for the bond we share. You have been my confidant, my supporter and my best friends.
And to my family, I cannot say enough. My husband, Ali, who has loved and supported me no matter what. His patience, encouragement, understanding and scientific advice have been greatly appreciated. He made the rough times more bearable and the good times even better. To our two sons, Amin and Nima, I am in awe of their wisdom, patience and unbelievable understanding. You two have been a blessing to us and look forward to watching you grow. I love you and your father very much and I could not be a person who I am today without you.
Finally, I thank God, the most compassionate, the most merciful, who makes all things possible. Table of Contents
1. General Introduction 1
1.1 An introduction to omega-3 fatty acids 1
1.2 The importance of omega-3 fatty acids 2 1.2.1 Omega-3 fatty acids and cardiovascular disease 4 1.2.2 Omega-3 fatty acids and cancer 6 1.2.3 Omega-3 fatty acids and diabetes 8
1.3 Sources of omega-3 fatty acid and recommended dietary intakes for the omega-3 fatty acids 9
1.4 An Introduction to Shewanella Species 10
1.5 The genus Shewanella and omega-3 fatty acid production 12
1.6 Mechanism of polyunsaturated fatty acid biosynthesis in bacteria 13
1.7 Genetic Studies in Shewanella Species 17
1.8 Methods for cloning the omega-3 fatty acid genes 20
1.9 Rationale 26
1.10 Research objectives 28
2. Reclassification of Shewanella putrefaciens MAC1 and detection of EPA/DHA production by Shewanell baltica strains 29
2.1 Abstract 29
2.2 Introduction 30
2.3 Materials and Methods 33 2.3.1 Bacterial strains 33 2.3.2 Preparation of Shewanella baltica cultures 34 2.3.3 Growth conditions and biochemical characteristics 34 2.3.4 DNA extraction from Shewanella baltica strains 35
ii 2.3.5 Polymerase Chain Reaction (PCR) procedure for 16S rRNA sequence determination 36 2.3.6 Separation of DNA fragments 36 2.3.7 Purification of amplified DNA fragment from PCR reactions 38 2.3.8 Growth conditions for fatty acid extraction 38 2.3.9 Extraction of fatty acids 38 2.3.10 Gas Chromatographic Analysis 39 2.3.11 Gas Chromatographic-Mass Spectrometry Analysis 40 2.3.12 Statistical Analysis 42
2.4 Results and discussion 42 2.4.1 Physiological properties and phenotypic characteristics of S. baltica strains 2.4.2 16S rRNA sequence analysis 42 2.4.3 Fatty acid composition of Shewanella baltica strains at different temperatures 45 2.4.4 EPA production by Shewanella baltica strains at different temperatures .... 48
2.5 Conclusions 56
3. Cloning omega-3 fatty acid genes from Shewanella balticaMACl by constructing a Bacterial Artificial Chromosomelibrary 57
3.1 Abstract 57
3.2 Introduction 58
3.3 Material and Methods 61 3.3.1 Bacterial strains and growth conditions 61 3.3.2 Genomic DNA isolation 62 3.3.2.1 DNA extraction using commercial kits 62 3.3.2.2 Extraction of DNA in plug agarose 63 3.3.3. Partial digestion of isolated chromosomal DNA with kits and in plugs 63 3.3.4 Pulsed Field Gel Electrophoresis 65 3.3.5 DNA recovery from the gel slices 65 3.3.5.1 Recovering of digested DNA from the gel slices using Gelase 65
iii 3.3.5.2 Recovering of DNA from gel slices by Electroelution 66 3.3.6. Construction of Bacterial Artificial Chromosome (BAC) Libraries 67 3.3.6.1 Ligation of DNA fragments into the CopyControl pCCBAC Cloning- Ready_Vector 67 3.3.6.2 Transformation, Plating and Selecting the BAC Clones 69 3.3.6.3 Sizing the BAC clones and storage of the BAC DNA 69 3.3.7 Screening of the Library 71 3.3.7.1 Growth condition 71 3.3.7.2 Fatty acid extraction 71 3.3.7.3 Gas Chromatographic Analysis 71 3.3.8 Analysis of EPA/DHA positive clones 72 3.3.8.1 Bacterial strain and growth conditions 72 3.3.8.2 BAC DNA isolation from positive clones 73 3.3.8.3 Complete digestion of BAC DNA 73 3.3.9 Determination of EPA/DHA production by positive clones in Skim milk... 75 3.3.9.1 Bacterial strains and growth condition 75 3.3.9.2 Fatty acid extraction and GC analysis 75
3.4 Results and discussion 75 3.4.1 Preparation of a BAC library of S. baltica MAC 1 genomic DNA 75 3.4.2 Screening of the BAC library 81 3.4.3 Isolation, digestion and end sequence of EPA/DHA positive BAC DNA ... 91 3.4.5 Assessment of Recombinant production of EPA/DHA in LB Broth and Skim Milk at different temperatures 94
3.5 Conclusion 98
4. Cloning omega-3 fatty acids genes from Shewanella balticaMACl by constructing afosmid library 99
4.1 Abstract 99
4.2 Introduction 100
4.3 Material and Methods 103
iv 4.3.1 Amplification of genes responsible for EPA/DHA production 103 4.3.1.1 Bacterial strains and growth conditions 103 4.3.1.2 Genomic DNA isolation of S.baltica MAC1 104 4.3.1.3 Polymerase Chain Reaction (PCR) procedure for EPA/DHA genes 104 amplification 104 4.3.1.4 Separation of DNA fragments 109 4.3.1.5 Purification of amplified DNA fragment from PCR reactions for sequencing 109 4.3.2 Construction of Fosmid library from S. baltica MAC1 genome 110 4.3.2.1 Bacterial strains and growth conditions 110 4.3.2.2 Genomic DNA isolation from 5. baltica MAC1 110 4.3.2.3 Shearing S.baltica MAC1 genomic DNA and sizing the fragmented DNA by_pulse field gel electrophoresis Ill 4.3.2.4 End-repair of the sheared DNA to blunt 5'-phosphorylated ends Ill 4.3.2.5 Sizing the end-repair DNA in gel electrophoresis 112 4.3.2.6 Ligation of DNA fragments into the pCClFOS fosmid vector 113 4.3.2.7 Packaging the ligated DNA and infecting EPI 300-77 E.coli 113 4.3.2.8 Sizing the fosmid clones and storage of the fosmid clones 114 4.3.3. Screening of the fosmid library by colony hybridization 115 4.3.3.1 Polymerase Chain Reaction (PCR) procedure to amplify pfaA (EPA5) and pfaD (EPA8) genes 115 4.3.3.2 Labeling the purified PCR products (probe) 116 4.3.3.3 Determination of labeling efficiency 116 4.3.3.4 Chemiluminescent detection with CSPD 117 4.3.3.5 Colony hybridization procedure 118 4.3.4 Analysis of fosmid clones positive for pfaA and/or pfaD genes 120 4.3.4.1 Bacterial strains and growth conditions 120 4.3.4.2 DNA isolation and PCR for pfaA and pfaD genes 121 4.3.4.3 Digestion of FOS DNA positive for both pfaA andpfaD genes 121 4.3.4.4 End sequence of fosmid DNA isolated from positive clones 122 4.3.5 Detection of EPA/DHA production in positive FOS clones 122
V 4.3.5.1 Bacterial strains and growth conditions 122 4.3.5.2 Fatty acid extraction and GC analysis 123 4.3.5.4. Gas Chromatographic-Mass Spectrometry Analysis 123 4.3.7 Complete sequence of the insert containing EPA/DHA genes 123 4.3.7.1 Bacterial strains and growth conditions 123 4.3.7.2 Fosmid DNA isolation from a positive clone 124 4.3.7.3 Sequencing of the insert 124
4.4 Results and Discussions 124 4.4.1 Characterization of 5. baltica MAC1 genes homologous to the pfaA, pfaB, pfaC andpfaD synthase gene cluster 124 4.4.2 Amplification of 37 kb EPA gene cluster using thermo stable DNA taq polymerase 129 4.4.3. Cloning of the Shewanella baltica MAC1 EPA/DHA gene cluster by constructing a Fosmid Library 130 4.4.4 Detection of recombinant production of EPA/DHA by positive FOS clones for pfaA andpfaD genes 133 4.4.5 Assessment of recombinant production of EPA/DHA by FOS clones in LB broth at different growth temperatures 135 4.4.6 Sequence analysis of the 5. baltica MAC1 EPA/DHA gene cluster 144
4.5 Conclusions 147
5. General Discussion 149 6. Future directions 155 7. References 156
VI List of Tables
Table 2.1 Primers used to isolate the 16S rRNA from S. baltica LMG 2250, LMG
22253, LMG 2263, MAC1 and S. putrefaciens LMG 2369 37
Table 2.2 Conditions used for PCR 37
Table 2.3 Gas chromatography parameters for fatty acid separation of Shewanella.
baltica strains 41
Table 2.4 Selected phenotypic characteristics of the S. baltica strains 43
Table 2.5 Fatty acid compositions of S. baltica strains at different temperatures 46
Table 3.1 Digestion conditions for BAC DNA 74
Table 3.2 EPA concentration in dry cells of transgenic E. coli cultured at different
media and temperatures 95
Table 3.3 DHA concentration in dry cells of transgenic E. coli cultured at different
media and temperatures 95
Table 4.1 Primers used to locate EPA/DHA genes in the 5. baltica MAC1 genome... 105
Table 4.2 Conditions that were used in PCR 107
Table 4.3 Primers used to amplify the 37 kb EPA gene cluster in the S. baltica MAC1
genome 108
Table 4.4 Fatty acid composition of fosmid clone harboring EPA/DHA gene cluster. 142
Vll List of Figures
Figure 1.1 Polyunsaturated fatty acid synthesis in bacteria 15
Figure 1.2 pEPA plasmid and its restriction map. 22
Figure 2.1 Schematic representation of the 16S rRNA phylogenetic tree for the genus
Shewanella 32
Figure 2.2 Gel electrophoresis of PCR product 44
Figure 2.3 Gas chromatography of total fatty acid methyl esters prepared from
Shewanella baltica 2263 49
Figure 2.4 Mass chromatogram of EPA produced by S. baltica 2263 50
Figure 2.5 EPA concentration in cell dry weight of S. baltica MAC1, LMG 2250, LMG
2263 and LMG 22253 at different temperatures 52
Figure 3.1 Diagram of pIndigoBAC-5, a derivative of pBeloBAC 60
Figure 3.2 Map of pCClBAC cloning vector 68
Figure 3.3 PFGE of partially digested genomic DNA isolated from S. baltica
MAC1 78
Figure 3.4 PFGE of BAC DNA extracted from randomly selected clones from the BAC
Library 80
Figure 3.5 Gas chromatography of three positive clones 82
Figure 3.6 Gas chromatography of positive EPA/DHA clone and standard 83
Figure 3.7 Mass spectra for EPA/DHA standard 85
Figure 3.8 Mass spectra for EPA.DHA positive clone 86
Figure 3.9 Comparision of mass spectra of EPA and DHA standard with mass spectra
viii of EPA and DHA peaks from positive clone 88
Figure 3.10 Mass spectra of S. baltica MAC1 89
Figure 3.11 PFGE of digested and undigested EPA/DHA positive and
negative clones 92
Figure 3.12 Comparision of DHA concentration in transgenic E. coli grown at
different temperatures and different media 96
Figure 4.1 Diagram of a fosmid cloning vector 102
Figure 4.2 Gel electrophoresis of PCR products using different sets of primers 126
Figure 4.3 Gel electrophoresis of mechanically sheared genomic DNA of S. baltica
MAC1 and PCR products using pfaA and pfaD primers 131
Figure 4.4 Gel electrophoresis of undigested and digested fosmid DNA with Not 1.... 132
Figure 4.5 Gas chromatography of the fatty acid profile of positive EPA/DHA
clone and standard 134
Figure 4.6 Mass spectra for EPA/DHA standard 136
Figure 4.7 Mass spectra for EPA/DHA positive clone 137
Figure 4.8 EPA production in EPA/DHA positive clones at 10°C and 15°C 139
Figure 4.9 EPA production in EPA/DHA positive clones at 20°C and 25°C 140
Figure 4.10 Effect of growth temperature on the production of EPA by fosmid
clone 143
Figure 4.11 Shewanella baltica MAC1 EPA/DHA gene cluster and flanking DNA.... 145
IX List of Abbreviation
AA Arachidonic acid
ACC Acetyl coenzyme A carboxylase
ACP Acyl carrier protein
ALA a-Linolenic acid a Alfa
ATP Adenosine tri phosphate
AT AcylTransferase
BAC Bacterial Artificial Chromosome bp Base pair
°C Centigrade
C Carbon
Ca Calcium
CLF Chain Length Factor
Cm Centimeter
COA Coenzyme A
CVD Cardiovascular disease
DHA Docosahexaenoic acid
Dig Digoxigenin
DNA Deoxyribonucleic acid dNTP 2' -deoxyribonucleoside 5"-triphosphate
DPA Docosapentaenoic acid E. coli Escherichia coli
EB Elution buffer
EDTA Ethylenediaminetetraacetic Acid
EPA Eicosapentaenoic acid
ETB Ethidium Bromide
FA Fatty acid
Fab Fatty acid biosynthesis
FAS Fatty acid synthesase
FID Fume ionization detector
FOS Fosmid
F-plasmid Fertility plasmid g Gram g Gravity
GPCT GenomicPrep Cells and Tissue
Y Gamma
GC Gas chromatograph
GC-MS Gas chromatographic-Mass Spectrome h Hourse
H2 Hydrogen
HCL Hydrogen Chloride
HD Hydroxydecarbonyl Dehydratase
HDL-C High-density lipoproteins-cholesterol
HPETEs Hydroperoxyeicosatetraenoic
XI HETEs Hydroxyeicosatetraenoic
HMW Highe Molecular Weight
KAS 3-oxoacyl carrier protein synthase
Kb Kilobase
KC1 Potassium Chloride
KR Ketoacyl Reductase
KS Ketoacyl Synthase
I Lambda
1 Litter
LA Linoleic acid
LB Loading buffer
LBA Lb agar
LBB Lb broth
LDL-C Low-density lipoproteins-cholesterol
LMP Low melting point
LNA Linolenic acid
LPUFA Long Chain Polyunsaturated fatty
LTs Leukotrienes
Lxs Lipoxins m Meter
M Mole
MA Marine agar
MB Marine broth
Xll MCT Malonyl CoA-ACP Transferase mg Milligram
MgCk Magnesuium Chloride
MgSo4 Magnesium Sulfate
Min Minute ml Milliliter mm Milimeter mM Milimole
MUFA Monounsaturated Fatty Acid
NaCl Sodium chloride ng Nano-gram nm Nano meter
OD Optical density
OL Oleic acid
CO Omega
ORFs Open reading frames
Par Partitioning
PCR Polymerase chain reaction
PGs Prostaglandin
Pfa Polyunsaturate fatty acid pmol Pico-molar
PKS Polyketide Synthase
PPS Phosphate Buffered Saline
Xlll PPtase Phosphopantetheinyl Transferase
PUFA Polyunsaturated Fatty Acids
PFGE Pulse Field Gel Electrophoresis
RNA Ribonucleic acid rRNA ribosomal Ribonucleic acid rpm Rotation per minute
S Shewanella
Sec Second
SFA Saturated Fatty Acid
SSC Sodium Chloride-Sodium Citrate
SUM Summary
TAE Tris/acetate buffer
Tag Thermus aquaticus
TB Terrific Broth
TBE Trice/borate buffer
TE Tris- Ethylenediaminetetraacetic Acid
TG Triglycerides
Tn5 Transposon five
Tris-HCl Tris hydrochloride
TXs Thromboxanes
U Unite
UCM Ultra Clean Microbial
UV ultraviolet
XIV (0.g Microgram
(0,1 Microlitter
\iM MicroMole
V Volte
XV 1. General Introduction
1.1 An introduction to Omega-3 fatty acids
a-Linolenic acid (ALA; 18:3co3), eicosapentaenoic acid (EPA; 20:5co3), docosapentaenoic acid (DPA; 22:5co3) and docosahexaenoic acid (DHA; 22:6co3) are long chain polyunsaturated fatty acids (PUFA) which are known as omega-3 fatty acids
(CO-3 FAs). The term (0-3 signifies the position of the first double bond which is in the third carbon-carbon bond from the terminal methyl end of the carbon chain. All naturally occurring co-3 FAs are in the cis- configuration. The co-3 FAs are classified as essential components of the human diet because mammals cannot synthesize them. Linoleic acid
(AL) and ALA are precursors for (0-6 and (0-3 fatty acids, respectively. a-Linolenic acid can be partially converted to the long chain co-3 FAs; albeit at a very slow rate. The conversion rate of ALA to EPA in humans is between 8 to 20%, while its conversion to
DHA is 0.5-9% (Stark et al, 2008). In humans, several desaturase and elongase enzymes are responsible for the synthesis of EPA and DHA. Firstly, ALA is converted to EPA when desaturation enzymes add 2 double bonds to the carbon chain. This is followed by addition of 2 carbon atoms to the chain by elongation enzymes. Eicosapentaenoic acid is capable of being elongated to DPA, which finally can be converted to DHA (Stark et al,
2008). Today co-3 FAs are known as the "good fat" and they have generated much media attention. They are also a hot topic in the research area. What has made these polyunsaturated fatty acids so important? 1.2 The importance of omega-3 fatty acids
co-3 FAs have been the topic of extensive scientific research for over 25 years.
Much has been written in recent years on the remarkably wide ranging physiological effects of the co-3 FAs and their roles in human health (Krebs et al, 2006; Siddiqui et al,
2004; Siddiqui et al, 2008b). They have been described in terms such as the 'missing link in nutrition' (Borek, 1994) with 'miraculous' clinical effects. To date, most clinical trials with human subjects consistently show that consuming co-3 FAs impacts diverse physiological functions such as: eicosanoid synthesis, alteration of membrane fluidity, effects on signal transduction, changes in gene expression and effects on intraluminal bacteria. Among these, alteration of eicosanoids has received more attention.
Eicosanoids are highly active lipid mediators in physiological and pathological processes.
They are a family of 20 carbon-oxygenated derivatives of ALA, arachidonic (AA;
20:4co6), and EPA. Eicosanoids include prostaglandins (PGs) and thromboxanes (TXs) which are termed prostanoids, as well as leukotrienes (LTs) lipoxins (Lxs), hydroperoxyeicosatetraenoic acids (HPETEs) and hydroxyeicosatetraenoic acids
(HETEs) (Sellmayer & Koletzko, 1999). These compounds affect blood pressure, blood clotting, inflammation, immune function and coronary spasms. The eicosanoid metabolic products from AA such as prostagalandin E2; thromboxane A2, a potent platelet aggregator and vasoconstrictor; and leukotriene B4, a strong inducer of inflammation; contribute to the development of inflammatory disorders and cell proliferation
(Simopoulos, 2000; Simopoulos, 2002). The ingestion of EPA partially replaces the AA in cell membranes, particularly those of neutrophils, monocytes, platelets and erythrocytes. Therefore, the production of prostagalandin E2, thromboxane A2 and
2 leukotriene B4 decrease. It is well documented that o>3 FAs produce a series of eicosanoids such as thromboxane A3, a weak platelet aggregator; prostagalandin PGI3, an inhibitor of platelet aggregation; and leukotriene B5, a weak inducer of inflammation, that decreases the risk of heart disease and inflammation (Fontani et al, 2005; Jolly, 2005).
Further studies demonstrated the involvement of DHA in the normal function of the brain and retina (SanGiovanni & Chew, 2005). DHA is the major PUFA accumulated in the cell membrane of the brain and constitutes more than 30% of fatty acids in the brain. Therefore, it is required at high levels in the brain to provide an optimal brain function (Innis, 2008). In the retina, DHA comprises over 60% of the total fatty acids.
Several research and clinical studies have proved that DHA is essential for the growth and development of the brain in infants (Innis, 2008; Mitmesser & Jensen, 2007). EPA plus DHA also have beneficial effects in human growth and development (Heird &
Lapillonne, 2005; Innis, 2008; Singh, 2005) and in the immune system (Nakamura et al,
2005; Vedin et al, 2008). The beneficial health effects of EPA and DHA have also been well documented with respect to obesity (Delarue et al, 2004; Parra et al, 2008; Voss et al, 2008) and depression (Bourre, 2005; McNamara et al, 2008). Furthermore, there are some reports that co-3 FAs have beneficial effects on patients with HrV (Metroka et al,
2007; Wohl, 2007). According to the 2008 report of the World Health Organization
(Cohen et al), 68% of people die from three diseases in which the involvement of fatty acids has been implicated; viz. cardiovascular disease (CVD) (43.8%), cancer (22,4%) and diabetes (1.8%). EPA plus DHA have beneficial effects on the prevention, management and treatment of cardiovascular diseases (Yamagishi et al, 2008), cancer
3 (Jolly, 2005; Khal & Tisdale, 2008) and type 2 diabetes (Hartweg et ai, 2008; Nettleton
& Katz, 2005; Steyn et ai, 2004).
1.2.1 Omega-3 fatty acids and cardiovascular disease
Heart disease includes any conditions that compromise heart function. Terms closely related to heart disease, having specific medical definitions include cardiovascular disease, coronary artery disease, ischemic heart disease, atherosclerosis, thrombosis and heart attack. Cardiovascular disease is a multifactorial disease in which genetics and environment interact.
Omega-3 FA, particularly EPA and DHA, appear to reduce the risk of dying from
CVD. There are several possible ways that omega-3 FA may reduce the risk of CVD.
Firstly, they appear to have a modest blood pressure lowering effect (Mori et ai, 1999;
Nestel et ai, 2002). It has been proposed that co-3 FAs are incorporated into cell membranes, modifying their physiological properties and affecting ion transport and signal transduction (Sassen et ai, 1994). Secondly, co-3 FAs seem to improve blood lipid profiles. Lipid attaches to a variety of water-soluble substances in blood and tissues. By attachment to proteins and phospholipids, lipids become soluble in water and are transported in the blood and across cell membranes. The most important lipids are triglycerides, cholesterol and other sterols, phospholipids, and fatty acids. All these lipids are constituents of lipoproteins and cell membranes. Lipoproteins are spherical particles with a lipid core composed mainly of triglyceride, cholesteryl ester and a surface layer containing phospholipids, cholesterol and protein. There is no longer doubt that high
4 blood cholesterol levels greatly increase the risk of CVD and premature death. Total
blood cholesterol measurements, however, do not reveal the distribution of cholesterol
between the low- and high- density fractions. The distribution of cholesterol among the
major lipoproteins determines the atherogenicity of circulatory cholesterol. High levels of
low-density lipoprotein-cholesterol (LDL-C) greatly increase the risk of CVD (Hulthe et
al, 2000). These lipids narrow the blood vessel diameter and restrict blood flow. In
contrast high levels of high-density lipoprotein-cholesterol (HDL-C) protect against
CVD. Omega-3 FA in the form of fish or fish oil usually decrease blood triglycerides
(TG) and LDL-C levels, while HDL-C remains unaltered. EPA and DHA reduce blood
triglycerides by inhibiting both TG and apolipoprotein B synthesis in the liver (Laidlaw
& Holub, 2003; Stark et al, 2000). Thirdly, in regard to blood coagulation and platelet
function, it is reported that the vasoactive and aggregatory factors (thromboxane A2,
ADP and serotonin) liberated in the process are involved in platelet aggregation and
thrombus formation through activation of the coagulation cascade (Semplicini & Valle,
1994; Wada et al, 2007). o>-3 FAs alter vascular function by additional mechanisms
including changes in the release of ADP, a potent platelet aggregator, and also inhibit the
production of TXA2 (Caramia, 2008).
Finally, the impact of co-3 FAs in stabilizing myocardial membranes and reducing
susceptibility to ventricular arrhythmias is a primary mechanism by which they reduce sudden cardiac death (Mozaffarian et al, 2008). The latter effect may be related to heart rate reduction by 0>-3 FA (Mozaffarian et al, 2005). Other effects of (0-3 FAs include improvement of the oxygen supply to tissues, a moderate reduction in arterial blood
5 pressure, and relaxation of coronary arteries, all of which may contribute to the prevention of atherosclerosis.
1.2.2 Omega-3 fatty acids and cancer
There is both epidemiological and experimental evidence to suggest that (0-3 FA exert protective effects against some common cancers, notably those of the breast, colon, and prostate. Multiple mechanisms are involved in cancer preventive activity using
EPA/DHA, including suppression of neoplastic transformation, cell growth inhibition and enhanced apoptosis, and antiangiogenicity (Berquin et ah, 2008). EPA and DHA have effects on both cancer cells (proliferation, invasion, metastasis and apoptosis) and host cells (inflammation, immune response, and angiogenesis) (Berquin et ah, 2008). The exact mechanism by which dietary omega-3 FA inhibits cancer development is not completely understood.
It is currently believed that the omega-3 FA changes the composition of tumor membrane lipids by partial replacement of AA. Therefore, the biosynthesis of co-6 series eicosanoids is reduced. Eicosanoids in general modulate cell growth and differentiation, inflammation, immunity, platelet aggregation, and angiogenesis (Rose & Connolly,
2000). In particular PGE2 appear to suppress the anti-tumor activity of the host defense.
Prostaglandins suppress immune responses that could facilitate cell proliferation and oppose the immune system's usual antitumor response (Kobayashi et ai, 2006).
Reducing PGE2 production by omega-3 FA would be expected to facilitate restoration of
6 antitumor activity and retard tumor development (Kobayashi et ai, 2006). A further role for eicosanoids is suggested by the involvement of thromboxane A2 in lymphocyte transformation. Lipoxygenase products such as 5-HETE inhibit lymphocyte transformation, whereas other polyhydroxy eicosanoids may promote transformation
(Calder et ai, 1992). Omega-3 FA also reduces the production of the cytokines interleukin-1 and -2 and tumor necrosis factor from mononuclear cells (Guijarro et al,
2006). These are part of the immune response and host defense system.
Omega-3 and omega-6 FAs have also different effects on the signaling pathways which are relevant to carcinogenesis and tumor progression. It is reported that high intakes of G>-6 FA up-regulate and activate cellular signaling mediators such as protein kinase C and NF-kB, while co-3 FA has the opposite effect (McCarty, 1996). Changes in the signaling pathway have an effect on the transcription factor, which alters gene expression. However, the exact mechanism of alteration in signaling and transcription is not clearly established. It is suggested that changes in the lipid composition of the membrane may affect membrane fluidity and the way that growth factors, cytokines and hormones interact with their receptors. Changes in membrane composition have been specifically linked to the activity of the membrane-bound enzyme, protein kinase C that has been implicated in tumor promotion (Guillem et ai, 1987). Omega-3 may also regulate the translation machinery. It is reported that EPA affects intracellular homeostasis. EPA induces Ca2+ release from the intracellular Ca2+ stores and simultaneously inhibits Ca2+ flow through store operated Ca2+ channels in the membrane.
This results in reduction of the intracellular Ca2+ stores and activation of IF2a kinase,
7 which can inhibit translation initiation. It is also reported that the ability of EPA and
DHA to induce apoptosis in tumor cells is a result of increased susceptibility of the tumor cells to lipid peroxidation (Stoll, 2002). The tumor suppressive effect of EPA/DHA in human breast cancer and colon cancer cell lines was blocked in the presence of anti oxidants (Berquin et al, 2008).
In general, activation of the immune system, prostaglandin synthesis, membrane fluidity, mammogenic hormone secretion, hormone and growth factor activity, and intercellular communication are some effects of EPA/DHA resulting in suppression of tumor cells.
1.2.3 Omega-3 fatty acids and diabetes
Many studies have been conducted to investigate the effect of co-3 FA in diabetic patients, particularly in type II diabetes mellitus. This type of diabetes increases the level of tryglyceride and LDL-cholesterol in blood and also elevates insulin resistance, leading to increased glucose levels (Cannon, 2008). In some studies, it was reported that co-6 and co-3 FAs may play a beneficial role in insulin resistance (Hartweg et al, 2008). These
FAs direct glucose toward glycogen storage, and direct fatty acids away from triglyceride synthesis and assimilation and toward fatty acid oxidation (Woodman et al, 2002).
Moreover, co-3 FAs have the unique ability to increase thermogenesis, decrease body fat position and improve glucose clearance. The effects of FAs on lipid metabolism and thermogenesis are exerted by up regulating the transcription of mitochondrial uncoupling protein-3, and inducing genes encoding proteins involved in fatty acid oxidation. On the
8 other hand, EPA/DHA simultaneously down-regulate the transcription of genes encoding proteins involved in lipid synthesis (Clarke, 2001; Minami et al, 2002). Another study indicated that EPA, in particular, reduced plasma lipids, hepatic triacylglycerols, insulin resistance and abdominal fat deposits (Minami et al, 2002). In addition, EPA/DHA have beneficial effects on serum HDL-cholesterol, lipid peroxidation and antioxidant enzymes, which may lead to a decreased rate of occurrence of vascular complications in diabetes
(Barre, 2007).
1.3 Sources of omega-3 fatty acid and recommended dietary intakes
for the omega-3 fatty acids
Linseed oil, rapeseed oil, flax seed, walnuts and soybeans are good dietary sources of ALA. The primary source of EPA and DHA are fungi, marine algae, such as
Porpyridium and Nannochloropsis (Patil et al, 2007), diatoms and certain strains of marine bacteria (Gentile et al, 2003) which, upon consumption, are passed up the food chain into fish. According to Hau and Gralnick, some marine bacteria, such as those belonging to the genus Shewanella, are the major EPA and DHA suppliers for higher organisms in aquatic environments (Hau & Gralnick, 2007). Currently, the main human dietary sources of EPA and DHA are fish oil and marine fish such as salmon, tuna, halibut, mackerel, sardine and herring.
Depending on gender, age, culture and health-status, the recommended consumption of ALA, EPA and DHA is quite variable. For example, the U.S. Food and
Nutrition Board recommendation for ALA intake by healthy children, adolescents and
9 adults is 0.5 gram/day, 0.8 gram/day and 1.3 gram/day, respectively. For EPA and DHA, the recommended intake varies from 500 mg/day for children to 700 mg/day for healthy adults. The American Dieteitic Association and the Dietitians of Canada recommends
500 mg/day EPA plus DHA. The American Heart Association and Food and Drug
Administration recommendation for EPA/DHA is 2 to 3 servings of fish per week or 900 mg/day for those with coronary heart disease. For healthy individual, the recommendation intake is 650 mg/day combined EPA/DHA for males and 410 mg/day for females. The European Commission has set no minimum or maximum intake for
ALA consumption while they believe that a maximum safe level for EPA and DHA intake must be established. The U.S. Food and Drug Administration recommends that omega-3 intake should not exceed more than a total of 3 grams/day. In North America
EPA and DHA are consumed at the level of approximately 50 and 80 mg/day, respectively. In addition, the low intake of 19 mg/day DHA in young children in North
America is a particular concern.
1.4 An Introduction to Shewanella Species
The Shewanella genus was named after James Shewan for his work in fisheries microbiology (Shewan et al., 1960). The original member of the genus, now called
Shewanella putrefaciens, was isolated from rancid butter in 1931 by Derby & Hammer.
They identified the isolated microorganism as Achromobacter putrefaciens. Ten years later it was renamed Pseudomonas putrefaciens (Hammer & Long, 1941) according to its biochemical characteristics. It was reassigned from Pseudomonas putrefaciens to the genus Alteromonas putrefaciens (Lee et al, 1977) based on moles percent guanine plus
10 cytosine (G+C) of DNA. Finally, MacDonell & Col well (1985) reclassified Alteromonas putrefaciens into a new genus, Shewanella, on the basis of 5S rRNA sequence. Since
1985, the birth date of Shewanella, no further reclassifications have been made. The genus Shewanella, originally included the species Shewanella putrefaciens, S. benthica,
S. baltica, S. hanedai, S. colwelliana and S. frigidimarina (Bowman et al., 1997; Hau &
Gralnick, 2007; MacDonell & Colwell, 1985; Weiner et al., 1988). However, advanced taxonomic techniques, such as PCR technology, have allowed the identification of new
Shewanella species. A number of novel species have been recently introduced, including:
S. affinis (Ivanova et al., 2004) ,5. pneumatophori (Hirota et ah, 2005), S. hafniensis and
S. morhuae (Satomi et al., 2006). On the basis of DNA-DNA hybridization and 16S rRNA approximately 45 species are assigned to the genus Shewanella. Members of the genus Shewanella are widely distributed in nature, especially in marine and fresh water environments and sediment; in the deep sea, especially cold regions such as polar areas; and in the intestine of marine fish (Nichols et al, 1997; Valentine & Valentine, 2004).
The genus Shewanella includes a diverse group of gram-negative, facultative anaerobic, rod-like (2-3[xm in length and 0.4-0.7[Am in diameter), and motile bacteria
(Hau & Gralnick, 2007). The species of Shewanella are known as food spoilage organisms, symbionts, epibionts, opportunistic pathogens and beneficial microorganisms.
In addition, a vast variety of organic and inorganic compounds can be respired by
Shewanella. It is one of the most important characteristics of Shewanella, which allows them to exist everywhere and to survive in harsh environments. This genus has been studied for at least 70 years. The Shewanella species have been of interest to both applied
11 and environmental microbiologists. In addition, this genus has been studied for their important roles in the dissimilatory reduction of manganese and iron oxides (Myers &
Nealson, 1988) and in the spoilage of proteinaceous foods (Vogel et al, 2005), but the most recent studies have investigated the ability of this genus to produce omega-3 FAs
(Frolova et al., 2005; Satomi et al., 2003; Yazawa, 1996; Yu et al., 2000).
1.5 The genus Shewanella and Omega-3 fatty acid production
EPA and DHA production in bacteria isolated from deep sea water and sediment was first documented in 1986 (Delong & Yayanos, 1986). Several strains of marine bacteria belong to the genus Shewanella, Colwellia, Vibrio and Flexibacter (Bowman et al, 1998a; Bowman et al, 1998b; Russell & Nichols, 1999) are recognized as strong biological primary sources of EPA and DHA. Colwellia and Shewanella are the two main genera contributing to the production of (0-3 FAs in the marine environment. However, several Shewanella strains have been found to produce high levels of EPA and DHA
(Allen & Bartlett, 2002; Russell & Nichols, 1999). Wilkinson and Caudwell concluded that the fatty acid compositions of all S. putrefaciens groups contained some unusual fatty acyl components; the major ones being 16:ln7cis, isol3:0, isol5:0, 15:0 and 17:ln8cis, but co-3 FA production was not found (Wilkinson & Caudwell, 1980). EPA-production by S. putrefaciens was first reported in 1987 (Wirsen et al, 1987). A report in the following year (Yazawa et al, 1988) described the isolation of an EPA-producing marine bacterium (strain SCRC-2738) which was classified as S. putrefaciens. Furthermore, later publications indicated production of EPA and DHA by S. putrefaciens strains (Yazawa,
1996; Jostensen, 1996). Shewanella frigidimarina, S. gelidimarina, S. benthica, S.
12 hanedai, S.marinintestina, S.schlegeliana, S.sairae, S.pneumatophori and S.baltica were found to contain varying proportions of EPA and DHA (Bowman et al., 1997; Hirota et al, 2005; Russell & Nichols, 1999; Satomi et al, 2003).
Since co-3 FAs-producing bacteria were mostly isolated from deep, cold sea environments, it was suggested that low temperature and high pressure act as two important factors for the evolution of bacteria that incorporate co-3 FAs, particularly EPA and DHA, into their membrane phospholipids (Allen & Bartlett, 2002). In addition, the synthesis of co-3 FA is more prevalent in species with low growth temperature optima, particularly S. hanedai and 5. benthica (Bowman et al, 1997; Hau & Gralnick, 2007).
The production of EPA/DHA by some species of Shewanella increases as temperature decreases; leading to the hypothesis that EPA/DHA may be essential for growth at low temperature (Wang et al., 2008). This phenomenon could be an adaptation to low temperature by modulating the lipid composition of the membranes in order to maintain the homeoviscosity of cellular membranes and to make them more permeable in cold and high-pressure environments (Gentile et al., 2003; Russell & Nichols, 1999).
1.6 Mechanism of polyunsaturated fatty acid biosynthesis in bacteria
Pathways for Poly-Unsaturated Fatty Acid (PUFA) synthesis are common in all microorganisms. All bacteria use the fatty acid synthase (FAS) pathway and acetyl-CoA carboxylase (ACC) for fatty acid production (Ratledge, 2004). The first committed reaction in fatty acid synthesis is the carboxylation of acetyl-CoA to malonyl-CoA.
Palmitate can then be synthesized by a series of reactions catalyzed by a FAS system
13 from one molecule of acetyl-CoA (the primer) and 7 molecules of malonyl-CoA. After cycles of condensation, reduction, dehydration and reduction for each 2C-unit added from malonyl-CoA, the end product of fatty acid synthase, palmitic acid (C16:0), is made which is subsequently elongated to stearic acid (CI8:0). This fatty acid becomes a backbone for PUFA production and it is then modified through a sequence of desaturases and elongases to produce both unsaturated and polyunsaturated fatty acids. The position of the double bound is fixed relative to the carboxyl end of the substrate molecule. In bacteria that produce PUFA, there are distinct desaturases for inserting double bonds at different positions which are separated by a -CH2 unit (Calder, 1997). The desaturases would have to act in concert with an elongation system consisting of a set of specific elongases to make EPA and DHA. A series of elongation and desaturation steps result in the production of co-3 and co-6 polyunsaturated fatty acids (Pereira et al, 2004). In addition, it is the location of first double bond, counting from the methyl end of the fatty acid molecule, which distinguishes these classes. The synthesis of EPA and DHA from acetyl-CoA requires approximately 30 distinct enzyme activities and 70 reactions (Metz et al, 2001). The pathway for EPA and DHA production is shown in Figure 1.1
Metz et al. (Metz et al, 2001) described the polyketide synthase (PKS) pathway for EPA and DHA biosynthesis in Shewanella pneumatophori SCRC-2738. Unlike the
FAS pathway, it does not require desaturation and elongation of saturated fatty acids.
PKS carry out some of the same reactions as FAS and use acyl carrier protein as a
14 Oleic acid (18:ln9)
co-3 Fatty acids o>6 Fatty acids
1 02, A12 desaturase oc-Linolenic acid (18:3n3) Al5 desaturase Linoleic (18:2n6) I " 1
A6-Desaturase OctadecatetraenoiI c (18:4) y-Linolenic (18:3) A6-Elongase
Eicosatetraenoic (20:4) Dihomo-y-Linolnic (20:3)
A5-Desaturase
Eicosapentaenoic (20:5) o>3 Desaturase Arachidonic (20:4)
Docosapentaenoic (22:5) Docosatetraenoic (22:4)
• A4-Desaturase •
Docosahexaenoic (22:6) Docosapentaenoic
co-3 Desaturase
Figure 1.1. Polyunsaturated fatty acid synthesis in bacteria (Pereira et al., 2004).
15 covalent attachment site for the growing carbon chain. However, PKS enzymes catalyze the synthesis of PUFA by the repetitive condensation and the cycle of reduction, dehydration and reduction is shorter than that of FAS. Other recent reports indicate that
EPA and DHA are synthesized de novo in bacteria via the PKS pathway (Metz et al,
2001; Orikasa et al, 2006a; Orikasa et al, 2006b). It is suggested that the PKS pathway employs several small protein domains, which are homologues to FAS enzymes. These domains of PKS are: 3-ketoacyl synthase (KS), malonyl coenzyme A:acyl carrier protein, acyltransferase, 3-ketoacyl ACP reductase (KR), acyltransferase (AT), 3- hydroxydecarbonyl-ACP dehydratases (HD), enoyl reductase (ER), Chain Length Factor
(CLF) and phosphopantetheinyl transferase (PPTase) (Metz et al, 2001; Orikasa et al,
2006a). Despite considerable advances in our understanding of the mechanism of PUFA production, the regulation of PUFA synthesis is still not defined completely.
Mammals such as human are able to convert 18:0 to 18:1 (n-9) using a membrane
-bound 18:0 desaturase, however, they lack A4 and A12 desaturase activities. Therefore, they require LA (18:2n6) and ALA (18:3n3) in their diet as substrates for the elongation machinery. These fatty acids are converted to AA, EPA and DHA via a series of desaturation and elongation reactions in the endoplasmic reticulum. Morover, the synthesis of DHA is known to consist of two succeeding elongation cycles, a A6 desaturation and a p-oxidation chain-shortening.
16 1.7 Genetic Studies in Shewanella Species
As mentioned previously, fatty acids in bacteria are synthesized by the FAS or
PKS routes; meaning that the individual reactions are catalyzed by separate enzymes that are encoded by different genes. Half of the fatty acid biosynthetic genes are scattered around the chromosome, the remaining half are present as a fab gene cluster (Morita et al, 2000). As the field of molecular biology advanced and techniques for finding gene(s) and manipulating them were developed, researchers all around the world started to apply this knowledge to marine bacteria in the hopes of finding the EPA and DHA gene(s) and expressing the gene(s) in industrially relevant organisms. To date, the genes responsible for EPA and DHA synthesis have not been fully defined. However, a 38 kb cluster of genes involved in the biosynthesis of EPA (pfa genes) in Shewanella putrifaciens SCRC-
2738 has been identified (Yazawa, 1996). This organism has been reclassified as
Shewanella pneumatophori SCRC-2738 (Hirota et al., 2005). The 38 kb fragment carries at least 18 open reading frames (ORFs), of which 9 were thought to be necessary for EPA production (Yazawa, 1996). Following cloning and sequencing of the 38 kb genomic
DNA, it was suggested that 8 open reading frames in the 38 kb genome DNA are involved in EPA synthesis (Yazawa, 1996). In the following year, the 38 kb gene cluster involved in EPA synthesis was cloned and expressed in the marine bacterium
Cyanobacterium synechococcus (Takeyama et al, 1997). The DNA sequence of the EPA biosynthesis gene cluster has the accession number U73935 (www.ncbi.nlm.nih.gov/).
Since 1996, several attempts have been made to isolate the EPA gene cluster from EPA- producing bacteria using the EPA published sequence. In 2002, Allen and Bartlett (Allen
& Bartlett, 2002) prepared a genomic DNA library of Photobacterium profundum SS9,
17 using Yazawa's method. They constructed primers based on the ORF9-3 and ORF9-2 sequences derived from Shewanella SCRC-2738. They obtained a 33 kb locus which had four ORFs for EPA biosynthesis. However, they were unable to achieve EPA biosynthesis in E. coli following recombination. In a more recent study Orikasa et al. reported that five ORFs, now named pfaA, pfaB, pfaC, pfaD and pfaE, are essential for
EPA production (Orikasa et al, 2004). It is important to mention that the sequential arrangement of these pfa genes is broken by the two ORFs unrelated to EPA biosynthesis.
Gentile et al. (Gentile et al, 2003) partially cloned pfa A from Shewanella sp.
GA-22. Moreover, a fab gene cluster, homologous to EPA genes, was cloned from the
DHA-producing bacterium, Moritella marina strain MP-1 (Morita et al, 2000). Since the fab gene cluster cloned from Moritella marina MP-1 had only pfaABCD, their attempt at recombinant production of EPA and DHA in E. coli was not successful. Several attempts have been made to clone EPA and DHA genes from other bacteria such as
Pseudoalteromonas sp. strain DS-12, Shewanella marinintestina IK-1, Shewanella sp. strain SC2A, Shewanella oneidensis MR-1, and Colwellia psychrerythraea 34H (Methe et al, 2005; Okuyama et al, 2007; Venkateswaran et al., 1999). In these studies, a cluster of pfa genes similar to the pfa genes from Shewanella SCRC-2738 in size, in the number of ORFs and domain structure of individual ORFs was cloned. However, transgenic E. coli was not able to synthesize any recombinant EPA or DHA (Okuyama et al, 2007).
18 Yazawa (Yazawa, 1996) reported the only successful recombinant production of
EPA in E. coli in 1996. Although the pja genes of Moritella marina MP-1 were the first genes to be cloned from DHA-producing bacteria, the lack of pfaE prevented recombinant production of DHA in E. coli (Morita et al, 2000). However, recently, the pfaE gene was cloned from Moritella marina MP-1 and, when expressed with pfaABCD, resulted in the recombinant production of DHA (Orikasa et al., 2006a; Orikasa et al.,
2006b). It is well documented that the presence of the pfaE gene, which encodes for phosphopantetheinyl transferase (PPTase), is necessary for the production of EPA and
DHA (Orikasa et al, 2006a; Sugihara et al., 2008). Transgenic E. coli carrying the plasmid containing genes pfaABCD but lacking the pfaE gene resulted in no EPA nor
DHA production. When the pfaE gene cloned from the EPA-producing Photobacterium profundum strain SS9 was co- expressed with pfaA-D genes derived from Moritella
Marina MP-1, the transgenic E. coli was able to produce DHA (Sugihara et al., 2008).
On the basis of these findings, it is suggested that the DHA biosynthesis gene cluster should have the same gene organization as that of the EPA cluster (Orikasa et al., 2006b).
Therefore, the genes pfaA, pfaE, pfaC, pfaD and pfaE are involved in biosynthesis of both EPA and DHA.
According to a recent article (Okuyama et al, 2007), the pfaA gene encodes a multifunctional protein with domains of 3-ketoacyl synthase (KS), Acetyl transferase
(AT), malonyl coenzyme A: acyl carrier protein, 6 repeats of ACP and 3-ketoacyl -ACP reductase (KR). The gene pfaB encodes the protein domain of AT. The pfaC gene encodes proteins with 2 KS repeats, 3 repeats of 3-hydroxydecanoyl-ACP dehydratases
19 (HD), and chain length factor (CLF). The pfaD encodes a protein with the domain of enoyl reductase (ER). The gene pfaE encodes PPTase. Although the basic structure and the domain structure of the EPA and DHA gene cluster are the same, there are some slight differences among the pfaA-D genes of Moritella marina MP-1, Shewanella sp.
SCRC2738 and Photobacterium profundum SS9. ThepfaB of M. marina MP-1 has a KS domain which is not present in the pfaB gene product of Shewanella sp. SCRC2738 and
P. profundum SS9. Moreover, (Tanaka et al, 1999) reported that the pfa A gene has five repeats for ACP domains in MP-1 while Orikasa et al. (2004) reported 6 repeats of the
ACP domain for Shewanella SCRC 2738. However, these differences did not affect the final production of EPA or DHA by MP-1 and SCRC 2738 because the pfa A gene of P. profundum SS9, an EPA-producing bacterium, has 5 repeats for ACP (Allen & Bartlett,
2002). In conclusion, all the five genes (pfaA, pfaB, pfaC, pfaD and pfdE) and their domains are necessary for the biosynthesis of both EPA and DHA.
1.8 Methods for cloning the omega-3 fatty acid genes
In 1996, Yazawa was the first to clone the EPA genes (Yazawa, 1996). He constructed a genomic DNA library of the chromosomal DNA of Shewanella pneumatophori SCRC-2738. The DNA was partially digested with the restriction enzyme
Sau3AI. The fragment, which was larger than 20kb, was recovered and ligated to the cosmid pWE15. The cosmid size was 8.8kb. The cosmid with the ligated DNA was packaged into the phage heads using GIGAPACK II XL packaging extract and then transfected into E. coli AGI. The transformants were chosen according to their ampicillin resistance and they were tested for EPA production. After analyzing 390 clones, only one
20 clone was able to produce EPA; indicating that the clone had all the genes required for
EPA biosynthesis. The plasmid carrying the gene cluster was designated pEPA. The 38 kb genomic DNA fragment obtained after digestion of the plasmid pEPA was cloned in the BamHI site of the cosmid pWE15. The restriction map of pEPA is shown in Figure
1.2. In the following year, the 38 kb EPA gene cluster was excised from pEPA using the
Sau3Al restriction enzyme (Takeyama et ah, 1997). The excised fragment was ligated to a broad host range cosmid vector, pJRD215, packaged into bacteriophage lambda particles and then transfected into E. coli NM554. The transformants were selected based on kanamycin resistance. The plasmid pJRDEPA (48kb) was transferred to a marine
Cyanobacterium synechococcus by conjugation. The production of EPA by the cyanobacterial transconjugants grown at 17, 23 and 29°C was less than 0.5, 0.4 and 0.1 % of total fatty acid, respectively.
In 1999, several attempts were made to isolate the EPA gene cluster from an
EPA-producing bacterium identified as Shewanella putrefaciens MAC1 based on the published sequence of the EPA biosynthetic gene cluster (Wang et al., unpublished data).
They designed many primers based on the published DNA sequence and PCR was performed under different conditions. In addition, the entire 38 kb DNA sequence published by Yawaza, (1996) was scanned for restriction sites, and several unique sites such as Pmel, Aatll and Xhol were found. Consequently, a restriction map was created and all unique restriction enzymes were used to directly isolate the DNA fragment
21 8.8 kb cosmid vector i
37.8 kb DNA fragment from Shewanella SCRC 2738
$2
Xb 6 £ Xb K4 S4 S3 S3
' i m*m ' KI.T3
Figure 1.2. pEPA plasmid and its restriction map (Yazawa 1996; Takeyama et
ai, 1997). Restriction sites: S2, Sau3Al; S3, Spel; S4, Sphl; Xb, Xbal;
Xh, Xhol; E, EcoRl; N, Noil.
22 responsible for EPA biosynthesis from the genomic DNA of Shewanella putrefaciens
MAC1. Furthermore, partial digestion of the genomic DNA with Sau3AI was studied, using the same method as Yawaza (Wang et al, unpublished data). However, the EPA gene cluster was not detected in Shewanella putrefaciens MAC1 by any of the above methods (Wang et al, unpublished data).
In another study (Tanaka et al, 1999), a cosmid library was constructed from the genomic DNA of Vibrio marinus MP-1, a DHA-producing bacterium. Genomic DNA was digested partially using SauiAl as described by Yazawa (Yazawa, 1996). The digested DNA was fractionated and fragments with a size of 50 kb were ligated to the
Lorist 6 cloning vector and then packaged into the X phage particles. E. coli Xl-1 Blue and DH5a were used as host strains for transformation. Kanamycin-resistant colonies were selected for the library. To screen the cosmid library, primers were synthesized based on the conserved sequences in the 3-oxoacyl carrier protein synthase (KAS) and malonyl CoA-ACP transacylase (MCT) genes of E. coli. The expected KAS/MCT fragment with the size of 710bp was amplified and the whole sequence showed 39% homology to ORF 5 of the EPA cluster from Shewanella pneumatophori SCRC-2738.
The cosmid library was screened by colony hybridization using the radio-labeled
KAS/MCT fragment. They obtained 2 positive colonies out of 245 colonies screened
(Tanaka et al, 1999). One of them contained a 40kb insert, designated as p3D5. The organization of the ORFs in the 40kb fragment was similar to that of the EPA cluster.
23 In 2000, Morita et al. prepared a ADASH II phage library and a Lorist 6 cosmid library from chromosomal DNA of Moritella marina MP-1, a DHA-producing bacterium
(Morita et al, 2000). Two positive clones were obtained. One of them had an insert of approximately 40 kb. Twenty-two putative ORFs were present in the insert DNA. Among them ORFs 8, 9, 10 and 11 were similar to ORFs 5, 6, 7 and 8 of Shewanella pneumatophori SCRC-2738.
In 2002, Allen and Bartlett prepared a genomic fosmid library of Photobacterium profundum SS9 (Allen & Bartlett, 2002). They extracted a high molecular mass genomic
DNA in agarose plugs, which was partially digested with Sau3Al. DNA fragments between 35-45 kb were purified and were ligated into the BamHI site of the pFOSl vector. The fosmid was packaged and then transfected to E. coli DH10B. Approximately
960 fosmid clones resistant to chloramphenicol were selected. Two probes were prepared using the published sequence of the EPA gene cluster for primer design. The PCR products with the size of 571 bp and 885 bp showed high similarity to the SCRC-2738 pfaA and pfaD genes. Probes were made with the amplified fragments and used in colony hybridization. From 42 clones that hybridized to both probes, a 33 kb clone was selected for sequencing.
In 2003 Gentile et al., designed a set of primers from the conserved regions within the pfaA-pfaC gene cluster of Shewanella SCRC-2738, Moritella marina MP-1 and
Photobacterium profudum SS9. The primers were used to amplify the pfaA and pfaC
24 gene from genomic DNA of Shewanella GA-22. They were able to amplify a part of putative pfaA andpfaC genes (Gentile et al., 2003).
In 2004 Orikasa et al. described Yazawa's work in detail and, additionally, they subcloned a 27kb region from Xhol to Spel sites (ORF1-9) of the 38 kb fragment isolated by Yazawa, into pBluescript and pNEB193 vecctors (Orikasa et al., 2004). In total, six types of deletion clone of pEPA were constructed and E. coli JM 109 was used to examine the production of EPA. Their aim was to study the involvement of each ORFs necessary for EPA biosynthesis.
In conclusion, all the groups involved in the hunt for PUFA synthetic genes either fragmented the genomic DNA to smaller sizes (from 25 to 50 kb) using restriction enzymes or amplified some parts of the genes and ligated the DNA fragments to a vector
(plasmid, cosmid and fosmid), which was transferred to different strains of E. coli. They screened the genomic DNA library by testing the clones for the production of EPA and
DHA, making probes and performing hybridization, and finally using PCR to identify the genes which were involved in EPA/DHA production. However, in spite of all the efforts that have been made by different researchers worldwide over more than a decade, only
Yazawa (1996) was able to clone EPA gene cluster; although recently, Orikasa et al.
(2006) co-expressed the pfaA-D cluster, cloned by Morita et al. (2000) with pfaE in E. coli. The result was the production of recombinant DHA by E. coli in the amount of 3% of total fatty acids. They have reported that the combination of two plasmid vectors carrying pfaA-D and pfaE are required for DHA production. However, the exact
25 sequence of the DHA producing gene cluster showing all the ORFs involved in the production of DHA has not been reported yet.
1.9 Rationale
Long chain polyunsaturated fatty acids, particularly EPA and DHA, are essential to human health. Oils extracted from fish and microalgae are the main source of co-3
FAs. Fish populations are declining worldwide and fish oil may have an unpleasant smell and taste, which limits its use by the food industry. In addition, most human exposure to mercury occurs through the intake of fish. Contamination of some fish with methyl mercury and lead has raised safety concerns. High levels of lead and mercury may result in brain damage, kidney damage, depression, anxiety and psychological disorders
(Castoldi et ah, 2003). Marine microalgae are another suitable candidate as a dietary source of EPA/DHA because of their lack of fishy flavor. However, due to very slow growth rate and complicated harvesting process, commercial production of EPA/DHA from microalgae in large scale is very costly. For instance, only the harvesting costs can acount for up to 33% of total production costs because of the large volume of water that must be processed to harvest the algae. There are omega-3 containing eggs and milk on the market but since chickens and cows have been fed with flaxseed or fishmeal, they cannot be considered as primary sources for EPA/DHA. In this case, the declining fish population would be an issue again. However, these sources are not always economic and convenient for human use.
26 Genetic engineering may enable EPA/DHA synthesis to be acquired by other microorganisms or plants in order to provide other primary sources for EPA/DHA production. Much research has been done in the past 15 years to produce AA, EPA and
DHA in transgenic oilseed crops like sunflower and linseed. Several elongase and desaturase genes (A4' A5' A6) involved in AA/EPA/DHA synthesis have been cloned from algae, fish and bacteria (Hastings et al, 2004; Tonon et al, 2004; Zhou et al, 2007). In
2004, the production of EPA by a transgenic plant was reported for the first time (Napier et al, 2004; Sayanova & Napier, 2004). However, the concentration of EPA present was less than 1% of total fatty acids. Moreover, the accumulation of EPA was reported in leaf tissue not in seed-oils. Marine bacteria, particularly Shewanella spp., are recognized for their high production of EPA and DHA (Bowman et al, 1997; Nichols et al, 1997;
Satomi et al, 2003). Bacterial production of 00-3 FAs is of interest for several reasons.
Firstly, the relative ease of genetic manipulation in bacteria makes them suitable candidates for studying enzyme regulation in order to increase or modify their co-3 FAs production capacity. Secondly, co-3 FA genes can be cloned from EPA/DHA producing bacteria and transferred into other microorganisms. Thirdly, genetically modified bacteria have the potential to be used in the food industry. Fourthly, EPA and DHA obtained from bacteria are free of heavy metals and fishy flavors.
27 1.10 Research objectives
In order to discover an alternative primary dietary source for EPA/DHA, the following studies were carried out:
Investigate Shewanella sp. MAC1, an EPA/DHA producing-marine bacterium, genotypically and phenotypically.
Clone the omega-3 fatty acids genes from Shewanella sp. MAC1 by construction of a genomic DNA library.
Detect recombinant production of EPA and/or DHA in transgenic E. coli.
Determine concentration of EPA/DHA produced by transgenic E. coli at different temperatures.
Sequence the EPA/DHA gene cluster and identify the exact genes responsible for
EPA/DHA production
28 2. Reclassification of Shewanella putrefaciens MAC1 and
detection of EPA/DHA production by Shewanell baltica
strains
2.1 Abstract
Shewanella baltica has been reported in the literature as non-omega-3 fatty acid- producing marine bacteria. Four Shewanella baltica strains, designated S. baltica MAC1,
S. baltica LMG 2263, S. baltica LMG 22253 and S. baltica LMG 2250 isolated from mackerel, cuttlefish, Baltic sea and oil brine, respectively, were tested for the production of omega-3 fatty acids, using a modified fatty acid extraction protocol. All S. baltica strains were able to produce eicosapentaenoic acid (EPA; 1.7-12.6% of the total fatty acids) at 4°C, 10°C, and 25°C. The production of docosapentaenoic acid (DPA), which is a known precursor for docosahexaenoic acid (DHA), was also detected in 5. baltica strains grown at 4°C. Analysis of the complete 16S rRNA gene sequences of these strains indicated that they are very close phylogenetically (sequence similarity >99%).
Furthermore, phenotypic characterization and biochemical analysis of these strains confirmed their close relationship and placed them within the Shewanella baltica species. Based on these data, we propose to reclassify the above four S. baltica strains as
EPA-producing Shewanella in the phylogenetic tree.
29 2.2 Introduction
In 1996, a bacterium was isolated from mackerel entrails at the University of
Guelph (Cadieux, 1996), which was initially identified as Shewanella putrefaciens
MAC1 based on its fatty acid composition and its biochemical profile using the biolog
GN MicroPlate (Cadieux et al, 1998). Shewanella putrefaciens MAC1 was characterized as a gram-negative, aerobic, single flagellated, rod-shaped marine bacterium. S. putrefaciens MAC1 was found to grow optimally at 25°C (Cadieux, 1996). In addition, it was reported that this bacterium could grow at temperatures ranging from 0°C to 30°C, but not at 37°C. This bacterium was of interest because it can synthesize EPA, DPA and
DHA (Amiri-Jami et al., 2006; Cadieux et al, 1998). Shewanella putrefaciens MAC1 is able to produce DPA at a concentration of 0.2% of total fatty acids (Cadieux et al, 1998).
It produces EPA in significant amounts ranging from 2.5% to 12.6% of total fatty acids and DHA from traces to 0.4% upon manipulation of the growth temperature and growth media (Amiri-Jami et al, 2006; Cadieux et al, 1998). Decreasing the growth temperature resulted in an increase in EPA/DHA production. Temperature is one of the most important environmental factors affecting the growth and fatty acid composition of this bacterium. Early reports highlighted that most Shewanella species capable of synthesizing EPA were found in cold deep-sea waters and sediments (Delong &
Yayanos, 1986; Wirsen et al, 1987). For instance, Shewanella benthica grew optimally at 4°C, S. hanedai at 15-25°C, S. gelidimarina at 15°C, S. frigidimarina at 10°C and other species at 25-35°C (Venkateswaran et al, 1999).
30 The major fatty acids produced by 5. putrefaciens MAC1 are EPA as the PUFA, and palmitoleic acid (16:1) as the monounsaturated fatty acid (Amiri-Jami et ai, 2006;
Cadieuxefa/., 1998).
To enhance EPA/DHA production by S. putrefaciens MAC1, the transposon TN5 was used to generate random mutations (Amiri-Jami et ai, 2006). Three mutants were identified as high EPA/DHA producers. They were able to produce 3-5 times more EPA and DHA compared to the wild type at 0, 4 and 10°C. Interestingly, one of the mutants produced 0.3 mg EPA g 4 (0.2% of total fatty acid) when grown at high temperature
(30°C). Preliminary analysis of the complete 16S rRNA gene sequence of Shewanella.
MAC1 showed high homology (sequence similarity >99%) to Shewanella baltica (Amiri
& Griffiths, unpublished data). Therefore, the name was changed from Shewanella putrefaciens MAC1 to Shewanella baltica MAC1. This coupled with the production of
EPA by S. sp. MAC1 in substantial amounts contradicts other reports that described S. baltica as a non- EPA-producing Shewanella (Bowman, 1999; Kato & Nogi, 2001; Owen et ai, 1978; Satomi et al, 2003). It was demonstrated (Owen et ai, 1978) that
Shewanella species was comprised of at least four clearly separated genomic groups numbered I to IV. They were named as Shewanella putrefaciens, Shewanella benthica,
Shewanella hanedai and Shewanella colwelliana. Shewanella putrefaciens was further divided into four sub-groups. Shewanella putrefaciens groups II and IV were renamed as
Shewanella baltica and Shewanella algae, respectively (Ziemke et ai, 1998) (Fig. 2.1).
31 -$. putrefaciens ATCC 8071(T) S. putrefaciens Mft-30 57i. putrefaciens S. putrefaciens ATCC8071(T) . pufmfacims A31 Owen's Grriup I —S. pulrefadens NCIMB B768 -5. putmfmiens NCW6 12S77 5/). putrefaciens J.Wrefactem ATCC8073 jSc , putrefaciens-5.0 GC-29 Owen's Group III r» ruermfxelKmt 5P-7 ' S, fwtnefaciens SP-JZ
Sh. baliica - 137 •5. toteM ATCC 10735(T, -5. Oa/tfca OS195 5. trigiaiimrimi NTBSLe 5. fvgktirtigrtizf NTBSLb
.( X, triQidiimrma 930rS *-£ friakSmtrim NTPt SLJ —5. m&dimmm MfPiS-SU 5. ftiCjtttimatinz 5 a s. immmim AI 69 5. frmaift&ftra ACAM 593 S, flkfiitfltmtmil ACAW 531{Tj & ftfgidimswtgi ^eSIOAJ "5'. fraidinmrms ACAM .388 " MgKftmtifrns AC^MSOO *—S. tmtfimmm SC2* S. fimdimaww NQM8 400 S. a^ae BrY•— — -c ftigMimartrti) ACAM 1 ?? S. ingsciwmms Sc/.A 1 L S. a/gae ATCC 5119E
Figure 2.1. Schematic representation of the 16S rRNA phylogenetic tree for the genus
Shewanella. Lines with green color show the species known to produce EPA/DHA. Black
Black lines show the species known not to produce EPA/DHA. For lines with blue color no data are available (Bowman, 1999).
32 Among these, it was reported that Shewanella baltica and Shewanella algae did not produce EPA (Russell & Nichols, 1999). Recently, Shewanella spp. were divided into two groups; members of group 1 were characterized as high pressure cold-adapted species which produce a great amount of EPA, while Shewanella group 2 are mesophilic and cannot produce EPA (Kato & Nogi, 2001; Satomi et al., 2003). Shewanella baltica is placed in Group 2. No EPA production has been reported by 5. baltica so far and in the phylogenetic tree for the genus Shewanella S. baltica is placed in the non-EPA producing branch (Skerratt et al, 2002).
In this study, Shewanella MAC1 was reclassified on the basis of 16S rRNA sequence data. We further analyzed strain MAC 1 and three strains of S. baltica obtained from a European culture collection both genotypically and phenotypically; including their ability to produce EPA and/or DHA. Finally their total fatty acid profiles were compared at different temperatures.
2.3 Materials and Methods
2.3.1 Bacterial strains
To identify the production of EPA by Shewanella baltica, four strains of this species were tested i.e., 5. baltica LMG 2250, S. baltica LMG 22253 (OS 155), S. baltica
LMG 2263, and S. baltica MAC1. The first three strains were obtained from Culture
Collection of the Laboratorium voor Microbiologic Universiteit Gent, whereas S. baltica
MAC1 was procured from Canadian Research Institute for Food Safety (CRIFS). S.
33 putrefaciens LMG 2369, which was used as a negative control, was also obtained from
LMG. The strains analyzed had different origins. S. baltica 2250 was isolated from oil brine in Japan, S. baltica MAC1 and S. baltica 2263 were isolated from fish intestine, while 5. baltica 22253 was isolated from seawater of Central Baltic Sea.
2.3.2 Preparation of Shewanella baltica cultures
Stock cultures of S. baltica MAC1 were maintained frozen at -80°C in 15% glycerol. Bacterial cultures for use in experiments were prepared by inoculation of frozen stock cultures onto Marine Agar (MA) 2216 (Difco, Detroit, MI, U.S.A). S. baltica LMG
2250, S. baltica LMG 22253 (OS 155), 5. baltica LMG 2263 and S. putrefaciens LMG
2369 were obtained as freeze-dried cultures in vacuum-sealed ampoules. To prepare bacterial culture for the experiments, 0.5ml Marine Broth (MB) 2216 (Difco, Detroit, MI,
U.S.A) were added to the dried bacteria. The contents was gently mixed and transferred to the MA plates. Plates with S. baltica MAC1 and those with S. baltica LMG 2250, S. baltica LMG 22253 (OS 155), S. baltica LMG 2263, S. putrefaciens LMG were incubated at 30°C overnight. Thereafter, all the plates were stored at 4°C.
2.3.3 Growth conditions and biochemical characteristics
Temperature requirements were determined for all the strains by growing the strains on MA at different temperatures. Inoculated plates were incubated for up to 2 weeks at temperatures of 4°C, 10°C, 25°C, 30°C and 37°C. Growth was considered positive if the strains were able to form visible colonies on the solid medium.
34 Biochemical tests were performed with API 20 NE and API 20 E kits (BioMerieux,
Lyon, France), which were prepared according to the manufacturer's instructions. The
API strips were incubated for 48 hours at 25°C.
2.3.4 DNA extraction from Shewanella baltica strains
To extract the genomic DNA of S. baltica LMG 2250, LMG 22253, LMG 2263,
S. baltica MAC1 and 5. putrefaciens LMG 2369, an isolated colony of each purified
Shewanella on the MA was inoculated in 50 ml tubes containing 5 ml MB then incubated at 30°C overnight while shaking at 200 rpm/min. One milliliter of each sample (overnight culture) was added to an autoclaved 1.5 ml microcentrifuge tube. The tubes were centrifuged at 15,000 g for 3 minutes in a Microfuge (Beckman Coulter Mississauga,
Ontario) to pellet the cells. After removing the supernatant, the bacterial pellet was used for genomic DNA extraction.
The extraction of genomic DNA was conducted with the GenomicPrep Cells and
Tissue DNA Isolation Kit (Amersham Pharmacia Biotech, NJ, U.S.A). DNA concentration was measured by absorbance at 260nm. A total amount of 60 |ig DNA was obtained. Isolated genomic DNA from S. baltica strains and S. putrefaciens LMG 2369 were stored at 4°C.
35 2.3.5 Polymerase Chain Reaction (PCR) procedure for 16S rRNA sequence
determination
Two oligonucleotide primers, described in Table 2.1, synthesized by Laboratory
Services Division, University of Guelph (Guelph, Ontario, Canada) were used to amplify the 16S rRNA gene from the genomic DNA of S. baltica. Forty five microliters of PCR
SuperMix (Invitrogen Life technologies, Burlington, Ontario) containing 22 mM Tris-
HC1 (pH 8.4), 55 mM KC1, 1.65 mM MgCl2, 220 uM dNTP and 22 U recombinant Taq
DNA polymerase/ml, were added to a sterile 250 |il microcentrifuge PCR tube. One microliter of 50 pmol/|il forward primer and l|il of 50 pmol/|jl reverse primer were added to 45 fil SuperMix and then 0.5 \ig DNA were added to the mixture of SuperMix and the primers. The final solution was gently mixed and PCR was performed in a Mastercycler
(Eppendorf Scientific, N.Y, U.S.A) under the conditions shown in Table 2.2.
2.3.6 Separation of DNA fragments
The standard method used to separate DNA fragments is electrophoresis through agarose gels. To perform agarose gel electrophoresis, a slab of 1% agarose gel was prepared by melting 0.5 g ultra pure agarose powder (Life Technologies, N.Y., U.S.A) in
50 ml of 1 x TAE buffer. The mixture was heated to boiling until a clear solution was observed. Then the solution was cooled to 40-45 °C and, after adding 0.5 Jul 10 mg/ml
Ethidium Bromide (ETB) (Sigma, MO, U.S.A), it was poured in a 10 cm tray and solidified at room temperature (23°C). The gel was placed in a gel electrophoresis submarine cell containing 250 ml 1 x TAE buffer.
36 Table 2.1
Primers used to isolate the 16S rRNA from S. baltica LMG 2250, LMG 22253, LMG
2263, S. baltica MAC1 and S. putrefaciens LMG 2369
Primer Sequence Target sequence
Forward primer 5' - AGTTTGATCCTGGCTC AG-3' 16S rRNA gene
Reverse primer 5'-GGTTACCTTGTTACGACTT- 16S rRNA gene
3'
Table 2.2
The conditions that were used in PCR
Temperature Duration Number of Cycles
94°C 4 min 1
94°C 30 sec 30
50°C 30 sec 30
72°C 2 min 30
72°C 10 min 1
Min: minutes; Sec: second; °C: centigrade
37 2.3.7 Purification of amplified DNA fragment from PCR reactions
QIAquick PCR purification kit (Qiagen, Mississaga, On, Canada) was used to purify the amplified product following the manufacturer's protocol. At the end, the DNA retained in the resin column was eluted with 30 [i\ elution buffer (EB). The DNA concentration of each sample was determined by absorbance at 260nm- The total amount of extracted DNA was 3-7 (ig. Purified PCR products were sequenced at the Guelph
Molecular Supercentre, Guelph, ON, Canada.
2.3.8 Growth conditions for fatty acid extraction
S. baltica strains and S. putrefaciens LMG 2369 were streaked on fresh MA plates and incubated at 30°C overnight. A single colony of each strain was grown in triplicate in a 50 ml tube containing 5 ml of MB and then incubated overnight at 30°C while shaking at 200 rpm/min. Autoclaved 250ml Erlenmeyer flasks containing 150 ml MB were inoculated with 5 ml of each inoculum. The cells were incubated at 4°C, 10°C, 25°C,
30°C for 7, 5, 2 and 1 day, respectively, on a rotary shaker (200-rpm) until an absorbance at 600 nm of 2 was obtained for each.
2.3.9 Extraction of fatty acids
The cultures were harvested by centrifugation using a Beckman J2-MC centrifuge at 9,000 g and 4°C for 15 minutes. The harvested cells were washed with 200 ml ultra pure water, and centrifuged under the same conditions. This was repeated twice to ensure that all salts and media components were removed from the cells. The cell pellets
38 obtained from different cultures were lyophilized overnight in a Stoke Freeze-Drier
(Equipment Division Pennsalt Chemicals Crop, Philadelphia, PA, U.S.A) and ground to a fine powder form in a clean sterilized mortar with a pestle. The powders from each sample were weighed and were transferred to 20 ml screw cap test tubes. Five milliliters of 14% borontriflouride in methanol (Sigma, Oakville, ON, Canada) were added to the test tubes and then the cells were esterified in a boiling water bath for 15 minutes.
Samples were cooled to room temperature (23°C) followed by the addition of 5 ml ultra pure water saturated with NaCl. Two milliliter of hexane were added and the mixture was vortexed for 10 seconds. The phases were separated using a Beckman J2-MC centrifuge at 600 g and 4°C for 2 minutes. The upper hexane layer was transferred to a clean screw cap tube by using a 1 ml Pasteur pipette. The reaction mixture was re-extracted 4 times with 2 ml of hexane and each time the upper layer was transferred to the same screw cap tube. The collected hexane in each screw cap tube was evaporated under nitrogen (4.8 grade) at room temperature. The fat at the bottom of each tube was dissolved in 100 \il of iso-octane (Fisher Scientific, Napean, ON). In the final step, 5 mg of sodium sulfate was added to the fatty acid samples to remove any water present.
2.3.10 Gas Chromatographic Analysis
The fatty acid methyl esters were analyzed using a Shimadzu Gas Chromatograph
GC-14A. Each fatty acid sample (0.3 [A1) was injected into the GC and integrated using a
Shimadzu C-R4A integrator {Shimadzu, INC, Kyoto, Japan). The samples were injected using a clean l^il Hamilton syringe (Hamilton Company, Reno, NV, U.S.A). A fused
39 silica BPX 70 capillary column (60 m x 0.22 mm ID) was used to separate the fatty acids under the specific conditions shown in Table 2.3. The flow rates of the carrier gas, air, and hydrogen for the flame were 1 ml/min, 400 ml/min and 30 ml/min, respectively. The range of the flame-ionization detector was set at 102. The fatty acid methyl esters were identified by comparison of their retention times with those of known standards, including 37 fatty acid methyl esters from C4:0 to C22: 6n3 (Sigma, Oakville, On,
Canada), pure EPA, DPA, DHA, oleic (OL), LA and ALA, standards (Sigma, Oakville,
On, Canada). Internal standards for EPA and DHA peaks were also used. In addition, before injecting the first sample, the initial temperature of the GC was increased to 280°C for 15 minutes in order to remove any remaining fatty acid residue and then the GC column was cleaned by injecting 1 \i\ iso-octane and then running the GC for 30 minutes under the same conditions shown in Table 2.3. The syringe was washed 50 times in clean solvent, either iso-octane or hexane, between each injection.
2.3.11 Gas Chromatographic-Mass Spectrometry Analysis
EPA/DHA production was detected by Gas Chromatographic-Mass Spectrometry
(GC-MS) in the Chemistry Department, University of Guleph, Guelph, ON. Individual
EPA and DHA peaks were confirmed with a Varian Saturn 2000 ion trap mass spectrometer in external EI mode (Varian Canada, Mississauga, ON, Canada). EPA and
DHA were identified with target ion mass to charge ratios (m/z) of 287 +15= 302 (EPA molecular weight) and 313 + 15= 328 (DHA molecular weight). The fatty acid methyl esters in iso-octane were injected onto a 30 m x 0.25 mm capillary column. The oven was programmed to increase from 125 °C (3 min wait) to 250 °C for 4 min"1. Injection
40 Table 2.3
Gas Chromatography Parameters for fatty Acid Separation of Shewanella baltica strains
Program Duration/Temperature
Detector temperature 280°C
Injector temperature 260°C
Initial temperature 110°C
Initial time 2min
Program rate 4°C/min
Final temperature 230°C
Final time lOmin
41 volume and temperature were 1 [xl and 250°C, respectively. Compounds were identified by comparison of relative retention time and mass spectra with that of pure standards.
2.3.12 Statistical Analysis
To find the significant differences in EPA production by different S. baltica strains at different temperatures a two factor ANOVA was used. The between 5. baltica strains and the between temperatures variations were considered as analytical factors.
Groups with significantly different means were identified using Tukey's test (p < 0.05).
2.4 Results and discussion
2.4.1 Physiological properties and phenotypic characteristics of 5. baltica strains
All four Shewanella strains produced circular, light to dark salmon pink- pigmented colonies of 2 - 4mm diameter on plates of MA at temperatures of 4°C, 10°C,
25°C and 30°C, while no growth occurred at 37°C. Colonies formed following 7 days, 5 days, 2 days and 24 hours incubation at 4°C, 10°C, 25°C and 30°C, respectively. S. putrefaciens LMG 2369, used as a negative control, grew at 37°C but not at 4°C. All the strains were positive for nitrate reduction. Biochemical properties are summarized in
Table 2.4.
2.4.2 16S rRNA sequence analysis
Figure 2.2 shows the PCR results on 1% agarose gel when forward and reverse primers were used for S. baltica strains, MAC1 and S. putrifaciens. One strong fragment with a size of -1500 bp was obtained for all S. baltica strains, MAC1 and 5. putrifaciens.
42 Table 2.4. Selected phenotypic characteristics of the 5. baltica strains Strains: 1, S.baltica MAC1; 2, S.baltica LMG 2250; 3, S.baltica LMG 2263; 4, S.baltica LMG 22253. +, Test is positive; -, test is negative.
Characteristic S. baltica S. baltica S. baltica S. baltica MAC1 LMG2250 LMG2263 LMG22253 Growth at: 4°C + + + + 30°C + + + + 37°C - - - - Production of: Lipase + + + + Amylase - - - - Gelatinase + + + + P-glucosidase + + + + Urea - - - - H2S production + + + + Utilization of: D-Glucose + + + + D-Gluconate + + + + L-Arabinose - - - - Maltose + + + + Sucrose + + + + Citrate + + + + Caprate - - - - Glucosamine + + + +
43 M_ 1 2 3 4 5 6
Figure 2.2 Gel electrophoresis (1%) of PCR products. M: X Eco 471 DNA marker (New England Biolabs). Lanes 1, S. putrefacins; lane 2, S. baltica LMG 2250; lane 3, S. baltica LMG 22253; lane 4, 5. fca/rica LMG 2263, lanes 5 & 6, 5. fca/f/ca MAC 1.
44 The almost-complete 16S rRNA gene sequence (-1500 bp) for S. baltica LMG 2250, S. baltica LMG 22253, S. baltica LMG 2263, and S. sp. MAC1 were obtained and compared to the GenBank nucleotide database using an online BLAST search. S. sp.
MAC1 showed 99% homology to S. baltica OS155 (LMG 22253) and S. baltica
SBAJ216. This suggests that S. MAC1 can be considered as a member of S. baltica. S. baltica LMG 2250, S. baltica LMG 22253 and S. baltica LMG 2263 showed 99.7-100% homology to S. baltica NCTC 10735, S. baltica OS 155 and 5. baltica NCTC 10737.
Since the 16S rRNA sequence confirmed that all four strains were S. baltica, we examined them for co-3 FA production.
2.4.3 Fatty acid composition of Shewanella baltica strains at different
temperatures
The fatty acid profiles of all four S. baltica strains at four different temperatures are shown in Table 2.5. The fatty acids were within the range of C12 - C22, commonly found in many marine bacteria (Hirota et ai, 2005; Satomi et al, 2003). They were dominated by palmitoleic acid (CI6:1) and cis-10-heptadecenoic acid (C17:l) as
MUSFA. Palmitoleic acid was the most abundant monounsaturated fatty acid found in all four S. baltica strains. The presence of this monounsaturated fatty acid is also reported to be present in large amounts in almost all Shewanella species (Gentile et al., 2003; Kato &
Nogi, 2001; Venkateswaran et al., 1999). Polyunsaturated fatty acids found to be present were LA, ALA, DPA and EPA.
45 Table 2.5 Fatty acid compositions (shown as percentage of total detected fatty acids) of
S. baltica strains at different temperatures. SFA, saturated fatty acid; MUFA, monounsaturated fatty acid; PUFA, polyunsaturated fatty acid.
Fatty acid composition (%) Fatty acid 4°C 10°C 25°C 30°C 12:0 1.3 1.5 2.2 3.1 13:0 0.9 1.6 1.8 2.7 14:0 1.3 1.3 1.3 2 15:0 3.2 6 9.1 11.8 16:0 2 4.2 4.5 3.7 17:0 5 6.1 6.8 6 18:0 2.4 4.9 4.4 4.8
Sum SFA 16.1 25.5 30.1 34 15:1 1.5 1.3 1.9 1.5 16:1 37 32.1 30.9 32 17:1 8.6 5.6 13.8 16.5 18:ln9c 3.2 5.2 2.2 2.3 C22:ln9 0.5 0.2 0.11 0.01
Sum 51 44.4 48.9 52.3 MUFA C18:2n6c 8.6 6.6 0.4 0.06 C18:3n3 1.15 0.8 0.3 0.05 C20:5n3 7.8 2.5 0.95 0.06 C22:5n3a 0.05 - -
Sum 17.6 10 1.7 0.2 PUFA Others" 15.3 20.1 19.3 13.5
a only detected in S.baltica MAC1 "includes: 14:1; 15:ln8c; 16:ln9c; 17:ln8c; 18:ln9t; 18:2n6t; 18:3n6, 20:0; 20:ln9, and some traces which were less than 1% (traces of DHA was only found for S. baltica MAC1). This was calculated from the area under each fatty acid peak, which were added together to give a total peak area. The area of each fatty acid peak was divided by the total, and multiplied by 100 to give percent of total fatty acid for each individual fatty acid.
46 Table 2.5 shows the percentage of each detected fatty acid for all four strains of S. baltica at different temperatures. Changes in the growth temperature clearly resulted in quantitative and qualitative changes in the lipid composition of the membrane. The presence of docosapentaenoic acid (DPA, 22:5n3) at 4°C and also higher levels of linoleic acid, linolenic acid and EPA at lower temperatures demonstrate the effect of temperature on the PUFA composition of the bacterial cell membrane. It has been well documented that most bacteria change their membrane lipid composition as a response to an alteration in environmental factors such as temperature, pressure and salinity
(Valentine & Valentine, 2004). Henderson et al. (Henderson et al, 1993) reported that temperature had a more pronounced effect on the lipid composition of Vibrio sp. than did other growth conditions such as salinity. The maintenance of membrane fluidity is one of the critical characteristics for bacterial survival when they live at low temperature or high pressure. Bacterial cells modulate the composition of their lipid membrane when the temperature increases or decreases as a response to regulate membrane fluidity (Russell
& Nichols, 1999). The ability of bacteria to adapt their membrane lipid composition in response to changes in growth temperature usually involves changes such as acyl chain length, methyl branching and, particularly a change in fatty acyl unsaturation (Foot et al,
1983). Moreover, the changes in unsaturation are mediated by desaturase enzymes or, in some bacteria, by an alteration in the anaerobic pathway of fatty acid biosynthesis. It is believed that the maintenance of membrane lipid bilayer fluidity is necessary to ensure the proper functioning of membrane transport, intracellular signaling, gene regulation, and protein-protein interaction within the bilayer. Moreover, changes in the lipid composition influence the physical properties of a membrane such as temperature-
47 phase behavior (gel or liquid-crystalline) and the viscosity of the lipid bilayer.
At higher temperatures (30°C), S.baltica strains produced about 2 times more short chain saturated fatty acid compared to those grown at 4°C. It is well documented that psychrophilic bacteria respond to changes in growth temperature by altering not the unsaturation but the chain length of its fatty acids (Allen et ai, 1999). It is also reported that other marine bacteria such as Vibrio marinus adapt to higher growth temperatures by increasing the number and the amounts of saturated fatty acids. In addition, smaller amounts of long chain-saturated fatty acids such as C16:0 and C18:0 were detected at
4°C compared to 30°C. The amount of C16:0 and C18:0 increased when the cells were grown at higher temperatures. It was also reported that the amount of C16 was higher when Vibrio sp. was grown at higher temperature (Hamamoto et ai, 1994).
2.4.4 EPA production by Shewanella baltica strains at different temperatures
Previous research indicated that Shewanella baltica did not produce EPA.
However, all S. baltica strains used in this study were found to be positive for EPA production at temperatures of 4°C, 10°C, and 25°C. Among strains tested S. baltica
MAC1 was able to produce EPA, DPA and traces of DHA. Pure EPA standard (applied alone as an internal standard) was used to confirm the putative EPA peak obtained in gas chromatograms. The presence of EPA was also confirmed by comparison of electron impact mass spectra of the samples with that of the known standard. Figure 2.3 illustrates the presence of EPA in 5. baltica 2263 at 4°C by GC. The GC-MS result confirms the production of EPA by S. baltica 2263 at 4°C (Fig. 2.4).
48 < t.Cv <~- c-J *-£> CD h ! r- - OO 06 irc : :! ir L f r— f *Jl> T ti I CO c«-\4 i CI r-J 1 ir% LL
Figure 2.3 Gas Chromatograph of total fatty acid methyl esters prepared from S.
baltica 2263. Fatty acids were extracted from cells grown for 7 days at
4°C and then analyzed using a Shimadzu 14A Gas Chromatograph.
49 ion: ZB7all epa.SId.3nB
(ore 267 bit «&ijl,tmi»
J '' ' ' ' d.fes ' afeb " b.k> ' ahs:'
5tt>3i mn. scan: SWCJ tjnah: \ ton: 1«J us RtC; 21O&309 BC
Sffifc-
oar m 11 Jlli AHAlJLuiL &G3GnitX Son Sa2ChWV 1 ttxv. 461 us RtC: Ue3$0^EC
10G3&3
1 ll U I :a>*
4 The mass spectra for EPA standard and EPA in 5. baltica 2263. A)
Chromatograph of EPA standard analyzed by GC-MS; B) Chromatograph
of EPA in sample at retention time of 9.62; C) Mass spectra of EPA
standard; D) Mass spectra of EPA in sample.
50 Similar results were obtained for S. baltica 22253, S. baltica 2250 and S. baltica MAC1.
As oc-linolenic acid is a precursor for EPA, DPA, and DHA, the presence of ALA at different temperatures was studied (Table 2.5). S. baltica strains were able to produce this fatty acid at 4°C, 10°C, 25°C and 30°C but in different amounts. A possible explanation of this phenomenon is that the activities of A15 and A6 desaturases (both of which desaturate linoleic acid) were affected differently by the growth temperature. At lower growth temperatures, the amounts of linoleic acid metabolized by A15 desaturase might be increased to form more ALA. This change might lead to an increased level of EPA. In the case of S. baltica strains, the amount of EPA was significantly lowest at 30°C (p <
0.05 ) and increased gradually at 4°C (Fig. 2.5). This suggests that S.baltica strains adapted to the colder temperature (4°C) by synthesizing more EPA. It is also well documented that the degree of unsaturation affects the melting temperature of fatty acids, which is directly related to membrane fluidity (Rotert et al., 1993). Furthermore, the proportion of polyunsaturated fatty acids, particularly EPA, in membrane lipids indicates that EPA is likely to be part of a homeoviscous adaptive response to regulate membrane fluidity. The cell membranes of bacteria containing more PUFA, especially EPA and/or
DHA, are more permeable at cold temperatures (Gentile et al., 2003; Valentine &
Valentine 2004,). This characteristic is common to the Shewanella spp. found in cold, deep sea water and Polar Regions which provides them with the ability to grow well at the low temperatures encountered in these environments (Russell & Nichols 1999; Kato
& Nogi 2001). Most Shewanella strains are psychrotolerant and a large number of them isolated from cold environments have the capacity to produce EPA and DHA. Production
51 MAC1 2250 2263 22253 bacterial strain
Figure 2.5 EPA concentration in 0.2 g cell dry weight of S. baltica MAC1, LMG
2250, LMG 2263 and LMG 22253 cultured at 4 (•), 10 (H ), 25 (jl ) and
30°C (D) in 3 independent experiments. EPA concentration is expressed
in mg and was calculated from the area under the GC chromatogram peak
corresponding to EPA. Results show mean ± SD from 3 independent
experiments.
52 of EPA and DPA by S. baltica strains increases in the cell membrane as a function of decreased temperature, indicating that the presence of these components may be important for growth at low temperatures. The same phenomenon has been recently reported (Wang et al, 2008) in a psychrotolerant deep-sea bacterium, Shewanella piezotolerans WP3. They reported that Shewanella piezotolerans WP3 possess genes or gene clusters, which help it to adapt to life at low temperatures by enhancing its capacity for energy production, protein function and nutrient acquisition. It is unknown how environmental changes such as temperature and pressure modulate synthesis of EPA and
DHA in bacteria. Despite the hypothesis that marine bacteria produce omega-3 fatty acids at cold temperature to maintain their membrane fluidity, it is still unclear why these bacteria should produce these specific fatty acids since there are some cold, deep sea bacteria such as Photobacterium profundum that do not require either EPA or DHA to grow at low temperature (Allen et ai, 1999).
A possible explanation for our ability to detect EPA in Shewanella baltica might be the modification in the method we employed to extract the fatty acids. The method of lipid extraction by Cadieux et al. (1998) was modified by incorporating a NaCl saturated solution, together with re-extraction of the fatty acid methyl ester from the reaction mixture with 2ml hexane (which was repeated 4 times) and separation of the phases by a brief centrifugation.
In most of the articles describing lipid extraction from membranes of Shewanella or other marine bacteria (Gentile et ai, 2003; Morita et al, 2005; Nichols et al, 1997;
53 Yano et al, 1994; Yazawa et al, 1988; Yazawa, 1996), the methods of Folch or Bligh and Dyer were used for total lipid extraction (Bligh & Dyer, 1959; Folch et al, 1957).
These methods are rapid and effective methods for detecting total lipid in fish muscle and other biological tissues. The sample is first homogenized with chloroform/methanol/water, filtered under suction, the phases are allowed to separate and then the lower phase, which contains chloroform, is transferred to another tube and evaporated under nitrogen. The only difference between the two methods is the chloroform/methanol/water ratio and solvent/sample ratio. The advantage of the Bligh and Dyer protocol is a reduction in solvent/sample ratio. The Bligh and Dyer method is one of the most recommended methods for total lipid extraction from biological tissues.
In several publications related to the total lipid extraction from Shewanella, many modifications to Bligh and Dyer method have been reported to improve the efficiency of lipid recovery from cell membranes (Gentile et al, 2003; Nichols et al, 1997; Yano et al, 1994). However, these researchers did not describe what modifications have been implemented. Recently, another method of extraction was published by Orikasa et al.,
(2006). Since then some researchers have used Orikasa's method to extract total lipid from bacterial cells (Orikasa et al, 2006b; Sugihara et al, 2008). In this method, the cells are directly methanolyzed using 2 M HCL in methanol at 80°C for 60 min, then the fatty acid methyl ester is extracted with hexane and subjected to GC after concentration. This method is similar to Takeyama's method with minor modifications (Takeyama et al,
1997). There are some obvious differences between the above methods and our method of extraction. Firstly, only 20 - 40 mg wet weight or lyophilized cells were used for lipid extraction, while we used 200 mg dry cells. Secondly, in all the methods lyophilized cells
54 or wet cells were used directly for esterification while we disrupted the cell membranes by grinding them to a fine powder. This may enhance the contact of the solvents with the lipids in the samples. Thirdly, a smaller volume of solvent was used previously for estification compared to our method. Using a larger volume of hexane increases the concentration of the extracted fatty acids. Finally, in all the previous extraction methods after esterification either pure water or weak salt solution 0.8% was added to the samples and then lipid was extracted with solvent, while we used water saturated with NaCl instead of pure water. In principle a solvent mixture must be polar enough to remove the lipid from cell debris and since NaCl increases the polarity of the solution, it could be the key factor leading to a better separation of total fatty acids from the cell membrane. All the above modifications in our method might contribute to our ability to detect EPA in
Shewanella baltica strains.
55 2.5 Conclusions
Our results provide evidence of the presence of EPA in Shewanella baltica strains. Currently, S. baltica is considered to be a member of Shewanella group 2 in the phylogenetic tree. This group is described as mesophilic marine bacteria that lack the ability to produce EPA (Kato & Nogi, 2001; Satomi et al, 2003). Our data confirm that
S. baltica can grow very well at 4°C but not at 37°C; a phenotypic property that distinguishes them from related species (Ziemke et al, 1998). In addition, use of fatty acid composition is an effective tool in the identification of new microorganisms and in their taxonomic classification. We present evidence to support the reclassification of
Shewanella baltica in the phylogenetic tree. The ability to detect EPA in S. baltica may be due to an improved method for fatty acid extraction from freeze-dried bacteria.
56 3. Cloning omega-3 fatty acid genes from Shewanella baltica
MAC1 by constructing a Bacterial Artificial Chromosome
library
3.1 Abstract
Omega-3 fatty acids, particularly eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), are well documented for their beneficial effects on human health. Shewanella baltica MAC1 synthesizes EPA and DHA. To identify EPA/DHA genes in S. baltica MAC1 and to obtain recombinant EPA/DHA, a large DNA fragment library was constructed. The 200 kb genomic DNA fragment of S. baltica MAC1 was cloned into a bacterial artificial chromosome (BAC) vector. This resulted in 490 clones with average size of 100-200 kb. One hundred and twenty BAC clones were chosen randomly from the library and tested for EPA/DHA production. Recombinant production of EPA (in a trace amount; < 0.02 % of total fatty acid) and DHA (0.85% of total fatty acid) was detected by GC and GC-MS in one clone carrying the BAC vector with the insert size of 200 kb. Attempts to subclone the EPA/DHA gene cluster into a food grade, broad host range vector were not successful due to an inability to digest and to isolate the insert from the BAC vector. Based on this study, constructing a BAC library with large
DNA fragments increases the chance of chimerism that prevents isolation of the insert.
57 3.2 Introduction
Omega-3 fatty acids are essential fats required for human health. Currently, the main dietary source is cold-water fish. Due to a substantial decline in the fish population, alternative dietary sources for omega-3 fatty acids have been sought. The production of omega-3 fatty acids, particularly EPA and DHA, by bacteria from cold deep-sea water and sediments was reported first by DeLong and Yayanos (1986). Their discovery resulted in extensive research into EPA/DHA producing bacteria. The production of
EPA/DHA by Shewanella strains is well documented (Gentile et al, 2003; Hirota et al,
2005; Nichols et al, 1997; Sato et al, 2008; Yazawa et al, 1988). Yazawa (1996) reported the first evidence for a 38 kb gene cluster containing 9 ORFs involved in the biosynthesis of EPA in the marine bacterium, Shewanella pneumatophori SCRC-2738.
So far, this is the only report regarding recombinant production of EPA. In a follow up study using the same plasmid (pEPA) generated by Yazawa, Orikasa et al (2004) reported that among the 9 ORFs in the sequence, only five (ORFs 2 and 5-8) were essential for EPA biosynthesis. The same group later added the phosphopantetheinyl transferase (PPTase) gene to the original plasmid and claimed the production of recombinant DHA. Many attempts have been made by other independent researchers to clone the EPA/DHA genes in order to produce recombinant EPA/DHA. However, they were unable to clone all genes responsible for EPA/DHA production, nor did they obtain any recombinant EPA/DHA. To isolate the EPA/DHA gene cluster, researchers generated genomic DNA libraries in which 35-50 kb fragments were insert into a cosmid vector. In addition, they designed primers using the available published sequence for
58 EPA/DHA synthesis genes. To date, their attempts to isolate all the EPA/DHA biosynthesis genes have been unsuccessful.
To identify all the ORFs responsible for EPA and/or DHA production, the construction of a genomic DNA library with larger DNA fragments was helpful.
Bacterial Artificial Chromosome (BAC) cloning vectors are extensively used for preparation of genomic DNA libraries with large inserts. There are reports outlining the construction of BAC libraries from fungi (Diaz-Perez et al, 1996), plants (Mozo et al,
1998; Song et al, 2000), marine macroalgae (Deng et al, 2004), mammals (Schibler et al., 1998; Sparwasser & Eberl, 2007; Walter et al., 2005), and human DNA (Asakawa et al., 1997). Large DNA fragments are highly stable in BAC clones. In addition, the BAC cloning system provides the user with total control over the clone copy number. BACs are synthetic vectors and they are constructed based on an E. coli single-copy fertility plasmid or F- plasmid. The F plasmid is relatively large and vectors derived from that have a higher capacity compared to the other plasmid vectors. The F-factor plays an important role because it contains partition genes (parA and parB), which are part of a partitioning (par) system that ensures at least one copy of the plasmid segregates into each daughter cell during bacterial cell division. The insert size for BAC ranges from
100-300 kb; 130 kb is the average. The sizes of the BAC vectors are 7-10 kb. Several derivatives of BAC vectors are commercially available. Figure 3.1 is a diagram of a BAC vector which is a derivative of pBeloBAC (7.4 kb), one of the most widely used BAC vectors (Song et al., 2000; Wild et al., 2002).
59 BamH i 353 P'B /Hind lit 383 HpaI 6996 Pel 16849 __ __ Sea I 793 >4pa I 6339 Bsa 16177
parB plndigoBAC-5 redf ^— SsfZ17 11832
7506 bp ori2 X/?ol2380
^^ __ '£0^12836 estX I 4452 Afe I 3933
Figure 3.1. Diagram of plndigoBAC-5, a derivative of pBeloBAC.
60 It is well documented that DH10B and EPAI300 are suitable hosts for BAC vectors (Deng et al, 2004; Song et al, 2000; Walter et al, 2005). These E. coli strains have been specifically engineered to carry BAC vectors. They can take up large DNA fragments, block restriction of DNA containing methylated DNA and also block restriction of foreign DNA by endogenous restriction endonucleases. The EPI300 E. coli contains the trfA gene, whose protein product is required for initiation of replication.
In this chapter, construction of a genomic DNA library with large DNA fragments prepared from S. baltica MAC1 using a BAC vector is described. This BAC library was screened using GC to detect clones that produce EPA and/or DHA. Attempts were made to isolate the insert containing the EPA/DHA synthesis genes from the EPA/DHA producing clone in order to digest it to smaller sizes and subclone it into a food grade vector. Finally, the production of EPA/DHA by positive clones was tested in skim milk.
3.3 Material and Methods
3.3.1 Bacterial strains and growth conditions
Stock cultures of S. baltica MAC1 were maintained frozen at -80°C in 15% glycerol. Bacterial cultures for use in experiments were prepared by inoculation of frozen stock cultures onto Marine Agar (MA) 2216 (Difco, Detroit, MI, U.S.A). Plates with S. baltica MAC1 were incubated at 30°C overnight. To extract the chromosomal DNA of S. baltica MAC1, an isolated colony from an overnight culture was inoculated in 50 ml tubes containing 5 ml Marine Broth (MB) 2216 (Difco) and incubated at 30°C overnight
61 while shaking at 70 rpm/min on a New Brunswick CI platform shaker (New Brunswick
Scientific, Edison, NJ, U.S.A).
3.3.2 Genomic DNA isolation
3.3.2.1 DNA extraction using commercial kits
One milliliter of each sample (overnight culture) was added to an autoclaved 1.5 ml microcentrifuge tube. The tubes were centrifuged at 15,000 g for 3 minutes in a
Microfuge (Beckman Coulter, Mississauga, ON) to pellet the cells. After removing the supernatant, the bacterial pellet was used for genomic DNA extraction.
The extraction of genomic DNA was conducted with the GenomicPrep Cells and
Tissue DNA Isolation Kit (Amersham Pharmacia Biotech, Piscataway, NJ, U.S.A). The
UltraClean Microbial DNA isolation kit (Mo Bio Laboratories, San Diego, CA, U.S.A) was also used to isolate DNA. The DNA concentration was measured by absorbance at
260nm and 280nm. A total amount of 25-30 |ig DNA was obtained using the GenomicPrep
Cells and Tissue DNA Isolation (GPCT) kit and UltraClean Microbial DNA Isolation
(UCM) kit. In addition, the ratio of A260/A280 was calculated to determine the DNA purity. Isolated genomic DNA from S. baltica MAC 1 was stored at 4°C. The DNA was isolated several times with a minor modification of the extraction method. This included resuspending DNA in 50 ul instead of 100 [il elution buffer, until the DNA of the desired size and purity was obtained.
62 3.3.2.2 Extraction of DNA in plug agarose
One milliliter of overnight culture was transferred to an autoclaved 1.5 ml microcentrifuge tube (Fisher Scientific, Ottawa, ON, Canada). The tubes were centrifuged at 14,000 g for 3 minutes in a BeckMan Coulter Microfuge 22R centrifuge
(Beckman Coulter, Lawrence, KS, U.S.A) to pellet the cells. Pellets were washed with
500 ul cold SE buffer (25ml 0.5M EDTA + 7.5ml 5M NaCl + 467.5ml MQ water). After washing, the cells were resuspended in 500 ul cold SE buffer. Two hundred microlitter of
1% low melting point agarose solution were mixed with 200 ul of sample in Biorad plug molds and the plugs were cooled at 4°C for 20 min. Each plug was removed from the mold into 2 ml of lysis buffer (0.5ml 1M Tris + 2ml 0.5M EDTA + 1ml 10% Sarkosyl +
0.5ml 20mg/ml proteinase K + 6ml water) and incubated at 52°C for 24 hours. Following incubation, the lysis buffer was replaced with 2 ml TE (500ml lOmM Tris + 1ml 0.5M
EDTA) buffer and incubated at 37°C with vigorous shaking for 1 hr. This step was repeated three times. Fresh TE buffer was added each time. Plugs containing large, non- sheared chromosomal DNA were stored at 4°C in the final rinse solution. The concentration and size of the genomic DNA in the plugs was determined by pulsed field gel electrophoresis (PFGE) as described below.
3.3.3. Partial digestion of isolated chromosomal DNA with kits and in plugs
HindHI restriction endonuclease (NewEngland BioLab, Mississauga, Canada) at a concentration of lOU/ul was diluted 10, 100, 1000, and 10000 times in lx buffer 2. The
DNA (l-8[ig) extracted by the GPCT and UCM isolation kits was digested with 1-8 jxl of the diluted enzyme, and 5-20 ul lOx restriction buffer and water to a final volume of 50
63 to 200 ul were added. The reaction mixture was incubated at 37°C in a water bath for 5,
10, 15, 20, 25 and 30 minutes in 9 independent experiments. To inactivate the enzyme after partial digestion, all tubes were incubated at 65°C in a water bath for 20 min. To achieve complete inactivation of the enzyme, 5 u,l 0.5M EDTA was added following heat inactivation.
Partial digestion of DNA isolated in plugs was carried out to obtain large DNA fragments of good quality and concentration. To achieve this, varying concentrations of
Hindm (lU/|il and 10U/uJ) and different incubation times (5, 10, 15, 30, 45, 60, 90, 120,
180 or 240 minutes and 12 or 15 hours) were examined. The effect of agarose concentration used to make the plugs and gel for pulsed field gel electrophoresis was also examined. The optimal protocol was as follows. Plugs were removed from the tubes using a sterile spatula and placed on the inside of the lid of a sterile Petri dish. Two hundred ul of sterile Milli Q (MQ) water were pipette into each sterile Eppendorf tube and half a plug was placed into each tube. The tubes were agitated and then the water was replaced with 200 ul of lx restriction buffer and incubated at room temperature for 15 min. In the next step, the buffer in each tube was replaced with 200 ui of the digestion solution (89.2ul MQ water + lOul lOx NEBuffer2 + 0.8 ul 1U Hindm) and incubated at
37°C for 15, 30, 45 and 60 min, respectively. After incubation, 20 ul 0.5M EDTA were added to the tube to inactivate the enzyme and the digestion solution in each tube was removed. The plugs were washed with 1 ml TE buffer three times and stored in 1ml TE buffer at 4°C for 30 min.
64 3.3.4 Pulsed Field Gel Electrophoresis
Low melting point agarose gel (1%) was prepared by melting 1.2 g agarose powder (Invitrogen, Burlington, ON, Canada) in 120 ml of 0.5 x Tris borate (TBE) buffer. The mixture was boiled in a microwave until a clear solution was observed. The solution was cooled to 40-45°C and then was poured into a casting stand (14 cm wide x
13 cm long) and solidified at room temperature (23°C). Each digested plug, undigested plug and markers were placed into the wells and the wells were closed with a drop of molten agarose gel. The gel was placed into 2.2 L of 0.5x TBE buffer in the gel holding template in the electrophoresis chamber. Several different running conditions and running times were tested until a good separation of DNA was obtained. The best PFGE conditions were: Initial time 10 Sec, Final time 60 Sec, Run time 15 hr, Volts 6 cm,
Included angle 120, Running temperature 14°C. Half of the gel containing the marker, undigested and digested DNA was stained in 500 ml distilled water contained 40 [il 10 mg/ml ethidium bromide for 45 min and then destained for 25 min in 500 ml distilled water. The gel was analyzed using the Bio-Rad Gel Doc system (BioRad, Mississauga,
ON, Canada). Bands corresponding to fragment sizes of 90-120 kb, andl20-200 kb orl00-388 kb and 242-582 kb were excised from the other half of the gel.
3.3.5 DNA recovery from the gel slices
3.3.5.1 Recovering of digested DNA from the gel slices using Gelase
The gel slices were weighed after trimming the excess agarose around each slice and then each 200 mg portion of gel was transferred to a 1.5 ml microcentrifuge tube.
One microliter 50x gelase buffer was added to the tube for each 50 mg of gel. The gel
65 slices were melted by incubating at 70°C for 10 min. Each tube containing clear liquid was transferred to a 45°C water bath and equilibrated at 45°C for 2 min. To digest the gel slice, 2 U gelase enzyme (EpiCentre, Madison, WI, U.S.A) were added to each tube and the tubes were incubated at 45°C for 15min. The clear liquid did not gel or become viscous on cooling to 0°C. DNA was used to construct a library. Since the concentration of the recovered DNA from the gel slice using gelase was too low for constructing a library and also DNA was sheared to smaller sizes, electroelution was explored as a method to recover DNA from the gel.
3.3.5.2 Recovering of DNA from gel slices by Electroelution
CelluSep T-series dialysis membrane in combination with conventional electrophoresis was used for electroelution. The dialysis membrane (Millipore,
Massachusetts, U.S.A) was cut into a 5-10 cm long piece. It was soaked in cold water for
5 min and then in 0.5x TBE for 3 min. The tubes were rinsed with ice-cold water and then with ice-cold 0.5x TBE. A gel section containing the size-selected DNA fragments was placed into the dialysis tube. One end of the tube was closed with membrane tubing closer and it was filled with 150 u.1 ice-cold 0.5x TBE. The gel was moved to one side of the tube, all bubbles were removed and the other end of the tube was closed. The dialysis tube was submerged in a gel electrophoresis submarine cell containing 500 ml 0.5x TBE and electrophoresis was carried out for 2 hrs at 75V. The dialysis tube had rotated 180° and electrophoresis was continued for exactly one minute to force the eluted DNA off the dialysis tube wall. The DNA in the dialysis tube was carefully collected with a wide bore pipette tip and transferred to a 1.5 ml microcentrifuge tube. To remove small DNA
66 fragments that possibly were trapped between the large DNA segments, a second size selection was performed. The collected DNA was loaded into the wells of a 1% low melting point agarose CHEF gel and the same PFGE conditions described in section 3.2.4 were applied. Fragments of 100-300 kb were excised from the gel and the DNA was electroeluted from the gel slices as described above. This DNA was used to prepare the library.
3.3.6. Construction of Bacterial Artificial Chromosome (BAC) Libraries
3.3.6.1 Ligation of DNA fragments into the CopyControl pCCBAC Cloning-Ready
Vector
Three sets of DNA were used for making the library. These were i) fragmented
DNA obtained from partial digestion of genomic DNA isolated with the kits and recovered from the gel slices by gelase; ii) DNA of different sizes recovered from digested plugs using electroelution; iii) DNA recovered using electroelution from excised gel slices following a second size selection. pCClBAC vector (EpiCentre, Madison, WI,
U.S.A) (Figure 3.2), a derivative of pBeloBAC, was used as the cloning vector. One hundred ng (80 u.1) of DNA, 6 u,l sterile water and 1 ul pCClBAC vector were mixed and incubated at 55°C for 10 min. The solution was allowed to cool at room temperature for
15 min. Ten ul lOx fast-link ligation buffer, 1 ul lOOmM ATP and 2 (xl fast link DNA ligase (EpiCentre, Madison, WI, U.S.A) were added to the cooled solution in a total reaction volume of 100 |il. The ligation reaction mixture was incubated at 16°C overnight and then heated at 65°C for 15 min to inactivate the ligase. The ligation reaction was
67 Ecofi 1332 Ban iH I 3 53 Hind IN 3B3 Hpa I 7618 1 T?7/ Pci I T471 \ Sea I 793
(3sd I 6739 _ \
pjiJ CopyControl"1 ,-ecT ^—OJIZI7MS52 PCC1BAC"*
' Xfco i 2330 S>MB I 5620
SiK I 5074 EcoN 1 3<158 A/e I AS5=
Note: Not all restriction enzymes that cut only once are indicated above.
33? T7 ftjmHI Htna III p:DFP-1 pe RP-1 230-236 3I1-IU1 JU II I 300
FP = pCCI "VpEpiFOS™ Foward Sequencing Primer S GGATGTGCTGCAAGGCGATTAAGTTGG 3' RP = pCC 1 "•/pEpFOS1" Reverse Sequencing Primer S CTCGTATGTTGTGTGGAATTGTGAGC 3' 77 = T7 Promoter Primer S TAATACGACTCACTATAGGG 3'
Figure 3.2. The map of pCClBAC cloning vector (Epicentre, Madison, WI, U.S.A).
68 desalted using an agarose cone (EpiCentre) and this mixture was used for transformation.
3.3.6.2 Transformation, Plating and Selecting the BAC Clones
TransforMax EPI300 Electrocompetent E. coli (EpiCentre, Madison, WI, U.S.A) cells were thawed on ice. Three microliters of desalted ligation reaction were mixed with
50 {xl of cells in a pre-chilled microcentrifuge tube. The BioRad gene pulser XCell device
(BioRad, Mississauga, Canada) was set for bacterial (E. coli) electroporation at manufacturer's recommendations. The cell/DNA mix was transferred to the pre-chilled electroporation cuvette and it was placed into the electroporator. The electric pulse of
2500 voltage was applied and then immediately 950 ul SOC medium at room temperature were added to the cuvette and mixed gently. The cells were transferred to a 15 ml tube and incubated at 37°C with shaking at 220-230 rpm for 1 h to recover the cells and allow the expression of the antibiotic resistance marker. One hundred microliters of transformation reaction were plated on LB + chloramphenicol (12.5 (ig/ml) + X-Gal (40
Hg/ml) + IPTG (0.4mM) plates and incubated overnight at 37°C. The remaining 900 ul of the transformation reaction were stored at 4°C until the size and the quality of the BAC clones were assessed.
3.3.6.3 Sizing the BAC clones and storage of the BAC DNA
Ten clones were chosen randomly. One ml of LB broth supplemented with 12.5 ug/ml chloramphenicol was inoculated with an isolated single BAC clone from an overnight plate. Following overnight incubation, 800 ul of culture medium were removed
69 and replaced with 800 ul fresh LB broth supplemented with 12.5 ug/ml chloramphenicol.
The culture was incubated for 30 min at 37°C with shaking at 250 rpm. One microliter of lOOOx induction solution added to the culture and incubated for 2 h at 37°C with vigorous shaking. To isolate DNA from the induced culture, the BAC DNA purification kit provided by EpiCentre was used. The isolated BAC DNA was stored at 4°C overnight. Five microliters of isolated BAC DNA were loaded in a 0.7% agarose gel and electrophoresed for 4h at 50 V/cm. The size was confirmed by using EpiBlue and
EpiLyse Solution (Epicentre, Madison, WI, USA). In this method, a colony was picked and put into a microfuge tube. EpiLyse solution (25 \il) was added to the tube. The tube was vortexed to completely resuspend the cells. Ten microliters of EpiBlue solution was added to the tube. The tube was vortexed for 2 sec and then centrifuged for 30 sec at 600 g. Twenty microliters of sample were loaded in 1% low melting agarose gel and PFGE was performed under the same conditions described in section 3.2.4. When it was confirmed that BAC contained the large DNA insert, the remaining 900 jxl of the transformed cells (from section 3.6.2) were plated on LB agar supplemented with chloramphenicol (12.5 ug/ml) + X-Gal (40 ug/ml) + IPTG (0.4 mM) and incubated at
37°C overnight.
For short term storage, two replicates of each plate were prepared, incubated at
37°C overnight and then stored at 4°C. For long term storage, each clone was inoculated in 50 ml tubes containing 5 ml MB supplemented with 5 u,l (12.5mg/ml) chloramphenicol and then incubated at 37°C overnight while shaking at 200 rpm. One milliliter of overnight culture was mixed with 1 ml freezing media (EpiCentre) and stored at -80°C.
70 3.3.7 Screening of the Library
3.3.7.1 Growth condition
One hundred and twenty recombinant E. coli were chosen randomly from the library and a single colony of each was cultured in 50 ml tubes containing 5 ml LB broth supplemented with chloramphenicol (12.5 mg/ml). Cultures were incubated at 37°C overnight. Five milliliter inoculums of each clone were transferred to 250 ml LB broth supplemented with 250 \il chloramphenicol (12.5 mg/ml) and incubated at 10°C for 10 days with shaking at 180 rpm.
3.3.7.2 Fatty acid extraction
Cells were harvested by centrifugation using a Beckman J2-MC centrifuge at
10000 g for 17 min at 10°C. The cells were washed two times with MQ water, freeze- dried overnight in a Stokes Freeze-Drier (Equipment Division Pennsalt Chemicals Crop,
Philadelphia, PA, U.S.A) and ground to a fine powder in a clean sterile mortar with a pestle. The powders of each sample were weighed and were transferred to 20 ml screw cap test tubes. Fatty acid was extracted according to the method described in Section
2.3.9.
3.3.7.3 Gas Chromatographic Analysis
The fatty acid methyl esters were analyzed by using an automated Agilent 6890
GC system (Agilent, Palo Alto, CA, U.S.A). Two microliters were injected into the GC.
A fused silica BPX 70 capillary column (60 m x 0.22 mm ID) and a flame ionization detector were used to separate the fatty acids. The flow rates of the carrier gas
71 (hydrogen), air, and hydrogen for the flame were 1 ml/min, 400 ml/min and 30 ml/min, respectively. The GC analysis conditions are described in Table 2.3. The fatty acid methyl esters were identified using known standards of 37 fatty acid methyl esters from
C4:0 to C22: 6n3 and pure EPA/DHA methyl esters (Sigma, Oakville, On, Canada). To prevent any carry over from previous samples, the initial temperature of the GC was increased to 280°C for 15 min to remove any remaining fatty acid residue and after that the column was cleaned by iso-octane for 45 min. The syringe used to load the column was also washed in clean iso-octane 5 times between each injection. To confirm
EPA/DHA production by clones, fatty acids of positive clones were sent to the McMaster
Regional Centre for Mass Spectrometry, McMaster University (Hamilton, ON, Canada).
3.3.8 Analysis of EPA/DHA positive clones
3.3.8.1 Bacterial strain and growth conditions
Fresh colonies prepared from E. coli transformants positive for EPA/DHA,
EPI300 E. coli and blue clones (containing the BAC vector without insert) were stored at
-80°C in freezing media. An isolated colony from each plate was transferred to a 50 ml
Erlenmeyer flask containing 10 ml LBB supplemented with 10 [A1 12.5 mg/ml chloramphenicol for the blue clone; 50 ml Erlenmeyer flask containing 10 ml LBB + 12.5
(xg/ml chloramphenicol + 60 \i\ BAC autoinduction solution for positive clones; and 50 ml flask containing 10 ml LBB for EPI300 E. coli. The cultures were incubated at 37°C with shaking at 250 rpm overnight. To increase the BAC DNA concentration, Terrific
Broth (TB) was used in addition to LBB to obtain more cells. Moreover, larger culture
72 volumes were prepared by inoculating colonies in 50, 100, 250 and 500 ml LBB/TB supplemented with 12.5 fig/ml chloramphonicol + 6u,l/ml autoinduction solution.
3.3.8.2 BAC DNA isolation from positive clones
BAC DNA was isolated from 1 ml, 3 ml, 5 ml, 10 ml, 50 ml, 100 ml, 250 ml and
500 ml cultures using the BACMAX DNA purification kit (EPICENTRE, Madison, WI,
U.S.A), Qiagen Large-Construct kit (Qiagen, Mississauga, ON), or Nucleobond BAC isolation kit (Clontech, CA, U.S.A) according to the manufacturer's instruction. In addition, published methods for BAC DNA extraction were also employed (Villalobos et ai, 2004). The extracted BAC DNA from positive clones was sent for end sequencing to the Guelph Molecular Supercentre (Guelph, ON, Canada). It was also subjected to complete digestion.
3.3.8.3 Complete digestion of BAC DNA
To isolate the insert from BAC DNA, the complete digestion of BAC DNA was performed using Hindlll. The reaction mix consisted of 5 u.1 of BAC DNA + 2 jxl
NEBuffer 2 + 1 n 1U Hindlll + 12 \il H20 and this was incubated at 37°C for 225 min. In other experiments, NotI, BamHI, Apal and EcoRI restriction enzymes were used to digest the BAC DNA. The amount of BAC DNA, enzymes and incubation times (3 h to overnight) were variable in each set of experiment (Table 3.1).
73 Table 3.1. Digestion conditions for BAC DNA
Restriction Buffer (lOx) BAC DNA Water Incubation time
Enzyme at 37°C
Hindm (l-3|il) Buffer 2 (2 jil) 1-10 nl 4-16 |il 3-24hrs
Mrtl(l-3[il) Buffer 3 (2 |il) 1-10 nl 4-16 |il 3-24hrs
BamRl (\-3\x\) Buffer 3 (2 |il) 1-3 |il 12-16 |il 24hrs
Apal(l-3|il) Buffer 4 (2 |il) 1-3 |il 12-16 |il 24hrs
EcoRl (l-3|il) Buffer 1 (2 fil) 1-3 |il 12-16 |il 24hrs
74 3.3.9 Determination of EPA/DHA production by positive clones in Skim milk
3.3.9.1 Bacterial strains and Growth condition
EPI300 E. coli as a negative control, EPA/DHA positive clones and S. baltica
MAC1 as a positive control were streaked on LB agar, LB agar + 12.5 ng/ml chloramphenicol and Marine Agar, respectively. An isolated colony of each of E. coli,
EPA/DHA positive clone and S. baltica MAC1 was inoculated into 5 ml LB broth, 5 ml
LB broth + chloramphenicol and Marine Broth, respectively. Cultures of E. coli and the positive clone were incubated at 37°C overnight while shaking at 200 rpm. 5. baltica
MAC1 was incubated at 30°C overnight with shaking at 70 rpm. Five milliliters of these cultures of E. coli, the positive clone and S. baltica MAC1 were transferred to 250 ml
LBB, 250 ml Skim milk ± chloramphenical and 250 ml MB, respectively and incubated at 10°C, 15°C, 25°C or 30°C on a shaker at 200 rpm for 10 days, 5 days, 2 days and 1 day, respectively.
3.3.9.2 Fatty acid extraction and GC analysis
Cells were harvested, freeze-dried, ground to a powder and then subjected to the fatty acid extraction and GC analysis as described in sections 2.3.9 and 3.3.7.3.
3.4 Results and discussion
3.4.1 Preparation of a BAC library of S. baltica MAC1 genomic DNA
The genomic DNA of Shewanella baltica MAC1 was extracted using different commercial kits in order to get a high molecular weight (HMW) DNA. The size of
75 genomic DNA obtained was 194 kb and 130 kb using GenomicPrep Cells and Tissue
DNA isolation and UltraClean Microbial isolation kit, respectively. After a series of preliminary experiments, an optimized method was established for partial digestion of the DNA. The DNA fragments (50-120 kb) were recovered from the gel using gelase.
The recovered DNA was used for construction of the BAC library. The transformation efficiency was too low and only 20 recombinant clones were recovered following each electroporation.
In order to obtain high-quality HMW DNA, genomic DNA of S. baltica MAC1 was isolated from the low melting point agarose plugs. The size of HMW DNA in LMP agarose plugs was 900-1000 kb. The plugs containing HMW DNA were partially digested under different conditions until the size of the digested DNA ranged from 100-
900 kb. The DNA fragments with sizes 100-200 kb and 200-350 kb were excised from the gel and recovered from the gel slices using gelase. The recovered DNA was used directly to make a BAC library. After electrotransformation, a total of 35 colonies were collected from 2 ligations. The efficiency of the library was still low. The experiment was repeated incorporating the following modifications: the DNA was digested in 1% low melting agarose prepared with lx TAE instead of 0.5x TBE; the initial condition for
DNA separation changed from l-10s pulses over 16 h to 10-60 pulses over 20 h for
PFGE; and the recovered DNA was concentrated and purified by ethanol precipitation. A total of 61 clones were collected from 2 ligations. The efficiency of the BAC library was still not optimal. It is reported that nucleic acids of any size, from less than 50 kb to 1 megabase DNA can be easily purified intact and in high yield and can be used directly for
76 cloning. However, the residual electrophoresis buffer salts and oligosaccharides generated by digestion of the agarose decreased the efficiency of ligation and transformation (Folkertsma et al, 1999). It is suggested that purification and concentration of DNA following gelase digestion can improve the efficiency.
Nevertheless, DNA will shear as a result of many centrifugations during the purification
(Chen et al, 1994). Our results show that using gelase for recovering large DNA fragments is not a good choice since the efficiency of ligation and transformation was affected by salt contamination and the DNA purification procedure. It was shown by other researchers that electroelution resulted in more intact DNA than agarose digestion
(Strongs al, 1997).
To recover high quality and quantities of DNA fragments from a gel, a dialysis membrane was used. DNA fragments in the 100-585 kb size range were excised from the gel (Figure 3.3) and electroeluted into the dialysis membrane. The concentration of electroeluted DNA was 20 ng/[il. This DNA was ligated and transformed to EPI 300
E.coli. Nine hundred and forty white colonies were obtained on the plates. In addition, to remove all possible small DNA fragments (< 100 kb) that usually are trapped among the large DNA fragments, a second size DNA selection was performed by loading the electroeluted DNA in 1% low melting agarose gel and varying the PFGE conditions for better separation of HMW DNA from smaller fragments. After the second size selection,
DNA fragments with a size of 100-300 kb were excised from the gel. The DNA was electroeluted from the gel slices and the solution desalted using an agarose-cone. DNA
77 M M 1 2 3 4
B
300 kb i
100 kb'
C
582 kb
242 kb
' <- wMSrMm
Figure 3.3. PFGE of partially digested genomic DNA isolated from S.baltica MAC1 in 1% LMP agarose. A) columns M show the PFG marker with size of 1020 kb; 1 is undigested DNA; 2, 3, and 4 are partially digested DNA with 1 unit of Hind III/100 ul reaction; B) shows sizes 100-388 kb which were cut out of the gel; C) shows sizes 242=582 kb.
78 was ligated to the BAC vector and transferred to the electrocompetent EPI300 E. coli. A total of 490 clones from second size selection were obtained from 5 ligations. The second
PFGE eliminates the smaller fragments, however, it greatly decreased the DNA concentration and recovery, which results in a low efficiency BAC library. When a second PFGE was conducted, 2 ng/^1 DNA were obtained, which is 10-times less than following the first PFGE. A decrease in DNA concentration after the second PFGE was also reported when a BAC library was prepared from the marine macroalga Porphyra yezoensis (Beja et al, 2000; Deng et al, 2004).
Ten clones were chosen randomly from the library to size the BAC clones. BAC
DNA was extracted from the clones and subjected to PFGE on 1% low melting agarose gel (Figure 3.4). Two fragments were obtained with sizes of 50 kb and 200 kb. This indicated that BAC clones had the large DNA insert. All clones were stored in freezing media at -80°C.
The preparation of high-quality DNA for construction of a BAC library is critical.
When the quality of DNA is poor, the ligation and transformation steps will fail or will be at low efficiency (Deng et al, 2004). In this study, we found that preparing HMW DNA and particularly, recovering HMW DNA from the gel slices were the major obstacles for the construction of a BAC library. To reduce DNA degradation and impurities, each time we used the prepared plugs immediately for partial digestion or we maintained the prepared plugs in 0.5M EDTA at 4°C for less than 48 hrs. It is suggested that the plugs can be maintained in 0.5M EDTA at 4°C for several months (Zhang et al, 1995) or in
79 M M 1 2 3 4 5
• '. \ * 339 kb,
291 kb' • 1 tfB _ 242 kb. y* •» "l. "' *.•'- . >"••;*; "*v -'TV:
„ * A ** -.- . • -.-^
. • ;^ 48 kb * ;. >if •"• - "
• ..it? • . *.".«a«fc.!&.
Figure 3.4. PFGE (1% LMP) of BAC DNA extracted from randomly selected clones
from the BAC library. M shows the PFGE markers; lane 1 shows BAC
tracker supercoiled marker; 2, 3,4, and 5 are BAC plasmid containing the
insert isolated from the clones.
80 70% ethanol at -20°C for a year (Luo et al, 2001). Most of the BAC libraries have been constructed from land plants, humans, animals, insects and fungi, but few from bacteria, particularly marine bacteria, have been reported (Beja et al, 2000; Sabehi et al, 2005;
Suzuki etal., 2004).
We were able to establish a method to prepare HMW DNA in megabase sizes as well as large DNA fragments of 100-300 kb, which were needed to construct a BAC library from S. baltica MAC1 in order to clone all the genes required for EPA/DHA production.
3.4.2 Screening of the BAC library
The best way to screen a library when there is no information available about the gene(s) of interest is to look for the product of the gene. One hundred and twenty BAC
DNA clones were chosen randomly from the library. A blue clone (containing only BAC vector) and electrocompetent E. coli were the negative controls and Shewanella MAC1 was used as a positive control. Fresh samples of all clones and controls were prepared and the fatty acids of all samples were extracted and analyzed by GC. The fatty acid profile of all clones were almost the same as negative controls except for three clones (,
#34 and #48) which gave two extra peaks at retention times corresponding to eicosapentaenoic and docosahexaenoic acid peaks. The fatty acid profiles of these 3 clones are shown in Figure 3.5. Retention times of the suspected EPA/DHA peaks in the fatty acid chromatograph of clone # 17 is compared with the retention time of an
EPA/DHA standard in Figure 3.6.
81 s pA i r- I - 36-1 > 3s Q DHA 35-3 en 34- CM 5 4 -29.17 : 36.61 7 33- C\o i a i EPA j CO 32- 31- 30- 29: o CM CD DHA LOO T? C0> T-0> o CO o CM co CO CM CO c\j CO CO IU^JLLJ a UJJL V-j5ly_^ Figure 3.5. GC of 3 clones selected from the library. Gas chromatography of A) clone # 17; B) clone # 34; C) clone # 48. GC condition is outlined in Table 2.3. The cultures were grown at 10°C and FAs extracted from cells and analyzed using an Agilent 6890 GC. 82 DHA B r^c\j —-A -U DHA CM EPA 35 i 30-j Figure 3.6. GC of the fatty acid profile of positive EPA/DHA clone and Standard. A) Clone # 17 before injection of internal standard; B) clone # 17 after injection of internal standard; C) EPA/DHA standard. Clone # 17 was grown at 10°C for 10 days and extracted FAs form cells was analyzed using an Agilent 6890 GC. 83 To verify the EPA/DHA peaks in clone # 17, the sample was divided into two portions and 1 \il EPA/DHA standard with a concentration of 10 mg in 200 ml iso-octane was injected into one portion of the sample. The fatty acid profile of sample containing the internal standard and the one without internal standard were analyzed under the same conditions (Figure 3.6). The suspected peaks became larger while other peaks stayed the same or were a bit smaller because they were diluted when they were mixed with EPA/DHA standard. These three EPA/DHA-producing clones were stored at -80°C, refreshed, cultured in duplicate in LB broth supplemented with chloramphenicol and incubated at 10°C for 10 days. Subsequently, their fatty acids were extracted and analyzed by GC. The production of EPA/DHA was reconfirmed for all 3 clones. The EPA/DHA positive clones, MAC1 and pure EPA/DHA standards were sent to McMaster Regional Centre for Mass Spectrometry, McMaster University, Hamilton for confirmation. The standard sample showed the EPA and DHA eluting at 29.4 and 33.3 min, respectively. Figure 3.7 shows the chromatogram for the EPA/DHA standard, the mass spectra for the 29.4 and 33.3 minute peaks and the results of library database searches for the two mass spectra: 5,8,11,14,17 eicosapentaenoic acid methyl ester and 4,7,10,13,16,19-docosahexaenoic acid methyl ester. The results showed similar spectra for EPA and DHA. Differences in the retention time in the GC and GC-MS was due to the different columns used in each instrument. The chromatogram and mass spectra for clone 17 is shown in Figure 3.8. The library database search gave similar spectra for EPA and DHA peaks in clone 17. There is an intense peak at 33.3 minutes (DHA retention time), as well as a very weak peak at 29.4 minutes (EPA retention time). The spectra are 84 DILUTED DHA+EPA MWG5973B 3329 ItKh 29.40 J L i|iiii|iiii|iiii|iiiii'ii>iiiii|ii"i,,M I"11 i""i""M 'I""!'"1!1111!11"! 'I MWG5974 DILUTED DHA+EPA MWG5973B 3164 (29.362) Cm (3162:3164-3144:3146x1.500) 100n 79.0533 191.0529 93.0683 29.4 MIN .106.0755 133.0975 41.0386 145.1001 161-1167 1 201.1637 ' 247.1673 £I|I li'll-.liVtL U el- MWG5973B 3629 (33.237) Cm (3625:3629-3608:3612x1.500) 1«h 79r7.91.0505 33.3 MIN 93.0670 119.0824 67.0528 131.0830 145.1006 A ,J,i. rri!":.0^81^i6-.?2^a^.i7B?^4i.i785 /731881 31 M 3.2171 40 60 80 100 12M0 140 160 180 200 220 240 260 280 300 320 340 360 79 100- 91 67 '93f.119 41 55 „133 r147 180 201215^220 247 JU 1 i i i i | i i r i I l"l | l'"l I "I'| JI")U II "| I T I I'| I I 1 I [ I I I I | I I I l'| I I I I | I I I I | I I I I | I I I I | I I F:764 MIST 30776: 5.8.11.14.17-EIC0SAPENTAEN0IC ACID. METHYL ESTER. 100- 7? 91 93105119 67 / / / 131 166 145 15 <55 /"20U06/ 241255 273 313314 yju W^ riVi'l'M I'I'I'I r 1111 n'lTrTrp'TH'irn1 m rrf4*r w rr\ \ n ip'i'n F:826 NIST 30778: 4,7,10,13,16,19-DOCOSAHEXAENOIC ACID, METHYL ESTER Figure 3.7. The mass spectra for EPA/DHA standard. A) EPA/DHA chromatograph analysed by GC-MS; B) mass spectra for EPA peak at retention time of 29.4; C) mass spectra of DHA peak at retention time of 33.3; D) mass spectra for EPA obtained from FAs library database search; E) mass spectra for DHA obtained from FAs library database search. 85 IVC IOC MWG5974 DHA 100-1 EPA IVC 10C MWQ5974 3169 (29.410) Cm (3167:3171-3124:3128x1.500) 79.0544 100-, 191.0535 93.0698 29.4 MIN 105.0699 67.0543 / 1481163 180.1167 201.1607 243.2135 257 2072 Ld W I ,1 / | &IJUU 303.2397 402.9735. MWG5974 3630 (33.252) Cm (3624:3630-3589:3595x1.500nJlH LJlLi ) '.Iwlhi I , I i,,<, i ,,i 79.0503 100-] .£1.0497 33.3 MIN 93 0664 67.0522 / 119.0813 131.0828 145.0998 C | r173.1278 201.1517 241.1749 273.1884 327.2538 313.2175 I'l'l'l'lp1 LMI' , v i'l'l'l'l'ITl hi C\ .,,.m C •, <,,,„;, < 40 60 80 100 u12 0 140 160 180 200 220 240 260 280 300 320 340 360 '380 Figure 3.8. Mass spectra for clone # 17 (EPA/DHA positive clone). A) chromatograph of peaks at retention time 29.4 and 33.3 analysed by GC-MS; B) mass spectra for EPA peak at retention time of 29.4; C) mass spectra of DHA peak at retention time of 33.3. Clone # 17 was grown at 10°C for 10 days and fatty acids extracted from 0.1 gr dry cells. Extracted FAs were analysed by GC-MS in McMaster Regional Center for Mass Spectrometry. 86 visually quite similar to the standard spectra (Figure 3.9), and also yield the same library search results. For clones 34 and 48 only a very weak peak at 29.4 minutes was observed. However, no useful spectral data were obtained from the 29.4 minute peak for these clones. No peak was detected for clones 34 and 48 at the DHA retention time. In addition, the GC-MS result for the positive control (S. baltica MAC1) gave an intense peak at 29.3 minutes and a relatively weak peak at 33.5 minutes. The chromatograph and corresponding spectra for S. baltica MAC1 are shown in Figure 3.10. The library search for the peak at 33.5 minutes returned EPA as the top hit, and the spectrum closely resembled that of EPA and DHA; indicating that EPA and DHA are very closely related compounds. EPA and DHA were not detected by GC-MS for clone 34 and 48, this could be due to the very low concentration of these compounds in those samples. Clone # 17 was the only one among the 3 clones in which production of both EPA and DHA was confirmed by GC and GC-MS. Recombinant production of EPA in E. coli was first reported by Yazawa (1996) and recently Orikasa et al. (2006) reported recombinant production of DHA in E. coli. So far, no reports have been published in which recombinant production of both EPA and DHA in E. coli has been described. Results from this study show, for the first time that both EPA/DHA genes can be expressed in E. coli at the same time. It was previously reported that only five genes, p/aABCDE, are generally mandatory for the biosynthesis of EPA and DHA. Even though the basic structures of all pfa genes for EPA and DHA biosynthesis are the same, the domain structure of some pfa genes is not quite the same. 87 IVC 10C MWG5974 3169 (29.410) Cm (3167:3171-3124:3128x1.500) 100n 79.0544 191.0535 93.0698 29.4 MIN 105.0699 67.0543 / 133.1007 66.0472 148.1163 180.1167 201.1607 243 2135 o^omo I , , y Z4J.^1J5 257.2072 3032397 342.0197 402.S • I , Wl Hill ill I^Xl ,., I I'l'l'iii'irliru'il'hfiI. I •• if.I i MWG5974 3630 (33.252) Cm (3624:3630-3589:3595x1.500) 79.0503 33.3 MIN 93.0664 / 119.0813 131.0828 241.1749 273.1884 327 2538 .173.12781201.1517 / 313.2175 / '•'" •••'. i.r.i. .. . . W* \rA \jkr Il'i'j-, i'.'I'l',I, , J..• | |W|.|, JL.'I I I"'i •''||'"| '") ~ ' "I" 40 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 DILUTED DHA+EPA MWG5973B 3164 (29.362) Cm (3162:3164-3144:3146x1.500) 100-1 79.P"'3 191.0529 93.0683 29.4 MIN / 67.0538 -106.0755 133.0975 41.0386 145.1001 1611167 215 1604 „ III / ,,i 201.1637 ^b'1bU4 247.1673 II, ,1 *WJJ. (A. r MJ 'ill I'l'l'l'l'l I I'I'I '•'•'I 'l ' MWG5973B 3629 (33.237) Cm (3625:3629-3608:3612x1.500u ) c 1«h 79Cf507.91.0505 33.3 MIN 93.0670 119.0824 67.0528 131.0830 1451006 41.0385 (. , -166.0981 731881 206.1296^215.1782,241.1785 ! 313.2171 'I' i I 'I lit'',I iJiLUI k'" ' 1 1 ri'i'l'''''I1i' '! • "'i Ii ••t'l i'iI "•''• I1 1I'I1' •' ! I I^''• I• • •• I 40 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 36i Figure 3.9. Comparison of mass spectra of EPA and DHA standard with mass spectra of EPA and DHA peaks from clone # 17. A) mass spectra of EPA peak in clone # 17; B) mass spectra of DHA peak in clone # 17; C) mass spectra of EPA peak in standard; D) mass spectra of DHA peak in standard. 88 EPA 29.44 DHA 33.53 MACI+EPA B MWG5976 3184 (29.535) Cm (3184:3185-3140:3141x1.500) 100i 79.0437 91.0436 -93.0631 119.0798 29.4 MIN 133.0978 147.1107 180.1187 201.1673 •220.15642,47-1798 ll , f i . .., . 287?173 316.2599 It I|IP'I |l)l|'lvI| | VI'A'itwii ..•!•(•• ,•,••.')•! .....•,).. MWG5976 3663 (33.527) Cm (3660:3666-3671:3677x1.500h ) iC 100! 79"528 33.5 MIN /93.068 5 119.0834 133.0988 147.0975 208.1457 275.2028 348.189C 1 1 1 u l|,iili| ll A 11 '''I' " ' !' •'" |1 1 jp~1 W'" ' •' I I • •' • »• 11 • • •=» 40 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 Figure 3.10. Mass spectra of FAs of S.baltica MAC1 (positive control). A) chromatograph of peaks at retention time 29.4 and 33.5 analysed by GC-MS; B) mass spectra for EPA peak at retention time of 29.4; C) mass spectra of DHA peak at retention time of 33.5. S.baltica MAC1 was grown at 10°C for 3 days and fatty acids extracted from 0.2 gr dry cells. Extracted FAs were analysed by GC-MS in McMaster Regional Center for MS. 89 An extra repeat of A:acyl carrier protein acyltransferase in pfaA gene; a 3-keto-acyl synthase (KS) in pfaB; in pfaC, 2 domains 3- hydroxydecanoyl-ACP dehydratases (HD) similar to the Fab A and 1 HD similar to the FabZ/FabA; are the extra factors for the production of DHA (Okuyama et al, 2007). In addition, the PPTase gene or pfaE may have slightly different domains for the synthesis of EPA or DHA in recombinant E. coli (Orikasa et al, 2006a). They reported 4 protein domains designated as P0, PI, P2 and P3. The PI domain was recognized separately as two subdomains of Pla and Plb. They suggested that P2 and P3 participates in Mg2+ binding; Pla, Plb and P3 might be involved in substrate binding and catalysis; and the P0, Pla and Plb domains might be involved in the recognition of the tertiary structure of the substrates carrying repeated ACP domains in the pfaA gene. They cloned the PPTase gene (pfaE) from Moritella marina MP-1, a DHA-producing bacterium, and the gene was expressed in E. coli along with pfaA-D genes derived from S. pneumatophori SCRC2738. E. coli produced only EPA (Orikasa et al., 2006a). In another study, the PPTase gene was cloned from an EPA-producing bacterium and expressed in E. coli with pfaA-D genes derived from Moritella marina MP-1, a DHA producing bacterium. The recombinant E. coli produced DHA (Sugihara et al, 2008). However, it is not well defined which protein domains in the pfaE (PPTase) gene are responsible for EPA and which ones are needed for DHA production. We speculate that clone #17 with a large insert of 200 kb probably is carrying all pfaA-E genes responsible for EPA and DHA production with their complete protein domains. 90 3.4.3 Isolation, digestion and end sequence of EPA/DHA positive BAC DNA To analyze and further study the large insert DNA in the positive clone, a series of preliminary experiments using conventional methods (Sambrook & Russell, 2001), different protocols adapted from manuscripts (Lijavetzky et al, 1999; Sambrook & Russell, 2001; Thomas et al, 1988; Villalobos et al, 2004) and several commercial kits (with slight modifications) were used to isolate BAC DNA. There were two obstacles encountered when using different extraction methods: firstly, the large insert was susceptible to physical damage and secondly, the yield of DNA was low. A BAC DNA at a reasonable concentration was obtained from 100 ml freshly induced EPA/DHA positive clones using the Qiagen kit with a few modifications (using TBB instead of LBB, decreasing centrifugation speed, incubating DNA in ethanol and sodium acetate at -80°C overnight). The yield of BAC DNA ranged from 0.5-5 [ig/ml of culture, depending on the culture volume and the autoinduction solution used. DNA samples were digested with Hindlll and NotI and then were analyzed by PFGE on 1% low melting point agarose gels to check their insert sizes. Surprisingly, BAC DNA was not digested with any of the enzymes. Digested and undigested samples showed two fragments (Figure 3.11). The lower fragment is the supercoiled form of the BAC DNA and the upper one is the nick form of the BAC DNA. In the next step, the BAC DNA of positive clones, negative clones, and a blue clone, which contained only the BAC vector, was isolated and it was cleaned up from chromosomal DNA by Plasmid-safe DNase. The purified BAC DNA was subjected to digestion using different restriction enzymes such as EcoRl, Bamiil, Apal, NotI, Xbal and pstl that cut pCCClFOS vector in 1 to 3 sites. In addition, for each set of digestion using different enzymes, different incubation times starting from 3hrs to 91 MM 12 3 4 5 6 7 8 9 10 11 200 kb 48kb 9kb iS(Sj*jn--sj-* ; _..*.•_., - -. Figure 3.11. PFGE (1% LMP) of digested and undigested BAC DNA from clones that were positive for EPA/DHA production and those that were negative for EPA/DHA production. Lanes M, PFG markers; 1, digested clone # 17; 2, digested clone # 34; 3, digested clone # 37; 4, digested clone # 48; 5, digested blue clone with Hind III; 6, digested blue clone with Not I; 7, undigested clone # 17; 8, undigested clone # 34; 9, undigested clone # 37; 10, undigested clone # 48; 11, BAC tracker supercoiled (EpiCentre). 92 48 hrs, and different enzyme and DNA concentrations were used. No digestion was observed except for the BAC vector. The qualities of the Hindlll, and Not 1 were checked by using lu.1 enzyme to digest uncut BAC vector or l(Ag genomic DNA isolated from S. baltica MAC1 with a reaction time of 3 h. The activities of enzymes were satisfactory when genomic DNA and vector were digested. Different hosts were tried for BAC DNA to eliminate the possible inhibitors that might have been produced by EPI300 E. coli and affect enzyme activity. The purified BAC DNA was transferred to DH5oc and DH10B and then plated on the LB agar supplemented with chloramphenicol, Xgal and IPTG. The BAC DNA was isolated from the cells and subjected to digestion. The enzymes were not able to cut the BAC DNA. Since we were not able to digest the BAC DNA, we checked the insert by end sequencing. The results for end sequence of BAC DNA came back negative, no signal was obtained for the insert despite the use of different primers and repetition of the experiments several times. This suggested that the positive clone might have lost the insert. To ensure that the clone has the insert, fresh culture was prepared on LB plates + chloramphenicol + X-gal + IPTG, and this resulted in good growth of the positive clone and the white color of the clone indicated that it still has the insert. Moreover, the size of the isolated BAC DNA from the clone was 100-200 kb. In addition, the EPA/DHA positive clone was rechecked for EPA/DHA production and the result was positive both with GC and GC-MS. At this point, we concluded that we probably had a chimeric clone. It is reported that 1-6% of BAC library clones could be chimeric (Berton et al., 2003; Crooijmans et al, 2000; Osoegawa et al, 2000). Methylation of the Hindlll site might be 93 another reason that prevents the restriction enzyme from digesting the BAC DNA. Nevertheless, the other enzymes that were used for partial digestion also did not digest the BAC DNA. 3.4.5 Assessment of Recombinant production of EPA/DHA in LB Broth and Skim Milk at different temperatures Skim milk was used to examine whether the EPA/DHA positive clone can produce EPA/DHA in a food environment. The production of EPA/DHA in EPA/DHA- producing clone 17 was tested at 10°C, 15°C, 25°C and 30°C in LB broth and skim milk. EPI300 E. coli and 5. baltica MAC1 were negative and positive controls, respectively. DHA production in LBB and skim milk ± chloramphenicol was detected at all temperatures. Traces of EPA were detected in LBB and a very small amount was produced in skim milk both in the presence and absence of chloramphenicol (Table 3.2). The DHA concentration in LBB, skim milk with choloramphenicol and skim milk without chloramphenicol at different temperatures is shown in Table 3.3. The production of DHA in EPI300 E. coli cells carrying the BAC vector with an insert size of 200 kb at 10°C, 15°C, 25°C and 30°C was 0.85%, 0.7%, 0.3% and 0.05% of total fatty acids, respectively. The level of DHA increased with decreasing growth temperature. The DHA concentration when transgenic E. coli was cultured in LB broth and grown at 10°C, 15°C, 25°C and 30°C was compared with DHA levels of this clone when cultured in skim milk with and without chloramphenicol at 10°C, 15°C, 25°C and 30°C (Figure 3.12). DHA production by S. baltica MAC1 is 0.4% and EPA is 4-10% at the lowest temperature. The level of EPA and DHA in S. baltica decreased to 0% when the growth temperature was 94 Table 3.2 EPA concentration in dry cells (mg/0.2 g) of transgenic E. coli (clone 17) cultured at different media and temperatures. Media/Temperautre 10°C 15°C 25°C 30°C LB Broth tr tr tr Tr Skim Milk ± Chloramphenicol 0.02 0.031 0.013 0.017 tr: traces of EPA Table 3.3 DHA concentration in dry cells (mg/0.2 g) of transgenic E. coli (clone 17) cultured at different media and temperatures. Media/Temperautre 10°C 15°C 25°C 30°C LB Broth 0.47 0.48 0.17 0.057 Skim Milk + Chloramphenicol 0.08 0.098 0.056 0.018 Skim Milk - Chlorampphenicol 0.073 0.097 0.062 0.012 95 H10°C H15°C Q25°C S30°C m &a_ LBB SM SM+Chl Growth media Figure 3.12 Comparision of DHA concentration (mg/g) in transgenic E. coli grown at different temperatures and different media. DHA concentration extracted from transgenic E.coli, when cultured in LB broth and grown at 10°C, 15°C, 25°C and 30°C was compared with DHA levels of this clone when cultured in skim milk with and without chloramphenicol at 10°C, 15°C, 25°C and 30°C. DHA was extracted from 0.2 g dry weight of transgenic E. coli and analyzed using an Agilent 6890 GC. Results show mean ± SD from 3 independent experiments. 96 increased to 30°C (Amiri-Jami et al, 2006). When autoinduction solution was added to the growth medium, the production of DHA increased by 1.4%. The autoinduction solution increases the copy number of the BAC vector carrying the DHA gene cluster, which results in a higher production of DHA. According to Orikasa et al. (2004), more EPA was produced when E. coli cells were transformed with a high copy number plasmid carrying the EPA gene cluster. The recombinant production of DHA in transgenic E. coli DH5a at 10°C, 15°C, 20°C and 25°C was reported to be 3%, 5%, 1.3% and 0%, respectively (Orikasa et al, 2006b). They coexpressed the PPTase gene with pfaA-D genes in E. coli. It was also reported that EPA yield increased by 1.2% when the cyanobacterium Synechococcus harboring the pEPA plasmid was cultured at lower temperature (18°C) (Yu et al, 2000). Moreover, there is a significant difference (p < 0.05) in DHA concentration when EPI300 E. coli grown in LB broth was compared with DHA levels produced when it was cultured in skim milk at different temperatures. No significant differences (p > 0.4) were observed between DHA concentrations in cells grown in skim milk supplemented with chloramphenicol and skim milk without chloramphenicol. The growth media components have a direct effect on EPA/DHA biosynthesis. S. baltica MAC1 produced the smallest amount of EPA (10-33 mg/1) on buffered peptone water and Pseudomonas broth because of the low concentrations of sodium chloride in these media, but MAC1 produced 105.5 mg/1 when it was cultured in unsalted sweet whey containing 1% NaCl (Cadieux et al, 1998). It was suggested by Cadieux et al. (1998) that modifying the growth medium increased the synthesis of EPA and DHA in S. baltica MAC1. It is also reported (Morita 97 et al, 2005) that when DHA-producing Moritella marina MP-1 was cultured in the medium containing cerulenin, the content of DHA in the total FA increased from 5.9% to 19.4%. In addition, cerulenin treatment increased the EPA yield in EPA-producing S.marinintestina IK-1 (Morita et al, 2005). The levels of EPA and DHA production could be increased if the insert containing the EPA/DHA genes was ligated to a broad host range plasmid and transfered to another host or by appropriate selection of growth media components and growth temperatures. 3.5 Conclusion We successfully cloned the EPA/DHA gene cluster by constructing a BAC library with large genomic DNA inserts. The transgenic E. coli obtained was able to produce EPA in a trace amount and DHA corresponding to 0.85% of the total fatty acids in LB broth. The recombinant E. coli also produced EPA/DHA in skim milk. Our attempt to subclone the EPA/DHA gene cluster into a food grade broad host range vector was not successful due to an inability to digest the BAC DNA and isolate the insert. In our experience, although, constructing the BAC library with large inserts increased the chance of cloning EPA/DHA gene cluster, this proved to be a challenging and time consuming technique. In addition, a BAC library with large DNA fragments significantly increases the chance of chimerism that prevents isolation of the insert for subcloning. However, the EPA/DHA clone can be used to study gene expression and the enzymes involved in EPA/DHA production. 98 4. Cloning omega-3 fatty acids genes from Shewanella baltica MAC1 by constructing a fosmid library. 4.1 Abstract Polyunsaturated fatty acids such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) are essential for human health. Shewanella baltica MAC1, a marine bacterium isolated from mackerel entrails, produces EPA and DHA. To clone the EPA/DHA gene cluster of S. baltica MAC1, 40 kb genomic DNA fragments of S. baltica MAC1 were cloned into the fosmid vector (pCClFOS). A fosmid library of 12,000 clones was obtained with insert sizes ranging from 34-41 kb. Two thousand five hundred fosmid clones were screened by colony hybridization for pfaA (ORF 5) and pfaD (ORF 8) genes. In total, 31 clones were positive for either pfaA or pfaD genes. These clones were further subjected to PCR for both pfaA and pfaD genes of which eight were positive for both genes. These eight clones were examined for EPA/DHA production. One clone was detected to produce both EPA (13% of total fatty acids) and DHA (0.3% of total fatty acids). Six clones were found to produce only EPA (4.8- 14% of total fatty acids), and one clone did not produce either EPA or DHA. We were able to isolate fosmid DNA of positive clones, digest them and perform end-sequencing to confirm the presence and the size of the inserts. The complete sequence of the 35 kb gene cluster isolated from the clone positive for EPA/DHA production had homology to the published pfa genes required for EPA/DHA production. 99 4.2 Introduction The therapeutic effects of omega-3 fatty acids such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) have been well documented (La Guardia et al, 2005; Siddiqui et al, 2008a; Siddiqui et al, 2008b). As a result, markets for EPA and DHA have increased in the areas of health supplements and food enrichment. The use of fish and fish oils, the main dietary sources of EPA and DHA, as a food additive or food supplement have some drawbacks due to odor, taste and stability problems. Marine algae, the primary EPA/DHA producers, and marine bacteria are capable of producing EPA and DHA. The use of microalgae can resolve the odor and taste issues associated with fish oil. However, commercial production of EPA/DHA from microalgae on a large scale is reported to be costly (Barclay et al, 1998; Barclay et al, 1994; Chi et al, 2008; Sijtsma & de Swaaf, 2004; Wen & Chen, 2003) Therefore, EPA/DHA extracted from microalgae is currently applied mainly in infant formula and pharmaceutical products (Sijtsma & de Swaaf, 2004). Bacterial production of EPA/DHA is not suitable due to low accumulation of triacylglycerols and the existence of some unusual fatty acids that are not found in other organisms (Ratledge, 2004). Cloning of genes involved in the biosynthesis of EPA/DHA from marine bacteria will enable us to use it as a tool for production of EPA/DHA in other beneficial microorganism such as lactic acid bacteria or in higher organism such as plants. A marine bacterium isolated from mackerel entrails was identified as Shewanella baltica MAC1, and is able to produce EPA and DHA. Recently, we were able to clone and express the EPA/DHA biosynthesis genes from S. baltica MAC1, in E. coli using the Bacterial Artificial Chromosome (BAC). Although production of EPA and DHA was detected, the yield of EPA (0.1%) and DHA (0.8%) 100 was low and we were also unable to excise the 200kb fragment containing all the genes responsible for biosynthesis of EPA and DHA from the BAC plasmid. Construction of a BAC library with large DNA fragments increases the chance of chimerism, causing a rearrangement of the sequence, which makes it difficult to excise the insert. Constructing a genomic DNA library with smaller inserts (35-40kb) decreases the risk of getting chimeric clones. A fosmid cloning vector is able to hold an insert up to 40 kb. Cloning genomic DNA fragments (-40 kb) using fosmid vectors has been widely reported (Ammiraju et al, 2005; Beja et al, 2002; Huang et al, 2008; Smailus et al, 2007). Fosmid clones are large enough to carry the entire (37kb) EPA/DHA gene cluster but still small enough to be sequenced and manipulated easily. The fosmid vector is derived from single-copy F-factor of E. coli containing the cos site for in vitro lambda packaging. Figure 4.1 shows a diagram of the fosmid vector. It contains chloramphenicol resistance as an antibiotic selectable marker, E. coli F-factor based partitioning and single copy origin of replication, inducible high copy origin of replication (oriV) and bacteriophage lambda cos site for lambda packaging. Fosmid cloning utilizes high efficiency lambda packaging that eliminates background and false positives (Smailus et al, 2007). A fosmid library usually consists of a large number of clones. An appropriate and rapid detection method is necessary to screen a fosmid library. Partial amplification of each EPA/DHA gene and designing specific probes can be helpful too. 101 EeciFZl te# COS favZ. pafC pro CQp^QomtOF* ffl£F' pC€1FOSwancl pCC2FOS •• vectors 1,1 Kb m0 Figure 4.1 Diagram of a Fosmid Cloning Vector. (EpiCentre, Madison, WI, U.S.A) 102 In one study, a fosmid libray of Photobacterium profundum SS9, an EPA- producing bacterium, was constructed and screened using a partial fragment of SS9 pfaA and pfaD (Allen & Bartlett, 2002). However, no recombinant EPA/DHA production was obtained when they analyzed the clone which hybridized both probes (Allen & Bartlett, 2002). In this study, each pfa (pfaA, pfaB, pfaC, pfaD, pfaE) gene of 5. baltica MAC1 responsible for EPA/DHA production was amplified partially in order to make probes for rapid detection of positive clones in a fosmid library. A fosmid library was constructed from sheared S. baltica MAC1 genomic DNA. The fosmid library was tested for the presence of EPA/DHA biosynthesis genes by colony hybridization. Clones positive for the EPA/DHA genes were screened for recombinant production of EPA/DHA. All positive clones for recombinant production of EPA/DHA were end-sequenced. The complete sequence of the insert for the clone positive for both EPA and DHA was obtained and studied further. 4.3 Material and Methods 4.3.1 Amplification of genes responsible for EPA/DHA production 4.3.1.1 Bacterial strains and growth conditions Stock cultures of S. baltica MAC1 were maintained frozen at -80°C in 15% glycerol. Bacterial cultures for use in experiments were prepared by inoculation of frozen stock cultures onto Marine Agar (MA) 2216 (Difco, Detroit, MI, U.S.A). Plates with S. 103 baltica MAC1 were incubated at 30°C overnight. An isolated colony from an overnight culture was inoculated in 50 ml tubes containing 5 ml Marine Broth (MB) 2216 (Difco) and incubated at 30°C overnight while shaking at 70 rpm on a New Brunswick CI platform shaker (New Brunswick Scientific, Edison, NJ, U.S.A). 4.3.1.2 Genomic DNA isolation of S.baltica MAC1 To extract genomic DNA of S. baltica MAC1, one milliliter of each sample (overnight culture) was added to an autoclaved 1.5 ml microcentrifuge tube (Fisher Scientific, Ottawa, ON, Canada). The tubes were centrifuged at 14,000 g for 3 minutes in a BeckMan Coulter Microfuge 22R centrifuge (Beckman Coulter, Lawrence, KS, U.S.A) to pellet the cells. After removing the supernatant, the bacterial pellet was resuspended in 200 nl lx Phosphate Buffered Saline (PBS) (Invitrogen, Burlington, ON, Canada) to eliminate the formation of a salt pellet during precipitation of DNA. Genomic DNA was extracted from the cell suspension with the Easy DNA Kit (Invitrogen) with the following modifications: 600 ^1 lysis solution was added instead of 350 ul, during the phase separation and DNA precipitation, centrifugation was reduced from 20 min to 8 min and the DNA pellet was resuspended in 50 \il instead of 100 u,l. DNA concentration was measured by absorbance at 260nm. A total amount of 115-135 [ig DNA was obtained. 4.3.1.3 Polymerase Chain Reaction (PCR) procedure for EPA/DHA genes amplification To amplify each individual gene responsible for EPA/DHA production, a set of oligonuclotide primers, described in Table 4.1, 104 Table 4.1 primers used to locate EPA/DHA genes in the S.baltica MAC1 genome Primer Sequence(5'-3') Target EPA5-1 Forward, GCTTTAATGCCGACATGGTT pfaA gene Reverse, TTCTCCGCAGAAAGACCATT EPA5-2 Forward, ATCAAGCGCGTTGAAATTCT pfaA gene Reverse, GCGCTATCGGTAACATCCAT EPA5-3 Forward, GTTGGCATGGCGAGTATTTT pfaA gene Reverse, TGCAGTACCTGTTCCGTGAG EPA5-4 Forward, TCACCGGTGGTGTGTGTACT pfaA gene Reverse, GTAGCTGCCATGCGTTATCA pfaA.S Forward, GGTGTSGGYGGTGGTCAR pfaA gene Reverse, CTCACCRAARCTRTGRCC EPA6-1 Forward, GTGTGGAACTTGGCCCTAAA pfaB gene Reverse, GCCGGTGTTTTGTATTGGAT EPA6-2 Forward, CTGAGCTAAATGCCCTGCTT pfaB gene Reverse, TTTTGCGTTGATAGGCACTG pfaB2 Forward, GGTGAAGCATCRATGTGGGC pfaB gene Reverse, TCSGCRCCAATTTCAACAA pfaB6 Forward, GGTGAAGCATCRATGTGGGC pfaB gene Reverse, GTTMCGGAAGAACAGCTC EPA7-1 Forward, AAAGGCGGCTACATTGAGAA pfaC gene Reverse, CTCAATGGCTACAGGTGCAA EPA7-2 Forward, TAGACGGACAAATCCCTTGG pfaC gene Reverse, TAACCGGATCTTGGTGGAAG pfaC3 Forward, TTGATGGTCARATCCCTTGG pfaC gene Reverse, GTTMCGGAAGAACAGCTC pfaC4 Forward, TTGATGGTCARATCCCTTGG pfaC gene Reverse, GTGACTTKACCGTTATCTAGGCC EPA8-1 Forward, TGGCGCAACTTGTACCTATG pfaD gene Reverse, CATCTAGGCTTGAGCGGAAT EPA8-2 Forward, GATTTTAGCCGGGCATGTTA pfaD gene Reverse, AGTTTTACGCCCATCTCGAA EPA2-1 Forward, TATCAGCAGCGTTCATTTGC pfaE gene Reverse, CAGTCAGGCAAGCAAGGAG EPA2-2 Forward, TTGTGATCCGCAATAGCATC pfaE gene Reverse, TGAAATACGCCCCAATGAA EPA2-3 Forward, TGCAATTTTGAACTAGCTAGTCT pfaE gene Reverse, CTCAAGGTTTAATGGTAAGAGGC EPA2-4 Forward, CTAGCTAGTCTTAGCTGAAGCT pfaE gene Reverse, ATGGTAAGAGGCTATTTGCGCG PPTex Forward, GTATCCATTCTACATATGTACAG pfaE gene Reverse, AAATAGTCTCGAGCTTCACTC PPTsh Forward, GGCGATAAAGGYAARCCK pfaE gene Reverse, CAACGHTCRATRTCWCCACC 105 were synthesized by Laboratory Services Division, University of Guelph (Guelph, ON, Canada). Forty five microliters of PCR SuperMix (Invitrogen Life Technologies, Burlington, ON) containing 22 raM Tris-HCl (pH 8.4), 55 mM KC1, 1.65 mM MgCl2, 220 |JM dNTP and 22 U recombinant Taq DNA polymerase/ml, were added to a sterile 250 |il microcentrifuge PCR tube. One microliter of 25 pmol/jil forward primer and l(J.l of 25 pmol/jil reverse primer were added to 45 \i\ SuperMix and then 1 |ig DNA were added to the mixture of SuperMix and the primers. The final solution was gently mixed and PCR was performed in a MasterCycler (Eppendorf Scientific, Westbury, N.Y, U.S.A) under the conditions shown in Table 4.2. To amplify the 37 kb gene cluster (ORF1-9), a series of primers was designed from ORFsl, 2, 8, and 9 (Table 4.3) and were synthesized by Laboratory Services Division, University of Guelph (Guelph, ON, Canada). Three to five microliters of genomic DNA isolated from 5. baltica MAC1 were used for the amplification of a large fragment using Expand 20 kb PCR System (Roche Diagnostics, Quebec, Canada), iProof High-Fidelity DNA polymerase (BioRad, Mississauga, On, Canada) and MasterAmp extra long PCR kits (EpiCentre, Madison, WI, U.S.A). The preparation of the PCR and conditions were performed according to the manufacturers' protocols. 106 Table 4.2 Conditions that were used in PCR Primers Temperature Duration Number of Cycle pfaA 95°C lOmin 1 94°C 1 min 40 EPA5-4 56.4°C 1 min 40 72°C 3 min 40 72°C 10 min 1 pfaB 95°C 10 min 1 EPA6-1 94°C 1 min 45 pfaC3 42°C 1 min 45 pfaC4 72°C 3 min 45 EPA7- 72°C 10 min 1 EPA8-2 EPA2-1 95°C 10 min 1 94°C 1 min 40 46.4°C 1 min 40 72°C 3 min 40 72°C 10 min 1 Vlin: minute, °C: centigrade 107 Table 4.3 Primers used to amplify the 37kb EPA gene cluster in the S.baltica MAC1 genome Primer Sequence (5'-3') Target ORF1-8 Forward, TTATGAAACAGACTCTAATGGCTATC EPA genes Reverse, GTGTATTAGGCCATTCTTTGGTTTG ORF1-9 Forward, ACCCACTCGAGTAAACAACTCTTCTC EPA genes Reverse, GGCTTAATGACCGATTTTATTTGCTT ORF2-8 Forward, CTAGCTAGTCTTAGCTGAAGCTCGA EPA genes Reverse, GTGTATTAGGCCATTCTTTGGTTTG 108 4.3.1.4 Separation of DNA fragments To detect the amplification of each gene by PCR, the standard method used to separate DNA fragments in electrophoresis through agarose gels was used. To perform agarose gel electrophoresis, a slab of 1 % agarose gel was prepared by melting 0.5 g ultra pure agarose powder (Invitrogen) in 50 ml of 1 x TAE buffer. The mixture was boiled in a microwave until a clear solution was observed. The solution was cooled to 40-45°C and, after adding 0.5 |il 10 mg/ml Ethidium Bromide (ETBR) (Sigma, Oakville, ON, Canada), it was poured into a 10 cm tray and solidified at room temperature (23°C). The gel was placed in a gel electrophoresis submarine cell containing 250 ml 1 x TAE buffer. Five microliters of PCR product were mixed with 1 [xl 6x gel loading dye (Fermentase, Burlington, ON, Canada) and loaded in each well then subjected to electrophoresis at 75V for 2 h. To detect the amplification of the 37 kb cluster of genes, a 0.7% agarose gel was prepared in a 15 cm tray and electrophoresis was performed at 55V for 6 h. 4.3.1.5 Purification of amplified DNA fragment from PCR reactions for sequencing The QIAquick PCR purification kit (Qiagen, Mississaga, ON, Canada) was used to purify the amplified product following the manufacturer's protocol. At the end, the DNA retained in the resin column was eluted with 32 (J,l elution buffer (EB). The DNA concentration of each sample was determined by absorbance at 260nm. The total amount of purified DNA was 0.7-3 (ig. Purified PCR products were sequenced at the Guelph Molecular Supercentre, Guelph, ON, Canada. 109 4.3.2 Construction of Fosmid library from 5. baltica MAC1 genome 4.3.2.1 Bacterial strains and growth conditions S. baltica MAC1 was prepared as described in section 4.2.1.1. The EPI300-T1 E. coli strain was supplied as a glycerol stock (Epicentre, Madison, WI, U.S.A). Prior to beginning the fosmid library construction, EPI300-T1 was streaked out on LB plates without antibiotic and incubated at 37°C overnight. Thereafter, plates were stored at 4°C. The day before packaging the fosmid clones, a single colony of EPI300-T1 was cultured in 50 ml LB broth +10 raM MgSCu and incubated at 37°C overnight while shaking at 200 rpm on a New Brunswick Scientific C24 incubator shaker (New Brunswick Scientific). On the day of packaging, 50ml of LB broth + 10 mM MgS04 were inoculated with 5 ml of the EPI300-T1 overnight culture and incubated at 37°C while shaking at 200 rpm to an OD600 = 0.9. The cells were stored at 4°C until use (the maximum storage time at4°Cis72h). 4.3.2.2 Genomic DNA isolation from S. baltica MAC1 A 50 ml tube containing 5 ml Marine Broth (MB) was inoculated with a single colony of an overnight culture of S. baltica MAC1. The culture was incubated at 30°C overnight while shaking at 70 rpm. Cells of 3 ml culture were harvested and resuspended in 400 |xl IX PBS. Genomic DNA was extracted using the Easy-DNA kit (Invitrogen) according to the manufacturer's protocol with the modification described in section 4.2.1.2. Isolated genomic DNA from 5. baltica MAC1 was stored at 4°C. 110 4.3.2.3 Shearing S.baltica MAC1 genomic DNA and sizing the fragmented DNA by pulse field gel electrophoresis Genomic DNA isolated from S. baltica MAC1 was sheared mechanically to approximately 40 kb fragments. To shear the DNA, different concentrations of DNA (5, 10, 15 or 20 fig) were passed through a 200 ul small bore pipette tip (Fisher) for 60 times. The fragmented DNA was separated in a 1% agarose gel prepared with lx TAE buffer. The gel was placed on the gel holding template in the electrophoresis chamber and covered with 2.2 L lx TAE buffer. Five microliters of fragmented DNA were mixed with 1 ^1 6x gel loading dye and loaded in the gel. The CHEF-DR III pulse field gel electrophoresis (PFGE) (BioRad, Mississauga, ON, Canada) was set as follows: initial switch time 2.2 sec, final switch time 54.2 sec, run time 15 h, voltage 6V, include angle 120, running temperature 14°C. The gel was stained in 500 ml distilled water contained 40 (xl lOmg/ml ethidium bromide (Sigma, Oakville, ON, Canada) for 45 min and then destained for 25 min in 500 ml distilled water. The size of the fragmented DNA was also observed on a 1% agarose gel prepared with lx TAE by conventional gel electrophoresis at 40V for 14 h. The gel was observed in a Bio-Rad Gel Doc system (Bio-Rad). 4.3.2.4 End-repair of the sheared DNA to blunt 5'-phosphorylated ends After obtaining DNA fragments with the approximate size of 40 kb, the two ends of DNA fragments were blunt ended. Twenty two micrograms of sheared DNA were transferred to sterile 1.5 ml microcentrifuge tubes and were end repaired as follows: 2 |xl water + 8 jxl lOx end-repair buffer + 8[il 2.5 raM dNTP mix + 8 \il 10 mM ATP + 22 [xg sheared DNA + 4u.l end-repair enzyme (total volume of 80 \il) were mixed together and ill incubated at room temperature for 50 min. To inactivate the end repair enzyme, 16 ul 6x gel loading dye buffer were added to the reaction mixture, which was then incubated at 70°C for 12 min. Repaired DNA was purified and concentrated as follows: 9 \il of 3 M sodium acetate (Sigma, Oakville, On, Canada) were added to the 96 ul reaction mix and mixed by gentle inversion; 250 \i\ absolute ethanol (Fisher) were added and the tube was inverted very gently. The DNA was allowed to precipitate at 4°C for 2 h; DNA was collected by centrifugation in a Beckman Coulter microcentrifuge (Beckman Coulter) for 20 min at 9000 g. The supernatant was aspirated from the DNA pellet carefully and the DNA was washed with 70% cold ethanol two times. Finally, the DNA was air-dried for 3 min and then resuspended in 30 |xl TE buffer (Sigma). The size of the purified, concentrated, end-repaired DNA was determined as set out in the following section. 4.3.2.5 Sizing the end-repair DNA in gel electrophoresis A 15 cm 1% agarose gel was prepared by melting 1.5 g ultra pure agarose powder in 150 ml of lx TAE buffer. The mixture was boiled in a microwave until a clear solution was observed. The solution was cooled to 40-45°C and 0.9 (0.1 10 mg/ml ethidium bromide were added. The cooled solution was poured in a 15 cm tray and solidified at room temperature (23°C). The gel was placed in a gel electrophoresis submarine cell containing 1500 ml lx TAE buffer. Five microliters of purified, concentrated, end-repaired DNA were mixed with l(xl of 6x gel loading dye and loaded in the gel's wells. The electrophoresis was performed for 17 h at 40V (2.5 volts/cm). 112 4.3.2.6 Ligation of DNA fragments into the pCClFOS fosmid vector The concentrated, purified, end-repaired 40 kb DNA fragments and 36 kb fosmid control DNA (as a control) were used to construct a fosmid (FOS) library. Linearized, dephosphorylated pCClFOS cloning-ready vector (EpiCentre) was used as the cloning vector. In two separate tubes (one used as a control and the other one for sample), 1 \il lOx fast-link ligation buffer (EpiCentre) + 1 ul 10 mM ATP (Sigma) + l^xl (0.5 fig) pCClFOS + 6 |o,l (0.25 ng) concentrated insert DNA + 1 nl fast-link DNA ligase (EpiCentre), were mixed and incubated at room temperature for 2.5 h. The DNA ligase was inactivated at 70°C for 12 min. 4.3.2.7 Packaging the ligated DNA and infecting EPI300-77 E.coli A tube of 50 u.1 MaxPlax lambda packaging extract (EpiCentre) that had previously been stored at -80°C in a Thermo scientific freezer (Fisher) was thawed on ice. It was divided into two 1.5 ml microfuge tubes. One-half of the packaging extract (25 [xl) was kept on ice and the second half was transferred to the -80°C freezer for use in the next step. The 10 \x\ of ligation reaction mixture described in section 4.2.2.5 were mixed with 25 \i\ of lambda packaging extract by pipetting the solutions several times and this packaging reaction was then incubated at 30°C in an ISOTemp water bath (Fisher Scientific Ltd) for 2 h. After 2 h, the remaining 25 |xl of MaxPlax lambda packaging extract were thawed on ice and added to the packaging reaction. The packaging reaction was incubated for an additional 2 h at 30°C in a water bath. At the end of the second 2 h incubation period, 950 fil phage dilution buffer (PDB) (EpiCentre) were added to the tube containing phage particles (packaged fosmid clones) and mixed by gentle inversion. 113 Twenty five microliters of chloroform were added to the tube, the solution was mixed gently and was stored at 4°C. Varying volumes (2, 5, 8, 10, 15, and 20 \i\) of packaged phage particles were added, individually, to 100 ul of the prepared EPI300-T1 host cells, which were prepared as described in section 4.2.2.1. Each tube containing the different amount of phage particles and host cells was incubated at 37°C in a water bath for 22 min. Infected cells were spread on LB plates supplemented with chloramphenicol (12.5 (ig/ml; Sigma) and incubated at 37°C overnight to select for the FOS clones. 4.3.2.8 Sizing the fosmid clones and storage of the fosmid clones Plates containing 200 to 250 FOS clones were selected. Six FOS clones were randomly chosen from 6 plates and streaked on a fresh LB agar supplemented with chloramphenicol (12.5 mg/ml). Plates were incubated at 37°C overnight. Six 50 ml tubes, each tube containing 5ml LB broth + 5 ^.1 chloramphenicol (12.5 ng/ml) + 10 [xl 500X fosmid autoinduction solution, were inoculated with a single colony of each overnight culture. The tubes were incubated at 37°C for 16 h while shaking at 230 rpm. FOS DNA was extracted from 3 ml overnight cultures using FosmidMax DNA purification kit (InterScience, Markham, ON, Canada). The manufacturer's protocol was followed for the FOS DNA extraction. The FOS DNA was stored at 4°C. To determine the insert size, 2.5 \il (l[xg) of each sample + 2 \i\ buffer H + 1 ^1 restriction enzyme (Notl) + 15 \i\ water were mixed together and digested in a 37°C water bath overnight (21h). In addition, 2.5 ul of the fosmid control DNA were digested as described above. Digested and undigested FOS DNA isolated from 6 clones and 114 Fosmid control DNA were separated in a 1% agarose gel prepared with lx TAE as described in section 4.2.2.4. The presence of the 40 kb insert size was confirmed in the randomly selected clones. The packaged DNA (phage particles) as a primary library and infected cells with the phage particles (packaged fosmid clones) were stored for further use as follows: 200 ui of sterile glycerol were added to the tube containing 800 ul phage particles (glycerol was added to a final concentration of 20%), mixed and stored at -80°C in a freezer and used as a primary library. To each plate containing FOS clones, 2 ml LB broth were added and all the clones from each plate were resuspended in the LB broth using a sterile glass rod. The resuspended cells were transferred to 2 ml Corning tubes (Fisher Scientific Ltd) and sterile glycerol was added to each tube to a final concentration of 20%, mixed and stored at -80°C in a freezer. 4.3.3. Screening of the fosmid library by colony hybridization 4.3.3.1 Polymerase Chain Reaction (PCR) procedure to amplify pfaA (EPA5) and pfaD (EPA8) genes Genomic DNA of S. baltica MAC1 was isolated as described in section 4.2.1.2 and subjected to PCR, in order to amplify pfaA and pfaD genes, as described in section 4.2.1.3. PCR reactions were purified using a QIAquick PCR purification kit (Qiagen). The purification was done according to the manufacturer's protocol. The concentration of the purified PCR products at 260nm was 1.4-1.7 \ig. The PCR product stored at -20°C for further use. 115 4.3.3.2 Labeling the purified PCR products (probe) The purified PCR products were used as probes. The two probes were labeled using digoxigen (Dig) (Roche Diagnostics, Quebec, Canada), a steroid hapten. To label the two probes, 1 u.g (16 \il) DNA (probe) was transferred to a 250 ul PCR tube and the DNA was denatured in a MasterCycler PCR machine (Eppendorf Scientific) for 13 min at 95°C and placed in ice immediately after denaturation. To the denatured DNA, 4 u.1 Dig were added, mixed thoroughly, centrifuged briefly and incubated at 37°C in a water bath overnight (20 h). The reaction was stopped by heating to 65°C for 12 min. The labeled DNA was stored at -20°C until further use. 4.3.3.3 Determination of labeling efficiency The yield of Dig-labeled DNA is important for optimal and reproducible hybridization results. To determine the efficiency of the Dig-labeled DNA, a series of dilutions of Dig-labeled probes pfaA, pfaD and a Dig-labeled control DNA (Roche Diagnostics) was prepared. One microliter of the labeled DNA was added to 9 \il DNA dilution buffer (TE buffer), dilution was continued up to 10"8. One microliter of the labeled probes or the labeled DNA control was applied to the nylon membrane (Roche Diagnostics). The labeled DNA was fixed to the membrane by cross linking with UV- light for 1 min for each side using a Transilluminator FBTIV-88 (Fisher Scientific Ltd) and then was subjected to chemiluminescent detection. 116 4.3.3.4 Chemiluminescent detection with CSPD The membranes with fixed Dig-labeled probes or Dig-labeled control DNA were transferred into a plastic container with 20 ml maleic acid buffer (Appendix A) and incubated at room temperature for 5 min under shaking. The membrane was transferred to 10 ml blocking solution (Roche Diagnostics) and was incubated at room temperature for 1 h; following which 1 \il antibody solution (Roche Diagnostics) was added to the 10 ml blocking solution and the membrane was incubated at room temperature for 30 min. The membrane was washed twice for 15 min with 10 ml washing buffer (Appendix A) and then equilibrated in 10 ml detection buffer (Appendix A) for 5 min. After the washing steps, a 30 x 30 cm sheet of plastic wrap was placed on the lab bench and 4 drops of CSPD-ready-to-use (Roche Diagnostics) was loaded onto the plastic wrap. The membrane was placed on the CSPD drops with the DNA side face down on the plastic to spread the substrate evenly. The membrane was covered with a second sheet of plastic so that it was sandwiched between the plastic wraps. After 5 min incubation at room temperature, the excess amount of CSPD was squeezed out and the edges of the plastic were sealed. In a dark room with red light, the membrane was placed in an autoradiography cassette (Fisher Scientific Ltd), which was exposed to a Kodak BioMax XAR film (Sigma) for 10 min. The film was transferred into 500 ml developer (Sigma) for 5 min and then it was washed with water for 1 min. After this washing step, the film was transferred to the 500 ml fixer (Sigma) and the images were fixed for 3 min. The film was washed with water for 1 min in the final step and, after this step, the images of the labeled DNA on the film were visualized. 117 4.3.3.5 Colony hybridization procedure 4.3.3.5.1 Colony lift, DNA transfer and fixation To screen the FOS library, the stock tube containing FOS library that had been stored at -80°C was thawed on ice and in each set of experiments, 80 [xl of infected cells were mixed with 120 ul of LB broth. The suspension was divided into two equal volumes and each was spread onto two LB agar plates supplemented with chloramphenicol (12.5 mg/ml). The plates were incubated at 37°C overnight. Approximately 250 isolated clones were seen on each plate. A copy of each plate was made on another LB plate + chloramphenicol (12.5 u.g/ml) using a velveteen square (Fisher Scientific). Both plates, the original and the copy, were incubated at 37°C for 4-6 h and overnight, respectively. Plates were pre-cooled after 4-6 h incubation at 4°C for 45 min. To transfer colonies from LB plates to a nylon membrane (Roche, Quebec, Canada), a circular nylon membrane (82 mm) was placed on top of the clones in the original plate (the orientation was marked on both plates and membranes in order to be able to identify positive clones). When the nylon membrane was attached onto the surface of the clones, the membrane was left for 2 min. The nylon membrane was removed very carefully with filter tweezers (BioRad), placed (colonies upside) on fresh LB agar supplemented with chloramphenicol (12.5 (xg/ml) and then incubated at 37°C overnight (22 h). The membrane with grown colonies was lifted with tweezers from the LB plate and blotted on a Whatman 3 MM paper (Fisher Scientific). The membrane was placed upside on Whatman paper soaked with denaturation solution (Appendix B) for 15 min. The membrane was air-dried for 30 sec and was placed onto the Whatman paper soaked with neutralization solution (Appendix B) for 15 min. The membrane was finally transferred onto a Whatman paper soaked with 118 2x SSC (Sodium Chloride and Sodium Citrate) (Sigma) for 2 min. The DNA on the membranes was cross linked with UV-light for 1 min and was treated with 2 ml proteinase K solution (Roche Diagnostics) for lh at 37°C. Cell debris was washed off the membrane by placing a pre-wet Whatman paper with water on top of the membrane and pressing firmly across the paper using a ruler. Cell-debris was removed by gently pulling off the Whatman paper. This step was repeated 2 to 3 times until all debris came off the nylon membrane. 4.3.3.5.2 Hybridization, washing steps and detection Prior to hybridization, 20 ml Dig Easy Hyb (Roche Diagnostics) for addition to each membrane was pre-heated to 42°C in a Thermo Scientific Hybaid MAXi 14 hybridization oven (Fisher Scientific). The membrane was placed in a hybridization tube and 20 ml of the preheated Dig Easy Hyb (pre-hybridization solution) were transferred to the hybridization tube for each membrane. The membrane was pre-hybridized in the hybridization oven for 6 h at 42°C with gentle agitation. Labeled probes were thawed on ice and denatured at 95°C for 12 min in a MasterCycler PCR machine. Tubes were put in ice immediately. Sixteen microliters of each denatured labeled probe were transferred to 20 ml pre-heated (42°C) Dig Easy Hyb and mixed gently (probe/hybridization mixture). After 6 h, the prehybridization solution was poured off and 20 ml of the probe/ hybridization mixture were added to the membrane. It was incubated in the hybridization oven overnight with gentle agitation. In total, hybridization with two labeled probes (pfaA, pfaD) was carried out for each of the 7 individual membranes containing FOS clone DNA in 7 independent experiments. After hybridization, each membrane was 119 washed twice for 10 min in 100 ml 2x SSC and 0.1% SDS under constant agitation, it was followed by another washing sequence (2 x 30 min in 0.5x SSC and 0.1% SDS at 68°C under constant agitation). For chemiluminescent detection, membranes were incubated at room temperature with a gentle agitation in 20 ml washing buffer for 5 min, 100 ml blocking solution for 2 h, 20 ml antibody solution for 30 min and 100 ml washing buffer 2x15 min and, finally, it was equilibrated in 20 ml detection buffer for 5 min. The chemiluminescent detection with CSPD was performed as described in section 4.2.3.4. 4.3.4 Analysis of fosmid clones positive for pfaA and/or pfaD genes 4.3.4.1 Bacterial strains and growth conditions Thirty-one FOS clones selected on the basis that their DNA hybridized to either probe pfaA or pfaD were streaked on fresh LB agar supplemented with chloramphenicol (12.5 fig/ml) and incubated at 37°C overnight. Thirty-one 50 ml tubes, each tube containing 5ml LB broth + 5(xl chloramphenicol (12.5 [ig/ml) + 10 \il 500x fosmid autoinduction solution were inoculated with a single colony of each overnight culture. Tubes were incubated at 37°C for 16 h while shaking at 230 rpm. Two 50 ml tubes containing either LB broth or Marine broth (MB), were inoculated with an overnight single colony of EPI 300-T1 E. coli (negative control) as described in section 4.2.1.1 and S. baltica MAC1 (positive control), respectively. The tube with EPI 300-T1 E. coli was incubated at 37°C overnight while shaking at 200 rpm and the one with S. baltica MAC1 was incubated at 30°C overnight while shaking at 70 rpm. 120 4.3.4.2 DNA isolation and PCR for pfaA and pfaD genes Cells from 3 ml cultures were harvested in a Beckman microcentrifuge at 14,000 g. The cells of S. baltica MAC1 and EPI300-T1 E. coli were resuspended in PBS (Invitrogen) and their genomic DNA was extracted as described in section 4.2.1.2. The cells of FOS clones were resuspended in buffer PI provided by the QIAprep Miniprep Qiagen large plasmid isolation kit (Qiagen). FOS DNA was isolated according to the manufacturer's protocol. In order to confirm the presence of the pfaA and pfaD genes in FOS clones, isolated DNA from FOS clones, along with negative and positive controls, was subjected to PCR, as described in section 4.2.1.3. Five microliters of PCR product were mixed with 1 of 6x gel loading dye, loaded in 1% lx TAE and subjected to electrophoresis at 90V for 2.5 h. 4.3.4.3 Digestion of FOS DNA positive for both pfaA and pfaD genes To determine the insert size of 9 positive clones for both pfaA and pfaD, 3 \il DNA of each sample + 2 \il buffer H + 1 of Notl + 14 \il water were mixed together and digested in a 37°C water bath overnight (21h). In addition, 2 jxl fosmid control DNA was digested as described above. Digested and undigested FOS DNA isolated from the 9 clones and Fosmid control DNA were separated in a 1% agarose gel prepared with lxTAE as described in section 4.2.2.4. 121 4.3.4.4 End sequence of fosmid DNA isolated from positive clones Isolated FOS DNA from positive samples for both pfaA and pfaD were sent to the Laboratory Service Division, Guelph Molecular Supercentre (Guelph, ON, Canada) for end sequencing. 4.3.5 Detection of EPA/DHA production in positive FOS clones 4.3.5.1 Bacterial strains and growth conditions The positive FOS clones for pfaA (EPA5) and pfaD (EPA8) were tested for EPA and/or DHA production. A FOS clone negative for pfaA and pfaD and EPI300-T1 E.coli were used as negative controls and S. baltica MAC1 was used as a positive control. Inocula of FOS clones and negative controls were prepared by culturing an overnight single colony of each isolate into 5 ml of LB broth supplemented with 5 of chloramphenicol (12.5 ng/ml) and LB broth (for EPI300-T1), respectively. Cultures were incubated at 37°C overnight while shaking at 200 rpm. Five milliliter inocula of each clone and EPI300T1 were transferred to 1000 ml Erlenmeyer flasks containing 200 ml LB broth supplemented with 200 pil chloramphenicol (12.5 mg/ml) and 200 ml LB broth, respectively. All cultures were incubated at 10°C (8 days), 15°C (5 days), 20°C (3 days) and 25°C (1 day) on a shaker (200 rpm). In addition, a single colony of S. baltica MAC1 was cultured in 5 ml MB at 30°C overnight. Five milliliter inocula of S. baltica MAC1 were transferred to 1000 ml Erlenmeyer flasks containing 200 ml MB and incubated at 4°C (5 days) and 10°C (3 days). 122 The cells were transferred to 250 ml sterile plastic bottles and then were harvested by centrifugation using a Beckman J2-MC centrifuge at 9000 g for 17 min. 4.3.5.2 Fatty acid extraction and GC analysis The harvested cells were resuspended in 200 ml ultra pure water, vortexed for 30 seconds and then centrifuged under the same condition described in section 4.2.5.1. The cells were freeze-dried overnight in a Stokes Freeze-Drier (Equipment Division Pennsalt Chemicals Crop, Philadelphia, PA, U.S.A) and ground to a fine powder in a clean sterile mortar with a pestle. The powders of each sample were weighed and were transferred to 20 ml screw cap test tubes. Fatty acid was extracted and GC analysis was done as described in sections 2.3.9 and 3.3.7.3. 4.3.5.4. Gas Chromatographic-Mass Spectrometry Analysis Samples which were positive for EPA/DHA production were sent to the Chemistry Department, University of Guelph (Guelph, ON, Canada) for GC-mass spectrometry. 4.3.7 Complete sequence of the insert containing EPA/DHA genes 4.3.7.1 Bacterial strains and growth conditions A clone, which was confirmed to produce EPA and DHA by GC and GC-MS, was streaked on LB agar supplemented with chloramphenicol (12.5 [xg/ml) and incubated at 37°C overnight. Five milliliters of LB broth supplemented with 5 jxl chloramphenicol 123 (12.5 ng/ml) and 10 nl of 500 x fosmid autoinduction solution were inoculated with a single colony and incubated at 37°C for 20 h. 4.3.7.2 Fosmid DNA isolation from a positive clone Three milliliter of cells were harvested using a Beckman microcentrifuge and FOS DNA was extracted using a QIAprep Miniprep large plasmid isolation kit (Qiagen). FOS DNA was isolated according to the manufacturer's protocol. DNA in the column was eluted with 35[xl of elution buffer at 70°C. 4.3.7.3 Sequencing of the insert Fifty micrograms of FOS DNA from the clone positive for EPA and DHA production were sent to Laboratory Service Division, Guelph Molecular Supercentre (Guelph, On, Canada) for complete sequencing of the insert. 4.4 Results and Discussions 4.4.1 Characterization of S. baltica MAC1 genes homologous to the pfaA, pfaB, pfaC and pfaD synthase gene cluster Omega-3 fatty acid genes in the EPA/DHA producing bacterium, S. baltica MAC1 may have a similar gene organization with other EPA/DHA-producing bacteria. PCR was carried out using S. baltica MAC1 as a template with 21 primer sets. The primers were designed using available EPA/DHA gene cluster sequences from the GeneBank database from Moritella marina MP-1 (accession # AB025342), 124 Photobacteriwn profundum SS9 (AF409100) and Shewanella pneumatophori SCRC- 2738 (U73935). We designed a different set of primers from conserved and non- conserved regions using the Primer3 Output program. For the pfaA gene, a set of primers targeted the conserved 3-ketoacyl synthase domain. Among all primers for pfaA described in Table 4.1, only primer pfaA.S and EPA5-4 gave a band at size 1800bp (Figure 4.2A). The forward and reverse sequence of the pfaA gene (1800bp) was obtained and compared to the GenBank nucleotide database using an online BLAST search. It gave the highest homology of 79% to S. pneumatophori SCRC-2738 EPA synthesis gene cluster (Appendix C); a homology of 77% to Shewanella livingstonensis gene for P-ketoacyl synthase (KS) and Photobacterium profundum SS9 EPA synthesis gene; 75% homology to Shewanella sp. GA-22 partial pfaA gene for omega-3 fatty acid and 70% homology to Moritella marina MP-1 DHA synthesis gene and Pseudoalteromonas sp DS-12 EPA synthesis genes. For pfaB (EPA 6), primers pfaB2 and pfaB6, described in Table 4.1 were adapted from those described by Gentile et al., which were designed from the conserved area of the acyl CoA:ACP transacylase (AAT) domain (Gentile et al, 2003). The EPA 6-1 and EPA 6-2 primer sets were designed by the Primer3 output program using the sequence of the EPA gene cluster from S. pneumatophori SCRC 2738 to amplify the complete pfaB gene (2kb) or a small part of the gene (500bp). Figure 4.2B shows the PCR results on 1% agarose gel when forward and reverse EPA 6-1 primers were used for S. baltica MAC1. Only this primer gave a strong fragment with a size of ~ 500bp while other primers did 125 A 'ifttStS t 15' •fesssatf f f F ' 1$ S*^:'?f ¥ ;:« ft** M 1 2 3 4 5 /• ,.* S***^w. t fuses' Tnirjr™"1 '•W C D Fngmir© 4o2 Gel electrophoresis (1%) of PCR products using primers pfaA.S, -4, EPA6-1, EPA7-2, EPA8-2, EPA2-1, pfaC3 and pfaC4. First lane in all gels is (New England Biolabs). A: lane 1 and 2 are s of pfaA.S pri product of 1, lanes 5 and 6 product of EPA7-2, lanes 7 product of EPA8' Cs lanes 4 and 5 a products of EPA2-L D: lane 126 not amplify any part of the pfaB gene in the genome of S. baltica MAC1. The BLAST search using the sequence of the 500bp DNA fragment showed 96% homology to S. baltica strains. These data suggest that either the sequence of the pfaB gene in S. baltica MAC1 is completely different from the pfaB genes of other EPA/DHA- producing bacteria or the amplified DNA is from the non-conserved area. It is also reported by other researchers that some primers for pfaB genes used with genomic DNA of Shewanella sp. GA-22 and the proteobacterium Oleispira antarctica were not able to amplify the pfaB gene or the amplified segment(s) did not show any specific homology with the EPA/DHA biosynthesis gene cluster (Gentile et ai, 2003; Yakimov et ai, 2003). Figure 4.2 shows the PCR result on 1% agarose gel when forward and reverse pfaC3, pfaC4 and EPA7-2 primer sets were used to amplify the pfaC gene in the genomic DNA of S. baltica MAC1. PfaC3 and pfaC4 forward and reverse primer sets were synthesized from the published sequence (Gentile et ai, 2003). These two primers were designed from the conserved area of the gene to target the 3-hydroxydecaoyl-ACP dehydratase (HD) domain in the pfaC gene. EPA7-2 was designed using Primer3 output program targeting 2 kb of the pfaC gene close to the end of the ORF 7. Forward and reverse pfaC3 primers gave a strong single band with a size of 1700 bp (Figure 4.2D), pfaC4 gave a strong single band with a size of 1900 bp (Figure 4.2D) and EPA7-2 primer gave a very clear strong band with a size of 2100bp (Figure 4.2B). The BLAST search of the sequence for these DNA fragments gave the highest homology of 77% to S. pneumatophori SCRC-2738 EPA synthesis gene cluster (Appendix C); a homology of 74% to Photobacterium profundum SS9 EPA synthesis gene; Shewanella sp. GA-22 127 partial pfaC gene for omega-3 fatty acid; 70% homology to Moritella marina MP-1 DHA synthesis gene and 68% homology to Pseudoalteromonas sp. DS-12 EPA synthesis genes. The amplification of the pfaC gene from S. sp. GA-22 and Oleispira antarctica has been reported using the pfaC3 and pfaC4 primer sets (Gentile et ai, 2003; Yakimov etai, 2003). For gene pfaD (EPA 8), forward and reverse primers were designed using the Primer3 output program to amplify the complete pfaD gene (1.6 kb) and conserved enoyl reductase (ER) domain. PCR, using the EPA 8-2 primer set and S.baltica MAC1 genomic DNA as template, gave a strong band with a size of 1100 bp (Figure 4.2B). The BLAST search of the sequence for this fragment gave the highest homology of 77% to S. pneumatophori SCRC-2738 EPA synthesis gene cluster (Appendix C); a homology of 76% to Photobacterium profundum SS9 EPA synthesis gene and 66% homology to Moritella marina MP-1 DHA synthesis gene. Figure 4.2C shows the PCR results when forward and reverse pfaE (EPA2) primers were used to amplify the pfaE gene in the genomic DNA of S. baltica MAC1. A set of oligonucleotide primers were used as described in Table 4.1 to target phosphopantetheinyl transferase (PPTase) gene (pfaE) in 5. baltica MAC1. Primers PPTex and PPTsh were adapted from published articles (Allen & Bartlett, 2002; Orikasa et ai, 2006a). EPA2-1, EPA2-2, EPA2-3 and EPA2-4 were designed using the Primer3 output program. Among all these primers, only primer EPA2-1 gave a strong band with a size of 650bp (Figure 4.2C). The BLAST search of the sequence for the PCR product 128 using EPA2-1 gave the highest homology of 96% to S. baltica strains complete genome (Appendix C). EPA2-1 was designed using the sequence of PPTase gene (ORF2) of S. pneumatophori SCRC-2738 (833 bp). No homology was found in an alignment between the ORF2 of 5. pneumatophori SCRC-2738 and the obtained sequence using the primer set EPA2-1. Allen and Bartlett (2002) used template DNA isolated from a variety of EPA-producing bacteria to amplify the PPTase gene using the sequence of ORF 2 of SCRC-2738 to design PCR primers, however, only DNA from Shewanella sp. strain SC2A yielded an amplification product with 55% similarity to ORF2 of the SCRC-2738. Recently, the complete PPTase gene was cloned from Photobacterium profundum SS9, an EPA-producing bacterium (Orikasa et al, 2006b; Sugihara et al, 2008). According to these reports, there are two groups of PPTase gene in this bacterium; one PPTase is mainly used for omega-3 synthesis while the other is involved in polyketide and non- ribosomal peptide synthesis. When this PPTase gene was coexpressed with pfaA-D derived from Moritella marina MP-1, a recombinant E. coli capable of synthesizing DHA was produced (Orikasa et al., 2006b). However, it is not clear whether the PPTase gene ipfaE) cloned from an EPA-producing bacterium encodes the same protein as the PPTase gene from DHA-producing bacteria. 4.4.2 Amplification of 37 kb EPA gene cluster using thermo stable DNA taq polymerase In order to clone the EPA/DHA gene cluster, a set of forward and reverse primers (Table 4.3) from ORF1-9, ORF1-8 and ORF 2-8 was designed using the sequence of the EPA gene cluster from Shewanella pneumatophori SCRC-2738 (U73935). Genomic 129 DNA of S. baltica MAC1 was used as template. Although several attempts (20 times) were made using different kits to amplify the 38 kb, 26 kb and 24 kb gene cluster corresponding to ORF1-9, ORF1-8 and ORF 2-8, respectively, no PCR fragments were obtained. To our knowledge, there is no report regarding amplification of the complete EPA/DHA gene cluster using PCR methodology. 4.4.3. Cloning of the Shewanella baltica MAC1 EPA/DHA gene cluster by constructing a Fosmid Library A fosmid library was constructed with large DNA inserts (~40kb) from S. baltica MAC1 in order to clone the EPA/DHA biosynthesis gene cluster. Forty kilo-base-pair DNA fragments were obtained when the genomic DNA of 5. baltica MAC1 was sheared mechanically (Figure 4.3A). After end repairing the DNA fragments, a fosmid library of 12,000 clones was obtained. Ten plates, each containing 200-250 clones, were screened by colony hybridization for pfaA and pfaD genes. In total 31 clones were positive for either pfaA or pfaD genes. These clones were subjected to PCR for both pfaA and pfaD genes of which 8 were positive for both genes (Figure 4.3B & C). The FOS DNA of 8 clones were extracted and subjected to digestion by Notl. All FOS DNA were digested with Notl. Four digested FOS DNA gave 3 bands, one strong band with a size of ~ 36 kb, one at 8 kb, which is the vector, and the other at 6 kb (Figure 4.4) and 4 digested FOS DNA gave only two bands, one with a size of ~ 36 kb, which is the size of inserts and the other band is at size 8 kb which is the vector size (Figure 4.4). These 8 FOS clones were designated as FOS clone 2, 3,4, 5, 6,7, 8, and 9. 130 1 2 3 4 M mmmmm iWPP wmmmmm 1 36kb fa.f Wi<^W^.^;<."fti<>ifcj*t.aflKlAaau«rttt M123456789 10 1112M M123456789 10 1112M 2000bp 1200bp B Figure 4.3. Gel electrophoresis (1%) of mechanically sheared genomic DNA of S. baltica MAC1 (A) and PCR products using pfaA and pfaD primers (B, C). A: Lanes 1, 2, 3 and 4 are sheared genomic DNA, lane M is 36kb fosmid marker DNA (EpiCentre). B: Lanes M are GeneRuler lOObp DNA ladder (New England Biolabs), Lane 1, 2, 3, 4, 5, 6, 7, 8, and 9 are PCR products of positive clones for the pfaA gene; lanes 10 & 11 are PCR products of S.baltica MAC1 for pfaA gene (as positive control); lane 12 is PCR product of E.coli for pfaA gene (as negative control). C: Lanes M are GeneRuler lOObp DNA ladder (New England Biolabs), Lane 1, 2, 3, 4, 5, 6, 7, 8, and 9 are PCR products of positive clones for pfaD gene; lanes 10 & 11 are PCR products of S. baltica MAC1 for pfaD gene (as positive control); lane 12 is PCR product of E. coli for pfaD gene (as negative control). 131 C DC U2 D2 U3 D3 U4 D4 U5 D5 U6 D6 U7 D7 U8 D8 U9 D9 Figure 4.4 Gel electrophoresis (1%) of undigested and digested FOS DNA with Noil (NewEngland BioLab) isolated from positive clones. Lane C is a 36kb fosmid control DNA (EpiCentre), lane DC is digested fosmid control DNA, lanes U2, U3, U4, U5, U6, U7, U8, U9 are undigested FOS DNA; lanes D2, D3, D4, D5, D6, D7, D8, D9 are digested FOS DNA with Notl. 132 The extracted FOS DNA from these 8 clones was end-sequenced. The sequence of the two ends of the inserts showed 94-99% homology to S. baltica strains. This confirmed the existence of the insert for all the clones. The estimated size of the inserts for each of the FOS clones was determined when the sequence of the two ends were aligned with the complete available sequence of the genome of S. baltica strains. FOS DNA of clone 2 was 40kb, clone 3 (38 kb), clone 4 (38 kb), clone 5 (35 kb), clone 6 (35 kb), clone 7 (41 kb), clone 8 (38 kb) and clone 9 (36 kb). 4.4.5 Detection of recombinant production of EPA/DHA by positive FOS clones for pfaA and pfaD genes Fatty acids from positive FOS clones for both pfaA and pfaD genes, 5. baltica MAC1 (positive control), a FOS clone negative for pfaA and pfaD genes and EPI 300-T1 E. coli (negative controls) were extracted from cells grown at 15°C and analyzed by GC. New peaks appeared in the GC analysis of fatty acids from FOS clones at retention times corresponding to EPA (32.3min) and/or DHA (36.5min). Clone # 6, which was positive for both pfaA and pfaD genes, did not produce any EPA or DHA. Clone # 5 produced EPA and DHA. Clones 2, 3, 4, 7, 8 and 9 produced only EPA. No peaks at retention times of EPA/DHA peaks were observed for the negative controls. The fatty acid profile of clone # 5 is shown in Figure 4.5 as an example. Retention times of the suspected EPA/DHA peaks in the fatty acid chromatograph of clone # 5 are compared with the retention time of an EPA/DHA standard in Figure 4.5. 133 EPA CO DHA JL m B EPA DHA »egj) S to EPA A ^HA Figure 4.5. GC of the fatty acid profile of positive EPA/DHA clone and Standard. A) FOS clone 5. B) FOS clone plus internal standard QEPA/DHA standard. FOS clone 5 was grown at 15°C for 8 days and extracted FAs form cells was analyzed using an Agilent 6890 GC. 134 To verify the EPA/DHA peaks in FOS clones (2, 3, 4, 5, 7, 8, 9), the samples were divided into two portions and 1 (xl EPA/DHA standard with a concentration of 10 mg in 200 ml iso-octane was injected into one portion of the sample. The fatty acid profile of the sample containing the internal standard and the one without internal standard were analyzed under the same conditions. The suspected peaks became larger while other peaks stayed the same or were a bit smaller because they were diluted when they were mixed with EPA/DHA standard. In addition, EPA/DHA production in FOS clones (2, 3, 4, 5, 7, 8, 9) was detected by Gas Chromatographic-Mass Spectrometry (GC-MS). The GC-MS results confirm the production of EPA by all FOS clones. The standard sample showed the EPA and DHA eluting at 11.28 and 13.69 min, respectively. Figure 4.6 shows the chromatogram for the EPA/DHA standard, the mass spectra for the 11.28 and 13.68 minute peaks. Differences in the retention time in the GC and GC-MS was due to the different columns used in each instrument. The GC-MS results for FOS clones (2, 3, 4, 5, 7, 8, 9) gave an intense peak at 11.29 minutes (EPA retention time), as well as a very weak peak at 13.8 minutes (DHA retention time) only for clone # 5. The chromatogram and mass spectra for FOS clone # 5 is shown in Figure 4.7 as an example. The spectra are visually quite similar to the standard spectra. 4.4.6 Assessment of recombinant production of EPA/DHA by FOS clones in LB broth at different growth temperatures The production of EPA/DHA in EPA/DHA-producing FOS clones was tested at 10°C, 15°C, 20°C and 25°C in LB broth. EPA was produced by all FOS clones at all temperatures. DHA was produced only by clone # 5 at 10°C, 15°C and 20°C. Among all 135 B Hki lIllillllllllLUlillllllllH.ltl.lillu.LLl.g.J L.,,u m illllJilllllliJiUjjjLkLlUlJ, 3d o Figure 4.6. The mass spectra for EPA/DHA standard. A) EPA/DHA chromatograph analysed by GC-MS; B) mass spectra for EPA peak at retention time of 11.29; C) mass spectra of DHA peak at retention time of 13.7. 136 RIC all 5-e.sms u ii B mil llllll III lllllll mJ,Ulu[lM,iklJ,l,hl.l,i].,lm.ll,iJ,l..i.l,l ..»;]„• 293 311 Ill UHH tlillillibli.L.lii.il.iJ.ili • U.I. I Figure 4.7. Mass spectra for clone 5 (EPA/DHA positive clone). A) chromatograph of peaks at retention time 11.28 and 13.8 analysed by GC-MS; B) mass spectra for EPA peak at retention time of 11.28; C) mass spectra of DHA peak at retention time of 13.7. Clone 5 was grown at 15°C for 8 days and fatty acids extracted from 0.1 g dry cells. Extracted FAs were analysed by GC-MS in Chemistry department, University of Guelph. 137 tested FOS clones, Fosmid 6 (insert size of 35 kb) was found to positively hybridize to both probes (pfaA and pfaD). However, neither EPA nor DHA was produced by FOS 6 at any temperatures. This indicates that the complete sequence of pfaA and pfaD are essential for EPA/DHA production. It is likely that fosmid 6 did not have the entire pfaA and/or pfaD genes. In two other studies, it is reported that a fosmid clone (33kb) and a cosmid clone (40kb) containing pfaA-D genes were not able to produce recombinant EPA/DHA in E. coli (Allen & Bartlett, 2002; Tanaka et al, 1999). They suggested that an additional gene (PPTase) whose product (phosphopantetheinyl) is essential for the synthesis of acyl carrier protein domain present in pfaA was needed. PPTase gene (ORF2) is located within close vicinity of the pfaA-D operons (Metz et al., 2001). According to this, we speculate that fosmid 6 also may lack the PPTase gene. The production of EPA by all FOS clones at 10°C, 15°C, 20°C and 25°C are shown in figures 4.8 and 4.9. At 10°C all the clones produced 2.5- to 8-fold more EPA compared to S. baltica MAC1. At 15°C they produced 4.2- to 9.8-times more EPA than S. baltica MAC1. At 20°C, EPA production by FOS clones was 3.4- to 4.8-times more than S. baltica MAC1. At 25°C, they produced 3.7- to 6.6-fold more EPA than S. baltica MAC1. All FOS clones produced EPA, 3.6%-10.5% of total fatty acids at 10°C, 4.8-14% at 15°C, 4.1-6% at 20°C and 3-5.7% at 25°C. More EPA was produced in all FOS clones at 15°C. The recombinant production of EPA by transgenic E. coli EPI3000-T1 (FOS clones) does not correspond with that in S. baltica MAC1, where the level of EPA increased with decreasing growth temperature (Amiri-Jami et al., 2006). The same 138 10C 25 20 ui 15 O) 1(H III S.b.M FC2 FC3 FC4 FC5 FC6 FC7 FC8 FC9 E.coli Fosmid Clones 15C 20 15 UJ "5 10 en S.b,M FC2 FC3 FC4 FC5 FC6 FC7 FC8 FC9 E.coli Fosmid Clones Figure 4.8. EPA production (mg/g cell dry weight) by clones at 10°C and 15°C EPA extracted from S. baltica MAC1, EPI300-T1 E. coli and 8 fosmid clones, when cultured in LB broth and grown at 10°C and 15°C. Total fatty acid was extracted from freeze-dried cell and analyzed using and analyzed using an automated Agilent 6890 GC. This is the result of two independent experiments. 139 20C 12 10 2 8 iu •S 6H ? 4^ S.b.M FC2 FC3 FC4 FC5 FC6 FC7 FC8 FC9 E.coli Fosmid Clones 25C 10 . UJ O) 2J S.b.M FC2 FC3 FC4 FC5 FC6 FC7 FC8 FC9 E.coli Fosmid Clones Figure 4.9. EPA production (mg/g cell dry weight) by clones at 20°C and 25°C EPA extracted from S. baltica MAC1, EPI300-T1 E. coli and 8 fosmid clones, when cultured in LB broth and grown at 20°C and 25°C. Total fatty acid was extracted from freeze-dried cell and analyzed using an automated Agilent 6890 GC. This is the result of two independent experiments. 140 phenomenon is reported for recombinant production of DHA in E. coli DH5a (Orikasa et al, 2006b) where the level of DHA was increased by increasing the temperature to 15°C. It is also reported that E. coli DH5a carrying a pEPA plasmid (containing ORF2-9) produced most EPA at 15°C (Orikasa et al, 2004). In addition, the amount of EPA produced in FOS clones carrying variations of the EPA biosynthesis gene cluster may be related to their construction. It is reported that removal of ORF9 may enhance expression of the EPA biosynthesis genes (Orikasa et al, 2004). When Orikasa et al (2004) deleted different ORFs in the 38 kb EPA biosynthesis gene cluster, a different amount of EPA was produced (1.4-21.6%). It is also reported that the relative ratio of PPTase protein to the other EPA biosynthesis genes is an important factor in the production of EPA (Orikasa et al, 2006b; Sugihara et al, 2008). The percentage of major detected fatty acid in FOS clone # 5 that produces both EPA and DHA at different temperatures are shown in Table 4.4. The production of short chain fatty acids (CI2, C14, CI6) was increased slightly when temperature was increased. This is in contrast with the situation in psychrophilic bacteria, which respond to changes in growth temperature by increasing levels of short chain fatty acids at lower temperatures (Hamamoto et al, 1994). This suggests that the EPA/DHA gene cluster only had an effect on EPA/DHA production in the FOS clone. The level of EPA in FOS clone # 5 was 8.9%, 14%, 7.04% and 3.9% of total fatty acids at 10°C, 15°C, 20°C, and 25°C, respectively (Figure 4.10). The highest production was at 15°C. This suggests that the relationship between temperature and EPA production in E. coli might not be controlled only by EPA/DHA biosynthesis enzymes but some other unknown factor(s) 141 Table 4.4 Fatty acid composition of FOS clone 5 harboring EPA/DHA gene cluster % of total fatty acids at different temperatures Fatty acid 10°C 15°C 20°C 25°C 12:0 1.04 2.4 3.2 4.2 14:0 5.6 6.1 6.7 7 16:0 36.2 36.8 40 40.8 16:1 16.2 17.8 14.6 18.2 18:ln9c 12.4 15 14.5 14.2 20:5n3 8.9 13 7.04 3.9 22:6n3 0.34 0.202 0.105 tr Others8 19.3 8.69 13.8 11.7 a includes: 17:0, 17:1, 18:2n6, 18:3n3, 18:3n6, 20:0, 20:ln9, 21:0, 20:3n6, 20:3n3, 22:0 and22:ln9. 142 20°C and25°C.Totalfattyaciwasextractefromfreeze-driecellsanalyzeusing Figure 4.10EffectofgrowthtemperaturnthproductioEPAbyfosmidclon5 an automatedAgilent6890GC.Thisitheresul t oftwindependenexperiments. (transgenic E.coliEPI300-T1).CellswereculturedinLBbrothangrowat10°C,15°C D) o E EPA 20 1 10 - 12- 14 - 16- 18- 4 - 0 - 2 - 6 8- 10°C 15°20°25° 143 Temperature may be involved as well. The level of DHA increased with decreasing growth temperature. This phenomenon is similar to what we have observed for S. baltica MAC1. Moreover, it is reported that transgenic E. coli AGI and JM109 harboring the EPA gene cluster produced 3 and 1.4% EPA at 25°C, respectively (Orikasa et al, 2004). Recombinant production of EPA in FOS clone 5 is 4% of total fatty acid at 25°C. This suggests that the production of EPA depends on the host strain of E. coli. The reason for this is not yet clear, however the level of enzyme activity or the expression level of the biosynthesis gene cluster might be different in different host strains of E. coli. 4.4.7 Sequence analysis of the 5. baltica MAC1 EPA/DHA gene cluster The isolation of Shewanella baltica MAC1 gene cluster required for EPA/DHA biosynthesis involved the construction of a large insert (~ 40 kb) fosmid library of S. MAC1 genomic DNA and the subsequent probing of the library with the partial fragments of pfaA and pfaD genes. The entire DNA insert from the clone positive for EPA/DHA production was sequenced. The insert size was 35373 bp. We then analysed the entire sequence for all possible ORFs and the position of the ORFs potentially responsible for EPA/DHA production. A total of 16 predicted ORFs were identified within the sequence (figure 4.11) as well as the size and BLAST searches show the similarity of these ORFs to other sequences available in GenBank. Of the 16 ORFs identified, ORF4 (1004 bp), ORF5 (8114 bp), ORF6 (2250 bp), ORF7 (3367 bp) and ORF8 (1705 bp) gave similarity to ppTase, pfaA, pfaB, pfaC and pfaD genes, respectively. These five ORFs span a region of 20668 bp. 144 ORF Size (bp) Putative function 1 840 Transcriptional regulator, LysR 2 1987 Beta-lactamase domain-containing protein 3 550 Unknown hypothetical protein 4 1004 Phosphopantetheinyl transferase (ppTase gene) 5 8114 pfaA of EPA and DHA gene cluster 6 2250 pfaB of EPA gene cluster 7 3367 pfaC of EPA and DHA gene cluster 8 1705 pfaD of EPA and DHA gene cluster 9 424 Conserved hypothetical protein 10 6695 Conserved hypothetical protein 11 1005 Conserved hypothetical protein 12 1055 Major royal jelly protein 13 1448 Conserved hypothetical protein 14 493 Specific transcriptional regulator 15 560 Conserved hypothetical protein 16 430 Conserved hypothetical protein Figure 4.11 Shewanella baltica MAC1 EPA/DHA gene cluster and flanking DNA. Graphic map showing the organization of 16 ORFs identified from sequencing of the 35 kb insert of the fosmid DNA. The size of each ORF and their similarity to sequences obtained from GenBank on BLAST searches. 145 We then compared the homology of these five ORFs to the pfa genes from Shewanella pneumatophori SCRC, Photobacterium profundum SS9 and Moritella marina MP-1. The nucleotide or amino acid similarity was as follow: 67% to Shewanella SCRC pfaA, 68% to SS9 pfaA, 69% to Moritella pfaA; 68% to Shewanella SCRC pfaB, 65% to SS9 pfaB, no homology was found for Moritella pfaB; 69% to Shewanella SCRC pfaC, 70% to SS9 pfaC, 67% to Moritella pfaC; 77% to Shewanella pfaD, 74% to SS9 pfaD, 67% to Moritella pfaD. No homology was found between the ppTase (pfaE) gene of Shewanella baltica MAC1 and Shewanella pneumatophori SCRC. Moreover, the gene cluster isolated from Photobacterium profundum SS9 and Moritella marina had only pfaABCD but lacked pfaE {ppTase) gene (Okuyama et al, 2007). The ORF4 of Shewanella baltica MAC1 showed 97% nucleotide identity to the phosphopantetheinyl transferase {ppTase) gene of Shewanella baltica OS223. The EPA biosynthesis genes from Shewanella pneumatophori SCRC include five essential ORFs (2, 5, 6, 7, and 8) (Orikasa et al, 2004). In this study, ORFs5, 6, 7, and 8 of S. baltica MAC1 showed partial homology to ORFS5, 6, 7, and 8 of Shewanella pneumatophori SCRC. In the EPA/DHA gene cluster isolated from S. baltica MAC1, ORF4 was identified as the ppTase gene while in the EPA gene cluster from S. pneumatophori ORF2 was identified as the ppTase gene. Furthermore, unlike the EPA gene cluster isolated from S. pneumatophori SCRC that has two other ORFs (ORF3 and ORF4) between ppTase and pfaA gene, there are no ORFs between the ppTase and pfaA gene in the S. baltica MAC1 EPA/DHA gene cluster. Upstream of the ORF4 {ppTase gene) and down stream of ORF8 (pfaD gene) of the EPA/DHA gene cluster isolated from S. baltica MAC1 are conserved hypothetical proteins with unknown functions (figure 4.11). Recombinant production of EPA and 146 DHA has not been detected when the EPA and DHA gene cluster were isolated from Photobacterium profundum SS9 and Moritella marina MP-1, respectively, most probably due to the lack of a ppTase ipfaE) gene in their gene cluster (Allen & Bartlett, 2002; Morita et al, 1999). In another study when the ppTase gene was cloned and coexpressed with pfaA-D genes, recombinant production of DHA was reported (Sugihara et al, 2008). These reports and our findings suggest that the ppTase gene is essential for EPA/DHA production. The protein domains identified in the gene cluster were 3-ketoacyl synthase (KS), acyl carrier protein (ACP), P-hydroxyacyl-ACP dehydratase (HD), acyl transferase (AT), 3-ketoacyl reductase (KR), short-chain dehydrogenase reductase (DR) and phosphopantetheinyl transferase (ppTase). The BLAST search showed that the pfaA gene encodes KS, AT, KR and DR domains; pfaB gene encodes HD and KS; pfaC encodes HD and KS domains; and pfaD gene encodes a HD domain. The pfaE gene encodes KS, ppTase and amino acid/peptide transporter. The domain structure of our EPA/DHA gene cluster is different from that of the Shewanella pneumatophori SCRC. The EPA/DHA gene cluster isolated from S. baltica MAC1 has an extra DR domain in pfaA gene (not found in S. SCRC), an extra HD domain in pfaB and pfaD, and a KS domain in pfaE in addition to ppTase. 4.5 Conclusions We successfully cloned the EPA/DHA gene cluster from S. baltica MAC1 by constructing a fosmid library contained large genomic DNA inserts (~40kb). To screen 147 the large number of the FOS clones for EPA/DHA genes, we established a fast detection method for EPA/DHA gene cluster by partial amplification of pfaA, pfaB, pfaC, pfaD and pfaE genes of S. baltica MAC1. Screening of fosmid clones with pfaA and pfaD probes enabled us to detect inserts containing EPA/DHA genes. We obtained a transgenic E. coli with the ability to produce recombinant EPA and DHA. Unlike BAC DNA, FOS DNA could be isolated, digested and end-sequenced easily. This suggests that the size of insert has a direct effect on the manipulation and future application of the insert containing the EPA/DHA genes. In addition, preparing a library with smaller DNA inserts decreases the risk of getting chimeric clones. We obtained an EPA/DHA gene cluster that could be ligated into a food grade broad host range vector in order to get recombinant EPA and DHA in other organisms as alternatives for fish oils. Furthermore, having the complete sequence of EPA/DHA gene cluster cloned from S. baltica MAC1 enabled us to identify pfaA-E genes responsible for EPA/DHA production in S. baltica MAC1. 148 5. General Discussion Several strains of marine bacteria, particularly those belonging to the genus Shewanella, has been the focus of extensive recent studies because of their ability to produce EPA and DHA (Bowman et al, 1997; Hau & Gralnick, 2007; Hirota et al, 2005; Nichols et al, 1997; Russell & Nichols, 1999; Sato et al, 2008; Satomi et al, 2003). Shewanella baltica MAC1 isolated from mackerel entrails produces EPA and DHA (Cadieux, 1996). This bacterium was initially identified as Shewanella putrefaciens MAC1 based on its fatty acid composition and its biochemical profile (Cadieux, 1996). In the present study Shewanella MAC1 was reclassified on the basis of 16S rRNA sequence data. The complete 16S rRNA gene sequence of 5. sp. MAC1 showed 99% homology to S. baltica strains. Therefore, S. putrefaciens MAC1 was renamed S. baltica MAC1. S. baltica is described as a non-EPA producing Shewanella (Bowman, 1999; Kato & Nogi, 2001; Satomi et al., 2003). Four S. baltica were examined for EPA/DHA production. All S. baltica strains used in this study were positive for EPA production. Our ability to detect EPA in 5. baltica strains may be due to the improved method used for the extraction of the total fat from dried bacteria. The method of Folch or Bligh and Dyer was used by other researchers for lipid extraction from Shewanella or other marine bacteria (Gentile et al, 2003; Morita et al, 2005; Yano et al, 1994). In the modified method that we used to extract the lipid; i) more cells were used, ii) cell membrane was disrupted by grinding which increases the contact of the solvent with the lipid in the sample, iii) saturated salt increases the solution polarity and helps to remove the lipid from debris, and iv) a larger volume of organic solvent was used to separate the fat from 149 cell debris. All these modifications might have contributed to our ability to detect EPA in 5. baltica strains. Production of omega-3 fatty acids by S. baltica MAC1 made this bacterium a prime candidate for genetic studies. Cloning EPA/DHA genes from 5. baltica MAC1 and ligating the gene cluster to a food grade, broad host range plasmid may provide alternative sources which can be used in the food industry. S. putrefaciens SCRC-2738, a bacterium isolated from mackerel entrails (Yazawa et ah, 1988) produced only EPA. The EPA gene cluster was cloned from S. putrefaciens SCRC-2738 and expressed in the marine cyanobacterium Synechococcus sp. (Takeyama et ai, 1997). However, the yield of EPA was low (0.12 mg g"1 cell dry weight) and it was not considered as a suitable alternative source for EPA production. Constructing a BAC DNA library with large DNA fragments of S. baltica MAC1 resulted in recombinant production of DHA (0.85 % of total fatty acid) and EPA (0.02% of total fatty acid) in E. coli. The amount of EPA produced by E. coli was much lower than that of S. baltica MAC1 (12.6%). One reason may be a low copy number of the BAC DNA. When the copy number of the BAC vector carrying DHA gene cluster was increased using autoinduction solution, the production of DHA was increased by 1.4%. In one study, when the copy number of the plasmid carrying the EPA gene cluster was increased, more EPA was produced in E. coli cells (Orikasa et ai, 2004). In another report, the amount of EPA produced by a cyanobacterial transconjugant was much lower than that of E. coli due to a lower copy number of the cosmid vector in the cyanobacterium as compared to E. coli (Takeyama et ai, 1997). Moreover, BAC replication of large DNA fragments may not follow cell division, and 150 cells without BAC may have been enriched when lipid was extracted from cells for EPA/DHA analysis. We prepared a BAC library with large DNA insert (200 kb) in order to isolate EPA/DHA genes. Although the EPA/DHA gene cluster was cloned and expressed in E. coli, however, due to having a chimeric clone we were unable to isolate the 200 kb DNA fragment containing all EPA/DHA genes. There are reports that 1-6% of BAC library clones could be chimeric (Berton et ai, 2003; Crooijmans et al, 2000; Osoegawa et al, 2000). Constructing a library with smaller DNA fragment (< 200 kb) decreases the chance of getting chimeric clones. However, this significantly increases the number of clones in the library. As a result, a fast and reliable screening method was needed to detect the EPA/DHA gene cluster. We designed primers targeting the sequences within highly conserved domains of the EPA/DHA gene cluster. Five primers were applied successfully for amplification of pfaA, pfaB, pfaC, pfaD and pfaE genes from genomic DNA of S. baltica MAC1. The specificity of amplified fragments was confirmed by sequencing the purified PCR products. The products with the primers EPA5-2, EPA7-2 and EPA 8-2 were specific fragments of the pfaA, pfaC and pfaD genes harbouring p-ketoacyl synthase, 3-hydroxydecanoyl-ACP dehydratases and enoyl reductase domains, respectively. It is reported that the specific products were obtained when primers pfaA.S and pfaC were used (Gentile et al, 2003; Yakimov et al, 2003). The specific fragments of pfaA and pfaC genes contained C-terminal P-ketoacyl synthase, N-teriminal acyl coenzyme A acyl carrier protein acyltransferase and dehydratase /isomerase domains, respectively (Gentile et al., 2003). 151 Probes were made with amplified fragments of the pfaA and pfaD genes. Colony hybridization of 25,00 fosmid clones containing ~40kb DNA insert resulted in 8 clones positive for both pfaA and pfaD genes. Six clones produced EPA and one clone produced EPA and DHA. This suggests that all five genes (pfaA, pfaB, pfaC, pfaD, pfaE) are required for EPA/DHA production. It is reported that only five pfaABCDE genes are necessary for the biosynthesis of EPA and DHA (Okuyama et al, 2007; Orikasa et al., 2004). In addition, no EPA/DHA production was detected in one clone which was positive for both pfaA and pfaD genes. It is reported that the EPA and DHA gene clusters cloned from Photobacterium profundum SS9, an EPA-producing bacterium and Moritella marina MP-1, a DHA-producing bacterium, contained pfaA-pfaD genes, however, no recombinant production of EPA and DHA was detected in E. coli (Allen & Bartlett, 2002; Morita et al., 2000). It was suggested that the gene clusters lacked pfaE (PPTase) gene, the product of which (phosphopantetheinyl transferase) is required for the post- translational modification of the constituent acyl carrier protein, acyltransferase domains present in the pfaA gene (Allen & Bartlett, 2002; Metz et al, 2001). In another study, when the PPTase gene was cloned from Photobacterium profundum SS9 and coexpressed with pfaA-D genes, recombinant production of DHA was obtained (Sugihara et al, 2008). This suggests that our 7 fosmid clones possessing the ability to produce EPA and/or DHA carry all the genes required for EPA/DHA production. Nevertheless, one clone carrying pfaA-D genes may lack the pfaE (PPTase) gene, which resulted in no EPA/DHA production in E. coli. The end-sequence of isolated fosmid DNA from 8 clones showed the size of the insert ranging from 34-41 kb. Unlike BAC DNA, the fosmid DNA from positive clones for EPA/DHA was digested easily with Notl. 152 We obtained an EPA/DHA gene cluster carrying all the genes required for EPA/DHA production that could be cloned into a food grade, broad host range vector in order to get recombinant EPA and DHA in other organisms as alternatives for fish oils. Interestingly, positive clones produced 4.2-9.8 times more EPA compared to the S. baltica MAC1 at 15°C in a simple media (LB broth). The level of DHA production was the same as S. baltica MAC1 (0.4% of total fatty acids). It was expected that only a small amount of DHA would be produced in E. coli, because the original productivity of DHA in S. baltica MAC1 is also much lower than that of EPA. However, the production of DHA could be improved in E. coli. It has been shown that addition of cerulenin to the Moritella marina MP-1 growth medium increased the production of DHA (from 1.6 to 5.7 \ng I dry weight of cells) (Morita et al, 2000). Furthermore, the cerulenin treatment increased the EPA yield in Shewanella marinintestina IK-1 (from 1.6mg to 8 mg l"1) (Morita et al, 2005). The reason why cerulenin inhibits monounsaturated fatty acid biosynthesis while at the same time increasing the polyunsaturated fatty acids biosynthesis is not known. The addition of cerulenin is considered an advantage to increase the bacterial intracellular DHA. Moreover, the production of DHA could be improved by ligating the EPA/DHA gene cluster to an expression plasmid containing a stronger promoter. The complete sequence analysis of the insert isolated from the fosmid clone positive for EPA/DHA production revealed that this pfa gene cluster is unique to Shewanella baltica MAC1 in several aspects. First, only a small segment of pfaA, B, C and D had some degree of similarity to those of Shewanella pneumatorophi SCRC EPA 153 gene cluster. Secondly, there was no homology for the key enzyme (ppTase) between our gene cluster and that of the published gene cluster. Thirdly, the domain structures of genes in our clone were quite different compared to those published. Finally, in our clone there are no extra ORFs between pfaE (ppTase) and the pfaA gene while the only clone reported (Orikasa et al, 2004) to produce recombinant EPA has two extra ORFs between pfaE and pfaA of unknown function(s). These differences in the sequence and gene structure of our clone could account for the production of both EPA and DHA. 154 6. Future directions This thesis provides a strong foundation for future studies in new directions. These are: 1) To subclone the EPA/DHA gene cluster in order to study the involvement of individual ORFs essential for EPA/DHA production. 2) To clone the exact ORFs involved in EPA/DHA production to a food grade vector and express them into other organism such as yeast and plants. 3) To study the effect of clone positive for EPA/DHA in prevention of cancer and cardiovascular disease in animal models. 4) To study EPA/DHA biosynthetic enzymes in order to identify the total domain organization of the 5. baltica MAC1 pfaA-E genes and to define the fatty acid biosynthesis pathways. 5) To test the expression of EPA/DHA gene cluster in other E.coli hosts such as E.coli DH10B and E.coli JM 109. 6) To study the expression of the EPA/DHA gene cluster in E.coli when it is ligated to different vectors. 7) To increase the production of DHA using a stronger promoter and an expression vector. 8) To study the effect of cerulenin and glucose on the levels of EPA/DHA production. 155 7. References Allen, E. E., Facciotti, D. & Bartlett, D. H. (1999). Monounsaturated but not polyunsaturated fatty acids are required for growth of the deep-sea bacterium Photobacterium profundum SS9 at high pressure and low temperature. Appl Environ Microbiol 65,1710-1720. Allen, E. E. & Bartlett, D. H. (2002). Structure and regulation of the omega-3 polyunsaturated fatty acid synthase genes from the deep-sea bacterium Photobacterium profundum strain SS9. Microbiology 148, 1903-1913. Amiri-Jami, M., Wang, H., Kakuda, Y. & Griffiths, M. W. (2006). Enhancement of polyunsaturated fatty acid production by Tn5 transposon in Shewanella baltica. Biotechnol Lett 28, 1187-1192. Ammiraju, J. S., Yu, Y., Luo, M. & other authors (2005). Random sheared fosmid library as a new genomic tool to accelerate complete finishing of rice (Oryza sativa spp. Nipponbare) genome sequence: sequencing of gap-specific fosmid clones uncovers new euchromatic portions of the genome. TheorAppl Genet 111, 1596-1607. Asakawa, S., Abe, I., Kudoh, Y. & other authors (1997). Human BAC library: construction and rapid screening. Gene 191,69-79. 156 Austria, J. A., Richard, M. N., Chahine, M. N., Edel, A. L., Malcolmson, L. J., Dupasquier, C. M. & Pierce, G. N. (2008). Bioavailability of alpha-linolenic acid in subjects after ingestion of three different forms of flaxseed. J Am Coll Nutr 27, 214-221. Barclay, W., Abril, R., Abril, P., Weaver, C. & Ashford, A. (1998). Production of docosahexaenoic acid from microalgae and its benefits for use in animal feeds. World Rev Nutr Diet 83, 61-76. Barclay, W. R., Meager, K. M. & Abril, J. R. (1994). Heterotrophic production of long chain omega-3 fatty acids utilizing algae and algae-like microorganisms. Journal of Applied Phycology 6, 123-129. Barre, D. E. (2007). The role of consumption of alpha-linolenic, eicosapentaenoic and docosahexaenoic acids in human metabolic syndrome and type 2 diabetes—a mini-review. J OleoSci 56, 319-325. Beja, O., Suzuki, M. T., Koonin, E. V. & other authors (2000). Construction and analysis of bacterial artificial chromosome libraries from a marine microbial assemblage. Environ Microbiol 2, 516-529. Beja, O., Koonin, E. V., Aravind, L. & other authors (2002). Comparative genomic analysis of archaeal genotypic variants in a single population and in two different oceanic provinces. Appl Environ Microbiol 68, 335-345. 157 Berquin, I. M., Edwards, I. J. & Chen, Y. Q. (2008). Multi-targeted therapy of cancer by omega-3 fatty acids. Cancer Lett 269, 363-377. Berton, G., Delannay, R., Richard, P., Taberlet, N. & Valance, A. (2003). Two- dimensional inclined chute flows: transverse motion and segregation. Phys Rev E Stat Nonlin Soft Matter Phys 68, 051303. Bligh, E. G. & Dyer, W. J. (1959). A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37, 911-917. Borek, C. (1994). Omega 3 fatty acids as anticarcinogens: cellular and molecular studies. World Rev Nutr Diet 76, 66-69. Bourre, J. M. (2005). Dietary omega-3 Fatty acids and psychiatry: mood, behaviour, stress, depression, dementia and aging. J Nutr Health Aging 9, 31-38. Bowman, J. P., McCammon, S. A., Nichols, D. S., Skerratt, J. H., Rea, S. M., Nichols, P. D. & McMeekin, T. A. (1997). Shewanella gelidimarina sp. nov. and Shewanella frigidimarina sp. nov., novel Antarctic species with the ability to produce eicosapentaenoic acid (20:5 omega 3) and grow anaerobically by dissimilatory Fe(III) reduction. Int J Syst Bacteriol 47, 1040-1047. 158 Bowman, J. P., Gosink, J. J., McCammon, S. A., Lewis, T. E., Nichols, D. S., Nichols, P. D., Skerratt, J. H., Staley, J. T. & McMeekin, T. A. (1998a). Colwellia demingiae sp. nov., Colwellia hornerae sp. nov., Colwellia rossensis and Colwellia psychrotopica sp. nov. : psychrophilic Antarctic species with the ability to synthesize docosahexaenoic acid Int J Syst Bacteriol 48, 1171 -1180. Bowman, J. P., McCammon, S. A., Lewis, T., Skerratt, J. H., Brown, J. L., Nichols, D. S. & McMeekin, T. A. (1998b). Psychroflexus torquis gen. nov., sp. nov., a psychrophilic species from Antarctic sea ice, and reclassification of Flavobacterium gondwanense (Dobson et al. 1993) as Psychroflexus gondwanense gen. nov., comb. nov. Microbiology 144 ( Pt 6), 1601-1609. Bowman, J. P. (1999). Genus Shewanella. Bergey's Manual of Systematic Bacteriology. Cadieux, S. U. (1996). Characerization of a marine bacterium for the production of eicosapenataenoic acid (EPA) in unsalted sweet whey, theis, 38-71. Cadieux, S. U., Kakuda, Y., Ward, O. P. & Griffiths, M. W. (1998). Bacterial production of Omega-3 fatty acids in whey. Milchwissenschaft 53, 547-551. Calder, P. C, Bevan, S. J. & Newsholme, E. A. (1992). The inhibition of T-lymphocyte proliferation by fatty acids is via an eicosanoid-independent mechanism. Immunology 75, 108-115. 159 Calder, P. C. (1997). n-3 polyunsaturated fatty acids and cytokine production in health and disease. Ann Nutr Metab 41, 203-234. Cannon, C. P. (2008). Combination therapy in the management of mixed dyslipidaemia. J Intern Med 263, 353-365. Caramia, G. (2008). [The essential fatty acids omega-6 and omega-3: from their discovery to their use in therapy]. Minerva Pediatr 60, 219-233. Castoldi, A. F., Coccini, T. & Manzo, L. (2003). Neurotoxic and molecular effects of methylmercury in humans. Rev Environ Health 18, 19-31. Chen, L., Kosslak, R. & Atherly, A. G. (1994). Mechanical shear of High Molecular Weight DNA in agarose plugs. Biotechniques 16, 228. Chi, Z., Liu, Y., Frear, C. & Chen, S. (2008). Study of a two-stage growth of DHA- producing marine algae Schizochytrium limacinum SR21 with shifting dissolved oxygen level. Appl Microbiol Biotechnol. Clarke, S. D. (2001). Polyunsaturated fatty acid regulation of gene transcription: a molecular mechanism to improve the metabolic syndrome. J Nutr 131, 1129-1132. 160 Cohen, B. E., Garg, S. K., AH, S., Harris, W. S. & Whooley, M. A. (2008). Red blood cell docosahexaenoic acid and eicosapentaenoic acid concentrations are positively associated with socioeconomic status in patients with established coronary artery disease: data from the Heart and Soul Study. JNutr 138, 1135-1140. Crooijmans, R. P., Vrebalov, J., Dijkhof, R. J., van der Poel, J. J. & Groenen, M. A. (2000). Two-dimensional screening of the Wageningen chicken BAC library. Mamm Genome 11, 360-363. Delarue, J., LeFoll, C, Corporeau, C. & Lucas, D. (2004). N-3 long chain polyunsaturated fatty acids: a nutritional tool to prevent insulin resistance associated to type 2 diabetes and obesity? Reprod Nutr Dev 44, 289-299. Delong, E. F. & Yayanos, A. A. (1986). Biochemical Function and Ecological Significance of Novel Bacterial Lipids in Deep-Sea Procaryotes. Appl Environ Microbiol 51, 730-737. Deng, D. y., Zhao, G., xuan, J. S., Yang, J. L., Duan, D. L., Weng, M. L. & Wang, B. (2004). Construction and characterization of a Bacterial Artificial Chromosome library of Marine Macroalgae Porphyra yezoensis (Rhodophyta). Plant Molecular Biology Reporter 22, 375-386. 161 Diaz-Perez, S. V., Crouch, V. W. & Orbach, M. J. (1996). Construction and Characterization of a Magnaporthe grisea Bacterial Artificial Chromosome Library. Fungal Genet Biol 20, 280-288. Folch, J., Lees, M. & Sloane-Stanley, G. (1957). A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem 226, 497-509. Folkertsma, R. T., Spassova, M. I., Prins, M., Stevens, M. R., Hille, J. & Goldbach, R. W. (1999). Construction of a bacterial artificial chromosome (BAC) library of Lycopersicon esculentum cv. Stevens and its application to physically map the Sw-5 locus Molecular Breeding 5, 197-207. Fontani, G., Corradeschi, F., Felici, A. & other authors (2005). Blood profiles, body fat and mood state in healthy subjects on different diets supplemented with Omega-3 polyunsaturated fatty acids. Eur J Clin Invest 35,499-507. Foot, M., Jeffcoat, R., Barratt, M. D. & Russell, N. J. (1983). The effect of growth temperature on the membrane lipid environment of the psychrophilic bacterium Micrococcus cryophilus. Arch Biochem Biophys 224, 718-727. Frolova, G. M., Pavel, K. G., Shparteeva, A. A., Nedashkovskaia, O. I., Gorshkova, N. M., Ivanova, E. P. & Mikhailov, V. V. (2005). [Lipid composition of novel Shewanella species isolated from far Eastern seas]. Mikrobiologiia 74, 766-771. 162 Gentile, G., Bonasera, V., Amico, C, Giuliano, L. & Yakimov, M. M. (2003). Shewanella sp. GA-22, a psychrophilic hydrocarbonoclastic antarctic bacterium producing polyunsaturated fatty acids. JAppl Microbiol 95, 1124-1133. Guijarro, A., Laviano, A. & Meguid, M. M. (2006). Hypothalamic integration of immune function and metabolism. Prog Brain Res 153, 367-405. Guillem, J. G., O'Brian, C. A., Fitzer, C. J., Forde, K. A., LoGerfo, P., Treat, M. & Weinstein, I. B. (1987). Altered levels of protein kinase C and Ca2+-dependent protein kinases in human colon carcinomas. Cancer Res 47, 2036-2039. Hamamoto, T., Takata, N., Kudo, T. & Horikoshi, K. (1994). Effect of temperature and growth phase on fatty acid composition of the psychrophilic Vibrio sp. strain no.5710 FEMS microbiology letters 119, 77-82. Hammer, B. W. & Long, H. F. (1941). Factors Influencing Bacterial Growth in Butter. Bacteriol Rev 5, 337-374. Hartweg, J., Perera, R., Montori, V., Dinneen, S., Neil, H. A. & Farmer, A. (2008). Omega-3 polyunsaturated fatty acids (PUFA) for type 2 diabetes mellitus. Cochrane Database Syst Rev, CD003205. 163 Hastings, N., Agaba, M. K., Tocher, D. R., Zheng, X., Dickson, C. A., Dick, J. R. & Teale, A. J. (2004). Molecular cloning and functional characterization of fatty acyl desaturase and elongase cDNAs involved in the production of eicosapentaenoic and docosahexaenoic acids from alpha-linolenic acid in Atlantic salmon (Salmo salar). Mar Biotechnol (NY) 6,463-474. Hau, H. H. & Gralnick, J. A. (2007). Ecology and biotechnology of the genus Shewanella. Annu Rev Microbiol 61, 237-258. Heird, W. C. & Lapillonne, A. (2005). The role of essential fatty acids in development. Annu Rev Nutr 25, 549-571. Henderson, R. J., Millar, R. M., Sargent, J. R. & Jostensen, J. P. (1993). Trans- monoenoic acid and polyunsaturated fatty acids in phospholipids of a Vibrio species of bacterium in relation to growth conditions. Lipids 28, 389-397. Hirota, K., Nodasaka, Y., Orikasa, Y., Okuyama, H. & Yumoto, I. (2005). Shewanella pneumatophori sp. nov., an eicosapentaenoic acid-producing marine bacterium isolated from the intestines of Pacific mackerel (Pneumatophorus japonicus). Int J Syst Evol Microbiol 55, 2355-2359. 164 Huang, Y., Lai, X., He, X., Cao, L., Zeng, Z., Zhang, J. & Zhou, S. (2008). Characterization of a Deep-Sea Sediment Metagenomic Clone that Produces Water- Soluble Melanin in Escherichia coli. Mar Biotechnol (NY). Hulthe, J., Bokemark, L., Wikstrand, J. & Fagerberg, B. (2000). The metabolic syndrome, LDL particle size, and atherosclerosis: the Atherosclerosis and Insulin Resistance (AIR) study. Arterioscler Thromb Vase Biol 20, 2140-2147. Innis, S. M. (2008). Dietary omega 3 fatty acids and the developing brain. Brain Res. Ivanova, E. P., Nedashkovskaya, O. I., Sawabe, T., Zhukova, N. V., Frolova, G. M., Nicolau, D. V., Mikhailov, V. V. & Bowman, J. P. (2004). Shewanella affinis sp. nov., isolated from marine invertebrates. Int J Syst Evol Microbiol 54, 1089-1093. Jolly, C. A. (2005). Diet manipulation and prevention of aging, cancer and autoimmune disease. Curr Opin Clin Nutr Metab Care 8, 382-387. Kato, C. & Nogi, Y. (2001). Correlation between phylogenetic structure and function: examples from deep-sea Shewanella. FEMS Microbiol Ecol 35, 223-230. Khal, J. & Tisdale, M. J. (2008). Downregulation of muscle protein degradation in sepsis by eicosapentaenoic acid (EPA). Biochem Biophys Res Commun 375, 238-240. 165 Kobayashi, N., Barnard, R. J., Henning, S. M. & other authors (2006). Effect of altering dietary omega-6/omega-3 fatty acid ratios on prostate cancer membrane composition, cyclooxygenase-2, and prostaglandin E2. Clin Cancer Res 12,4662-4670. Krebs, J. D., Browning, L. M., McLean, N. K., Rothwell, J. L., Mishra, G. D., Moore, C. S. & Jebb, S. A. (2006). Additive benefits of long-chain n-3 polyunsaturated fatty acids and weight-loss in the management of cardiovascular disease risk in overweight hyperinsulinaemic women. Int J Obes (Lond) 30, 1535-1544. La Guardia, M., Giammanco, S., Di Majo, D., Tabacchi, G., Tripoli, E. & Giammanco, M. (2005). Omega 3 fatty acids: biological activity and effects on human health. Panminerva Med 47, 245-257. Laidlaw, M. & Holub, B. J. (2003). Effects of supplementation with fish oil-derived n-3 fatty acids and gamma-linolenic acid on circulating plasma lipids and fatty acid profiles in women. Am J Clin Nutr 77, 37-42. Lee, J. V., Gibson, D. M. & Shewan, J. M. (1977). A numeroca taxonomic study of some Pseudomonas-like marine bacteria. / Gen Microbiol 98,439-451. Lijavetzky, D., Muzzi, G., Wicker, T., Keller, B., Wing, R. & Dubcovsky, J. (1999). Construction and characterization of a bacterial artificial chromosome (BAC) library for the A genome of wheat. Genome 42, 1176-1182. 166 Luo, M., Wang, Y. H., Frisch, D., Joobeur, T., Wing, R. A. & Dean, R. A. (2001). Melon bacterial artificial chromosome (BAC) library construction using improved methods and identification of clones linked to the locus conferring resistance to melon Fusarium wilt (Fom-2). Genome 44, 154-162. MacDonell, M. T. & Colwell, R. R. (1985). Phylogeny of the Vibrionaceae, and recommendation for two new genera, Listonella and Shewanella. Syst Appl Microbiol 6, 171-182. McCarty, M. F. (1996). Fish oil may impede tumour angiogenesis and invasiveness by down-regulating protein kinase C and modulating eicosanoid production. Med Hypotheses 46, 107-115. McNamara, R. K., Jandacek, R., Rider, T., Tso, P., Stanford, K. E., Hahn, C. G. & Richtand, N. M. (2008). Deficits in docosahexaenoic acid and associated elevations in the metabolism of arachidonic acid and saturated fatty acids in the postmortem orbitofrontal cortex of patients with bipolar disorder. Psychiatry Res 160, 285-299. Methe, B. A., Nelson, K. E., Deming, J. W. & other authors (2005). The psychrophilic lifestyle as revealed by the genome sequence of Colwellia psychrerythraea 34H through genomic and proteomic analyses. Proc Natl Acad Sci USA 102, 10913-10918. 167 Metroka, C. E., Truong, P. & Gotto, A. M., Jr. (2007). Treatment of HIV-associated dyslipidemia: a role for omega-3 fatty acids. AIDS Read 17, 362-364, 367-373. Metz, J. G., Roessler, P., Facciotti, D. & other authors (2001). Production of polyunsaturated fatty acids by polyketide synthases in both prokaryotes and eukaryotes. Science 293, 290-293. Minami, A., Ishimura, N., Sakamoto, S., Takishita, E., Mawatari, K., Okada, K. & Nakaya, Y. (2002). Effect of eicosapentaenoic acid ethyl ester v. oleic acid-rich safflower oil on insulin resistance in type 2 diabetic model rats with hypertriacylglycerolaemia. Br J Nutr 87, 157-162. Mitmesser, S. H. & Jensen, C. L. (2007). Roles of long-chain polyunsaturated fatty acids in the term infant: developmental benefits. Neonatal Netw 26, 229-234. Mori, T. A., Bao, D. Q., Burke, V., Puddey, I. B. & Beilin, L. J. (1999). Docosahexaenoic Acid but Not Eicosapentaenoic Acid Lowers Ambulatory Blood Pressure and Heart Rate in Humans Hypertension 34, 253-260. 168 Morita, N., ueno, A., Tamaka, M., Ohgiya, S., Hoshino, T., Kawasaki, K., Yumoto, I., Ishizaki, K. & Okuyama, H. (1999). Cloning and sequencing of clustered genes involved in fatty acid biosynthesis from the docosahexaenoic acid-producing bacterium, Vibrio marinus strain MP-1. Biotechnology letters 21, 641-646. Morita, N., Tanaka, M. & Okuyama, H. (2000). Biosynthesis of fatty acids in the docosahexaenoic acid-producing bacterium Moritella marina strain MP-1. Biochem Soc Trans 28, 943-945. Morita, N., Nishida, T., Tanaka, M., Yano, Y. & Okuyama, H. (2005). Enhancement of polyunsaturated fatty acid production by cerulenin treatment in polyunsaturated fatty acid-producing bacteria. Biotechnol Lett 27, 389-393. Mozaffarian, D., Geelen, A., Brouwer, I. A., Geleijnse, J. M., Zock, P. L. & Katan, M. B. (2005). Effect of fish oil on heart rate in humans: a meta-analysis of randomized controlled trials. Circulation 112, 1945-1952. Mozaffarian, D., Stein, P. K., Prineas, R. J. & Siscovick, D. S. (2008). Dietary fish and omega-3 fatty acid consumption and heart rate variability in US adults. Circulation 117, 1130-1137. Mozo, T., Fischer, S., Shizuya, H. & Altmann, T. (1998). Construction and characterization of the IGF Arabidopsis BAC library. Mol Gen Genet 258, 562-570. 169 Myers, C. R. & Nealson, K. H. (1988). Bacterial Manganese Reduction and Growth with Manganese Oxide as the Sole Electron Acceptor. Science 240, 1319-1321. Nakamura, K., Kariyazono, H., Komokata, T., Hamada, N., Sakata, R. & Yaniada, K. (2005). Influence of preoperative administration of omega-3 fatty acid-enriched supplement on inflammatory and immune responses in patients undergoing major surgery for cancer. Nutrition 21, 639-649. Napier, J. A., Sayanova, O., Qi, B. & Lazarus, C. M. (2004). Progress toward the production of long-chain polyunsaturated fatty acids in transgenic plants. Lipids 39, 1067-1075. Nestel, P., Shige, H., Pomeroy, S., Cehun, M., Abbey, M. & Raederstorff, D. (2002). The n-3 fatty acids eicosapentaenoic acid and docosahexaenoic acid increase systemic arterial compliance in humans. Am J Clin Nutr 76, 326-330. Nettleton, J. A. & Katz, R. (2005). n-3 long-chain polyunsaturated fatty acids in type 2 diabetes: a review. J Am Diet Assoc 105,428-440. Nichols, D. S., Nichols, P. D., Russell, N. J., Davies, N. W. & McMeekin, T. A. (1997). Polyunsaturated fatty acids in the psychrophilic bacterium Shewanella 170 gelidimarina ACAM 456T: molecular species analysis of major phospholipids and biosynthesis of eicosapentaenoic acid. Biochim Biophys Acta 1347, 164-176. Okuyama, H., Orikasa, Y., Nishida, T., Watanabe, K. & Morita, N. (2007). Bacterial genes responsible for the biosynthesis of eicosapentaenoic and docosahexaenoic acids and their heterologous expression. Appl Environ Microbiol 73, 665-670. Orikasa, Y., Yamada, A., Yu, R., Ito, Y., Nishida, T., Yumoto, I., Watanabe, K. & Okuyama, H. (2004). Characterization of the eicosapentaenoic acid biosynthesis gene cluster from Shewanella sp. strain SCRC-2738. Cell Mol Biol (Noisy-le-grand) 50, 625- 630. Orikasa, Y., Nishida, T., Hase, A., Watanabe, K., Morita, N. & Okuyama, H. (2006a). A phosphopantetheinyl transferase gene essential for biosynthesis of n-3 polyunsaturated fatty acids from Moritella marina strain MP-1. FEBS Lett 580, 4423- 4429. Orikasa, Y., Nishida, T., Yamada, A., Yu, R., Watanabe, K., Hase, A., Morita, N. & Okuyama, H. (2006b). Recombinant production of docosahexaenoic acid in a polyketide biosynthesis mode in Escherichia coli. Biotechnol Lett 28, 1841-1847. 171 Osoegawa, K., Tateno, M., Woon, P. Y., Frengen, E., Mammoser, A. G., Catanese, J. J., Hayashizaki, Y. & de Jong, P. J. (2000). Bacterial artificial chromosome libraries for mouse sequencing and functional analysis. Genome Res 10, 116-128. Owen, R. J., Legors, R. M. & Lapage, S. P. (1978). Base composition, size and sequence similarities of genoma deoxyribonucleic acids from clinical isolates of Pseudomonas putrefaciens. J Gen Microbiol 104, 127-138. Parra, D., Ramel, A., Bandarra, N., Kiely, M., Martinez, J. A. & Thorsdottir, I. (2008). A diet rich in long chain omega-3 fatty acids modulates satiety in overweight and obese volunteers during weight loss. Appetite 51, 676-680. Patil, V., Kallqvist, T., Olsen, E., Vogt, G. & Gisler0d, H. R. (2007). Fatty acid composition of 12 microalgae for possible use in aquaculture feed Aquaculture International 15, 1-9. Pereira, S. L., Huang, Y. S., Bobik, E. G., Kinney, A. J., Stecca, K. L., Packer, J. C. & Mukerji, P. (2004). A novel omega3-fatty acid desaturase involved in the biosynthesis of eicosapentaenoic acid. Biochem J 378, 665-671. Ratledge, C. (2004). Fatty acid biosynthesis in microorganisms being used for Single Cell Oil production. Biochimie 86, 807-815. 172 Rose, D. P. & Connolly, J. M. (2000). Regulation of tumor angiogenesis by dietary fatty acids and eicosanoids. Nutr Cancer 37, 119-127. Russell, N. J. & Nichols, D. S. (1999). Polyunsaturated fatty acids in marine bacteria—a dogma rewritten. Microbiology 145 (Pt 4), 767-779. Sabehi, G., Loy, A., Jung, K. H., Partha, R., Spudich, J. L., Isaacson, T., Hirschberg, J., Wagner, M. & Beja, O. (2005). New insights into metabolic properties of marine bacteria encoding proteorhodopsins. PLoS Biol 3, e273. Sambrook, J. & Russell, D. W. (2001). Molecular Cloning: A laboratory Manual. . Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. SanGiovanni, J. P. & Chew, E. Y. (2005). The role of omega-3 long-chain polyunsaturated fatty acids in health and disease of the retina. Prog Retin Eye Res 24, 87- 138. Sassen, L. M., Lamers, J. M. & Verdouw, P. D. (1994). Fish oil and the prevention and regression of atherosclerosis. Cardiovasc Drugs TherS, 179-191. Sato, S., Kurihara, T., Kawamoto, J., Hosokawa, M., Sato, S. B. & Esaki, N. (2008). Cold adaptation of eicosapentaenoic acid-less mutant of Shewanella livingstonensis Ac 10 173 involving uptake and remodeling of synthetic phospholipids containing various polyunsaturated fatty acids. Extremophiles. Satomi, M., Oikawa, H. & Yano, Y. (2003). Shewanella marinintestina sp. nov., Shewanella schlegeliana sp. nov. and Shewanella sairae sp. nov., novel eicosapentaenoic- acid-producing marine bacteria isolated from sea-animal intestines. Int J Syst Evol Microbiol 53, 491-499. Satomi, M., Vogel, B. F., Gram, L. & Venkateswaran, K. (2006). Shewanella hafniensis sp. nov. and Shewanella morhuae sp. nov., isolated from marine fish of the Baltic Sea. Int J Syst Evol Microbiol 56, 243-249. Sayanova, O. V. & Napier, J. A. (2004). Eicosapentaenoic acid: biosynthetic routes and the potential for synthesis in transgenic plants. Phytochemistry 65, 147-158. Schibler, L., Vaiman, D., Oustry, A., Guinec, N., Dangy-Caye, A. L., Billault, A. & Cribiu, E. P. (1998). Construction and extensive characterization of a goat bacterial artificial chromosome library with threefold genome coverage. Mamm Genome 9, 119- 124. Sellmayer, A. & Koletzko, B. (1999). Long-chain polyunsaturated fatty acids and eicosanoids in infants—physiological and pathophysiological aspects and open questions. Lipids 34, 199-205. 174 Semplicini, A. & Valle, R. (1994). Fish oils and their possible role in the treatment of cardiovascular diseases. Pharmacol Ther 61, 385-397. Shewan, J., Hobbs, G. & Hodgkiss, W. (1960). A determinative scheme for the identification of certain genera of gram-negative bacteria, with special reference to the Pseudomonadaceae. J Appl Bacteriol 23, 379-390. Siddiqui, R. A., Shaikh, S. R., Sech, L. A., Yount, H. R., Stillwell, W. & Zaloga, G. P. (2004). Omega 3-fatty acids: health benefits and cellular mechanisms of action. Mini Rev Med Chem 4, 859-871. Siddiqui, R. A., Harvey, K. & Stillwell, W. (2008a). Anticancer properties of oxidation products of docosahexaenoic acid. Chem Phys Lipids 153,47-56. Siddiqui, R. A., Harvey, K. A. & Zaloga, G. P. (2008b). Modulation of enzymatic activities by n-3 polyunsaturated fatty acids to support cardiovascular health. / Nutr Biochem 19,417-437. Sijtsma, L. & de Swaaf, M. E. (2004). Biotechnological production and applications of the omega-3 polyunsaturated fatty acid docosahexaenoic acid. Appl Microbiol Biotechnol 64, 146-153. 175 Simopoulos, A. P. (2000). Human requirement for N-3 polyunsaturated fatty acids. Poult Sci 79, 961-970. Simopoulos, A. P. (2002). Omega-3 fatty acids in inflammation and autoimmune diseases. J Am Coll Nutr 21, 495-505. Singh, M. (2005). Essential fatty acids, DHA and human brain. Indian J Pediatr 72, 239- 242. Skerratt, J. H., Bowman, J. P. & Nichols, P. D. (2002). Shewanella olleyana sp. nov., a marine species isolated from a temperate estuary which produces high levels of polyunsaturated fatty acids. Int J Syst Evol Microbiol 52, 2101-2106. Smailus, D. E., Warren, R. L. & Holt, R. A. (2007). Constructing large DNA segments by iterative clone recombination. Syst Synth Biol 1, 139-144. Song, J., Dong, F. & Jiang, J. (2000). Construction of a bacterial artificial chromosome (BAC) library for potato molecular cytogenetics research. Genome 43, 199-204. Sparwasser, T. & Eberl, G. (2007). BAC to immunology-bacterial artificial chromosome-mediated transgenesis for targeting of immune cells. Immunology 121, SOS- 176 Stark, A. H., Crawford, M. A. & Reifen, R. (2008). Update on alpha-linolenic acid. NutrRev 66, 326-332. Stark, K. D., Park, E. J., Maines, V. A. & Holub, B. J. (2000). Effect of a fish-oil concentrate on serum lipids in postmenopausal women receiving and not receiving hormone replacement therapy in a placebo-controlled, double-blind trial. Am J Clin Nutr 72, 389-394. Steyn, N. P., Mann, J., Bennett, P. H., Temple, N., Zimmet, P., Tuomilehto, J., Lindstrom, J. & Louheranta, A. (2004). Diet, nutrition and the prevention of type 2 diabetes. Public Health Nutr 7, 147-165. Stoll, B. A. (2002). N-3 fatty acids and lipid peroxidation in breast cancer inhibition. Br J Nutr 87, 193-198. Strong, S. J., Ohta, Y., Litman, G. W. & Amemiya, C. T. (1997). Marked improvement of PAC and BAC cloning is achieved using electroelution of pulsed-field gel-separated partial digests of genomic DNA. Nucleic Acids Res 25, 3959-3961. Sugihara, S., Orikasa, Y. & Okuyama, H. (2008). An EntD-like phosphopantetheinyl transferase gene from Photobacterium profundum SS9 complements pfa genes of Moritella marina strain MP-1 involved in biosynthesis of docosahexaenoic acid. Biotechnol Lett 30, 411-414. 177 Suzuki, M. T., Preston, C. M., Beja, O., de la Torre, J. R., Steward, G. F. & DeLong, E. F. (2004). Phylogenetic screening of ribosomal RNA gene-containing clones in Bacterial Artificial Chromosome (BAC) libraries from different depths in Monterey Bay. Microb Ecol 48, 473-488. Takeyama, H., Takeda, D., Yazawa, K., Yamada, A. & Matsunaga, T. (1997). Expression of the eicosapentaenoic acid synthesis gene cluster from Shewanella sp. in a transgenic marine cyanobacterium, Synechococcus sp. Microbiology 143 ( Pt 8), 2725- 2731. Tanaka, M., Ueno, A., Kawasaki, K., Yumoto, I., Ohgiya, S., Hoshino, T., Ishizaki, K., Okuyama, H. & Morita, N. (1999). Isolation of clustered genes that are notably homologous to the eicosapentaenoic acid bisynthesis gene cluster from the docosahexaenoic acid-producing bacterium Vibrio marinus strain MP-1. Biotechnol Lett 21, 939-945. Thomas, N. R., Koshy, S., Simsek, M. & Abraham, A. K. (1988). A precaution when preparing very large plasmids by alkaline lysis procedure. Biotechnol Appl Biochem 10, 402-407. 178 Tonon, T., Harvey, D., Qing, R., Li, Y., Larson, T. R. & Graham, I. A. (2004). Identification of a fatty acid Deltall-desaturase from the microalga Thalassiosira pseudonana. FEBS Lett 563, 28-34. Valentine, R. C. & Valentine, D. L. (2004). Omega-3 fatty acids in cellular membranes: a unified concept. Prog Lipid Res 43, 383-402. Vedin, I., Cederholm, T., Freund Levi, Y. & other authors (2008). Effects of docosahexaenoic acid-rich n-3 fatty acid supplementation on cytokine release from blood mononuclear leukocytes: the OmegAD study. Am J Clin Nutr 87, 1616-1622. Venkateswaran, K., Moser, D. P., Dollhopf, M. E. & other authors (1999). Polyphasic taxonomy of the genus Shewanella and description of Shewanella oneidensis sp. nov. Int J Syst Bacteriol 49 Pt 2, 705-724. Villalobos, D. P., Bautista, R., Canovas, M. & Claros, M. G. (2004). Isolation of bacerial Artificial Chromosome DNA by means of improved alkaline lysis and double potassium acetate precipitation. Plant Molecular Biology Reporter 22,419-425. Vogel, B., Venkateswaran, K., Satomi, M. & Gram, L. (2005). Identification of Shewanella baltica as the most important H2S-producing species during iced storage of Danish marine fish. Appl Environ Microbiol 71, 6689-6697. 179 Voss, A. C, Maki, K. C, Garvey, W. T., Hustead, D. S., Alish, C, Fix, B. & Mustad, V. A. (2008). Effect of two carbohydrate-modified tube-feeding formulas on metabolic responses in patients with type 2 diabetes. Nutrition 24, 990-997. Wada, M., DeLong, C. J., Hong, Y. H. & other authors (2007). Enzymes and receptors of prostaglandin pathways with arachidonic acid-derived versus eicosapentaenoic acid-derived substrates and products. J Biol Chem 282, 22254-22266. Walter, J., Mangold, M. & Tannock, G. W. (2005). Construction, analysis, and beta- glucanase screening of a bacterial artificial chromosome library from the large-bowel microbiota of mice. Appl Environ Microbiol 71, 2347-2354. Wang, F., Wang, J., Jian, H. & other authors (2008). Environmental adaptation: genomic analysis of the piezotolerant and psychrotolerant deep-sea iron reducing bacterium Shewanella piezotolerans WP3. PLoS ONE 3, el937. Weiner, R. M., Coyne, V., Brayton, P., West, P. & Raiken, S. F. (1988). Alteromonas colwelliana sp. nov., an isolate from oyster habitates. Int J Syst Bacteriol 38, 240-244. Wen, Z. Y. & Chen, F. (2003). Heterotrophic production of eicosapentaenoic acid by microalgae. Biotechnol Adv 21, 273-294. 180 Wild, J., Hradecna, Z. & Szybalski, W. (2002). Conditionally amplifiable BACs: switching from single-copy to high-copy vectors and genomic clones. Genome Res 12, 1434-1444. Wilkinson, S. G. & Caudwell, P. F. (1980). Lipid composition and chemotaxonomy of Pseudomonas putrefaciens (Alteromonas putrefaciens). J Gen Microbiol 118, 329-341. Wirsen, C. O., Jannasch, H. W., Wakeham, S. G. & Canuel, E. A. (1987). Membrane lipids of a psychrophilic and baropilic deep-sea bacterium. Curr Microbiol 14, 319-322. Wohl, D. A. (2007). Editorial comment: fish oil to keep the cardiologist away? AIDS Read 17, 371. Woodman, R. J., Mori, T. A., Burke, V., Puddey, I. B., Watts, G. F. & Beilin, L. J. (2002). Effects of purified eicosapentaenoic and docosahexaenoic acids on glycemic control, blood pressure, and serum lipids in type 2 diabetic patients with treated hypertension. Am J Clin Nutr 76, 1007-1015. Yakimov, M. M., Giuliano, L., Gentile, G., Crisafi, E., Chernikova, T. N., Abraham, W. R., Lunsdorf, H., Timmis, K. N. & Golyshin, P. N. (2003). Oleispira antarctica gen. nov., sp. nov., a novel hydrocarbonoclastic marine bacterium isolated from Antarctic coastal sea water. Int J Syst Evol Microbiol 53, 779-785. 181 Yamagishi, K., Iso, H., Date, C, Fukui, M., Wakai, K., Kikuchi, S., Inaba, Y., Tanabe, N. & Tamakoshi, A. (2008). Fish, omega-3 polyunsaturated fatty acids, and mortality from cardiovascular diseases in a nationwide community-based cohort of Japanese men and women the JACC (Japan Collaborative Cohort Study for Evaluation of Cancer Risk) Study. J Am Coll Cardiol 52, 988-996. Yano, Y., Nakayama, A., Saito, H. & Ishihara, K. (1994). Production of docosahexaenoic acid by marine bacteria isolated from deep sea fish. Lipids 29, 527-528. Yazawa, K., Araki, K., Okazaki, N., Watanabe, K., Ishikawa, C, Inoue, A., Numao, N. & Kondo, K. (1988). Production of eicosapentaenoic acid by marine bacteria. J Biochem 103, 5-7. Yazawa, K. (1996). Production of eicosapentaenoic acid from marine bacteria. Lipids 31 Suppl, S297-300. Yu, R., Yamada, A., Watanabe, K., Yazawa, K., Takeyama, H., Matsunaga, T. & Kurane, R. (2000). Production of eicosapentaenoic acid by a recombinant marine cyanobacterium, Synechococcus sp. Lipids 35, 1061-1064. Zhang, H., Zhao, X., Ding, X., Paterson, A. & Wing, R. (1995). Preparation of megabase-size DNA from plant nuclei. Plant J 7, 175-184. 182 Zhou, X. R., Robert, S. S., Petrie, J. R., Frampton, D. M., Mansour, M. P., Blackburn, S. I., Nichols, P. D., Green, A. G. & Singh, S. P. (2007). Isolation and characterization of genes from the marine microalga Pavlova salina encoding three front- end desaturases involved in docosahexaenoic acid biosynthesis. Phytochemistry 68, 785- 796. Ziemke, F., Hofle, M. G., Lalucat, J. & Rossello-Mora, R. (1998). Reclassification of Shewanella putrefaciens Owen's genomic group II as Shewanella baltica sp. nov. Int J Syst Bacteriol 48 Pt 1, 179-186. 183 Appendix A Preparation of maleic acid, washing buffer and detection buffer. Maleic Acid Buffer: Mix 0.1 M maleic acid with 0.15 M NaCl and adjust to pH=7.5 and then autoclave at 1210Cforl5min. Washing Buffer: Mix 0.1 M maleic acid with 0.15 M NaCl adjust the pH to 7.5 and then add 0.3% (v/v) Tween 20. Detection Buffer: Mix 0.1 M Tris-HCL with 0.1 M NaCl and adjust the pH to 9.5 184 Appendix B Preparation of Denaturation and Neutralization solutions. Denaturation solution: Add 87.75 gram NaCl (1.5 M) and 50 ml NaOH (0.5 N) to 1000 ml autoclaved water, mix and store in a clean autoclaved bottle. Neutralization solution: Add 175.5 gram NaCl (3 M) and 250 ml Tris-HCL (0.5 M) to 1000 ml autoclaved water, mix and adjust the pH to 7.5. 185 Appendix C Blast search for amplified fragments with EPA5-4 (pfaA), EPA 7-2 (pfaC), EPA 8-2 (pfaD) and EPA2-1 primers. pfaA Shewanella sp. SCRC-2738 eicosapentaenoic acid (EPA) synthesis gene cluster, complete sequence Length=37895 alignments for this subject sequence by: Score = 848 bits (940), Expect = 0.0 Identities = 768/966 (79%), Gaps = 1/966 (0%) Strand=Plus/Plus Query 26 GTATTCNC-ANCAGCGGCCTGAGCGATGAAGACGGC^TAATGCTGATCAAAAAGTTCCAA 84 MINI ! I MUM I M M Ml M MMM MMI M MMM Sbjct 14416 GTATTCGCCAATAGCGGCATTAGTGACACCGACAGCGAAATGCTTATCAAGAAATTCCAA 14475 Query 85 GACCAATATATCCACTGGGAAGAAAACTCTTTCCCAGGGTCCTTAGGCAACGTGATTGCA 144 MMMMI I MMMMMMMMI MMMM M I M MMI Mill Sbjct 14476 GACCAATATGTACACTGGGAAGAAAACTCGTTCCCAGGTTCACTTGGTAACGTTATTGCG 14535 Query 145 AGCCGTATCGCCAACCGTTTTGATTTTGGCGGCATGAACTGTGTGGTCGATGCTGCCTGC 204 llllllllllllllll II I I lllllll I I I I I I Ml I INI I II MIMIIMM Sbjct 14536 GGCCGTATCGCCAACCGCTTCGATTTTGGCGGCATGAACTGTGTGGTTGATGCTGCCTGT 14595 Query 205 GCGGGCTCACTTGCCGCTATGCGTATGGCGTTGACTGAACTGACCGAAGGTCGCAGCGAC 264 II II IIMIMI MMMMIIIMM I II II II II MMMMI M Sbjct 14596 GCTGGATCACTTGCTGCTATGCGTATGGCGCTAACAGAGCTAACTGAAGGTCGCTCTGAA 14655 Query 265 ATGATGATCACCGGCGGCGTCTGTACCGACAACTCGCCGTCCATGTATATGAGCTTCTCA 324 MMMMIIMM II II Mill II Mill II M MIMIMMMM III Sbjct 14656 ATGATGATCACCGGTGGTGTGTGTACTGATAACTCACCCTCTATGTATATGAGCTTTTCA 14715 Query 325 AAAACGCCTGCCTTCACTACCAATGAAACCATTCAACCCTTTGATATCGATTCAAAGGGC 384 MMMM Mill II II II MIMIIMM II MIMIIMM Mill III Sbjct 14716 AAAACGCCCGCCTTTACCACTAACGAAACCATTCAGCCATTTGATATCGACTCAAAAGGC 14775 Query 385 ATGATGATCGGCGAAGGTATTGGCATGGTAGCACTTAAGCGCCTTGAAGATGCCGAGCGC 444 MMMM II MMMMMMMMI II II Mill MIMIIMM MMM Sbjct 14776 ATGATGATTGGTGAAGGTATTGGCATGGTGGCGCTAAAGCGTCTTGAAGATGCAGAGCGC 14835 Query 445 GATGGCGACCGGATTTATGCCGTCATCAAAGGCGTTGGCGCCTCATCGGACGGTAAATTT 504 MMMM! Mill I II II Mill II II II Mill MMMM III Sbjct 14836 GATGGCGACCGCATTTACTCTGTAATTAAAGGTGTGGGTGCATCATCTGACGGTAAGTTT 14895 Query 505 AAGAGTATTTATGCGCCGCGCCCTGAAGGCCAAGCTAAAGCATTGGAGCGCGCCTACGAC 564 II II Mill II Mill llllllllllllllll I I M Mill II Sbjct 14896 AAATCAATCTATGCCCCTCGCCCATCAGGCCAAGCTAAAGCACTTAACCGTGCCTATGAT 14955 Query 565 GACGCGGGTTTTGCCCCGCACAGCATTGGCTTAGTTGAAGCCCATGGCACGGGCACTGCC 624 Mill MMMM Mill I I I II II lllllll II II II II Mill Sbjct 14956 GACGCAGGTTTTGCGCCGCATACCTTAGGTCTAATTGAAGCTCACGGAACAGGTACTGCA 15015 Query 625 GCAGGTGATGTGGCCGAATTCAATGGCTTAAAATCGGTATTTGCCCAAGGCAACGACACC 684 186 Sbjct 15016 GCAGGTGACGCGGCAGAGTTTGCCGGCCTTTGCTCAGTATTTGCTGAAGGCAACGATACC 15075 Query 685 AATCAACATATCGCGTTAGGTTCAGTGAAATCCCAAGTGGGCCACACTAAATCAACCGCA 744 II lllll II III MINIMI Mill III I II II MMIIIMI III Sbjct 15076 AAGCAACACATTGCGCTAGGTTCAGTTAAATCACAAATTGGTCATACTAAATCAACTGCA 15135 Query 745 GGTACTGCTGGGGTAATCAAAGCCGCGCTGGCGCTGCACCACAAGGTATTACCTGCGACC 804 lllll II II MM lllll II II II MM lllllllll I II Mill Sbjct 15136 GGTACAGCAGGTTTAATTAAAGCTGCTCTTGCTTTGCATCACAAGGTACTGCCGCCGACC 15195 Query 805 ATTAACGTCAGCAAGCCTAATCCAAAACTGAATATCGAAAGCTCACCATTCTATTTAAAT 864 llllllll II MM I II lllll IMMIIM MUM II III MM Sbjct 15196 ATTAACGTTAGTCAGCCAAGCCCTAAACTTGATATCGAAAACTCACCGTTTTATCTAAAC 15255 Query 865 ACCGAAACGCGCCCTTGGCTGCAACGCACTGACGGTACGCCGCGCCGTGCTGGCATAAGT 924 II II II II II III I I III III I ! I I I I I I I I I I 1 I II II II II Sbjct 15256 ACTGAGACTCGTCCATGGTTACCACGTGTTGATGGTACGCCGCGCCGCGCGGGTATTAGC 15315 Query 925 TCCTTTGGTTTTGGCGGCACTAACTTCCATCTCGTATTAGAAGAATACAAACCCGAGCAC 984 II 11111111111 IMIIIIIMM! I III Mlllll lllll I II III Sbjct 15316 TCATTTGGTTTTGGTGGCACTAACTTCCATTTTGTACTAGAAGAGTACAACCAAGAACAC 15375 Query 985 AGCCGT 990 MUM Sbjct 15376 AGCCGT 15381 pfaC Shewanella sp. SCRC-2738 eicosapentaenoic acid (EPA) synthesis gene cluster, complete sequence Length=37895 Score = 729 bits (808), Expect = 0.0 Identities = 734/949 (77%), Gaps = 35/949 (3%) Strand=Plus/Plus Query 1 ATGCGACTTGATGCTGATCAGCTACTTAGGTATCGACTTTGAAAACAAGGGCGAGCGCGT 60 III IMMIMMI II lllll I MMMMMIMI lllll llllllll II Sbjct 28273 ATGTGACTTGATGCTTATTAGCTATCTCGGTATCGACTTTGAGAACAAAGGCGAGCGGGT 28332 Query 61 TTATCGCCTGCTCGATTGCACCCTGACCTTCCTTGGCGACTTACCGCGCGGTGGCGACAC 120 llllll II llllllll Mill llllllll llllllll II II II II II II Sbjct 28333 TTATCGACTACTCGATTGTACCCTCACCTTCCTAGGCGACTTGCCACGTGGCGGAGATAC 28392 Query 121 CCTGCGCTACGATATTTCAATCAATCACTTTGCCCGCAATGGCGATACCTTGTTGTTCTT 180 III II Mill III llllll III III lllll lllll III II Mlllll Sbjct 28393 CCTACGTTACGACATTAAGATCAATAACTATGCTCGCAACGGCGACACCCTGCTGTTCTT 28452 Query 181 CTTCTCCTACGAATGTTTCGTGGGCGACAAGCTGATCCTGAAAATGGATGGCGGCTGTGC 240 llllll II II lllll II lllllllll Mlllll II MMMMMIMI II Sbjct 28453 CTTCTCGTATGAGTGTTTTGTTGGCGACAAGATGATCCTCAAGATGGATGGCGGCTGCGC 28512 Query 241 CGGCTTCTTCACCGATAAAGAACTAGCCGACGGCAAAGGCGTTATTCACACCGAAGCCGA 300 IIIIMIMM III MM II IMIIMI llllllll MM III MM II Sbjct 28513 TGGCTTCTTCACTGATGAAGAGCTTGCCGACGGTAAAGGCGTGATTCGCACAGAAGAAGA 28572 Query 301 AATCAAAGCGCGCAACCTCGCCTTGAACAATCCGAATAAGCCGCGCTTTAATCCGTTACT 360 II lllll MM III I II III MM MMIMMMIIMM Sbjct 28573 GATTAAAGCTCGCAGCCTAG TG--CAA- AAGCAACGCTTTAATCCGTTACT 28620 Query 361 CAACTGCGCGCAAAACCAATTTGATTACAGCCAAATCCATAAACTGCTCGGCGCCGATAT 420 I II I III Mlllll III I I II Mill II I II Mill Sbjct 28621 AGATTGTCCTAAAACCCAATTTAGTTATGGTGATATTCATAAGCTATTAACTGCTGATAT 28680 Query 421 CGGTGGCTGTTTTGGCG GCGCACACGCGGCGCATCAAGCCCAATATGGTTTGCAGCC 477 187 I II MINN II llll I MM III Mill Sbjct 28681 TGAGGGTTGTTTTGGCCCAAGC-CACA-GTGGCGT-- CCAC CAGCC 28722 Query 478 CTCTTTATGTTTTGCATCTGAAAAATTCCTGATGATTGAACAAGTCAGCAATCTTGAGGT 537 M I Mill lllllllllllllll MMMMMMMMMIMI llll Sbjct 28723 GTCACTTTGTTTCGCATCTGAAAAATTCTTGATGATTGAACAAGTCAGCAAGGTTGA-TC 28781 Query 538 GCA-TGGCGGCGCGTGGGGCTTAGGCTCAGTTCAAGGCCATAAGCAGCTCGAAGCCGATC 596 III Mill I Mill I llll I II I II I I I I I I I I I I I Mill II I Sbjct 28782 GCACTGGCGGTACTTGGGGACTTGGCTTAATTGAGGGTCATAAGCAGCTTGAAGCAGACC 28841 Query 597 ATTGGTATTTCCCGTGTCATTTCAAGGGCGACCAAGTGATGGCGGGGTCGTTAATGGCCG 656 I Mill Mill 11 llilll 11II III 1111111 llllll I II Ml Mill I Sbjct 28842 ACTGGTACTTCCCATGTCATTTCAAGGGCGACCAAGTGATGGCTGGCTCGCTAATGGCTG 28901 Query 657 AAGGCTGTGGTCAATTACTGCAATTCTTTATGCTACATATTGGTATGCACCTCGGTGTTA 716 MM Mill II III llll llll llllll II I! 11111111 I II Sbjct 28902 AAGGTTGTGGCCAGTTATTGCAGTTCTATATGCTGCACCTTGGTATGCATACCCAAACTA 28961 Query 717 AAGATGGTCGTTTCCAACCGCTCGAAAACGCGTCACAAAAAGTGCGTTGTCGCGGTCAAG 776 II MMIMMMMIM II llllll Mill MM II 1111! 1111 i I i I Sbjct 28962 AAAATGGTCGTTTCCAACCTCTTGAAAACGCCTCACAGCAAGTACGCTGTCGCGGTCAAG 29021 Query 777 TGTTGCCGCAATCAGGCCTGCTCACCTATCGTATGGAAATCACTGAAATCGGTATGAGCC 836 II llll IMMIIII llll II II IMMIIII I 11 M 11111 i 1 I II I Sbjct 29022 TGCTGCCACAATCAGGCGTGCTAACTTACCGTATGGAAGTGACTGAAATCGGTTTCAGTC 29081 Query 837 CGCGCCCGTATGCTAAGGCGAATATCGATATTCTGCTCAATGGTAAAGTGGTTGTGGACT 896 I Mill MINIM II II llllllll MM Mill llll III Mill I Sbjct 29082 CACGCCCATATGCTAAAGCTAACATCGATATCTTGCTTAATGGCAAAGCGGTAGTGGATT 29141 Query 897 TCCAAAACCTTGGGGTGATGATCAAAGAAGAAGCCGAATGCACCCGCTA 945 IMMIIIII MIMIIMM Mill llll II II II II II Sbjct 29142 TCCAAAACCTAGGGGTGATGATAAAAGAGGAAGATGAGTGTACTCGTTA 29190 PfaD Shewanella sp. SCRC-2738 eicosapentaenoic acid (EPA) synthesis gene cluster, complete sequence Length=37895 Score = 699 bits (774), Expect = 0.0 Identities = 684/881 (77%), Gaps = 3/881 (0%) Strand=Plus/Plus Query 66 CCCGTCAGCGCCTTTGCGCCCGCCCTTGGCACCCAAAGTTTAGGCGACAGTAATTTTCGC 125 II II II II III I II II I II III llll IMMIIII llllll III Sbjct 30937 CCTGTTAGTGCTTTTACTCCTGCATTAGGTACCGAAAGCCTAGGCGACAATAATTTCCGC 30996 Query 126 CGCGTACACGGGGTTAAATACGCTTACTACGCTGGCGCTATGGCTAACGGTATTGCCTCA 185 Mill Mill llllllllllllll Mill IMMIIIII IMMIIII I II Sbjct 30997 CGCGTTCACGGCGTTAAATACGCTTATTACGCAGGCGCTATGGCAAACGGTATTTCATCT 31056 Query 186 GAAGAACTGGTTATCGCGCTGGGCCAAGCGGGCATTTTGTGT TCGTTTGGCGCGGCG 242 Mill II II II II II II Mill 111111111111 MIMIII II II Sbjct 31057 GAAGAGCTAGTGATTGCCCTAGGTCAAGCTGGCATTTTGTGTGGTTCGTTTGGAGCAGCC 31116 Query 243 GGGTTAATCCCATCCCGCGTTGAAGCGGCCATTACTCGCATTCAAGCGGCGCTGCCTAAT 302 II I II III I I I I I I ! I I I I I I I llll II llllllll llllllll III Sbjct 31117 GGTCTTATTCCAAGTCGCGTTGAAGCGGCAATTAACCGTATTCAAGCAGCGCTGCCAAAT 31176 Query 303 GGTCCTTACGCCTTTAATTTAATTCACAGCCCAAGCGAGCCCGCATTAGAGCGCGGCAGT 362 188 II Mill Mill I II II II II MINIM i i I j 1111111 Mill Sbjct 31177 GGCCCTTATATGTTTAACCTTATCCATAGTCCTAGCGAGCCAGCATTAGAGCGTGGCAGC 31236 Query 363 GTTGAGTTGTTCTTAAAACATAAAGTGCGCACGGTCGAAGCCTCGGCATTTTTAGGTTTA 422 II III I II Mill Mill II Mill II Mill II II II MUM II Sbjct 31237 GTAGAGCTATTTTTAAAGCATAAGGTACGCACCGTTGAAGCATCAGCTTTCTTAGGTCTA 31296 Query 423 ACGCCACAAATCGTCTATTACCGCGCAGCAGGTTTGAGCCGCGACGCACATGGCGACATC 482 II MIIMMMIMMIMM llllllll llllllll IMIIMI II I I Sbjct 31297 ACACCACAAATCGTCTATTACCGTGCAGCAGGATTGAGCCGAGACGCACAAGGTAAAGTT 31356 Query 4 83 GTCATTGGCAACAAAGTCATAGCCAAAATCAGTCGCACCGAAGTCGCGACTAAGTTTATG 542 II MM Mill II II II III I 11111111111 M I II IMIIMM Sbjct 31357 GTGGTTGGTAACAAGGTTATCGCTAAAGTAAGTCGCACCGAAGTGGCTGAAAAGTTTATG 31416 Query 543 GAGCCGGCGCCTGCCAAAATTCTGCAGCAATTAGTCAGTGAAGGCCTTATCAGCCAAGAT 602 III Mill II Mill II II II MM III II MM II Sbjct 31417 ATGCCAGCGCCCGCAAAAATGCTACAAAAACTAGTTGATGACGGTTCAATTACCGCTGAG 31476 Query 603 CAAATGGCGATGGCGCAACTTGTACCCATGGCGGACGATATCACGGCCGAAGCCGATTCT 662 MINI I MMIMMMIMM Mill Mill Mill II II llllllll Sbjct 31477 CAAATGGAGCTGGCGCAACTTGTACCTATGGCTGACGACATCACTGCAGAGGCCGATTCA 31536 Query 663 GGCGGCCATACCGACAATCGTCCACTGGTCACGCTATTGCCGACGATTTTGGCGCTCAAA 722 II llllllll II II llllll I II II I MM II Mill Mill Ml Sbjct 31537 GGTGGCCATACTGATAACCGTCCATTAGTAACATTGCTGCCAACCATTTTAGCGCTGAAA 31596 Query 723 GATGAAATCCAAGCTAAGTATCAATACAAGACGCCCATCCGTGTGGGAGCAGGCGGCGGC 782 II Mill llllllll II llllll I II II II Mill II II Mill Sbjct 31597 GAAGAAATTCAAGCTAAATACCAATACGACACTCCTATTCGTGTCGGTTGTGGTGGCGGT 31656 Query 783 GTTGGTACCCCCGACGCAGCATTAGCCACCTTCAACATGGGCGCGGCCTTTATCGTCACA 842 II Mill II II Mill I II II II MMIMMIMM I III II II Sbjct 31657 GTGGGTACGCCTGATGCAGCGCTGGCAACGTTTAACATGGGCGCGGCGTATATTGTTACC 31716 pfaQuerEy 843 GGTTCAATCAACCAAGCGTGTGTGGAGGCGGGCGCGAGCGAACACACACGTAAGTTACTC 902 II II IMIIMM Mill II llllllll II II Mill Mill Mill ShewanellSbjct 3171a7 balticGGCTCTATCAACCAAGCTTGTGTTGAAGCGGGCGCAAGTGATCACACTCGTAAATTACTa OS185, complete genome T 31776 Length=522968Query 903 GCCACCACAGAAATGGCCGATGTGACTATGGCACCCGCCG6 C 943 IIIIIIII iiiiiiiiiiiiiiiiiiiiimii ii ii FeatureSbjct 3177s 7i n GCCACCACTGAAATGGCCGATGTGACTATGGCACCAGCTGthis part of subject sequence: C 31817 Score = 572 bits (634), Expect = 3e-160 Identities = 370/405 (91%), Gaps = 2/405 (0%) Strand=Plus/Minus Query 8 AGCTCANANCGAACGTCTTGCATTCCTTTGATATTTTGCTGCTA-GGGATTACTTGNTCG 66 llllll I Mill MMMMMMMIMMIMM I I Mill Mill III Sbjct 3008766 AGCTCAGAGCGAACATCTTGCATTCCTTTGATATTTTGCAGTAAAGGGATCACTTGTTCG 3008707 Query 67 CTTANNTTAATCACCTCNGAGGTTGAGCGACCTGTGAGCGTCACGCGCACACCAGAGTTG 126 MM Mill Ml MMIMMMMMMMMMIMMMMIMMIMM Sbjct 3008706 CTTAAGTTAATCAGCTCAGAGGTTGAGCGACCTGTGAGCGTCACGCGCACACCAGAGTTG 3008647 Query 127 TCATTACCCCCCCCAACTTGCGGTTTGGCGATTGAATACTTAAGGAACCCGCTGCGGATT 186 Mill MM MM III 111111118 M9 11 IMIIMM MM 11II MM 1111 Sbjct 3008646 TCATTGCCCCAACCAAATTGTGGTTTGGCGATCGAATACTTAGGGAAGCCGCTGCGGATT 3008587 Query 187 TTCTTTTTCAACTCATCCAGTGGCATAGGGAAGTCTTTCTTCGCCCATATCNCTGATGAA 246 lllllllllllllllll MINIM llllllllllll I Ml I MINI Sbjct 3008586 TTCTTTTTCAACTCATCCAATGGCATAGGTAAGTCTTTCTTCAGCAATATCACCGATGAA 3008527 Query 247 CCATCGTCGGATGAGTAGTAGCTGTAGACTGAATCAATATGAAATTCCTCTTTATTCTTA 306 IMMMMMMIMMMMMMM 111111111111111111111111111111 Sbjct 3008526 CCATCGTCGGATGAGTAGTAGCTGTAGACAGAATCAATATGAAATTCCTCTTTATTCTTA 3008467 Query 307 TAAAGATACTCTTCCTTTTGATTGACCATTGCCTCANTGACATTGAGATTATGGCGACCT 366 111111111111 f 11 111111! M ! 11111111 M llllllllllllllll MM Sbjct 3008466 TAAAGATACTCTTCCATTTGATTGACCATTGCCTCAGTGACATTGAGATTATGCCGACCT 3008407 Query 367 TCCACTTGATAGTTGATGTAAATCCGCTCCTTGCT-TGCCTGACT 410 > I I I I I I I I I I M I I I I I I I I I I ! I I 1 I I I I I I M || Ml II Sbjct 3008406 TCCACTTGATAGTTGATGTAAATCCGCTCCTTGCTCTGGCTGTCT 3008362 190