Investigation of the Role of the Extracellular B-Agarase, Produced

The copyright of this thesis vests in the author. No quotation from it or information derived from it is to be published without full acknowledgementTown of the source. The thesis is to be used for private study or non- commercial research purposes only. Cape Published by the University ofof Cape Town (UCT) in terms of the non-exclusive license granted to UCT by the author.

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INVESTIGATION OF THE ROLE OF THE EXTRACELLULAR -AGARASE, PRODUCED BY THE BACTERIAL EPIPHYTE PSEUDOALTEROMONAS SP. LS2I, IN THE VIRULENCE RESPONSE TOWARDS THE AGAROPHYTE GRACILARIA GRACILIS.

Carin Gildenhuys

Town

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University A thesis submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of Molecular and Cell Biology, Faculty of Science, University of Cape Town, South Africa.

Cape Town February 2008 2

ACKNOWLEDGEMENTS

I would like to thank my supervisor Professor Vernon Coyne for his excellent guidance and support which he provided throughout the duration of this project, as well as for proof- reading my thesis.

Thank you to all my lab mates over the years for your friendship, advice and encouragement. I really enjoyed working with you.

Thanks to Di James for sequencing my DNA samples and to Mo Jaffer for his patience, expert advice and service rendered with the electron microscopy work.

To all the academic staff, students and departmental assistants, thank you for your assistance throughout this project. Town Thank you to my family and friends for your love and support and to Patricia for helping me with my two children while I was writing this thesis. Cape Special thanks go to my husband Anton, for providingof constant encouragement and support throughout this endeavor. Without your help it would not have been possible for me to complete this.

I am grateful to God for giving me the strength and ability to do this.

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TABLE OF CONTENTS

Abstract II

Abreviations V

CHAPTER 1 General introduction 1

CHAPTER 2 Cloning and sequencing of the agarase gene from 37 Pseudoalteromonas sp. LS2i

CHAPTER 3 Purification and characterization of the agarase protein expressed 66 from the recombinant plasmid pEB3 in E. coli

CHAPTER 4 Isolation and characterization of Pseudoalteromonas sp. LS2i 86 mutants with reduced agarolytic activity

CHAPTER 5 Investigation of the role of the extracellular agaraseTown of 107 Pseudoalteromonas sp. LS2i on virulence in the seaweed Gracilaria gracilis

CHAPTER 6 General Discussion Cape 132

APPENDIX A Media and Solutions of 136

APPENDIX B Standard methods 156

Literature cited 167 University

1. CHAPTER 1 ...... 1 2. CHAPTER 2 ...... 37 3. CHAPTER 3 ...... 66 4. CHAPTER 4 ...... 86 5. CHAPTER 5 ...... 108 6. CHAPTER 6 ...... 136 7. APPENDIX A ...... 140 8. APPENDIX B ...... 160 9. LITERATURE CITED ...... 171

INVESTIGATION OF THE ROLE OF THE EXTRACELLULAR -AGARASE, PRODUCED BY THE BACTERIAL EPIPHYTE PSEUDOALTEROMONAS SP. LS2I, IN THE VIRULENCE RESPONSE TOWARDS THE AGAROPHYTE GRACILARIA GRACILIS.

Carin Gildenhuys February, 2008

Department of Molecular and Cell Biology, University of Cape Town, Private Bag, Rondebosch, 2201, South Africa

ABSTRACT

Gracilaria gracilis that grows naturally at Saldanha Bay, South Africa is economically important as a source of agar. The Gracilaria yields from natural beds at Saldanha Bay are however unreliable, and consequently the South African GracilariaTown industry has experienced a number of setbacks over the years. The only way a consistent supply can be assured is by mariculture to supplement the natural harvests. In 1993 the Seaweed Research Institute (SRU) found that mariculture of G. gracilis in SaldanhaCape Bay is feasible but that there is potential to improve yields by technical researchof and development (Anderson et al.1996a). Jaffray and Coyne (1996) developed a pathogenicity assay that demonstrated that agarolytic bacteria isolated from Saldanha Bay Gracilaria induced disease symptoms such as thallus bleaching, while non-agarolytic isolates did not. It is thought that unfavorable environmental conditions such as elevated water temperature and nutrient depletion, which occur during the summer months in the surface layers of the water column in Saldanha Bay, induce the onset of agarase productionUniversity or result in changes in the bacterial community structure in which agarase-producers become more dominant.

By using the pathogenicity assay, Jaffray and Coyne (1996) identified the highly agarolytic Gracilaria gracilis pathogen, Pseudoalteromonas sp. LS2i. The aim of this study was to characterize the bacterial pathogen, Pseudoalteromonas sp. LS2i to further our understanding of virulence regulation and specifically, the role of the agarase enzymes in the process of seaweed-pathogen interaction.

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Two agarolytic clones, pEB1 and pJB1, were obtained after constructing and screening a Pseudoalteromonas sp. LS2i genomic library in Esherichia coli. Restriction enzyme mapping suggested that both clones contain the same agarase gene. Southern hybridization studies confirmed the origin of the cloned DNA and sequencing studies revealed the 1062 bp ORF, putative promoter region, putative ribosome binding site and putative transcriptional start point of the cloned agarase gene. The ORF showed sequence identity to several other β- agarases and was identified as a member of the GH-16 family of glycoside hydrolases.

The agarase was purified from the E. coli JM109 (pEB3) transformant. The molecular weight was estimated to be 39 kDa by SDS-PAGE. Zymogram analysis confirmed that the purified protein is agarolytic and TLC analysis revealed that the predominant end-products of agar hydrolysis are neoagarohexaose and neoagarobiose, which indicates the same mode of action as that observed for the agarase produced extracellularly by Pseudoalteromonas sp. LS2i. Town Sixteen Pseudoalteromonas sp. LS2i mutants with reduced extracellular agarolytic activity were isolated after chemical mutagenesis. Two of these mutants (R9 and R16), which had similar growth rates to strain LS2i, were selected forCape further study. Strain LS2i exhibited an extracellular agarase activity after 2 h of incubation, while no agarolytic activity was detected for either R9 or R16 over a 24 h growth periodof when a ferrecyanide assay was used to detect agarolytic activity. TLC and zymogram analysis did however show that while significantly less active than the wild type agarase, strain R9 does produce an active extracellular agarase with the same mode of action and identical size to the agarase produced by Pseudoalteromonas sp. LS2i. No active extracellular agarase was detected in mutant R16 using either TLC analysisUniversity or zymogram detection.

The role played by the extracellular agarase produced by Pseudoalteromonas sp. LS2i during G. gracilis infection was investigated. Firstly, mutants R9 and R16 exhibited reduced virulence in G. gracilis compared to the wild type strain or purified agarase protein. Secondly, histology studies indicated that thallus bleaching caused by the agarase leads to weakening of the cell structure. Finally, immuno-gold labeled antibodies showed that the agarase was localized to the cell wall. Enumeration of the gold particles showed a 44% reduction in the number of gold particles located in the cell walls of thalli infected with mutant R16 and an 83% reduction of gold particles in the cell walls of thalli infected with IV mutant R9 in comprisson to LS2i-infected thalli. Western hybridization confirmed that the antibodies raised against the β-agarase purified from E. coli JM109 (pEB3) are specific to the 38 kDa agarase expressed from E. coli JM109 (pEB3). In this study evidence is thus provided that the extracellular agarase produced by an agarolytic bacterium causes disease symptoms in the seaweed Gracilaria gracilis and that the agarase can be located in the cell wall of the infected seaweed.

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ABBREVIATIONS

α alpha β beta κ kappa ug microgram ul microlitre um micrometer(s) ˚E degrees east ˚S degrees south ˚C degrees Celsius % percentage US$ United States dollars Town A adenine

A600 absorbance at 600 nm Cape BMB basal marine broth of bp base pairs BSA bovine serum albumin c cell C cytosine cm University centimetre(s) CTAB cetyltrimethylammonium bromide cw cell wall

DNA deoxyribonucleic acid Da Dalton(s)

EDTA ethylenediaminetetra-acetic acid EM electron microscope et al. and others (et alii) VI

ETOH ethanol g gram(s) G guanine Glu glutamic acid h hour(s) k kilo K thousand kb kilobase(s) kDa kilodalton(s) kg kilogram(s) km kilometer(s) Town l litre(s) LA Luria agar Cape LB Luria broth of m metre(s) M molar mA milli-Amperes MA marine agar MB University marine broth MCS multiple cloning site mg milligram min minute(s) ml milliliter mM millimolar mm millimeter mol mole(s) MOPS (3-[N-morpholino]propane-sulfonic acid)

VII ng nanogram(s) NNS nine salts solution nm nanometer(s) NTG nitrosoguanidine

O/N overnight OD optical density ORF open reading frame p plasmid PAGE polyacrylamide gel electrophoresis PBS Phosphate buffered saline PCR polymerase chain reaction PIPES piperazine-N-N’-bis(2-ethanesulfonic acid)Town rDNA ribosomal DNA RNA ribonucleic acid Cape RNAse ribonuclease rpm revolutions per minuteof rRNA ribosomal RNA RT room temperature s second(s) SDS University sodium dodecyl sulphate SE standard error sp. species SRU Seaweed Research Unit of Marine and Coastal Management SSC sodium chloride tri-sodium citrate buffer SSW sterile seawater STE sodium chloride tris-EDTA buffer

T thymine TAE Tris-acetate-EDTA buffer VIII

TCA Trichloroacetic acid TE Tris-EDTA buffer TEMED N,N,N’,N’-tetramethylethylenediamine TLC thin-layer chromatography Tris tris(hydroxymethyl)aminomethane

U unit(s) UV ultraviolet

V volt(s) v volume

W Watt(s) w weight Town

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1. CHAPTER 1

GENERAL INTRODUCTION

CONTENTS

1.1. Gracilaria ...... 3 1.1.1. The taxonomic position...... 3 1.1.2. The life history ...... 4 1.1.3. The structure of agar ...... 5 1.1.4. The importance of Gracilaria as a source of agar ...... 6 1.1.5. Applications of agar obtained from Gracilaria ...... 6 1.1.6. Other applications of Gracilaria ...... 7 1.1.7. The economic importance of Gracilaria ...... 8 1.2. Gracilaria gracilis at Saldanha Bay, South Africa ...... Town ...... 9 1.2.1. Location of Saldanha Bay ...... 9 1.2.2. History of the Gracilaria gracilis population in Saldanha Bay ...... 10 1.2.3. Thermal stratification in Saldanha Bay...... Cape 11 1.2.4. Other relevant studies that were ofconducted in Saldanha bay ...... 12 1.2.5. Bacterial epiphytes of Gracilaria gracilis from Saldanha Bay ...... 13 1.3. Agarases ...... 15 1.3.1. Agarolytic bacteria ...... 15 1.3.2. Applications of agarases ...... 15 1.3.3. Characterized agarases ...... 16 1.3.3.1. PseudoalteromonasUniversity atlantica ...... 16 1.3.3.2. Pseudoalteromonas antarctica ...... 19 1.3.3.3. Pseudomonas sp. W7 ...... 19 1.3.3.4. Pseudomonas sp. SK38 ...... 20 1.3.3.5. Pseudomonas aeruginosa AG LSL-11 ...... 20 1.3.3.6. A Pseudomonas-like bacterium ...... 21 1.3.3.7. Alteromonas agarlyticus strain GJ1B ...... 21 1.3.3.8. Alteromonas sp. strain C-1 ...... 23 1.3.3.9. Alteromonas sp. E-1 ...... 23 2

1.3.3.10. Alteromonas sp. SY37-12 ...... 23 1.3.3.11. Bacillus sp. MK03 ...... 23 1.3.3.12. Vibrio sp. strain JT0107 ...... 24 1.3.3.13. Vibrio sp. AP-2 ...... 26 1.3.3.14. Vibrio sp. PO-303 ...... 26 1.3.3.15. Cytophaga flevensis ...... 27 1.3.3.16. Streptomyces coelicolor A3(2) ...... 27 1.3.3.17. A Microbulbifer – like bacterium, strain JAMB-A94 ...... 28 1.3.3.18. Microbulbifer strain JAMB-A7 ...... 29 1.3.3.19. Microscillla sp. strain PRE1 ...... 29 1.3.3.20. Agarases from a mixed microbial population ...... 29 1.3.3.21. Thalassomonas strain JAMB-A33 ...... 29 1.3.3.22. Agarivorans sp. JAMB-A11 ...... 30 1.3.3.23. Acinetobacter sp. AG LSL-1 ...... Town 30 1.3.3.24. Saccharophagus degradans 2-40 ...... 30 1.3.3.25. Zobellia galactanivorans Dsij...... 32 1.4. Concluding remarks and aim of this study ...... Cape 35

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1.1. Gracilaria

Seaweeds can be classified into three broad groups based on pigmentation: Brown (Phaeophyceae), red (Rhodophyceae) and green (Chlorophyceae) (McHugh, 2003). Gracilaria gracilis is classified as red seaweed, and has economic importance mainly as a source of food grade agar (Armisen, 1995). Gracilaria is distributed widely, with some species adapted to warm tropical water while others grow in colder water. Natural populations of Gracilaria have been reported to grow along the coast of France (Mediterranean sea), southern coast of Chile, the Atlantic coast of Canada, the southern provinces of China, the southern coast of Thailand as well as along the coasts of Taiwan, Viet Nam, Indonesia, Argentina, Brazil, Luderitz in Namibia and Saldanha Bay in South Africa (McHugh, 2003; Marinho-Soriano et al., 2005).

1.1.1. The taxonomic position Town

Gracilaria verrucosa (Hudson) Papenfuss from the British Isles was reported as the type species of the economically important family of theCape Gracilariaceae (Fredericq and Hommersand, 1989). However in 1995, Steentoftof and co-workers (1995) reclassified both the species G. verrucosa (Hudson) Papenfuss and G. confervoides (Stackhouse) Greville as Gracilaria gracilis (Stackhouse) Steentoft, Irvine et Farnham. Characterization based on morphological characteristics indicated that these species were previously misidentified. The current taxonomic position of Gracilaria gracilis as described by Armisen et al. (1995) is as follows: University Division: Rhodophyta Class: Florideophyceae Order: Gracilariales Family: Gracilariaceae Genus: Gracilaria Species: gracilis

1.1.2. The life history

The study of the genus Gracilaria in the laboratory has revealed a life history of the ‘Polysiphonia-type’ in most species, i.e. an alternation of diploid and haploid generations (Marinho-Soriano et al., 1998). Haploid and diploid individuals are morphologically indistinguishable at the vegetative stage but during the reproductive period individuals are distinguished by their sexual organs (Destombe et al., 1993). In the wild, attached populations of Gracilaria generally exhibit a haploid and a diploid phase, while drifting populations are mostly tetrasporophytic or infertile. Infertile thalli can grow indefinitely and regenerate from fragmentation, which makes Gracilaria one of the most used in mariculture (Marinho-Soriano et al., 1998). Isaac and Molteno (1952) found that the sterile vegetative form of Gracilaria gracilis was mostly observed in Saldanha Bay, South Africa, while only a few fertile forms were reported. Town Contrary to the theoretical expectation of predominantly endogamous mating systems (fertilization from gametes on the same plant) in haploid-diploid organisms, Gracilaria gracilis showed a clearly allogamous mating systemCape (cross-fertilization in plants) (Engel et al., 2004).The life history of Gracilaria is depicted in Figure 1. The female gamete is fertilized in situ and the resulting zygote developsof as a diploid carposporophyte. The latter is a spore-producing structure entirely dependent on the female gametophyte. The resulting carpospores produced by the carposporophyte are numerous and genetically identical. Each spore can develop into a diploid tetrasporophyte in which reproduction involves meiosis in order to produce haploid genetically variable tetraspores. This life history of Gracilaria has been completed in culture,University taking 5-12 months (Kain and Destombe, 1995).

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Figure 1. A diagram of the life history of Gracilaria (Kain and Destombe, 1995) Cape of 1.1.3. The structure of agar

Agars and carrageenans are the main components of the cell wall of red algae (Graigie, 1990). This family of hydrocolloids consists of linear chains of galactose, with disaccharide repeating units of 3-linked β-D-galactose and 4-linked -galactose units. The latter occurs as the L-isomer in agar Universityand the D-isomer in carrageenan (Yaphe, 1984). In the carrageenans the galactose units are substituted in a characteristic manner with sulphate groups. In contrast, the galactose units in agar are non-substituted or substituted with either O-methylether, pyruvic acid, ketal or sulphate groups. These substituent groups can alter in a number of ways the structural regularity of agar (Murano, 1995; Yaphe, 1984). Agar composition varies, depending not only on the particular strain or species, but also on the variations in nutrient levels, light levels and season (Cote and Hanisak, 1986). The quality of agar can be predicted from the type and content of sulphate esters, pyruvate ketals and methyl ethers (Murano, 1995). 6

1.1.4. The importance of Gracilaria as a source of agar

The main use of red seaweeds is as food and as a source of two hydrocolloids, agar and carrageenan. Both agar and carrageenan are water-soluble carbohydrates that are used to thicken aqueous solutions, to form gels of varying degrees of firmness, to form water-soluble films and to stabilize some food products (McHugh, 2003). Seaweeds as a source of these hydrocolloids dates back to 1658 when the gelling properties of agar was discovered in Japan after Minoya Tarazaemon extracted agar for the first time from a red seaweed, using a freeze- thaw method (Armisen, 1995). Today agar is mainly extracted from two genera of red seaweed, Gelidium and Gracilaria. Gelidium gives the higher quality agar but unlike Gracilaria, all the Gelidium used for commercial agar extraction comes from natural resources. Gelidium is a small, slow growing plant and efforts to cultivate it have generally proved to be uneconomic. Gracilaria were however once consideredTown unsuitable for agar production because the gel strength of the agar was to low. However in the 1950s it was discovered that pre-treatment of the seaweed with alkali before extraction lowered the yield but gave a good quality agar (McHugh, 2003). BecauseCape agar from Gracilaria has a higher degree of sulphation, the gelling ability of Gracilariaof agar is considerably improved by an alkali pretreatment with sodium hydroxide that converts -L-galactose-6-sulphate into

3,6-anhydro-- L-galactose (Murano, 1995; Armisen, 1995). Unlike Gelidium, Gracilaria must however be processed within a few months and cannot be allowed to remain in storage for use during years of lower availability (Armisen, 1995). Pre-treatment with alkali allowed expansion of the agar industry and led to the harvesting of a variety of wild species of Gracilaria, but soon Universitythere was evidence of overharvesting of the wild crop and cultivation methods where then developed, both in ponds and in the open waters of protected bays. Today the supply of Gracilaria still comes mainly from natural populations, with the degree of cultivation depending on price fluctuations (McHugh, 2003).

1.1.5. Applications of agar obtained from Gracilaria

Agars obtained from Gracilaria can however not be classified as bacteriological grade agar since these agars have a high content of methoxyls and consequently high gelling 7 temperatures. Gracilaria is however considered to be the most important source of food and sugar-reactive grade agars (Murano, 1995). In contrast to gelatin that melts around 37˚C, agar does not melt until heated to 85˚C or higher. In food applications, this means there is no requirement to keep agar products refrigerated in hot climates (McHugh, 2003).

About 90% of all the agar produced is for food applications. Agar has been classified as GRAS (Generally Recognized As Safe) by the United States of America Food and Drug Administration, which has set maximum usage levels that depends on the application (McHugh, 2003). Gracilaria agar can thus be used for the preservation of canned meats, as a clarifying agent in the manufacturing of wines, beers and coffee and as a covering for pharmaceutical capsules (Shang, 1976). Agar is also used as a stabilizer and thickener in pie fillings, icings and meringues. Some agars can also be used in confectionary, jellies, gelled meat and fish products (McHugh, 2003). Town 1.1.6. Other applications of Gracilaria

Gracilaria consumed as food, is usually gathered andCape sold fresh, locally. This practice is most common in South-East Asian countries such as Indonesia, Malaysia, the Philippines and southern Thailand (McHugh et al., 2003). Seaweedsof that are however used as sea vegetables or ingredients by the food industry in Europe or Japan are usually dried first in mild conditions (Rouxel et al., 2001). Gracilaria verrucosa, also known as Ogo-nori, and Gracilaria changgi are two of the species used for food consumption (Rouxel et al., 2001; Norziah and Ching, 2000). Gracilaria is also quite popular in Hawaii and is sold fresh in Honolulu markets asUniversity ‘limu manauea’ or ‘limo ogo’ (McHugh et al., 2003).

Results from experiments conducted by Zhou et al. (2006) and Yang et al. (2006) indicated that Gracilaria lemaneiformis has the potential to remove excess nutrients, such as inorganic nitrogen and phosphorus from sea water. This alga could be effective in controlling eutrophication in Chinese coastal waters especially when grown in an integrated mariculture system in fed fish farms. Gracilaria chilensis is another strain that was shown to be successful in the removal of ammonium from fish effluent produced by salmonids cultivated in tanks (Bushmann et al., 1996). Gracilaria gracilis was also reported to grow faster when 8 cultivated close to a site where fish waste is released in Saldanha Bay, South Africa (see section 1.2.4), which makes G. gracilis also ideal for integrated seaweed/fish mariculture.

1.1.7. The economic importance of Gracilaria

In 1989 Santelices and Doty reported that approximately 5000 tonnes of agar is processed annually from 25 000 to 30 000 tonnes of Gracilaria, harvested mainly from the wild in Chile, Argentina, Brazil and South Africa and from fishpond culture in Taiwan, Hainan Island, China and mainland China. In 2003 McHugh reported that agar production from both Gracilaria and Gelidium has an estimated total annual value of US$ 132 million. Chile is the largest supplier of Gracilaria. In Chile the harvest of wild seaweed has fluctuated from 121 000 wet tonnes in 1996 down to 73 000 tonnes in 1998 and back up to 137 000 wet tonnes in 2000, while cultivation has yielded about 33 000 wet tonnes during 1999 and 2000. Buschmann et al. (2001) reported total landings of Gracilaria inTown Chile being 120 000 wet tonnes in 2001. China, Indonesia, Namibia and Viet Nam all supplied between 12 000 and 18 000 wet tones each from a mixture of wild and cultivated seaweed. In Argentina the harvest of dried Gracilaria varied from 1 700 to 3 100 tonnesCape between 1985 and 1995. In India, harvests of wild Gracilaria and Gelidiella have varied between 750 and 1 300 dry tonnes from 1996 to 1999. Chile is thus the largest exporterof of Gracilaria, but also uses appreciable quantities itself. Indonesia and Viet Nam each uses about one third of its production, exporting the remainder. Other exporters of agar include Namibia and South Africa (McHugh et al., 2003).

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1.2. Gracilaria gracilis at Saldanha Bay, South Africa

1.2.1. Location of Saldanha Bay

Saldanha Bay forms the northern part of the Saldanha-Langebaan bay system and is situated on the west coast of South Africa, about 100 km north of Cape Town (Figure 2a and 2b). While the shallow southern portion (Langebaan Lagoon) is a nature reserve, a man-made ore jetty divides Saldanha Bay into Big Bay and Small Bay (Figure 2b and 2c) (Anderson et al., 1996a; Anderson et al., 1996b). Saldanha Bay is the only large sheltered bay on the temperate west coast of South Africa which supports natural populations of Gracilaria gracilis and where mariculture of this seaweed can also be conducted. It is thus the obvious site for development by the local industry.

S.B. B.B. b a Town

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Figure 2. a) Location of Saldanha Bay on South African west coast. b) Map of the Saldanha- Langebaan lagoon system. Small Bay is indicated as S.B.and Big Bay as B.B. c) Detailed map of Saldanha Bay showing depth contours in Small Bay (adapted from Anderson et al., 1992).

1.2.2. History of the Gracilaria gracilis population in Saldanha Bay

In Saldanha Bay, beach-cast Gracilaria gracilis has been collected commercially since the early 1940s. Due to the unavailability of agar in Japan during World War II, industrial agar was extracted from Gracilaria gracilis harvested from Saldanha Bay and the industry proved to be economically successful (Simons, 1977; Rothmann, 1990). In 1951, 20 tons of agar was produced locally, almost all being Gracilaria collected at Saldanha Bay, and 90% of this was used in the meat-canning industry (Anderson et al., 1989). Local production of agar ceased shortly afterwards for economic reasons, but it restarted in the 1960s when two factories for the production of agar were constructed in Kraaifontein and Vredendal in the Western Cape. In 1974, both factories had to close down when the supply of beach-cast Gracilaria collapsed as a result of dredging and the construction of a large ore-loading jetty and breakwater in the northern part of the bay (Rothmann, 1990; Anderson et al., 1989). The Gracilaria resource seemed to be recovering in the 1980s and in 1988 about 400 dry Towntons were obtained from beach cast in the northern area of Saldanha Bay (Anderson et al., 1992). Although the northern and northeastern shores of Saldanha Bay traditionally yielded large beach-casts of Gracilaria (Isaac, 1956), only the northern enclosedCape part of the Bay (Smalll Bay: Figure 2c) now yields commercially useful quantities of Gracilaria as a consequence of the construction of the jetty (Anderson et al., 1992). Constructionof of an agar-processing factory in Saldanha Bay was under consideration in 1988 (Anderson et al., 1996a).

However, at the end of 1988 the Gracilaria resource collapsed once again. Grazer-exclusion experiments were carried out during 1989, and it was found that sea urchins (Parechinus angulosus) and keyholeUniversity limpets (mainly De ndrofissurella scutellata, with some Fissurella mutabilis) together consumed more than 10% of transplanted Gracilaria within 24 hours at depths greater than about 4 m, while in shallow water (1-2 m), fish (Sarpa salpa) consumed about 80% of transplanted Gracilaria overnight. It was therefore concluded that these grazers were the main cause of the reduction in the Gracilaria during 1989, but their role in causing the initial collapse at the end of 1988 was uncertain (Anderson et al., 1992; Anderson et al., 1993). Anderson and co-workers (1992) however suggested the possibility that the exceptionally large swells during July, that produced the biggest beach-cast that year, may have unbalanced the system by reducing the Gracilaria beds and allowing the effects of grazers to outweigh production of biomass. 11

No further beach-casts were obtained until 1992. In the summer of 1993-1994, localized eutrophication due to fish-processing waste caused a problematic bloom of Ulva lactuca in Saldanha Bay (Anderson et al., 1996a). Large wash-ups of Ulva lactuca occurred in Small Bay for the first time. The Ulva was usually mixed in with the Gracilaria and it was practically impossible to separate the two, rendering much of the commercial Gracilaria beach-cast useless (Anderson et al., 1996b).

In 1996 Anderson et al. (1996a) reported that regular beach-casts of Gracilaria were again being obtained and conducted experimental investigation of mariculture of Gracilaria in Saldanha Bay. The authors used rope rafts to evaluate the growth of Gracilaria in Small Bay over a period of four years. Although warm oligotrophic water severely reduced the growth of the Gracilaria in Saldanha Bay during late summer, they found that growth would be optimized by growing the plants as close as possible to the surfaceTown (0.2 m) where water motion and nutrient uptake is higher. They concluded that suspended cultivation of Gracilaria in Saldanha Bay is feasible and that there is potential to improve yields by technical research and development, but that the mainCape limiting factor is the oligotrophic surface water (discussed in section 1.2.3), particularly in late summer. of 1.2.3. Thermal stratification in Saldanha Bay

For much of the 10-month upwelling season, which lasts from August to May, Saldanha Bay is thermally stratified with a surface warmed upper layer and a lower layer of cold upwelled water that is advectedUniversity into the bay (Monteiro et al., 1999). In spring (September – October), a thermocline develops as the surface water of the bay warms with increasing insulation and is trapped in the bay by southerly winds (Anderson et al., 1996a). During summer (November – March), increased insulation and prevailing southerly winds reinforce this effect, and strong stratification is maintained with a well-developed thermocline overlaid by a persistent warm oligotrophic (17 – 20 ˚C) surface layer (Anderson et al., 1996b). Stratification during summer is also enhanced by surface heating of the bay (Monteiro et al., 1999). In autumn (April – May), the bay de-stratifies as upwelling decreases. In winter (June – July), strong northerly winds and low air temperatures keep the water column well mixed at 12-14˚C throughout.

There is however an inverse relationship between the water temperature and nutrient concentrations. The cold upwelled water is high in nitrate, while the warm surface layer, which is effectively maintained in the bay throughout summer, is very low in nutrients. The thermocline is also not static in position throughout the summer. When cold bottom water moves into the bay, the depth of the warm surface layer decreases as it flows out, but when warm surface water flows in, it can drive the thermocline deeper than 10 m and thus below the range of the Gracilaria population. The thermocline thus moves from deeper to shallower water on an approximately 6 – 7 day cycle (Anderson et al., 1996a).

1.2.4. Other relevant studies that were conducted in Saldanha bay

Small Bay receives about 650 tons of nitrogen annually over and above natural fluxes. This nitrogen is in the form of liquid fish-processing waste that is discharged from two factories on the west shore. A study was conducted to test the hypothesis thatTown Gracilaria grown close to the site of waste release (1.5 km away and in the waste plume) would grow faster than at the control site 3.5 km away. It was found that proximity to the waste site may sometimes benefit the Gracilaria, but that the fish waste does Capein fact provide a significant source of nitrogen for seaweed cultivated throughout the northern area of Small Bay. This could however be particularly beneficial for the cultivationof of Gracilaria on suspended rafts during the summer months when the surface layer of the water becomes warm and oligotrophic, and thus is not naturally as nitrogen-rich as the cold bottom layer (Anderson et al., 1999).

During certain times of the year, raft-cultivated Gracilaria in Saldanha Bay becomes infected with the epiphyte CeramiumUniversity diaphanum and occasionally C. diaphanum which is also a red alga reaches greater biomass than G. gracilis, consequently lowering the yield and quality. Raft-cultivated Gracilaria in Saldanha Bay is also inhabited by large numbers of the invertebrate, Paridotea reticulata. P. reticulata is an isopod and can attain extremely high densities, comprising 10% of the harvested biomass. A study was conducted to establish the effect that high densities of P. reticulata may have on cultivated Gracilaria crops and their associated epiphytes. It was found that when given a choice, P. reticulata cosumed C. diaphanum rather than G. gracilis. The amount of Gracilaria consumed was thus negligible. If P. reticulata is therefore not present on the Gracilaria rafts, C. diaphanum may be able to reach pest status rapidly. It seems that the importance of P. reticulata may be in retarding the 13 initial spread of C. diaphanum, and keeping the Gracilaria relative epiphyte-free for a longer period of time (Anderson et al., 1998).

1.2.5. Bacterial epiphytes of Gracilaria gracilis from Saldanha Bay

Jaffray and co-workers (1997) suggested another reason for the collapse of the Saldanha Bay Gracilaria population at the end of 1988. Since plants examined during partial die-offs of the Saldanha beds (e.g. in January 1993) exhibited symptoms such as lesions and thallus bleaching, they reasoned that the collapse of the Gracilaria at the end of 1988 could have been the result of disease. Farming Gracilaria in open water provides an ideal submerged surface for the attachment of a variety of epiphytes such as bacteria, fungi, algae, nematodes and diatoms. Jaffray et al. (1997) hypothesized that although some secrete enzymes such as agarases which are potentially detrimental to Gracilaria, these epiphytes can exist on the surface of Gracilaria gracilis as non-pathogenic symbionts. It hasTown indeed been demonstrated by Weinberger et al. (1994) that the agarases secreted by epiphytic bacteria on the surface of the seaweed Gracilaria conferta are responsible for disease symptoms in the alga. It is thus possible that changes in environmental factors, suchCape as nutrient depletion and elevated water temperature, could alter the balance between epiphytes and the host plant, inducing pathogenesis. In order to determine the potentialof of the bacterial epiphytes for pathogenicity, a study was conducted to characterize the bacterial epiphytes colonizing a natural Gracilaria population from Saldanha Bay (Jaffray et al., 1997). It was found that 40% of the bacterial epiphytes exhibited varying degrees of extracellular agarolytic activity. This high proportion of agarolytic bacteria isolated from the surface of the Gracilaria suggests that their ability to produce agarases mightUniversity be important in addressing the nutritional requirements of these bacteria, by enabling them to exploit the agar in the cell wall of the Gracilaria as a nutrient source. It was therefore concluded that the nutrient depleted surface layer of the water, which occurs during the summer months at Saldanha Bay, may force agarolytic epiphytic bacteria growing on raft-cultivated Gracilaria to exploit the agar in the alga’s cell wall as a primary nutrient source during these unfavorable growth conditions. Lesions, bleaching and other signs of disease have indeed been observed in benthic and cultivated Gracilaria following more than 2 -3 weeks of warm oligotrophic water conditions in Saldanha Bay (Jaffray et al., 1997).

Jaffray and Coyne (1996) then developed an in situ assay to detect bacterial pathogens of Gracilaria gracilis. In short, an antibiotic ‘cocktail’ was used to generate axenic Gracilaria, whereafter 5 cm pieces of this axenic thalli were injected with the bacterial strain to be tested. The thalli were then scored for the appearance of symptoms, such as lesions and thallus bleaching, after a 5 day incubation period in liquid media at the elevated temperature of 30˚C. This assay was found to produce very consistent results and also satisfied the criteria of Koch’s postulates. The assay was used to identify several of the agarolytic bacterial epiphytes of Gracilaria gracilis as being pathogens during unfavorable environmental conditions. One of the pathogens that Jaffray and Coyne (1996) identified through this procedure, was the highly agarolytic strain Pseudoalteromonas sp. LS2i that was characterized in this study.

Later Schroeder (2001) also isolated a few culturable bacterial isolates from G. gracilis that was obtained from Saldanha Bay. He reported that 44 % of the isolates were agarolytic. The pathogenicity assay, devised by Jaffray and Coyne (1996), was thenTown used to proof that one of these isolates, Pseudoalteromonas gracilis B9, were indeed a pathogen of Gracilaria. An agarolytic gene was cloned from Pseudoalteromonas gracilis B9 and the enzyme from this gene was purified and used as a tool to elucidate itsCape role in the virulence mechanism employed by the bacterium in eliciting disease in G. gracilis (Schroeder, 2001; Schroeder et al., 2003) of

University

1.3. Agarases

1.3.1. Agarolytic bacteria

Red algae (Rhodophyta) produce carrageenans and agars, which are the main components of their cell wall (Graigie, 1990). The bioconversion of these two components is essentially performed by marine bacteria (Michel et al., 2006). Agar degrading bacteria were first described in 1902 by Gran (Morrice et al., 1983b). Since then, a number of bacteria have been reported to degrade agars. They were mainly isolated from the marine environment, either in the water column, in coastal marine sediments or associated with red algae (Michel et al., 2006). Agarolytic bacteria have however also been identified from other sources, such as soil (Suzuki et al., 2002; Kirimura et al., 1999; Voget et al., 2003; Lakshmikanth et al., 2006a), sewage (Hofsten and Malmqvist, 1975), fresh water (Van der Meulen et al., 1974), the rhizosphere of spinach (Hosada et al., 2003) and plant roots (HosadaTown and Sakai, 2006). In comparison to agar-degrading bacteria, a lot less bacteria have been reported to hydrolyze carrageenans and all these bacteria were isolated from the marine environment (Michel et al., 2006). Cape of 1.3.2. Applications of agarases

Seaweeds contain useful substances such as vitamins, fatty acids, carotenoids and betaine. Degradation of the polysaccharides in the cell wall of seaweeds is however required during purification of these substances and degradation methods such as acid hydrolysis is so severe that the labile substancesUniversity are not extracted intact. Extraction with enzymes such as agarases is a good alternative, because labile substances may be quite tolerant of the mild conditions produced during agarolytic degradation (Sugano et al., 1993a). Agarases can also be used to degrade the cell walls of marine algae for the generation of protoplasts. The production of large amounts of enzyme is needed for these purposes (Sugano et al., 1995). Protoplasts can be used for the performance of biotechnology on agarophyte red algae (Araki et al., 1998).

Agarases have potential applications in the food, cosmetic and medical industries for the production of oligosaccharides from agar (Lakshmikanth et al., 2006). Oligosaccharides that are obtained by enzymatic hydrolysis of agarose have several properties, i.e. they are potent 16 anti-oxidants and reactive free radical scavengers and has the ability to inhibit lipid peroxidation (Wang et al., 2004); they can inhibit the growth of bacteria, slow the rate of degradation of starch and are often used as low-calorie additives to improve food quality (Otha et al., 2004c); they have been shown to have antitumorigenic activity in mice (Fernandez et al., 1989); they have macrophage-stimulating activity and are suitable as a source of physiologically functional foods with protective and immunopotentiating activity (Yoshizawa et al., 1995). Neoagarobiose, one of the hydrolysis products of agarose following β-agarase degradation, has both whitening and moisturizing effects on melanoma cells (Kobayashi et al., 1997). Another breakdown product of agar hydrolysis, neoagarohexaose, may possibly be useful as a more effective skin moisturizer than smaller oligosaccharides, because the viscosity of hexamers is higher than that of smaller oligosaccharides (Otha et al., 2004a).

1.3.3. Characterized agarases Town

1.3.3.1. Pseudoalteromonas atlantica Cape The agarase system of the marine bacterium Pseudoalteromonas atlantica (Figure 3) initially discovered by Yaphe (1957), was extensivelyof characterized and provides a model for the study of other marine agarase systems. The pathway of agar breakdown by this bacterium begins with the initial cleavage of agar at the -1,4 linkage by the extracellular -agarase I to produce a mixture of oligosaccharides with neoagarotetraose [O-3,6-anhydro--L- galactopyranosyl(1-3)-O-- D-galactopyranosyl(1-4)-O-3,6-anhydro-- L- galactopyranosyl(1-3)-University D-galactose] as the predominant product. Neoagarotetraose permeates into the cell, while the larger agar oligomers cannot cross the cell wall barrier. Neoagarotetraose is then cleaved by the membrane-bound -agarase II, also known as neoagarotetraose hydrolase, to produce neoagarobiose [O-3,6-anhydro--L- galactopyranosyl(1-3)- D-galactose]. Ultimately, the neoagarobiose is hydrolyzed by the intracellular neoagarobiose hydrolase to the metabolically useful carbon sources, D-galactose and 3,6-anhydro-L-galactose (Belas et al., 1988; Day and Yaphe, 1975; Morrice et al., 1983a, 1983b, 1983c and 1983d).

AGAROSE

β – AGARASE I (HYDROLYSIS OF β (1-4) BONDS)

NEOAGAROTETRAOSE

β – AGARASE II (HYDROLYSIS OF β (1-4) BONDS)

Town

NEOAGAROBIOSE

Capeα – NEOAGAROBIOSE

HYDROLASE of (HYDROLYSIS OF α (1-3) BONDS)

University 3,6 – ANHYDRO – L - GALACTOSE D - GALACTOSE

Figure 3. Enzymology of agar digestion in P. atlantica strain ATCC 19262. Oligomeric agar is initially cleaved by the exported β-agarase I. This enzyme cleaves agar at the β-1,4 linkage to produce neoagarotetraose. This tetramer is then broken down to neoagarobiose through the action of β-agarase II (neoagarotetraose hydrolase). The latter enzyme is also capable of digesting oligomeric agar, cleaving it at the β-1,4 linkage to produce neoagarobiose. The enzyme neoagarobiose hydrolase cleaves neoagarobiose at the -1,3 linkage to produce D- galactose and 3,6-anhydro-L-galactose (adapted from Belas et al., 1988).

-agarase I of P. atlantica strain ATCC 19292 (NCMB 301) was purified by Morrice et al. (1983d). SDS-PAGE analysis indicated a homogeneous protein with a molecular weight of 32 000 Da. This enzyme specifically recognizes regions containing a minimum of one unsubstituted neoagarobiose unit [3,6-anhydro-alpha-L-galactopyranosyl-(1-3)-D-galactose] and hydrolyzes the reducing side of the moiety (Morrice et al., 1983b). Yaphe (1957) demonstrated that agar was degraded by -agarase I to neoagarooligosaccharides where the tetramer is the predominant end-product. The nucleotide sequence of the above mentioned - agarase I was submitted to the database in 1996 as the dagA gene. The dagA gene has an open reading frame (ORF) of 873 nucleotides. The nucleotide sequence of the agrA gene from P. atlantica strain T6c, also encoding an extracellular -agarase I, was determined after it was cloned into E. coli. This gene has an open reading frame of 1515 nucleotides which predicts a primary translation product of 504 amino acids with a molecular weight of 57 486 Da. Probing the genome of strain ATCC 19262 with the agrA gene from strain T6c indicated that the homolog of agrA is not present in strain ATCC 19262 (BelasTown et al., 1988; Belas, 1989).

Belas and co-workers (1988) also constructed a P. Capeatlantica strain T6c mutant with reduced agarolytic activity by gene replacement. Thisof was accomplished through the insertion of a selectable drug (kanamycin) resistance gene into the coding sequence of the cloned agrA gene (encoding -agarase I) and the subsequent introduction of the agrA::kan plasmid into P.atlantica in which the wild-type locus was displaced by homologous recombination. The wild type strain produced colonies that were larger and more mucoid than those of the agrA – strain. Although the wild type strain grew better and produced more exopolysaccharide, the mutant strain could stillUniversity give rise to colonies on a medium where agar was the sole carbon source. Therefore, although growth of the mutant was impaired, the minor agarase activity produced by the mutant was still sufficient to sustain growth of this strain on agar. They concluded that since the minor agarolytic activity still present in the mutant was much weaker than that of the primary exported agarase, and since the pitting were more closely associated with the cells, it may be the result of the second agarase, the membrane-bound -agarase II.

Morrice et al. (1983d) partially purified the -agarase II protein from the soluble fraction of disrupted P. atlantica strain ATCC 19292 cells (NCMB 301).This enzyme rapidly degraded isolated neoagarotetraose and neoagarohexaose to the disaccharide (Morrice et al. 1983b). - agarase II is probably the ‘-neoagarotetraose hydrolase’ described by Groleau and Yaphe (1977). -neoagarotetraose hydrolase was purified from P. atlantica (National Culture Type Collection (NCTC) 19262) and was localized on or outside the cytoplasmic membrane, in the cell wall region. The authors found that this enzyme cleaves the central -linkage near the nonreducing end in neoagarohexaose and neoagarooctaose to yield neoagarobiose (Groleau and Yaphe, 1977).

The neoagarobiose hydrolase enzyme was purified from P. atlantica NCTC 18685 by Day and Yaphe (1975). The size of the enzyme was determined by sucrose density gradient centrifugation to be 10 kDa. Neoagarobiose hydrolase was found in the outer region of the cell, on or outside the cytoplasmic membrane. It only cleaved theTown -linkage in neoagarobiose and 3,6-anhydro-L-galactose appears to be a requirement for activity (Day and Yaphe, 1975).

1.3.3.2. Pseudoalteromonas antarctica Cape of Pseudoalteromonas antarctica strain N-1 was isolated from decomposing algae from the southern Chilean coast. An extracellular agarase was purified from this bacterium and the size of the protein was determined to be 33 kDa. The protein was found to be homogeneous on the basis of SDS-PAGE. The enzyme hydrolyzed the β-1,4-glycosidic linkages of agar, yielding neoagarotetraose and neoagarohexaose as the main products. P. antarctica strain N- 1 also produced a red-brownUniversity diffusible pigment after incubation on agar for more than 48 h at 25 C (Vera et al., 1998).

1.3.3.3. Pseudomonas sp. W7

The agar-degrading bacterium, Pseudoalteromonas sp. W7, was isolated from an aquacultured red alga. A β-agarase gene, designated pjaA, was cloned from this strain into E. coli JM83 (Kong et al., 1997). Sequence analysis of pjaA revealed that this agarase gene consists of an open reading frame of 1926 bp (Lee et al., 2000). The recombinant agarase was 20 expressed in E. coli JM83 and purified from the culture supernatant. The purified β-agarase migrated as a single band with a molecular mass of 59 kDa on SDS-PAGE and was found to hydrolyze the β-1,4-linkage of agarose to yield neoagarotetraose as the main product (Ha et al., 1997). The results of a biochemical study done by Ha and co-workers (1997) indicated that the agarolytic activity of PjaA was increased by the addition of NaCl and they therefore postulated that the Na+ and the Cl- ions are involved in enzyme activation. After determining the sequence of the pjaA gene, Lee and his co-workers (2000) found that the three conserved chloride-binding ligands which are essential to the -amylase from A. haloplanctis for the binding of chloride ions and subsequent enzyme activation could be found within the PjaA enzyme. Therefore they suggested that these conserved binding ligands within PjaA could help regulate the enzymatic activity in the same manner as the A. haloplanctis -amylase.

1.3.3.4. Pseudomonas sp. SK38 Town This yellow-pigmented bacterium that causes green spot rot in the seaweed Porphyra dentata was isolated and identified by Ryu et al. (2001). The pagA gene encoding a β-agarase from Pseudomonas sp. SK38 was cloned and expressed Capein E. coli by Kang and co-workers (2003). The structural gene consists of 1011 bp and encodes a protein with a predicted molecular weight of 37326 Da and a signal peptide of 18of amino acids. The recombinant β-agarase was also expressed and purified from E. coli (Kang et al., 2003).

1.3.3.5. Pseudomonas aeruginosa AG LSL-11

This strain of PseudomonasUniversity aeruginosa was isolated from seawater through an enrichment culture technique. Lakshmikanth and his co-workers (2006) optimized the culture medium and culture conditions for maximal production of extracellular agarases from this bacterium. Three agarases, LSL-11a, LSL-11b and LSL-11c, whose molecular weights (determined by SDS-PAGE) were estimated to be 76, 64 and 46 kDa respectively, were partially purified from P. aeruginosa AG LSL-11. The agarases secreted by this bacterium were inducible by agar. This strain was also able to utilize a number of other sugars as sole source of carbon and energy. Unlike previously reported agarolytic Pseudomonas strains, P. aeruginosa AG LSL- 11 did not have a specific requirement for sodium chloride, and could grow well and produce extracellular agarases even in its absence (Lakshmikanth et al., 2006). 21

1.3.3.6. A Pseudomonas-like bacterium

This gram-negative rod, only described as a Pseudomonas-like bacterium, was isolated from sewage by Hofsten and Malmqvist (1975). Two different agarases, agarase I and agarase IIb, were purified from this bacterium. The two proteins had similar amino acid compositions and both were found to be glycoproteins. Molecular weight determinations indicated agarase I to be a dimer of 210 kDa, while the molecular weight of agarase II was determined to be 63 kDa. Both of these agarases were cell bound during exponential growth, but were released into the medium during the stationary growth phase. Agarase I hydrolyzed the β-linkages in neoagarooctaose to produce two moles of neoagarotetraose or one mole of neoagarohexaose and one mole of neoagarobiose. Agarase IIb hydrolyzed only the central β-linkage in neoagarooctaose to form two moles of neoagarotetraose (Malmqvist, 1978). Town 1.3.3.7. Alteromonas agarlyticus strain GJ1B

Alteromonas agarlyticus strain GJ1B is an agarolyticCape marine bacterium. The capability of this strain to degrade agar is however more pronounced in comparison to the majority of other characterized agarolytic bacteria. Instead of pittingof the agar around the colony like other agar decomposers, this strain liquefies the gel and makes holes until the colonies reach the bottom of the agar plate (Potin et al. 1993). An -agarase was first isolated and purified from A. agarlyticus strain GJ1B by Young and co-workers (1978). Later Potin et al. (1993) also purified the -agarase from the culture supernatant of this strain to homogeneity. They detected the purified University-agarase as a single band with a molecular mass of 180 kDa with SDS- PAGE. The native molecular mass of the enzyme was however determined to be 360 kDa after affinity-chromatography, which suggested that the native enzyme is a dimer.

Town Figure 4. The agarolytic system of A. agarlyticus. Agaro-oligosaccharides are first formed through the specific cleavage of -1,3 linkages in agarose by an -agarase. These oligosaccharides are then partially degraded by a β-galactosidase acting at the reducing end, yielding oligosaccharides of the agarotriose family; i.e., with an odd number of saccharide units. Saccharide units and linkages are symbolizedCape as: , galactose; , reducing galactose; , 3,6-anhydrogalactose; , 3,6-anhydrogalactosehydrate (adapted from Potin et al., 1993). of

An -agarase is specific for the -1,3 linkages in agarose and Both Potin and Young found that the major end product of agarose hydrolysis by this specific -agarase is agarotetraose. Potin and co-workers (1993) co-purified a β-galactosidase in the -agarase affinity- chromatography fractionUniversity (first of the two steps in the purification process), probably as part of a complex with the -agarase. The degradation of agarose by this agarase complex yielded a mixture of agarotetraose and agarotriose. The agarolytic system of A. agarlyticus strain GJ1B is illustrated in Figure 4, where agaro-oligosaccharides are initially formed by specific cleavage of the -1,3 linkages in agarose. These are then partially degraded by the β-galactosidase acting at the reducing end to yield oligosaccharides of the agarotriose family.

1.3.3.8. Alteromonas sp. strain C-1

This marine bacterium was isolated from the Bay of San Vicente in Chile. An extracellular agarase was purified from it to homogeneity. The molecular weight of the agarase as determined by SDS-PAGE is 52 kDa. The purified agarase hydrolyzed agar to yield neoagarotetraose as the main product (Leon et al., 1992).

1.3.3.9. Alteromonas sp. E-1

Alteromonas sp. E-1 was isolated from soil. Unlike the majority of characterized β-agarases that are produced extracellularly, a β-agarase was purified from the cell-free extract of this strain after sonication. The molecular weight was estimated to be 82 kDa by SDS- polyacrylamide gel electrophoresis. The final product of agar hydrolysis was found to be neoagarobiose. This β-agarase was the first agarase reported to produceTown only neoagarobiose and not a mixture of neoagarobiose and neoagarotetraose as the final product (Kirimura et al., 1999). Cape 1.3.3.10. Alteromonas sp. SY37-12 of This bacterium was isolated from the southern ocean of China. An extracellular agarase was purified from it and the purified protein exhibited a single band of 39.5 kDa on SDS-PAGE. The enzyme was found to hydrolyze the β-1,4-glycosidic linkages of agar to yield neoagarotetraose and neoagarohexaose as the main products (Wang et al., 2005). University 1.3.3.11. Bacillus sp. MK03

Unlike the majority of agarolytic bacteria that have been characterized so far, this bacterium was not isolated from a marine environment, but from soil collected from the Gifu prefecture in Japan (Suzuki et al., 2002). An -neoagarooligosaccharide hydrolase was purified from the culture fluid of Bacillus sp. MK03 and the purified enzyme appeared as a single band on a SDS-PAGE gel. An estimation of the molecular mass by SDS-PAGE and gel filtration gave values of 320 kDa and 42 kDa, respectively indicating that the enzyme is octameric. The -neoagarooligosaccharide hydrolase cleaved the -1,3 linkage in neoagarobiose to produce 24

3,6-anhydro-L-galactose and D-galactose. It was also found to cleave the -1,3 linkage at the nonreducing end in neoagarotetraose or neoagarohexaose to give 3,6-anhydro-L-galactose and agarotriose or agaropentaose (Figure 5). This enzyme has thus the same mode of action as the -neoagarooligosaccharide hydrolase that was purified from Vibrio sp. strain JT0107 (section 1.3.3.12). However the latter enzyme acts intracellularly, while this enzyme acts extracellularly (Suzuki et al., 2002).

An extracellular β-agarase was also purified from Bacillus sp. MK03. The purified enzyme appeared as a single band of 92 kDa on SDS-PAGE. The predominant hydrolysis product generated by this agarase was neoagarotetraose, indicating cleavage of the β-1,4 linkage (Suzuki et al., 2003).

Town

Cape of

Figure 5. The mode of action of -neoagarooligosaccharide hydrolase. N2, neoagarobiose; N4, neoagarotetraose;University N6, neoagarohexaose; closed circles, 3,6-anhydro-L-galactose; open circles, D-galactose; , -1,3 linkage; β, β-1,4 linkage. The nonreducing end is indicated on the left-hand side and the reducing end is indicated on the right-hand side. The linkage selectively cleaved by the -neoagarooligosaccharide hydrolase from Bacillus sp. MK03 is indicated by an inverted closed triangle (adapted from Suzuki et al., 2002).

1.3.3.12. Vibrio sp. strain JT0107

Vibrio sp. strain JT0107 is a marine strain that decomposes the walls of some seaweeds, such as Undaria pinnatifida and a Laminaria sp. (Sugano et al., 1993a). Three agarases as well as 25 an -neoagarooligosaccharide hydrolase, have been characterized from Vibrio sp. strain JT0107. Agarase 0107 or agaA was the first agarase to be characterized (Sugano et al., 1993a). The latter enzyme was isolated and purified from the culture medium and was found to migrate as a single band of 107 kDa on a SDS-PAGE gel. This novel enzyme hydrolyzed the β-1,4 linkage of agarose to yield neoagarotetraose and neoagarobiose (Sugano et al., 1993a). Agarase 0107 (agaA) not only hydrolyzed agarose, but also neoagarotetraose to yield neoagarobiose. Tha agarase 0107 gene (agaA) was then cloned into Escherichia coli before it was sequenced. The gene had an open reading frame of 2985 nucleotides which gave a primary translation product of 975 amino acid residues, with a predicted molecular weight of 105271 Da which agrees well with the molecular weight determined through SDS-PAGE of the purified protein (Sugano et al., 1993b).

Analysis of the genomic DNA sequence just upstream of agaA revealed an open reading frame of 955 amino acids coding for another agarase gene (agaBTown). A comparison of the deduced amino acid sequence of agaB to that of agaA revealed the existence of highly homologous regions. A portion of the agaB gene expressed in E. coli yielded a protein with agarolytic activity. The substrate specificity of agaBCape was however different to that of agaA as agaB expressed in E. coli did not hydrolyze neoagarotetraose whereas agaA could. The authors concluded that since agaA and agaB ofare located in tandem, the genes may constitute an agarase operon (Sugano et al., 1994a).

A third agarase was found to be produced under different culture conditions, such as increased aeration and a sufficient concentration of agarose. This agarase designated agarase 0072 was isolated andUniversity purified to homogeneity. The purified protein migrated as a 72 kDa band on a SDS-PAGE gel. The N-terminal amino acid sequence of agarase 0072 was not identical to either that of agaA or agaB or to any integral sequence of agaA or agaB, and was therefore considered to be a third agarase isolated from Vibrio sp. strain JT0107. This novel enzyme was found to be a β-agarase that catalyzes the hydrolysis of the β-1,4 linkage of agarose to yield neoagarotetraose and neoagarobiose (Sugano et al., 1995).

Sugano and his co-workers (1994) also described for the first time the purification and characterization of an enzyme that can hydrolyze oligosaccharides from agarose to monosaccharides. This -neoagarooligosaccharide hydrolase, which was not secreted into the 26 culture medium, was purified from cultured cells of Vibrio sp. strain JT0107. Using SDS- PAGE analysis, the purified protein was found to give a single band of 42 kDa. However, estimation of the molecular weight of the protein by gel filtration gave a value of 84 kDa, indicating that the enzyme is a homodimer. The enzyme was found to hydrolyze the -1,3 linkage of neoagarooligosaccharides to yield agaropentaose, agarotriose, agarobiose, 3,6- anhydro-L-galactose and D-galactose. The -neoagarooligosaccharide hydrolase recognizes the -1,3 linkage between D-galactose and 3,6-anhydro-L-galactose at the nonreducing end.

1.3.3.13. Vibrio sp. AP-2

This pophyran-degrading bacterium was isolated from an alga collected in the coastal sea of the Fukuoka Perfecture of Japan (Aoki et al., 1990). This strain produces three extracellular agarases. Two of these agarases act on hexoses or larger saccharides while the third agarase hydrolyzed neoagarotetraose, larger neoagarooligosaccharides andTown agar to give neoagarobiose as the predominant product. The latter agarase was purified to homogeneity from the culture fluid of Vibrio sp. AP-2, whereafter the molecular mass was determined through SDS-PAGE analysis to be 20 kDa (Aoki etCape al., 1990).

1.3.3.14. Vibrio sp. PO-303 of

Vibrio sp. PO-303 was isolated from sea mud collected from the coastal sea area in Japan (Araki et al., 1998). Three agarases were purified to homogeneity from this bacterium. Through SDS-PAGE, the molecular mass of agarase-a, -b, and –c was estimated to be 87.5 kDa, 115 kDa and 57University kDa respectively. The three agarases differed from each other in their N-terminal amino acid sequences. In comparison with amino acid sequences of other reported agarases, only the sequence of agarase-b showed significant homology (75% identity in the first 20 amino acids) with the N-terminal region of agarase 0107 from Vibrio sp. strain JT0107. Agarase-a hydrolyzed agarose to give neoagarotetraose and neoagarohexaose as predominant products, while the main hydrolysis product of agarase-b was neoagarobiose from agarose. Agarase-c was found to be a new enzyme with different characteristics to those of the agarases that have been reported so far, because the main hydrolysis products from agar were neoagarooctaose and neoagarodecaose (Araki et al., 1998).

1.3.3.15. Cytophaga flevensis

This yellow-pigmented bacterium was isolated from a fresh-water lake in the central part of the Netherlands (Van der Meulen et al., 1974). An extracellular agarase produced by Cytophaga flevensis was partially purified and the molecular weight of the enzyme was determined to be 26 kDa. This enzyme was determined to be a β-agarase which hydrolyzed agarose to produce mainly neoagarotetraose and neoagarobiose (Van der Meulen and Harder, 1975). Later, Van der Meulen and Harder (1976) reported the presence of two additional enzymes, a neoagarotetra-ase and neoagarobiase, whose enzyme activities were localized in the cytoplasm of C. flevensis. They proposed that neoagarotetra-ase was found to hydrolyze neoagarotetraose by cleavage of the central β-galactosidic linkage, while neoagarobiose, produced both by the action of neoagarotetra-ase and agarase, is then hydrolyzed by neoagarobiase through the cleavage of its -galactosidic linkageTown into D-galactose and 3,6-anhydro-L-galactose.

1.3.3.16. Streptomyces coelicolor A3(2) Cape

An extracellular agarase gene from Streptomycesof coelicolor A3(2) was cloned into Streptomyces lividans 66 using the multicopy plasmid vector pIJ702 from which the agarase was overexpressed up to 500 times (Kendall and Cullum, 1984). Butner et al. (1987) determined the sequence of the coding and regulatory regions of this cloned agarase that was designated dagA. The sequence predicted a primary translation product of 309 amino acids with a predicted molecularUniversity weight of 35 kDa. The agarase protein produced by the S. lividans 66 strain that contained the recombinant plasmid was purified to near homogeneity. A comparison of the molecular weight of the purified extracellular dagA protein with that of the purified intracellular form and the in vitro transcription-translation product of this protein implies that the agarase is synthesized as a pre-protein possessing a signal peptide which is removed during secretion of the protein into the extracellular medium to give protein with a molecular weight of 29 kDa in size (Bibb et al., 1987).

Two S. coelicolor A3(2) mutant strains which exhibited a loss in the production of diffusible agarase when grown on an agar plate have also been reported (Hopwood et al., 1973a). One 28 of these mutations had resulted from the insertion of a plasmid into the dag region of the chromosome and the subsequent inactivation of this gene. The other dag mutation may have resulted from a spontaneous or UV- induced mutation of the dag gene (Hodgson and Chater, 1981). Several mutants deficient in their ability to utilize agar were also identified in S. coelicolor A3(2). These included mutations in the aga locus which prevented the strain from growing on agar as a sole carbon source, but did not affect the extracellular agarolytic activity (Hopwood et al., 1973a).

1.3.3.17. A Microbulbifer – like bacterium, strain JAMB-A94

Strain JAMB-A94 was isolated from Suruga Bay, Japan, at a depth of 2 406 m and the 16S rDNA of the isolate had the closest match (94.8% homology) to a Microbulbifer species (Ohta et al., 2004c). Two agarases, AgaA and AgaO, were cloned and sequenced from this strain (Ohta et al., 2004c; Ohta et al., 2004a). The AgaA gene encodedTown a protein with a calculated molecular mass of 48,2 kDa, and like the majority of other characterized agarases, the deduced amino acid sequence showed substantial homology with other agarases that are members of the glycoside hydrolase family 16 (GH-16).Cape The AgaA gene was cloned into Bacillus subtilis and the recombinant enzyme that was over-produced extracellularly, was purified from the latter strain. The purified AgaAof was identified as a β-agarase that yielded neoagarotetraose as the main final product (Ohta et al., 2004c).

The calculated molecular mass of the mature AgaO protein, deduced from its amino acid sequence, was 126.9 kDa. The catalytic module of AgaO resembled a glycoside hydrolase family 86 (GH-86) βUniversity-agarase. The AgaO gene was also cloned into Bacillus subtilis and overproduced extracellularly before it was purified. This is the first glycoside hydrolase family 86 enzyme that was homogeneously purified and characterized. AgaO hydrolyzed agarose to yield neoagarohexaose as the main product. This is also the first reported agarase to produce neoagarohexaose effectively. There was no homology between the catalytic domain of the family GH-16 agarase AgaA and that of the family GH-86 AgaO agarase (Ohta et al., 2004a).

1.3.3.18. Microbulbifer strain JAMB-A7

This agar-degrading bacterium was isolated at a depth of 1174 m from the sediment in Sagami Bay, Japan (Otha et al., 2004b). A β-agarase gene was cloned and sequenced from Microbulbifer strain JAMB-A7. It was determined to encode a protein with a calculated molecular mass of 49 kDa, belonging to family 16 of the glycoside hydrolases. The agarase enzyme was purified to homogeneity after it was overproduced extracellularly in Bacillus subtilis. The purified agarase hydrolyzed agar to yield neoagarotetraose as the main product (Otha et al., 2004b).

1.3.3.19. Microscillla sp. strain PRE1

This bacterium was isolated from coastal California marine sedimentTown and was found to harbour a single 101-kb circular DNA plasmid. Sequence analysis of the plasmid indicated the presence of five putative agarase genes. Loss of the plasmid was associated with loss of agarolytic activity, which supported the sequence analysisCape results (Zhong et al., 2001).

1.3.3.20. Agarases from a mixed microbialof population

In order to isolate genes that encode novel biocatalysts, the authors of this paper used a combined cultivation and direct cloning strategy to exploit the metagenome of a mixed microbial population from soil that was collected from an unplanted field (Voget et al., 2003). Total genomicUniversity DNA from this bacterial community was used to construct cosmid DNA libraries, from which four clones encoding 12 putative agarase genes were identified. Most of the agarase genes were organized in clusters consisting of two or three genes (Voget et al., 2003).

1.3.3.21. Thalassomonas strain JAMB-A33

Thalassomonas strain JAMB-A33 was isolated from the sediment off Noma Point, Japan, at a depth of 230 m (Otha et al., 2005a). A novel -agarase from this bacterium was purified to homogeneity. The molecular mass of the purified protein, designated agarase A33, was 30 determined to be 85 kDa by both SDS-PAGE and gel-filtration chromatography. Agar hydrolysis yielded predominantly tetramers with a concomitant production of hexamers and dimers. This is the second report on the characterization of an -agarase (Otha et al., 2005a).

1.3.3.22. Agarivorans sp. JAMB-A11

This strain was isolated from a sediment sample from the the Kuril Trench in Japan at a depth of 4152 m (Otha et al., 2005). A β-agarase gene from Agarivorans sp. JAMB-A11, designated agaA11 was expressed in Bacillus subtilis and the sequence of this gene determined. The efficiency of production of this agarase in Bacillus subtilis was found to be 30-fold greater than that in the original strain. The AgaA11 enzyme had a molecular mass of 105 kDa and is one of the few β-agarases that hydrolyzes not only agarose, but also neoagarotetraose, to yield neoagarobiose as the final main product. This recombinant agarase would be useful for industrial production of neoagarobiose (OthaTown et al., 2005).

1.3.3.23. Acinetobacter sp. AG LSL-1 Cape This agar-liquefying bacterium was isolated from soil samples. The bacterium produced an extracellular β-agarase that was partially purifiedof and found to produce a 100 kDa agarase as determined by native-PAGE and SDS-PAGE. Agar hydrolysis yielded neoagarobiose as the final end product (Lakshmikanth et al., 2006a).

1.3.3.24. Saccharophagus degradans 2-40 University A marine bacterium strain 2-40, was isolated from the surface of decomposing saltwater cord grass, Spartina alterniflora, in the lower Chesapeake Bay and was initially classified as a species within the Alteromonas genus (Andrykovitch and Marx, 1988). Later, based upon 16S rDNA analysis and phenotypic criteria, it was shown to be closely related to the genus Microbulbifer and was renamed to Microbulbifer degradans (Gonzalez and Weiner, 2000). Recently the genome sequence of this bacterium was completed and the strain was renamed to Saccharophagus degradans 2-40 (Ekborg et al., 2006).

Whitehead and co-workers (2001) reported that S. degradans 2-40, which is able to degrade a number of different complex carbohydrates, synthesized an agarase system that consisted of at least three enzymes. End-product analysis revealed that a β-agarase I, a β-agarase II and an

-neoagarobiose hydrolase, act in concert to degrade polymeric agar to D-galactose and 3,6- anhydro-L-galactose. They also purified the principal agarase, β-agarase I and determined the molecular mass of this agarase to be 98 kDa by SDS-PAGE analysis. The latter enzyme was found to degrade agarose to neoagarobiose and neoagarotetraose. According to Whitehead et al. (2001) the mode of action of the agarase system of S. degradans 2-40 is similar in part to that of Pseudoalteromonas atlantica (see section 1.3.3.1). Both strains synthesize β-agarase systems, consisting of a β-agarase I, a β-agarase II and an -neoagarobiose hydrolase, but in S. degradans 2-40, enzyme activity resided with the cell in the earlier stages of growth, and subsequently in the spent medium, while in P. atlantica the β-agarases appear to be extracellular while the -neoagarobiose hydrolase is cell-associated (Whitehead et al., 2001). Town After the completion of the genome sequence of S. degradans 2-40, Ekborg and his co- workers (2006) identified five agarases encoded by this bacterium. The identification of these agarases was based upon sequence similarity, conservedCape structural features, agarase activity of expressed genes in E. coli and/or the phenotype of gene replacement mutants. Aga16B resembled the β-agarase I that were purified byof Whitehead et al. (2001). Ekborg et al. (2006) demonstrated however that the latter agarase is produced extracellularly and belongs to glycoside hydrolase family 16 (GH-16). Although the enzymatic activity of Aga50A has yet to be demonstrated directly, this agarase exhibits sequence similarity to several known agarases as well as a GH-50 domain, which thus far has only been found in agarases. University Enzymatic activity of Aga86C could also not be demonstrated but again this enzyme exhibits sequence similarity to known agarases and has a GH-86 domain in its conserved region. Like GH-50 domains, GH-86-domains have also been identified only in agarases thus far. Aga86C was also found to be expressed by agarose-grown cells but not by glucose-grown cells. The identification of Aga50D as an agarase is based solely upon conserved sequence features. The latter gene has extensive sequence similarities to a GH-50 domain, but whether Aga50D is expressed or not, remains to be established. Aga86E exhibited sequence similarity to several other agarases, as well as to a probable GH86 domain. A purified His-tagged derivative of the latter gene was found to be active in the degradation of agarose, releasing almost exclusively 32 neoagarobiose, which suggests that Aga86E acts similarly to the β-agarase II from P. atlantica. An Aga86E mutant also exhibited a diminished ability to grow on agar as a carbon source, indicating Aga86E having a role in agarose metabolism. Although cell-free lysates of S. degradans 2-40 released D-galactose from polymeric agarose, the sequence of a neoagarobiose hydrolase enzyme has not been identified as yet (Ekborg et al., 2006).

1.3.3.25. Zobellia galactanivorans Dsij

This bacterium has the ability to degrade both agar and carrageenan and was isolated from the red alga, Delesseria sanguinea, on the sea-shore of Roscoff, Brittany, France (Potin et al., 1991). It was originally classified as Cytophaga drobachiensis Dsij but was later renamed to Zobellia galactanivorans Dsij (Barbeyron et al., 2001). Z. galactanivorans Dsij produces three different agarases, designated AgaA, AgaB and AgaC. AgaA and AgaC are extracellular, while AgaB is not secreted. The two β-agarases, AgaATown and AgaB, were functionally cloned into E. coli, overexpressed and purified to homogeneity. The sequence of the AgaA and AgaB genes was determined and they were found to encode proteins with theoretical masses of 60 kDa and 40 kDa respectively.Cape Sequence analysis indicated that both AgaA and AgaB feature homologous domains belonging to glycoside hydrolase family 16 (GH-16). It was further determined that AgaAof consists of the catalytic domain (AgaAc) and two C-terminal domains which are processed during secretion of the enzyme, while AgaB has a catalytic module and a signal peptide. Electrospray MS experiments and gel filtration demonstrated that AgaB is a dimer in solution, while AgaAc is a monomer. Both enzymes cleave the -1,4 linkages of agarose in a random manner, with AgaAc producing essentially neoagarotetraose andUniversity neoagarohexaose and AgaB producing neoagarotetraose and neoagarobiose as end-products. Although these enzymes behave similarly in the presence of liquid agarose, AgaAc was twice as efficient as AgaB in the degradation of agarose gels. The fact that AgaB is likely to be membrane-bound while AgaA is produced extracellularly, together with the organizational and catalytic differences between the two enzymes, suggests that AgaA is specializes in the initial attack on solid-phase agarose, while AgaB is involved with the degradation of agarose fragments (Jam et al., 2005).

In 2003, Alouch and co-workers (2003) reported the crystal structures of AgaA and AgaB, which are the first crystal structures of any agarases to be characterized. The structure of AgaA was solved by the multiple anomalous diffraction method, while the structure of AgaB was solved with molecular replacement using AgaA as model. To avoid the problems associated with the crystallization of multimoduler proteins, the authors expressed only the catalytic models of the two enzymes. Both their structures adopted a jelly roll fold with a deep active site channel harboring the catalytic machinery. Structure similarities together with sequence similarities between members of family GH-16 allowed the identification of the catalytic machinery, i.e. the catalytic nucleophile and acid/base residue that are specific to GH-16 members. Glu-147 and Glu-184 were identified as the nucleophilic residues and Glu- 152 and Glu-189 as the acid/base residues in AgaA and AgaB respectively. As expected, the overall fold of both agarases is similar to the other members of family GH-16 with known structures (Alouch et al., 2003). Town The three-dimensional structures of the two agarases helped define the amino acid residues involved in the recognition and cleavage of agarose and to understand the determinants of substrate specificity by comparison with the knownCape structures of other GH-16 members such as the K-carrageenase of P. carrageenovora and the β-1,3-1,4-glucanases of B. licheniformis and P. macerans (Alouch et al., 2003). Moreof precise details of agarose recognition did however require the production of a catalytically inactive mutant and co-crystallization with a substrate molecule. An inactive AgaA mutant was therefore constructed by site-directed replacement of the catalytic nucleophile with a serine, and the crystal structure of this AgaA mutant in complex with an agaro-octaose substrate provided the authors with detailed information on agaroseUniversity recognition in the catalytic site. The information obtained also suggested that the agarase might be able to unwind the double-helical structure prior to catalytic cleavage. Finally, alignment of the sequence of AgaA with the other family GH-16 β-agarases indicated that the residues involved in substrate binding are not conserved in the other agarases whose sequences are available. A model of AgaA in complex with an agarose substrate is shown in Figure 6 (Alouch et al., 2004).

Town

Cape

of Figure 6. Overall view of the constructed model of β-agarase A from Zobellia galactanivorans Dsij in interaction with an unwinding double helix of agarose (adapted from Allouch et al., 2004).

University

1.4. Concluding remarks and aim of this study

Most agar is extracted from species of Gelidium and Gracilaria (McHugh, 2003). Gracilaria gracilis that grows naturally at Saldanha Bay, South Africa is thus economically important as a source of agar. The Gracilaria yields from natural beds at Saldanha Bay is however unreliable and as a result the South African Gracilaria industry has experienced a number of setbacks over the years. The only way a consistent supply can be assured is by mariculture to supplement the natural harvests in order to support an agar factory (Stegenga et al., 1997). Saldanha Bay is the largest natural deep-water bay on the west coast of South Africa and is thus suitable for mariculture of Gracilaria gracilis. In 1993 the Seaweed Research Institute (SRU) conducted experimental investigations in Saldanha Bay for the mariculture of G. gracilis, where they evaluated the growth of Gracilaria on rope rafts. Even though they experienced a number of partial die-offs of the raft-cultivated Gracilaria during the summer months each year, they found that mariculture of Gracilaria in SaldanhaTown Bay is feasible but that there is potential to improve yields by technical research and development (Anderson et al.1996a). However due to the setbacks and problems (described in section 1.2) that the industry has experienced, investors are reluctant toCape establish commercial scale farms at Saldanha Bay. Therefore investors, such as Questof International and Taurus Products (Transkei) (Pty) Ltd., will only be interested in mariculture of Gracilaria gracilis at Saldanha Bay if the financially devastating Gracilaria collapses can be prevented.

Farming Gracilaria in open water provides an ideal submerged surface for the attachment of a variety of epiphytes. Jaffray et al. (1997) found that 40% of the bacterial epiphytes isolated from Saldanha Bay GracilariaUniversity have the ability to produce extracellular agarases. It is thus thought that unfavorable environmental conditions such as elevated water temperature and nutrient depletion, which occur during the summer months in the surface layer of the water in Saldanha Bay, may induce the onset of agarase production by these agarolytic bacterial epiphytes, causing them to become pathogenic to the raft-cultivated Gracilaria by utilizing the agar in the seaweed’s cell wall as a nutrient source. Jaffray and Coyne (1996) indeed developed a pathogenicity assay that satisfied Koch’s postulates. The assay demonstrated that agarolytic bacteria isolated from Saldanha Bay Gracilaria induced disease symptoms such as thallus bleaching on the seaweed, while non-agarolytic isolates did not. By using this 36 pathogenicity assay, Jaffray and Coyne (1996) identified the highly agarolytic Gracilaria gracilis pathogen, Pseudoalteromonas sp. LS2i.

The aim of this study was to characterize the bacterial pathogen, Pseudoalteromonas sp. LS2i to deepen our understanding of virulence regulation and specifically, the role of the agarase enzymes in the process of seaweed pathogen interaction. In order to determine the involvement of a specific gene in pathogenesis, the candidate gene must be mutated before it can be observed whether this mutation leads to a significant virulence defect in the host. Mutagenesis of the primary agarase gene in Pseudoalteromonas sp. LS2i following pathogenicity assays on Gracilaria gracilis will thus be performed during the course of this study. Antibodies raised against the primary agarase enzyme will also be used as tool to further explain the virulence mechanism in Gracilaria gracilis, i.e. to prove that the amount of agarase produced by this bacterium is linked to the severity of the disease symptoms in Gracilaria and to locate the agarase enzyme within an infected GracilariaTown thallus. In order to achieve this, the gene responsible for the primary agarolytic activity in Pseudoalteromonas sp. LS2i must firstly be cloned and characterized, whereafter the agarase be overexpressed in E. coli (JM109) from where it can be purified and usedCape to raise antibodies against it. Knowledge gained from this study could then hopefully help to detect and prevent disease in Gracilaria and will be invaluable when consideringof mariculture of Gracilaria in Saldanha Bay or it can also be helpful to other Gracilaria farmers cultivating Gracilaria in other areas or in on shore tanks.

University 37

2. CHAPTER 2

CLONING AND SEQUENCING OF THE AGARASE GENE FROM PSEUDOALTEROMONAS sp. LS2i

CONTENTS

2.1. Introduction ...... 39 2.2. Materials and Methods ...... 41 2.2.1. Bacterial strains and plasmids ...... 41 2.2.2. Media and culture conditions ...... 42 2.2.3. Construction of a Pseudoalteromonas sp. LS2i gene bank ...... 42 2.2.4. Identification of two agarolytic E. coli clones, pEB1 and pJB1 ...... 43 2.2.5. Restriction endonuclease mapping, deletion analysis and subcloning of the recombinant plasmids ...... Town 43 2.2.6. Southern hybridization analysis of the recombinant plasmid pEB3 ...... 44 2.2.7. Heinekoff shortening of pEB3 and pEB3a ...... 44 2.2.8. DNA sequencing of the region responsibleCape for agarase production ...... 45 2.3. Results ...... of ...... 46 2.3.1. Isolation and restriction enzyme analysis of the constructs pJB1 and pEB1 ...... 46 2.3.2. Deletion analysis and subcloning of pEB1 ...... 46 2.3.3. Southern hybridization studies ...... 54 2.3.4. DNA sequencing of the agarase gene ...... 54 2.3.5. Homology searches ...... 57 2.4. Discussion ...... University 62

Summary

A Pseudoalteromonas sp. LS2i genomic library was constructed and screened for agarolytic activity. Two agarolytic E. coli JM109 transformants, pEB1 and pJB1 were obtained. Restriction enzyme maps were constructed for the two plasmids. Based on the restriction maps of pEB1 and pJB1, a ‘common region’ was identified and it was therefore deduced that both constructs contain the same agarase gene. Deletion and subcloning of pEB1 enabled us to narrow down the region responsible for agarase production to the 1.8 kb insert of plasmid pEB3. Southern hybridization studies performed on pEB3 confirmed that the cloned DNA was of Pseudoalteromonas sp. LS2i origin. The nucleotide sequence of the pEB3 insert revealed an 1062 bp ORF, as well as putative promoter regions, a putative ribosome binding site and a putative transcriptional start point. A BLAST search of the GENBANK database showed that the ORF located in the sequenced Pseudoalteromonas sp. LS2i DNA had sequence identity to several -agarases and other proteins includedTown in the GH-16 family of glycoside hydrolases. It also showed significant sequence identity to an -agarase.

Cape of

University 39

2.1. Introduction

Even though β-agarases are functionally similar, genetic and biochemical studies of these enzymes show high degrees of heterogeneity in terms of their molecular masses, amino acid sequences, catalytic properties and substrate specificities (Otha et al., 2004c). In order to understand the molecular basis for these differences, agarase genes have been cloned and analyzed from the following genera: Agarivorans (Ohta et al., 2005), Microbulbifer (Ohta et al., 2004a; Ohta et al., 2004b; Ohta et al., 2004c), Pseudomonas (Kang et al., 2003; Lee et al., 2000a; Kong et al., 1997), Pseudoalteromonas (Belas et al., 1987; Belas, 1989), Streptomyces (Kendall and Cullum, 1984; Buttner et al., 1987), Vibrio (Sugano et al., 1993b; Sugano et al., 1994a), Zobellia (Jam et al., 2005) and Microscilla (Zhong et al., 2001). Voget and his co-workers also cloned 12 putative agarase genes by using a combined cultivation and direct cloning strategy on a mixed microbial population (Voget et al., 2003). Town A few cases have been reported where more than one agarase were cloned from a singular bacterium (Sugano et al., 1994a; Kong et al., 1997; Jam et al., 2005; Zhong et al., 2001) Sugano et al. (1994a) reported that the two agaraseCape genes, agaA and agaB that have been cloned from Vibrio sp. strain JT0107 were locatedof in tandem, while nine of the agarase genes cloned from mixed cultures as described by Voget et al (2003) were located in clusters of three. A similar observation has been made for Microscilla sp. strain PRE1 which harbors a single 101-kb circular plasmid that contains five putative agarase genes located in two clusters (Zhong et al., 2001). These findings suggest that some agarase genes may be organized in an operon. University Glycoside hydrolases are defined as a widespread group of enzymes which hydrolyze the glycosidic bond between a carbohydrate and a non-carbohydrate moiety. Henrissat (1991) proposed a new classification system for glycosyl hydrolases which was based on amino acid sequence similarities and not substrate specificity. An advantage of this system is that a protein or the translated sequence of even a domain can be classified before the enzyme activity is known (Henrissat et al., 1993). By using this system agarases can be classified into three families of glycoside hydrolases namely GH-16, GH-50 and GH-86 (afmb.cnrs- mrs.fr/CAZY) (Henrissat et al., 1996). GH-16 is the most abundant family which includes 40

-agarases from Pseudoalteromonas gracilis B9 (submitted to database as Aeromonas sp. B9) (U61972), Streptomyces coelicolor A3 (X05811), Pseudomonas sp. ND137 (AB063259 and AB200919), Pseudomonas sp. BK1 (AF534639), Pseudoalteromonas sp. CY24 (AY150179), Pseudoalteromonas atlantica ATCC 19262 (M73783), Microbulbifer sp. JAMB-A3 (AB158516), Microbulbifer sp. JAMB-A7 (AB107974), Microbulbifer sp. JAMB- A94 (AB124837), Saccharophagus degradans 2-40 (AY653535), Zobellia galactovorans Dsij (AX008608 and AX008610) as well as the -agarase from Pseudoalteromonas sp KJ2-4 (AY488029). Family GH-50 contains the two β-agarases from Vibrio sp. JT0107 (D14721 and D21202) and the β-agarase from Agarivorans sp. JAMB-A11 (AB178483), while family GH-86 includes the β-agarases from Pseudoalteromonas atlantica T6c (M22725) and Microbulbifer sp. JAMB-A94 (AB160954).

Both Jaffray (Jaffray et al., 1997; Jaffray, 1999) and Schroeder (Schroeder, 2001; Schroeder et al., 2003) have shown that Gracilaria gracilis from Saldanha TownBay supports a diverse community of bacterial epiphytes which includes many agarolytic strains. Jaffray isolated the agarolytic bacterial strain Pseudoalteromonas sp. LS2i from the surface of Saldanha Bay G. gracilis and has shown that LS2i is a pathogen of thisCape red alga ( Jaffray and Coyne, 1996; Jaffray et al., 1997; Jaffray, 1999). Schroeder confirmed a relationship between the disease symptoms exhibited by infected G. gracilis andof the agarolytic phenotype of the bacterial epiphyte Pseudoalteromonas gracilis B9 (Schroeder, 2001; Schroeder et al., 2003). In order to start investigating a relationship between the extracellular agarase produced by Pseudoalteromonas sp. LS2i and the occurrence of disease in the seaweed G. gracilis we describe the characterization of the gene responsible for the extracellular agarolytic activity in Pseudoalteromonas sp.University LS2i in this chapter. The construction of a Pseudoalteromonas sp. LS2i genomic library in Escherichia coli JM109, the screening for agarolytic transformants and the subsequent isolation of two agarolytic clones are described. The sequencing and similarity to other agarases are also described for the Pseudoalteromonas sp. LS2i gene present in one of the agarolytic E. coli clones.

2.2. Materials and Methods All media and solutions used in this study are listed in Appendix A.

2.2.1. Bacterial strains and plasmids

The bacterial strains and plasmids that were used to clone and characterize the agarase gene from Pseudoalteromonas sp. LS2i are listed in Table 1.

Table 1. Bacterial strains and plasmids Strain/plasmid Genotype/relevant feature Reference Strains E. coli JM109 RecA1 supE44 endA1 hsdR17 gyrA96 relA1 Sambrook et al. thi(lac-proAB) F’(traD36 proAB’ lacIq (1989) lacZM15) Town Pseudoalteromonas Saldanha Bay, South Africa Jaffray and Coyne sp. LS2i (1996) Plasmids Cape pBluescript SK Ampr, -galactosidase of Short et al. (1988) pEcoR251 Ampr, EcoRI endonuclease Zabeau and Stanley (1982) pEB1 pEcoR251 containing ~7.5 kb This work Pseudoalteromonas sp. LS2i genomic DNA pEB1a pEB1 with a 1.75 kb SphI-SphI fragment This work Universitydeleted from it pEB1c pEB1 with a 7 kb PstI-PstI fragment This work deleted from it pJB1 pEcoR251 containing ~9.5 kb This work Pseudoalteromonas sp. LS2i genomic DNA pEB2 pBluescript containing 2.4 kb ClaI-SacI This work fragment from pEB1 pEB3 pBluescript containing 1.8 kb EcoRV This work fragment from pEB2 42 pEB3a ~4.3 kb Heinekoff deletion mutant This work generated from the XhoI site of pEB3 pEB31 ~3.9 kb Heinekoff deletion mutant This work generated from the XhoI site of pEB3 pEB33 ~3.3 kb Heinekoff deletion mutant This work generated from the XhoI site of pEB3 pEB3a1 ~4.0 kb Heinekoff deletion mutant This work generated from the EcoRI site of pEB3a pEB3a2 ~3.5 kb Heinekoff deletion mutant This work generated from the EcoRI site of pEB3a pEB3a3 ~3.2 kb Heinekoff deletion mutant This work generated from the EcoRI site of pEB3a

Town 2.2.2. Media and culture conditions

Escherichia coli JM109 was either grown in Luria broth (LB) (Appendix A.1.1) or on Luria agar (LA) (Appendix A.1.2) at 37C. E. coli JM109Cape transformants that harbored recombinant pEcoR251 and pBluescript SK plasmids wereof grown in LB or on LA containing 100 ug/ml ampicillin (Appendix A.2.1). Wild-type Pseudoalteromonas sp. LS2i was grown in marine broth (MB) (Appendix A.1.3) or on marine agar (MA) (Appendix A.1.4) at 22C.

2.2.3. Construction of a Pseudoalteromonas sp. LS2i gene bank

Chromosomal DNA Universitywas isolated from Pseudoalteromonas sp. LS2i (Appendix B.1). In order to determine the optimum conditions (in terms of the ratio between enzyme concentration and DNA concentration) to generate the ideal size DNA fragments for a genomic library, a pilot restriction reaction was performed. This was done through digesting a fixed amount of genomic DNA with varying amounts of Sau3AI enzyme at 37C before separating the fragments by electrophoresis through a 0.8% agarose gel. The LS2i DNA was then partially digested with the restriction enzyme Sau3AI by incubating 0.22 U enzyme/2.8 ug DNA at 37C before electrophoresis on a 0.8% agarose gel (Appendix B.2). Fragments ranging between 7.0 kb and 10.0 kb were extracted from the gel using a BIO 101, Inc. Geneclean III 43 kit. The plasmid vector pEcoR251 was linearized using the restriction endonuclease Bgl II. The purified LS2i fragments were then ligated into the vector (Appendix B.3.2), and transformed into competent E. coli JM109 cells (Appendix B.4). Transformants were incubated at 37C for 16 h on LA agar plates containing ampicillin.

2.2.4. Identification of two agarolytic E. coli clones, pEB1 and pJB1

The genomic library was screened for active agarases, which are exported by E. coli JM109, by visual detection of colonies pitting the agar. Two agarolytic clones, designated pEB1 and pJB1, were isolated.

2.2.5. Restriction endonuclease mapping, deletion analysis and subcloning of the recombinant plasmids

Plasmid DNA was extracted from pEB1 and pJB1 by using a NucleobondTown AX100 kit (Macherey-Nagel, Düren). In order to determine whether the plasmids pEB1 and pJB1 contain similar LS2i DNA both plasmids were initially mapped. The following restriction enzymes were used in the construction of plasmid Caperestriction maps of pEB1 and pJB1: PstI, EcoRI, BamHI, NdeI, BclI, PvuII, HaeIII, ClaofI, Sph I, StuI, SacI, ApaI . Most of these enzymes were also used in combination with one another. (Appendix B.5).

In order to narrow down the region within pEB1 that is responsible for agarase production, deletion analysis was performed. The enzyme SphI was used to perform a complete digest and a partial digest was performed with the enzyme PstI. The resultant constructs pEB1a and pEB1c were religatedUniversity and transformed into E. coli JM109 and subsequently assayed for their ability to hydrolyze agar as described in section 2.2.4.

In order to locate the region responsible for agarase production on pEB1, the 2.3 kb ClaI-SacI fragment of pEB1 was subcloned into pBluescript SK. The resultant agarolytic construct designated pEB2, was then subjected to further restriction enzyme mapping using the following enzymes: StyI, BsteII, KpnI, SalI, BglI, SspI, NsiI, EcoRV.

The 1.8 kb EcoRV fragment of pEB2 was subcloned into pBluescript SK. The resultant construct that still contained the region responsible for agarase production was designated pEB3. Restriction enzyme digestions (completed and partial), agarose gel electrophoresis, ligation and transformation procedures were performed as described in Appendix B.5, B.2, B.3 and B.4

2.2.6. Southern hybridization analysis of the recombinant plasmid pEB3

In order to confirm that the cloned DNA of pEB3 was of Pseudoalteromonas sp. LS2i origin, the 806 bp NsiI-BsteII fragment of pEB3 was used as a probe against equal amounts of LS2i and E. coli genomic DNA in a Southern hybridazation procedure. After restriction enzyme digestion and agarase gel electrophoresis (Appendix B.5 and B.2), the 0.8 kb pEB3 fragment was extracted from the gel using a BIO 101, Inc. Geneclean III kit. The LS2i genomic DNA was digested with the restriction enzymes HindIII, SspI, BsteII, NsiI, and the E. coli genomic DNA was digested with HindIII (Appendix B.5). The genomic DNATown fragments were then separated on a 1% agarose gel. The transfer of the DNA onto a nitrocellulose membrane is described in Appendix B6. The southern hybridization procedure was performed using a NEBlot phototope kit (New England Biolabs) as wellCape as a Phototope-Star detection kit (New England Biolabs). The former kit was used toof incorporate biotin into the 0.8 kb hybridization probe, while the latter kit contains the chemiluminescent reagent (CDP-star) that was used to detect the biotinylated probe that were immobilized on the membrane

2.2.7. Heinekoff shortening of pEB3 and pEB3a

In order to obtain theUniversity nucleotide sequence of the cloned agarase gene, Heinekoff-shortening was employed to generate several deletion plasmids from pEB3 for forward sequencing and from pEB3a for reverse sequencing.

The restriction enzymes used to generate the 3’ and 5’ overhangs for forward sequencing in pEB3 were KpnI and XhoI, respectively. In pEB3a PstI was used to generate the 3’ overhang while EcoRI was used to generate the 5’ overhang for reverse sequencing. Heinekoff shortening was performed as described in Appendix B.7.

2.2.8. Sequence analysis of the region responsible for agarase production

An automated sequencer (ALFexpressTM DNA Sequencer AM version 3.01, Pharmacia Biotech) was used to obtain double stranded sequence for a 1658 bp region of pEB3. This included the agarase gene as well as 582 bp of the region upstream of the agarase’s start codon. Plasmids used for forward sequencing were pEB3 (4.8 kb), pEB3a (4.3 kb), pEB31 (3.9 kb) and pEB33 (3.3 kb), while plasmids used for reverse sequencing were pEB3a (4.3 kb), pEB3a1 (4kb), pEB3a2 (3.5kb) and pEB3a3 (3.2 kb).

All the sequence data generated was assembled using the DNASIS software version 2.1 (Hitachi Software Engineering), while the assembled sequences were analyzed using DNAMAN version 4.13 (Lynnon Biosoft). Homology searches with protein sequences were carried out using the BLAST algorithm (Altschul et al., 1990) provided by the Internet service of the National Centre for Biotechnology Information (http://www.ncbi.blast.nlm.nih.gov/BLAST/). Town

Cape of

University 46

2.3. Results

2.3.1. Isolation and restriction enzyme analysis of the constructs pJB1 and pEB1

In order to isolate an agarase gene from Pseudoalteromonas sp. LS2i, a gene bank was constructed in Escherichia coli JM109. LS2i genomic DNA was isolated and partially digested with the restriction enzyme Sau3AI. Fragments of genomic DNA ranging from 7 kb to 10 kb in size were cloned into the BglII site of the plasmid vector pEcoR251. About 4750 E. coli JM109 transformants were screened for agarolytic activity by visual detection of colonies hydrolyzing the agar; i.e. we searched for small indentations around the colonies growing on the agar. Two agarolytic colonies were isolated and the recombinant plasmids were designated pJB1 (13 kb) and pEB1 (11 kb) (Figure 1). Restriction enzyme mapping performed on the two constructs revealed that both constructs had a similar restriction enzyme pattern and that compared to the vector’s restriction enzymeTown pattern, the inserts were cloned in opposite orientations. The ‘common region’ of the two recombinant constructs was ~7.5 kb in length (which is the size of the pEB1 insert), and contained PstI, ClaI, PvuII, SacI as well as two SphI enzyme sites. Cape

2.3.2. Deletion analysis and subcloning ofof pEB1

Since pJB1 and pEB1 had similar restriction enzyme patterns, it was assumed that both constructs contained the same agarase gene and we therefore decided to only characterize the slightly smaller plasmid pEB1. Since the pEB1 construct still contained a fairly large insert of 7.5 kb, it was necessary to locate the region responsible for agarase production. Deletion analysis was thereforeUniversity performed by deleting two different fragments from the pEB1 insert and scoring for agarolytic activity. The deletion strategies that were employed in order to obtain the resultant constructs pEB1a and pEB1c are shown in Figure 2. Deletion of the 1.75 kb SphI fragment from pEB1 yielded the plasmid pEB1a, which upon transformation into E. coli JM109 still had the ability to hydrolyze the agar. Deletion of the 7 kb PstI fragment produced the construct pEB1c. E. coli JM109 transformed with pEB1c lost its agarolytic activity. From these results the agarase gene was expected to be located on either the 4.0 kb BglII-SphI fragment or the 1.9 kb SphI-BglII fragment, located on either side of the deleted 1.75 kb SphI fragment (Figure 2). 47

BglII (11000) BamHI (460) PstI (190) NdeI (695)

SphI (9100) PstI (2010)

pEB1

EcoRI (2740) Town 11000bp

BglII (3350)

SacI (7500) Cape

SphI (7350) of

PstI (4250)

ClaI (5200) PvuII (5650) University

Figure 1a. Restriction enzyme map of the recombinant plasmid pEB1. The thick blue line represents cloned Pseudoalteromonas sp. LS2i DNA and the thin line represents pEcoR251 DNA. The size of the plasmid is shown in base pairs (bp). The numbers inside the brackets indicate the various positions in bp of the restriction enzyme sites. The position of the - lactamase gene is indicated with an arrow.

BglII (13000) BamHI (460) PstI (12600) PstI (190) NdeI (695)

EcoRI (11600) ClaI (11500) BamHI (11100) PstI (2010)

EcoRI (2740) PstI (10100) pJB1

BglII (3350) Town 13000bp

ClaI (9150)

Cape PvuII (8700) of

SphI (5250)

SphI (7000) SacI (6850) University

Figure 1b. Restriction enzyme map of the recombinant plasmid pJB1. The thick blue line represents cloned Pseudoalteromonas sp. LS2i DNA and the thin line represents pEcoR251 DNA. The size of the plasmid is shown in base pairs (bp). The numbers inside the brackets indicate the various positions in bp of the restriction enzyme sites. The position of the - lactamase gene is indicated with an arrow. 49

The region responsible for agarase production was narrowed down to the 2.3 kb ClaI-SacI fragment (which included a large portion of the 4.0 kb BglII-SphI fragment of pEB1a) by the construction of pEB2 where the 2.3 kb ClaI-SacI fragment was cloned into the multiple cloning site (MCS) of pBluescript SK as depicted in Figure 3. In order to construct pEB2 the vector pBluescript was linearized with the enzymes ClaI and SacI. This procedure generated two fragments sized, 2,9 kb and 72 bp. When ligation of pEB2 was performed, a 72 bp fragment was cloned on either side between the 2.3 kb ClaI-SacI fragment and the 2.9 kb linearized vector (Figure 3).

Further restriction enzyme analysis of pEB2 identified additional restriction enzyme sites as shown in Figure 4. Using this information further subcloning of the region responsible for agarase production was performed by cloning the 1.8 kb EcoRV fragment of pEB2 into pBluescript SK (Figure 5). The resultant construct (pEB3) is shown in Figure 6. Town

NdeI pEB1 BglII BamHI Cape SacI PstI PstI EcoRI BglII PstI ClaI PvuII SphI SphI BglII (11000 of bp) NdeI BglII BamHI SacI pEB1a PstI PstI EcoRI BglII PstI ClaI PvuII SphI SphI BglII

(9250 bp) NdeI University BglII BamHI SacI pEB1c PstI PstI EcoRI BglII PstI ClaI PvuII SphI SphI BglII

(4000 bp) Figure 2. Deletion analysis of pEB1. Two different regions of Pseudoalteromonas sp. LS2i DNA were deleted in pEB1. The thick blue line represents cloned Pseudoalteromonas sp. LS2i DNA whereas the thin line represents pEcoR251 DNA. The thick red line represents the fragments deleted from pEB1a and pEB1c. The position of the -lactamase gene is indicated with an arrow.

Sph I Sph I

Sac I Bgl II Pst I Cla I Pvu II Bgl II pEB1 insert

SspI

KpnI PvuII XhoI SalI PvuI ClaI HindIII

pBluescript SK EcoRV EcoRI PstI Cla I Sac I BglI 2959bp BamHI BstXI SacI PvuII 2,3 kb ClaI – SacI fragment ori

Sac I Cla I Cla I Sac I

Town

SspI KpnI PvuI PvuII XhoI Cape SalI BglI ClaI

of SacI SphI

pEB2 ori

University

SacI ClaI PvuII

Figure 3. Construction of pEB2. The thick yellow line represents the 2.3 kb ClaI-SacI fragment cloned from pEB1 into the MCS of pBluescript SK. The positions of the - lactamase ( ) gene, the -galactosidase ( ) gene as well as the two origins of replication, i.e. M13 ori ( ) and ori ( ), in the pBluescript cloning vector are indicated. The red line represents the 72 bp fragments generated from linearizing pBluescript SK with ClaI and SacI. A 72 bp fragment was ligated into pEB2 on either side between the 2.3 kb insert and the linearized vector. 51 KpnI (648) RsaI (649) ApaI (654) AvaI (663) XhoI (663) HincII (669) SalI (669) AccI (669) ClaI (678) SspI (5221) HindIII (684) ApaLI (5086) DraIII (220) EcoRV (690) XmnI (5014) NaeI (326) EcoRI (696) AcyI (4955) PstI (702) RsaI (4898) BglI (464) AvaI (708) ScaI (4897) PvuI (495) SmaI (708) XmaI (708) PbuI (4786) PvuII (525) BamHI (714) AvaII (4779) SpeI (720) XbaI (726) MstI (4639) NotI (732) XmaIII (733) AvaII (4557) BstXI (739) BglI (4533) DsaI (742) SacIII (742) SacI (750) SphI (900) NsiI (940)

BspHI (4246)

BglI (1173) pEB2 SspI (1192)

Town

AlwNI (3937) 5337 bp

ApaLI (3840) SalI (1516) Cape

of SspI (1639)

BstEII (1746)

PvuII (3348)

SacI (3128) SacIII (3120) DsaI (3120) BstXI (3117) XmaIII (3111) NotI (3110) University XbaI (3104) SpeI (3098) SspI (2455) BamHI (3092) XmaI (3086) StyI (2800) EcoRV (2500) SmaI (3086) PvuII (2600) AvaI (3086) BstEII (2750) PstI (3080) EcoRI (3074) KpnI (2750) EcoRV (3068) HindIII (3062) ClaI (3056)

Figure 4. Restriction enzyme map of the recombinant plasmid pEB2. The thick yellow line represents the insert DNA whereas the thin line represents pBluescript SK DNA. The positions of the -lactamase gene (black), the -galactosidase gene (dark blue) as well as the two origins of replication, i.e. M13 ori (green) and ori (light blue), in the cloning vector pBluescript are indicated. The size of the plasmid is shown in base pairs (bp). The numbers inside the brackets indicate the various positions in bp of the restriction enzyme sites. 52

EcoRV PvuII StyI SacI ClaI SacI SphI BglI SalI BstEII EcoRV BstEII ClaI pEB2 insert 200bp

EcoRV EcoRV SspI 100bp 1,8kb EcoRV - EcoRV fragment KpnI PvuII XhoI SalI PvuI ClaI HindIII pBluescript SK EcoRV EcoRI PstI BglI 2959bp BamHI BstXI SacI PvuII ori

EcoRV EcoRV SspI Town

PvuI PvuII KpnI XhoI SalI BglI ClaI Cape HindIII EcoRV SphI of pEB3 BglI

ori

SalI

BstEII SacI BstXI UniversityBamHI PstI EcoRI EcoRV

Figure 5. Construction of pEB3. The thick yellow line represents the 1.8 kb EcoRV-EcoRV fragment cloned from pEB2 into the MCS of pBluescript SK. The positions of the - lactamase ( ) gene, the -galactosidase ( ) gene as well as the two origins of replication, i.e. M13 ori ( ) and ori ( ), in the cloning vector pBluescript are indicated. The red lines represent the 72 bp fragments generated from linearizing pBlueskript SK as depicted in Figure 3. 53

KpnI (648) RsaI (649) ApaI (654) SspI (4653) XhoI (663) DraIII (220) AvaI (663) ApaLI (4518) SalI (669) XmnI (4446) NaeI (326) AccI (669) AcyI (4387) HincII (669) RsaI (4330) ClaI (678) BglI (464) HindIII (684) ScaI (4329) PvuI (495) PvuII (525) EcoRV (690) PvuI (4218) EcoRI (696) PstI (702) AvaII (4211) SmaI (708) XmaI (708) AvaI (708) MstI (4071) BamHI (714) SpeI (720) AvaII (3989) XbaI (726) NotI (732) BglI (3965) XmaIII (733) BstXI (739) SacIII (742) DsaI (742) SacI (750) SphI (900) NsiI (940)

pEB3 BspHI (3678)

BglI (1173)

SspI (1192)

Town 4769bp

AlwNI (3369)

Cape ApaLI (3272) SalI (1516) of

SspI (1639)

BstEII (1746)

SacI (2560) DsaI (2552) SacIII (2552) BstXI (2549) PvuII (2780) XmaIII (2543) NotI (2542) UniversityXbaI (2536) SpeI (2530) Bam HI (2524) SspI (2455) XmaI (2518) SmaI (2518) AvaI (2518) PstI (2512) EcoRI (2506) EcoRV (2500)

Figure 6. Restriction enzyme map of the recombinant plasmid pEB3. The thick yellow line represents the insert DNA whereas the thin line represents pBluescript SK DNA. The positions of the -lactamase gene (black), the -galactosidase gene (dark blue) as well as the two origins of replication, i.e. M13 ori (green) and ori (light blue), in the pBluescript cloning vector are indicated. The size of the plasmid is shown in base pairs (bp). The numbers inside the brackets indicate the various positions in bp of the restriction enzyme sites. 54

2.3.3. Southern hybridization studies

The 806 bp NsiI-BsteII fragment from pEB3, located within the region responsible for the production of the agarase enzyme, was isolated and used as a probe against Pseudoalteromonas sp. LS2i chromosomal DNA. The Southern hybridization study, to verify that the agarase gene carried by the recombinant plasmid pEB3 was of LS2i origin, is shown in Figure 7. Indeed the 806 bp NsiI-BsteII fragment did not hybridize to E. coli chromosomal DNA (lane 1) that served as a control but hybridized to the following fragments of LS2i chromosomal DNA: a ~3.5 kb Hind III fragment (lane 2), two SspI fragments of 1.0 kb and 816 bp in size (lane 3) and a 806 bp NsiI-BsteII fragment (lane 4). Since there are no HindIII sites in the 1,8 kb insert of the pEB3 construct, a hybridization band larger than 1.8 kb i.e. 3.5 kb was expected in lane 2. According to the map of pEB3 (Figure 6) digestion with the restriction enzyme SspI should yield a fragment of unknown size, which is the 1.0 kb fragment, as well as a 816 bp band that is in fact present in lane 3 and a 447 bp fragment. Due to its small size the 447 bp fragment was probably electrophoresedTown off the gel. The 806 bp NsiI-BsteII fragment shown in lane 4 was as expected also present in lane 5. The cloned agarase is thus present as a single copy on the chromosome of Pseudoalteromonas sp. LS2i as no unknown bands were present in any of the lanes.Cape of 2.3.4. DNA sequencing of the agarase gene

For sequencing purposes, insert DNA of pEB3 was sequentially deleted to generate constructs for forward sequencing while the first deletion plasmid obtained from shortening pEB3, designated pEB3a, was sequentially deleted to generate constructs for reverse sequencing. The plasmid,University pEB3a is approximately 300 bp shorter than pEB3. The 300 bp that were deleted in pEB3a included one of the duplicate 72 bp fragments that were also cloned into pEB2 as described in 2.3.2. Because of the way the pEB2 plasmid was constructed with the 72bp fragment cloned on either side between the vector DNA and the LS2i DNA, all the enzyme sites that could be used for creating overhangs (required by exonuclease III for shortening) for reverse sequencing were present in duplicate in pEB3 and therefore pEB3a had to be used for reverse sequencing. Various nested deletions were sequenced. Double stranded DNA sequence was obtained for 1658 bp of the 1810 bp insert of pEB3.

1 2 3 4 5

3.5kb

1.0kb

816bp 806bp

Town Figure 7. Southern hybridization of the 806 bp Nsi I-Bste II restriction fragment of pEB3 against chromosomal DNA isolated from Pseudoalteromonas sp. LS2i and E. coli as well as plasmid DNA from pEB3. E. coli DNA was digested with Hind III (lane1) and Pseudoalteromonas sp. LS2i DNA was digested with Hind III (lane 2), Ssp I (lane3) and Nsi I-Bste II (lane 4). The construct pEB3 was digestedCape with Nsi I-Bste II (lane 5) of

A single complete open reading frame (ORF) was identified within the 1658 bp double stranded nucleotide sequence (Figure 8). The ORF begins with an ATG initiation codon at position 583 and ends with a TGA stop codon at position 1645. It therefore codes for a protein of 354 amino acids, with a putative size of 38,9 kDa. The putative ribosome-binding site, otherwise knownUniversity as the Shine Dalgarno sequence, AAGAGG was observed at position 572-577 just upstream from the initiation codon (Shine and Dalgarno, 1975). A putative –35 promoter region, TTGACA, was identified at position 490-495. Since the –10 promoter and the –35 promoter regions in the majority of cases has an interregion spacing of 16-18 bp, a putative –10 promoter region could be the sequence, CTTATT, 17 bases downstream of the putative –35 promoter, at position 513-518 (Harley and Reynolds, 1987). The cytosine at position 528, which is exactly 10 bases downstream of the putative –10 promoter, could be the transcriptional start point (Harley and Reynolds, 1987) (Figure 8).

1 GCTATTGTTAACTTTGAAGGTGAGGGCGGTGTCACCTAGTTGAGGAAATTCTACTAATGC

61 ATTAAAAGTGTGTTTGCCGGTGCGGGCTATGTTGAGGTTTTCAAATAATAGCGTGCCGTT

121 GGCATAAATATTTAAAGTTGCGTAGTCACCGGGGTTTGATATGACAAGGTTTAATTTTTT

181 AATCTTACTCCCAGCCACATGATAATTAGTAGTTAGTGGTTCAGTCGTTTTAATAAGGTG

241 TGTTTGTAGATTCTGTAAATGAACTGAATTGACATTATTTTTGCTATCACTACACCCCGT

301 CACCATTAGCGCACCAAAAGTTAGCATTATAAGAGGGGGTATCCACTTAATTATTCTATT

361 TTTTAGCATCATGAGAGCCCTTCCATTGTTGTATTATCTTTTGATTGAGACGCGCATAAC

421 CAATGAGTTATACACAACGTGATGCAAAACTGACCATAGCGCATTCAACTCATTGTTAAA

481 AGTATTTTGTTGACAATTTATTTATTTTGTACCTTATTTGTACTAATCTTTGTTGGCTAC -35 -10 541 ATTTTGTGGCTAAAATGACAACATTAACTCAAAGAGGCAAAAATGAACAAAACAACATTG 1 RBS M N K T T L

601 TTTATCGGGTGTTTACTCACTACTACCAACTTGTTTGCAAATGATTGGGACTCAATCCCT 7 F I G C L L T T T N L F A N D W D S I P

661 CTACCAGTTTCACCTGATGATGGAAAAGTATGGCAGCTACAAGAAGACTACTCAGATTCA 27 L P V S P D D G K V W Q L Q Town E D Y S D S

721 TTTAATTACACTGGGAAACCAGCGGCATTTACCAGTAAATGGAATGATACTTACTTTAAT 47 F N Y T G K P A A F T S K W N D T Y F N

781 AGTTGGACAGGGCCTGGCTTGACCTATTGGCAGCAGGACGAGTCTTGGGTTGCAGACGGT 67 S W T G P G L T Y W CapeQ Q D E S W V A D G

841 AACCTTATAATTAGTGCTTCGCGTCGTGCCGGCACAGATAAGGTGAACGCAGGGGTTATCof 87 N L I I S A S R R A G T D K V N A G V I

901 ACCTCGAAAACAAAAGTTAGTTTTCCAATCTTTTTAGAGGCAAATATTAAGGTCAGTAAT 107 T S K T K V S F P I F L E A N I K V S N

961 CTGGAATTATCTTCAAATTTTTGGCTACTTAGTGACAATGATGAACGCGAAATTGATGTG 127 L E L S S N F W L L S D N D E R E I D V

1021 CTAGAGGTATACGGTGGGGCACGTGATGATTGGTTTGCTAAAAACATGTCGACTAACTTT 147 L E UniversityV Y G G A R D D W F A K N M S T N F

1081 CATGTATTTATTCGTGATCAACAATCTAATCAAATAATTAGTGATTACAACGACCAAACG 167 H V F I R D Q Q S N Q I I S D Y N D Q T

1141 CATAATACGCCTAGTTGGGGTACTTATTGGCGTGAGGGTTTTCATCGTTTTGGCGTGTAT 187 H N T P S W G T Y W R E G F H R F G V Y

1201 TGGAAAAGCCCAACAGAAGTCACATTTTACATAGATGGCCAGCAAACGCCTGATGGTTCG 207 W K S P T E V T F Y I D G Q Q T P D G S

1261 TGGGCACAGGTGGTAATGAAAGACAAAGACTACACTGGGGCGACGTTAAAAAAGGACACA 227 W A Q V V M K D K D Y T G A T L K K D T

1321 CATAATATGGATCAATCTGCTTATATCATTATTGATACAGAAGATCATGATTGGCGCTCA 247 H N M D Q S A Y I I I D T E D H D W R S

1381 GAGGCTGGAAATATTGCTACAGATGCCGATTTGGCGGACGGTAGTAAAAATAAAATGTAT 267 E A G N I A T D A D L A D G S K N K M Y

1441 GTCGATTGGGTGCGAGTTTATAAACCTGTTAATGCGCCGAACACAAACAATGTTAGTAAT 287 V D W V R V Y K P V N A P N T N N V S N

1501 GGCGGGCAGCTCAAAGCTAAGCATAGTCAAAAGTGTATTGATATAACAGGTGGCGCAATG 307 G G Q L K A K H S Q K C I D I T G G A M

1561 AGTAATGGCTCTTATTATCAGCAGTGGGGTTGTGGCTCTGATAATACTAACCAACAATTT 327 S N G S Y Y Q Q W G C G S D N T N Q Q F

1621 AACCTTGTTGAGTTAAGTAATGCATGAATATGCAATTA 347 N L V E L S N A *

Figure 8. Nucleotide sequence of the coding region of the agarase gene cloned from Pseudoalteromonas sp. LS2i and the deduced amino acid sequence. The putative –10 and –35 promoter regions are underlined, while the putative tsp and the initiation codon, ATG, are in bold. The Shine Dalgarno sequence is underlined and labeled as RBS (ribosomal binding site). The termination codon, TGA, is indicated with an asterisk.

2.3.5. Homology searches Town

The putative protein produced by the 1062 bp ORF, shares sequence identity with a number of other family GH-16 agarase type proteins (TableCape 2) as well as several non-agarase members of the family GH-16 (Table 3). This information was obtained through a BLAST search of the translated sequence of the 1062of bp ORF against both the protein database (blastx) and the translated sequence database (tblastx) of the GENBANK database. The ORF was found to have the greatest sequence similarity, i.e. 76% and 68% at the DNA and amino acid levels, respectively to an extracellular agarase precursor (AY150179) from Pseudoalteromonas sp. CY24. Several other β-agarases as well as an -agarase from Pseudoalteromonas sp.University KJ 2-4 (Table 2) were identified with significant amino acid identity to the putative agarase cloned from Pseudoalteromonas sp. LS2i (E-values of 4e-148 to 2e-46). Several other family GH-16 enzymes (Table 3) also appeared on the blast search but except for the Endo-β-1,3-glucanase precursor from the archaea Pyrococcus furiosus (E-value of 7e- 06) had poor E-values. The short sequences of similarity of the latter enzymes were essentially limited to a few conserved regions as can be seen from the multiple sequence alignment depicted in Figure 9.

The multiple sequence alignment showed significant sequence similarities occurring at specific regions of the peptide chain, with Glu143 and Glu148 of the 1062 bp ORF (Figure 9) 58 from Pseudoalteromonas sp. LS2i aligning with the two glutamic acid residues that are strictly conserved in family 16 hydrolases as described for Glu160 and Glu165 of the Kappa- Carrageenase from Zobellia galactanivorans (Barbeyron et al., 1998) as well as Glu147 and Glu152 of β-agarase A and Glu184 and Glu189 of β-agarase B from Zobellia galactanivorans (Allouch et al., 2003).

Table 2. Similarity between the putative protein expressed by the cloned agarase gene from Pseudoalteromonas sp. LS2i and agarases from other bacteria Accession Protein/Organism Sequence identity number (%) AY150179 Extracellular agarase precurser (AgaA)/ 68 Pseudoalteromonas sp. CY24 ABO63259 Agarase (AagA)/ Pseudomonas sp. ND137 Town 42 AB124837 Agarase (AgaA)/ Microbulbifer sp. JAMB-A94 41 AF098955 β-agarase B precursor (AgaB)/ Zobellia 40 galactanivorans Cape AF098954 β-agarase A precursor (AgaA)/ Zobellia 39 galactanivorans of AB158516 Agarase (agaA3) / Microbulbifer sp. JAMB-A3 38 AY653535 -agarase I (agaB) / Saccharophagus degradans 38 2-40 X05811 Extracellular agarase precurser(dagA) / 38 StreptomycesUniversity coelicolor A3(2) AB107974 Agarase (AgaA7)/ Microbulbifer sp. JAMB-A7 37 U61972 -agarase (aagA) / Pseudoalteromonas gracilis 37 B9 (submitted as Aeromonas) M73783 -agarase I (dagA) / Pseudoalteromonas atlantica 36 AY488029 -agarase / Pseudoalteromonas sp. KJ 2-4 36 AF339846 Putative -agarase precursor (MS116) / 35 Microscilla sp. PRE1 plasmid psD15

Table 3. Similarity between the putative protein expressed by the cloned agarase gene from Pseudoalteromonas sp. LS2i and non-agarase proteins from other bacteria Accession Protein/Organism Sequence identity number (%) CP000282 Arabinogalactan endo-1,4-β-galactosidase / 32 Saccharophagus degradans 2-40 ABO38772 Endo--galactosidase C / Clostridium 25 perfringes AF007559 Kappa-carrageenase precurser / Zobellia 24 galactanivorans AF013169 Endo-β-1,3-glucanase precursor / Pyrococcus 23 furiosus ABO78775 1,3-(1,3;1,4)--D-glucan 3(4)-glucanohydrolase 23 / Bacillus circulans Town AF052745 -1,3-glucanase II / Cellulosimicrobium 23 cellulans LLG109 Cape of

University 60

Ls2i(aga) 25 IPLPVSPDDGKVWQLQEDY....SDSFNYTGKP....AAFTSKWNDTYF.NSWTG 70 P.CY(aga) 29 IPIPAELDPGQSWELQESY....SDSFNYSGKP....SSFTSKWKDAYF.HNWTG 74 P.ND(aga) 20 .DWDNTPVPANAGNGKVWELQAVSDDFNYSSSLDNYHSEFTRRWHEGFI.NPWTG 72 38 YPVPAAPGGNRSWQLLPSH....SDDFNYTGKP....QTFRGRWLDQHK.DGWSGI 83 M.A94(aga) Z(agaA) 28 VPANPGNGMTWQLQDNV...... SDSFNYTSSEGNRPTAFTSKWKPSYI.NGWTG• I 75 M.A7(aga) 21 .DWDGTPVPADAGPGNTWELHPLSDDFNYSAPASGKSATFFERWSEGFI.NPWLG 73 M.A3(aga) 21 .DWDNIPVPADAGAGNTWELHSLSDDFNYAAPPVGKSATFFERWSEGFI.NPWLGI 73 Z(agaB) 58 VDWKDIPVPADAGPNMKWEFQEISDNFEYEAPADNKGSEFLEKWDDFYH.NAWAG 111 S(agaB) 16 .DWDGIPVPADPGNGNTWELQSLSDDFNYAAPANGKRTTFYSRWSEGFI.NAWLG 68 M.PRE(aga) 43 VPASAGQGKTWQLQSAA...... SDDFNYTFNETSQLTNFGSNKWYNFYHNGWDGI I • 91 S(dagA) 38 YPVPAAPGGNRSWQLLPSH....SDDFNYTGKP....QTFRGRWLDQHK.DGWSG 83 A(aagA) 23 .DWDAYSIPASAGSGKTWQLQTVSDQFNYQAGTSNKPAAFTNRWNASYI.NAWLG 75 P(dagA) 23 .DWSSFSIPAQAGAGKSWQLQSVSDEFNYIAQPNNKPAAFNNRWNASYI.NAWLG 75 P.KJ(aga) 23 .DWSPFSIPAQAGAGKSWQLQSVSDEFNYIAQPNNKPAAFNNRWNASYI.NAWLGI I 75 Glucanase(P) 56 GSEVNKEYWTFEKGNGIAYGIPGWGNGELEYYTE...NNTYIVNGTLVIEARKEI 107 Galactos.(C) 402 IDENKWTIIDG.MV...... NHGAIYNRGAVSIKKDGNNSYLA.INTKNFNSTEE 448 Glucanohydr. 44 LNRANWTPEIG.TG...... SGGWGNNELQYYTDRAQNVQVTG.GNLVITAQKE.I 89 Glucanase(C) 68 GSAPNPAVWNHETG...... AHGWGNAELQNYTASRANSALDGQGNLVITARRE. 115 Carrageenase 48 SDEFNKNDPDWAKWIKTGNLPNTS...... AWKW 75 Galactos.(S) 45 YTFYNDAGQQQ.DV...... LQILKDHGMDSIRLRVWVNPAGGWYSSINDVIEKA• • 92 Ls2i(aga) 71 P.....GLTYWQQ 78 79 DESWVA.DGNLIISASRRAGTDK.....VNAGVI 106 P.CY(aga) 75 P.....GLTYWSS 82 83 DESWVG.DGNLIISASRRQGTNQ.....VNAGVV 110 P.ND(aga) 73 P.....GLTEWIDI 80 81 GHAYVT.DGNLGIAATRKPGTDK.....VRAGSI 108 M.A94(aga) 84 P.....ANSLYSA 91 92 RHSWVA.DGNLIVEG.RRAPDGR.....VYCGYV 118 Z(agaA) 76 P.....GSTIFNA 83 84 AQAWTN.GSQLAIQAQPA.GNGK.....SYNGII 110 M.A7(aga) 74 P.....GETEYYG 81 82 PNSSVE.SGNLVIKASRKAGTTK.....IHAGAI 109 M.A3(aga) 74 P.....GETEYYA 81 82 PNSYVE.GGNLVIKASRKPGTIK.....VHTGAI 109 Z(agaB) 112 P.....GLTEWKR 119 120 DRSYVA.DGELKMWATRKPGSDK.....INMGCI 147 S(agaB) 69 P.....GQTEFYG 76 77 PNASVE.GGHLIIKATRKPGTTQ.....IYTGAI 104 M.PRE(aga) 92 P.....GTTYWQY 99 100 NHVSVS.GGNLVLRASRNPSTTKMGVPGVNAGCI 132 S(dagA) 84 P.....ANSLYSAI 91 92 RHSWVA.DGNLIVEG.RRAPDGR.....VYCGYV 118 A(aagA) 76 P.....GDTEFSS 83 84 GHSYTT.GGALGLQATEKAGTNK.....VLSGIV 111 P(dagA) 76 P.....GDTEFSA 83 84 GHSYTT.GGALGLQATEKAGTNK.....VLSGII 111 P.KJ(aga) 76 P.....GDTEFSA 83 84 GHSYTT.GGALGLQATEKAGTNK.....VLSGIITown 111 Glucanase(P) 108 ITDPNEGTFLYT.I 119 130 FSPPVVVEARIKLPKGKGLWPAF.....WMLGSN 158 Galactos.(C) 449 LIKAVGVDNYLGQ 461 482 FQFGRM.AVRAKVNDSQGIWPAI.....WML... 506 Glucanohydr. 90 ...SYGGMN.YT. 97 108 FTYGKV.EARIKLPSGQGLWPAF.....WMLGSN 135 Glucanase(C) 116 ...... GDGSYT.I 121 132 PQYGRI.EARIQIPRGQGIWPAF.....WMLGGSII 159 Carrageenase 76 N.....NQKNVKI 83 92 MRHNANNTPPD...... GGTY.....FTSGIF 112 Galactos.(S) 93 QRAKAAGMRIMID• 105 106 FHYSDS.WADPGKQYKPAAWTNY.....TLDGLM• .. 133 Cape * * Ls2i(aga) 107 TSKTKVSFPIFLEANIKVSNLEL.SSNFWLLSDNDER...... EIDVLEVY 150 P.CY(aga) 111 TSKTKVKYPIFLEANIKVSNLEL.SSNFWLLSENDQR...... EIDVLEVY 154 P.ND(aga) 109 TSHDTFTYPLYVETKAKISKLVL.ASDVWLLSADSTQ...... EIDVLEAY 152 M.A94(aga) 119 TSRTPVEYPLYTEVLMRVSGLKL.SSNFWLLSRDDVN...... EIDVIECYof 162 Z(agaA) 111 TSKNKIQYPVYMEIKAKIMDQVL.ANAFWTLTDDETQ...... EIDIMEGYII. · 154 M.A7(aga) 110 HSNESVTYPLYMEARVQVTNLTM.ANAFWLLSSDSTQ...... EIDVLESY 153 M.A3(aga) 110 HSKESMTYPLFMEARVKITNLTL.ANAFWLLSSDSTE...... EIDVLESY 153 Z(agaB) 148 TSKTRVVYPVYIEARAKVMNSTL.ASDVWLLSADDTQ...... EIDILEAY 191 S(agaB) 105 HSNESFTYPLYLEARTKITNLTL.ANAFWLLSSDSTE...... EIDVLESY 148 M.PRE(aga) 133 TSNNRVKYPVFVEASVSVANIAL.ASDVWLLSPDDTQ...... EIDIIECY 176 S(dagA) 119 TSRTPVEYPLYTEVLMRVSGLKL.SSNFWLLSRDDVN...... EIDVIECYII I I 162 A(aagA) 112 SSKATFTYPLYLEAMVKPSNNTM.ANAVWMLSSDSTQ...... EIDAMESY 155 P(dagA) 112 SSKATFTYPLYLEAMVKPTNNTM.ANAVWMLSADSTQ...... EIDAMESY 155 P.KJ(aga) 112 SSKATFTYPLYLEAMVKPTNNTM.ANAVWMLSADSTP...... EIDAMESY 155 Glucanase(P) 159 IREVGWPNCGEIDIMEFLGHEPRTIHGTVHGPGYSGS...... KGITRAYTII I I 203 Galactos.(C) 507 ..SQDETGHDEIDVLEYLGQDPW.GAWTTNHFG...... ILGKNKA 543 Glucanohydr. 136 ISSVGWPKSGEIDIMERVNNNPY.VNGTVHWDA...... GGHADFG.. I 174 Glucanase(C) 160 FPGTPWPSSGEIDIMENVGFEPHRVHGTVHGPGYSGG...... SGITGMYQ• • University• • •• • 204 Carrageenase 113 KSYQKFTYGYFEAKIQGADIGEG.VCPSFWLYSDFDYSVANGETVYSEIDVVELQ 166 Galactos.(S) 134 SAVWWHTYDSLVALKNAGITPEW.VQVGNETNN...... GMLWEEG• . _. -I 172 Ls2i(aga) 151 GGAR..DDW.....FAKN..MSTNFHVFIRDQQSNQIISDYNDQTHNTPSWGTYW•• 196 P.CY(aga) 155 GGAR..QDW.....FAKN..MSTNFHVFFRNND.NSISSDFNDQTHNTPTWGNYW• 199 P.ND(aga) 153 GSDRAGQEW.....FAER..IHLSHHVFIRDPF.QDYQPTDAGSWY.TDGQGTVWI I I 198 M.A94(aga) 163 GNES..LHG...... KH..MNTAYHIFQRNPF.TELARSQKGYFADGSYGYNGE• 205 Z(agaA) 155 GSDRGG.TW.....FAQR..MHLSHHTFIRNPF.TDYQPMGDATWY.YNG.GTPW 198 M.A7(aga) 154 GSDRPSETW.....FDER..LHLSHHVFIREPF.QDYQPKDDGSWY.PNPNGGTW 199 M.A3(aga) 154 GSDRPSETW.....FDER..LHLSHHVFIREPF.QDYQPKDAGSWY.PNPDGGHW 199 Z(agaB) 192 GADYSESAGKDHSYFSKK..VHISHHVFIRDPF.QDYQPKDAGSWF.EDG..TVW 240 S(agaB) 149 GSDRATETW.....FDER..LHLSHHVFIRQPF.QDYQPKDAGSWY.PNPDGGTW 194 M.PRE(aga) 177 GGAGSNNAY.....FAQF..IHLSHHSFVRNPF.QDYQPRDLNSWW.GKSGVSSW 222 S(dagA) 163 GNES..LHG...... KH..MNTAYHIFQRNPF.TELARSQKGYFADGSYGYNGEI I 205 A(aagA) 156 GSDRVGQEW.....FDQR..MHVSHHVFIREPF.QDYQPKDAGAWV.YNSGETYR 201 P(dagA) 156 GSDRIGQEW.....FDQR..MHVSHHIFIRDPF.QDYQPKDAGSWV.YNNGETYR 201 P.KJ(aga) 156 GSDRIGQEW.....FDQR..MHVSPHVFIRDPF.QDYQPKDAGSWV.YNNGETYR 201 Glucanase(P) 204 ....LPEGVPDFTED.FH..VF.GIVWYPDKI...... KWI 229 Galactos.(C) 544 SNGIRNSNYE.AWSQDFH..VF.EVEWDPEFI...... KWI 573 Glucanohydr. 175 ••....RVSGNL.DFSQ.FH..VY.SIEWDSKYI...... RW • 199 Glucanase(C) 205 ....HPQGWS.FADT.FH..TF.AVDWKPGEI...... TW• • 229 Carrageenase 167 QFDWYEGHQ....DDIYDMDLNLHAVVKENGQG.VWKRPKMYPQEQ.LNKWRAMDI 215 Galactos.(S) 173 ....RASANM.QNYA.WL..VN.SGYDAVKEVF.PNTKAVVHLANC.HDNANFRW- . I. • _ •• I 216 • 61

Ls2i(aga) 197 ...... R E G F H ...... R F G V Y W K S P T E V T F Y I D G Q Q T P D G S 226 P.CY(aga) 200 ...... R E G F H ...... R F G V Y W K S P T E V T F Y I N G Q K T T K G A 229 P.ND(aga) 199 ...... S D D F H ...... R I G V H W K D P W N L D Y Y I D G Q L V R S V . 227 M.A94(aga) 206 T G Q V F G D G A G Q P L L R N G F H ...... R Y G V H W I S A T E F D F Y F N G R L V R R L N 249 Z(agaA) 199 ...... R S A Y H ...... R Y G C Y W K D P F T L E Y Y I D G V K V R T V . 227 M.A7(aga) 200 ...... R D Q W I ...... R I G T Y W V D P W T L E Y Y V N G E H V R T V . 228 M.A3(aga) 200 ...... R D Q F F ...... R I G V Y W I D P W T L E Y Y V N G E H V R T V . 228 Z(agaB) 241 ...... N K E F H ...... R F G V Y W R D P W H L E Y Y I D G V L V R T V . 269 S(agaB) 195 ...... R D Q F F ...... R I G V Y W I D P W T L E Y Y V N G E L V R T V . 223 M.PRE(aga) 223 ...... G D Y C W N N G N R K Y V R V G V N W V G P K H F E Y Y I D G E L V R V L Y 260 S(dagA) 206 T G Q V F G D G A G Q P L L R N G F H ...... R Y G V H W I S A T E F D F Y F N G R L V R R L N 249 A(aagA) 202 ...... N K . F R ...... R Y G V H W K D A W N L D Y Y I D G V L V R S V . 229 P(dagA) 202 ...... N K . F R ...... R Y G V H W K D A W N L D Y Y I D G V L V R S V . 229 P.KJ(aga) 202 ...... N K . F R ...... R Y G V H W K D A W N L D Y Y I D G V L V R S V . 229 Glucanase(P) 230 ...... Y V D G T ...... F Y H ...... 237 Galactos.(C) 574 ...... Y I D G K ...... E V F ...... 581 Glucanohydr. 200 ...... F V D G Q ...... Q F N ...... 207 Glucanase(C) 230 ...... F V D G Q ...... Q F H ...... 237 Carrageenase 216 ...... P S K D F ...... H I Y G C E V N Q N E I I W Y V D G V E V A R K . 244 Galactos.(S) 217 ...... I F D G L ...... Q A N G G K W D V I G A S I Y P T N A S G Y S W . 245

Ls2i(aga) 253 . A Y I I I D T E D H D W R S E ...... A G N I A . T D A D L A D G S K N K M Y V D W V R V Y K P V N 297 P.CY(aga) 256 . M F I I L D T E D H S W R S E ...... A G H I A . T D A D L A D G D K N K M Y V D W I R V Y K P T G 300 P.ND(aga) 247 P M H L I I N T E D Q D W R S D ...... N G I S P . T D A E L A N T N K S I Y W V D W I R V Y K P V D 292 M.A94(aga) 266 . M H L I L N T E S H Q W R V D ...... R G I E P . T D A E L A D P S I N N I Y Y R W V R T Y Q A V . 309 Z(agaA) 246 A T N I I I D C E N Q T D W R P A A T Q E ...... E L A D D S K N I F W V D W I R V Y K P V A 288 M.A7(aga) 248 P M Q V I F D A E H Q P W R D T ...... Q G T A P P T D E E L A D P S R N K F L V D W V R F Y K P V P 294 M.A3(aga) 248 P M Q V I F D A E H Q P W R D A ...... Q G T A P P T D E E L A D P S R N K F L V D W V R F Y K P V A 294 Z(agaB) 300 E M D I I I N T E D Q T W R S S P A S G L Q S N T Y T P . T D N E L S N I E N N T F G V D W I R I Y K P V E 352 S(agaB) 243 P M Q V I F D A E H Q P W R D E ...... Q G T A P P T D A E L A D S S R N Q F L I D W V R F Y K P V A 289 M.PRE(aga) 333 E L D I I I N V E S Q N W H V E A G R T P S ...... D A D L N D P A K N K M K V D W I R V Y K P V T 378 S(dagA) 266 . M H L I L N T E S H Q W R V D ...... R G I E P . T D A E L A D P S I N N I Y Y R W V R T Y Q A V . 309 A(aagA) 249 P M H I I L D M E H Q P W R D V ...... K . . . P . N S T E L A D S N K S I F W I D W V R V Y K A N . 290 P(dagA) 249 P M H I I L D M E H Q P W R D V ...... K . . . P . N A S E L A D P N K S I F W V D W I R V Y K A Q . 290 P.KJ(aga) 249 P M H I I L D M E H Q P W R D V ...... K . . . P . N A S E L A D P N K S I F W V D W L R V Y K A Q . 290 Glucanase(P) 239 ...... Town...... E V T K E Q V E A M G 249 Galactos.(C) 283 ...... Q S T Q G R D D G R D 293 Glucanohydr. 209 ...... E F Y I E N G T G N T 219 Glucanase(C) 239 ...... R V T R A S V G A N A 249 Carrageenase 287 ...... E K L S D I P T S M Y V D Y V R V W E K S A 308 Galactos.(S) 288 ...... A Q N A G A T G V F Y W E P Q A S N W Q G Y T 310

Cape Figure 9. Multiple sequence alignment of agarases and other representatives of family GH- 16. The amino acids highlighted in pink, blueof and yellow represent sequence homology of >75%, >50% and >33%, respectively. The asterisks indicate the positions of the strictly conserved Glu residues in family 16 hydrolases. Amino acid sequence codes are LS2i(aga): β-agarase from Pseudoalteromonas sp. LS2i; P.CY(aga): Agarase (AgaA) from Pseudoalteromonas sp. CY24; P.ND(aga): Agarase (AagA) from Pseudomonas sp. ND137; M.A94(aga): Agarase (AgaA) from Microbulbifer sp. JAMB-A94; Z(agaA): β-agarase A (AgaA) from Zobellia galactanivorans; M.A7(aga): Agarase (AgaA7) from Microbulbifer sp. JAMB-A7; M.A3(aga): Agarase (agaA3) from Microbulbifer sp. JAMB-A3; Z(agaB); β- agarase B (AgaB) fromUniversity Zobellia galactanivorans ; S(agaB): -agarase I (agaB) from Saccharophagus degradans; M.PRE(aga): -agarase (MS116) from Microscilla sp. PRE1; S(dagA): Agarase (dagA) from Streptomyces coelicolor; A(aagA): -agarase (aagA) from Pseudoalteromonas gracilis B9 (submitted as Aeromonas); P(dagA): -agarase I (dagA) from Pseudoalteromonas atlantica; P.KJ(aga): -agarase from Pseudoalteromonas sp. KJ 2-4; Glucanase(P): Endo-β-1,3-glucanase from Pyrococcus furiosus; Galactos.(C): Endo-- galactosidase C from Clostridium perfringes; Glucanohydr.: 1,3-(1,3;1,4)--D-glucan 3(4)- glucanohydrolase from Bacillus circulans; Glucanase(C): -1,3-glucanase II from Cellulosimicrobium cellulans; Carrageenase: Kappa-carrageenase precurser from Zobellia galactanivorans; Carrageenase: Kappa-carrageenase precurser from Zobellia galactanivorans. 62

2.4. Discussion

A Pseudoalteromonas sp. LS2i genomic library was constructed in E. coli JM109. About 4750 colonies were screened for agarolytic activity by visual inspection of the agar surrounding the transformants for signs of pitting. Two agarolytic E. coli JM109 transformants, pEB1 and pJB1, were isolated. The two clones were subjected to restriction enzyme mapping, which revealed a 7.5 kb common region based on the restriction enzyme patterns. It was therefore concluded that both constructs harbored the same agarase gene and further work was continued on plasmid pEB1 which is 11kb in size.

Deletion analysis of pEB1 gave an indication of the location of the agarase gene, allowing a 2.3 kb fragment of pEB1 to be subcloned into pBluescript SK. The resultant construct still contained the region responsible for agarase production and was designated pEB2. Additional enzyme sites were identified on pEB2 through further restrictionTown enzyme mapping, which made the subcloning of a 1.8 kb insert into pBluescript SK possible. The subsequent plasmid, pEB3, contained the agarase-encoding region. Southern hybridization studies performed on pEB3 confirmed that the cloned agarase was of PseudoalteromonasCape sp. LS2i origin and that the gene is present as a single copy on the chromosomeof of Pseudoalteromonas sp. LS2i.

Heinekoff shortening performed on pEB3 as well as on the first deletion plasmid obtained from shortening pEB3, i.e. pEB3a, resulted in sequentially deleted plasmids from both ends of the pEB3 insert DNA. The resultant deletion plasmids were sequenced and assembled. The completed 1658 bp double stranded sequence was then analyzed. Upon translation of this DNA sequence a putativeUniversity protein of 354 amino acids was identified. A BLAST search of the GENBANK database revealed that the ORF situated in the Pseudoalteromonas sp. LS2i DNA cloned into pEB3 had sequence identity, which ranged between 35% and 68%, to a number of -agarases as well as a 36% homology to an -agarase (Table 2). The ORF also had a sequence similarity that ranged between 23% and 32% to several other types of proteins (Table 3). An extracellular agarase precursor (AY150179) cloned from the marine bacterium Pseudoalteromonas sp. CY24 had the greatest sequence identity of 68% to the ORF from Pseudoalteromonas sp. LS2i.

Agarases can be separated into three families of glycoside hydrolases namely GH-16, GH-50 and GH-86 (see section 2.1) with most agarases being classified into family GH-16 (afmb.cnrs-mrs.fr/CAZY). Based on their substrate specificities, the GH-16 family can be further divided into several subgroups i.e. agarases, laminarinases, lichenases, K-carrageenases, xyloglucan endotransferases and endo-β-galactosidases. The ORF sequenced from Pseudoalteromonas sp. LS2i displayed no sequence similarity to members of families GH-50 and GH-86, but showed significant sequence similarity to other agarases belonging to family GH-16 (Table 2). The sequence identity between the 1062 bp ORF and members of the other subgroups of family GH-16 (Table 3) was limited to a few conserved residues (Figure 9). The latter result agrees with the findings of Jam et al. (2005) for -agarase A and -agarase B of Zobellia galactanivorans. According to Allouch et al. (2003), the overall sequence identity between members of a GH-16 subfamily is typically better than 30-35% and the inter-subfamily sequence identity is low (10-25%) with the similarity restricted to a few invariant residues. This is in accordanceTown with our results.

Glycosyl bond hydrolysis in family GH-16 members occurs via an overall retention of the anomeric configuration of the protein. This takes placeCape via a two -step mechanism involving two catalytic residues, which in the case of family GH-16 agarases, are the two glutamic acid residues referred to as the catalytic machinery.of In the first step (glycosylation), one of the glutamic acid residues acts first as a general acid (proton donor), which protonates the glycosidic oxygen and thereby facilitates aglycone departure. Simultaneously, the other catalytic residue acts as a nucleophile, attacking the anomeric carbon and forming a glycosyl- enzyme intermediate. In the second step (deglycosylation), the general acid now acts as a general base wherebyUniversity it deprotonates a water molecule, in order to hydrolyze the glycosyl- enzyme intermediate and to release the product. Both steps occur via transition states (Davies et al., 1995; Alouch et al., 2003). The two glutamic acid residues which act as the catalytic residues are usually separated by three amino acids in bacterial lichenases and plant xyloglucan endotransglycosylases, or by four residues in the galactanases, laminarinases and agarases (Barbeyron et al., 1998). Site directed mutagenesis was used to demonstrate the catalytic function of Glu134 and Glu138 in the β-1,3-1,4-glucanase of Bacillus licheniformis (Juncosa et al., 1994) and of Glu103 and Glu107 in the endo-1,3-1,4-β-glucanase of Bacillus macerans (Hahn et al., 1995) while selective inhibition was used to identify Glu105 as a catalytic residue in the β-1,3-1,4-glucanase of Bacillus amyloliquefaciens (HØj et al., 1992). 64

All of the above mentioned enzymes are family GH-16 representatives. By performing multiple sequence alignments, glutamic acid residues have also been identified in several other family GH-16 members such as: Glu163 of the kappa-carrageenase from Alteromonas carrageenovora (Barbeyron et al., 1994); Glu160 and Glu165 of the kappa-carrageenase from Zobellia galactanivorans (Barbeyron et al., 1998); Glu147 and Glu152 of β-agarase A and Glu184 and Glu189 of β-agarase B from Zobellia galactanivorans (Allouch et al., 2003). Multiple sequence alignments (Figure 9) with all the family GH-16 proteins that share sequence identity with the 1062 bp ORF from Pseudoalteromonas sp. LS2i showed a strong conservation of glutamic acid residues in relation to Glu143 and Glu148 of the 1062 bp ORF. The latter two residues aligned with the catalytic residues identified in the kappa- carrageenase, β-agarase A and β-agarase B from Zobellia galactanivorans described above as well as to glutamic acid residues in all of the eleven other agarases in Figure 9. Except for Glu160 of the kappa-carrageenase from Zobellia galactanivorans no other conservation to Glu143 of the 1062 bp ORF was identified amongst the six non–agarolyticTown family GH-16 members while conserved glutamic acid residues that aligned with Glu148 from the Pseudoalteromonas sp. LS2i ORF were identified in two (Glu165 of the kappa-carrageenase from Zobellia galactanivorans and Glu170 of the arabinogalactanCape endo-1,4-β-galactosidase from Saccharophagus degradans) out of the six non-agarolytic family GH-16 members shown in Figure 9. In contrast to the highly conservedof catalytic residues in family GH-16 agarases, the residues involved in surface binding are however not conserved in the agarases whose sequences are available (Allouch et al., 2004).

Upon analysis of the sequence upstream of the identified 1062 bp ORF, the putative ribosome-binding siteUniversity was observed 6 bases upstream of the ATG initiation codon. A putative –35 promoter region as well as a putative –10 promoter region was identified at positions 490-495 and 513-518 respectively. The base, C, at position 528 was identified as a putative transcriptional start point. Primer extension analysis can be employed in the future to determine the exact locations of the promoter regions and the transcriptional start site.

As described in this chapter the construction and screening of a Pseudoalteromonas sp. LS2i gene bank resulted in the successful cloning of an agarase gene from this G. gracilis pathogen. Since the agarase is exported in E. coli, as can be seen from the pitting on agar around an E. coli JM109 transformant containing one of the constructs with the agarase gene, 65 we concluded that an extracellular agarase has been cloned from Pseudoalteromonas sp. LS2i. The nucleotide sequence, deduced amino acid sequence, putative regulatory regions as well as sequence identity of the cloned agarase to several other proteins from the database was also described in this chapter. Our results also suggest that the agarase cloned from Pseudoalteromonas sp. LS2i is a novel member of family GH-16 of the glycoside hydrolases. It is known from literature that agarolytic bacteria often contain more than one agarase (see section 2.1). Both Zong et al. (2001) and Sugano et al. (1994) also concluded that microorganisms appear to degrade agar by using a series of enzymes with narrow specificities rather than a single enzyme with broad specificity. It is thus very likely that Pseudoalteromonas sp. LS2i contains other agarases, which can be cloned in future. The overexpression of the cloned agarase in Escherichia coli JM109 will however simplify the purification (described in chapter 3) of the enzyme produced by this gene. The work done in this chapter has thus provided a basis for further analysis of the role this extracellular agarase, which has been cloned from Pseudoalteromonas sp. LS2i, mightTown play in eliciting disease symptoms in the red alga Gracilaria gracilis.

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University 66

3. CHAPTER 3

PURIFICATION AND CHARACTERIZATION OF THE AGARASE PROTEIN EXPRESSED FROM THE RECOMBINANT PLASMID pEB3 IN E. COLI

CONTENTS

3.1. Introduction ...... 68 3.2. Materials and Methods ...... 71 3.2.1. Determination of the optimum conditions for protein export from E. coli JM109 (pEB3) ...... 71 3.2.2. Purification of the agarase from E. coli JM109 (pEB3) ...... 71 3.2.2.1. Ammonium sulphate precipitation ...... 71 3.2.2.2. Ion-exchange chromatography...... 72 3.2.2.3. Gel filtration chromatography ...... 72 3.2.3. Bradford assay for protein quantitation ...... Town...... 73 3.2.4. Ferricyanide assay for quantitation of agarolytic activity...... 73 3.2.5. SDS-polyacrylamide gel electrophoresis ...... 73 3.2.6. Zymogram detection of the extracellularCape agarase ...... 73 3.2.7. Thin-Layer chromatographic analysis of the purified agarase ...... 73 3.3. Results ...... of ...... 75 3.3.1. Growth characteristics of E. coli JM109 (pEB3) ...... 75 3.3.2. Purification of the agarase from E. coli JM109 (pEB3) ...... 75 3.3.3. Characterization of the agarase from E. coli JM109 (pEB3) ...... 79 3.4. Discussion ...... 82 University

Summary A -agarase that was cloned from Pseudoalteromonas sp. LS2i into E. coli JM109 was isolated from the transformant E. coli JM109 (pEB3). The extracellular -agarase was purified using ammonium sulphate precipitation as well as anion-exchange and gel filtration chromatography. SDS-PAGE analysis revealed that the size of the purified protein is in the region of 39 kDa and zymogram analysis confirmed that it is agarolytic. Thin-layer chromatography (TLC) of the digestion products produced by the purified agarase showed that this enzyme hydrolyzed the -1,4 linkage of agarose to predominantly produce neoagarohexaose and neoagarotetraose. The enzyme was however unable to hydrolyze neoagarotetraose and neoagarobiose. TLC data also revealed that the extracellular agarase produced by Pseudoalteromonas sp. LS2i has the same mode of action as the purified -1,4 agarase, suggesting that the primary agarase of Pseudoalteromonas sp. LS2i was cloned into E. coli JM109 and purified from the extracellular media of the E. coli JM109 (pEB3) transformant. Town

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3.1. Introduction

If the epiphytic bacteria present on the surface of red alga utilize agarose as a carbon source, agarose should be degraded to its constitutive monosaccharides, D-galactose and 3,6- anhydro-L-galactose. Two types of agarolytic enzymes, - and -agarases, are involved in the degradation of agar. -agarases, which cleave the -1,4 linkage in agarose to produce a series of neoagarooligosaccharides have been purified from several bacterial strains belonging to the genera Alteromonas (Kirimura et al., 1999; Leon et al., 1992; Wang et al., 2005), Cytophaga (van der Meulen et al., 1975; Duckworth et al., 1969), Pseudomonas (Malmqvist, 1978; Ha et al., 1997; Yamaura et al., 1991; Kang et al., 2003), Pseudoalteromonas (Morrice et al., 1983d; Vera et al., 1988; Schroeder et al., 2003), Streptomyces (Bibb et al., 1987; Parro et al., 1997), Vibrio (Araki et al., 1989; Sugano et al., 1995; Sugano et al., 1993a; Aoki et al., 1990; Fukasawa et al., 1987), Bacillus (Suzuki et al., 2003), Zobellia (Allouch et al., 2003; Jam et al., 2005), Microbulbifer (Whitehead et al., 2001; OhtaTown et al., 2004a, 2004b and 2004c) and Agarivorans (Ohta et al., 2005). -agarases cleave the -1,3 linkage in agarose leading to the production of a series of agarooligosaccharides. In contrast to the many purified -agarases, only two -agarases have beenCape purified from Alteromonas agarlyticus (Potin et al., 1993; Young et al., 1978) and Thalassomonasof sp. JAMB-A33 (Otha et al., 2005a). However three -neoagarooligosaccharide hydrolases that are capable of cleaving the

-1,3 linkage in neoagarobiose to produce D-galactose and 3,6-anhydro-L-galactose have been purified from Vibrio sp. Strain JT0107 (Sugano et al., 1994), Pseudoalteromonas atlantica (Day and Yaphe; 1975) and Bacillus sp. MK03 (Suzuki et al., 2002).

The agarase system ofUniversity Pseudoalteromonas atlantica, consisting of at least three enzymes, was most extensively characterized and provides a model for the study of other marine agarase systems. In this system agar is hydrolyzed through cleavage of the -1,4 linkage by the extracellular -agarase I to produce a mixture of oligosaccharides with neoagarotetraose as the predominant product. Neoagarotetraose permeates into the cell while the larger agar oligomers cannot cross the cell wall barrier. Neoagarotetraose is then further hydrolyzed by the membrane-bound -agarase II, also known as neoagarotetraose hydrolase, to produce neoagarobiose (Morrice et al., 1983a, 1983b, 1983c and 1983d). Finally the neoagarobiose is 69 hydrolyzed by the intracellular neoagarobiose hydrolase to the metabolically useful carbon sources, D-galactose and 3,6-anhydro-L-galactose (Day and Yaphe, 1975).

Several other agarases have been characterized and a number of grouping systems were proposed for these enzymes. Schroeder (2001) divided all the characterized extracellular agarases into three groups based on the mode of action of the agarases. The first group included -agarases with the inability to hydrolyze neoagarotetraose and neoagarobiose, while the second group included all the -agarases that were not able to hydrolyze neoagarobiose. The third group was defined by the ability to hydrolyze the -1,3 linkage in agarose with the only member of that group being the -agarase isolated from Alteromonas agarlyticus (Potin et al., 1993; Young et al., 1978). Suzuki et al. (2002) described three types of -agarases, which were again defined by their mode of action on neoagarooligosaccharides. The first group was similar to group I described by Schroeder (2001) and was defined by the ability to hydrolyze neoagarohexaoseTown to produce neoagarotetraose and neoagarobiose, and the inability to hydrolyze the latter products. The second group included agarases with the ability to hydrolyze neoagarotetraose and neoagarohexaose to produce neoagarobiose, whichCape is similar to group II described by Schroeder (2001). The third group described ofby Suzuki et al. (2002) contained the extracellular agarase-c from Vibrio sp. PO-303 (Araki et al., 1998). This group was defined by the ability to hydrolyze agarose to produce neoagarooctaose and neoagarodecaose as the main products, and the inability to hydrolyze neoagarotetraose and neoagarohexaose. Vera et al. (1998) proposed a grouping system for -agarases according to size, where the first group contained agarases with sizes that ranged between 31 and 33 kDa. Agarases with molecular weights between 52 andUniversity 59 kDa were sorted into the second group, while the third group contained one -agarase with a molecular weight of 105 kDa. Vera et al. (1998) further explained that agar-clearing, softening and depressions around the agarolytic colony is characteristic of bacteria that synthesize agarases belonging to the first and second groups and that this effect is related to the production of low-molecular-weight agarases that can diffuse through the gel pores.

The aim of this chapter was to firstly purify the agarase from the recombinant E. coli JM109 (pEB3) to a level of purity where it could be used to raise polyclonal antibodies (Chapter 5). 70

Secondly the size of the purified agarase will be determined and compared to the theoretical size as determined from the sequence of the agarase gene cloned from Pseudoalteromonas sp. LS2i as described in Chapter 2. The third aim was to determine the mode of action of the agarase cloned from Pseudoalteromonas sp. LS2i into E. coli JM109. This will confirm whether the cloned agarase, which had significant homology to several -agarases (Chapter 2), is in fact a -agarase. Finally, in order to predict whether it was the primary agarase that was cloned from Pseudoalteromonas sp. LS2i into E. coli JM109 (pEB3) (Chapter 2), it was necessary to determine whether the agarase produced by E. coli JM109 (pEB3) had the same mode of action as the agarase produced extracellularly by Pseudoalteromonas sp. LS2i.

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3.2. Materials and Methods All media and solutions used in this study are listed in Appendix A.

3.2.1. Determination of the optimum conditions for protein export from E. coli JM109 (pEB3)

In order to obtain a sufficient yield of agarase protein for purification from the E. coli JM109 (pEB3) culture supernatant, the optimum conditions for protein export into the supernatant had to be determined. E. coli JM109 containing the pEB3 construct was thus inoculated from an O/N culture into LB (a 1% inoculum) (Appendix A.1.1) containing 100 ug/ml ampicillin (Appendix A.2.1) either with or without IPTG (120 ug/ml) (Appendix A.2.1). These cultures were grown O/N at both 22C and 30C on an orbital shaker (100 rpm), after which the supernatants were collected. The agarolytic activity of each cultureTown was determined as described in section 3.2.4.

3.2.2. Purification of the agarase from E. coli JM109 (pEB3) Cape In order to inhibit microbial growth sodium azideof (200 ug/ml) (Appendix A.2.2) was added to all buffers and solutions used in this section of the study.

3.2.2.1. Ammonium sulphate precipitation

Unless stated otherwise the following procedures were performed at 22C. Three, 1 l E. coli JM109 (pEB3) culturesUniversity were grown for 36 h in LB containing 100 ug/ml ampicillin and 120 ug/ml IPTG. The cells were removed by centrifugation at 8K rpm for 5 min. A total of 2.5 l of supernatant was collected. The collected supernatant was adjusted to a final ammonium sulphate saturation of 85% (559 g/l) by adding ammonium sulphate (Merck) in increments of 250 g (Englard and Seifter, 1990). The precipitate was collected by centrifugation (8K rpm for 15 min) and resuspended in 60 ml of 20 mM Tris-Cl buffer (pH7) (Appendix A.2.2). The collected precipitate was dialyzed twice for 2 h against 4 l of 20 mM Tris-Cl buffer (pH7) at 4C.

3.2.2.2. Ion-exchange chromatography

The following procedures were performed at 4C. DEAE-Sephadex A-50 (Pharmacia) was activated with 5 M NaCl (Appendix A.2.2) for 48 h before it was poured into a column (2.5 cm x 45 cm) to a final bed volume of 133 ml. The column was equilibrated with 500 ml 20 mM Tris-Cl buffer (pH7). The dialyzed ammonium sulphate concentrate was applied to the column and the column was washed with 300 ml 20 mM Tris-Cl buffer (pH7). In order to elute the proteins from the column a gradient mixer was used to prepare two gradient solutions. The two NaCl gradient solutions, i.e. 0 – 0.2 M (400 ml) (Appendix A.2.2) and 0.2 – 0.4 M (400 ml) (Appendix A.2.2) were passed through the column respectively. Five ml fractions were collected using a Gilson FC 204 Fraction collector (Gilson Medical Electronincs, Inc.). Every 5th fraction was assayed for agarolytic activity (section 3.2.4) and the absorbance at 280 nm was read for every 2nd fraction with a Beckman spectrophotometer. In order to exclude as much of the E. coli JM109 proteins presentTown in the active fractions as possible, only active fractions with an absorbance (280 nm) reading below 2.5 were pooled and the resultant total volume (180 ml) was reduced with an Amicon Centricon PM10 filter system to a final volume of 5 ml. Cape

3.2.2.3. Gel filtration chromatography of

The following procedures were also performed at 4C. A column (2.7 cm x 45 cm) of Sephadex G75 (Pharmacia) with a final bed volume of 160 ml was prepared. The column was equilibrated with 160 ml of 20 mM Tris-Cl (pH 7) before the active concentrate (5 ml) was applied to it. The proteinUniversity was eluted with 20 mM Tris-Cl (pH 7) buffer and 10 ml fractions were collected using a Gilson FC 204 Fraction collector (Gilson Medical Electronincs, Inc.). Every 3rd fraction was assayed for agarolytic activity (section 3.2.4) and the absorbance at 280 nm was read for every 2nd fraction with a Beckman spectrophotometer. In order to increase the purity as much as possible, the active fractions with an absorbance (280 nm) reading below 1.0 were pooled and the resultant total volume (90 ml) was reduced with an Amicon Centricon PM10 filter system to a final volume of 5 ml. Finally, the active concentrate was dialyzed against 10 mM phosphate buffer, pH 7 (Appendix A.2.2).

3.2.3. Bradford assay for protein quantitation

Protein concentrations were determined by the Bradford method as described in Appendix B.10.

3.2.4. Ferricyanide assay for quantitation of agarolytic activity

Agarase activity was determined by the Ferricyanide reducing sugar assay as described in Appendix B.11. The incubation period used to assay the activity of the chromatography fractions was reduced to 30 min. The agarolytic activity (U) of the chromatography fractions was expressed as ug galactose per ml per min at 37C.

3.2.5. SDS-polyacrylamide gel electrophoresis Town SDS-PAGE analysis was performed with a 12% gel according to the Laemmli method (Appendix B.12). Protein concentrations were determined by the Bradford method as described in 3.2.3. Cape

3.2.6. Zymogram detection of the extracellularof agarase

Zymogram detection was performed as described in Appendix B.13. A sample containing 6.5 ug purified protein, as determined with the Bradford method described in 3.2.3, was prepared and loaded onto the gel. University 3.2.7. Thin-Layer chromatographic analysis of the purified agarase

LS2i and JM109 (pEB3) were inoculated from 5 ml O/N cultures into 200 ml and 100 ml broths, respectively, and incubated for 24 h at 22C on an orbital shaker at 100 rpm. The cells were harvested (8K rpm for 10 min at 4C) and the culture supernatants collected. The extracellular proteins present in the supernatants were precipitated with 50% Trichloroacetic acid (Merck) (Appendix B.14) and the pellets were each resuspended in 1ml 10 mM phosphate buffer (pH 7). Protein concentrations were determined using the Bradford assay described in 3.2.3. 74

Extracellular proteins from Pseudoalteromonas sp. LS2i (50 ug) and E. coli JM109 (pEB3) (50 ug), collected as described in the previous paragraph, as well as purified agarase (700 ng) were each added to 100 ul 1% agarose substrate (Appendix A.2.4) that had been freshly prepared. Similarly 50 ug extracellular proteins from strain LS2i, E. coli JM109 (pEB3) and 700 ng purified agarase protein were each added to 50 ul of each of the two oligosaccharides, neoagarotetraose (0.5 ug/ul; final concentration) and neoagarobiose (0.5 ug/ul; final concentration). The reaction mixes were made up to a final volume of 200 ul with 20 mM PIPES solution (Appendix A.2.4), and incubated at 37C for 1 h. The reaction mixes were analyzed by TLC as described in Appendix B.15. The two oligosaccharides (0.75 ug of each), neoagarotetraose and neoagarobiose, were combined and used as molecular size markers.

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3.3. Results

3.3.1. Growth characteristics of E. coli JM109 (pEB3)

In order to obtain the optimum yield of extracellular agarase protein for purification from E. coli JM109 (pEB3), the bacterium was cultured at different conditions as described in 3.2.1. The agarolytic bacterium E. coli JM109 (pEB3) produced the highest amount of extracellular agarase when grown in LB containing 100 ug/ml ampicillin and 120 ug/ml IPTG at 22C (Figure 1).

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Agarolytic activity (U) activity Agarolytic of 1

0 22°C (IPTG) 22°C 30°C (IPTG) 30°C Culture conditions

Figure 1. The effect of different incubation temperatures and IPTG on the activity of the E. coli JM109 (pEB3) extracellular agarase. Agarolytic activity (U) is expressed as ug galactose/ml/min at 37UniversityC.

3.3.2. Purification of the agarase from E. coli JM109 (pEB3)

The supernatant collected from 3 l of E. coli JM109 (pEB3) culture medium was saturated to 85% with ammonium sulphate. The collected precipitate was resuspended in and dialyzed against 20 mM Tris-Cl (pH 7). The active precipitate was then subjected to anion-exchange chromatography using DEAE-Sephadex A-50. Using a 0.2 – 0.4 M NaCl gradient, proteins were eluted from the column in five ml fractions which were collected in 68 tubes. The 76 agarolytic activity eluted in fractions 25 to 68 while most of the other proteins present in the active concentrate eluted between fractions 11 and 33 (Figure 2). Therefore in order to increase the purity of the agarase, only fractions 33 to 68 were pooled and subsequently concentrated to a final volume of 5 ml. The amount of total protein recovered after this purification step was 27.5% (Table 1). Since agarase activity was still present in the last fraction (no 68), further elution with a 0.4 M NaCl solution probably would have resulted in the elution of more agarase protein and a subsequent higher yield of agarase after this purification step.

Gel filtration chromatography was performed by passing the active concentrate through a Sephadex G75 column. The proteins eluted in 10 ml fractions that were collected in 31 tubes. The active fractions eluted between tubes 15 and 30, while the bulk of the other proteins still present in the concentrate eluted in fractions 13 to 19 (Figure 3). In order to exclude as much of the E. coli JM109 (pEB3) proteins as possible, fractions 1 to 18Town and fractions 28 to 31 were discarded in an attempt to keep the agarase concentration in the active concentrate as high as possible. Only fractions 19 to 27 were pooled and further concentrated to a final volume of 5 ml before dialysis against 10 mM phosphateCape buffer, pH 7. The amount of total protein recovered after this purification step was 1.5% (Table 1). of An aliquot of the culture supernatant of E. coli JM109 (pEB3) and the final concentrate were analyzed by SDS-PAGE. Multiple bands of different sizes were present in the crude extract (Figure 4, lane 3), while the final active concentrate exhibited a protein band of 39 kDa in size (Figure 4, lane 2). University 77

12 3.5 10 3 8 2.5 2 6 1.5 4 1 2

Absorbance (280nm) Absorbance 0.5 0 0 Activity Agarolytic (U) 0 10 20 30 40 50 60 70 Fraction Number protein agarase activity

Figure 2. Ion-exhange chromatography of the ammonium sulphate concentrate of the E. coli JM109 (pEB3) culture supernatant on DEAE-Sephadex A-50 eluted with a 0.2 – 0.4 M NaCl gradient solution. The protein was monitored by absorbance at 280 nm and the agarolytic activity was assayed as described in section 3.2.4. Agarolytic activity (U) is expressed as ug galactose/ml/min at 37C. Town

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0.5 Agarolytic activity Agarolytic (U) Absorbance (280nm) Absorbance 0.5 0 0 0 5 10 15 20 25 30 35 UniversityFraction Number protein agarase activity

Figure 3. Gel filtration chromatography of the active concentrate (post DEAE-Sephadex A50 chromatography) from E. coli JM109 (pEB3) on Sephadex G75. The protein was monitored by absorbance at 280 nm and the agarolytic activity was assayed as described in section 3.2.4. Agarolytic activity (U) is expressed as ug galactose/ml/min at 37C.

Table 1. Purification of extracellular agarase from Pseudoalteromonas sp. LS2i Purification step Total protein (mg) Recovery of total protein (%) Culture supernatant 212.674 100 Ammonium sulphate preparation 135.686 63.80 Sephadex A50 58.542 27.52 Sephadex G75 3.258 1.54

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Figure 4. SDS-PAGE of samples obtained before and after purification of the extracellular agarase of E. coli JM109 (pEB3). Lane 1: molecular mass markers; lane 2: 0.5 ug final concentrate; lane 3: 15 ug culture supernatant. The proteins were visualized by staining with Coomassie blue dye. The sizes of the molecular mass markers are shown in kDa.

3.3.3. Characterization of the agarase from E. coli JM109 (pEB3)

A zymogram was performed to determine whether the single band visible in Figure 4, lane 2 (final concentrate) was agarolytic. Due to the lack of sensitivity of the latter assay thirteen times more of the final concentrate (Figure 5, lane 3) had to be loaded onto the gel in order to detect agarolytic activity. When the same amount of protein was subjected to SDS-PAGE it exhibited two bands, running close to each other on the gel, in the region of 39 kDa (Figure 5, lane 2). The zymogram confirmed that at least one of these two bands represented an agarolytic enzyme (Figure 5, lane 3). Since the two bands were so close to each other it was impossible to determine whether both bands or only one of the two bands were represented by the 39 kDa band exhibited on the zymogram.

TLC was used to firstly determine the substrate specificity of the purified extracellular agarase from E. coli JM109 (pEB3) and secondly to determine whetherTown the cloned agarase expressed in E. coli JM109 (pEB3) has the same mode of action as the extracellular agarase(s) expressed in Pseudoalteromonas sp. LS2i. TLC revealed that the extracellular agarase(s) expressed in Pseudoalteromonas sp. LS2iCape hydrolyze agarose to produce neoagarohexaose and neoagarotetraose (Figure 6, lane 4). The purified agarase, which is expressed extracellularly in E. coli JM109 (pEB3),of hydrolyzed agarose to predominantly yield neoagarohexaose and neoagarotetraose (Figure 6, lanes 3 and 5). The extracellular agarase present in the supernatant of E. coli JM109 (pEB3) did however also yielded a small amount of neoagarobiose when agarose was used as a substrate (Figure 6, lane 5). The agarase(s) present in the extracellular fraction of strain LS2i and E. coli JM109 (pEB3), as well as the purified agarase,University did not hydrolyze neoagarotetraose or neoagarobiose (Figure 6, lanes 6 – 11)

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Figure 5. SDS-PAGE and Zymogram detection ofCape the extracellular agarase of E. coli JM109 (pEB3) in the final concentrate. Lane 1: molecular mass markers; Lane 2: 6.5 ug final concentrate; Lane 3: Zymogram of final concentrateof (6.5 ug). The proteins were visualized by staining with Coomassie blue dye and the enzymatically active bands were detected by staining with Gran’s Iodine. The sizes of the molecular mass markers are shown in kDa

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Figure 6. TLC analysis of the oligosaccharides formed following degradation of the substrate by the purified agarase, the culture supernatant from E. coli JM109 (pEB3) and the culture supernatant from Pseudoalteromonas sp. LS2i. Lane 1: standard markers; Lane 2: agarose incubated with buffer; Lane 3: agarose incubated with purified agarase; Lane 4: agarose incubated with supernatantUniversity from strain LS2i; Lane 5: agarose incubated with supernatant from E. coli JM109 (pEB3); Lane 6: tetraose incubated with purified agarase; Lane 7: tetraose incubated with supernatant from strain LS2i; Lane 8: tetraose incubated with supernatant from E. coli JM109 (pEB3); Lane 9: biose incubated with purified agarase; Lane 10: biose incubated with supernatant from strain LS2i; Lane 11: biose incubated with supernatant from E. coli JM109 (pEB3); Lane 12: standard markers.

3.4. Discussion

The extracellular agarase was purified from E. coli JM109 (pEB3) by performing the following procedures: ammonium sulphate precipitation, anion-exchange chromatography through DEAE-Sephadex A-50 and gel filtration chromatography through Sephadex G75. SDS-PAGE revealed that the size of the purified protein is approximately 39 kDa, which is consistent with the theoretical size of 38.9 kDa predicted from the nucleotide sequence of the mature agarase (Chapter 2, section 2.2.4).

Zymogram analysis confirmed that the 39 kDa protein purified from E. coli JM109 (pEB3) was agarolytic. There are a number of possible reasons for the two similarly-sized bands that were observed when an increased amount of the final concentrate was analyzed by SDS- PAGE (Figure 5, lane 2). According to Englard and Seifter (1990) a number of proteins with unusual compositions, such as various lipoproteins and glycoproteins,Town can migrate “anomalously” during SDS-PAGE and consequently form multiple bands. Malmqvist (1978) found that two different agarases, agarase I and II, purified from a Pseudomonas-like bacterium are glycoproteins. Malmqvist (1978) alsoCape observed a broad band, as compared to other proteins, when agarase I from this Pseudomonasof -like bacterium was analyzed by SDS- PAGE. This indicated microheterogeneity, which is a phenomenon that occurs when a particular glycoprotein is present in different forms due to a difference in structure of one or more of its carbohydrate units (Malmqvist, 1978; McNaught and Wilkinson, 1997). Thus one explanation for the two bands observed in Figure 5 might be that the agarase purified from E. coli JM109 (pEB3) is a glycoprotein that is present in two different forms. Another possible explanation Universityfor the two bands might be that the agarase purified from E. coli JM109 (pEB3) is a heterodimer, consisting of two non-identical subunits. To our knowledge no agarase heterodimers have been reported so far but at least four agarases and one -neoagarooligosaccharide hydrolase have been isolated where the native molecular mass, as determined by gel filtration chromatography, was double that (Potin et al., 1993; Sugano et al., 1994), slightly less than double that (Malmqvist, 1978) or slightly more than double that (Jam et al., 2005; Kirimura et al., 1999) of the molecular mass determined from the band observed upon SDS-PAGE analysis. This suggests that the latter enzymes are homodimers, each consisting of two identical subunits. Another agarase purified from Bacillus sp. MK03 83 with a native molecular mass of 320 kDa, was discovered to be an octamer consisting of eight identical subunits (Suzuki et al., 2002). A third possible reason for the occurrence of the two bands might be that the protein was only partially purified and that one of the bands is not an agarase. This is however unlikely because it would imply that both proteins were not only similar in size but also had the same ion-exchange affinity.

The reason for the one band observed in Figure 4, lane 2 as opposed to the two bands observed in Figure 5, lane 2, is due to a difference in concentration of the protein loaded. Thirteen times (6.5 ug) more of the final concentrate was loaded in lane 2 of the gel shown in Figure 5 than what was loaded (0.5 ug) in lane 2 of Figure 4. This also suggests that the proteins represented by the two bands in Figure 5, lane 2 are not present in equal amounts.

TLC revealed that the agarase purified from E. coli JM109 hydrolyzed agarose to predominantly produce neoagarohexaose and neoagarotetraose (FigureTown 6, lane 3), and that it was unable to hydrolyze neoagarotetraose or neoagarobiose (Figure 6, lanes 6 and 9). According to this data the purified agarase can be classified as a -1,4 agarase and it falls into the first group of agarases according to the groupingCape systems described by Schroeder (2002) and Vera et al. (1998) which is discussed in section 3.1. In addition to the neoagarohexaose and neoagarotetraose, a small amount of neoagarobioseof was also produced by the agarase present in the supernatant of E. coli JM109 (pEB3) when agarose was used as the substrate (Figure 6, lane 5). The exact concentration of the agarase present in the extracellular fraction of E. coli JM109 (pEB3) and Pseudoalteromonas sp. LS2i are however unknown, but it seems like more agarase was present in the supernatant of E. coli JM109 (pEB3) than in the final concentrate (FigureUniversity 6, lane 3) and in the supernatant of Pseudoalteromonas sp. LS2i (Figure 6, lane 4), since only neoagarohexaose and neoagarotetraose and not neoagarobiose was produced from agarose by the latter two enzyme samples. A longer reaction time or increased amount of the purified agarase or the proteins present in the supernatant of Pseudoalteromonas sp. LS2i probably would have also resulted in the production of neoagarobiose from agarose. TLC data thus further suggested that the extracellular agarase(s) expressed in Pseudoalteromonas sp. LS2i had the same mode of action as the agarase purified from E. coli JM109 (pEB3). This result strongly suggests that the only extracellular or primary agarase produced by Pseudoalteromonas sp. LS2i was cloned into E. coli (JM109) (Chapter 2). 84

Most reported agarases are endotype enzymes that hydrolyze agar and agarose to produce hydrolysis products in the following order: neoagarotetraose = -hexaose  -biose, and they do not cleave neoagarotetraose (Araki et al., 1998). Since the agarase purified from E. coli JM109 (pEB3) also showed these characteristics, it is likely to also be an endotype enzyme. The mode of action of the latter enzyme is also consistent with that of the extracellular -agarase I from Pseudoalteromonas atlantica (Morrice et al., 1983b). Based on the model agarase system of Pseudoalteromonas atlantica as described in section 3.1, it is expected that Pseudoalteromonas sp. LS2i has at least one additional agarase, which would probably be similar to the -agarase II of Pseudoalteromonas atlantica. Other characterized β-agarases that also predominantly produce neoagarohexaose and neoagarotetraose as end products are pjaA from Pseudomonas sp. W7 (Ha et al., 1997), agarase-a from Vibrio sp. PO-303 (Araki et al., 1998), AgaA from Microbulbifer strain JAMB-A94 (Otha et al., 2004) and AgaA from Zobellia galactanivorans Dsij (Jam et al., 2005) as well as agarasesTown from Pseudoalteromonas Antarctica (Vera et al., 1998), Alteromonas sp. strain C-1 (Leon et al., 1992), Alteromonas sp. SY37-12 (Wang et al., 2005), Bacillus sp. MK03 (Suzuki et al., 2003) and Microbulbifer strain JAMB-A7 (Suzuki et al., 2003). To our knowledgeCape none of the latter enzymes were reported to not be secreted, but several were ofhowever reported to be expressed extracellularly. The sizes of the -agarases reported to have the same mode of action as that of the agarase from Pseudoalteromonas sp. LS2i, ranged between 32 kDa for the dagA from Pseudoalteromonas atlantica and 92 kDa for the -agarase expressed by Bacillus sp. MK03 (Morrice et al., 1983b; Suzuki et al., 2003). Of these enzymes, the extracellular -agarase expressed by Alteromonas sp. SY37-12 was most similar in size (39.5 kDa) to the agarase expressed from E. coliUniversity JM109 (pEB3) which is 39 kDa (Wang et al., 2005). Literature is only available on some of the agarases present in the database that exhibited sequence similarity to the agarase cloned from Pseudoalteromonas sp. LS2i (Table 2, Chapter 2). Of these agarases, the only one reported to not be secreted, and to have a different mode of action to the extracellular agarase from strain LS2i, is AgaB from Zobellia galactanivorans Dsij which was found to have 40% identity to the agarase from Pseudoalteromonas sp. LS2i (Jam et al., 2005).

We have succeeded in addressing the main aim of this chapter, which was to purify the primary agarase of Pseudoalteromonas sp. LS2i so that it could be used to raise polyclonal antibodies (Chapter 5). Future work on this protein might however entail the extraction of each of the two bands observed in Figure 5, lane 2, from a polyacrylamide gel after SDS- PAGE analysis of the final concentrate, and subsequent determination of the N-terminal amino acid sequence of each of the proteins. A comparison of the N-terminal sequences of these proteins to the DNA sequence of the agarase cloned into E. coli JM109 (pEB3) (Chapter 2) will reveal whether both bands represent this particular agarase or whether one of the bands is due to impurities from E. coli JM109 (pEB3).

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4. CHAPTER 4

ISOLATION AND CHARACTERIZATION OF PSEUDOALTEROMONAS sp. LS2i MUTANTS WITH REDUCED AGAROLYTIC ACTIVITY

CONTENTS

4.1. Introduction ...... 88 4.2. Materials and Methods ...... 92 4.2.1. Determination of the effect of nitrosoguanidine on the viability of Pseudoalteromonas sp. LS2i ...... 92 4.2.2. Identification of Pseudoalteromonas sp. LS2i mutants with reduced agarolytic activity...... 92 4.2.3. Characterization of sixteen Pseudoalteromonas sp. LS2iTown mutants ...... 93 4.2.3.1. Monitoring of agarase production and pigmentation on agar ...... 93 4.2.3.2. Agarolytic activity and cell density in 24 h liquid cultures ...... 93 4.2.4. Further characterization of mutants PseudoalteromonasCape sp. R9 and Pseudoalteromonas sp. R16 ...... of ...... 94 4.2.4.1. Quantitation of growth and agarase activity over a 24 h period ...... 94 4.2.4.2. Zymogram detection of agarolytic activity ...... 94 4.2.4.3. Thin-layer chromatography ...... 94 4.3. Results ...... 96 4.3.1. Mutagenesis of Pseudoalteromonas sp. LS2i ...... 96 4.3.2. CharacterizationUniversity of mutants R1-R16 ...... 97 4.3.3. Characterization of the agarolytic phenotype of strains LS2i, R9 and R16...... 99 4.4. Discussion ...... 103

Summary

In order to study the role that the extracellular agarase produced by Pseudoalteromonas sp. LS2i plays in the infection process of the seaweed G. gracilis, the aim of this chapter was to isolate one or more LS2i mutants with a defective extracellular agarase gene. Sixteen Pseudoalteromonas sp. LS2i mutants with reduced extracellular agarolytic activity were isolated after chemical mutagenesis. Six of the mutants had similar growth rates to strain LS2i, and of these, two mutants designated R9 and R16 were selected for further study. No agarolytic activity was detected for either R9 or R16 over a 24 h growth period, while strain LS2i exhibited an extracellular agarase activity after 2 h of incubation. Zymogram analysis revealed that mutant R9 produced a significantly less active extracellular agarase of the same size to that produced by LS2i. Thin-layer chromatography (TLC) also showed that while significantly less active than the enzyme produced by LS2i, the extracellular agarase from mutant R9 produced the same breakdown products to those generatedTown by the wild type strain, i.e. neoagarohexaose and neoagarotetraose. However, no active extracellular agarase was detected in mutant R16 using either zymogram detection or TLC analysis. Cape

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4.1. Introduction

To our knowledge, mutant strains with a deficiency or reduction in agarase production or agarose utilization have been reported only for Pseudoalteromonas atlantica, Streptomyces coelicolor and Saccharophagus degradans 2-40 (Belas et al., 1987; Hodgson and Chater, 1981; Ekborg et al., 2006). The construction of the Pseudoalteromonas atlantica mutant was accomplished through the inactivation of its agarase gene (agrA) by inserting a selectable drug (kanamycin) resistance gene into the coding sequence of the cloned agrA gene and the subsequent introduction of the agrA::kan plasmid into P. atlantica in which the wild-type locus was displaced by homologous recombination. Construction of this mutant enabled Belas and his co-workers to test the possibility that the extracellular agarase conferred a survival advantage to P. atlantica by providing the bacterium with the ability to break down and utilize agar as a sole carbon source. They found that while the P. atlantica agrA mutant had lost its primary agarase activity, minor agarolytic activity wasTown still present and although the wild type strain grew better, the mutant strain was still able to grow on a medium where agar was the sole carbon source (Belas et al., 1987). Cape Wild type Streptomyces coelicolor A3(2) is alsoof one of the few bacterial species that has the ability to utilize agar as a sole carbon source. Several mutants deficient in their ability to utilize agar were previously identified in Streptomyces coelicolor A3(2). These included mutations in the aga locus, which prevented the strain from growing on agar as sole carbon source but did not affect the extracellular agarolytic activity. The aga gene product appears therefore to be responsible for further processing or scavenging of agar degradation products (Hopwood et al., 1973a;University Hopwood and Wright, 1973b; Hodgson and Chater, 1981; Bibb et al., 1987). Two independent dag mutations that mapped some distance from the aga gene resulted in the inactivation of extracellular agarase production in S. coelicolor (Hodgson and Chater, 1981; Bibb et al., 1987). One of these mutations had resulted from the insertion of plasmid SCP1 into the dag region of the chromosome and the subsequent inactivation of this gene (Hodgson and Chater, 1981). The plasmid SCP1 that codes for methylenomycin A exists in Streptomyces coelicolor A3(2) and has the ability to integrate into the host chromosome (Kendall and Cullum, 1986). The other dag mutation may have resulted from a spontaneous or UV- induced mutation of the dag gene. Since neither of the two dag – strains could be cross-fed by a dag + strain, it was concluded that the dag – strains must be defective 89 in some functions additional to the agarase, since the products of agarase action should have been available to any cell in the vicinity. This implies that another gene or genes essential for growth on agar was affected by the mutagenesis (Hodgson and Chater, 1981).

Saccharophagus degradans 2-40 is an agarolytic marine bacterium containing five agarases. In an attempt to characterize the agarolytic system of this bacterium, Ekborg and co-workers (2006) have constructed gene replacement mutants for both the aga50A and the aga86E genes of Saccharophagus degradans 2-40. Since the genome sequence for this bacterium is known, mutagenic constructs were assembled by fusing the 1-kb segments flanking each side of the gene of interest to an antibiotic resistance cassette (kanamycin) by using splicing PCR. Each of the resulting linear mutagenic constructs was then introduced into Saccharophagus degradans 2-40, before potential transformants were selected on media containing kanamycin. Agarase activity was reduced but not abolished in any of the two mutants and both mutants have lost the ability to utilize agar as sole carbon source.Town

In order for Allouch and co-workers to study the crystal structure of the β-agaA protein from Zobellia galactanivorans in complex with its substrateCape of agaro-oligosaccharides, an inactive form of this agarase had to be used (Allouch et al., 2004). β-agaA is one of two β-agarases from Z. galactanivorans that were cloned intoof an E. coli strain and subsequently purified from it (Jam et al., 2005; Allouch et al., 2003). The mutation in the agaA gene was introduced by site directed replacement of the catalytic nucleophile (Glu147) by a serine on the recombinant plasmid which contains the cloned agaA gene. The resultant plasmid was then transformed into E. coli and the non-agarolytic mutant agarase was expressed in and purified from E. coli in the sameUniversity way as the native agarase (Allouch et al., 2004). The agaA mutation was however not introduced into wild type Zobellia galactanivorans.

Chemical as well as transposon mutagenesis has in the past successfully been used on other gram-negative bacteria to generate mutants defective in the excretion of certain extracellular proteins; i.e. Aeromonas hydrophila (Howard et al., 1983); Aeromonas salmonicida (Sakai, 1985); Escherichia coli (Oliver et al., 1981); Erwinia chrysanthemi (Andro et al., 1984); Erwinia carotovora (Reeves et al., 1993; Laasik et al., 2005); Pseudomonas aeruginosa (Wretlind et al., 1984); Ralstonia solanacearum (Huang et al., 1997; Tans-Kersten et al., 1998; Liu et al., 2005); Serratia marcescens (Hines et al., 1988); Xanthomonas campestris 90

(Dow et al., 1989). The mutations in some of these mutants often caused pleiotropic effects with decreased activities of several or all of the extracellular enzymes (Wretlind et al., 1984; Andro et al., 1984; Oliver et al., 1981; Howard et al., 1983; Reeves et al., 1993; Laasik et al., 2005; Dow et al., 1989). The synthesis of the extracellular proteins in these pleiotropic mutants was usually not impaired but they were accumulated in active forms inside the mutant cells. Certainly not all exoprotein deficient mutants are pleiotropic; i.e. using transposon Tn5, bacteriophage Mu or NTG mutagenesis, at least 23 mutants were isolated from Serratia marcescens where the mutation in each strain affected only a single extracellular protein (Hines et al., 1988). Mutations where only a single exoprotein is affected are thus likely to be located in the structural gene of that specific protein, while the mutation(s) in pleiotropic mutants seem to be located in global regulators of extracellular gene expression or in expression gene(s) monitoring one or more steps of the secretion process of different extracellular proteins. Some of the global regulators (i.e. GacA and KdgR) and genes involved in the secretion process (i.e. RsmA-RsmBTown system) have been identified and characterized in the plant pathogen Erwinia carotovora (Hyytiäinen et al., 2001; Koiv et al., 2001). Cape The evaluation of a single enzyme in pathogenicity is usually difficult because of the retention of residual enzyme activity in the mutantsof or because of the possible existence of other pathogenicity factors that can compensate for the lack of a single enzyme (Isshiki et al., 2001). However, in order to investigate the role that the primary or extracellular agarase produced by Pseudoalteromonas sp. LS2i plays in the virulence of this bacterium on the seaweed Gracilaria gracilis, the aim of this chapter was to construct a LS2i mutant with an inactive extracellularUniversity agarase. Different fragments of the agarase gene from E. coli JM109 (pEB3) were cloned into the vector pGp704. The resulting four constructs were then each transformed into E. coli (SM10) before conjugation was unsuccessfully attempted with Pseudoalteromonas sp. LS2i. Attempts to electroporate the constructs into Pseudoalteromonas sp. LS2i were also unsuccessful and we therefore decided to use chemical mutagenesis. We chose to use the chemical NTG as mutagen since it was readily available.This chapter describes the isolation and basic characterization of sixteen Pseudoalteromonas sp. LS2i mutants with reduced extracellular agarolytic activity, after chemical mutagenesis was applied. The further characterization of two of these agarase mutants with similar growth characteristics to wild type LS2i is also described here. This 91 includes the determination of the level of extracellular agarolytic activity compared to the wild type as well as the mode of action or the absence thereof of these two mutants with a reduced extracellular agarolytic activity.

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4.2. Materials and Methods All media and solutions used in this study are listed in Appendix A.

4.2.1. Determination of the effect of nitrosoguanidine on the viability of Pseudoalteromonas sp. LS2i

An O/N culture of Pseudoalteromonas sp. LS2i grown in VNSS medium (Appendix A.1.5) was inoculated (a 2% inoculum) into five 500 ml flasks, each containing 100 ml VNSS medium. (Appendix A.1.5). The cultures were grown at 22C on an orbital shaker (100 rpm) for 5 h. The cells were harvested by centrifugation (5K rpm for 5 min). Collected cells were resuspended in a nine salts solution (NNS) (Appendix A.1.5), washed twice with NNS, whereafter each pellet was resuspended in 33 ml VNSS medium (Appendix A.1.5). Nitrosoguanidine (NTG), resuspended in 20mM Tris-maleate buffer (pH9) (Appendix A.2.16), was added to four of the cultures to obtain a final concentrationTown of 50 ug NTG/ml. An equal volume of 20mM Tris-maleate buffer (pH9) (Appendix A.2.16) was also added to the control flask (no exposure to NTG). The experimental cultures were mutagenized for 10, 20, 30 and 40 min respectively at 30C and the controlCape culture was incubated for 40 min at 30C. After the incubation each culture wereof washed twice in 33 ml NNS (Appendix A.2.16) and resuspended in 100 ml VNSS (Appendix A.1.5). Serial dilutions of each culture were plated onto VNSS agar (Appendix A.1.5). The plates were incubated for 2 days at 22C, whereafter the number of colonies growing on each agar plate were counted and recorded in order to determine the survival rate of Pseudoalteromonas sp. LS2i at each time point. The method used for NTG-mutagenesis, as described above, was adapted from the experimental procedure used by LeeUniversity et al. (2000) for mutating Pseudoalteromonas sp. strain A28 in order to select for mutants lacking algicidal activity.

4.2.2. Identification of Pseudoalteromonas sp. LS2i mutants with reduced agarolytic activity

The mutagenized cultures that were washed and resuspended in 100 ml VNSS, as explained in section 4.2.1, were incubated O/N with shaking (100 rpm) at 22C. After incubation the cells were harvested by centrifugation (5K rpm for 5 min). Each pellet was resuspended in 30 93 ml VNSS (Appendix A.1.5) and 15 ml of glycerol (50%) (Appendix A.2.1) was added. The glycerol stock cultures were stored at -70C. The glycerol stock cultures that were mutagenized for 10, 20 and 30 min respectively were then defrosted and plated onto both marine agar (Appendix A.1.4) and VNSS agar (Appendix A.1.6). LS2i mutants which failed to produce indentations on the agar or which produced significantly smaller indentations on the agar compared to wild type Pseudoalteromonas sp. LS2i were identified after either a 24 h or 48 h incubation period at 22˚C. Approximately 15480 colonies were screened and sixteen mutants with reduced agarolytic activity were identified and designated R1-R16.

4.2.3. Characterization of sixteen Pseudoalteromonas sp. LS2i mutants

4.2.3.1. Monitoring of agarase production and pigmentation on agar

Mutants R1 – R16 as well as wild type Pseudoalteromonas sp. LS2iTown were streaked out onto VNSS agar (Appendix A.1.6) and incubated at 22C for three days. The amount of pitting on agar around the colonies as well as the colour of the colonies and the colour of the area around the colonies were compared to the wild typeCape on a daily basis.

4.2.3.2. Agarolytic activity and cell densityof in 24 h liquid cultures

The wild type as well as each one of the sixteen mutants was inoculated from O/N broth cultures into flasks containing 20 ml volumes of BMB (Appendix A.1.7) to obtain starting

OD600 values of 0.01. These strains were cultured for 24 h at 22C with agitation at 100 rpm. The growth of the strainsUniversity was determined after 24 h by measuring the OD600 (0.5 ml samples) in duplicate.

After measuring the cell density (OD600), 500 ul aliquots of the culture supernatants were collected (14000 rpm for 5 min) and used to quantitate agarase production. Agarase activity was determined in duplicate by the ferricyanide reducing sugar assay as described in Appendix B.11. The incubation period at 37 C, used to assay the activity of the samples, was reduced to 30 min. The agarolytic activity (U) of the samples was expressed as ug galactose per ml per min at 37C.

4.2.4. Further characterization of mutants Pseudoalteromonas sp. R9 and Pseudoalteromonas sp. R16

4.2.4.1. Quantitation of growth and agarase activity over a 24 h period

Strains LS2i, R9 and R16 were inoculated (in duplicate) from O/N cultures into flasks containing 100 ml BMB (Appendix A.1.7) to obtain starting OD600 values of 0.01. The cultures were grown at 22C on an orbital shaker (100 rpm) for 24 h. In order to determine the culture OD600 and the agarase activity of the strains, the cultures were sampled in duplicate from each flask every 2 h (1 ml volumes) over the 24 h growth period. After measuring the absorbance at each time interval, the rest of the sampled culture medium (0.5 ml) was centrifuged to collect the culture supernatant which was used to quantitate agarase activity. Agarase activity was determined in duplicate by the ferrecyanide reducing sugar assay as described in Appendix B.11. The incubation period at 37Town C, used to assay the activity of the samples, was reduced to 30 min. The agarolytic activity (U) of the samples was expressed as ug galactose per ml per min at 37C. Cape 4.2.4.2. Zymogram detection of agarolytic activity of Zymogram detection was performed in order to determine the presence or absence of an active extracellular agarase in LS2i, R16 and R9. In order to obtain extracellular proteins, the supernatants of 100 ml (BMB) (Appendix A.1.7) cultures of LS2i, R9 and R16 were collected following incubation for 24 h at 22C. The proteins present in the supernatants were concentrated (5-10 mlUniversity final volume) using an Amicon filter system. The zymogram procedure was performed as described in Appendix B.13. Each sample that was loaded onto the gel contained 0.5 ug extracellular protein. Protein concentrations were determined by the Bradford method described in Appendix B.10.

4.2.4.3. Thin-layer chromatography

LS2i, R16 and R9 (200 ml BMB) (Appendix A.1.7) were cultured at 22C with agitation at 100 rpm for 24 h. The cells were harvested (8K rpm for 10 min at 4C) and the culture 95 supernatants collected. The extracellular proteins present in the supernatants were precipitated with 50% Trichloroacetic acid (Merck) (Appendix B.14) and the pellets were each resuspended in 1 ml 10 mM phosphate buffer (pH 7) (Appendix A.2.2). Protein concentrations were determined using the Bradford assay as described in Appendix B.10.

Less than 100 ul protein from each of the three strains (50 ug/strain) was added to 100 ul amounts of a 1% agarose substrate (Appendix A.2.4) that had been freshly prepared. Similarly 50 ug amounts of protein from LS2i, R9 and R16 were added to 50 ul of neoagarotetraose (0.5 ug/ul; final concentration) as well as to 50 ul of neoagarobiose (0.5 ug/ul; final concentration). The reaction mixes were made up to a final volume of 200 ul with 20 mM PIPES solution (Appendix A.2.4) and incubated at 37C for 1 h. TLC was then performed as described in Appendix B.15. The two oligosaccharides (0.75 ug of each), neoagarotetraose and neoagarobiose, were combined and used as molecular size markers. Town

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4.3. Results

4.3.1. Mutagenesis of Pseudoalteromonas sp. LS2i

In order to mutagenise strain LS2i without killing all the viable cells in the culture, the optimum time period for exposing strain LS2i to NTG had to be determined. Therefore strain LS2i (in stationary phase) was exposed to NTG (50 ug/ml) at 30C for 0, 10, 20, 30 and 40 min respectively. Since the combination of rather high NTG concentrations and an alkaline pH causes the rapid breakdown of NTG to the effective mutagen diazomethane (Delic et al., 1970), Tris-maleate buffer adjusted to pH 9 and NTG at a final concentration of 50 ug/ml were used. After the treatment, equal volumes from each culture were plated and incubated on VNSS agar. The number of colonies on each agar plate represents the survival of strain LS2i at that specific time point and is shown in Table 1. Town

As can be seen from the data represented in Table 1, NTG definitely has an effect on the viability of strain LS2i during the first 40 min of exposure. There were however still viable colonies after 40 min of exposure. LS2i mutants mightCape therefore be present in any of the cultures that has been exposed to NTG for lessof than 40 min. LS2i cultures exposed to NTG for 10, 20, 30 and 40 min respectively were thus amplified, stored and screened for agarase mutants as explained in 4.2.2. A total of approximately 15 480 colonies were screened and sixteen agarase mutants, which had a reduced ability or an inability to pit the agar, were identified. University Table 1. LS2i survival following exposure to NTG Time (min) Number of colonies (cfu / 0.1 ml) % Survival 0 2.88 x 106 100 10 998 0.035 20 390 0.014 30 180 0.006 40 61 0.002

4.3.2. Characterization of mutants R1-R16

Agarolytic activity and growth in BMB after a 24 h incubation period was measured for LS2i and the sixteen mutants (Figure 1). Strain R10 (0.33 U) still produced only 33% of the agarolytic activity measured in LS2i (0.98 U) and is the only mutant for which agarolytic activity was detected after 24 h in liquid broth. Since mutants R4 and R8 did not grow in the basal marine broth (BMB) used in this experiment, the agarolytic activity of these strains could not be measured. Growth after 24 h of strains R7 (OD600 = 2.3), R9 (OD600 = 2.6), R10

(OD600 = 2.2), R11 (OD600 = 2.3), R13 (OD600 = 2.7) and R16 (OD600 = 2.5) was unaffected by the mutations, whereas growth of strains R1 (OD600 = 1.8), R12 (OD600 = 1.5) and R14

(OD600 = 1.7) was somewhat lower compared to wild type LS2i (OD600 = 2.3). Strains R2

(OD600 = 0.7), R3 (OD600 = 0.7), R5 (OD600 = 1.0) and R15 (OD600 = 0.8) were obviously struggling to grow since their OD600 values were less than 50 % of that of strain LS2i, while strain R6 (OD600 = 0.2) had the largest reduction in growth (11.5Town fold) over a 24 h period compared to the wild type strain.

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0.9 2.5 0.8

0.7 2 0.6

1.5 0.5

University 0.4 1

Absorbance(600nm) 0.3 Agarolytic activity (U) activity Agarolytic

0.2 0.5 0.1

0 0 LS2i R 1 R 2 R 3 R 4 R 5 R 6 R 7 R 8 R 9 R 10 R 11 R 12 R 13 R 14 R 15 R 16

Strain OD600 Agarolytic activity (U) Figure 1. Growth and agarolytic activity of strains LS2i and R1 - R16 after a 24 h incubation period in BMB at 22C. The cell density was monitored by absorbance at 600 nm and the agarolytic activity was assayed as described in section 4.2.3.4. Agarolytic activity (U) is expressed as ug galactose/ml/min at 37C.

The amount of pitting and pigmentation of LS2i and mutants R1-R16 were also monitored on agar plates (Table 2). Strain LS2i does not produce a pigment under normal growth conditions but the mutants R2 and R9 produced brown pigments (also produced pigments when grown in liquid broth) after 2 days on agar at 22C. All the mutants, except for R7 and R16, pitted the agar to some extent after 3 days of incubation. Mutants R2, R3 and R9 did however only start to pit the agar on the third day, while strains R5, R6, R12, R13, R14 and R15 began pitting the agar on the second day of incubation. Only strains R1, R4, R8, R10 and R11 started pitting the agar, to a lesser extent than LS2i, after 1 day of incubation.

Table 2. Characteristics of mutants R1-R16 in comparison to strain LS2i

1 day at 22C 2 days at 22C 3 days at 22TownC pitting pigmented pitting pigmented pitting pigmented R1 L − L − L − R2 − − − + L + R3 − − − − CapeL − R4 L − S − of S − R5 − − L − L − R6 − − L − L − R7 − − − − ¯ − R8 L − L − L − R9 − − − + L + R10 L −University S − S − R11 L − L − L − R12 − − L − L − R13 − − L − L − R14 − − S − S − R15 − − L − L − R16 − − − − − − _ S: same as LS2i, L: less than LS2i, +: pigmented, : not detected

4.3.3. Characterization of the agarolytic phenotype of strains LS2i, R9 and R16

Two of the mutant strains, R9 and R16 (Figure 2), that exhibited a significant reduction in extracellular agarolytic activity and did not seem to be growth impaired, as determined from the preliminary experiments described in 4.3.2 were characterized more extensively. The growth characteristics (Figure 3) and agarolytic activity (Figure 4) of strain LS2i and the mutant strains R9 and R16 were examined in BMB over a 24 h growth period. The growth characteristics of mutants R9 and R16 are unaffected by their specific mutations and were similar to the growth rate of LS2i (Figure 3). The mutations in R9 and R16 did however affect agarase activity significantly. No agarolytic activity was measured during the 24 h incubation period for either R9 or R16, while low levels of agarase activity for LS2i were already detectable after 2 h of incubation. Agarase activity for LS2i increased significantly after 8 h and reached a maximum enzyme activity value of 0.75 TownU after 24 h (Figure 4).

A B Cape C of

Figure 2. A: LS2i, B: R9 and C: R16 cultured on a VNSS agar plate. The arrows indicate the zone of agar hydrolysisUniversity around LS2i

100

0.1 Absorbance (600nm) Absorbance

0.01 0 4 8 12 16 20 24

Time (h) LS2i R9 R16

Figure 3. Growth of LS2i, R9 and R16 cultured in BMB at 22C. The optical density at 600 nm represents the mean of 4 values, while the error bars represent the standard error from the mean.

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0.2 Agarolytic activity (U) activity Agarolytic

0 0 5 10 15 20 25 Time (h) LS2i R9 R16

Figure 4. Agarolytic activity of LS2i, R9 and R16 cultured in BMB at 22C. Agarase activity is the mean of 4 values,University while the error bars represent the standard error from the mean. Agarolytic activity (U) is expressed as ug galactose/ml/min at 37C.

In order to confirm the presence or absence of an active extracellular agarase in LS2i, R9 and R16, the supernatants of 24 h cultures of the latter strains were concentrated and subjected to zymogram analysis (Figure 5). The zymogram exhibited a clear 30 kDa band for LS2i (Figure 5, lane 2), a very faint 30 kDa band for R9 (Figure 5, lane 3), while no bands were detected for R16 (Figure 5, lane 4). The zymogram therefore demonstrated that during the first 24 h of growth, LS2i produced an active extracellular agarase, while reduced 101 extracellular agarase activity and no extracellular agarase activity were detected for R9 and R16, respectively when all three strains were grown for 24 h at 22C.

1 2 3 4

97kDa

66kDa

45kDa

30kDa

20,1kDa Town

Figure 5. Zymogram detection of extracellular agarase activity in the concentrated supernatants of strain LS2i and mutants R9 and R16. Lane 1: molecular mass markers; Lane 2: LS2i supernatant; Lane 3: R9 supernatant; Lane 4: R16 supernatant. The molecular mass markers were visualized by staining with CoomassieCape blue dye and the enzymatically active bands were detected by staining with Gran’s Iodine. The sizes of the molecular mass markers are shown in kDa. of

In order to determine the substrate specificity of strain LS2i, mutant R9 and mutant R16, TLC analysis was performed. Equal amounts of extracellular protein collected from 24 h cultures of strain LS2i and the mutant strains R9 and R16 were allowed to react with various substrates before TLCUniversity analysis (Figure 6). TLC confirmed that the extracellular agarase expressed in LS2i is active and that it hydrolyzes agarose to produce the oligosaccharides neoagarohexaose and neoagarotetraose (Figure 6, lane 3) (same result obtained in Chapter 3, Figure 6). Consistent with the results obtained from the zymogram (Figure 5), extracellular protein from strain R9 exhibited slight agarolytic activity, but significantly less than that produced by strain LS2i (Figure 6). Faint spots, representing neoagarohexaose and neoagarotetraose produced by an extracellular agarase expressed by strain R9, were visible in lane 4 (Figure 6). No extracellular agarolytic activity was however detected by TLC analysis in strain R16 (Figure 6, lane 5). Also consistent with the results obtained in Chapter 3, the extracellular agarase produced by strain LS2i did not hydrolyze either neoagarotetraose or 102 neoagarobiose. As can be expected, the proteins present in the extracellular fractions of strain R9 and strain R16 did not hydrolyze either neoagarotetraose or neoagarobiose.

1 2 3 4 5 6 7 8 9 10 11 12

Town neoagarobiose

neoagarotetraose Cape of neoagarohexaose

University Figure 6. TLC analysis of the oligosaccharides formed following incubation of the culture supernatants from strain LS2i, strain R9 or strain R16 with various substrates. Lane 1: molecular size markers; Lane 2: agarose incubated with buffer; Lane 3: agarose incubated with supernatant from strain LS2i; Lane 4: agarose incubated with supernatant from strain R9; Lane 5: agarose incubated with supernatant from strain R16; Lane 6: tetraose incubated with supernatant from strain LS2i; Lane 7: tetraose incubated with supernatant from strain R9; Lane 8: tetraose incubated with supernatant from strain R16; Lane 9: biose incubated with supernatant from strain LS2i; Lane 10: biose incubated with supernatant from strain R9; Lane 11: biose incubated with supernatant from strain R16; Lane 12: molecular size markers.

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4.4. Discussion

In an attempt to isolate one or more Pseudoalteromonas sp. LS2i mutants with an inability or reduction in ability to produce extracellular agarases, the chemical nitrosoguanidine (NTG) was used to mutate Pseudoalteromonas sp. LS2i. Conditions giving less than 1% survival were used to generate a population of mutant cells. A total of 15480 colonies were screened visually for reduced or no extracellular agarolytic activity and 16 mutants were isolated. The growth rates of most of the mutants were reduced by the mutations and only six of the mutants (R7, R9, R10, R11, R13 and R16) had similar OD600 values to LS2i after 24 h of growth at 22˚C in basal marine broth (BMB). Of these six mutants agarolytic activity was detected only for strain R10 (34% of the activity measured for LS2i) after 24 h in BMB (22˚C), while reduced extracellular agarolytic activity was visible for both strains R10 and R11 after incubation for 1 day on VNSS agar (22˚C). After 2 days of incubation on solid media, strain R10 produced agarolytic activity at similar levels toTown that of the wild type, while strain R11 still produced less agarolytic activity than the wild type after 3 days of incubation on VNSS agar. Mutant R13 started to produce agarolytic activity on the second day, while strain R9 started to pit the agar on the third day, followingCape incubation on VNSS agar. Mutants R7 and R16 were however the only strains thatof did not show any signs of extracellular agarase production after being incubated for 3 days on VNSS agar. Compared to the other strains that had similar growth rates to LS2i, strains R7, R9 and R16 appeared to have the greatest reduction in extracellular agarolytic activity over a three day incubation period on VNSS agar.

We were unable to measureUniversity the extracellular agarolytic activity of strains R4 and R8 since these two strains could not be cultured in BMB. Basal marine broth (BMB) is a low nutrient medium (Appendix A.1.7), ideal for inducing agarase production in LS2i as shown by Jaffray (1999). Strains R4 and R8 did however grow on the more nutrient rich VNSS agar. It is likely that the mutations in R4 and R8 have affected the ability of the two mutants to grow without certain nutrients. The growth of mutant strains of the fungal pathogen Cochliobolus carbonum with mutations in the SNF1 gene that is required for cell wall-degrading enzyme expression and virulence on maize, was affected to varying degrees depending on the carbon source that was present in the growth medium (Tonukari et al., 2000). One such carbon 104 source (of which the absence thereof might have affected the ability of R4 and R8 to grow) that is present in VNSS agar but not in BMB, is glucose. Both R4 and R8 were amongst the few strains that exhibited extracellular agarolytic activity, but to a lesser extent than the wild type, after incubation on VNSS agar for 1 day at 22˚C. It is thus also possible that the mutations in mutants R4 and R8 as well as R1, R5, R6, R12, R14 and R15, which were growth impaired when cultured in liquid broth but produced agarolytic activity after 24h or 48h on solid media, did not affect extracellular agarase production directly, but that the later onset of agarase production is a result of a reduction in the growth rate.

Except for a reduction in agarolytic activity, mutants R2 and R9 also produced brown pigment. The data presented in Figure 1 suggests that strain R2 was also growth impaired, while R9 had similar growth characteristics to the wild type. It is not unusual for agarolytic bacteria to produce pigments; i.e. three different pigmenting strains of agarolytic bacteria belonging to the species Cellulophaga pacifica were recently isolatedTown from a marine environment (Nedashkovskaya et al., 2004); the agarolytic bacterium Pseudoalteromonas gracilis B9 that was, like Pseudoalteromonas sp. LS2i, isolated from the red seaweed Gracilaria gracilis is also able to produce a pigmentCape (Schroeder et al., 2001). It has been documented that in some bacteria like Vibrio cholerae, Streptomyces griseus and Synechococcus cedrorum, pigment productionof occurs in response to conditions that are stressful to the bacteria (Coyne et al., 1992; Lee et al., 2001; Bisen et al., 1993). The mutations in strains R2 and R9 might thus be causing stress in these strains which leads to pigment production. A medium dependant expression of pigment, where the bacterium lose the pigment when grown in rich media, has been shown in other Pseudoalteromonas species (Holmström et al., 2002;University Egan et al., 2002). This suggests that pigmentation may be switched on and off depending on the available nutrient sources which in turn suggests that an inability to metabolize the breakdown products of agarose due to a lack of β-agarase I production in R2 and R9 might limit the available nutrient sources for these strains which then causes nutritional stress and leads to pigment production.

Consistent with the results obtained from the preliminary experiments (Figure 1), mutants R9 and R16 displayed similar growth rates to LS2i over a 24 h period (Figure 3). Mutants R9 and R16 are thus different from the Pseudoalteromonas atlantica mutant that was constructed through a gene replacement strategy, since the P. atlantica mutant was found to be growth 105 impaired (Belas et al., 1987). By using the ferricyanide assay we detected that LS2i already started to produce extracellular agarase after 2 h of growth, while no extracellular agarolytic activity was measured for either mutant R9 or mutant R16 during the 24 h growth period (Figure 4).

Both TLC and zymogram analysis revealed that although significantly less than the extracellular agarase produced by LS2i, mutant R9 produces an active extracellular agarase. The extracellular agarase produced by both LS2i and R9 is 30 kDa in size (Figure 5) and hydrolyzes agarose to produce neoagarohexaose and neoagarotetraose (Figure 6). The reason for detecting agarolytic activity in the extracellular fraction of 24 h cultures of mutant R9 using TLC (Figure 6) and zymogram (Figure 5) analysis and not during the 24 h preliminary growth (Figure 1) and growth experiments (Figure 4) is due to a difference in the agarase concentration in the extracellular fractions used in these experiments. Agarase activity was measured during the growth experiments by performing a ferricyanideTown assay on the non- concentrated culture supernatant of the strains while the supernatant used in both the zymogram and TLC experiments was concentrated 10 to 200 fold. In fact, when a ferricyanide assay was performed on concentrated Capesupernatant (25ug total protein/sample) of 24 h cultures of LS2i, R9 and R16, reduced agarolytic activity were detected for R9 (14% of activity measured for LS2i) but no activity wasof detected for strain R16 (results not shown). It is also possible that if a ferricyanide assay were done on an even more concentrated volume of supernatant from a 24 h culture of mutant R16, that agarolytic activity would have been detected as well. The results from the zymogram and TLC analysis therefore suggests that like LS2i, strain R9 produces the same extracellular agarase during the first 24 h of growth, but at significantly reducedUniversity levels.

When Belas and his coworkers (1987) constructed a Pseudoalteromonas atlantica mutant, they found that although the mutant had lost its primary agarase activity, minor agarolytic activity was still present. They concluded that this secondary agarase activity may be due to the enzyme described by Morrice et al (1983b; 1983c) as β-agarase II and by Groleau and Yaphe (1977) as neoagarotetraose hydrolase. After having constructed gene replacement mutants for two of the five agarase genes, aga50A and aga86E, of Saccharophagus degradans 2-40, Ekborg et al (2006) found that when grown on media supplemented with glucose, the mutants pitted the agar slowly, but less so than the wild-type. Redundancy in 106 extracellular enzymes is also common in bacterial and fungal plant pathogens. All of the pathogens that have been studied in detail have multiple genes for any particular enzyme activity (Brito et al., 2005; Tonukari et al., 2000). No extracellular agarase or agarolytic activity was detected during the first 24 h of growth for mutant R16 using either TLC or zymogram analysis, but since minor agarolytic activity was detected after growing strain R16 for several days on agar (results not shown), it is possible that this effect might be due to another agarase i.e. β-agarase II. It is also possible that R16, as seems to be the case for R9, still produces the 30 kDa extracellular agarase like LS2i, but at a further reduced rate than mutant R9. As mentioned before, it is not unusual for bacterial or fungal mutants, like R9 and R16, with altered extracellular enzyme production to still produce residual enzyme activity. The latter was the case for Serratia marcescens mutants where each mutant was affected in the production of a single extracellular protein, the mutants from Erwinia chrysanthemi defective in the secretion of pectinase and cellulose as well as the mutants from Pseudomonas aeruginosa which are defective in the formation of certain exoproteinsTown (Hines et al., 1988; Andro et al., 1984; Wretlind et al., 1984).

Since the mutations in strains R9 and R16 were inducedCape through chemical mutagenesis, the location of these mutations in the chromosome is uncertain. There are however several possibilities as to what might have happened ofduring mutagenesis. One possibility is that the mutation is located in a global regulator or in one of the genes involved in monitoring the secretion process of the agarase and possibly the secretion process of other extracellular enzymes as well. Bacterial or fungal mutants with the latter characteristics are usually pleiotropic and therefore produce reduced levels of several extracellular enzymes (see Section 4.1). Since theUniversity agarolytic Pseudomonas sp. SK38 isolated from seaweed, was found to produce more than one extracellular enzyme such as agarase, xylanase, protease and carboxymethylcellulase (Kang et al., 2003), it is entirely possible that Pseudoalteromonas sp. LS2i also produce multiple extracellular enzymes. Future characterization of mutants R9 and R16 can therefore involve resolving whether the production or the secretion process of the agarase, as well as that of other extracellular enzymes, was affected by the mutation. If other extracellular proteins as well as the agarase are found to be absent or produced at reduced levels, the mutation is likely to be located in a global regulator. If the agarase and/or other extracellular enzymes are found to be cell-bound in the mutants, the mutation is most likely located in a gene involved in the secretion process. Another possibility is that the specific 107 mutation could be in the β-agarase I gene itself. Since the sequence of the β-agarase I gene of LS2i is known (see Chapter 2), oligonucleotide primers can be designed to PCR amplify and sequence the gene from R9 and R16. This will determine whether the mutation is in the structural gene or not. If the mutation is found not to be in the structural gene, the method of Ryding and co-workers (1999) can possibly be used to locate the mutation on the chromosome. They used used NTG to identify new sporulation loci in Streptomyces coelicolor A3(2). Sporulation mutants appear white because they are defective in the synthesis of the grey polyketide spore pigment. The method they used to locate the mutation on the chromosome was in short as follows: A chromosomal library of wild type DNA was constructed. Clones from this library were arrayed on master plates and mated with each mutant in turn by replica plating, whereafter complimenting clones were identified. In each case, plasmid linkage of the phenotype was confirmed by reintroduction of the cloned DNA into the corresponding mutants. Complementing clones were identified for 12 of the 19 mutants that were subjected to complementation studies. AnalysisTown of the wild type chromosomal DNA in the complementing clones can then give an indication of the specific gene that was affected by the mutation. Previous attempts of electroporation and conjugation in order to perform transposon mutagenesis in PseudoalteromonasCape sp. LS2i with the vector pGp704 were however unsuccessful. Therefore, provided that a suitable vector can be found for conjugation studies, this procedure can potentiallyof be used to locate the mutations in R9 and R16. For the purpose of this study it was however sufficient to isolate a LS2i mutant or mutants that exhibited a significant reduction in extracellular activity. Therefore mutants R16 and R9 were used in further studies to determine the effect of the extracellular agarase of Pseudoalteromonas sp. LS2i on virulence in the seaweed Gracilaria gracilis. University 108

5. CHAPTER 5

INVESTIGATION OF THE ROLE OF THE EXTRACELLULAR AGARASE OF PSEUDOALTEROMONAS sp. LS2I ON VIRULENCE IN THE SEAWEED GRACILARIA GRACILIS

CONTENTS

5.1. Introduction ...... 110 5.2. Materials and Methods ...... 112 5.2.1. Antibody production against the purified agarase ...... 112 5.2.2. Western hybridization analysis to detect the presence of antibodies in the serum ...... 112 5.2.3. Western hybridization analysis to determine antibody specificity ...... 113 5.2.4. Generation of axenic Gracilaria gracilis ...... Town...... 113 5.2.5. Infection of Gracilaria gracilis ...... 114 5.2.6. Ultrastructure evaluation and colloidal gold immunolabeling ...... 114 5.2.6.1. Sample fixation and embedding schedule...... Cape 114 5.2.6.2. Post-embedding schedule for transmission electron microscopy ...... 115 5.2.6.3. Quantification of labeling ...... of 116 5.3. Results ...... 117 5.3.1. Specificity of the anti-agarase antibodies ...... 117 5.3.2. Pathogenicity of strain LS2i and mutants R9 and R16 ...... 118 5.3.3. Transmission electron microscopy of cross-sections from infected G. gracilisUniversity thalli...... 119 5.3.4. In situ localization of the extracellular Pseudoalteromonas sp. LS2i agarase in infected G. gracilis...... 124 5.4. Discussion ...... 132

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Summary

Polyclonal antibodies against the β-agarase purified from E. coli JM109 (pEB3) were generated in a rabbit. Western hybridization confirmed that the antibodies are specific to the 39 kDa agarase expressed from E. coli JM109 (pEB3). A pathogenicity assay showed that mutants R9 and R16, which have reduced extracellular activity, also exhibited reduced virulence in Gracilaria gracilis. It also confirmed that the agarase cloned from Pseudoalteromonas sp. LS2i into E. coli JM109 and purified from the latter does cause disease symptoms in G. gracilis. Histology studies indicated that bleaching of the G. gracilis thalli caused by the agarase leads to weakening of the cell structure. Immuno-gold labeled antibodies were used to localize and quantify the agarase inside the wild type and mutant infected thalli. The agarase was localized to the cell wall and enumeration of the gold particles in the electron micrographs showed a 44% reduction in the number of gold particles located in cell walls of mutant R16 infected thalli and an 83% reductionTown in the cell walls of mutant R9 infected thalli when compared to LS2i infected thalli.

Cape of

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5.1. Introduction

Cell walls of seaweeds are hydrophilic and soft macromolecular assemblies which are composed of fibrillar and matrix polysaccharides with minor structural proteins. In red alga, the fibrillar network is made of low crystalline cellulose, xylan or mannan and represents only 10% of the cell wall weight (Craigie et al., 1990; Lahaye et al., 2003). The polysaccharide component in red alga can consist of sulfated galactans, carrageenans or as in the case of Gracilaria, agars (Craigie et al., 1990). The mucilaginous (polysaccharide) component is regarded as a non- or para-crystaline matrix in which the fibrillar network is imbedded (Mackie and Preston, 1974).

A number of seaweed diseases caused by bacteria or fungi have been reported. Red rot disease which is caused by a fungus belonging to the Pythium genus is prevalent throughout the Porphyra cultivation areas in Japan (Woo and Kamei, 2003).Town In the Phillipines, ice-ice disease has been identified in Euchema denticulatum and Kappaphycus alvarezii. Agarolytic bacteria belonging to the genera, Cytophaga and Vibrio were identified as pathogens and were found to promote the ice-ice disease (Largo etCape al., 1995; Largo et al., 1998). Friedlander and Gunkel (1992) isolated several agarolyticof bacteria from the tips of Gracilaria conferta with white tip disease and found that several bacterial strains were involved in disease development on this seaweed. Weinberger and co-workers (1994) identified a bacterial strain designated OR-11 that was found to be responsible for white tip disease caused in Gracilaria conferta. A few years later they reported that tip necrosis on Gracilaria conferta was related to the excretion of active agents from bacteria belonging to the Corynebacterium- Arthrobacter-group andUniversity to the Flavobacterium-Cytophaga -group (Weinberger et al., 1997). Agar-digesting bacteria belonging to the Vibrio genus were also associated with a white to pinkish discoloration and thallus disintegration of tank-held stocks of a Gracilaria species (Lavilla-Pitogo, 1992)

Red algae produce sulfated galactans, agars and carrageenans, which are the main components of their cell walls (Craigie et al., 1990) and several bacteria, isolated from the surface of red seaweeds produce extracellular enzymes that have the ability to degrade these polysaccharides. A number of these epiphytic bacteria and their extracellular products have been characterized but the involvement of bacterial and fungal extracellular enzymes in the 111 virulence response of the host has been more extensively studied in plants. The standard assay to determine the involvement of a specific gene in pathogenisis is to mutate the candidate gene and observe whether the mutation resulted in a significant virulence defect in the host. A number of bacterial mutants from Erwinia carotovora (Hyytlainen et al., 2001; Laasik et al., 2005), Pseudomonas syringae (Fogliano et al., 2002), Ralstonia solanacearum (Huang et al., 1997; Tans-Kersten et al., 1998; Liu et al., 2005) and fungal mutants from Alternaria citri (Isshiki et al., 2001), Botrytis cinerea (Gronover et al., 2003; Brito et al., 2005), Cochliobolus carbonum (Tonukari et al., 2000), Fusarium oxysporum (Di Pietro et al., 2001) and Magnaporthe grisea (Wu et al., 2005), with reduced production in certain or all of their extracellular enzymes, have been isolated and used in pathogenicity studies on their respective host plants.

Therefore in order to better understand the role of the agarase in the appearance of disease symptoms in Gracilaria gracilis, we constructed PseudoalteromonasTown sp. LS2i mutants with reduced agarolytic activity (Chapter 4) and infected the seaweed with two of the mutants as well as the wild type to examine their ability to cause disease in the seaweed. We also managed to locate and quantitate the agarase insideCape the cell wall of the seaweed by using colloidal gold immunolabeling. of

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5.2. Materials and Methods

All media and solutions used in this study are listed in Appendix A

5.2.1. Antibody production against the purified agarase

Polyclonal antibodies against the agarase purified from E. coli JM109 (pEB3) as described in Chapter 3, were raised by immunizing a rabbit with 2 ml antigen emulsion. The antigen emulsion consisted of 50% purified agarase (110 ug/ml) and 50% Freund’s incomplete adjuvant (Ausubel et al., 1989). Four subsequent injections, the first two containing 2 ml antigen emulsion each and the last two containing 1 ml antigen emulsion each, were given intravenously every 7 days over a period of four weeks. The rabbit was bled before any injections (prebleed) were given and then again on the day of the second injection (bleed 2), continuing on a weekly basis (bleeds 3-8) over a seven-week period.Town The latter was done in order to obtain serum containing polyclonal antibodies against the purified agarase.

5.2.2. Western hybridization analysis to detect Capethe presence of antibodies in the serum of The prebleed as well as the 2nd, 3rd, 4th and 5th bleeds were tested for the presence of anti- agarase antibodies. Serial dilutions of the purified agarase were ‘dot-blotted’ onto five nitrocellulose membranes and the membranes were subsequently air-dried before western blot analysis was performed. The membranes were then blocked by immersion in 100 ml blocking solution (Appendix A.2.17) for 2 h at room temperature. The five bleeds were each first absorbed againstUniversity E. coli JM109 whole cell extracts (1 h at 37˚C with shaking) and the resultant protein conjugants were collected by centrifugation (10K rpm for 5 min at RT). The preabsorbed bleeds were then diluted 1:100 in blocking solution and each membrane was incubated separately in each bleed for 1 h at room temperature. The membranes were then each washed 2x in 100 ml 1x PBS for 15 min per cycle. Alkaline phosphatase-goat anti rabbit IgG conjugate (Sigma) were diluted 1:2000 in blocking solution before the membranes were incubated in it for 30 min at room temperature. The membranes were again washed 2x in 100 ml 1x PBS for 15 min per cycle. Developing substrate solution (Appendix A.2.17) was freshly prepared before it was added to the membranes. Colour development was allowed to 113 proceed for a few minutes, before the reaction was stopped by rinsing the membranes in water. The colour development on all the membranes was stopped simultaneously.

5.2.3. Western hybridization analysis to determine antibody specificity

O/N cultures of E. coli JM109 (pEB3) and E. coli JM109 (pBluescript SK) were inoculated into 100 ml flasks of LB (Appendix A.1.1) containing 100 ug/ml ampicillin and 120 ug/ml IPTG. These flasks were also incubated for 24 h at RT before the supernatants were collected (10K rpm for 15 min at 4˚C). The extracellular proteins present in the supernatants were precipitated with 50% Trichloroacetic acid (Merck) (Appendix B.14) for each culture and the pellets were each resuspended in 3 ml 10 mM phosphate buffer (pH 7). Protein concentrations were determined using the Bradford assay described in Appendix B.10.

A total of 7 ug from E. coli JM109 (pEB3) and E. coli JM109 (pBluescriptTown SK) and 0.5 ug of the purified agarase were separated on a 12% SDS-PAGE gel in accordance with the Laemmli method (Appendix B.12). The resulting proteins were transferred to a nitrocellulose membrane as outlined in Appendix B.16. In order toCape purify the anti-agarase antibodies present in bleed V as much as possible, Bleed V was preabsorbed against E. coli JM109 whole cell extracts as described in 5.2.2 beforeof it was precipitated with PEG as described in Appendix B.17. The membrane was blocked, incubated in bleed V, exposed to alkaline phosphatase-goat anti rabbit IgG, washed and developed as described in 5.2.2.

5.2.4. Generation of axenic Gracilaria gracilis University The method used for generating axenic G. gracilis was adapted from the procedure described by Jaffray and Coyne (1996). Healthy G. gracilis that was dark red and free of any epiphytic algae was collected from Saldanha Bay and was cut into 5 cm pieces. The algae pieces were then incubated in 100 ml sterile seawater (SSW) containing an ‘antibiotic cocktail’ (Appendix A.2.20) for 24 h at 15˚C with air bubbling through the flask. The pieces were then washed at least 3x in SSW.

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5.2.5. Infection of Gracilaria gracilis

Strain LS2i, mutant R9, mutant R16 as well as strain SS5g were cultured in 5 ml BMB (Appendix A.1.7) at 22˚C for 24 h at 100 rpm. Strain SS5g is a non-agarolytic bacterial strain that like LS2i was isolated from the surface of Saldanha Bay G. gracilis by Jaffray (1999). The axenic thalli were then injected at both ends with 100 ul of the particular bacterial suspension (LS2i, R9, R16 or SS5g) or with 15 ul purified agarase (0.24 ug/ul). The injections were performed with a sterile needle to a depth of approximately 1 cm. Four thalli were injected per experiment. The injected thalli were placed in Erlenmeyer flasks containing 100 ml BMB (Appendix A.1.7) and incubated at 22˚C for 5 days at 50 rpm. The infected thalli were monitored daily for the appearance of symptoms up to 5 days, whereafter they were prepared for light and transmission electron microscopy as described below.

5.2.6. Ultrastructure evaluation and colloidal gold immunolabelingTown

5.2.6.1. Sample fixation and embedding schedule Cape The procedure for the preparation of pathogenicity assay samples was adapted from the process described by Dykstra (1993). About 2mmof of the infected end of a thallus from each experiment was excised and washed in base buffer (bb) (Appendix A.2.21) to remove any excess material. The samples were immersed in 5-10 times the sample volume of a 2.5% fixative of glutaraldehyde in bb (Appendix A.2.21) and were incubated O/N at 22˚C. The samples were then rinsed twice for 5 min each in bb. The tissues were post-fixed in 1% osmium tetroxide (AppendixUniversity A.2.21) for 1 h at 22˚C, before rinsing twice (5 min each) in bb. The samples were then dehydrated by passing them through the following alcohol dilution series: 30% ethanol, 5 min 50% ethanol, 5 min 70% ethanol, 5 min 80% ethanol, 5 min 90% ethanol, 5 min 95% ethanol, 5 min 100% ethanol, 10 min (twice) 115

100% acetone, 10 min (twice)

The tissues were then infiltrated with Spurr resin (Spurr, 1969) as follows: Spurr resin and 100% acetone (1:1), O/N Spurr resin and 100% acetone (4:1), 5 h 30 min 100% Spurr resin O/N New 100% Spurr resin the next morning.

The samples were then individually placed in moulds which were then covered with Spurr resin. The tissues were polymerized for two days at 60˚C after which the polymerized wedges were stored at 22˚C.

5.2.6.2. Post-embedding schedule for transmission electron microscopy Town The Leica ultracuts ultramicrotone (Leica, Cambridge Ltd) was used to slice ultra-thin cross sections of the polymerized thalli, before the sections were mounted on nickel grids. Grids that were used to study the histology of the G. gracilisCape infected cells were stained as described in the last paragraph of this section, while grids that were used in immunogold labeling experiments were treated as describedof in the next paragraph before the grids were stained.

The procedure for the post-embedding schedule was adapted from the procedure described by Slot and Geuze in Beesley (1989). The grids were floated, section downwards, on phosphate buffered saline (PBSUniversity containing 1 % BSA) (PBS BSA) (Appendix A.2.21) for 5 min. The grids were then transferred to PBS containing glycine (Appendix A.2.21) for 3 min before they were washed twice for 1 min each with PBS BSA. Duplicate grids from each experiment were floated on a 1:100 dilution of bleed V (containing anti-agarase antibodies) in PBS BSA for 12 h. The grids were then washed five times (1 min each) with PBS BSA containing 0.1 % Tween (Appendix A.2.21), followed by three washes (1 min each) with PBS BSA. The grids were then floated for 2 h on a 1:50 dilution of 15 nm gold anti-rabbit probe in PBS BSA. The grids were rinsed five times (1 min each) with PBS BSA containing 0.1 % Tween and washed three times (1 min each) with PBS BSA. The conjugant label complexes were 116 fixed for 3 min at 22˚C with 1 % glutaraldehyde in PBS (Appendix A.2.21). The grids were then rinsed five times (1 min each) in ultrapure water.

Staining of the grids was performed with 2% uranyl acetate for 10 min (Appendix A.2.21), followed by five washes (1 min each) with ultrapure water. The grids were then stained with a second stain, Reynolds lead citrate (Appendix A.2.21), for 5 min before they were washed in a stream of ultrapure water for 2 min. The samples from each experiment were then visualized on a JEM-200CX transmission electron microscope (JEOL).

5.2.6.3. Quantification of labeling

To evaluate labeling density in the cell walls of all the different samples, 2 - 4 micrographs per sample were analyzed after colloidal gold immunolabeling. The labeling density in either the cytoplasm or on the grid where no cell components were presentTown was taken as background. Squares of 0.5 μm x 0.5 μm were randomly selected and drawn in the specific areas. The number of individual gold particles localized in each selected square was counted by hand and labeling density was measured as the numberCape of gold particles / 0.25 μm2 ± standard error. The gold particles located in the cell walls were counted in 24 – 32 squares / sample, while 20 – 37 squares / sample were ofcounted for background labeling. Background labeling was subtracted from the cell wall labeling for each specific sample before the data was represented in a histogram.

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5.3. Results

5.3.1. Specificity of the anti-agarase antibodies

To determine which bleeds had antibodies that were specific for the purified agarase from Pseudoalteromonas sp. LS2i, a preliminary screening of the prebleed as well as the 2nd, 3rd, 4th and 5th bleeds was done. The agarase, purified from E. coli JM109 (pEB3), was “dot- blotted” onto five nitrocellulose membranes and each membrane was probed with one of the bleeds. No signal was produced when the membrane was probed with antiserum obtained from the prebleed (Figure 1a). A positive reaction was detected with antisera from bleed 2, 3, 4 and 5 (Figure 1b, 1c, 1d and 1e). Bleed 5 was subsequently chosen for use in further experiments.

1 2 3 4 Town a Cape b of

Universityd

Figure 1. A preliminary ‘dot-blot’ western hybridization to determine which bleed contains anti-agarase antibodies. a) Prebleed; b) Bleed 2; c) Bleed 3; d) Bleed 4; e) Bleed 5. Position 1: 60 ng of purified agarase; Position 2: 30 ng of purified agarase; Position 3: 15 ng of purified agarase; Position 4: 7.5 ng of purified agarase.

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In order to determine the specificity of the polyclonal antibodies against the agarase purified from E. coli JM109 (pEB3), extracellular fractions of E. coli JM109 (pEB3) and E. coli JM109 (pBluescript SK) as well as a sample of the purified agarase were used in western hybridization studies. The extracellular fractions were separated on a 12% SDS-PAGE and transferred onto a nitrocellulose membrane. The nitrocellulose membrane was subjected to western hybridization, which showed that the anti-agarase antibodies obtained from the 5th bleed cross-reacted strongly to both the agarase purified from E. coli JM109 (pEB3) (Figure 2, lane 2) and the extracellular sample from E. coli JM109 (pEB3) (Figure 2, lane 3) by detecting a 39 kDa band in these samples. The anti-agarase antibodies did not detect any bands in the extracellular sample from the control, E. coli JM109 (pBluescript SK) (Figure 2, lane1).

1 2 3 Town

Cape 39kDA of

Figure 2: Western blot analysis of the 12% SDS-PAGE containing the following: lane 1: extracellular extract of E. coli JM109 (pBluescript SK); lane 2: purified agarase from E. coli JM109 (pEB3); lane 3: extracellular extract of E. coli JM109 (pEB3).The size of the protein band is shown in kDaUniversity (arrow).

5.3.2. Pathogenicity of strain LS2i and mutants R9 and R16

The pathogenicity assay devised by Jaffray and Coyne (1996) was firstly used to evaluate the virulence of the wild type Pseudoalteromonas sp. LS2i compared to the virulence of the two mutants Pseudoalteromonas sp. R9 and Pseudoalteromonas sp. R16 on the seaweed Gracilaria gracilis. The assay was secondly used to determine whether the agarase cloned 119 from LS2i and purified from E. coli JM109 (pEB3) caused disease symptoms in G. gracilis thalli. After 2 days of incubation the ends of the thalli injected with purified agarase were completely bleached, while the ends of the thalli injected with LS2i were starting to bleach. The ends of the thalli injected with the mutant R9 appeared healthy, while slight bleaching (but less compared to thalli ends injected with LS2i) was observed in the thalli ends injected with mutant R16 after 2 days of incubation. The two controls, axenic thalli injected with sterile seawater (SSW) (Figure 3f) and axenic thalli injected with the non-agarolytic strain SS5g (Figure 3e), remained dark and healthy throughout the 5-day incubation period of the assay. After 5 days of incubation, the ends of the thalli injected with either the purified agarase (Figure 3a) or LS2i (Figure 3b) were completely bleached. However while the LS2i injected thalli ends still retained their original structure and looked more firm, the purified agarase injected thalli ends appeared disintegrated and frail. Although the R9 and R16 injected thalli ends were not as dark and healthy as the thalli ends of the two controls after 5 days of incubation, the ends of these thalli injected with either mutantTown R9 (Figure 3c) or mutant R16 (Figure 3d) had a similar appearance and were definitely less bleached than the ends injected with either LS2i or purified agarase. Cape 5.3.3. Transmission electron microscopy of cross-sections from infected G. gracilis thalli. of

Cross sections of the injected thallus areas, corresponding to the bleached areas in the LS2i, R9, R16 and the purified agarase injected thalli, and to the injected area in the SS5g and SSW injected thalli, were analyzed under a transmission electron microscope. Comparisons between the cross sectionsUniversity of the wild type LS2i (Figure 4b), mutant R9 (Figure 4d) and mutant R16 (Figure 4c) injected thalli revealed no apparent differences in the cell structure. However a comparison between any of these three cross-sections and that of the cross sections of the SSW (Figure 4f) and the SS5g (Figure 4e) injected thalli revealed a clear disruption in the algal cell structure of the former samples. In general the cell walls of the LS2i, R9 and R16 injected thalli appeared to have lost some of its firmness and structure and had a somewhat smoother appearance compared to the cell walls of the SSW and SS5g injected thalli. The cytoplasm in the LS2i, R9 and R16 injected thalli also appeared to be pulling away from the cell wall in certain cells, while the cytoplasm in the SSW and SS5g injected thalli generally appeared to be intact. A comparison between the purified agarase 120 injected thalli (Figure 4a) and any of the other samples revealed an even more severe disruption in the algal cell structure of the purified agarase injected thalli. Compared to the LS2i, R9 and R16 injected thalli, the cell walls of the purified agarase injected thalli appeared to have partially collapsed in certain areas which resulted in a wavy appearance.

a b c d e f

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of Figure 3: Results of pathogenicity assay after 5 days incubation at 22˚C. a) Axenic thallus injected with agarase purified from E. coli JM109 (pEB3); b) Axenic thallus injected with Pseudoalteromonas sp. LS2i; c) Axenic thallus injected with Pseudoalteromonas sp. R9; d) Axenic thallus injected with Pseudoalteromonas sp. R16; e) Axenic thallus injected with bacterium SS5g; f) Axenic thallus injected with SSW. University 121

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Figure 4. Transmission electron micrographs of cross sections through mature G. gracilis thallus injected with the following: a) Purified agarase b) LS2i c) R16 d) R9 e) SS5g f) SSW. (cw): cell wall and (c): cytoplasm. Arrows indicating: a) Partially collapsed cell wall; b, c and d) Cytoplasm pulling away from cell wall; e and f) cytoplasm intact.

124

5.3.4. In situ localization of the extracellular Pseudoalteromonas sp. LS2i agarase in infected G. gracilis.

Immuno-gold labeled antibodies revealed that the agarase was associated with the cell walls of bleached G. gracilis. Varying densities of immuno-gold was detected in the cell walls of purified agarase (Figure 5a), LS2i (Figure 5b), R16 (Figure 5c) and R9 (Figure 5d) infected thalli, while only very few gold particles corresponding to unspecific background binding was detected in the cell walls of SS5g (Figure 5e) and SSW (Figure 5f) infected thalli. In all of the samples similar levels of background labeling were also detected in other cell components, such as the cytoplasm as well as on the grid next to the cross sections.

In order to obtain a quantitative measure of the differences in immunogold labeling between thalli infected with wild type LS2i and thalli infected with mutants R9 and R16, the density of labeling in the cell walls of the different samples was determinedTown as described in section 5.2.6.3 and the results of the quantitative analysis are summarized in Figure 6. Compared to LS2i infected thalli a 44% reduction in the amount of gold particles was observed in the cell walls of thalli infected with mutant R16, while an 83%Cape reduction was observed for thalli that were infected with mutant R9. The greatest amount of gold particles was detected in the cell walls of thalli injected with the purified agarase.of The mean of the density labeling in the cell walls was equal to the mean of the density labeling in the cytoplasm or sections on the grid next to the mounted cross sections of thalli infected with either the SS5g control or the SSW control. The latter confirmed that the gold particles present in either the cell walls, the cytoplasm or on the grid next to the cross sections of the two control samples were due to unspecific backgroundUniversity labeling.

125

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University Figure 5. subcellular localization of the agarase in G. gracilis thalli as determined by immunogold labeling. Cross sections through the mature G. gracilis thallus injected with the following are shown: a) Purified agarase b) LS2i c) R16 d) R9 e) SS5g f) SSW. (cw): cell wall. Arrows: 15nm gold particles.

131

Gold particles / 0.25μm / particles Gold 5

0 Purified agarase LS2i R16 R9 Ssw SS5g

-5 Sample

Figure 6. Quantitative evaluation of labeling density (number of gold particles/0.25 μm2 ) in the cell walls of G. gracilis thalli infected with different samples.Town Data shows the mean and standard error.

Cape of

University 132

5.4. Discussion

The polyclonal antibodies raised against the β-agarase that was purified from E. coli JM109 (pEB3) cross-reacted with a 39 kDa protein in the extracellular fraction of E. coli JM109 (pEB3). The size corresponds to the predicted size of the agarase protein expressed from the only ORF detected on the 1,8 kb DNA insert from strain LS2i that was cloned into pBluescript SK and designated pEB3 (described in Chapter 2). The antibodies did however not cross-react with any proteins in the extracellular fraction of E. coli JM109 (pBluescript SK). This suggests that the antibody is specific to the 39 kDa agarase expressed from E. coli JM109 (pEB3) and does not react with any other proteins expressed from E. coli JM109 (pEB3).

Virulence on G. gracilis was assessed by visually comparing the symptoms caused on the thalli by the wild type and the two mutants R9 and R16 after injectionTown with the different strains (pathogenicity assay). A clear difference was observed between the symptoms caused by wild type LS2i (which were more severe) and the symptoms caused by the two mutants. Symptoms caused by mutant R16 seemed to be slightlyCape more severe than disease symptoms caused by mutant R9, since bleaching was alreadyof visible 2 days after injecting the thalli with R16 whereas R9 injected thalli still looked healthy two days after injection. No visual difference between the two could however be detected after 5 days of incubation. The latter results therefore showed that the mutations in R9 and R16 definitely reduced the virulence of these strains in G. gracilis. Since we have shown that mutants R9 and R16 had reduced extracellular agarase production (described in Chapter 4) it is reasonable to assume that the reduction in extracellularUniversity agarase production by these mutants caused the reduced virulence in G. gracilis. Several examples exist of bacterial and fungal mutants with a reduced production in plant cell wall-degrading enzymes that have been shown to also have reduced virulence on their respective host plants (see section 5.1). The observation that bleaching in thalli injected with purified agarase was more severe than that observed in thalli injected with LS2i, confirmed that the agarase purified from E. coli JM109 (pEB3) and expressed extracellularly in LS2i is responsible or at least partially responsible for causing disease in the alga G. gracilis.

133

Histology studies of the infected thalli cells revealed a clear disruption in the structure of the bleached cells. After the 5 day incubation period, a similar level of disruption was apparent in G. gracilis cells that were injected with either the wild type strain LS2i, or the mutants R9 and R16. A shorter incubation period after infection of the thalli with the different strains (i.e. before the appearance of disease symptoms on the mutant injected thalli) probably would have revealed a difference in the level of cell disruption between the wild type and mutant infected thalli. Since the cells of the unbleached SS5g or SSW injected thalli were not disrupted and still healthy in appearance, it appears as if cell disruption only occurs in bleached cells. Schroeder et al. (2003) hypothesized that agarase degradation of the mucilaginous component (agar) in the G. gracilis cell wall weakened the overall structure of the cell wall which resulted in the collapse of the fibrillar component of the thallus. This observation is consistent with our observations. The severely disrupted cells (more disrupted than LS2i and mutant infected thalli) of the purified agarase injected thalli confirmed the visual results of the pathogenicity assay where the purified agaraseTown injected thalli were so severely bleached that the thalli has lost its original structure and firmness.

Immunogold studies have been successfully used toCape localize fungal or bacterial proteins inside infected host cells which include various types of plant cells, guinea-pig and human alveolar macrophages (Wubben et al., 1994; ofRechnitzer et al., 1992; Benhamou, 1989; Barrasa, 1998). It has also been used for the in situ detection of the β(1-4) agarase of Pseudoalteromonas gracilis B9 in infected G. gracilis (Schroeder et al., 2003). Immuno-gold labeled antibodies raised against the agarase purified from LS2i localized varying levels of agarase in the cell walls of the thalli injected with the purified agarase, the wild type LS2i and the mutants R9 and R16.University Enumeration of gold particles in electron micrographs was used to obtain a quantitative measure of the differences in immunogold staining. A 44% reduction in the number of gold particles located in the cell walls of mutant R16 infected thalli and an 83% reduction in the cell walls of mutant R9 infected thalli was observed when compared to LS2i infected thalli. This result is consistent with the severity of the disease symptoms observed on the infected thalli during the pathogenicity assay and confirmed that the amount of agarase present in the cell walls of G. gracilis thalli is directly linked to the severity of the disease symptoms observed on the thalli. The highest amount of agarase was however detected by gold-labeled antibodies in the cell walls of purified agarase infected cells. This result is consistent with the severe bleaching observed during the pathogenicity assay and the 134 severely disrupted cells observed during transmission electron microscopy studies of purified agarase infected thalli. This observation again confirmed the relationship between the amount of agarase present in the cell wall and the severity of the observed disease symptoms. Similar results were also obtained by Schroeder et al. (2003) where G. gracilis thalli was infected with an agarolytic strain, Pseudoalteromonas gracilis B9 as well as the purified β-agarase produced by the latter strain.

Since the location of the mutations in R9 and R16 is unknown one could only speculate on why the agarolytic activity of the specific mutants is reduced. We know from the results of zymogram and TLC analysis (Chapter 4) that after being cultured for 24 h in liquid broth, mutant R9 produces an active extracellular agarase with the same size and the same mode of action as the extracellular agarase produced by wild type LS2i but with significantly reduced activity. This result is consistent with the observation that when mutant R9 was injected into G. gracilis, less bleaching of the thallus occurred and reduced levelsTown of agarase was detected in the thallus cell wall when compared to LS2i injected thalli after the 5-day incubation period. Cape No extracellular agarolytic activity was detected when zymogram and TLC analysis was performed on the supernatant of a 24 h cultureof of mutant R16 (Chapter 4). However after the 5-day incubation period, R16 infected thalli had a higher concentration of agarase in the cell wall than that detected in the cell walls of R9 infected thalli. One possible explanation for this occurrence might be that the mutation in R16 causes a delay in extracellular agarase production in this strain and that extracellular agarase activity possibly would have been detected by zymogramUniversity and TLC analysis after a longer than 24 h incubation period which would explain the detection of agarase in the cell walls of R16 injected thalli after the 5-day incubation period. Kim et al., (2003) found that a mutation in the luxSVv gene of the human pathogen Vibrio vulnificus resulted in a delay of extracellular protease production during stationary phase. The latter observation, together with other results obtained, lead the authors to conclude that the LuxS quorum-sensing system plays an important role in co-coordinating the expression of virulence factors such as protease production. It is therefore possible that a mutation in a similar kind of gene as the luxSVv gene in Vibrio vulnificus is present in mutant R16. Another possible explanation might be that the agarase protein in R16 is still produced but is inactive. The agarase detected in the cell wall by the gold-labeled antibody would then 135 be inactive. Since zymogram and TLC analysis detected no extracellular agarolytic activity in a 24 h culture of R16, disease symptoms occurring in the R16 injected seaweed as detected by the pathogenicity assay could then be attributed to other agarases produced at a detectable level by R16 after a 24 h incubation period. The latter hypothesis can only be true if R9 is then a pleiotropic mutant (see section Chapter 4, section 4.1) where the export of additional extracellular agarases produced after the 24 h incubation period is also reduced, thus explaining the more severe disease symptoms observed in R16 infected thalli compared to R9 infected thalli. If the latter hypothesis is true and the agarase detected in the cell wall of R16 infected thalli is in fact inactive, it is uncertain why reduced levels of agarase were detected in comparison to the wild type. A similar observation was however made for Bacillus megaterium where a mutation in the CcpA gene resulted in the production of reduced amounts of inactive CcpA protein (Krause et al., 1997).

In this chapter we have provided conclusive evidence that the extracellularTown agarase produced by an agarolytic bacterium causes disease symptoms in the seaweed Gracilaria gracilis and that the agarase can be located in the cell wall of the infected seaweed. The results of this work have initiated a number of subsequent studiesCape that ultimately will deepen our understanding of the molecular basis of virulence in the complicated process of seaweed pathogen interaction. The next step would beof to determine whether the extracellular agarase produced by mutant R16 is in fact inactive or whether the production thereof is just delayed. It must also be determined whether mutant R9 is a pleiotropic mutant and whether other agarases are produced in any of the strains after a longer than 24 h incubation period (zymogram and TLC analysis did not detect any other agarase activity in 24 h old R16 cultures). Other promisingUniversity directions for future research include exploring the influence of environmental cues such as water temperature and nitrogen content on agarase production by epiphytic bacteria. 136

6. CHAPTER 6

GENERAL DISCUSSION

The aim of this study was to obtain a better understanding of the interaction between the seaweed Gracilaria gracilis and the bacterial G. gracilis pathogen Pseudoalteromonas sp. LS2i. Therefore the role of the extracellular agarase enzyme produced by Pseudoalteromonas sp. LS2i in the virulence response was investigated.

In order to clone the primary agarase gene of Pseudoalteromonas sp. LS2i, a genomic library was constructed in E. coli JM109. This led to the successful cloning and sequencing of an extracellular agarase from Pseudoalteromonas sp. LS2i. An extracellular agarase precursor cloned from Pseudoalteromonas sp. CY24 had the greatest sequence identity to the cloned agarase. The agarase also showed sequence similarity to several Townother agarases, which are all representatives of family 16 of the glycoside hydrolases (GH-16), as well as to a number of other enzymes which are not agarases, but belong to family GH-16 subgroups. Two glutamic acid residues, Glu143 and Glu148, which may act as catalytic residues, were identified in the ORF through multiple sequence alignment. PutativeCape promoter regions, a putative transcriptional start site and a putative ribosome-bindingof site were identified upon analysis of the sequence upstream of the ORF of the agarase cloned from strain LS2i. Primer extension analysis should be employed to determine the exact location of the promoter regions and the transcriptional start site of the cloned agarase.

The agarase cloned into E. coli JM109 was purified from the extracellular fraction of E. coli JM109 (pEB3) and characterizedUniversity as a β-1,4 agarase. Although it was pure enough for raising polyclonal antibodies, two bands were visible when a high concentration of this protein was electrophoresed on a SDS-PAGE gel. Future work should involve the determination of the N-terminal amino acid sequence of each protein. A comparison of the N-terminal sequences of these proteins to the DNA sequence of the agarase cloned into E. coli JM109 (pEB3) will reveal whether both bands represent this particular agarase or whether one of the bands is due to impurities from the extracellular fraction of E. coli JM109 (pEB3).

137

The purified agarase had the same mode of action as the agarase produced extracellularly by Pseudoalteromonas sp. LS2i, suggesting that the primary extracellular agarase has been cloned. It can be attempted in future to clone other agarases from Pseudoalteromonas sp. LS2i. Characterization of the other agarases in conjunction with the results already obtained with the extracellular agarase will enable us to determine the role of each agarase in the infection process, which in turn will lead to a better understanding of the infection process and the subsequent virulence response in Gracilaria.

The size of the purified agarase that was produced extracellularly by E. coli JM109 (pEB3) is 39 kDa and is consistent with the theoretical size predicted from the nucleotide sequence of the cloned agarase. The size of the extracellular agarase produced by Pseudoalteromonas sp. LS2i is however only 30 kDa. This suggests firstly that the extracellular agarase is processed differently during secretion by Pseudoalteromonas sp. LS2i than when secreted by E. coli JM109 (pEB3), and secondly, that the size of the mature protein Townexcreted by Pseudoalteromonas sp. LS2i is smaller than the theoretical size of the agarase. It is not uncommon for the size of secreted proteins to be smaller than the theoretical size predicted from the nucleotide sequence. One example of thisCape is the AgaA agarase produced by Zobellia galactanivorans that has a theoretical mass of 60 kDa. The size of the agarase produced in the culture medium is however only 31 kDa. ofThis is due to the presence of two C-terminal domains of unknown function that are processed during secretion of the enzyme (Jam et al., 2005). Other examples are the AgaO agarase secreted by strain JAMB-A94 and the dagA agarase secreted by Streptomyces lividans 66 (Otha et al., 2004a; Bibb et al., 1987). The authors suggested in the latter two cases that AgaO and dagA have signal peptides which are removed during secretion.University

Several Pseudoalteromonas sp. LS2i mutants with reduced agarolytic activity were isolated. Two of these, mutants R9 and R16, which exhibited a significant reduction in agarolytic activity and had similar growth rates to the wild type, were characterized to some degree. The reduction in extracellular agarase production in mutants R9 and R16 caused a reduced virulence of these strains in G. gracilis. Although symptoms (bleaching of the G. gracilis thallus) were visible on both wild type and mutant injected thalli 5 days after the thalli had been injected with the different strains, symptoms caused by strain LS2i were more severe than symptoms caused by the mutants. A disruption in the structure of bleached cells was 138 apparent, while unbleached cells remained intact. The agarase was localized to the inside of the cell walls of infected G. gracilis thalli. A quantitative measure of the amount of agarase enzyme inside thalli infected with either the wild type, the mutants or the purified agarase confirmed that the amount of agarase present in the cell wall is directly linked to the severity of the disease symptoms. The experiment indicated that less agarase is present in mutant infected cells compared to cells that have been infected with strain LS2i.

While mutant R9 produced the extracellular agarase at significantly reduced levels compared to the wild type, it is uncertain whether the minor extracellular agarolytic activity detected in mutant R16 is due to reduced levels of the primary agarase or to the presence of other agarases produced by the mutant. The mutation in R16 may have caused a delay in extracellular agarase production in this strain. Future work can therefore involve TLC and zymogram analysis of samples taken at different time points between 24 h and 5 days to determine whether this is indeed the case. If the latter is true, theTown onset of extracellular agarase production in mutant R16 could be determined. It is also possible that the agarase protein in mutant R16 is still produced but is inactive. Thus, the agarase detected in the cell wall of the G. gracilis thalli 5 days after infection isCape inactive and disease symptoms present on the thalli are in fact caused by other agarases produced by mutant R16. In order to determine whether the agarase produced by mutantof R16 is inactive, the following experiment could be performed. Since the sequence of the extracellular agarase is known, the gene and its upstream region could be amplified by PCR from the chromosomal DNA of mutant R16 and cloned into an expression vector such as pBluescript. Expression of the cloned gene in E. coli could be determined by performing SDS-PAGE analysis where the expression pattern of the extracellular proteinsUniversity produced by E. coli harboring the recombinant plasmid is compared to the expression pattern of the extracellular proteins produced by E. coli without the foreign DNA. Agarolytic activity or the absence thereof could be determined by performing a ferricyanide reducing sugar assay on the extracellular fraction produced by E. coli harboring the recombinant plasmid.

Further characterization of mutants R9 and R16 should involve obtaining a better understanding of the nature of the mutations. It should therefore firstly be determined whether these mutants are pleiotropic, i.e. whether excretion of other extracellular enzymes by the mutants was also affected. If the latter is found to be the case, the mutation is most 139 likely located in a global regulator or in one of the genes involved in monitoring the secretion process of other extracellular enzymes as well. If the agarase and/or other agarases are found to be produced at significant levels but are rendered cell-bound, the mutation is probably in one of the genes involved in the secretion process. In order to determine whether the mutation is in the structural gene, oligonucleotide primers can be designed to PCR amplify and sequence the β-agarase gene from R9 and R16.

Jaffray et al. (1996) provided evidence that at least one of the modes of bacterial virulence towards G. gracilis involves the production and secretion of agarases. Schroeder et al. (2003) confirmed the relationship between disease symptoms exhibited by infected G. gracilis and the agarolytic phenotype of the epiphytic bacterium. By infecting Gracilaria with wild type LS2i, as well as mutants producing reduced levels of agarase, this study demonstrated that there is a direct relationship between the severity of thallus bleaching and the amount of agarase produced by the specific bacterial strain. Therefore this studyTown has provided conclusive evidence that the agarase produced by Pseudoalteromonas sp. LS2i causes disease symptoms in the seaweed Gracilaria gracilis. In accordance with the results obtained by Schroeder et al. (2003), the Pseudoalteromonas sp.Cape LS2i agarase was also shown to be associated with the cell wall of the infected thallus. of In order to conduct mariculture of Gracilaria successfully at Saldanha Bay, a way must be found to prevent the collapses of the Gracilaria beds. Gracilaria growing in the ocean will never be free of agarolytic bacterial epiphytes that can potentially cause disease. Knowledge gained from this study provided a better understanding of the disease caused by the epiphytic agarolytic bacteria onUniversity Gracilaria gracilis. Since we have established that the amount of agarase produced by the epiphytic bacterium Pseudoalteromonas sp. LS2i is directly linked to the severity of the disease symptoms appearing on the thallus of G. gracilis, it may be useful to develop a kit that can be used to quantitate the agarase produced by epiphytic bacteria on raft-cultivated G. gracilis. This should allow early harvesting of the crop before the appearance of symptoms on the thalli. 140

7. APPENDIX A

MEDIA AND SOLUTIONS

A. CONTENTS

A.1. Media ...... 142 A.1.1. Luria-Bertani broth (LB)...... 142 A.1.2. Luria-Bertani agar (LA) ...... 142 A.1.3. Marine Broth (MB) ...... 142 A.1.4. Marine Agar (MA) ...... 142 A.1.5. Basal marine broth (BMB)...... 143 A.1.6. VNSS medium (Egan et al., 2002; Marden et al., 1985) ...... 143 A.1.7. VNSS agar (Egan et al., 2002; Marden et al., 1985) ...... 143 A.2. Solutions ...... 143 A.2.1. General stock solutions ...... Town...... 143 A.2.2. Solutions for protein purification ...... 145 A.2.3. Solution for protein precipitation ...... 146 A.2.4. Solutions for ferrecyanide assays...... Cape 146 A.2.5. Solutions for chromosomal DNAof extractions ...... 147 A.2.6. Solutions for agarose gel electrophoresis ...... 148 A.2.7. Solutions for the preparation of competent cells ...... 149 A.2.8. Solution for electrophoresis of DNA on a agarose gel ...... 149 A.2.9. Solutions for transfer of DNA from agarose gel onto membrane ...... 150 A.2.10. Solutions for Heinekoff shortenings ...... 150 A.2.11. SolutionUniversity for Ammonium acetate precipitation ...... 151 A.2.12. Solutions for Bradford assays ...... 152 A.2.13. Solutions for SDS-PAGE gels ...... 152 A.2.14. Solution for zymogram detection...... 154 A.2.15. Solutions for TLC ...... 154 A.2.16. Solutions for performing NTG-mutagenesis ...... 154 A.2.17. Solutions for performing western blot analysis ...... 155 A.2.18. Solution for electroblotting of proteins onto a nitrocellulose membrane ...... 156 A.2.19. Solutions for precipitating antibodies with PEG ...... 156 141

A.2.20. Solutions for ‘antibiotic cocktail’ for axenic treatment of seaweed ...... 157 A.2.21. Solutions for electron microscopy ...... 158

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All media were autoclaved at 121 ˚C for 20 min prior to use, unless otherwise specified. Water used for making solutions, media and diluting buffers was purified using a Milli-RO Plus (Millipore) water purification system.

A.1. Media

A.1.1. Luria-Bertani broth (LB) Tryptone 10 g Yeast extract 5 g NaCl 5 g Water to 1 l

A.1.2. Luria-Bertani agar (LA) Tryptone 10 g Yeast extract 5 g Town NaCl 5 g Agar 15 g Water to Cape 1 l

A.1.3. Marine Broth (MB) of NaCl 30.0 g MgCl2.6H2O 2.3 g KCl 0.3 g Casamino acids 5.0 g Yeast extract 1.0 g Glucose (Saarchem)University 2.0 g Water to 1 l

A.1.4. Marine Agar (MA) NaCl 30.0 g MgCl2.6H2O 2.3 g KCl 0.3 g Casamino acids 5.0 g Yeast extract 1.0 g Glucose (Saarchem) 2.0 g 143

Agar 20.0 g Water to 1 l

A.1.5. Basal marine broth (BMB) NaCl 30.0 g

MgCl2.6H2O 2.3 g KCl 0.3 g Casamino acids 5.0 g Yeast extract 1.0 g Water to 1 l

A.1.6. VNSS medium (Egan et al., 2002; Marden et al., 1985) Peptone 1.0 g Yeast extract 0.5 g Glucose 0.5 gTown

FeSO4.7H2O 0.01 g

Na2HPO4 0.01 g NNS (Appendix A.2.16) Cape 1 l

A.1.7. VNSS agar (Egan et al., 2002; Mardenof et al., 1985) Peptone 1.0 g Yeast extract 0.5 g Glucose 0.5 g

FeSO4.7H2O 0.01 g

Na2HPO4 0.01 g Agar University 15 g NNS (Appendix A.2.16) 1 l

A.2. Solutions

A.2.1. General stock solutions  Ampicillin (100 mg/ml) Ampicillin (Sigma) 2 g Water to 20 ml 144

Filter sterilize and store aliquots at 4 ˚C. Dilute 1:1000 into media for final concentration of 100 ug/ml.

 IPTG (100 mM) IPTG 0.477 g Water to 20 ml Filter sterilize and store aliquots at 4 ˚C. Dilute 5 ml/1 l media for final concentration of 120 ug/ml.

 1 M Tris-HCl Tris 12.1 g Water to 100 ml Dissolve the Tris in 80 ml water and adjust pH to required level with concentrated HCL. Finally make up to a final volume of 100 ml. Town  0.5 M EDTA EDTA (Saarchem) 93.05 g NaOH (Saarchem) Cape 10 g Water to 500 ml Dissolve the EDTA and the NaOHof in 400 ml water. Adjust the pH to 8 and make up to a final volume of 500 ml.

 TE buffer (Tris-EDTA) 1 M Tris-HCl (pH 7.6) 1 ml 0.5 MUniversity EDTA 200 ul water to 100 ml

 70% EtOH Absolute EtOH (Merck) 70 ml Water to 100 ml Do not autoclave. Store at -20 ˚C.

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 50% Glycerol Glycerol (100%) 50 ml Water to 100 ml

A.2.2. Solutions for protein purification  Sodium azide (20%) Sodium azide 2 g Water to 10 ml Dilute 1 ml/1 l buffer for final concentration of 0.02%. Need not be autoclaved.

 20 mM Tris-HCl (pH 7) 1 M Tris-HCl (pH 7) (Appendix A.2.1) 20 ml Water to 1 l Town

 5 M NaCl NaCl Cape 292 g Water to 1 l of  1 M NaCl NaCl 58.4 g 20 mM Tris-HCl (pH 7) to 1 l

 0.2 MUniversity NaCl 1 M NaCl 200 ml 20 mM Tris-HCl (pH 7) to 800 ml

 0.2 M / 0.4 M NaCl Dilute 1M NaCl accordingly in 20 mM Tris-HCl (pH 7).

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 0.1 M Phosphate buffer (pH 7) (A) 0.2 M monobasic sodium phosphate solution

NaH2PO4 (Saarchem) 27.8 g Water to 1 l (B) 0.2 M dibasic sodium phosphate solution

NaH2PO4.7H2O (Saarchem) 53.65 g Water to 1 l Combine 39 ml of A with 61 ml of B and dilute with water to a total of 200ml.

 10 mM Phosphate buffer (pH 7) 0.1 M Phosphate buffer (pH 7) 100 ml Water to 1 l Town A.2.3. Solution for protein precipitation  TCA (50%) TCA (Merck) Cape 100 g Water to 200 ml of A.2.4. Solutions for ferricyanide assays  Mineral 1 solution

K2HPO4 (Saarchem) 6.0 g Water to 1 l University  Mineral 2 solution NaCl 12.0 g

(NH4)2SO4 (Roche) 12.0 g

KH2PO4 6.0 g

CaCl.2H2O (Saarchem) 2.5 g

MgSO4.7H2O (Merck) 2.5 g Water to 1 l

147

 20 mM PIPES solution PIPES (Merck) 0.6 g Mineral 1 solution 5.0 ml Mineral 2 solution 5.0 ml Water to 100 ml Adjust pH to 6.8 with 10 M NaOH to dissolve the PIPES. Adjust volume to 100ml.

 Agarose substrate (1%) 20 mM PIPES solution 90 ml agarose 1 g water to 100 ml

 Stop reagent Town Na2HPO4.7H2O (Saarchem) 1 g NaOH (Saarchem) 1 g Water to Cape 100 ml Replace monthly and need not be autoclaved. of  Colour reagent

K3[Fe(CN)6] (Merck) 116 mg NaOH 220 mg Water to 100 ml Store Universityin dark and replace monthly. Need not be autoclaved.

A.2.5. Solutions for chromosomal DNA extractions

 10% SDS (Sodium dodecyl sulphate) SDS (BDH) 10 g Water to 100 ml Stir on warm plate and do not overheat. Do not autoclave.

148

 Proteinase K (20 mg/ml) Proteinase K (Sigma) 20 mg Sterile water to 1 ml Do not autoclave. Store at -20˚C

 5 M NaCl NaCl 29.22 g Water to 100 ml

 CTAB/NaCl NaCl 4.1 g CTAB (USB) 10 g Water to 100 ml Dissolve the NaCl in 80 ml of water. Slowly add Townthe CTAB (cetyltrimethylammonium bromide). Heat while stirring slowly. If necessary, heat to 65 ˚C to dissolve. Adjust to a final volume of 100 ml with water. Cape  Chloroform (Merck) / isoamyl alcohol (Merck) Mix at ratio 24:1 of

 RNase (10 mg/ml) RNase A (Sigma) 0.1 g 1 M Tris-HCl (pH 7.5) 100 ul 5 M NaClUniversity 3 ml Water to 10 ml Heat for 15 min at 100 ˚C and allow to cool slowly to room temperature. Do not autoclave. Aliquot into eppendorf tubes and store at -20 ˚C.

A.2.6. Solutions for agarose gel electrophoresis

 50x TAE (Tris-acetate buffer) Tris 242 g Glacial acetic acid (Saarchem) 57.1 ml 149

0.5 M EDTA 100 ml Water to 1 l

 1x TAE 50x TAE 200 ml Water to 10 l

 EtBr 10 mg/ml (Ethidium Bromide) EtBr (Sigma) 0.1 g Water to 10 ml Shake well to dissolve. Do not autoclave. Powerful mutagen – wear gloves and clean spills with isopropanol.

A.2.7. Solutions for the preparation of competent cells Town

 0.1 M CaCl2

CaCl2.2H2O 14.7 g Water to Cape 1 l of

 0.1 M MgCl2

MgCl2.6H2O 20.3 g Water to 1 l

A.2.8. Solution Universityfor electrophoresis of DNA on a agarose gel

 Gel tracking dye Bromophenol blue 62.5 g Sucrose (Saarchem) 10 g 0.5 M EDTA 1 ml Water to 25 ml

150

A.2.9. Solutions for transfer of DNA from agarose gel onto membrane

 0.25 M HCl HCl (Saarchem) 21.35 ml Water to 1 l Do not autoclave

 0.4 M NaOH NaOH 16 g Water to 1 l

 0.4 M NaOH / 1 M NaCl NaOH 16 g NaCl 58.44Town g Water to 1 l

 20 x SSC (Sodium chloride tri-sodium citrate) NaCl Cape 17.5 g Tri-Na Citrate (Saarchem) of 8.82 g Water to 100 ml Dissolve NaCl and tri-Na citrate in 80 ml water, adjust pH to 7.4 with NaOH and make up to 100 ml with water

 2 x SSCUniversity 20 x SSC 100 ml Water to 1 l

A.2.10. Solutions for Heinekoff shortenings

 Exonuclease III buffer 1 M Tris-HCl (pH 7.6) 660 ul

1 M MgCl2 6.6 ul Water to 10 ml 151

 10 x S1 nuclease buffer KOA (Saarchem) 0.98 g NaCl 11.7 Glycerol (Saarchem) 50 ml

ZnSO4 (Saarchem) 0.3 g Water to 100 ml Adjust the pH to 4.6 with glacial acetic acid.

 S1 nuclease mix (enough for 15 tubes) 10 x S1 nuclease buffer 24.6 ul

sterile H2O 155.4 ul S1 nuclease (add prior to use) 36 U Town  S1 nuclease stop 1 M Tris base 3 ml 0.5 M EDTA Cape 1 ml Water to 10 ml of  Klenow mix 1 M Tris-HCl (pH 7.6) 0.2 ml

1 M MgCl2 70 ul Water to 10 ml University A.2.11. Solution for Ammonium acetate precipitation

 7.5 M Ammonium acetate Ammonium acetate (Saarchem) 262.8 g Water to 500 ml Dissolve ammonium acetate in 400 ml water, adjust pH to 7.5 using glacial acetic acid and adjust volume to 500 ml.

152

A.2.12. Solutions for Bradford assays

 Bovine serum albumin (BSA) (1 mg / ml) BSA 0.01 g Sterile water 10 ml Do not autoclave. Aliquot and store at – 20 ˚C

 0.15 M NaCl NaCl 0.88 g Water to 100 ml

 Coomassie brilliant blue solutions Town Coomassie brilliant blue G250 (Saarchem) 100 mg Absolute EtOH 50 ml Phosphoric acid (Saarchem) Cape 100 ml Water to 1 l In a volumetric flask dissolve ofCoomassie brilliant blue G250 in EtOH. Add 85 % phosphoric acid. Bring volume to 1 l with water. Filter through Whatman no 1 filter paper. Do not autoclave. Store at 4˚C.

A.2.13. Solutions for SDS-PAGE gels University  4 x Separating gel buffer Tris base 18.17 g 10 % SDS 4 ml Water to 100 ml Adjust pH to 8.8 with HCl and add water to a final volume of 100 ml.

 4 x Stacking gel buffer Tris base 6.06 g 10 % SDS 4 ml 153

Water to 100 ml Adjust pH to 6.8 with HCl and add water to a final volume of 100 ml.

 10 % Ammonium persulfate Ammonium persulfate (Pharmacia) 1 g Water to 10 ml Do not autoclave. Aliquot into eppendorf tubes and store at -20 ˚C.

 10 x SDS-PAGE Running buffer Tris base 30 g Glycine 144 g 10 % SDS 100 ml

 2 x SDS-PAGE sample buffer Town Glycerol 2 ml 10% SDS 2 ml Bromophenol blue Cape 0.25 mg 4 x Stacking gel buffer 2.5 ml β-mercaptoethanol (Merck) of 0.5 ml Water to 10 ml

 Coomassie blue dye staining solution CoomassieR brilliant blue R250 (Sigma) 2.5 g IsopropanolUniversity (Merck) 250 ml Glacial acetic acid 100 ml Water to 1 l Do not autoclave

 Destaining solution Glacial acetic acid 70 ml Water to 1 l Do not autoclave

154

A.2.14. Solution for zymogram detection

 Gran’s Iodine Iodine 5 g Potassium iodide 10 g Water to 500 ml Dissolve the iodine and potassium iodide in 100 ml water. Adjust the volume to 500 ml with water. Do not autoclave and store in dark.

A.2.15. Solutions for TLC

 Developing solvent n-butanol (Merck) 500 ml Glacial acetic acid 250 Townml Water 250 ml

 Naphthoresorcinol reagent Cape Solution A Naphthoresorcinol (Sigma)of 0.04 g ETOH to 20 ml Solution B ETOH 37.5 ml Sulphuric acid (Merck) 10 ml CombineUniversity solution A and B in 1:2 ratio (20 ml : 40 ml). Allow to cool and use immediately.

A.2.16. Solutions for performing NTG-mutagenesis

 Nine Salts Solution (NSS) (Egan et al., 2002; Marden et al., 1985) NaCl 17.6 g

Na2SO4 1.47 g

NaHCO3 0.08 g KCl 0.25 g 155

KBr 0.04 g

MgCl2.6H2O 1.87 g

CaCl2.2H2O 0.41 g

SrCl2.6H2O 0.008 g

H3BO3 0.008 g Adjust pH to 7 with HCl and add water to a final volume of 1 l.

 1 M Tris-maleate (pH 9) Tris 12.1 g Water to 100 ml Dissolve the Tris in 80 ml water and adjust pH to 9 with concentrated maleic acid. Finally make up to a final volume of 100 ml.

 20 mM Tris-maleate (pH 9) Town 1 M Tris-maleate (pH 9) 20 ml Water to 1 l Cape A.2.17. Solutions for performing western blot analysis of  Blocking solution Instant non-fat dried milk 1 g 10x PBS 10 ml Water to 100 ml Do notUniversity autoclave

 Developing substrate solution Nitroblue tetrazolium 75 mg 5-Bromo-4-chloro-3-indolyl phosphate 50 mg 1M Tris-HCl (pH 9.2) (Appendix A.2.1) 1 ml

1M MgCl2 50 ul 1M NaCl 1 ml Water to 100 ml Prepare fresh and do not autoclave 156

 1 M MgCl2

MgCl2.6H2O 203 g Water to 1 l

 1 M NaCl NaCl 58.4 g Water to 1 l

 10 x Phosphate buffered saline (PBS) NaCl 87 g

Na2PO4 22.5 g

KH2PO4 2 g Water to 1 l Town Dissolve NaCl, Na2PO4 and KH2PO4 in 900 ml water. Adjust pH to 7.4 and make up to 1 l with water. Cape A.2.18. Solution for electroblotting of proteins onto a nitrocellulose membrane of  Blotting buffer Tris 6 g Glycine 28.8 g Methanol 200 ml WaterUniversity to 2 l Do not autoclave

A.2.19. Solutions for precipitating antibodies with PEG

 1 x PBS 10 x PBS 100 ml Water to 1 l

157

 Borate buffered saline Boric acid 2.16 g NaCl 2.19 g NaOH 0.7 g 37 % HCl 620 ul Water to 1 l

Dissolve NaCl, NaOH and HCl in 900 ml water. Adjust pH to 8.6 with NaOH and make up to1 l with water.

A.2.20. Solutions for ‘antibiotic cocktail’ for axenic treatment of seaweed

 Penicillin G (Sigma) (100 mg/ml) Dissolve 1 g in 10 ml water. Filter sterilize and store aliquots at 4˚C. Dilute 1:200 into SSW for final concentration of 500Town ug/ml.

 Streptomycin (Roche) (100 mg/ml) Dissolve 2 g in 20 ml water. Filter sterilizeCape and store aliquots at 4˚C. Dilute 1:200 into SSW for final concentration of 500 ug/ml. of  Kanamycin (Sigma) (100 mg/ml) Dissolve 1 g in 10 ml water. Filter sterilize and store aliquots at 4˚C. Dilute 1:200 into SSW for final concentration of 500 ug/ml.

 NalidixicUniversity acid (Sigma) (0.1 mg/ml) Dissolve 1 g in 10 ml water. Filter sterilize and store aliquots at 4˚C. Dilute 1:200 into SSW for final concentration of 0.5 ug/ml.

 Cefotaxine (Claforan®, Roussel) (250 mg/ml) Dissolve 5 g in 20 ml water. Filter sterilize and store aliquots at -20˚C. Dilute 1:1000 into SSW for final concentration of 2.5 mg/ml.

158

 ‘Antibiotic cocktail’ Combine the above antibiotics in their stipulated dilutions into autoclaved seawater (SSW).

A.2.21. Solutions for electron microscopy

 Base buffer (bb) 10x PBS (Appendix A.2.17) 10 ml NaCl 2.34 g Water to 100 ml

 2.5% glutaraldehyde 25% glutaraldehyde (Sigma) 1 ml 10x bb 1 mlTown Sterile water to 10 ml Do not autoclave and prepare fresh. Cape  1% osmium tetroxide osmium tetroxide (Sigma) of 0.1 g Sterile water to 10 ml Do not autoclave

 PBS / 1% BSA (PBS BSA) 10x PBSUniversity (Appendix A.2.17) 1 ml BSA (Roche) 0.1 g Sterile water to 10 ml Filter sterilize and prepare fresh.

 PBS containing glycine (neutralizing buffer) 10x PBS (Appendix A.2.17) 1 ml glycine (Saarchem) 15 mg Sterile water 10 ml Filter sterilize and prepare fresh. 159

 PBS BSA containing 0.1% Tween 20 (v/v) 10x PBS (Appendix A.2.17) 1 ml BSA 0.1 g Tween 20 (Saarchem) 10 ul Sterile water to 10 ml Filter sterilize PBS / BSA (8 ml), add Tween and adjust volume to 10 ml. Prepare fresh.

 1% glutaraldehyde 25% glutaraldehyde 100 ul 10x PBS (Appendix A.2.17) 250 ul Sterile water to 2.5 ml Do not autoclave and prepare fresh. Town

 2% uranyl acetate (pH 5) solution uranyl acetate (Sigma) Cape 5 g 100% Methanol (BDH) 25 ml Filter sterilize and store at 4 ˚C.of

 Reynolds lead citrate Lead citrate (Sigma) 1.33 g Na citrate (Saarchem) 1.76 g SterileUniversity water 30 ml 1 M NaOH 8 ml Sterile water to 50 ml Dissolve lead citrate and sodium citrate in 30 ml water and shake for 1 min. Allow solution to stand for 30 min before adding NaOH. Make up to 50 ml. 160

8. APPENDIX B

STANDARD METHODS

B. CONTENTS

B.1. Large scale preparation of bacterial genomic DNA ...... 161 B.2. Agarose gel electrophoresis ...... 161 B.3. Ligations ...... 162 B.3.1. Intramolecular ligations ...... 162 B.3.2. Intermolecular ligations ...... 162 B.4. Transformation of E. coli ...... 163 B.4.1. Preparation of competent E. coli cells ...... 163 B.4.2. Transformation of competent cells ...... Town 163 B.5. Restriction endonuclease digestions ...... 163 B.6. Transfer of DNA from agarose gel onto nitrocellulose membrane ...... 164 B.7. Heinekoff shortenings ...... 164 B.8. Quantitation of DNA samples ...... Cape ...... 165 B.8.1. Spectrophotometric quantitationof of DNA ...... 165 B.8.2. Ethidium bromide fluorescent quantitation of DNA ...... 166 B.9. Ammonium acetate precipitation of DNA ...... 166 B.10. Bradford assay for protein quantitation ...... 167 B.11. Ferrecyanide assay for reducing sugars ...... 167 B.12. Preparing and electrophoresis of denaturing SDS-PAGE gels ...... 168 B.13. Zymogram detectionUniversity of agarase(s)...... 169 B.14. Protein precipitation with Trichloroacetic acid (TCA) ...... 169 B.15. Thin-Layer chromatography analysis (TLC) ...... 169 B.16. Electroblotting of proteins onto a nitrocellulose membrane ...... 170 B.17. Precipitating antibodies with PEG ...... 170

161

B.1. Large scale preparation of bacterial genomic DNA (Ausubel et al., 1989 unit 2.4)

 Grow a 100 ml culture of the bacterial strain O/N.  Pellet the cells for 10 min at 4 000 rpm and discard the supernatant.  Resuspend the cells in 9.5 ml TE buffer (Appendix A.2.1).  Add 0.5 ml 10% SDS (Appendix A.2.5) and 50 ul of 20 mg / ml proteinase K (Appendix A.2.5), mix and incubate 1 h at 37 ˚C.  Add 1.8 ml of 5 M NaCl (Appendix A.2.5) and mix thoroughly.  Add 1.5 ml CTAB / NaCl (Appendix A.2.5) solution and mix thoroughly.  Incubate for 20 min at 65˚C.  Extract with an equel volume of chloroform / isoamyl alcohol (Appendix A.2.5).  Centrifuge at 6 000 rpm for 10 min at room temperature and transfer aqueous phase to a clean tube. Town  Precipitate DNA by adding 0.6 volumes isopropanol (Merck).  Centrifuge at 10 000 rpm for 10 min.  Wash pellet with 1 ml 70 % EtOH (Appendix A.2.1) Resuspend DNA in 1 ml TE buffer with 10 ul RNAse (Appendix A.2.5)Cape  Measure the DNA concentration on aof spectrophotometer (Appendix B.8)

B.2. Agarose gel electrophoresis (Ausubel et al., 1989 unit 2.4)

 Melt agarose in 1x TAE (Appendix A.2.6) by heating in microwave and swirling to ensure even mixing.University Agarose concentrations can vary from 1 % for separating plasmid DNA fragments to 0.8 % for separating larger chromosomally restriction enzyme digested DNA fragments.  Add Ethidium bromide solution (Appendix A.2.6) to a final concentration of 0.5 ug / ml.  Cool the melted agarose to 55 ˚ C before pouring onto the gel platform.  Seal the gel-casting platform with masking tape if it is open at the ends and pour the melted agarose and insert the gel comb, ensuring that no bubbles are trapped underneath the comb. 162

 After the gel has hardened, remove the tape from the casting platform and withdraw the gel comb.  Place the gel-casting platform containing the set gel in an electrophoresis tank and add sufficient 1 x TAE to cover the gel.  Load DNA samples into the wells of the gel.  Attach leads so that DNA migrates into the gel toward the anode and electrophorese the gel at 1 to 10 V / cm until the dye in the loading buffer reaches the end of the gel.

B.3. Ligations (Coyne et al., 1996)

B.3.1. Intramolecular ligations

 Use approximately 1 pmol of DNA in order to re-circularize plasmid DNA.  Add 2 ul 10x ligation buffer (Roche) and adjust volume toTown 18 ul with sterile water.  Add 2 U of T4 ligase (Roche) to a final volume of 20 ul and incubate reaction mix overnight at 15 ˚ C.

B.3.2. Intermolecular ligations Cape of  In order to polymerize two distinct DNA fragments the total DNA concentration (vector plus insert) should not exceed 10 pmol. Use ratios of vector to insert in the order of 1:1 to 1:4 pmol.  To an eppendorf add the vector and insert DNA and add 2 ul of 10 x ligation buffer (Roche).

 Add 2 U of TUniversity4 ligase (Roche) to a final volume of 20 ul. When ligating DNA fragments with cohesive ends incubate reaction mixes overnight at 15 ˚ C. When joining blunt-ended DNA, use 10 x more enzyme and incubate overnight at room temperature.

163

B.4. Transformation of E. coli

B.4.1. Preparation of competent E. coli cells (Taken from Dagert and Ehrlich, 1979)

 Inoculate a single colony of freshly streaked E. coli into a 5 ml LB (Appendix A.1.1) and shake at 37 ˚C for 2.5 h.  Inoculate this starter culture into 100 ml pre-warmed LB and grow at 37 ˚C until the 7 OD600 reaches 0.35 (approximately 3.5 – 4.0 x 10 cells / ml).  Transfer the culture to a GSA tube and centrifuge for 5 min at 5 000 rpm, 4˚C.

 Decant the supernatant and resuspend the cells in 100 ml ice cold 0.1 M MgCl2 (Appendix A.2.7) and leave on ice for 1 min.

 Collect the cells as before and resuspend in 50 ml 0.1 M CaCl2 (Appendix A.2.7) and leave on ice for 2 h.

 Collect the cells as before and resuspend them in 10 ml 0.1Town M CaCl2.  Aliquot 100 ul into 1.5 ml eppendorf tubes and store at -70 ˚C.

B.4.2. Transformation of competent cells Cape (Taken from Dagert and Ehrlich, 1979) of  Add 1 to 50 ng of plasmid DNA to 100 ul of competent cells and leave on ice for 10 min.  Heatshock cells at 42 ˚C for 2 min or 37 ˚C for 5 min.  Add 0.9 ml LB (Appendix A.1.1) and allow expression at 37 ˚C for 30 to 60 min.  Plate 100 ul of cells on LA containing antibiotic selection.  Incubate overnightUniversity on LA plates (Appendix A.1.2) at 37 ˚C.

B.5. Restriction endonuclease digestions (Ausubel et al., 1989 unit 3.1)

 The restriction enzyme buffers which are supplied with their respective enzymes were obtained from Roche and Amersham.  Pipette 0.1 to 4 ug of either plasmid of chromosomal DNA into a clean eppendorf tube.  Add 2 ul restriction enzyme buffer and adjust the volume to 18 ul with sterile water. 164

 Add restriction enzyme nuclease (1 to 5 U / ug DNA) to a final volume of 20 ul.  Incubate the reaction mixture for 1 to 2 h at 37 ˚C for a complete digestion and for 5 to 30 min for a partial digestion.  Stop the reaction by adding 5 ul gel tracking dye (Appendix A.2.8).

B.6. Transfer of DNA from agarose gel onto nitrocellulose membrane

 Soak the agarose gel in 2x volumes 0.25 M HCl (Appendix A.2.9) for 5 min at room temperature.  Rinse the gel in 2x volumes of water.  Saturate 10 sheets (25 x 20 cm) Whatman 3MM paper with 0.4 M NaOH (Appendix A.2.9) and place the sheets on top of an inverted gel-casting tray, which has been placed in a tray covered with Saran wrap.  Add 0.4 M NaOH / 1 M NaCl (Appendix A.2.9) to the trayTown so that the ends of the Whatman paper are submerged.  Invert the gel and place on top of the saturated Whatman paper. Ensure that no air bubbles remain trapped. Cape  Cut Hybond N+ nylon membrane (Amersham) to the size of the gel and wet the membrane in water and place it on gel.of Ensure that all air bubbles are removed.  Cover the edges with Saran wrap and place 3x sheets (20 x 15 cm) Whatman 3MM paper over the membrane, followed by a 10 cm stack of dry absorbent paper towel.  Place a glass plate on top of the towels, followed by 0.2 to 0.4 kg weight and blot overnight.  Mark the wellsUniversity of the gel on the membrane with a pencil and rinse the membrane in 2x SSC (Appendix A.2.9) for 5 min at room temperature.  Air-dry the membrane on dry Whatman paper and store between 2 sheets of Whatman 3 MM sheets at 4˚C

B.7. Heinekoff shortenings

 Precipitate the plasmid DNA with ammonium acetate (Appendix B.9) and resuspend in exonuclease III buffer (Appendix A.2.10). 165

 Add 150 U of exonuclease III and remove 4.5 ul aliquots of DNA at 20 second intervals and transfer to tubes containing 12.5 ul SI nuclease mix / tube (Appendix A.2.10). Incubate for 30 min at room temperature.  Stop the reactions through the addition of 1.75 ul of S1 nuclease stop (Appendix A.2.10) and place at 70 ˚C for 10 min.  Check shortening by removing 2 ul from every 2nd time point and electrophorese on agarose gel as described in Appendix B.2  Fill in overhangs by adding 1.7 ul of klenow mix (Appendix A.2.10) and 1 ul of Klenow (1 U / ul) to each tube. Leave the reactions to proceed for 3 min at room temperature before the addition of 1 ul dNTP’s (0.5 mM)/ tube. Incubate for 5 min at room temperature.  Re-circularize the the shortened DNA fragments by performing blunt end ligations as described in Appendix B.3.1  Transform half of each ligation mix into competent E. coliTown JM109 (Appendix B.4)  Select the resulting transformants on LA (Appendix A.1.2) containing ampicillin (Appendix A.2.1)  Cleave isolated plasmid DNA from the transformantsCape with the restriction enzyme PvuII to identify pBluescript containingof the desired shortened inserts.

B.8. Quantitation of DNA samples (Coyne et al., 1996)

B.8.1. Spectrophotometric quantitation of DNA University  Perform a DNA scan of the DNA solution between 310-220 nm to determine the UV light absorbance of the sample  The absorbance peak at 260 nm allows the calculation of the concentration of the DNA since 1 OD unit at 260 nm is equivalent to 50 ug / ml for double stranded DNA and 40 ug / ml for single stranded DNA.

166

B.8.2. Ethidium bromide fluorescent quantitation of DNA

 Prepare three λ DNA standards with known concentrations: 5 ng / 10 ul, 10 ng / ul and 20 ng / 10 ul.  Load 10 ul from each standard with 2.5 ul gel tracking dye (Appendix A.2.8) into the wells of a 1 % TAE agarose gel (Appendix B.2).  Prepare several dilutions of DNA sample of unknown concentration in 10 ul and add 2.5 ul gel tracking dye and load next to the standards on the agarose gel.  Electrophorese the samples at 100 V for 5 min and visualize the DNA bands using a 254 nm UV transilluminator.  Determine the concentration of the DNA sample by comparing the intensity of the DNA band to that of the standards. If you load 10 ul of a 1/10 dilution of the DNA sample, which corresponds to an intensity equivalent to that of the 10 ng standard, the DNA sample will have a concentration of 10 ng / ul. Town

B.9. Ammonium acetate precipitation of DNA (Coyne et al., 1996) Cape  Add half the volume of a 7.5 M ammoniumof acetate, pH 7.5 (Appendix A.2.11) to the DNA suspension and incubate at room temperature for 15 min.  Centrifuge at 14 000 rpm for 15 min and transfer the supernatant to a clean eppendorf tube.  Add 2.5 x volumes 100 % EtOH.  Incubate at -20University ˚C for 30 min.  Centrifuge at 14 000 rpm for 30 min at room temperature.  Wash the DNA pellet with 70 % EtOH (Appendix A.2.1).  Resuspend DNA in 10 ul of TE buffer (Appendix A.2.1).  Determine the DNA concentration via the Ethidium bromide fluorescent quantitation method (Appendix B.8.2)

167

B.10. Bradford assay for protein quantitation (Ausubel et al., 1989 unit 10.1)

 Aliquot (in duplicate) the following amounts of BSA (Appendix A.2.12) and 0.15 M NaCl (Appendix A.2.12) into eppendorf tubes. Tube BSA NaCl 1 2.5 ul (2.5 ug / ml) 97.5 ul 2 5 ul (5 ug / ml) 95 ul 3 10 ul (10 ug / ml) 90 ul 4 15 ul (15 ug / ml) 85 ul 5 20 ul (20 ug / ml) 80 ul  Add 100 ul of protein sample with unknown concentration (in duplicate) to an eppendorf tube.  Add 1 ml of Coomassie Brilliant Blue (Appendix A.2.12) to the standard and sample tubes.  Vortex for 5 seconds and allows the tubes to stand at roomTown temperature for 5 minutes.  Determine the OD595 of all the samples and plot a standard curve of OD595 versus protein concentration, using the standards. Use the curve to determine the protein concentration of the sample. Cape B.11. Ferricyanide assay for reducing sugarsof (Park and Johnson, 1949)

 Add (in duplicate) 100 ul culture supernatant to 100 ul freshly-prepared 1% agarose substrate (Appendix A.2.4) and 200 ul 20 mM PIPES (piperazine-N-N’-bis(2- ethanesulfonic acid)) (Appendix A.2.4) solutions.  Incubate at 37University ˚C for 30 minutes.  Terminate the reaction by adding 200 ul of Stop reagent (Appendix A.2.4).  Add 300 ul Colour reagent (Appendix A.2.4) and incubate the tubes in a boiling water bath for 2.5 min.  Cool the tubes at 22 ˚C and determine the absorbance at 420 nm.  Agarolytic enzyme activity (agarolytic units: U) can be expressed as ug galactose produced per ml per hour or per minute. The concentration of galactose must be determined from a standard curve of absorbance at 420 nm versus galactose concentration (ug galactose/ml). 168

B.12. Preparing and electrophoresis of denaturing SDS-PAGE gels (Ausubel et al., 1989 unit 10.2)

 Combine the following reagents for a 12 % separating gel mix in a glass beaker. 40 % acrylogel 4 ml 4 x separating gel buffer (Appendix A.2.13) 2.5 ml Sterile water to 9.95 ml 10 % ammonium persulfate (Appendix A.2.13) 50 ul TEMED 15 ul  Pour the separating gel mix into the assembled gel plates, leaving sufficient space at the top for the stacking gel.  Gently overlay the gel mix with 0.1 % SDS.  After polymerization, remove the overlay and rinse the surfaceTown of the separating gel to remove unpolymerized acrylamide.  Prepare the 5 % stacking gel as follows: 40 % acrylogel Cape 625 ul 4 x stacking gel buffer (Appendixof A.2.13) 1.25 ml Sterile water to 4.97 ml 10 % ammonium persulfate (Appendix A.2.13) 25 ul TEMED 15 ul  Pour the stacking gel mix and insert the comb immediately. After the stacking gel has polymerized, remove the comb and rinse the wells to remove any unpolymerized acrylamide. University  Place the assembled gel into the electrophoresis apparatus and fill the tank with SDS- PAGE running buffer (Appendix A.2.13).  Prepare the protein samples by adding 5 ul of SDS-PAGE sample buffer (Appendix.2.13) to 5-15 ul of protein sample. Denature the protein samples by boiling for 3 min at 96 ˚C.  Load samples into the bottom of the wells and run the gel at constant current of 15 mA in the stacking gel and 30 mA in the separating gel. 169

 After electrophoresis, visualize the protein bands in the gel by staining with Coomassie blue dye (Appendix A.2.13) for 15 min at 37 ˚C.  Destain the gel in destaining solution (Appendix A.2.13)

B.13. Zymogram detection of agarase(s)

 Perform SDS-PAGE (Appendix B.12) with the following modification.  Incorporate a final concentration of 0.1% agarose into the separating gel matrix.  Prepare samples but do not denature the protein by boiling at 96 ˚C. Perform electrophoresis.  Soak the gel in 10 mM phosphate buffer (pH 7) at 22C for 3 h and replace the buffer hourly.  Incubate O/N in 10 mM phosphate buffer (pH 7) at 37C  Visualize the zones of hydrolysis by staining the gel withTown Gran’s Iodine (Appendix A.2.14)

B.14. Protein precipitation with TrichloroaceticCape acid (TCA) of  Add 4 ml of a 50% solution of TCA (Appendix A.2.3) to 100 ml supernatant  Incubate on ice for 1 h  Centrifuge at 10K rpm for 15 min at 4C  Wash pellets with 70% Ethanol  Centrifuge at 10K rpm for 15 min at 4C  Resuspend inUniversity buffer O/N at 4C

B.15. Thin-Layer chromatography analysis (TLC)

 Blot the reaction mixes onto a Silica gel 60 aluminium foil (20 cm x 20 cm) (Merck) in aliquots of 1 ul, until 3 ul is loaded per sample. Allow sample to dry after each 1 ul blot. 170

 Place the Silica gel in an upright position in a tank filled with a 100 ml solvent of n- butanol : acetic acid : water (Appendix A.2.15) and seal the tank. The reaction mixes that have been blotted on must be at the bottom of the gel.  Allow to develop until the solvent is 1 – 3 cm from the top of the gel.  Visualise the resulting saccharides by spraying with naphthoresorcinol reagent (Appendix A.2.15).

B.16. Electroblotting of proteins onto a nitrocellulose membrane

 Remove the SDS-PAGE gel from the glass plate and soak the gel in blotting buffer (Appendix A.2.18) for 1h.  Pre-wet the nitrocellulose membrane in blotting buffer (Appendix A.2.18) and place gel on membrane while ensuring that no air bubbles remain trapped.  Cut four sheets of Whatman 3MM filter paper (10 x 5 cm)Town and soak in blotting buffer.  Sandwich the membrane and gel between the filter paper.  Transfer the proteins to the nitrocellulose membrane using the Hoeffer electroblotting apparatus Cape of B.17. Precipitating antibodies with PEG

 Mix 500 ul of antibodies with 1 ml of borate-buffered saline (Appendix A.2.19).  Add 0.21 g of crushed PEG and gently dissolve by inversion.  Centrifuge theUniversity mixture at 14 000 rpm for 10 min.  Discard the supernatant and dissolve the pellet in 500 ul PBS (Appendix A.2.19).  Add 0.07 g of crushed PEG and gently dissolve by inversion.  Centrifuge the mixture at 14 000 rpm for 10 min.  Discard the supernatant and dissolve the pellet in 250 ul PBS (Appendix A.2.19). 171

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