Genetic Engineering-Based Approach to Explore the Bioactive Potential of Pseudoalteromonas Rubra S4059, a Prodigiosin-Producing Marine Bacterium

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Genetic engineering-based approach to explore the bioactive potential of Pseudoalteromonas rubra S4059, a prodigiosin-producing marine bacterium

Wang, Xiyan

Publication date: 2021

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Citation (APA): Wang, X. (2021). Genetic engineering-based approach to explore the bioactive potential of Pseudoalteromonas rubra S4059, a prodigiosin-producing marine bacterium. DTU Bioengineering.

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Genetic engineering-based approach to explore the bioactive potential of Pseudoalteromonas rubra S4059, a prodigiosin-producing marine bacterium

PhD thesis Xiyan Wang January 2021

PREFACE

This PhD study was conducted at the Department of Biotechnology and Biomedicine (DTU Bioengineering), The Technical University of Denmark from November 2017 to January 2021 under supervision of Professor Lone Gram and Post-doc Shengda Zhang.

The PhD was supported by the Chinese Scholarship Council (CSC scholarship No. 201706170066) and a stipend from the Technical University of Denmark.

The work resulted in three research articles/manuscripts which are included in this thesis.

January 2021

ABSTRACT

Antimicrobial resistance is a huge threat to global public health and we urgently need to discover novel antibiotics to treat infectious diseases. Most antibiotics are derived from natural products of microorganisms and they are typically encoded by biosynthetic gene clusters (BGCs) which are collections of the genes encoding biosynthetic enzymes that are responsible for natural compounds production. Mining of bacterial genomes has indicated that there is a large number of BGCs for which we do not yet know the chemistry; so a potentially untapped source of bioactive compounds in microorganisms. More recently, marine bacteria, for instance from the genus Pseudoalteromonas, have received attention due to their production of bioactive compounds, both enzymes and chemicals with drug activities with potential for pharmaceutical and industrial applications. The purpose of this project is to explore the bioactive potential of a pigmented bacterium Pseudoalteromonas rubra S4059 with genetic manipulation and cultivation-based approaches.

Genome mining on pigmented Pseudoalteromonas has shown that these species harbor large amounts of orphan BGCs and have a large potential for novel compounds production, however, these orphan BGCs are considered “silent” under laboratory conditions. Mimicking the natural environment using ecological niche associated nutrients such as chitin, can induce secondary metabolite production in some Vibrios and also activate unknown BGCs. Chitin degradation is a trait for pigmented Pseudoalteromonas species and thus, we hypothesized that there also in Pseudoalteromonas could a link between chitin degradation and bacterial secondary metabolites production. Using proteomic, metabolomic and genetic approaches, we found that chitin does not significantly influence secondary metabolites production in S4059 as compared to other carbon sources, however, its degradation products, chitin monomer can induce the overall high production of secondary metabolites in S4059. Surprisingly, we found that growth on natural chitin surface leads to spontaneous mutation of S4059 that abolishes pigment production. The non-pigmented mutants appeared to have a fitness advantage as compared to the wildtype as the degree of autolysis was lower. Thus, we here for the first time report that growth on chitin surface can provide genetic diversity driving bacterial evolution of pigmented Pseudoalteromonas. Furthermore, we used the developed genetic manipulation approach to verify a prodigiosin BGC and found that a blue pigment, dipyrrolyldipyrromethene prodigiosin, was produced by the pigC mutant of P. rubra

S4059. In conclusion, this work has contributed by developing a genetic approach to manipulate S4059 and thus has extended the genetic manipulation tools in this genus. Also, this work provides a novel sight on ecological niche driving pigmented Pseudoalteromonas evolution.

iii

RESUMÈ

Antibiotika-resistens er en stor trussel mod den globale folkesundhed, og vi skal hurtigst muligt finde nye antibiotika til behandling af smitsomme sygdomme. De fleste antibiotika er naturstoffer fra mikroorganismer, og deres syntese er ofte betinget af en række enzymer, der kodes for i såkaldte biosyntetiske genklynger (BGCer). Analyser af fuldgenomer fra bakterier har vist, at der er et stort antal BGCer, hvor vi endnu ikke véd hvilke kemiske stoffer, de resulterer i. Disse BGCer og deres kemiske fobindelser kan være en ny, endnu uudnyttet kilde, til bioaktive forbindelser fra mikroorganismer. Marine bakterier, for eksempel fra slægten Pseudoalteromonas, er i stand til a producere mange bioaktive forbindelser, både enzymer og kemikalier med potentiale for farmaceutiske og industrielle anvendelser. Formålet med dette projekt er at undersøge det bioaktive potentiale hos en pigmenteret bakterie Pseudoalteromonas rubra S4059 og udvikle genetiske teknikker, der kan anvendes til at udfolde dens bioaktive potentiale.

Analyse af genomer fra pigmenterede Pseudoalteromonas har vist, at disse arter har mange BGCer hvor den tilsvarende kemiske metabolit stadig ikke er opdaget. Disse arter har således et stort potentiale for produktion af nye bioaktive forbindelser, men de føromtalte BGCer antages ikke at være udtrykt, når bakterierne dyrkes under normale laboratorie-betingelser. Ved at efterligne det naturlige miljø, som bakterierne lever, fx ved ko-kultivering eller brug af naturlige carbon-kilder, kan man inducere produktionen af bioaktive stoffer (sekundære metabolitter) i nogle Vibrio stammer og også aktivere udtrykket af ukendte BGCer. Pigmenterede Pseudoalteromonas-arter er typisk stærke chitin- nedbrydere, og vi har derfor antaget , at der også i Pseudoalteromonas kunne være en forbindelse mellem kitin-nedbrydning og bakteriel sekundær metabolitproduktion. Ved hjælp af proteom- metabolom- og genomiske og genetiske analyse fandt vi, at kitin ikke signifikant påvirker produktionen af sekundære metabolitter i S4059 sammenlignet med andre kulstofkilder. Derimod kan kitinnedbrydningsprodukter, monomere, inducere den samlede høje produktion af sekundære metabolitter i S4059. Overraskende fandt vi, at vækst på naturlig kitinoverflade fører til spontan mutation af S4059, og fremvækst af mutanter, der ikke danner pigment. De ikke-pigmenterede mutanter synes at have en fitnessfordel sammenlignet med vildtypen, da graden af autolyse i stationær fase var lavere. Dette der den første beskrivelse af at vækst på kitin-overflader kan drive evolution af pigmenterede Pseudoalteromonas. Desuden brugte vi de udviklede genetiske værktøjer til at verificere en prodigiosin-BGC og fandt, at et blåt pigment, dipyrrolyldipyrromethen-prodigiosin, blev produceret af P. rubra S4059 pigC-mutanten. Afslutningsvis har dette arbejde bidraget ved at udvikle en genetisk tilgang til at manipulere S4059 og har således udvidet de genetiske

iv manipulationsværktøjer i denne slægt. Dette arbejde giver også et nyt syn på hvordan evolution af Pseudoalteromonas kan faciliteres i dens naturlige nicher.

PUBLICATIONS

Paper 1 – X Wang, T Isbrandt, ML Strube, SS Paulsen, MW Nielsen, Y Buijs, EM Schoof, TO Larsen, L Gram and SD Zhang 2021. Chitin degradation machinery and secondary metabolite profiles in the marine bacterium Pseudoalteromonas rubra S4059. Marine Drugs (accepted: 04 February 2021).

Manuscript 2– X Wang, T Isbrandt, EØ Christensen, TO Larsen, SD Zhang and L Gram 2021. Identification and verification of the prodigiosin biosynthetic gene cluster (BGC) in Pseudoalteromonas rubra S4059. Ready for submission.

Manuscript 3 –X Wang, T Isbrandt, TO Larsen, SD Zhang and L Gram 2021. Attachment to and growth on chitin surfaces results in loss of prodigiosin production in a pigmented Pseudoalteromonas rubra S4059. Manuscript in preparation.

Table of Contents

PREFACE ...... i ABSTRACT ...... ii RESUMÈ ...... iv PUBLICATIONS ...... vi 1. INTRODUCTION AND PURPOSE ...... 1 2. BIOPROSPECTING OF MARINE PROTEOBACTERIA ...... 4 2.1. Natural products from marine Proteobacteria...... 5 2.2. Lab-based procedures to explore the potential role of natural bioactive compounds from marine Proteobacteria ...... 9 2.3. Strategies to determine the bioactive potential of marine Proteobacteria ...... 12 3. GENUS PSEUDOALTEROMONAS ...... 16 3.1. Occurrence and ecological significance of Pseudoalteromonas ...... 16 3.2. Bioactive potential of Pseudoalteromonas ...... 19 4. BACTERIAL CHITIN DEGRADATION MACHINERY ...... 23 4.1. Different forms of chitin ...... 23 4.2. Bacterial chitinolytic enzymes ...... 24 4.3. The potential link between chitin degradation and secondary metabolites production ...... 30 5. GENETIC ENGINEERING OF MARINE PROTEOBACTERIA ...... 33 5.1. Natural and artificial transformation in marine Proteobacteria ...... 33 5.2. Conjugation-based method in marine Proteobacteria ...... 36 6. CONCLUSIONS AND FUTURE PERSPECTIVES ...... 41 7. ACKNOWLEDGEMENTS ...... 43 8. REFERENCES ...... 44 9. RESEARCH ARTICLES...... 62

1. INTRODUCTION AND PURPOSE

The rapid development and spread of antimicrobial resistance in pathogenic microorganisms are significant threats to humanity. We are thus in urgent need for novel drugs, including antibiotics. Natural products from both plants and microorganisms are important as antibiotics and other drugs, and the microbially derived drugs have, so far, mainly been isolated from terrestrial microorganisms, especially from the Actinobacteria group [1]. However, the discovery of novel antibiotics and drugs has dramatically declined since the 1960s and no novel class of antibiotics has been introduced over the last two decades [2]. Exploring microbes from new environments is a promising strategy to address this “innovation gap” on novel drug discovery [3].

The marine environment has attracted attention as a yet unexplored drug discovery reservoir that is rich in biologically active substances, both natural products and hydrolytic enzymes [4– 8]. In particular, marine Proteobacteria have appeared as an untapped source of novel natural compounds as revealed by both classical bio-assay guided fractionation and in silico analysis identifying biosynthetic gene clusters (BGCs) [9–11].

The focus of this thesis is the marine genus Pseudoalteromonas that belong to the class Gammaproteobacteria. The genus Pseudoalteromonas is phenotypically and genetically divided into two groups: pigmented and non-pigmented species [12]. Non-pigmented species produce a range of different enzymes, such as alginases and pseudoatlerin; the latter being able to degrade peptidoglycan [2,13]. On the other hand, a series of bioactive compounds with antibiotic, antiviral or antieukaryotic activity is produced by the pigmented species, and the compounds include violacein, indolmycin, prodigiosin, and tambjamines [12,14–16].

During the global research expedition Galathea 3 (www.galathea3.dk), our group has isolated and collected more than 500 marine bacterial strains with antibacterial activity. Approx. one fifth of these belong to the genus Pseudoalteronomas and half of these Pseudoalteronomas are pigmented [14,17]. The pigmented species represent a rich source of novel bioactive compounds [12] and some species dedicate up to 15% of their genomes to BGCs [10]. However, only few of these bioactive compounds have been chemically detected and characterized and it is believed that most of the BGCs are cryptic or silent when cultured

1 under laboratory conditions [18]. The reason why these species contain so high numbers of BGCs is not known.

Mimicking the natural growth conditions is one cultivation strategy to “unsilence” the mentioned BGCs [19,20] and it has been suggested that growing marine bacteria on chitin could be a way of enhancing production of natural products [20]. Chitin is the most abundant polysaccharide in marine environment and can be utilized by marine microorganisms as a carbon source [21] and can induce expression of BGCs in some marine vibrios [20,22]. Due to the insoluble property of chitin, its degradation relies on production and secretion of chitinolytic enzymes. Chitinases, the major chitinolytic enzymes, are glycoside hydrolases (GHs). They are mainly divided into two families: the GH18 family and the GH19 family [23]. The GH18 chitinases are common in bacteria, while the GH19 chitinases are mainly found in plants and fungi [24,25]. In addition, chitin degradation is facilitated by lytic polysaccharide monooxygenases (LPMOs) that are copper-dependent oxidative enzymes, which can degrade crystalline surface of chitin and facilitate access of chitinases [26,27]. LPMOs are reclassified to Auxiliary Activity family (AA) and AA10 family enzymes are mainly found in bacteria [28].

As mentioned, previous work has demonstrated that chitin can enhance production of secondary metabolites in several isolates of Vibrionaceae [20]. Genomic analysis on 157 Pseudoalteronomas sequences has revealed that pigmented Pseudoalteronomas have a highly developed chitin degradation machinery. This co-occurred with a large and untapped biosynthetic potential [10]. Therefore, the long-term purpose of this project is to determine if chitin metabolism can affect production of novel secondary metabolites in marine Pseudoalteromonas and potentially be used as a strategy to induce production of novel biologically active compounds with drug potential. To explore this, a second aim was to manipulate genetically the chitin degradation machinery on Pseudoalteromonas and determine if this affected the secondary metabolite profiles. Thus, the experimental aim was to develop methodology allowing genetic manipulation of pigmented Pseudoalteromonas. As mentioned, pigmented Pseudoalteromonas represent a rich source of novel bioactive compounds and contain a strong chitinolytic machinery. All chitinolytic Pseudoalteromonas (all pigmented and few non-pigmented) contain at least one GH19 chitinase and one conserved chitin degradation cluster (CDC) in their genome [10]. This PhD project focused on a pigmented Pseudoalteromonas rubra strain S4059, which is a very efficient chitin degrader

2 containing GH19 chitinases and two conserved chitin degradation clusters (LPMOs with two different types of GH18 chitinase). The bacterium has a high potential for secondary metabolites production with 19 BGCs (as described in article 1). The GH19 chitinase gene and two LPMOs genes were chosen as the first targets to explore the potential link between chitin degradation and secondary metabolites production. Protocols for generating in-frame deletion mutants were developed and the mutants were compared to wild type S4059 with respect to chitin degradation, growth kinetics on different carbon sources, biofilm formation on shrimp shells and secondary metabolome (as described in article 1). In order to compare the difference of chitinolytic enzymes between marine bacteria and other organisms, bioinformatics tools were used to analysis the phylogeny of chitinolytic enzymes. Subsequently, a putative prodigiosin BGC was identified by a combination of in silico analysis and genetic manipulation approach as described in paper 2. The last article (Paper 3) introduces a novel discovery that growth on chitin surface can induce spontaneous mutation in several pigmented Pseudoalteromons species.

In this PhD thesis, Chapter 2 gives an overview of marine Proteobacteria with potential for secondary metabolite production and the potential ecological role of natural products from marine Proteobacteria according to lab-based approaches is discussed. This includes approaches to access the bioactive potential of marine Proteobacteria. Chapter 3 describes the occurrence and ecological importance of Pseudoalteromonas and pharmaceutical and industrial applications of Pseudoalteromonas. Chapter 4 dedicates to bacterial chitin degradation machinery and the possible link between chitin degradation and secondary metabolites production. Chapter 5 contains an overview of genetic manipulation approaches used in marine bacteria.

2. BIOPROSPECTING OF MARINE PROTEOBACTERIA

Since the first antibiotic, penicillin, was discovered from a filamentous fungus Penicilum rubens [29], natural compounds have attracted significant attention as clinic drugs. Fungi and soil Actinobacteria have been the major sources of bioactive compounds during the past century [30,31]. However, there has been an innovation gap for novel antibiotic discovery starting from the 1960s and lasting for around 40 years [2,3].

One strategy for finding novel antibiotics is to explore new organisms in new environments, and the marine environment is becoming such an option. The oceans cover 71% of the Earth’s surface and contain an enormous ecological and biological diversity. For a long time, it was believed that microorganisms barely could survive in the harsh chemical and physical marine environment, with changes in temperature, salinity, pH, predator-prey interaction and poor nutrients [32]. However, we now know that the oceans harbor an enormous microbiological abundance and diversity, and these organisms are also considered a promising source for novel microbial and chemical diversity [33]. Over the past few years, marine microorganisms have appeared as significant sources of novel compounds as compared to marine eukaryotes (Figure 1) [34].

Figure 1. Sources of new compounds from the marine environment over the period 2012-2017. This figure was cited from [34].

Marine bacteria occur in all marine habitats, and marine waters contain on average approximately 104 to 106 bacterial cells per milliliter [17,35,36]. Marine Proteobacteria are the dominant microbial members of the marine microbial communities in the seawater (Figure 2) [37–42]. Furthermore, marine Proteobacteria are emerging as the potential source on novel bioactive compounds discovery [43,44].

Figure.2. Taxonomic classification of microbial reads retrieved from different environment water masses into Phylum levels using the EzTaxon database and this figure was cited from [40].

2.1. Natural products from marine Proteobacteria

Most bioactive compounds have been isolated from terrestrial actinobacteria and in particular the genus Streptomyces [45]. However, Actinobacteria have also been isolated from aquatic environments [46,47], and several natural compounds with multiple activities have been isolated from marine actinobacteria, including bonactin with antimicrobial activity [48], sesquiterpene with anticancer activity [49], salinamides with anti-inflammatory activity [50], salinosporamide A with anticancer activity [51,52] and trioxacarcins with antimalarial activity [53].

As mentioned, Gram-positive bacteria, especially the Actinobacteria, have been considered as the most potent producers of natural compounds as compared to Gram-negative bacteria [4]. More recently, however, several natural compounds have been isolated and identified from marine Gram-negative Proteobacteria, such as the genera Tistrella [54], the genera Thalassospira [55], the Roseobacter group [56], the genera Pseudoalteromonas [10], and the family Vibrionaceae [43].

Alphaproteobacteria. A few bioactive natural compounds from marine alphaproteobacterial have been identified and investigated. Some of the compounds have shown significant potential as clinical drugs. For instance, aplidine (Figure 3) that belongs to the didemnin family was produced by Tistrella sp. This compound was conferred for the treatment of acute lymphoblastic leukemia [57] and was continued in clinical trials as an anticancer agent [58]. Likewise, many compounds from thalassospiramide family are produced by Thalassospira sp and these display neuroprotective properties against human calpain proteases and exhibit promising potential for therapeutic applications. However, the clinical trials on some compounds were stopped due to compound toxicity. The first marine natural compound that entered clinical trials, Didemnin B was originally considered as a product from marine ascidians [59] and in 2011, a putative biosynthetic gene cluster was found and confirmed from the bacterium Tistrella mobilis [54,60]. Didemnin B was an antineoplastic agent (Figure 3), however, the trials were stopped due to toxicity of the compound during phase II trials [57,61]. Furthermore, tropodithietic acid (TDA) (Figure 3) is produced by marine alphaproteobacteria of the genus Phaeobacter [56], Ruegeria [62] and Pseudovibrio [63] has a promising potential on diverse applications. For example, TDA is a broad-spectrum antibiotic and TDA-producing bacteria have been explored as probiotics to protect fish larvae from vibriosis [64]. Also, pure TDA has been tested as a potential anticancer agent [65]. However, TDA has not been reported to start in clinical trials. Although, natural compounds that were produced by the class of marine alphaproteobacterial have been reported as toxins to normal cells during clinic trials, this class remains a large potential to produce natural compounds as clinic drugs.

Figure 3. Chemical structure of natural compounds from alphaproteobacterial.

Gammaproteobacteria. The marine gammaproteobacterial group is an enormous reservoir of natural products, especially in the genus Pseudoalteromonas [66]. Several natural compounds from this class are not unique to the marine environment and have also been isolated from bacteria from the non-marine environment. For example, prodigiosin (Figure 4), a red pigment, was first isolated from a soil bacterium Serratia marcescens [67]. Then the red pigment was detected in a marine bacterium Pseudoalteromonas rubra [68]. The prodigiosin encoding genes are very conserved between these two organisms although not 100% percent identical [69], suggesting that this BGC probably was transferred via horizontal gene transfer (HGT) from an ancestral BGC according to phylogenetic view [70]. Similar findings with the corresponding prodigiosin BGC are observed from other compounds, such as violacein [71,72], and holomycin [43,73] (Figure 4).

It is believed that the environmental factors in the marine environment, such as pressure and presence halides, lead to distinct molecular scaffolds and biosynthetic mechanisms [74,75]. Consequently, a great number of natural products are currently only isolated and identified from marine bacteria, especially marine gammaproteobacteria. This includes

7 bromoalterochromide [76], pentabromopseudilin [77] and thiomarinol [78]. Bromoalterochromide A and A’ are produced by Pseudoalteromonas maricaloris strain KMM 636 [76] and have been investigated for their antibiotic activity against Gram-positive bacteria [79]and cytotoxic activity against urchin sperm and eggs [76]. Another brominated natural product is pentabromopseudilin that was first isolated from Pseudoalteromonas spp and has antibiotic activities [80,81]. Furthermore, thiomarinol and its analogs are produced by Pseudoalteromonas spp and have significant antibiotic activities against a large number of antibiotic resistant bacteria, including Staphylococcus aureus [82]. In short, this class can produce unique marine natural compounds with particular molecular scaffolds and biosynthetic mechanisms, suggesting this class has a remarkable potential for novel drug discovery.

Figure 4. Chemical structures of holomycin, prodigiosin, and violacein.

Deltaproteobacteria. The marine myxobacteria that belong to deltaproteobacteria have long been known to produce natural compounds [83]. However, several factors have hampered the discovery of novel compounds from this group: The bacteria are in general difficult to culture and their metabolites productivity is poor [44].

A few bioactive compounds were isolated from marine myxobacteria. Haliangicin is an antifungal compound and produced by Haliangium ochraceum [84]. Chemical structure analysis suggested that haliangicin is a pure polyketide molecule [85]. Phenylnannolones are polyketide derived molecules produced by Nannocystis exedens [86]. P-glycoprotein, an ABC transporter with broad substrate acceptance mediated drug efflux, can be inhibited by phenylnannolones A. Furthermore, Haliamide is a polyketide-nonribosomal peptide hybrid natural product produced by Haliangium ochraceum and has cytotoxicity against tumor cells [87].

In summary, there are of course more classes in marine Proteobacteria. For instance, Sulfurospirillum barnesii from marine Epsilonproteobacteria was isolated from anoxic sediment and has been reported to produce Barnesin A which has antimicrobial activity. As indicated, quite a number of natural compounds have been detected and isolated from marine Proteobacteria, however, genome mining has revealed that there is still a large potential as indicated by a vast number of BGCs for which the chemistry is not known. For example, 23 BGCs are predicted by antiSMASH in P. luteoviolacea S2607, while only 5 BGCs have been matched with known compounds [10]. These unknown BGCs are orphan or cryptic under laboratory conditions and this indicates marine Proteobacteria are an untapped reservoir for novel compounds discovery [88].

2.2. Lab-based procedures to explore the potential role of natural bioactive compounds from marine Proteobacteria

As described, marine Proteobacteria produce a range of different bioactive compounds. However, the activity is often measured from laboratory grown cultures that reach high cell densities. The cell densities in natural marine environments are typically lower, hence the concentration of most natural compounds is probably lower in natural environments than in laboratory settings. The natural role of these compounds in complex and changing habitats remains largely unknown, however, lab-based studies may provide a sight on the potential role of the natural compounds.

Examples of antagonistic interactions. Given the antibiotic activity of many of the above described compounds, it is tempting to assume that such natural products play a role in competitive interactions. Bacteria-bacteria antagonistic interaction has been widely investigated and a range of evidence indicates that natural compounds can play major roles in antagonism between bacteria. For example, Lemos et al. [89] have reported that an antibiotic-producing strain from the genus Alteromonas, isolated from intertidal seaweeds, had a competitive advantage against non-producers. This finding suggested that the production of antibiotics could play a significant role during bacterial competition [89], although the authors do not document which compound (s) is (are) produced by the antibiotic-producing strain. Further, a Vibrio sp SWAT3 that produces a bioactive compound, andrimid can inhibit the colonization of V. cholerae on marine particles, while the andrimid deficient mutant of SWAT3 did not show inhibitory effect [90]. This finding indicated that bacterial antagonistic interaction affects the structure of marine microbial niches. Another example is the selective inhibition of Gram-negative bacteria by a Pseudoalteromonas strain that produces the antibiotic korormicin. This compound selectively inhibits the sodium- translocating NADH: quinone oxidoreductase in Gram-negative marine bacteria, but is not active against Gram-positive bacteria or non-salt-dependent strains [91]. This finding suggested that some natural compounds have specific antagonizing functions on the targets.

Examples of protection against UV radiation. Ultraviolet radiation is a risk factor for marine bacteria in the upper ocean waters as UV can damage intracellular structures and disrupt enzymatic reactions by inducing the formation of reactive oxygen species [92]. Several marine proteobacteria can produce many molecules with conjugated double bonds that can absorb UV light and thus protect against UV light. For example, prodigiosin, the red pigment mentioned above, has been found in several bacterial species, such as Serratia spp, Pseudoalteromonas rubra and Vibrio species [14,93,94]. Vibrio sp. DSM14379 was reported to produce this red pigment that provides a significant level of protection against UV radiation [95]. The authors demonstrated that the survival rate of wild type DSM14379 was significantly higher than that of the prodigiosin deficient mutant under high UV exposure (324 J/m2) and the survival rate of the mutant could be increased by addition of a culture extract from the wildtype [93]. This finding suggested that prodigiosin could potentially function as UV- protective pigment in DSM14379. Likewise, other pigments have been demonstrated to play

10 a role in UV-protection including melanins from Shewanella colwelliana and Vibrio cholerae [96].

Examples of bacterial communication. As previously described, the concentration of natural compounds probably rarely reaches the concentrations required to exert a killing or growth inhibiting function due to the low cell density in nature. The natural compounds may also be considered as signaling molecules that are involved in bacterial cell to cell communication [97]. Quorum sensing (QS) is a specialized bacterial communication setup that has been reported in certain bacterial species. In many cases, bacteria produce N-acylhomeserine lactones (AHLs) as an auto-inducer that can activate the expression of target genes and bacteria behaviors by binding to its transcriptional regulator proteins. Although AHLs are probably the most common (or most widely studied) intercellular signal molecules in bacteria, some natural products also were reported that can act as auto-inducers. For instance, tropodithietic acid (TDA) is a troplolone antibiotic that as described above is produced by several roseobacters, such as the genera Phaeobacter [98] and Silicibacter [99] can act as an autoinducer. The biosynthetic gene cluster of TDA consists of 12 essential genes, tdaABCDEF, paaIJK, cysI, malY and tdaH in Silicibacter sp. TM1040. Geng et al. have used a genetic approach by transferring a tdaCp::lacZ reporter fusion vector into 12 individual transposon insertion mutants of essential TDA genes. Cross-feeding experiment by placing wild type in close to the mutants results in the expression of tdaC gene, except for tdaA and tdaH. The reasons failed to restore the expression of tdaC in tdaA and tdaH mutant have not been clearly demonstrated, but this finding indicated that TDA can act as an auto-inducer to induce its own synthetic genes [99]. Likewise, TDA in Phaeobacter gallaeciensis also can act as an auto-inducer, however, it requires the presence of the response regulator PagR [98]. Bacterial communications occur not only between cells of the same species but also among different bacterial species. As previously mentioned, the concentrations of natural products may not be high enough to exert a killing or growth inhibiting function. Thus, these small molecules may act as signals to mediate bacterial communication. For instance, antimicrobial andrimid is produced by a marine bacterium Vibrio coralliilyticus. Using andrimid to challenge a bacterium Photobacterium galatheae results in the increasing production of holomycin in this strain. The authors suggested that the mechanism of andrimid-induced holomycin production in P. galatheae is due to bacterial stress response [100]. Bacteria can sense hazardous

11 compounds that were produced by competitors and thus respond with counter offensive, such as the increasing production of their own small molecules [100], altering virulence [101] or social behaviors for example biofilm formation [102].

2.3. Strategies to determine the bioactive potential of marine Proteobacteria

In the past, the main approach for natural compounds discovery has been bioassay-guided screening and fractionation. This approach is typically based on cultivation and subsequent testing of extracts of bacteria and culture media. However there are still large numbers of bacteria that cannot be cultured in the laboratory [103]. With the rapid advancement of sequencing technologies, mining of genome sequences has given a new approach to novel drug discovery. The increasing availability of genomic sequences provides us with a wealth of information to view the bioactive potential in microorganisms. The combination of conventional screening methods and sequencing approaches provides an unprecedented tool to exploit the large potential of novel bioactive compounds as described in the following.

Cultivation-based approaches. As mentioned, many (marine) microorganisms seem to harbor the ability to produce more natural products than that have been chemically detected. Several cultivation-based approaches are used to unlock this potential. Zeeck and colleagues reported the “one strain many compounds” approach (OSMAC) that microorganism has a large potential to produce distinct compounds under specific conditions [19,104] and on the secondary metabolism of a bacterium is often significantly influenced by the carbon and nitrogen sources available. For example, chitin is the most abundant polysaccharide in marine environment and can be utilized as a carbon source by marine microorganisms [105]. Growth on chitin can enhance secondary metabolite production as compared to growth on glucose in several isolates of Vibrionaceae [20]. In addition, different nitrogen sources have remarkable effects on secondary metabolites production. When an anti-MRSA strain, Pseudoalteromonas piscicida, grew with tryptone as a nitrogen source medium, its antimicrobial activity was enhanced as compared to yeast extract and peptone [106]. The authors also tried other nitrogen sources for example casein or meat extract, however, no inhibition activity was observed. Unfortunately, no chemical analysis was performed in this study, and the antibiotic compound (s) remain (s) unknown.

Another cultivation-based strategy is the co-culture of microorganisms. Some strains only produce bioactive compounds during competition or when requiring a defense [107]. Co- cultivation of the bacterium Sphingomonas strain KMK-001 with a fungal strain Aspergillus fumigatus KMC-901 resulted in the production of a novel diketopiperazine disulfide, glionitrin A. This compound cannot be detected in monoculture of either KMK-001 or KMC-901. Glionitrin A displayed a strong activity against a series of microorganisms, even methicillin- resistant Staphylococcus aureus. Interestingly, it also exhibited weak bioactivity against its “partner”, the A. fumigatus was used for co-cultivation. Although the biological origin of this compound remains unknown, it still indicates that co-cultivation is one of several strategies to be used for novel drug discovery.

Recently, many studies have reported that sub-inhibitory concentration (SIC) of antibiotics can modulate the production of secondary metabolites in bacteria [100,108–112]. Thus, a promising strategy to induce the expression of BGCs or even activate orphan BGCs is to add small molecule elicitors in bacterial cultures. Mitova et al. have reported that the presence of tetracycline or bacitracin in (SIC) can not only enhance the production of streptophenazines A-D but also induce the production of novel streptophenazines F-G in Streptomyces spp [109]. However, not all antibiotics can stimulate the production of secondary metabolites in bacteria, and further work is needed to determine which small molecules can serve as elicitors to modulate the production of natural compounds. Seyedsayamdost developed a high- throughput platform using a genetic reporter system for screening small molecule elicitors that can act as inducers of secondary metabolites productions [108]. Using this platform, a range of small molecule elicitors that have the potential to be antibiotics have been discovered mainly from Actinobacteria, such as cebulantin with bioactive activity against Vibrio species from Saccharopolyspora cebuensis [113]. These findings suggested that the presence of SIC of antibiotics in bacteria cultures can not only enhance the production of natural compounds but also activate orphan BGCs.

Genome-guided approaches. Generally, bacterial secondary metabolites are produced by genes encoding for enzymes that are tightly clustered together, so-called biosynthetic gene clusters (BGCs). Although the structure of these natural compounds is quite complex, the presence of conserved parts of the biosynthetic genes in their BGCs makes the predictions possible, especially the BGCs for the synthesis of polyketides (PKs), non-ribosomal peptides

(NRPs) and terpenes. Most NRPs share a common model of synthesis that contains an initiation module, iterative elongation modules and a termination module. Each module consists of a condensation domain, an adenylation domain and a peptidyl carrier protein domain, except that the termination module contains an extra domain, named thioesterase (TE) domain. The synthesis of PKs is structural modularity and consists of several conserved domains, such as acyl transferase domains, ketosynthase domains. Further, biosynthesis of terpenes is also a modular process, however, they were generally known as plant or fungal metabolites [114]. This common modular synthesis of those secondary metabolites can be used for the prediction of BGCs in bacteria genomes. According to the conserved biosynthetic genes, bioinformatics tools that allow identification of BGCs of natural products have been developed and provide an amazing opportunity to mine genome sequences in silico for the prediction of BGCs. The software antiSMASH is a widely used bioinformatics tool that allows us to rapidly identify, annotate and analysis BGCs in bacterial and fungal sequences [115,116]. For example, the Pseudoalteromonas strain used in this PhD project, Pseudoalteromonas rubra S4059, has 19 BGCs of which the chemistry of 15 BGCs is unknown as described in paper 1. There are many other bioinformatics tools that are based on identification and annotation polyketides (PKS) and non-ribosomal peptides (NRPS) domains to predict BGCs. For example, NP searcher is for identifying and annotating PKS and NRPS (http://dna.sherman.lsi.umich.edu/). Similar tools are natural product domain seeker and PKSIIpred [117].

Furthermore, mining metagenomes is a way to investigate DNA from a complex population of organisms and subsequently, DNA can be cloned either total or enriched environment DNA into easily cultivated hosts [118]. More recently, exogenous DNA (eDNA) has been isolated and analyzed directly from environmental samples by Next-Generation Sequencing (NGS) technique [119]. In many cases, the large number of short reads of Illumina sequencing data and the repetitive nature of the BGCs such as NRPS and PKS lead to rarely fully assembled of metagenomic data. This led to link BGCs to products more challenging. However, several studies have reported that they have succeeded in characterization of biosynthetic genes from environmental samples using metagenomic approaches [71,120]. One promising strategy is to assemble genomes from the metagenomes and then mine these metagenome- assembled genomes for BGCs [121,122]. Altogether, this is an effective strategy to access

14 bioactive potential of unculturable bacteria by cloning eDNA into suitable hosts and with the improvement of sequencing technique, mining metagenomic will be easier in the future.

Heterologous expression. In the age of synthetic biology, the approach based on heterologous expression of BGCs is obvious. However, it is also challenging for several reasons: selecting a suitable host and transferring and expressing the often large size of the BGCs. Furthermore, the biosynthesis pathways of BGCs may involve uncharacterized or non- clustered regulatory genes that are not always located at the BGC. These issues make heterologous expression of BGCs challenging [18,72]. Despite the challenges, several studies have demonstrated that BGCs can successfully be cloned into different hosts, such as Escherichia coli, Pseudomonas putida KT2440, Agrobacterium tumefaciens LBA4404 and Streptomyces species[72,123]. For instance, the violacein BGC from a culturable bacterium Pseudoalteromonas luteoviolacea 2ta16 has been cloned and expressed in two proteobacterial hosts, Pseudomonas putida KT2440 and Agrobacterium tumefaciens LBA4404 [72]. The authors demonstrated that the expression of the violacein BGC in these two proteobacterial hosts was much more robust than in the common laboratory strains of E. coli, such as BL21 (DE3), Top10 and DH5α [72]. This finding suggested that even a closed phylogenetic related host may not always be the good candidate for heterologous expression. More genetic manipulation tools will be introduced in chapter 5.

In summary, marine Proteobacteria are the dominant members in marine bacterial community and have a large genetic and chemical potential for natural bioactive compounds production. Several studies have used lab-based approaches to point the potential natural function of these small molecules, however, the natural role of these compounds is still being debating. Although large numbers of compounds have been reported with multiple activities and pharmaceutical potential from marine Proteobacteria, they still harbor a huge amount of “orphan” or “silence” BGCs which are untapped sources for novel compounds discovery. With the development of cultivation and genome sequencing techniques, these provide a light to further explore the unexploited sources of marine Proteobacteria.

3. GENUS PSEUDOALTEROMONAS

Pseudoalteromonas is a genus of Gram-negative bacteria belonging to the Gammaproteobacteria class. They are obligate aerobic and heterotrophic marine bacteria and found ubiquitously in ocean environments. Pseudoalteromonas species require sodium ions for growth and the rod-shaped cells are motile via polar flagella [12,124]. The genus Pseudoalteromonas has lately attracted attention for several reasons. First, this genus is frequently associated with eukaryotic hosts and can therefore be used as model stains for investigation of microbe-host interaction [125]. Second, the genus has a large potential for production of bioactive compounds and hydrolytic enzymes useful for biotechnological applications [126–128]. As mentioned, the genus is phenotypically and genotypically divided into pigmented and non-pigmented species [12]. Pigmented species harbor a high number of biosynthetic gene clusters (BGCs) as compared to non-pigmented ones. Both the groups produce a large diversity of glycosyl hydrolases and pigmented ones contain an elaborate chitin degradation machinery harboring several chitinolytic enzymes [10].

3.1. Occurrence and ecological significance of Pseudoalteromonas

Pseudoalteromonads are one of the most widely occurring bacteria that can be isolated from marine environments [129]. Pseudoalteromonas species have been isolated from multiple marine habitats, including open and coastal sea waters, deep-sea waters and sediments [130– 132] or associated with eukaryotic hosts, such as mussels [133], tunicates [134] and sponges [135]. The abundance of Pseudoalteromonas was 2.6% ( 2× 10 4 cells ml-1) in surface seawater during the Galathea3 expedition [136]. Likewise, Pseudoalteromonas species were found to account for 1.6% of the total abundance among all 11 marine samples from seawater and sea lettuce [137]. The bacterium Pseudoalteromonas rubra S4059 in this PhD study was isolated from seaweed from warm waters [17] and can produce a red pigment prodigiosin with antimicrobial activity [14].

The Pseudoalteromonas species are often colonizers of biotic or abiotic surfaces in marine environment. Surface colonization of this genus results in a range of both deleterious and beneficial effects to hosts. For example, the deleterious effects to hosts are secretion of extracellular enzymes with degradation of algal tissue activities [125,138] and toxin production [139]. On the other hand, members of this genus can provide beneficial effects to

16 their hosts, such as displaying antifouling activity. For example, six Pseudoalteromonas species including P. tunicata, P. rubra, P. citrea, P. aurantia, P. ulvae and P. piscicida, were tested on biofouling assays of inhibiting the settlement of two invertebrates, Hydroides elegans and Balanus amphitrite as well as the settlement of the common algae spores, Ulva lactuca and Polysiphonia sp.[140,141]. The P. tunicata, a dark pigmented marine bacterium, was highly inhibitory against all the tested invertebrate larvae and algal spores and the other five species also showed inhibition activities against different targets. For instance, P. rubra was inhibitory towards Balanus amphitrite and Polysiphonia sp, but not against Hydroides elegans and Ulva lactuca (Figure 5). The different inhibition activities may be due to various functions of the compounds and enzymes that were produced by these Pseudoalteromonas species. Furthermore, during this PhD study, we found that growth and attachment on natural chitin surface of P. rubra S4059 resulted in loss of pigment production and this phenomenon may occur in all chitinolytic pigmented Pseudoalteromonas species as described in paper 3, however, the reason or mechanism of this phenomenon is still unknown. This finding suggested that growth on the ecological niche, natural chitin surface, can provide genetic diversity driving evolution of pigmented Pseudoalteromonas species.

Figure 5. The biofouling assay on inhibition of invertebrate larvae and algal spore settlement by different Pseudoalteromonas species. This data was originally from [141] and the figure was cited from [12].

The pigmented Pseudoalteromonas species produce a range of bioactive compounds, such as prodigiosin produced by P. rubra S4059 as described in paper 1. Genome mining on 157 Pseudoalteromonas strains containing 62 pigmented and 95 non-pigmented demonstrated that the genomes of pigmented strains devoted to 7.6 ± 4.2% to BGCs and that the genomes of non-pigmented strains devoted to 1.1 ± 0.9% [10]. Notably, the genome of pigmented P. rubra S4059 dedicated 15% to BGCs with potential for secondary metabolites production; a level that is equal to the well-known bioactive bacteria, such as the genus Streptomyces and Myxococcus [142].

Figure 6. Phylogenetic tree of 157 Pseudoalteromonas species based on single nucleotide polymorphism (SNP). Square: chitinolytic activity; triangle: containing GH19 chitinase. The colored boxes were marked according to their colors of pigmentation and this figure was cited from [10].

Whilst not rich in BGCs, the non-pigmented species can produce a range of hydrolytic enzymes and this probably provides an advantage in an environment with many complex carbon-sources, such as algal polymers. Also, these enzymes are of industrial interest. For example, non-pigmented Pseudoalteromonas sp. CF6-2 that was isolated from marine sediment [143] that can produce pseudoalterin, a novel protease belonging to peptidase family M23. Pseudoalterin can kill Gram-positive bacteria by degrading peptidoglycan peptide chains of bacterial cell walls [13] and therefore has potential on biomedical applications. Furthermore, non-pigmented Pseudoalteromonas species may tolerate low temperatures by producing a number of cold-adapted enzymes [144,145] and those enzymes have a great potential on industrial biocatalysts by reducing the requirement of the reaction temperature.

3.2. Bioactive potential of Pseudoalteromonas

Due to the high potential for secondary metabolites and enzyme production, this genus is of interest for both pharmaceutical and industrial applications, including as probiotics in aquaculture or as producers of bioactive substances.

Pseudoalteromonas as probiotics in aquaculture. Aquaculture plays a significate role in the world food production, and also provides raw materials for industrial and pharmaceutical applications. To meet human consumption and avoid overfishing of wild population, the global production of fish from aquaculture has increased dramatically during the recent decades. However, this increase comes at a cost to the environment due to the heavy use of antibiotics in aquaculture [146] and alternative disease treatments are required. The World Health Organization has defined probiotics as “live microorganisms which when administered in adequate amounts, confer a health benefit on the host” [147]. Thus, developing and using probiotics in aquaculture is one way towards sustainable aquaculture.

Marine Pseudoalteromonas species have been investigated as probiotic candidates in aquaculture. One major disease challenge is acute hepatopancreatic necrosis disease (AHPND) affecting global shrimp production leading to loss a cost of the economy. Vibrio parahaemolyticus is one of the pathogens that can produce pirAB toxins causing AHPND. Two Pseudoalteromonas strains, CDM8 and CDA22, were isolated from healthy Penaeus vannamei

19 shrimp and both were antagonistic against Vibrio parahaemolyticus [148]. In vivo shrimp experiment using CDM8 or CDA22 as probiotics indicated that the cumulative mortalities of shrimp was significantly reduced and the copy numbers of pirA were decreased as compared to a control group fed commercial feed during a challenge with Vibrio parahaemolyticus [148]. Although the authors did not mention what commercial feed was used, the study indicated that Pseudoalteromonas species have the potential as probiotics in aquaculture. Likewise, Pseudoalteromonas sp. SLP1-MESO that was isolated from the gonads microbiota of the yellowtail Seriola lalandi had high activity against pathogenic bacteria Yersinia ruckeriI that causes enteric red-mouth disease in several species of fish [149]. These findings showed that this strain is a good candidate as a probiotic in aquaculture. Although many Pseudoalteromonas species have shown potential on biotechnological applications as probiotics, we must remind ourselves that some of Pseudoalteromonas species are pathogens and have adverse effects against the fish. For instance, P. piscicida is a known fish pathogen that can cause high mortality of damselfish Amphiprion clarkii eggs [150]. Pseudoalteromonas species have a large potential as probiotics in aquaculture and rarely Pseudoalteromonas species were reported as fish pathogens, but we still need to be prudent to consider their pathogenicity of these species when they were considered to be as probiotics.

In this PhD project, a common fish pathogen Vibrio anguillarum 90-11- 287 [151] was used as a target for bacterial antagonism. As described in paper 2, the filtered supernatant from the culture of a pigmented Pseudoalteromonas rubra S4059 was highly inhibitory against fish pathogenic bacterium Vibrio anguillarum 90-11- 287, suggesting that this strain could have potential as a probiotic in aquaculture if further experimental evidence can be provided that there are no adverse effects against the fish or other eukaryotic organisms.

Bioactive substances from Pseudoalteromonas. Natural bioactive substances can roughly be split into two major groups: low molecular weight substances which are mostly non- proteinaceous compounds and high molecular weight substances which are mainly proteins, including enzymes [12]. Notably, Pseudoalteromonas species produce substances of both types and in both, compounds of pharmaceutical and industrial applications can be found.

Several compounds with antibacterial activity are produced by Pseudoalteromonas. Some of the compounds have a broad spectrum of inhibition and affect both Gram-negative and Gram-positive bacteria. For instance, a hybrid antibiotic of a pseudomonic acid analogue and

20 holothin, thiomarinol, which is produced by Pseudoalteromonas sp. SANK 73390 has strong activity against a range of Gram-negative and Gram-positive bacteria and thromarinol can inhibit isoleucyl-transfer RNA synthetase and RNA synthesis [82]. Similarly, Pseudoalteromonas phenolica O-BC30 can produce anti- methicillin-resistant Staphylococcus aureus (MRSA) compounds, MC21-A and B. The MC21-A compound has broad spectrum antibacterial activity, while MC21-B only can inhibit Gram-positive strains [152,153]. Some Pseudoalteromonas species can produce compounds not only display antibacterial activities but also exhibit other activities. For instance, a yellow pigmented bacterium, Pseudoalteromonas tunicate produces a compound named YP1 with antifungal [154] and antifouling activity [155], which is a tambjamine-like alkaloid [154]. Another compound with structurally related with tambjiamine is prodigiosin, which is a red pigment. Prodigiosin and its analogous are produced by several bacterial species such as Serratia marcescens [67], Pseudoalteromonas rubra S4059 [14], and has pharmaceutical interest due to its antibacterial [156], anti-malarial [157] and anti-cancer [158] activities. In this PhD project (paper 2), a prodigiosin deficient mutant P. rubra S4059 was generated by deleting the pigC gene. The culture extraction of pigmented Pseudoalteromonas rubra S4059 wildtype was inhibitory against Staphylococcus aureus as compared to prodigiosin deficient mutant of S4059, and also the anti-Vibrio activity was reduced in the mutant. Collectively, suggesting that prodigiosin is a major antibacterial compound of S4059.

Figure 7. The chemical extraction from the culture of Pseudoalteromonas rubra S4059 wild type and prodigiosin deficient mutant ΔpigC after 1-day incubation in marine minimal medium containing

21 mannose. Wt: S4059 wild type; ΔpigC: prodigiosin deficient mutant S4059; C: negative control- medium extraction; 1, 2; bio-replicates of each strain.

Extracellular enzymes are essential in bacterial life as they degrade organic macromolecules providing nutrient sources for bacteria or they may serve as antimicrobial agents. For instance, an antimicrobial protein P-153 was isolated from Pseudoalteromonas piscicida and exhibited high antimicrobial activities against pathogenic vibrios and human pathogenic strains, S. epidermidis and Propionibacterium. The author suggested that this strain could be useful in aquaculture as a probiotic bacterium or as an application in human dermatologic diseases [159]. In this PhD thesis, the focus is not on the possible antibacterial activity of enzymes, but focus is on the hydrolytic degradative enzymes, specifically the chitin degradation machinery. This was identified in Pseudoalteromonas rubra S4059 by proteomic analysis as described in paper 1. An efficient chitinolytic enzyme cocktail was found from this strain, suggesting that this strain has a large potential on industrial application. The bacterial chitin degradation capacity will be introduced in the next chapter.

In conclusion, the genus Pseudoalteromonas is commonly occurring in marine environments. This genus contains a large potential on industrial and pharmaceutical applications. Pigmented Pseudoalteromonas species display a large potential on secondary metabolites production and these natural products are the promising candidates as clinic drugs, while non-pigmented ones can produce diverse proteins with industrial applications [12]. Although amounts of chemical compounds have been reported from this genus, these substances are from only a small subset of this genus for example a large amount of “silent” BGCs in this genus as forementioned. Therefore, the natural product diversity of this genus is likely untapped and needs to be further explored deeply.

4. BACTERIAL CHITIN DEGRADATION MACHINERY

Chitin is the most abundant polymer in the marine environment and was first isolated and characterized from the cell wall of mushrooms by Henri Braconnot [160,161]. Chitin has a crucial role in supporting some organisms as the skeleton and protecting the body, such as in insects and shellfish [162,163]. The global production of chitin is about 10 11 tons every year. Since large accumulations are not seen in the oceans, they must be hydrolyzed by marine (micro)organisms. Several microorganisms can use chitin both as a carbon and nitrogen source and it is thus incorporated into the biogeochemical C- and N-cycles [164]. Marine bacteria are believed to be the main mediators on chitin degradation in marine environment [165]. Chitinolytic bacteria can produce a range of chitinolytic enzymes which degrade complex chitin polymers into soluble oligomers and possess complex signal transduction systems which can (1) drive bacteria to the natural chitin surface, (2) attach and colonize on the surface, (3) degrade chitin polymer to oligomers, (4) transport chitin oligomers into cells, (5) utilize and catabolize the transported products [166].

4.1. Different forms of chitin

Chitin is the polymer of β-1,4-linked homopolymer of N-acetylglucosamine (GlcNAc) and the individual sugar units are rotated 180° to each other. Chitin is classified into three different crystalline allomorphs: α-, β- and γ-form. α-chitin is the most abundant allomorph in nature, which has antiparallel chains, while β-chitin has parallel chains that all chains are in the same direction. γ-chitin is the mixture of parallel and antiparallel chains [167] (Figure 8). Most commercial chitin is from crab and shrimp chitin which are the major source of α-chitin, and it also could be found in fungi and most protistans and invertebrate exoskeletons, while all three different crystalline allomorphs have been found in squid [164].

Figure 8. (A) Chemical structure of chitin and (B) three different crystalline allomorphs of chitin: α-, β- and γ-chitin. The figure was cited from [168].

4.2. Bacterial chitinolytic enzymes

Generally, bacterial chitin degradation consists of three steps: 1) breaking down the crystalline chitin into chitin oligomers; 2) hydrolyzing the chitin oligomers into dimers; and 3) cleaving of dimers into monomers. According to the Carbohydrate-Active enZymes database (CAZy), bacterial chitinolytic enzymes are mainly from the glycosyl hydrolases (GH) family 18, 19, and 20 [155]. The GH18 family is classified into four subfamilies: ChiA, ChiB, ChiC and ChiD. These chitinases have a common structure which contains a GH18 catalytic domain, a chitin- binding domain and a fibronectin type III like domain. However, such little information about the differences and properties of these enzymes from the GH18 subfamilies. GH18 chitinases are commonly found in bacteria and have been approved with genetic evidence to be essential enzymes for chitin degradation in several bacterial species, such as CjChi18D in Cellvibrio japonicus [170], ChiA 1 and ChiA 2 in Vibrio cholerae [171], while GH19 chitinases are commonly found in plants and believed to function as a weapon against fungal pathogens

[24,25]. GH18 and GH19 chitinases can catalyze the degradation of chitin polymers, while hexosaminidase from GH20 can break down the chitin dimers into monomers. The GH18 families are characterized by a catalytic region included a (α / β)8 barrel fold with an (α+β) insertion domain [172], while the catalytic domain of GH19 families has an a-helix-rich fold with a deep cleft containing a catalytic site [173]. Furthermore, lytic polysaccharide monooxygenases (LPMOs), which are from the auxiliary activities family 10 (AA10) can facilitate chitin degradation during bacterial chitin degradation process [26].

For a long time, it was thought that only higher plants can produce chitinases from the GH19 family, and that other organisms predominantly synthesized GH18 chitinases [174]. The first bacterial GH19 chitinase was reported in a bacterium Streptomyces griseus, and since then several GH19 chitinase genes have been identified in bacteria [175]. In plants, GH19 chitinases inhibit fungal pathogens and recently, a GH19 chitinase of the pigmented Pseudoalteromonas tunicata CCUG 44952T has been cloned and heterologously expressed in Escherichia coli and found to display both chitin degradation and antifungal activity [176].

Previously, Paulsen et al. have demonstrated that chitin degradation capacity is a significant trait in pigmented Pseudoalteromonas species and rare GH19 chitinases are consistently present in the pigmented species (as shown in Figure 6) [10]. To gain insights into the evolutionary relationship between GH19 chitinases from Pseudoalteromonas and other organisms, firstly, a total of 69 amino acid sequences of GH19 chitinase from for example plants or bacteria were used to construct a phylogenetic tree (Figure 10). According to the phylogenetic tree, these GH19 chitinases fell into three major clades (Figure 10). GH19 chitinases from the marine strains (colored in blue, including Vibrio and Pseudoalteromonas) clustered into a branch in Clade I (Figure 10). This clade also consisted of GH19 chitinases from fungi, viruses, and other aqua environmental or human-pathogenic Gammaproteobacteria such as strains from genera Aeromonas, Salmonella, etc. The marine bacterial cluster may be skewed as it predominantly contains sequences from two genera, Vibrio and Pseudoalteromonas. However, we speculated that GH19 chitinases in Vibrio spp and Pseudoalteromonas are likely in one clade. It is interesting to note that GH19s from pigmented Pseudoalteromonas clustered in a deeper branch together with most of the vibrio strains, while the GH19 of the non-pigmented Pseudoalteromonas (S1727 in Figure 10) fell into a slightly distinct subgroup. Clade II consisted of GH19 chitinases from other bacteria and

25 few plants, while clade III only contained GH19 chitinases from plants. In silico analysis on comparison of GH19 chitinases from different organisms demonstrated that GH19 chitinases in marine bacteria Pseudoalteromonas and Vibrio are different with soil bacteria and plants. Therefore, we could not speculate the function of GH19 chitinases in Pseudoalteromonas species based on soil bacteria and plants and we further need genetic approaches to explore the function of this enzyme in the marine producer.

Figure 10. Phylogenetic tree based on neighbor-joining analyses of amino acid sequences (bootstrap: 1,000) of GH19 chitinases in (marine) bacteria and higher organisms. Clade I: marine bacteria; Clade II: other bacteria; Clade III: plants. Scale bar: 0.1 amino acid substitutions per site.

Generally, chitinolytic bacteria contain several genes that encode chitinolytic enzymes, and this is also the case for P. rubra S4059 which is the main target bacterium in this PhD project (Figure 11). However, those enzymes do not seem to be equally important to bacteria on chitin degradation or they might target different forms of chitin. For instance, the genome of a soil bacterium Cellvibrio japonicus was predicted to encode four GH18 chitinases, one GH19 chitinase, one LPMO from AA10 family [177]. Secretome analysis showed that chitinolytic enzymes in C. japonicus, including GH18 chitinases and LPMOs, were abundant when grown both on α-chitin and β-chitin as compared to on glucose [178]. Interestingly, the expression of GH19 chitinase is highly upregulated when C. japonicus grown on β-chitin as compared to on α-chitin [178]. That could be because the major function of GH19 chitinase in C. japonicus is on β-chitin degradation. Further, the study generated a range of mutants on GH18 chitinases and the results indicated only one GH18 chitinase CjChi18D is essential for chitin degradation in C. japonicus. However, they did not target on GH19 chitinase and we could not refer the function of this enzyme. In this PhD project as described in paper 1, due to the consistent presence of GH19 chitinases in pigmented Pseudoalteromonas, a pigmented bacterium P. rubra S4059 was selected as a model strain to investigate the function of this enzyme. Firstly, proteomic analysis indicated that a rare GH19 chitinase only could be detected when grown on crystalline as compared to on mannose. Further, an in-frame deletion mutant on the GH19 chitinase was generated, however, S4059 wild type and the mutant did not show any phenotypic differences. This indicated that GH19 is not the predominant or essential enzyme on chitin degradation in S4059 under the tested conditions.

Figure 11. Heat map comparison of identified chitinolytic enzymes in the secretome and proteome of Pseudoalteromonas rubra S4059. Blank means that proteins are not detected under this condition. Overlap indicates those proteins present in both chitin and mannose- grown samples. Asterisk means those proteins are significantly upregulated when grown on chitin compared to on mannose. C-WT- cell: cell samples when S4059 grown on chitin; M-WT-cell: cell samples when S4059 grown on mannose; C-WT-sup: supernatant samples when S4059 grown on chitin; M-WT-sup: supernatant samples when S4059 grown on mannose. This figure was from Paper 1.

Hexosaminidases from the GH20 family can cleave intracellular GlcNAc2 from the non- reducing end of the water-soluble chitin oligomers into GlcNAc in the last step of chitin degradation and are often intracellular enzymes [179]. Generally, this step takes place in the cytoplasm or the periplasmic spaces of bacterial cells [166,180]. Hexosaminidases can, in some of bacteria, directly cleave chitin monomers from chitin oligomers and thus function as chitinases [181,182]. In this PhD project as described in paper 1, proteome analysis of P. rubra S4059 showed that hexosaminidases from the GH20 family could be detected in the culture supernatant. This indicated that hexosaminidases from the GH20 family in S4059 are secreted out of the bacterial cells as shown in Figure 11 and highly expressed when grown on crystalline chitin, suggesting that hexosaminidases in S4059 are secreted enzymes and may function as chitinases.

Lytic polysaccharide monooxygenases (LPMOs) are copper-dependent oxidative enzymes, which can degrade recalcitrant polysaccharides and thereby enable utilization of renewable sources for production of fuels and chemicals [183,184]. LPMOs were originally classified in carbohydrate binding module family 33 and now it was re-classified in the auxiliary activity family. According to the CAZy database, LPMOs are classified into four families: AA9, AA10, AA11 and AA13. The AA9 family enzymes are active on cellulose and hemicellulose, and therefore they are common in plants that function on degrading fungal cell walls [185], while the AA10 family was discovered as a chitin-degrading LPMO, however, the Eijsink group showed that LPMOs of the AA10 family could also degrade cellulose [186]. Fungal LPMOs belong to the AA11 family and AA13 family, and AA11 are chitin-specific LPMOs and AA13 are starch-specific LPMOs [187]. Bacterial LPMOs are mainly from the AA10 family and LPMOs were referred in this PhD project are all from this family.

In 2010, Vaaje-Kolstad et al. first reported an enzyme in a bacterium Serratia marcescens, LPMO, which degraded the surface of crystalline chitin by catalyzing the cleavage of glycosidic bonds and allowing the glycoside hydrolases more easily access and degrade the polysaccharide material [26]. After that, several studies have reported that LPMOs are crucial contributors on bacterial chitin degradation process [188,189]. An example is that an LPMO (previously was described as a chitin binding protein CBP21) was found in a soil bacterium Serratia marcescens with strongly promoting effects on chitin degradation. The deletion of CBP21 led to a significant reduction on promoting chitinase activities by measuring the total amount of degradation products (GlcNAc)2.[188,189]. Paulsen et al. have reported that a conserved chitin degradation cluster (CDC) was consistently found in chitinolytic Pseudoalteromonas strains and the CDC contains a ChiC, an LPMO and a ChiA as shown in Figure 12 [10]. In this PhD project, genome mining on CDC in P. rubra S4059 showed that S4059 harbors two CDCs and further we generated a single deletion mutant and a double deletion mutant on lpmo genes in the CDC. However, neither chitin degradation ability nor biofilm formation was influenced by the deletion of lpmo genes in S4059. Considering that commercial chitin has been pre-processed or even semi-degraded, and natural chitin is much more recalcitrant than the commercial one. In this PhD project, we used natural shrimp chitin as a substance to compare chitin degradation ability of S4059 wild type, a GH19 mutant and an LPMOs double mutant. However, we did not find any difference between S4059 wild type and the mutants on chitin degradation ability (Figure 13). It could be either that S4059 harbors an efficient chitinolytic enzyme cocktail and other enzymes may take over the “job” of GH19 and LPMOs in S4059, or that bacterial aggregation could lead to misestimating the numbers of bacteria cells and bacterial colony counting may not be sensitive enough to show the differences between S4059 wild type and the mutants under current conditions.

Figure 12. The chitin degradation cluster consistently present in all chitinolytic Pseudoalteromonas strains and this figure was cited from [10].

4 Log(CFU/mL) 3

0 0 1 2 3 4 5 6 7 8 9 10 Time (days)

Figure 13. Growth and attachment of Pseudoalteromonas rubra S4059 wild type and mutants (∆GH19, ∆∆LPMOs) on Vannamei shrimp shells. Open circle: wild type; open square diamond: ∆GH19; open triangle; ∆∆LPMOs. Black solid lines: samples from liquid surrounding shrimp shells before sonication. Grey solid lines: samples from liquid surrounding shrimp shells after sonication. Black dotted lines: samples from 3% Sigma sea salt without shrimp shells as a control. The points are bio-triplicates and error bars are standard deviation.

4.3. The potential link between chitin degradation and secondary metabolites production

Chitinolytic bacteria are as mentioned of interest due to their potential as antifungal and insecticidal agents in biocontrol [190]. Recently, it has also been reported that chitin can induce the expression of bacterial BGCs resulting in enhanced antimicrobial activities. Streptomycetes are, as described, prolific secondary metabolite producers and also produce a broad range of extracellular hydrolytic enzymes. A soil bacterium Streptomycetes coelicolor A3 (2) was cultured with or without 0.2% colloidal chitin in autoclaved soil. Microarray data indicated that 675 genes were significantly influenced by addition of chitin to the soil culture of S. coelicolor A3(2), including genes involved in chitin catabolic, carbohydrate metabolism, nitrogen and sulfur metabolism and amnio acid biosynthesis or secondary metabolites

30 pathways [191]. Further, they observed that chitin and its monomer enhanced the production of pigment in S. coelicolor A3 (2) as compared to other carbon sources. Similarly, the expression of BGCs and the production of antibiotics were increased in two marine bacteria Vibrio coralliityticus S2052 and Photobacterium galatheae S2753 when grown on chitin as compared to on glucose [20]. These findings suggested that growth on chitin is a special lifestyle for chitinolytic bacteria and that the increase of secondary metabolites production may provide an advantage during chitin colonization. The chitin-dependent influence on secondary metabolites production could be mediated by a transcriptional regulator, such as ChiS in Vibrio cholera [192]. ChiS could be activated when chitin is present in the environment and chitin and its oligosaccharides are likely to serve a regulatory role in Vibrio cholera [192]. Similarly, a transcriptional regulator DasR was found in a soil bacterium Streptomyces coelicolor A3 (2) and showed a similar function with ChiS in Vibrio cholera [191].

Figure 14. Comparison of identified proteins in supernatant (A) and cells (B) of Pseudoalteromonas rubra S4059 when grown on chitin compared to on mannose. Red dots represent upregulated proteins and blue dots represent downregulated proteins. This figure was cited from Paper 1.

In this PhD project, we explored the potential link between chitin degradation and secondary metabolites production in P. rubra S4059. Proteome analysis indicated that a high amount of proteins (Figure 14 and supplement material in paper 1) involved chitin degradation, bacterial chemotaxis, carbon metabolism, amino acid biosynthesis or secondary metabolites pathways were upregulated when S4059 grown on chitin as compared to on mannose. The result of

31 chemical analysis showed that growth in chitin monomer contained medium resulted in overall higher amounts of secondary metabolites production in S4059, however, not in mannose, crystalline and colloidal chitin medium. Although the utilization of chitin in Streptomyces coelicolor A3(2), some Vibrio and Photobacterium species showed the trend to induce the expression of BGCs [22], these studies did not include chitin monomer as a control. Thus, chitin monomer is likely to be the key factor to stimulate secondary metabolites production and chitin oligosaccharides are likely to serve as a regulatory role on transcriptional regulators in these strains and did not directly involve in secondary metabolites productions.

In conclusion, chitin is the most abundant polymer that can be utilized by microorganisms as a carbon source. Chitin degradation machinery is a key trait for pigmented Pseudoalteromonas [10] and pigmented species can produce a range of chitinolytic enzymes. However, chitinolytic enzymes are not equally important for bacterial chitin degradation and GH18 chitinase is likely to be an essential chitinolytic enzyme in bacteria as mentioned previously. Further, chitin can stimulate secondary metabolites production in few chitinolytic bacteria [191,193] and we observed that chitin monomer is likely to be the key factor to induce secondary metabolites production in P. rubra S4059.

5. GENETIC ENGINEERING OF MARINE PROTEOBACTERIA

Marine bacteria represent an untapped reservoir of bioactive potential. As described in previous chapters, many marine bacteria, especially marine Proteobacteria, have the potential for producing a range of natural compounds and enzymes with industrial and pharmaceutical applications. With the advancement of sequencing techniques, genetic manipulations of those marine bacteria have become possible. Genetic manipulation approaches play a significant role on exploring genetic properties or introducing genetic variants with phenotypic changes revealing the function of a particular gene or pathway [194,195]. One application of genetic manipulation is to determine which BGC is responsible for natural compound production in marine bacteria. Subsequently, the identified BGC could be expressed in other appropriate hosts. However, genetic manipulation of marine proteobacteria is challenging due to the lack of efficient and universal approaches. Luckily, several genetic approaches have succeeded in genetically manipulating some marine proteobacteria and in this PhD work, we established a conjugation-based approach allowing genetic manipulation in a pigmented bacterium P. rubra S4059. The aim of this chapter is to summarize the established genetic approaches for genetic manipulation on marine proteobacteria.

5.1. Natural and artificial transformation in marine Proteobacteria

Natural transformation of marine Proteobacteria. In marine Proteobacteria, natural competence has mainly been described in the Vibrionaceae family, particularly in the human pathogen Vibrio cholerae. V. cholera is a chitinolytic bacterium and inhabits in marine environments [196,197]. Meibom et al. have reported that chitin not only serves as nutrient sources in V. cholera but also plays a significant role on inducing exogenous DNA uptake [198]. Subsequently, the competence has been identified in several Vibrio species, such as V. fischeri [199], V. vulnificus [200] V. natriegens [201] and V. parahaemolyticus [202]. Although chitin can induce natural competence in several vibrios, the form of chitin capable of inducing the competence varies among the different species. For instance, the natural competence in V. fischeri can only be induced by the oligosaccharide chitin, while all forms of chitin can induce the competence in V. cholerae, V. parahaemolyticus and V. vulnificus [198,200,203]. Chitin-

33 induced competence has been utilized in the laboratory as a genetic manipulation approach on Vibrio species. For instance, P. Gulig et al. established a rapid and efficient protocol on chitin-based transformation in V. vulnificus and they succeeded in generating several mutants, for example deletion of the motAB genes which are involved in bacterial motility. However, this chitin-based approach is not a universal tool for marine Proteobacteria, and it requires natural competent cells as recipients. Furthermore, there are also some critical issues are that 1) it is a time- consuming approach, 2) need to provide huge amounts of DNA, and 3) bacterial restriction endonucleases, which play significant roles in protecting their DNA from bacteriophage infection, can degrade exogenous DNA leading to decrease the efficiency of natural transformation in Vibrio species [204].

Chemical transformation of marine Proteobacteria. Chemical transformation has been widely used in the laboratory on commercial E. coli strains and have been successfully used on marine Proteobacteria, however with challenges. For instance, Marcus et al. attempted to transfer the plasmid pBR322 into V. cholerae using an osmotic shock protocol and they only succeeded in transferring the plasmid into a deoxyribonuclease (DNase) - negative mutant, however, unsuccessful in the wildtype. The authors suggested that DNase which can degrade DNA was a major barrier for chemical transformation in V. cholerae [205]. Another example is the fast-growing bacterium V. natriegens which was successfully manipulated with both chemical transformation and electroporation [201]. Chemical transformation is an easy procedure to be performed in the laboratory, however, limitations of this approach are that recipients need to be chemical competent cells and the restriction endonucleases in the recipients highly decrease the efficiency of the transformation.

Electroporation of marine Proteobacteria. Electroporation is a well-established and quick method for introducing genetic material and has been used on a broad range of bacteria in the laboratory. Several marine Proteobacteria have been successfully transformed by electroporation, such as Vibrio, Pseudoalteromonas, and roseobacters [206–208]. For instance, a shuttle vector PVv3, which is a derivative from pVN-0126 plasmid, has been constructed and transferred into V. vulnificus for gene expression by electroporation [209]. However, the efficiency of this method is influenced by many factors, such as growth medium and buffer composition, restriction enzyme systems, growth conditions and phases, and the concentration of exogenous DNA.

The composition of growth medium which was used to culture bacterial cells before electroporation can significantly affect the efficiency of the transformation. For instance, most marine Proteobacteria require a high concentration of salts in the growth medium to mimic their natural habitats [210]. However, the presence of salts leads to arcing and impedes the electroporation process. Wang et al. have reported that electroporation is not effective for most of the Pseudoalteromonas strains because their growth is usually salts-dependent [68]. Furthermore, the concentration of salts also influences restriction enzyme systems. An example of this is that endonuclease I enzymes in V. salmonicida and V. cholerae were cloned, expressed in E. coli and purified. Those purified enzymes are more active when salts are present in a high concentration as compared to low concentration [211]. Although the enhancement of enzyme activity is in vitro, it provides an indication that salts can enhance the activity of restriction enzymes and further that may lead to a decrease in the efficiency of electroporation [211].

Delavat et al. developed a rapid and efficient electroporation protocol allowing the transfer of exogenous DNA in V. harveyi and two Pseudoalteromonas strains [212]. The results suggested that bacterial growth conditions and phases are very important for electroporation. For instance, they succeeded in obtaining transformants when using stationary-phase cells of V. harveyi grown at 37 °C, but no transformant was observed when V. harveyi grown at 28°C or using exponentially-phase cells grown at 37 °C [212]. The authors suggested that bacterial cells grown under optimal temperature (37°C in this case) could alter restriction enzyme systems to allow recipients more easily to accept exogenous DNA, as previously reported in Pseudomonas aeruginosa [212]. Generally, the efficiency of electroporation could be increased proportionally by increasing the concentration of exogenous DNA until a saturation level. For example, Piekarski et al. have demonstrated that the numbers of obtained transformants are proportional to the concentration of added plasmid during the electroporation of Caulobacter species [210].

Although electroporation has been successfully applied to several marine Proteobacteria, it is not always successful. Wang et al. failed to transfer the vector pWD2 into Pseudoalteromonas strains by electroporation. That means that the electroporation protocol is highly strain or species- dependent. The authors generated a conjugation based procedure

35 and succeeded in transferring both shuttle vector pWD2-oriT and other suicide plasmids into several Pseudoalteromonas species [68].

5.2. Conjugation-based method in marine Proteobacteria

Bacterial conjugation is mediated by cell-to-cell direct contact between the donor and recipient cells. The bacterial conjugation machinery consists of an oriT sequence and tra genes [213]. The oriT sequence, which is the origin of transfer, has to be provided in cis by vectors. The tra genes encode a relaxase, the type IV coupling protein and the mating pair formation complex [214]. The relaxase can recognize the oriT sequence and cause its cleavage in the donor to produce a single-strand DNA for transformation, and the transported DNA will be ligated in the recipient cells as the conjugated plasmid [214]. The type IV coupling protein is involved in the connection between the relaxase and the mating pair formation complex. The mating pair formation complex functions as a secretion machinery for the vector transformation during conjugation [214].

Bacterial conjugation is a well-established procedure and one of the major advantages of this method as compared to the three previously described methods is that the transformation happens on a single-strand DNA and it is not affected by the restriction endonuclease barrier of the recipients. However, it is a time-consuming method that requires selection steps that can be challenging. Wang et al. have reported that Pseudoalteromonas species are sensitive to chloramphenicol according to the result of antibiotic assay, however, a few false positive colonies were observed after the single crossover event. The solution was to change the selection marker to the erythromycin resistant gene [68]. Moreover, using antibiotic resistance genes as selection makers somehow could affect the conjugation efficiency, especially on salt-dependent marine bacteria. For example, the presence of high salt concentrations will decrease the activities of tetracycline and gentamicin [215]. Another issue is how to select the real mutants. Although the sacB gene is widely used as a counter-selection marker for allelic exchange in several marine proteobacteria, such as Vibrio [216–218] and Pseudoalteromonas species [68,219], the toxicity of sacB products is susceptible to the NaCl in the selective medium [220]. The growth of most marine bacteria is salt-dependent and that would lead to the inadequate toxicity of SacB, which results in decreasing the ratio of the deletion mutants on the selective plates [221]. Luo et al. have developed an efficient

36 conjugation-based method for rapid deletion mutation in several Vibrio species by using a novel counter-selectable marker, the vmi480 gene which can produce a strong toxic protein [221]. They succeeded in generating a range of mutants in V. cholerae, V. parahaemolyticus, and Vibrio alginolyticus by using vmi480 as a counter-selectable marker [221].

A tricky issue in the bacterial conjugation procedure is the selection of recipient colonies after mating, because both donor and recipient cells contain the same encoded traits. Many studies have reported different procedures to avoid this issue. Sawabe et al. labeled 39 Vibrio strains with a green fluorescent protein (GFP) by intergeneric conjugation with E. coli [222]. The mixture of donor and recipient cells was incubated on a 2216E agar allowing the growth of both strains. Then, the transconjugants were selected on the selective medium, the alginate- containing seawater medium [222]. The authors demonstrated that a critical step for the selection is that the gfp-transformed recipients must be significantly enriched on the selective medium to separate from the donor E. coli cells [222]. However, both of the strains can grow on the selective medium that is of course making the selection much difficult. In another study, the authors used an easy approach to avoid this selective outgrowth. Wang et al. have used an auxotrophic strain E. coli WM3064 in which growth relies on the supplementation of diaminopimelic acid (DAP). After the mating step, transconjugants were transferred on the 2216E agar plate without DAP and the donors cannot grow on the plates without the supplement of DAP. In conclusion, the combination of effective counterselection markers and auxotrophic donor strains could increase the efficiency of bacterial conjugation approaches on genetic manipulation of marine Proteobacteria. Besides these issues, the mating ratio of the donor to recipient cells can significantly affect the conjugation efficiency. Piekarski et al. have reported that the optimal ratios of conjugation at the donor E. coli to the recipient Roseobacter species are 5:1 or 10:1 [207], while the best conjugation efficiency for Pseudoalteromonas sp. SM9913 was obtained at a donor to recipient ratio of 100:1.

The successful conjugation-based approaches allow investigation of the bioactive potential in marine Proteobacteria and enable a procedure of “compound to gene” for example to identify BGC from its natural producer. Thiomarinol, an antimicrobial compound produced by marine bacteria, such as Pseudoalteromonas SANK 73390 [78] and Fukuda et al. used the suicide vector pAKE604 containing sacB gene as a counterselectable marker to facilitate allelic exchange and succeeded in generating three separate gene knockouts. The authors found

37 that the biosynthesis of thiomarinol is based on two independent pathways: the pseudomonic acid and the pyrrothine, which are responsible for thiomarinol production.

Conjugation-based transformation is not only useful for generating the deletion mutant but also allows other genetic manipulations. An example of this case is that the BGC for astaxanthin production from Alphaproteobacteria Paracoccus was cloned into the vector pBBR1-MCS2, and then the constructed vector was transferred into Paracoccus by conjugation with the donor E. coli strain to enhance the production of astaxanthin in its natural producer [223]. Likewise, the lysC gene, an aspartate kinase, was cloned into the vector pHS15N and the recombinant vector was transferred into a proteobacterium Halomonas elongata to improve the production of ectoine [224].

During this PhD project, we used a conjugational transformation approach in the pigmented Pseudoalteromonas rubra S4059. Although several studies have successfully developed conjugational transformation approaches for genetic manipulation Pseudoalteromonas species, several features of this genus challenge the genetic manipulation as summarized by Wang et al. [68]. Briefly, the active multidrug resistance genes and efflux pumps in Pseudoalteromonas, enable their resistance to a range of antibiotics and thus this, of course, makes it challenging to find a suitable antibiotic marker to use in genetic engineering. Secondly, the efficient restriction-modification systems in Pseudoalteromonas can degrade foreign DNA leading to reducing transformation efficiency. Lastly, the salt-dependent property of Pseudoalteromonas does not comply well with most donor organisms of non- marine origin. This characteristic further eliminates the use of electroporation as the reason described previously. In particular, most pigmented Pseudoalteromonas species can produce a range of antimicrobial compounds that result in decreasing the conjugation efficiency during mating with the donor E. coli. Fortunately, we finally established a conjugation-based approach according to previously described methods allowing genetic manipulation of a pigmented bacterium P. rubra S4059 as described in paper 1. Briefly, the suicide plasmids were constructed using pDM4 as the backbone [217,225]. The pDM4 plasmid contains a R6K replicon origin, a mob gene and oirT sequence for conjugation, and a chloramphenicol resistance gene cat and a sacB gene for counter selection [217,225]. An approximately 1-kb upstream and downstream region flanking of target genes were amplified and spliced by overlap PCR. The amplified recombining arms were inserted into pDM4 by RecET direct

38 cloning system [225]. The constructed plasmid was transferred into S4059 by intergenic conjugation with the donor E. coli WM3064 whose growth relies on the supplementation of DAP. After first crossover event, the suicide plasmid was inserted into the chromosome of S4059 by homologous recombination and the mutants were selected based on the antibiotic resistant gene, in this case is chloramphenicol resistance gene (Figure 15). Then sacB gene as a counter selection marker was used for second crossover event. The colonies of inserted mutants were transferred on a selection plate containing 10% sucrose to promote the second homologous recombination and the real in-frame deletion mutants were approved by both PCR results and DNA sequencing. Using the established genetic manipulation approach, we succeed in generating a range of mutants, such as a GH19 mutant in paper 1, LPMO single and double mutants (have not published) and a prodigiosin deficient mutant pigC in paper 2.

Figure 14. Schematics illustrating the use of suicide plasmid to generate an in-frame deletion mutant in Pseudoalteromonas strains. (A) the constructed suicide plasmid contains a sacB gene, an antibiotic resistant gene as counter selection markers and the recombining arms. (B) The suicide plasmid was inserted into the chromosome after first homologous recombination. (C) Sucrose counter selection for second homologous recombination. This figure was cited from Wang et al. [68].

Several genetic manipulation approaches, as aforementioned, have been applied to different marine bacteria. Nevertheless, it is challenging to develop a universal method for transforming exogenous DNA in any given marine bacterium. The parameters for genetic manipulation for example exogenous DNA concentration, electric voltage and incubation time or temperature of conjugation on different marine bacterial strains are quite diverse and testing these parameters is time-consuming. Therefore, it is necessary that further research for the given bacterium is needed to optimize protocols of genetic manipulation approaches.

6. CONCLUSIONS AND FUTURE PERSPECTIVES

The increase of microbial antibiotic resistance calls for rapid development of novel antibiotics. At present, marine bacteria have received significant attention due to the large potential on pharmaceutical and industrial substrates production. In this PhD thesis, pigmented marine Pseudoalteromonas rurba S4059 was selected as a model strain to explore its bioactive potential with a combination of genetic manipulation and cultivation-based approaches. We developed a conjugation-based approach allowing genetic manipulation in pigmented P. rubra S4059 and using this approach, we verified the prodigiosin BGC from this strain and found a blue pigment, dipyrrolyldipyrromethene prodigiosin.

Chitin as the ecological niche nutrient can induce secondary metabolites production in several Vibrio species and pigmented Pseudoalteromonas contain a highly chitinolytic machinery according to previous work in our group. Altogether, we tried to explore the potential link between chitin degradation and secondary metabolites production in S4059. Unfortunately, chitin did not significantly affect secondary metabolites production in S4059 as compared to other carbon sources. Interestingly, we observed that chitin monomer can induce overall secondary metabolites production in S4059 and is likely to be the key factor between chitin degradation and secondary metabolites production. However, we do not know how chitin monomers regulate gene expression or protein production in S4059. For future purposes, transcriptomic and proteomic analysis can unravel the mechanism in S4059.

Genetic manipulation in pigmented Pseudoalteromonas species is challenging, we finally developed a conjugation-based approach via homologous recombination allowing genetic manipulation in S4059 and succeeded in generated several mutants in S4059. However, this approach is still time- consulting and it takes at least one month to generate a mutant in S4059. For future work, it may good to develop CRISPR-cas9 systems for genetic manipulation in this strain.

The ecological-based bioprospecting is to mimic natural environment or its environmental niches to culture bacteria in the laboratory. Given the genetic potential for chitin degradation, natural chitin surfaces are likely an ecological niche for pigmented Pseudoalteromonas species. We observed that growth on natural chitin surface or chitin flakes lead to loss of pigment production in both S4059 and another yellow pigmented strain, P. piscicida S2724.

Growth on natural chitin surface can be an interesting setup to explore how an ecological niche affects gene expression or protein production in pigmented Pseudoalteromonas. In conclusion, this work not only has developed a genetic manipulation approach in pigmented Pseudoalteromonas rubra S4059, but also indicated that natural chitin surface can provide an environment facilitating genetic diversity in pigmented P. rurba.

7. ACKNOWLEDGEMENTS

First, I would like to thank my supervisor Lone Gram for her guidance during this research. Thank you so much for trusting me and providing me the opportunity to work on this promising project. Your insightful feedback pushed me to sharpen my thinking and brought my work to a higher level.

I would like to thank my co-supervisor Shengda Zhang, our “molecular wizard”. Thank you for guiding me to develop a genetic manipulation approach and providing your support, practical advice, and encouragement on both scientific and non-scientific alike.

I would like to thank my past and present colleagues of the GramLab for scientific discussions and social events.

A big thanks to all my collaborators Thomas Isbrandt, Emil Ørsted Christensen, Mikael Lenz Strube, Sara Skøtt Paulsen, Maike Wennekers Nielsen, Yannick Buijs, Erwin M. Schoof, Thomas Ostenfeld Larsen to support my PhD project. A special thanks to Emil Ørsted Christensen and my supervisor Lone Gram for translating my thesis abstract from English to Danish.

Finally, I would like to thank my family for supporting me to go abroad to study and thank my girlfriend Mengni Cui for encouraging me all the way.

8. REFERENCES

1. Khaw, K.T. Comment on Miettinen: Rose revisited. Eur. J. Epidemiol. 2004, 19, 745–748, doi:10.1023/B:EJEP.0000036615.16155.d2. 2. Xiong, Z.Q.; Wang, J.F.; Hao, Y.Y.; Wang, Y. Recent advances in the discovery and development of marine microbial natural products. Mar. Drugs 2013, 11, 700–717, doi:10.3390/md11030700. 3. Fischbach, M.A.; Walsh, C.T. Antibiotics for emerging pathogens. Science (80-. ). 2009, 325, 1089–1093, doi:10.1126/science.1176667. 4. Fenical, W. Chemical Studies of Marine Bacteria: Developing a New Resource. Chem. Rev. 1993, 93, 1673–1683, doi:10.1021/cr00021a001. 5. Molinski, T.F.; Dalisay, D.S.; Lievens, S.L.; Saludes, J.P. Drug development from marine natural products. Nat. Rev. Drug Discov. 2009, 8, 69–85, doi:10.1038/nrd2487. 6. Andryukov, B.; Mikhailov, V.; Besednova, N. The biotechnological potential of secondary metabolites from marine bacteria. J. Mar. Sci. Eng. 2019, 7, 1–16, doi:10.3390/jmse7060176. 7. Romano, G.; Costantini, M.; Sansone, C.; Lauritano, C.; Ruocco, N.; Ianora, A. Marine microorganisms as a promising and sustainable source of bioactive molecules. Mar. Environ. Res. 2017, 128, 58–69, doi:10.1016/j.marenvres.2016.05.002. 8. Parte, S.; Sirisha, V.L.; D’Souza, J.S. Biotechnological Applications of Marine Enzymes From Algae, Bacteria, Fungi, and Sponges. Adv. Food Nutr. Res. 2017, 80, 75–106, doi:10.1016/bs.afnr.2016.10.005. 9. Machado, H.; Sonnenschein, E.C.; Melchiorsen, J.; Gram, L. Genome mining reveals unlocked bioactive potential of marine Gram-negative bacteria. BMC Genomics 2015, 16, 1–12, doi:10.1186/s12864-015-1365-z. 10. Paulsen, S.S.; Strube, M.L.; Bech, P.K.; Gram, L.; Sonnenschein, E.C. Marine Chitinolytic Pseudoalteromonas Represents an Untapped Reservoir of Bioactive Potential . mSystems 2019, 4, 1–12, doi:10.1128/msystems.00060-19. 11. Jensen, P.R.; Chavarria, K.L.; Fenical, W.; Moore, B.S.; Ziemert, N. Challenges and triumphs to genomics-based natural product discovery. J. Ind. Microbiol. Biotechnol. 2014, 41, 203–209, doi:10.1007/s10295-013-1353-8. 12. Bowman, J.P. Bioactive compound synthetic capacity and ecological significance of marine bacterial genus Pseudoalteromonas. Mar. Drugs 2007, 5, 220–241, doi:10.3390/md504220. 13. Tang, B.L.; Yang, J.; Chen, X.L.; Wang, P.; Zhao, H.L.; Su, H.N.; Li, C.Y.; Yu, Y.; Zhong, S.; Wang, L.; et al. A predator-prey interaction between a marine Pseudoalteromonas sp. and Gram-positive bacteria. Nat. Commun. 2020, 11, 285, doi:10.1038/s41467-019- 14133-x. 14. Vynne, N.G.; Månsson, M.; Nielsen, K.F.; Gram, L. Bioactivity, Chemical Profiling, and 16S rRNA-Based Phylogeny of Pseudoalteromonas Strains Collected on a Global

Research Cruise. Mar. Biotechnol. 2011, 13, 1062–1073, doi:10.1007/s10126-011- 9369-4. 15. Pinkerton, D.M.; Banwell, M.G.; Willis, A.C. Total syntheses of tambjamines C, E, F, G, H, i and J, BE-18591, and a related alkaloid from the marine bacterium pseudoalteromonas tunicata. Org. Lett. 2007, 9, 5127–5130, doi:10.1021/ol7024313. 16. Nmr, C.; Nmr, C. A 2-Substituted Prodiginine , 2- ( p -Hydroxybenzyl ) prodigiosin , from Pseudoalteromonas rubra. 2008, 2, 1970–1972. 17. Gram, L.; Melchiorsen, J.; Bruhn, J.B. Antibacterial activity of marine culturable bacteria collected from a global sampling of ocean surface waters and surface swabs of marine organisms. Mar. Biotechnol. 2010, 12, 439–451, doi:10.1007/s10126-009-9233-y. 18. Rutledge, P.J.; Challis, G.L. Discovery of microbial natural products by activation of silent biosynthetic gene clusters. Nat. Publ. Gr. 2015, 13, 509–523, doi:10.1038/nrmicro3496. 19. Bode, H.B.; Bethe, B.; Höfs, R.; Zeeck, A. Big effects from small changes: Possible ways to explore nature’s chemical diversity. ChemBioChem 2002, 3, 619–627, doi:10.1002/1439-7633(20020703)3:7<619::AID-CBIC619>3.0.CO;2-9. 20. Giubergia, S.; Phippen, C.; Nielsen, K.F.; Gram, L. Growth on Chitin Impacts the Transcriptome and Metabolite Profiles of Antibiotic-Producing Vibrio coralliilyticus S2052 and Photobacterium galatheae S2753. mSystems 2017, 2, 1–12, doi:10.1128/msystems.00141-16. 21. Zobell, C.E.; Rittenberg, S.C. the Occurrence and Characteristics of Chitinoclastic Bacteria in the Sea. J. Bacteriol. 1938, 35, 275–87, doi:10.1128/jb.35.3.275-287.1938. 22. Giubergia, S.; Phippen, C.; Gotfredsen, C.H.; Nielsen, K.F.; Gram, L. Influence of niche- specific nutrients on secondary metabolism in Vibrionaceae. Appl. Environ. Microbiol. 2016, 82, 4035–4044, doi:10.1128/AEM.00730-16. 23. Gooday, G.W. The Ecology of Chitin Degradation. 1990, 387–430, doi:10.1007/978-1- 4684-7612-5_10. 24. Bai, Y.; Eijsink, V.G.H.; Kielak, A.M.; van Veen, J.A.; de Boer, W. Genomic comparison of chitinolytic enzyme systems from terrestrial and aquatic bacteria. Environ. Microbiol. 2016, 18, 38–49, doi:10.1111/1462-2920.12545. 25. Oyeleye, A.; Normi, Y.M. Chitinase: Diversity, limitations, and trends in Engineering for suitable applications. Biosci. Rep. 2018, 38, 1–21, doi:10.1042/BSR20180323. 26. Vaaje-Kolstad, G.; Westereng, B.; Horn, S.J.; Liu, Z.; Zhai, H.; Sørlie, M.; Eijsink, V.G.H. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science (80-. ). 2010, 330, 219–222, doi:10.1126/science.1192231. 27. Levasseur, A.; Drula, E.; Lombard, V.; Coutinho, P.M.; Henrissat, B. Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes. Biotechnol. Biofuels 2013, 6, 1–14, doi:10.1186/1754-6834-6-41. 28. Quinlan, R.J.; Sweeney, M.D.; Lo Leggio, L.; Otten, H.; Poulsen, J.C.N.; Johansen, K.S.;

Krogh, K.B.R.M.; Jørgensen, C.I.; Tovborg, M.; Anthonsen, A.; et al. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 15079–15084, doi:10.1073/pnas.1105776108. 29. Fleming A On the Antibacterial Action of Cultures of a Penicillium, with Special Reference to their Use in the Isolation of B. influenzæ. Br J Exp Pathol 1929, 226–236. 30. Patridge, E.; Gareiss, P.; Kinch, M.S.; Hoyer, D. An analysis of FDA-approved drugs: Natural products and their derivatives. Drug Discov. Today 2016, 21, 204–207, doi:10.1016/j.drudis.2015.01.009. 31. Hoffmeister, D.; Keller, N.P. Natural products of filamentous fungi: Enzymes, genes, and their regulation. Nat. Prod. Rep. 2007, 24, 393–416, doi:10.1039/b603084j. 32. Macleod, R.A. the Question of the Existence of Specific Marine Bacteria. Bacteriol. Rev. 1965, 29, 9–24, doi:10.1128/mmbr.29.1.9-23.1965. 33. Delong, E.F. Modern microbial seascapes. Nat. Rev. Microbiol. 2007, 5, 755–757, doi:10.1038/nrmicro1762. 34. Carroll, A.R.; Copp, B.R.; Davis, R.A.; Keyzers, R.A.; Prinsep, M.R. Marine natural products. Nat. Prod. Rep. 2019, 36, 122–173, doi:10.1039/c8np00092a. 35. Whitman, W.B.; Coleman, D.C.; Wiebe, W.J. Prokaryotes: The unseen majority. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 6578–6583, doi:10.1073/pnas.95.12.6578. 36. Campbell, L.; Vaulot, D. Photosynthetic picoplankton community structure in the subtropical North Pacific Ocean near Hawaii (station ALOHA). Deep. Res. Part I 1993, 40, 2043–2060, doi:10.1016/0967-0637(93)90044-4. 37. Britschgi, T.B.; Giovannoni, S.J. Phylogenetic analysis of a natural marine bacterioplankton population by rRNA gene cloning and sequencing. Appl. Environ. Microbiol. 1991, 57, 1707–1713, doi:10.1128/aem.57.6.1707-1713.1991. 38. Fuhrman, J.A.; McCallum, K.; Davis, A.A. Phylogenetic diversity of subsurface marine microbial communities from the Atlantic and Pacific Oceans. Appl. Environ. Microbiol. 1993, 59, 1294–1302, doi:10.1128/aem.61.12.4517-4517.1995. 39. Bowman, J.P.; Mccammon, sharee A.; Bbrown, M. V.; Nichols, D.S.; Mcmeekin, T.A. Diversity and Association of Psychrophilic Bacteria in Antarctic Sea Ice. Appl. Environ. Microbiol. 1997, 63, 3068–3078, doi:10.1103/PhysRevLett.40.1423. 40. Lee, S.-Y.; Eom, Y.-B. Analysis of Microbial Composition Associated with Freshwater and Seawater. Biomed. Sci. Lett. 2016, 22, 150–159, doi:10.15616/bsl.2016.22.4.150. 41. Nimnoi, P.; Pongsilp, N. Marine bacterial communities in the upper gulf of Thailand assessed by Illumina next-generation sequencing platform. BMC Microbiol. 2020, 20, 1–11, doi:10.1186/s12866-020-1701-6. 42. Porter, K.G.; Feig, Y.S. The use of DAPI for identifying and counting aquatic microflora. Limnol. Oceanogr. 1980, 25, 943–948, doi:10.4319/lo.1980.25.5.0943. 43. Mansson, M.; Gram, L.; Larsen, T.O. Production of bioactive secondary metabolites by

marine Vibrionaceae. Mar. Drugs 2011, 9, 1440–1468, doi:10.3390/md9091440. 44. Timmermans, M.L.; Paudel, Y.P.; Ross, A.C. Investigating the biosynthesis of natural products from marine proteobacteria: A survey of molecules and strategies. Mar. Drugs 2017, 15, doi:10.3390/md15080235. 45. Bérdy, J. Erratum: Thoughts and facts about antibiotics: Where we are now and where we are heading (Journal of Antibiotics (2012) 65 (385-395) DOI: 10.1038/ja.2012.27). J. Antibiot. (Tokyo). 2012, 65, 441, doi:10.1038/ja.2012.54. 46. Solanki, R.; Khanna, M.; Lal, R. Bioactive compounds from marine actinomycetes. Indian J. Microbiol. 2008, 48, 410–431, doi:10.1007/s12088-008-0052-z. 47. Dhakal, D.; Pokhrel, A.R.; Shrestha, B.; Sohng, J.K. Marine rare actinobacteria: Isolation, characterization, and strategies for harnessing bioactive compounds. Front. Microbiol. 2017, 8, doi:10.3389/fmicb.2017.01106. 48. Schumacher, R.W.; Talmage, S.C.; Miller, S.A.; Sarris, K.E.; Davidson, B.S.; Goldberg, A. Isolation and structure determination of an antimicrobial ester from a marine sediment-derived bacterium. J. Nat. Prod. 2003, 66, 1291–1293, doi:10.1021/np020594e. 49. Shao, J.W.; Fotso, S.; Li, F.; Qin, S.; Laatsch, H. Amorphane sesquiterpenes from a marine Streptomyces sp. J. Nat. Prod. 2007, 70, 304–306, doi:10.1021/np050358e. 50. Moore, B.S.; Trischman, J.A.; Seng, D.; Kho, D.; Jensen, P.R.; Fenical, W. Salinamides, antiinflammatory depsipeptides from a marine streptomycete. J. Org. Chem. 1999, 64, 1145–1150, doi:10.1021/jo9814391. 51. Fenicala, W.; Jensena, P.R.; Palladinob, M.A.; Lamb, K.S.; Lloydb, G.K.; Potts, B.C. Discovery and Development of the Anticancer Agent Salinosporamide A (NPI-0052). Bioorg Med Chem 2009, 17, 2175, doi:10.1161/CIRCULATIONAHA.110.956839. 52. Feling, R.H.; Buchanan, G.O.; Mincer, T.J.; Kauffman, C.A.; Jensen, P.R.; Fenical, W. Salinosporamide A: A highly cytotoxic proteasome inhibitor from a novel microbial source, a marine bacterium of the new genus Salinospora. Angew. Chemie - Int. Ed. 2003, 42, 355–357, doi:10.1002/anie.200390115. 53. Maskey, R.P.; Helmke, E.; Kayser, O.; Fiebig, H.H.; Maier, A.; Busche, A.; Laatsch, H. Anti- cancer and antibacterial trioxacarcins with high anti-malaria activity from a marine streptomycete and their absolute stereochemistry. J. Antibiot. (Tokyo). 2004, 57, 771– 779, doi:10.7164/antibiotics.57.771. 54. Tsukimoto, M.; Nagaoka, M.; Shishido, Y.; Fujimoto, J.; Nishisaka, F.; Matsumoto, S.; Harunari, E.; Imada, C.; Matsuzaki, T. Bacterial production of the tunicate-derived antitumor cyclic depsipeptide didemnin B. J. Nat. Prod. 2011, 74, 2329–2331, doi:10.1021/np200543z. 55. Oh, D.C.; Strangman, W.K.; Kauffman, C.A.; Jensen, P.R.; Fenical, W. Thalassospiramides A and B, immunosuppressive peptides from the marine bacterium thalassospira sp. Org. Lett. 2007, 9, 1525–1528, doi:10.1021/ol070294u. 56. Brinkhoff, T.; Bach, G.; Heidorn, T.; Liang, L.; Schlingloff, A.; Simon, M. Antibiotic

Production by a Roseobacter Clade-Affiliated Species from the German Wadden Sea and Its Antagonistic Effects on Indigenous Isolates. Appl. Environ. Microbiol. 2004, 70, 2560–2565, doi:10.1128/AEM.70.4.2560-2565.2003. 57. Lee, J.; Currano, J.N.; Carroll, P.J.; Joullié, M.M. Didemnins, tamandarins and related natural products. Nat. Prod. Rep. 2012, 29, 404–424, doi:10.1039/c2np00065b. 58. Lee, J.; Currano, J.N.; Carroll, P.J.; Joullié, M.M. Didemnins, tamandarins and related natural products. Nat. Prod. Rep. 2012, 29, 404–424, doi:10.1039/c2np00065b. 59. Rinehart, K.L.; Gloer, J.B.; Hughes, R.G.; Renis, H.E.; Mcgovren, P.; Swynenberg, E.B.; Stringfellow, D.A.; Kuentzel, S.L.; Porter, K.G. Didemnins: Antiviral and Antitumor Depsipeptides from a Caribbean Tunicate. Science (80-. ). 1981, 212, 933–935. 60. Xu, Y.; Kersten, R.D.; Nam, S.J.; Lu, L.; Al-Suwailem, A.M.; Zheng, H.; Fenical, W.; Dorrestein, P.C.; Moore, B.S.; Qian, P.Y. Bacterial biosynthesis and maturation of the didemnin anti-cancer agents. J. Am. Chem. Soc. 2012, 134, 8625–8632, doi:10.1021/ja301735a. 61. Williamson, S.K.; Wolf, M.K.; Eisenberger, M.A.; O’Rourke, M.; Brannon, W.; Crawford, E.D. Phase II evaluation of didemnin B in hormonally refractory metastatic prostate cancer - A southwest oncology group study. Invest. New Drugs 1995, 13, 167–170, doi:10.1007/BF00872867. 62. Porsby, C.H.; Nielsen, K.F.; Gram, L. Phaeobacter and Ruegeria species of the Roseobacter clade colonize separate niches in a Danish turbot (Scophthalmus maximus)-rearing farm and antagonize Vibrio anguillarum under different growth conditions. Appl. Environ. Microbiol. 2008, 74, 7356–7364, doi:10.1128/AEM.01738-08. 63. Harrington, C.; Reen, F.J.; Mooij, M.J.; Stewart, F.A. Characterisation of Non- Autoinducing Tropodithietic Acid (TDA) Production from Marine Sponge Pseudovibrio Species. Mar Drugs 2014, 5960–5978, doi:10.3390/md12125960. 64. D’Alvise, P.W.; Lillebø, S.; Wergeland, H.I.; Gram, L.; Bergh, Ø. Protection of cod larvae from vibriosis by Phaeobacter spp.: A comparison of strains and introduction times. Aquaculture 2013, 384–387, 82–86, doi:10.1016/j.aquaculture.2012.12.013. 65. Wilson, M.Z.; Wang, R.; Gitai, Z.; Seyedsayamdost, M.R. Mode of action and resistance studies unveil new roles for tropodithietic acid as an anticancer agent and the ?- glutamyl cycle as a proton sink. Proc. Natl. Acad. Sci. U. S. A. 2016, 113, 1630–1635, doi:10.1073/pnas.1518034113. 66. Offret, C.; Desriac, F.; Le Chevalier, P.; Mounier, J.; Jégou, C.; Fleury, Y. Spotlight on antimicrobial metabolites from the marine bacteria Pseudoalteromonas: Chemodiversity and ecological significance. Mar. Drugs 2016, 14, doi:10.3390/md14070129. 67. Williams, R.P. Biosynthesis of prodigiosin, a secondary metabolite of Serratia marcescens. Appl. Microbiol. 1973, 25, 396–402, doi:10.1128/aem.25.3.396-402.1973. 68. Wang, P.; Yu, Z.; Li, B.; Cai, X.; Zeng, Z.; Chen, X.; Wang, X. Development of an efficient conjugation-based genetic manipulation system for Pseudoalteromonas. Microb. Cell Fact. 2015, 14, 1–11, doi:10.1186/s12934-015-0194-8.

69. Sakai-Kawada, F.E.; Ip, C.G.; Hagiwara, K.A.; Awaya, J.D. Biosynthesis and bioactivity of prodiginine analogs in marine bacteria, Pseudoalteromonas: A mini review. Front. Microbiol. 2019, 10, 1–9, doi:10.3389/fmicb.2019.01715. 70. Fischbach, M.A.; Walsh, C.T.; Clardy, J. The evolution of gene collectives: How natural selection drives chemical innovation. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 4601– 4608, doi:10.1073/pnas.0709132105. 71. Brady, S.F.; Chao, C.J.; Handelsman, J.; Clardy, J. Cloning and Heterologous Expression of a Natural Product Biosynthetic Gene Cluster from eDNA. Org. Lett. 2001, 3, 1981– 1983, doi:10.1021/ol015949k. 72. Zhang, J.J.; Tang, X.; Zhang, M.; Nguyen, D.; Moore, B.S. Broad-host-range expression reveals native and host regulatory elements that influence heterologous antibiotic production in Gram-negative bacteria. MBio 2017, 8, 1–16, doi:10.1128/mBio.01291- 17. 73. Li, B.; Walsh, C.T. Identification of the gene cluster for the dithiolopyrrolone antibiotic holomycin in Streptomyces clavuligerus. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 19731–19735, doi:10.1073/pnas.1014140107. 74. Still, P.C.; Johnson, T.A.; Theodore, C.M.; Loveridge, S.T.; Crews, P. Scrutinizing the scaffolds of marine biosynthetics from different source organisms: Gram-negative cultured bacterial products enter center stage. J. Nat. Prod. 2014, 77, 690–702, doi:10.1021/np500041x. 75. Tambadou, F.; Lanneluc, I.; Sablé, S.; Klein, G.L.; Doghri, I.; Sopéna, V.; Didelot, S.; Barthélémy, C.; Thiéry, V.; Chevrot, R. Novel nonribosomal peptide synthetase (NRPS) genes sequenced from intertidal mudflat bacteria. FEMS Microbiol. Lett. 2014, 357, 123–130, doi:10.1111/1574-6968.12532. 76. Speitling, M.; Smetanina, O.F.; Kuznetsova, T.A.; Laatsch, H. Bromoalterochromides A and A′, unprecedented chromopeptides from a marine Pseudoalteromonas maricaloris strain KMM 636T. J. Antibiot. (Tokyo). 2007, 60, 36–42, doi:10.1038/ja.2007.5. 77. Chekan, J.R.; Moore, B.S. Preparation and Characterization of Tetrabromopyrrole Debrominase From Marine Proteobacteria; 1st ed.; Elsevier Inc., 2018; Vol. 605;. 78. Fukuda, D.; Haines, A.S.; Song, Z.; Murphy, A.C.; Hothersall, J.; Stephens, E.R.; Gurney, R.; Cox, R.J.; Crosby, J.; Willis, C.L.; et al. A natural plasmid uniquely encodes two biosynthetic pathways creating a potent anti-MRSA antibiotic. PLoS One 2011, 6, 1–9, doi:10.1371/journal.pone.0018031. 79. Sobolevskaya, M.P.; Smetanina, O.F.; Speitling, M.; Shevchenko, L.S.; Dmitrenok, P.S.; Laatsch, H.; Kuznetsova, T.A.; Ivanova, E.P.; Elyakov, G.B. Controlling production of brominated cyclic depsipeptides by Pseudoalteromonas maricaloris KMM 636T. Lett. Appl. Microbiol. 2005, 40, 243–248, doi:10.1111/j.1472-765X.2005.01635.x. 80. Burkholder, P.R.; Pfister, R.M.; Leitz, F.H. Production of a pyrrole antibiotic by a marine bacterium. Appl. Microbiol. 1966, 14, 649–653, doi:10.1128/aem.14.4.649-653.1966. 81. Lovell, F.M. The Structure of a Bromine-Rich Marine Antibiotic. J. Am. Chem. Soc. 1966, 88, 4510–4511, doi:10.1021/ja00971a040.

82. Shiozawa, H.; Kagasaki, T.; Takahashi, S.; Kinoshita, T.; Haruyama, H.; Domon, H.; Utsuib, Y.; Kodama, K. Thiomarinol, a new hybrid antimicrobial antibiotic produced by a marine bacterium fermentation, isolation, structure, and antimicrobial activity. J. Antibiot. (Tokyo). 1993, 46, 1834–1842, doi:10.7164/antibiotics.46.1834. 83. Gemperlein, K.; Zaburannyi, N.; Garcia, R.; Clair, J.J. La; Müller, R. Metabolic and Biosynthetic Diversity in Marine Myxobacteria. Mar Drugs 2018, 16, 314, doi:10.3390/md16090314. 84. Fudou, R.; Iizuka, T.; Yamanaka, S. Haliangicin, a novel antifungal metabolite produced by a marine myxobacterium. 1. Fermentation and biological characteristics. J. Antibiot. (Tokyo). 2001, 54, 149–152, doi:10.7164/antibiotics.54.149. 85. Sun, Y.; Feng, Z.; Tomura, T.; Suzuki, A.; Miyano, S.; Tsuge, T.; Mori, H.; Suh, J.W.; Iizuka, T.; Fudou, R.; et al. Heterologous Production of the Marine Myxobacterial Antibiotic Haliangicin and Its Unnatural Analogues Generated by Engineering of the Biochemical Pathway. Sci. Rep. 2016, 6, 1–11, doi:10.1038/srep22091. 86. Ohlendorf, B.; Leyers, S.; Krick, A.; Kehraus, S.; Wiese, M.; König, G.M. Phenylnannolones A-C: biosynthesis of new secondary metabolites from the myxobacterium Nannocystis exedens. Chembiochem 2008, 9, 2997–3003, doi:10.1002/cbic.200800434. 87. Sun, Y.; Tomura, T.; Sato, J.; Iizuka, T.; Fudou, R.; Ojika, M. Isolation and biosynthetic analysis of haliamide, a new pks-nrps hybrid metabolite from the marine myxobacterium haliangium ochraceum. Molecules 2016, 21, doi:10.3390/molecules21010059. 88. Buijs, Y.; Bech, P.K.; Vazquez-Albacete, D.; Bentzon-Tilia, M.; Sonnenschein, E.C.; Gram, L.; Zhang, S. Da Marine Proteobacteria as a source of natural products: Advances in molecular tools and strategies; Royal Society of Chemistry, 2019; Vol. 36, pp. 1333– 1350;. 89. Lemos, M.L.; Dopazo, C.P.; Toranzo, A.E.; Barja, J.L. Competitive dominance of antibiotic‐producing marine bacteria in mixed cultures. J. Appl. Bacteriol. 1991, 71, 228–232, doi:10.1111/j.1365-2672.1991.tb04452.x. 90. Long, R.A.; Rowley, D.C.; Zamora, E.; Liu, J.; Bartlett, D.H.; Azam, F. Antagonistic interactions among marine bacteria impede the proliferation of Vibrio cholerae. Appl. Environ. Microbiol. 2005, 71, 8531–8536, doi:10.1128/AEM.71.12.8531-8536.2005. 91. Yoshikawa, K.; Nakayama, Y.; Hayashi, M.; Unemoto, T.; Mochida, K. Korormicin, an antibiotic specific for gram-negative marine bacteria, strongly inhibits the respiratory chain-linked Na+-translocating NADH: Quinone reductase from the marine Vibrio alginolyticus. J. Antibiot. (Tokyo). 1999, 52, 182–185, doi:10.7164/antibiotics.52.182. 92. Danevčič, T.; Borić, M.; Stopar, D. Microbial ecophysiology of Vibrio ruber. Food Technol. Biotechnol. 2014, 52, 198–203. 93. Starič, N.; Danevčič, T.; Stopar, D. Vibrio sp. DSM 14379 Pigment Production-A Competitive Advantage in the Environment? Microb. Ecol. 2010, 60, 592–598, doi:10.1007/s00248-010-9671-0.

94. Allen, G.R.; Reichelt, J.L.; Gray, P.P. Influence of environmental factors and medium composition on Vibrio gazogenes growth and prodigiosin production. Appl. Environ. Microbiol. 1983, 45, 1727–1732, doi:10.1128/aem.45.6.1727-1732.1983. 95. Borić, M.; Danevčič, T.; Stopar, D. Prodigiosin from Vibrio sp. DSM 14379; A New UV- Protective Pigment. Microb. Ecol. 2011, 62, 528–536, doi:10.1007/s00248-011-9857-0. 96. Kotob, S.I.; Coon, S.L.; Quintero, E.J.; Weiner, R.M. Homogentisic acid is the primary precursor of melanin synthesis in Vibrio cholerae, a Hyphomonas strain, and Shewanella colwelliana. Appl. Environ. Microbiol. 1995, 61, 1620–1622, doi:10.1128/aem.61.4.1620-1622.1995. 97. Shank, E.A.; Kolter, R. New Developments in Microbial Interspecies Signaling. Curr Opin Microbiol 2009, 12, 205–214, doi:10.1038/jid.2014.371. 98. Berger, M.; Neumann, A.; Schulz, S.; Simon, M.; Brinkhoff, T. Tropodithietic acid production in Phaeobacter gallaeciensis is regulated by N-acyl homoserine lactone- mediated quorum sensing. J. Bacteriol. 2011, 193, 6576–6585, doi:10.1128/JB.05818- 11. 99. Geng, H.; Belas, R. Expression of tropodithietic acid biosynthesis is controlled by a novel autoinducer. J. Bacteriol. 2010, 192, 4377–4387, doi:10.1128/JB.00410-10. 100. Buijs, Y.; Isbrandt, T.; Zhang, S.-D.; Larsen, T.O.; Gram, L. The Antibiotic Andrimid Produced by Vibrio coralliilyticus Increases Expression of Biosynthetic Gene Clusters and Antibiotic Production in Photobacterium galatheae. Front. Microbiol. 2020, 11, 1– 13, doi:10.3389/fmicb.2020.622055. 101. Fajardo, A.; Martínez, J.L. Antibiotics as signals that trigger specific bacterial responses. Curr. Opin. Microbiol. 2008, 11, 161–167, doi:10.1016/j.mib.2008.02.006. 102. Lories, B.; Roberfroid, S.; Dieltjens, L.; De Coster, D.; Foster, K.R.; Steenackers, H.P. Biofilm Bacteria Use Stress Responses to Detect and Respond to Competitors. Curr. Biol. 2020, 30, 1231-1244.e4, doi:10.1016/j.cub.2020.01.065. 103. Stewart, E.J. Growing unculturable bacteria. J. Bacteriol. 2012, 194, 4151–4160, doi:10.1128/JB.00345-12. 104. Romano, S.; Jackson, S.A.; Patry, S.; Dobson, A.D.W. Extending the “one strain many compounds” (OSMAC) principle to marine microorganisms. Mar. Drugs 2018, 16, 1–29, doi:10.3390/md16070244. 105. Souza, C.P.; Almeida, B.C.; Colwell, R.R.; Rivera, I.N.G. The Importance of Chitin in the Marine Environment. Mar. Biotechnol. 2011, 13, 823–830, doi:10.1007/s10126-011- 9388-1. 106. Darabpour, E.; Roayaei Ardakani, M.; Motamedi, H.; Ronagh, M.T.; Najafzadeh, H. Purification and optimization of production conditions of a marine-derived antibiotic and ultra-structural study on the effect of this antibiotic against MRSA. Eur. Rev. Med. Pharmacol. Sci. 2012, 16, 157–165. 107. Marmann, A.; Aly, A.H.; Lin, W.; Wang, B.; Proksch, P. Co-cultivation - A powerful emerging tool for enhancing the chemical diversity of microorganisms. Mar. Drugs

2014, 12, 1043–1065, doi:10.3390/md12021043. 108. Seyedsayamdost, M.R. High-Throughput platform for the discovery of elicitors of silent bacterial gene clusters. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 7266–7271, doi:10.1073/pnas.1400019111. 109. Mitova, M.I.; Lang, G.; Wiese, J.; Imhoff, J.F. Subinhibitory concentrations of antibiotics induce phenazine production in a marine Streptomyces sp. J. Nat. Prod. 2008, 71, 824– 827, doi:10.1021/np800032a. 110. Okada, B.K.; Seyedsayamdost, M.R. Antibiotic dialogues: Induction of silent biosynthetic gene clusters by exogenous small molecules. FEMS Microbiol. Rev. 2017, 41, 19–33, doi:10.1093/femsre/fuw035. 111. Xu, F.; Nazari, B.; Moon, K.; Bushin, L.B.; Seyedsayamdost, M.R. Discovery of a Cryptic Antifungal Compound from Streptomyces albus J1074 Using High-Throughput Elicitor Screens. J. Am. Chem. Soc. 2017, 139, 9203–9212, doi:10.1021/jacs.7b02716. 112. Davies, J.; Spiegelman, G.B.; Yim, G. The world of subinhibitory antibiotic concentrations. Curr. Opin. Microbiol. 2006, 9, 445–453, doi:10.1016/j.mib.2006.08.006. 113. Moon, K.; Xu, F.; Seyedsayamdost, M.R. Cebulantin, a Cryptic Lanthipeptide Antibiotic Uncovered Using Bioactivity-Coupled HiTES. Angew. Chemie - Int. Ed. 2019, 58, 5973– 5977, doi:10.1002/anie.201901342. 114. Yamada, Y.; Kuzuyama, T.; Komatsu, M.; Shin-ya, K.; Omura, S.; Cane, D.E.; Ikeda, H. Terpene synthases are widely distributed in bacteria. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 857–862, doi:10.1073/pnas.1422108112. 115. Blin, K.; Shaw, S.; Steinke, K.; Villebro, R.; Ziemert, N.; Lee, S.Y.; Medema, M.H.; Weber, T. AntiSMASH 5.0: Updates to the secondary metabolite genome mining pipeline. Nucleic Acids Res. 2019, 47, W81–W87, doi:10.1093/nar/gkz310. 116. Medema, M.H.; Blin, K.; Cimermancic, P.; De Jager, V.; Zakrzewski, P.; Fischbach, M.A.; Weber, T.; Takano, E.; Breitling, R. AntiSMASH: Rapid identification, annotation and analysis of secondary metabolite biosynthesis gene clusters in bacterial and fungal genome sequences. Nucleic Acids Res. 2011, 39, 339–346, doi:10.1093/nar/gkr466. 117. Rocha-Martin, J.; Harrington, C.; Dobson, A.D.W.; O’Gara, F. Emerging strategies and integrated systems microbiology technologies for biodiscovery of marine bioactive compounds; 2014; Vol. 12; ISBN 3532142759. 118. Vikhnin, V.S.; Asatryan, H.R.; Zakharchenya, R.I.; Kutsenko, A.B.; Kapphan, S.E. Metagenomics: Application of Genomics to Uncultured Microorganisms. Microbiol. Mol. Biol. Rev. 2004, 68, 669–685, doi:10.1128/MBR.68.4.669. 119. Shokralla, S.; Spall, J.L.; Gibson, J.F.; Hajibabaei, M. Next-generation sequencing technologies for environmental DNA research. Mol. Ecol. 2012, 21, 1794–1805, doi:10.1111/j.1365-294X.2012.05538.x. 120. Brady, S.F.; Clardy, J. N-acyl derivatives of arginine and tryptophan isolated from environmental DNA expressed in Escherichia coli. Org. Lett. 2005, 7, 3613–3616,

doi:10.1021/ol0509585. 121. Youngblut, N.D.; Cuesta-zuluaga, J. De; Reischer, G.H. Large scale metagenome assembly reveals novel animal-associated microbial genomes, biosynthetic gene clusters, and other genetic diversity. 2020, 0–19. 122. Anderson, C.L.; Fernando, S.C. Insights into rumen microbial biosynthetic gene cluster diversity through genome-resolved metagenomics. bioRxiv 2020, doi:10.1101/2020.05.19.105130. 123. Ross, A.C.; Gulland, L.E.S.; Dorrestein, P.C.; Moore, B.S. Targeted capture and heterologous expression of the pseudoalteromonas alterochromide gene cluster in escherichia coli represents a promising natural product exploratory platform. ACS Synth. Biol. 2015, 4, 414–420, doi:10.1021/sb500280q. 124. Romanenko, L.A.; Zhukova, N. V.; Rohde, M.; Lysenko, A.M.; Mikhailov, V. V.; Stackebrandt, E. Pseudoalteromonas agarivorans sp. nov., a novel marine agarolytic bacterium. Int. J. Syst. Evol. Microbiol. 2003, 53, 125–131, doi:10.1099/ijs.0.02234-0. 125. Holmström, C.; Kjelleberg, S. Marine Pseudoalteromonas species are associated with higher organisms and produce biologically active extracellular agents. FEMS Microbiol. Ecol. 1999, 30, 285–293, doi:10.1016/S0168-6496(99)00063-X. 126. Holmström, C.; Steinberg, P.; Christov, V.; Christie, G.; Kjelleberg, S. Bacteria immobilised in gels: Improved methodologies for antifouling and biocontrol applications. Biofouling 2000, 15, 109–117, doi:10.1080/08927010009386302. 127. Sánchez-Porro, C.; Mellado, E.; Bertoldo, C.; Antranikian, G.; Ventosa, A. Screening and characterization of the protease CP1 produced by the moderately halophilic bacterium Pseudoalteromonas sp. strain CP76. Extremophiles 2003, 7, 221–228, doi:10.1007/s00792-003-0316-9. 128. Turkiewicz, M.; Kur, J.; Białkowska, A.; Cieśliński, H.; Kalinowska, H.; Bielecki, S. Antarctic marine bacterium Pseudoalteromonas sp. 22b as a source of cold-adapted β- galactosidase. Biomol. Eng. 2003, 20, 317–324, doi:10.1016/S1389-0344(03)00039-X. 129. Ivanova, E.P.; Flavier, S.; Christen, R. Phylogenetic relationships among marine Alteromonas-like proteobacteria: Emended description of the family Alteromonadaceae and proposal of Pseudoalteromonadaceae fam. nov., Colwelliaceae fam. nov., Shewanellaceae fam. nov., Moritellaceae fam. nov., Ferri. Int. J. Syst. Evol. Microbiol. 2004, 54, 1773–1788, doi:10.1099/ijs.0.02997-0. 130. Brian-Jaisson, F.; Ortalo-Magné, A.; Guentas-Dombrowsky, L.; Armougom, F.; Blache, Y.; Molmeret, M. Identification of Bacterial Strains Isolated from the Mediterranean Sea Exhibiting Different Abilities of Biofilm Formation. Microb. Ecol. 2014, 68, 94–110, doi:10.1007/s00248-013-0342-9. 131. Kim, E.H.; Cho, K.H.; Lee, Y.M.; Yim, J.H.; Lee, H.K.; Cho, J.C.; Hong, S.G. Diversity of cold-active protease-producing bacteria from arctic terrestrial and marine environments revealed by enrichment culture. J. Microbiol. 2010, 48, 426–432, doi:10.1007/s12275-010-0015-z. 132. Giudice, A. Lo; Caruso, C.; Mangano, S.; Bruni, V.; De Domenico, M.; Michaud, L. Marine

Bacterioplankton Diversity and Community Composition in an Antarctic Coastal Environment. Microb. Ecol. 2012, 63, 210–223, doi:10.1007/s00248-011-9904-x. 133. Ivanova, E.P.; Kiprianova, E.A.; Mikhailov, V. V.; Levanova, G.F.; Garagulya, A.D.; Gorshkova, N.M.; Yumoto, N.; Yoshikawa, S. Characterization and identification of marine Alteromonas nigrifaciens strains and emendation of the description. Int. J. Syst. Bacteriol. 1996, 46, 223–228, doi:10.1099/00207713-46-1-223. 134. Ivanova, E.P.; Kiprianova, E.A.; Mikhailov, V. V.; Levanova, G.F.; Garagulya, A.D.; Gorshkova, N.M.; Vysotskii, M. V.; Nicolau, D. V.; Yumoto, N.; Taguchi, T.; et al. Phenotypic diversity of Pseudoalteromonas citrea from different marine habitats and emendation of the description. Int. J. Syst. Bacteriol. 1998, 48, 247–256, doi:10.1099/00207713-48-1-247. 135. Holmstromfl, C.; James, S.; Neilan, B.A.; White, D.C. Pseudoalteromonas tunicata sp. nov,a Bacterium That Produces Antifouling Agents. Int. J. Syst. Bacteriol. 1998. 136. Wietz, M.; Gram, L.; Jørgensen, B.; Schramm, A. Latitudinal patterns in the abundance of major marine bacterioplankton groups. Aquat. Microb. Ecol. 2010, 61, 179–189, doi:10.3354/ame01443. 137. Skovhus, T.L.; Holmström, C.; Kjelleberg, S.; Dahllöf, I. Molecular investigation of the distribution, abundance and diversity of the genus Pseudoalteromonas in marine samples. FEMS Microbiol. Ecol. 2007, 61, 348–361, doi:10.1111/j.1574- 6941.2007.00339.x. 138. Vera, J.; Alvarez, R.; Murano, E.; Slebe, J.C.; Leon, O. Identification of a marine agarolytic Pseudoalteromonas isolate and characterization of its extracellular agarase. Appl. Environ. Microbiol. 1998, 64, 4378–4383, doi:10.1128/aem.64.11.4378- 4383.1998. 139. Ritchie, K.B.; Nagelkerken, I.; James, S.; Smith, G.W. A tetrodotoxin-producing marine pathogen. Nature 2000, 404, 354, doi:10.1038/35006168. 140. Holmström, C.; Egan, S.; Franks, A.; McCloy, S.; Kjelleberg, S. Antifouling activities expressed by marine surface associated Pseudoalteromonas species. FEMS Microbiol. Ecol. 2002, 41, 47–58, doi:10.1016/S0168-6496(02)00239-8. 141. Holmström, C.; James, S.; Neilan, B.A.; White, D.C.; Kjelleberg, S. Pseudoalteromonas tunicata sp. nov., a bacterium that produces antifouling agents. Int. J. Syst. Bacteriol. 1998, 48, 1205–1212, doi:10.1099/00207713-48-4-1205. 142. Cimermancic, P.; Medema, M.H.; Claesen, J.; Kurita, K.; Wieland Brown, L.C.; Mavrommatis, K.; Pati, A.; Godfrey, P.A.; Koehrsen, M.; Clardy, J.; et al. Insights into Secondary Metabolism from a Global Analysis of Prokaryotic Biosynthetic Gene Clusters. Cell 2014, 158, 412–421, doi:10.1016/j.cell.2014.06.034. 143. Zhou, M.Y.; Chen, X.L.; Zhao, H.L.; Dang, H.Y.; Luan, X.W.; Zhang, X.Y.; He, H.L.; Zhou, B.C.; Zhang, Y.Z. Diversity of both the cultivable protease-producing bacteria and their extracellular proteases in the sediments of the South China Sea. Microb. Ecol. 2009, 58, 582–590, doi:10.1007/s00248-009-9506-z. 144. Cotugno, R.; Rosaria Ruocco, M.; Marco, S.; Falasca, P.; Evangelista, G.; Raimo, G.;

Chambery, A.; Di Maro, A.; Masullo, M.; De Vendittis, E. Differential cold-adaptation among protein components of the thioredoxin system in the psychrophilic eubacterium Pseudoalteromonas haloplanktis TAC 125. Mol. Biosyst. 2009, 5, 519–528, doi:10.1039/b818467d. 145. Santiago, M.; Ramírez-Sarmiento, C.A.; Zamora, R.A.; Parra, L.P. Discovery, molecular mechanisms, and industrial applications of cold-active enzymes. Front. Microbiol. 2016, 7, doi:10.3389/fmicb.2016.01408. 146. Cabello, F.C. Heavy use of prophylactic antibiotics in aquaculture: A growing problem for human and animal health and for the environment. Environ. Microbiol. 2006, 8, 1137–1144, doi:10.1111/j.1462-2920.2006.01054.x. 147. FAO/WHO Guidelines for the Evaluation of Probiotics in Food. 2002, 1–11. 148. Wang, H.; Wang, C.; Tang, Y.; Sun, B.; Huang, J.; Song, X. Pseudoalteromonas probiotics as potential biocontrol agents improve the survival of Penaeus vannamei challenged with acute hepatopancreatic necrosis disease (AHPND)-causing Vibrio parahaemolyticus. Aquaculture 2018, 494, 30–36, doi:10.1016/j.aquaculture.2018.05.020. 149. Kumar, G.; Menanteau-Ledouble, S.; Saleh, M.; El-Matbouli, M. Yersinia ruckeri, the causative agent of enteric redmouth disease in fish. Vet. Res. 2015, 46, 1–10, doi:10.1186/s13567-015-0238-4. 150. Nelson, E.J.; Ghiorse, W.C. Isolation and identification of Pseudoalteromonas piscicida strain Cura-d associated with diseased damselfish (Pomacentridae) eggs. J. Fish Dis. 1999, 22, 253–260, doi:10.1046/j.1365-2761.1999.00168.x. 151. Skov, M.N.; Pedersen, K.; Larsen, J.L. Comparison of pulsed-field gel electrophoresis, ribotyping, and plasmid profiling for typing of Vibrio anguillarum serovar O1. Appl. Environ. Microbiol. 1995, 61, 1540–1545, doi:10.1128/aem.61.4.1540-1545.1995. 152. Isnansetyo, A.; Kamei, Y. MC21-A , a Bactericidal Antibiotic Produced by a New Marine against Methicillin-Resistant Staphylococcus aureus. Society 2003, 170, 481–490, doi:10.1128/AAC.47.2.480. 153. Isnansetyo, A.; Kamei, Y. Anti-methicillin-resistant Staphylococcus aureus (MRSA) activity of MC21-B, an antibacterial compound produced by the marine bacterium Pseudoalteromonas phenolica O-BC30T. Int. J. Antimicrob. Agents 2009, 34, 131–135, doi:10.1016/j.ijantimicag.2009.02.009. 154. And, N.L. 1; Fenical, W. New tambjamine class alkaloids from the marine ascidian Atapozoa sp. and its nudibranch predators. Origin of the tambjamines in Atapozoa. Experientia 1991, 47, 504–506, doi:10.4324/9781315657455-44. 155. Holmstromfl, C.; James, S.; Neilan, B.A.; White, D.C. a Bacterium That Produces Antifouling Agents. Int. J. Syst. Bacteriol. 1998. 156. OKAMOTO, H.; SATO, Z.; SATO, M.; KOISO, Y.; IWASAKI, S.; ISAKA, M. Identification of Antibiotic Red Pigments of Serratia marcescens F-1-1, a Biocontrol Agent of Damping- off of Cucumber, and Antimicrobial Activity against Other Plant Pathogens. Japanese J. Phytopathol. 1998, 64, 294–298, doi:10.3186/jjphytopath.64.294.

157. Kim, H.S.; Hayashi, M.; Shibata, Y.; Wataya, Y.; Mitamura, T.; Horii, T.; Kawauchi, K.; Hirata, H.; Tsuboi, S.; Moriyama, Y. Cycloprodigiosin hydrochloride obtained from Pseudoalteromonas denitrificans is a potent antimalarial agent. Biol. Pharm. Bull. 1999, 22, 532–534, doi:10.1248/bpb.22.532. 158. Kawauchi, K.; Tobiume, K.; Iwashita, K.; Inagaki, H.; Morikawa, T.; Shibukawa, Y.; Moriyama, Y.; Hirata, H.; Kamata, H. Cycloprodigiosin hydrochloride activates the Ras- PI3K-Akt pathway and suppresses protein synthesis inhibition-induced apoptosis in PC12 cells. Biosci. Biotechnol. Biochem. 2008, 72, 1564–1570, doi:10.1271/bbb.80064. 159. Longeon, A.; Peduzzi, J.; Barthélemy, M.; Corre, S.; Nicolas, J.L.; Guyot, M. Purification and partial identification of novel antimicrobial protein from marine bacterium Pseudoalteromonas species strain X153. Mar. Biotechnol. 2004, 6, 633–641, doi:10.1007/s10126-004-3009-1. 160. Kaya, M.; Lelešius, E.; Nagrockaite, R.; Sargin, I.; Arslan, G.; Mol, A.; Baran, T.; Can, E.; Bitim, B. Differentiations of Chitin content and surface morphologies of chitins extracted from male and female grasshopper species. PLoS One 2015, 10, doi:10.1371/journal.pone.0115531. 161. Park, B.K.; Kim, M.M. Applications of chitin and its derivatives in biological medicine. Int. J. Mol. Sci. 2010, 11, 5152–5164, doi:10.3390/ijms11125152. 162. Junges, Â.; Boldo, J.T.; Souza, B.K.; Guedes, R.L.M.; Sbaraini, N.; Kmetzsch, L.; Thompson, C.E.; Staats, C.C.; De Almeida, L.G.P.; De Vasconcelos, A.T.R.; et al. Genomic analyses and transcriptional profiles of the glycoside hydrolase family 18 genes of the entomopathogenic fungus Metarhizium anisopliae. PLoS One 2014, 9, doi:10.1371/journal.pone.0107864. 163. Merzendorfer, H. The cellular basis of chitin synthesis in fungi and insects: Common principles and differences. Eur. J. Cell Biol. 2011, 90, 759–769, doi:10.1016/j.ejcb.2011.04.014. 164. Gooday, G.W. Physiology of microbial degradation of chitin and chitosan. Biodegradation 1990, 1, 177–190, doi:10.1007/BF00058835. 165. Reay, R.C. Chitin in nature and technology; 1989; Vol. 25; ISBN 9781461292777. 166. Keyhani, N.O.; Roseman, S. Physiological aspects of chitin catabolism in marine bacteria. Biochim. Biophys. Acta - Gen. Subj. 1999, 1473, 108–122, doi:10.1016/S0304- 4165(99)00172-5. 167. Jagadish C. Roy, Fabien Salaün, S.G.; Ada Ferri, G.C. and J.G.; Ada Ferri, G.C. and J.G. Solubility of Chitin: Solvents, Solution Behaviors and Their Related Mechanisms. Intech 2016, 13. 168. Palanivel Rameshthangam1,*, Dhanasekaran Solairaj1, Gnanapragasam Arunachalam2, P.R. Chitin and Chitinases: Biomedical And Environmental Applications of Chitin and its Derivatives Palanivel. J. Enzym. 2018, 1–7, doi:10.14302/issn.2690. 169. Lombard, V.; Golaconda Ramulu, H.; Drula, E.; Coutinho, P.M.; Henrissat, B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 2014, 42, 490–495, doi:10.1093/nar/gkt1178.

170. Monge, E.C.; Tuveng, T.R.; Vaaje-Kolstad, G.; Eijsink, V.G.H.; Gardner, J.G. Systems analysis of the glycoside hydrolase family 18 enzymes from Cellvibrio japonicus characterizes essential chitin degradation functions. J. Biol. Chem. 2018, 293, 3849– 3859, doi:10.1074/jbc.RA117.000849. 171. Meibom, K.L.; Li, X.B.; Nielsen, A.T.; Wu, C.-Y.; Roseman, S.; Schoolnik, G.K. The Vibrio cholerae chitin utilization program. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 2524–9. 172. Umemoto, N.; Kanda, Y.; Ohnuma, T.; Osawa, T.; Numata, T.; Sakuda, S.; Taira, T.; Fukamizo, T. Crystal structures and inhibitor binding properties of plant class v chitinases: The cycad enzyme exhibits unique structural and functional features. Plant J. 2015, 82, 54–66, doi:10.1111/tpj.12785. 173. Kezuka, Y.; Ohishi, M.; Itoh, Y.; Watanabe, J.; Mitsutomi, M.; Watanabe, T.; Nonaka, T. Structural Studies of a Two-domain Chitinase from Streptomyces griseus HUT6037. J. Mol. Biol. 2006, 358, 472–484, doi:10.1016/j.jmb.2006.02.013. 174. Stoykov, Y.M.; Pavlov, A.I.; Krastanov, A.I. Chitinase biotechnology: Production, purification, and application. Eng. Life Sci. 2015, 15, 30–38, doi:10.1002/elsc.201400173. 175. Ohno, T.; Armand, S.; Hata, T.; Nikaidou, N.; Henrissat, B.; Mitsutomi, M.; Watanabe, T. A modular family 19 chitinase found in the prokaryotic organism Streptomyces griseus HUT 6037. J. Bacteriol. 1996, 178, 5065–5070, doi:10.1128/jb.178.17.5065-5070.1996. 176. García-Fraga, B.; da Silva, A.F.; López-Seijas, J.; Sieiro, C. A novel family 19 chitinase from the marine-derived Pseudoalteromonas tunicata CCUG 44952T: Heterologous expression, characterization and antifungal activity. Biochem. Eng. J. 2015, 93, 84–93, doi:10.1016/j.bej.2014.09.014. 177. DeBoy, R.T.; Mongodin, E.F.; Fouts, D.E.; Tailford, L.E.; Khouri, H.; Emerson, J.B.; Mohamoud, Y.; Watkins, K.; Henrissat, B.; Gilbert, H.J.; et al. Insights into plant cell wall degradation from the genome sequence of the soil bacterium Cellvibrio japonicus. J. Bacteriol. 2008, 190, 5455–5463, doi:10.1128/JB.01701-07. 178. Tuveng, T.R.; Arntzen, M.Ø.; Bengtsson, O.; Gardner, J.G.; Vaaje-Kolstad, G.; Eijsink, V.G.H. Proteomic investigation of the secretome of Cellvibrio japonicus during growth on chitin. Proteomics 2016, 16, 1904–1914, doi:10.1002/pmic.201500419. 179. Singh, S.; Scigelova, M.; Crout, D.H.G. Glycosidase-catalysed synthesis of α-galactosyl epitopes important in xenotransplantation and toxin binding using the α-galactosidase from Penicillium multicolor. Chem. Commun. 1999, 2065–2066, doi:10.1039/a906020k. 180. Yu, C.; Lee, A.M.; Basslers, B.L. Chitin Utilization by Marine Bacteria. J. Biol. Chem. 1991, 266, 24260–24267. 181. LeCleir, G.R.; Buchan, A.; Maurer, J.; Moran, M.A.; Hollibaugh, J.T. Comparison of chitinolytic enzymes from an alkaline, hypersaline lake and an estuary. Environ. Microbiol. 2007, 9, 197–205, doi:10.1111/j.1462-2920.2006.01128.x. 182. Hara, S.; Yamamura, Y.; Fujii, Y.; Mega, T.; Ikenaka, T. Purification and characterization of chitinase produced by Streptomyces erythraeus. J. Biochem. 1989, 105, 484–489, doi:10.1093/oxfordjournals.jbchem.a122691.

183. Bissaro, B.; Isaksen, I.; Vaaje-Kolstad, G.; Eijsink, V.G.H.; Røhr, Å.K. How a Lytic Polysaccharide Monooxygenase Binds Crystalline Chitin. Biochemistry 2018, 57, 1893– 1906, doi:10.1021/acs.biochem.8b00138. 184. Vaaje-Kolstad, G.; Forsberg, Z.; Loose, J.S.; Bissaro, B.; Eijsink, V.G. Structural diversity of lytic polysaccharide monooxygenases. Curr. Opin. Struct. Biol. 2017, 44, 67–76, doi:10.1016/j.sbi.2016.12.012. 185. Sani, R.K.; Krishnaraj, R.N. Extremophilic enzymatic processing of lignocellulosic feedstocks to bioenergy. Extrem. Enzym. Process. Lignocellul. Feed. to Bioenergy 2017, 1–308, doi:10.1007/978-3-319-54684-1. 186. Forsberg, Z.; Vaaje-kolstad, G.; Westereng, B.; Bunsæ, A.C.; Stenstrøm, Y.; Mackenzie, A.; Sørlie, M.; Horn, S.J.; Eijsink, V.G.H. Cleavage of cellulose by a cbm33 protein. Protein Sci. 2011, 20, 1479–1483, doi:10.1002/pro.689. 187. Hemsworth, G.R.; Johnston, E.M.; Davies, G.J.; Walton, P.H. Lytic Polysaccharide Monooxygenases in Biomass Conversion. Trends Biotechnol. 2015, 33, 747–761, doi:10.1016/j.tibtech.2015.09.006. 188. Hamre, A.G.; Eide, K.B.; Wold, H.H.; Sørlie, M. Activation of enzymatic chitin degradation by a lytic polysaccharide monooxygenase. Carbohydr. Res. 2015, 407, 166– 169, doi:10.1016/j.carres.2015.02.010. 189. Vaaje-Kolstad, G.; Horn, S.J.; Van Aalten, D.M.F.; Synstad, B.; Eijsink, V.G.H. The non- catalytic chitin-binding protein CBP21 from Serratia marcescens is essential for chitin degradation. J. Biol. Chem. 2005, 280, 28492–28497, doi:10.1074/jbc.M504468200. 190. A. Veliz, E.; Martínez-Hidalgo, P.; M. Hirsch, A. Chitinase-producing bacteria and their role in biocontrol. AIMS Microbiol. 2017, 3, 689–705, doi:10.3934/microbiol.2017.3.689. 191. Nazari, B.; Kobayashi, M.; Saito, A.; Hassaninasab, A.; Miyashita, K.; Fujiia, T. Chitin- induced gene expression in secondary metabolic pathways of Streptomyces coelicolor A3(2) grown in soil. Appl. Environ. Microbiol. 2013, 79, 707–713, doi:10.1128/AEM.02217-12. 192. Li, X.; Roseman, S. The chitinolytic cascade in Vibrios is regulated by chitin oligosaccharides and a two-component chitin catabolic sensor/kinase. Proc. Natl. Acad. Sci. 2004, 101, 627–631, doi:10.1073/pnas.0307645100. 193. Wietz, M.; Månsson, M.; Gram, L. Chitin stimulates production of the antibiotic andrimid in a Vibrio coralliilyticus strain. Environ. Microbiol. Rep. 2011, 3, 559–564, doi:10.1111/j.1758-2229.2011.00259.x. 194. Song, C.W.; Lee, J.; Lee, S.Y. Genome engineering and gene expression control for bacterial strain development. Biotechnol. J. 2015, 10, 56–68, doi:10.1002/biot.201400057. 195. Zeaiter, Z.; Mapelli, F.; Crotti, E.; Borin, S. Methods for the genetic manipulation of marine bacteria. Electron. J. Biotechnol. 2018, 33, 17–28, doi:10.1016/j.ejbt.2018.03.003.

196. Kirchberger, P.C.; Orata, F.D.; Barlow, E.J.; Kauffman, K.M.; Case, R.J.; Polz, M.F.; Boucher, Y. A small number of phylogenetically distinct clonal complexes dominate a coastal Vibrio cholerae population. Appl. Environ. Microbiol. 2016, 82, 5576–5586, doi:10.1128/AEM.01177-16. 197. Blokesch, M. Competence-induced type VI secretion might foster intestinal colonization by Vibrio cholerae: Intestinal interbacterial killing by competence-induced V. cholerae M. Blokesch. BioEssays 2015, 37, 1163–1168, doi:10.1002/bies.201500101. 198. Meibom, K.L.; Blokesch, M.; Dolganov, N.A.; Wu, C.Y.; Schoolnik, G.K. Microbiology: Chitin induces natural competence in vibrio cholerae. Science (80-. ). 2005, 310, 1824– 1827, doi:10.1126/science.1120096. 199. Pollack-Berti, A.; Wollenberg, M.S.; Ruby, E.G. Natural transformation of Vibrio fischeri requires tfoX and tfoY. Environ. Microbiol. 2010, 12, 2302–2311, doi:10.1111/j.1462- 2920.2010.02250.x. 200. Gulig, P.A.; Tucker, M.S.; Thiaville, P.C.; Joseph, J.L.; Brown, R.N. USER friendly cloning coupled with chitin-based natural transformation enables rapid mutagenesis of Vibrio vulnificus. Appl. Environ. Microbiol. 2009, 75, 4936–4949, doi:10.1128/AEM.02564-08. 201. Weinstock, M.T.; Hesek, E.D.; Wilson, C.M.; Gibson, D.G. Vibrio natriegens as a fast- growing host for molecular biology. Nat. Methods 2016, 13, 849–851, doi:10.1038/nmeth.3970. 202. Chen, Y.; Dai, J.; Morris, J.G.; Johnson, J.A. Genetic analysis of the capsule polysaccharide (K antigen) and exopolysaccharide genes in pandemic Vibrio parahaemolyticus O3:K6. BMC Microbiol. 2010, 10, 274, doi:10.1186/1471-2180-10- 274. 203. Udden, S.M.N.; Zahid, M.S.H.; Biswas, K.; Ahmad, Q.S.; Cravioto, A.; Nair, G.B.; Mekalanos, J.J.; Faruque, S.M. Acquisition of classical CTX prophage from Vibrio cholerae O141 by El Tor strains aided by lytic phages and chitin-induced competence. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 11951–11956, doi:10.1073/pnas.0805560105. 204. Blokesch, M.; Schoolnik, G.K. The extracellular nuclease Dns and its role in natural transformation of Vibrio cholerae. J. Bacteriol. 2008, 190, 7232–7240, doi:10.1128/JB.00959-08. 205. Marcus, H.; Ketley, J.M.; Kaper, J.B.; Holmes, R.K. Effects of DNase production, plasmid size, and restriction barriers on transformation of Vibrio cholerae by electroporation and osmotic shock. FEMS Microbiol. Lett. 1990, 68, 149–154, doi:10.1016/0378- 1097(90)90141-C. 206. Wang, H.; Griffiths, M.W. Mg2+-free buffer elevates transformation efficiency of Vibrio parahaemolyticus by electroporation. Lett. Appl. Microbiol. 2009, 48, 349–354, doi:10.1111/j.1472-765X.2008.02531.x. 207. Piekarski, T.; Buchholz, I.; Drepper, T.; Schobert, M.; Wagner-Doebler, I.; Tielen, P.; Jahn, D. Genetic tools for the investigation of Roseobacter clade bacteria. BMC Microbiol. 2009, 9, doi:10.1186/1471-2180-9-265. 208. Kurusu, Y.; Yoshimura, S.; Tanaka, M.; Nakamura, T.; Maruyama, A.; Higashihara, T.

Genetic transformation system for a psychrotrophic deep-sea bacterium: Isolation and characterization of a psychrotrophic plasmid. Mar. Biotechnol. 2001, 3, 96–99, doi:10.1007/s101260000067. 209. Klevanskaa, K.; Bier, N.; Stingl, K.; Strauch, E.; Hertwig, S. PVv3, a new shuttle vector for gene expression in Vibrio vulnificus. Appl. Environ. Microbiol. 2014, 80, 1477–1481, doi:10.1128/AEM.03720-13. 210. Gilchrist, A.; Smit, J. Transformation of freshwater and marine caulobacters by electroporation. J. Bacteriol. 1991, 173, 921–925, doi:10.1128/jb.173.2.921-925.1991. 211. Altermark, B.; Niiranen, L.; Willassen, N.P.; Smalås, A.O.; Moe, E. Comparative studies of endonuclease I from cold-adapted Vibrio salmonicida and mesophilic Vibrio cholerae. FEBS J. 2007, 274, 252–263, doi:10.1111/j.1742-4658.2006.05580.x. 212. Delavat, F.; Bidault, A.; Pichereau, V.; Paillard, C. Rapid and efficient protocol to introduce exogenous DNA in Vibrio harveyi and Pseudoalteromonas sp. J. Microbiol. Methods 2018, 154, 1–5, doi:10.1016/j.mimet.2018.09.022. 213. Le Roux, F.; Binesse, J.; Saulnier, D.; Mazel, D. Construction of a Vibrio splendidus mutant lacking the metalloprotease gene vsm by use of a novel counterselectable suicide vector. Appl. Environ. Microbiol. 2007, 73, 777–784, doi:10.1128/AEM.02147- 06. 214. Smillie, C.; Garcillán-Barcia, M.P.; Francia, M.V.; Rocha, E.P.C.; de la Cruz, F. Mobility of Plasmids. Microbiol. Mol. Biol. Rev. 2010, 74, 434–452, doi:10.1128/mmbr.00020-10. 215. Lambs, L.; Venturim, M.; Révérend, B.D. Le; Kozlowski, H.; Berthon, G. Metal ion- tetracycline interactions in biological fluids. Part 8. Potentiometric and spectroscopic studies on the formation of Ca(II) and Mg(II) complexes with 4-dedimethylamino- tetracycline and 6-desoxy-6-dem. J. Inorg. Biochem. 1988, 33, 193–209, doi:10.1016/0162-0134(88)80049-7. 216. Donnenberg, M.S.; Kaper, J.B. Construction of an eae deletion mutant of enteropathogenic Escherichia coli by using a positive-selection suicide vector. Infect. Immun. 1991, 59, 4310–4317, doi:10.1128/iai.59.12.4310-4317.1991. 217. Milton, D.L.; O’Toole, R.; Hörstedt, P.; Wolf-Watz, H. Flagellin A is essential for the virulence of Vibrio anguillarum. J. Bacteriol. 1996, 178, 1310–1319, doi:10.1128/jb.178.5.1310-1319.1996. 218. Karlsson, S.L.; Ax, E.; Nygren, E.; Källgård, S.; Blomquist, M.; Ekman, A.; Benktander, J.; Holmgren, J.; Lebens, M. Development of stable vibrio cholerae O1 Hikojima type vaccine strains co-expressing the Inaba and Ogawa lipopolysaccharide antigens. PLoS One 2014, 9, doi:10.1371/journal.pone.0108521. 219. Yu, Z.C.; Zhao, D.L.; Ran, L.Y.; Mi, Z.H.; Wu, Z.Y.; Pang, X.; Zhang, X.Y.; Su, H.N.; Shi, M.; Song, X.Y.; et al. Development of a genetic system for the deep-sea psychrophilic bacterium Pseudoalteromonas sp. SM9913. Microb. Cell Fact. 2014, 13, 1–9, doi:10.1186/1475-2859-13-13. 220. Blomfield, I.C.; Vaughn, V.; Rest, R.F.; Eisenstein, B.I. Allelic exchange in Escherichia coli using the Bacillus subtilis sacB gene and a temperature‐sensitive pSC101 replicon. Mol.

Microbiol. 1991, 5, 1447–1457, doi:10.1111/j.1365-2958.1991.tb00791.x. 221. Luo, P.; He, X.; Liu, Q.; Hu, C. Developing universal genetic tools for rapid and efficient deletion mutation in vibrio species based on suicide t-vectors carrying a novel counterselectable marker, vmi480. PLoS One 2015, 10, 1–17, doi:10.1371/journal.pone.0144465. 222. Sawabe, T.; Fukui, Y.; Stabb, E. V. Simple conjugation and outgrowth procedures for tagging vibrios with GFP, and factors affecting the stable expression of the gfp tag. Lett. Appl. Microbiol. 2006, 43, 514–522, doi:10.1111/j.1472-765X.2006.01992.x. 223. Ide, T.; Hoya, M.; Tanaka, T.; Harayama, S. Enhanced production of astaxanthin in Paracoccus sp. strain N-81106 by using random mutagenesis and genetic engineering. Biochem. Eng. J. 2012, 65, 37–43, doi:10.1016/j.bej.2012.03.015. 224. Tanimura, K.; Nakayama, H.; Tanaka, T.; Kondo, A. Ectoine production from lignocellulosic biomass-derived sugars by engineered Halomonas elongata. Bioresour. Technol. 2013, 142, 523–529, doi:10.1016/j.biortech.2013.05.004. 225. Wang, H.; Li, Z.; Jia, R.; Hou, Y.; Yin, J.; Bian, X.; Li, A.; Müller, R.; Stewart, A.F.; Fu, J.; et al. RecET direct cloning and Redαβ recombineering of biosynthetic gene clusters, large operons or single genes for heterologous expression. Nat. Protoc. 2016, 11, 1175–1190, doi:10.1038/nprot.2016.054.

9. RESEARCH ARTICLES

Paper 1: Chitin degradation machinery and secondary metabolite profiles in the marine bacterium Pseudoalteromonas rubra S4059.

Manuscript 2: Identification and verification of the prodigiosin biosynthetic gene cluster (BGC) in Pseudoalteromonas rubra S4059.

Manuscript 3: Attachment to and growth on chitin surfaces results in loss of prodigiosin production in a pigmented Pseudoalteromonas rubra S4059. Note: Due to the limitation time of 3 years PhD research, the works on manuscript 3 have not been completely finished and some of perspectives/ideas were included in the discussion.

Paper 1

Chitin degradation machinery and secondary metabolite profiles in the marine bacterium Pseudoalteromonas rubra S4059

Article Chitin degradation machinery and secondary metabolite profiles in the marine bacterium Pseudoalteromonas rubra S4059

Xiyan Wang 1, Thomas Isbrandt 1, Mikael Lenz Strube 1, Sara Skøtt Paulsen 1, Maike Wennekers Nielsen 1, Yannick Buijs 1, Erwin M. Schoof 1, Thomas Ostenfeld Larsen 1, Lone Gram 1 and Sheng- Da Zhang 1 *

1 Department of Bioengineering, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark; [email protected] (X.W.); [email protected] (T.I.); [email protected] (M.L.S.); [email protected] (S.S.P.); [email protected] (M.W.N.); [email protected] (E.M.S.); [email protected] (Y.B.); [email protected] (T.O.L); [email protected] (L.G.). * Correspondence: [email protected] (S.Z.); Tel.: +45-5011-7765.

Received:22 December 2020; Accepted: 04 February 2021; Published:

Abstract: Genome mining of pigmented Pseudoalteromonas has revealed a large potential for production of bioactive compounds and hydrolytic enzymes. The purpose of the present study was to explore this bioactivity potential in a potent antibiotic and enzyme producer, Pseudoalteromonas rubra strain S4059. Proteomic analyses (data are available via ProteomeXchange with identifier PXD023249) indicated that a highly efficient chitin degradation machinery was present in the red- pigmented P. rubra S4059 when grown on chitin. Four GH18 chitinases and two GH20 hexosaminidases were significantly upregulated under these conditions. GH19 chitinases, which are not common in bacteria are consistently found in pigmented Pseudoalteromonas and in S4059 GH19 was only detected when the bacterium was grown on chitin. To explore the possible role of GH19 in pigmented Pseudoalteromonas, we developed a protocol for genetic manipulation of S4059 and deleted the GH19 chitinase and compared phenotypes of the mutant and wild type. However, none of the chitin degrading ability, secondary metabolite profile or biofilm forming capacity was affected by GH19 deletion. In conclusion, we developed a genetic manipulation protocol that can be used to unravel the bioactive potential of pigmented pseudoalteromonads. An efficient chitinolytic enzyme cocktail was identified in S4059, suggesting that this strain could be a candidate with industrial potential.

Keywords: chitin; chitinase; chitin degradation machinery; Pseudoalteromonas; secondary metabolites

1. Introduction Marine microorganisms have emerged as a promising source of novel antimicrobial compounds or hydrolytic enzymes [1,2]. In particular, the marine genus Pseudoalteromonas harbors a wide range of bioactive compounds with antimicrobial, antifouling, and algicidal activities [3–6]. Based on both phenotypes and genome-wide analyses, Pseudoalteromonas can be divided into two groups: pigmented and non-pigmented species [3]. The genomes of pigmented species contain a high number of biosynthetic gene clusters (BGCs) as compared to those of non-pigmented species [7]. Both groups carry the genetic potential to produce a wide range of glycosyl hydrolases, and especially the pigmented Pseudoalteromonas harbors a powerful chitin degrading machinery containing several chitinolytic enzymes [7]. Chitin is the most abundant carbon source in the marine environment [8], where it is present in three crystalline allomorphs: α-, β-, and γ-chitin. α-chitin has antiparallel chains, β-chitin has parallel chains and γ-chitin has the mixture of the both chains [9]. Chitin in nature is predominantly degraded

by microorganisms [10] and chitin degradation depends on secreted extracellular chitinases (EC.3.2.1.14) and other chitinolytic enzymes/proteins, such as lytic polysaccharide monooxygenases (LPMOs) [11]. Chitinases are glycoside hydrolases (GHs) and are classified into GH18, GH19 and GH20 in the CAZy database [12,13]. The GH18 family chitinases are common in bacteria, whereas the GH19 chitinases are mostly found in plants and are believed to function as defense mechanism against fungal pathogens [14–16]. Chitinases of both families catalyze the degradation of chitin polymers [12]. GH 20 family β-N-acetylhexosaminidases hydrolyze amorphous chitin polymers [17] and the LPMOs are metalloenzymes that cleave glycosidic bonds in crystalline chitin and facilitate access of chitinase [18]. Paulsen et al. [7] found that pigmented and nonpigmented Pseudoalteromonas evolved divergent GH profiles in their genomic contents. Further, all pigmented Pseudoalteromonas species contain at least one GH19 chitinase, which is rarely reported in bacteria, however, only very few non-pigmented Pseudoalteromonase species contain a GH19 chitinase [7]. A GH19 chitinase of the pigmented Pseudoalteromonas tunicata CCUG 44952T has been heterologously expressed in Escherichia coli and displayed antifungal activity [19], however, whether this is the dominant role of GH19 in pigmented Pseodoalteromonas is yet to be investigated. The secondary metabolome of several bacteria is influenced by carbon-source and chitin may serve to enhance production of secondary metabolites as observed in strains of Vibrionaceae [20–22]. Likewise, the addition of chitin to Streptomyces coelicolor A3 (2) growing in autoclaved soil induced the expression of genes associated with secondary metabolites production [23]. Due to the potent chitinolytic machinery in Pseudoalteromonas [7], we speculated that there could be a link between chitin degradation and secondary metabolism. Since P. rubra S4059 dedicates 15% of its genome to secondary metabolites [7] and, as other pigmented pseudoalteromonads, contain GH19 chitinases [7], we further explored the bioactivity of this prodigiosin-producing strain as a model organism. The purpose of this study was to explore the chitin degradation machinery and secondary metabolite profiles when grown on chitin and to investigate the possible function of GH19 chitinase in P. rubra S4059.

2. Results

2.1. In silico analysis of chitin degrading machinery and bioactive potential in P. rubra S4059 The genome of S4059 consists of two circular chromosomes, of 4,595,233 bp and 1,348,119 bp, with an average G+C content of 47.71% and 46.93%. Fourteen putative chitinolytic enzymes were identified according to the prediction of the S4059 proteome in Uniprot (proteome ID UP000305729) including seven of the GH18 chitinase family, two of the GH19 chitinase family, and three of the GH20 hexosaminidase family, as well as two lytic polysaccharide monooxygenases (LPMOs) of the auxiliary activity (AA) family 10 (Table 1). All the chitinolytic enzymes contained a signal peptide at the N- terminal, except for one GH19 chitinase (A0A5S3UPT5). The genome of S4059 harbors nineteen predicted BGCs identified by antiSMASH 6.0, distributed with thirteen on chr I and six on chr II. BGC 2-5, 17, and 18 are non-ribosomal peptide synthetase clusters (NRPs), BGC 8, 13, and 16 are other unspecified ribosomally synthesised and post- translationally modified peptide products (RiPP) cluster, BGC 10, 11 and 15 are the hybrid of NRPs and Type I polyketide (PKS), BGC 7 is a hserlactone BGC, BGC 9 is prodigiosin BGC, and BGC 12, 13 belong to lanthipepride-class. Four of the BGCs located on chr I were predicted to produce indigoidine, kalimantacin A, amonabactin P 750 and prodigiosins (Table S1), however, only prodigiosin gene cluster was predicted to be conserved to the known BGC with a similarity score of 70%, while the others were below 40% as shown in Table S1. The products of the rest of the BGCs are not known.

Table 1. The chitinolytic machinery in Pseudoalteromonas rubra S4059 according to the prediction of proteome from Uniprot (Proteome ID UP000305729). The signal peptide was predicted using amino acid sequence by SignalIP-5.0 (http://www.cbs.dtu.dk/services/SignalP/).

Glycoside hydrolase type Accession Signal peptide

GH18 A0A5S3USE2 Y

A0A5S3V351 Y

A0A5S3V6T3 Y

A0A5S3V0U4 Y

A0A5S3USH6 Y

A0A5S3V3K3 Y

A0A5S3V378 Y

GH19 A0A5S3UX38 Y

A0A5S3UPT5 N

GH20 A0A5S3UX95 Y

A0A5S3UV09 Y

A0A5S3V1X9 Y

LPMO A0A5S3UTD1 Y

A0A5S3V4S2 Y

2.2. Global proteome profiles of P. rubra S4059 grown on chitin The proteomes of P. rubra S4059 grown on different carbon sources were analyzed by a liquid chromatography tandem mass spectrometry (LC-MS/MS)-based label-free quantitative proteomics approach. The analyses were done on both cells and culture supernatant (excreted proteins) of S4059 grown in crystalline chitin or mannose containing medium. A total of 2,813 proteins were identified in the global proteome of S4059, of which 738 and 142 proteins were up- and down-regulated in S4059 culture supernatant when grown on crystalline chitin as compared to growth on mannose (Figure 1A). Simultaneously, 1,861 and 141 proteins were up- and down regulated in S4059 cells when grown on crystalline chitin as compared to on mannose (Figure 1B). Proteins involved in bacterial chemotaxis, cell division, flagellar assembly and Type IV pilus assembly were upregulated when S4059 was grown on crystalline chitin (Table S3). Proteins associated with the core metabolism were also upregulated on chitin (Table S3). A protein involved in mannosidase from GH92 family was upregulated when growth on mannose as compared to when grown on crystalline chitin (Table S2).

Figure 1. Comparison of identified proteins in supernatant (A) and cells (B) of Pseudoalteromonas rubra S4059 when grown on chitin compared to on mannose. The dotted lines indicated that a threshold value for the cut off is a combination of p-value ≤ 0.05 and Log2 fold-change ≥ 1.5. Red dots represent upregulated proteins and blue dots represent downregulated proteins. Chitinolytic enzymes with significant fold changes were highlighted by enlarged dots with protein names in (A) and (B). Heat map comparison of identified chitinolytic enzymes in the supernatant and cells of Pseudoalteromonas rubra S4059 color-coded by increasing abundance (C). C-WT-cell: cells of S4059 grown on chitin; M-WT-cell: cells of S4059 grown on mannose; C-WT-sup: culture supernatant of S4059 grown on chitin; M-WT-sup: culture supernatant of S4059 grown on mannose. White denotes proteins not detected under this condition. ‘Overlap’ indicates these proteins were detected in both chitin and mannose-grown samples and asterisk highlights proteins significantly upregulated when grown on chitin compared to on mannose.

2.3. Comparative analysis of the expression of chitinolytic enzymes in P. rubra S4059 A total of twelve chitin utilization associated proteins, including seven GH18s, three GH20s, a GH19 and a LPMO, were identified with a confidence rate of 99% at the peptide and protein levels, in the P. rubra S4059 global proteome while a GH19 A0A5S3UPT5 and a LPMO A0A5S3V4S2 were not detected (Figure 1C). All chitinolytic enzymes could be detected in both cells and culture supernatant of S4059, except the enzyme A0A5S3UV09, from the GH20 family, that was not detected in the culture supernatant (Figure 1C). The GH18 chitinase A0A5S3USH6, annotated as ChiC, was the most highly expressed chitinolytic enzyme in both cells and culture supernatant during growth on chitin. Four GH18s

(A0A5S3USE2, A0A5S3V6T3 A0A5S3V3K3, and A0A5S3USH6) and two GH20s (A0A5S3V1X9 A0A5S3UX95) enzymes were significantly upregulated when S4059 was grown on crystalline chitin as compared to on mannose (Figure 1). Meanwhile, several chitin-utilization associated proteins were induced and could only be detected when grown on crystalline chitin, including a GH19 chitinase and three GH18 chitinases (A0A5S3V351, A0A5S3V0U4 and A0A5S3V378) (Figure 1 C). In addition, a LPMO from the AA10 family was identified under both growth conditions, but with no significant expression difference (Figure 1C).

2.4. Influence of chitin on the metabolome of S4059 To investigate the potential link between chitin degradation and secondary metabolite production, the S4059 wild type was grown on mannose, crystalline chitin, colloidal chitin, or N-acetyl glucosamine (NAG) containing medium in stationary phase. Chemical analysis using high-performance liquid chromatography coupled to diode array detection and high resolution mass spectrometry (HPLC- DAD-HRMS), showed that mannose and crystalline chitin resulted in largely the same chemical profiles, although two unknown tentative prodigiosin analogues were produced in higher amounts on mannose (Figure S1). NAG containing media resulted in an overall higher production of secondary metabolites compared to both mannose, crystalline and colloidal chitin, and NAG and colloidal chitin both led to increased production of two unknown non-prodigiosin derived (based on UV-Vis absorption) compounds (Figure S1). Prodigiosin, hexyl prodigiosin, and heptyl prodigiosin (Figure S1 B-D) where produced in varying amounts on all media. The identity of prodigiosin was confirmed using an authentic standard in combination with our in-house MS/MS library, and the two analogues were identified based on similarities with MS/MS and UV-Vis absorption spectra. Additionally, using our in-house MS/MS library, combined with a compound list generated from all Pseoudoalteromonas- derived secondary metabolites found in the Reaxys database, no other known secondary metabolites were identified in S4059.

2.5. The deletion of GH19 chitinase does not affect growth or chitin degradation To explore the function of GH19 chitinase in pigmented P. rubra S4059 and investigate a possible link between chitin degradation and secondary metabolites production, an in-frame deletion of GH19 chitinase gene (the GH19 (A0A5S3UX38) with a signal peptide) was generated in S4059 by homologous recombination (Figure 2A-B). Wild type S4059 and GH19 deletion mutant (ΔGH19) were grown in marine minimal medium (MMM) supplemented with crystalline chitin, colloidal chitin, NAG or mannose. The mutant grew similarly to the wild type in all substrates with the same growth rate and maximum cell density (Figure 2C-D and S2). The maximum cell density reached 109 CFU/mL in all substrates supplemented with casamino acid, while only reaching 108 CFU/mL when the strains grew in chitin containing MMM without casamino acid. The generation time of wild type and the mutant was 43.80 ± 8.46 mins in all substrates with casamino acid, while without casamino acid, the value was 91.27 ± 2.23 mins in crystalline chitin and 74.87 ± 0.30 mins in colloidal chitin contained MMM. The chitin degradation capacity of the P. rubra S4059 wild type and the GH19 mutant was also tested on chitin (crystalline and colloidal chitin) plates. As expected from the growth experiment, the mutant had the same chitinolytic ability (the size of clearing zone) as the wild type (Figure S2, E-F). According to previous study, the heterologously expressed GH19 chitinase in E. coli showed antifungal activity against Aspergillus niger. Therefore, the antifungal activity was also explored by co-cultivating both the strains with the fungus Aspergillus niger on MA agar plates, showing that the GH19 mutant displayed the same inhibitory effects as the wild type (data not shown).

Figure 2. The in-frame deletion of GH19 chitinase gene in Pseudoalteromonas rubra S4059 was verified by PCR with primers P1 and P2 that target left and right arm of the GH19 chitinase gene (A). The PCR products were analyzed by electrophoresis (B). M: 1-kb DNA ladder; NC: negative control with water as the PCR template; WT: PCR products with gDNA of wild type strain S4059 as template; ΔGH19: PCR product with gDNA of ΔGH19 as template. Growth kinetics of Pseudoalteromonas rubra S4059 wild type (red lines) and ΔGH19 (black lines) when grown in marine minimal medium containing (C) colloidal chitin or (D) crystalline chitin (shrimp chitin) without casamino acids at 25°C for 24h, 200 rpm. Data were analyzed on three biological replicates and the error bars represent the standard deviation.

2.6. Biofilm formation and chitin surface attachment of P. rubra S4059 was not affected by deletion of GH19 chitinase gene Many Pseudoalteromonas species are good biofilm formers that are able to colonize crustaceans in marine environments [25] and since chitin colonization has been linked to biofilm formation in other bacteria [26], the biofilm formation of S4059 wild type and the mutant were compared in chitin containing media. The surface formed biofilm of the mutant and wild type was detected at the same level in all media (Figure S3). The growth and attachment of the wild type and the mutant on natural chitin (shrimp shells) was assessed and both strains grew from an initial density of 104 CFU/mL to a final cell density of 107 CFU/mL in the liquid surrounding the shells (Figure 3). Surface-attached bacteria were removed from shrimp shells by sonication resulting in an increase in cell density to 108 CFU/mL with similar levels reached by wild type and the mutant (Figure 3).

4 Log(CFU/mL)

0 0 1 2 3 4 5 6 7 8 9 10 Time (days)

Figure 3. Growth and attachment of Pseudoalteromonas rubra S4059 wild type (red) and mutant ∆GH19 (black) on Vannamei shrimp shells. Long dash lines: samples from liquid surrounding shrimp shells before sonication. Solid lines: samples from liquid surrounding shrimp shells after sonication. Short dash lines: samples from 3% Sigma sea salt without shrimp shells as a control. Data were analyzed on three biological replicates and the error bars represent standard deviation.

2.7. Deletion of GH19 chitinase does not significantly influence the proteome of S4059 To investigate the impact of the GH19 chitinase deletion on the proteome of S4059, the GH19 mutant and the wild type were grown on chitin-based medium to stationary phase. Supernatant and cells were separated for proteome analysis. A total of 2,512 proteins were identified, but no significant changes of any of detected proteins were observed between wild type and GH19 mutant in either supernatant or cells when grown on crystalline chitin, except that GH19 chitinase could not be detected in the mutant cultures.

2.8. Similar secondary metabolome pattern in wild type strain and GH19 mutant Following the measurement of growth kinetics, the extracts of cultures grown in MMM were analyzed by HPLC-HRMS. Each strain was cultivated in three biological replicates in media containing one of the four carbon sources: mannose, NAG, colloidal chitin, or shrimp chitin. The wild type and the mutant had similar secondary metabolome profile as shown in Figure S1A. A screening for known natural products was also undertaken, revealing that only prodigiosin as well as three of its analogues could be identified (Figure S1B).

3. Discussion Chitinolytic bacteria have been widely studied due to the possible use as anti-fungal agents in biocontrol [27]. Also, the involvement of chitin in the ecology of Vibrio cholerae and its virulence regulation has been the focus of many studies [28–30]. Recently, a novel perspective on chitin degradation has been seen in some strains of Vibrionaceae, where growth on chitin as compared to on

glucose lead to enhanced expression of several BGCs [20] along with enhanced antibacterial activity [21]. Chitin is the most abundant polymer in the ocean and, hence, many marine bacteria are potent chitin degraders [10]. In silico analysis has demonstrated that pigmented species of the genus Pseudoalteromonas have an elaborate chitinolytic machinery containing at least one of the otherwise rare GH19 chitinase in their genomes and also, they devote up to 15% of their genome to BGCs with a vast potential for secondary metabolite production [7]. Here, we demonstrated that the pigmented bacterium P. rubra S4059 grew well using both purified chitin (crystalline and colloidal chitin) and natural shrimp shells as a sole source of nutrients. These results were further substantiated by proteome analyses that demonstrated that strain S4059 indeed produces an efficient chitinolytic enzyme cocktail including GH18, GH19, LPMO and GH20. GH18 chitinases are the predominant chitinolytic enzymes in bacteria [31–33] and the strain S4059 harbored seven putative GH18 chitinases as determined by genome analysis, four of which were significantly upregulated when grown on chitin. Secretome analysis of the soil-derived and chitinolytic bacterium Cellvibrio japonicus indicated that many chitinolytic enzymes including GH18 chitinases but also GH19 chitinases and LPMOs were highly upregulated when grown on β-chitin as compared to α- chitin [31]. Out of fourteen putative chitinolytic enzymes in S4059, only the two putative enzymes GH19 A0A5S3UPT5 and LPMO A0A5S3V4S2 were not detected when S4059 was grown on an α-chitin surface, suggesting that these two enzymes may have other functions. Proteome analysis of P. rubra S4059 culture supernatant confirmed the presence of all chitinolytic enzymes except for the two enzymes GH19 A0A5S3UPT5 and LPMO A0A5S3V4S2, and even higher amount of the enzymes in culture supernatant than in cells (Figure 1, C), implying that those detected proteins indeed are secreted enzymes able to degrade chitin. Secretome analysis of Cellvibrio japonicus showed that thousands of proteins were detected in the filtered culture supernatant, however, the authors mentioned that cell lysis can lead to overestimate the numbers of secreted proteins [31]. Further, they used the plate-based method that was developed by Bengtson et al. [34] to detect truly secreted proteins and the result indicated that only 267 secreted proteins were detected in α-chitin contained medium [31]. In our study, due to the insoluble property of chitin, secreted chitinases may bind to the small chitin particles and thus the secreted chitinases cannot be passed through sterile filters leading to misestimate the expression of secreted chitinases. Thus, we did not filter the culture supernatant for retaining bound chitinases which were on chitin particles. We found that 880 proteins were influenced by chitin in culture supernatant of S4059 as compared to on mannose and this high numbers of influenced proteins may due to a combination factor of the unfiltered culture supernatant and cell lysis during centrifugation [35]. Further, the agar plate-based method probably can point out true numbers of secreted proteins in S4059. One of the two GH19 chitinases in P. rubra S4059 was highly expressed when the bacterium was grown on chitin and whilst some GH19 chitinases have been linked to antifungal activity, their broader role in bacteria remains enigmatic [19,36,37]. The results suggesting antifungal activity have mainly relied on heterologous expression of GH19 in hosts such as E. coli, and this provides information about the enzyme per se, but not necessary about the actual role in its native host. We therefore chose to delete the GH19 that was highly expressed on chitin to further explore its possible role in S4059. Both the mutant and wild type displayed similar antifungal activity (data not shown). Despite the high expression of GH19 when S4059 was grown on chitin, the GH19 deficient mutant showed no difference in growth as compared to the WT when grown on chitin (Figure 4 and S1). Our results therefore indicate that GH19 chitinase is not the predominant chitin degrading enzyme in P. rubra S4059 given the wide array of other chitinolytic enzymes. Chitin utilization capacity has been investigated in a soil chitinolytic bacterium Cellvibrio japonicus [32]. Through a combination of secretome and genetic manipulation approaches, a highly expressed GH18 chitinase was identified as the enzyme being essential for the degradation of chitin in the strain [32]. Proteome analyses in S4059 demonstrated that two GH18 chitinases A0A5S3USH6 and A0A5S3USE2 are the most highly expressed chitinases in both cells and culture supernatant according to protein abundance when S4059 was grown on chitin (Table

S7), suggesting that the GH18 chitinases could be potentially essential enzymes for chitin degradation in S4059 under testing conditions. In conclusion, proteomic analysis showed that a highly efficient chitin degradation machinery was identified in pigmented P. rubra S4059 and four GH18 chitinases and two GH20 hexosaminidases in S4059 were significantly upregulated when grown on chitin. In contrast to different proteome profiles, the different growth conditions on chitin investigated here did not significantly alter the metabolite profile of S4059, in contrast to what has been reported in other Vibrio species, and Streptomyces coelicolor A3 (2) [20–23]. Although the deletion of GH19 chitinase did not influence of chitin degradation activity in S4059, we developed a conjugation-based approach allowing genetic manipulation in this bioactive bacterium S4059. Given the large potential for secondary metabolism as found by genome mining, and from uncharacterized compounds produced on especially chitin derived substrates, the genetic approach developed here, the genetic approach will allow exploration of the novel chemical space of this organism.

4. Materials and Methods

4.1. Bacterial strains, plasmids and growth conditions All bacterial strains and plasmids used in this study are listed in Table S5. P. rubra S4059 was isolated during the Galathea 3 expedition [38] and cultured in marine broth (MB, BD Difco 2216) or APY medium [39] at 25oC, 200 rpm. P. rubra S4059 carrying a chromosomal-integrated suicide plasmid was cultured in MB containing 30 µg/mL chloramphenicol (Sigma, C0378) at 25oC, 200 rpm. All chitin used in this study is α-chitin. P. rubra S4059 mutants and wild type were cultured in marine minimal medium (MMM) [40] supplemented with four different carbon sources (0.2% mannose, 0.2% crystalline chitin, 0.2% colloidal chitin or 0.2% NAG) and growth, biofilm formation and secondary metabolome determined. To compare chitin degradation of mutants and wild type S4059, they were grown on plates containing 2% Sea Salt (Sigma, S9883), 1,5% agar and 0,2% chitin (crystalline or colloidal). Colloidal chitin was prepared as described in previous study [41]. All Escherichia coli strains were cultured in Luria Bertani (LB) Broth (BD Difco 244520) at 37 ℃, 200 rpm. E. coli GB dir-pir116 [42] was used for constructing suicide plasmid. E. coli WM3064 was used as the donor strain in intergeneric conjugation experiments. E. coli WM3064 is a dapA mutant that requires exogenously supplied diaminopimelic acid (DAP, Sigma, D1377) with a final concentration of 0.3 µM for growth [43]. Plasmid pDM4 was used as the backbone of suicide vectors [44]. E. coli strains harboring pDM4 or its derivatives were grown in LB Broth with 10 µg/mL chloramphenicol or in LB agar with 15 µg/mL chloramphenicol.

4.2. Whole genome sequencing and assembly of P. rubra S4059 Genomic DNA of P. rubra S4059 was extracted using the Genomic DNA buffer set (QIAGEN, 19060) following the supplier’s instructions. The closed genome of P. rubra S4059 was obtained by minION sequencing using the EXP-FLP002 flow cell priming kit, SQK-RAD004 rapid sequencing kit, FLO-MINSP6 flow cell and the associated protocol (version RSE_9046_V1_revB-17Nov2017 and RSE_9046_V1_revB-14 AUG2019). lllumina reads were obtained from a previous study [7] were combined with the minION reads for hybrid assembly using the Unicycler package [45]. Prior to assembly, the minION reads were filtered using the Filtlong package, only keeping the top 500.000.000 bp. The genome was annotated using Prokka [46]. The genome is available at the National Center for Biotechnology Information (NCBI) under the accession number CP045429 and CP045430.

4.3. In silico analysis of chitin degrading genes and secondary metabolites The assembled genome was analyzed using the CLC Main Workbench 8.0.1 (CLC bio, Aarhus, Denmark) and the online platform MaGe MicroScope [47]. The chitinolytic enzymes and the prediction of chitin degradation pathway in S4059 were annotated according to the prediction of S4059 proteome

in Uniprot (UniProt ID UP000305729). Amino acid sequences of the candidate chitinases were also submitted to the SignalP 5.0 Server [48] to identify the signal peptides. In addition, genomes were submitted to antiSMASH version 6.0 (https://antismash.secondarymetabolites.org/#!/start) for the prediction of putative biosynthetic gene clusters involved in the production of secondary metabolites.

4.4. Growing bacteria and sample preparation for proteomic analyses. The protein samples were prepared from Chevallier et al. [49]. P. rubra S4059 wild type and GH19 mutant were grown in 20 mL MMM with crystalline chitin for 2 days at 25 °C, 200 rpm. All experiments are carried out in biological triplicates. 5 mL culture was harvested (5000 x g, 20 mins) and then the culture supernatant was transferred into a new 15 mL Falcon tube and ice- cold acetone (-20 °C) was added to the supernatant to a final concentration of 80%. Then the mixture was kept at -20 °C overnight and harvested at 2,000 x g for 20 mins. Acetone was carefully removed. The harvested bacterial cells were washed twice with ice cold phosphate-buffered saline (PBS) and the pellet was lysed using 20 µl of lysis buffer (consisting of 6 M Guanidinium Hydrochloride, 10 mM Tris (2-carboxyethyl) phosphine hydrochloride, 40 mM 2-chloroacetamide, 50 mM HEPES pH 8.5). Samples were inactivated at 95 °C for 5 minutes and were then sonicated on high for 3 times of 10 seconds in a 4°C Bioruptor sonication water bath (Diagenode). A Bradford assay (Sigma) was used to determine protein concentration, and 50 µg of each sample was used for digestion. Samples were diluted 1:3 with 10% Acetonitrile, 50 mM HEPES pH 8.5, LysC (MS grade, Wako) was added in a 1:50 (enzyme to protein) ratio, and samples were incubated at 37 °C for 4 hours. Samples were further diluted to 1:10 with 10% Acetonitrile, 50 mM HEPES pH 8.5, trypsin (MS grade, Promega) was added in a 1:100 (enzyme to protein) ratio and samples were incubated overnight at 37 °C. Enzyme activity was quenched by adding 2% trifluoroacetic acid (TFA) to a final concentration of 1%. Prior to mass spectrometry analysis, the peptides were desalted on SOLAu C18 plates (Thermo Fisher Scientific). After each solvent application, the plate was centrifuged for 1 minute at 350 x g. For each sample, the C18 material was activated with 200ul of 100% Methanol (HPLC grade, Sigma), then 200ul of 80% Acetonitrile, 0.1% formic acid. The C18 material was subsequently equilibrated 2 x with 200 µl of 1% TFA, 3% Acetonitrile, after which the samples were loaded. After washing the tips twice with 200ul of 0.1% formic acid, the peptides were eluted using 40% Acetonitrile, 0.1% formic acid, and transferred into clean 500 µL Eppendorf tubes. The eluted peptides were concentrated in an Eppendorf Speedvac, and re-constituted in 1% TFA, 2% Acetonitrile for Mass Spectrometry (MS) analysis.

4.5. Proteomic data acquisition The proteomic data acquisition approach was modified from Haddad Momeni et al [50]. Briefly, peptides were loaded onto a 2 cm C18 trap column (ThermoFisher 164705), connected in-line to a 15 cm C18 reverse-phase analytical column (Thermo EasySpray ES803) using 100% Buffer A (0.1% Formic acid in water) at 750 bar, using the Thermo EasyLC 1200 HPLC system, and the column oven operating at 30°C. Peptides were eluted over a 140 minutes gradient ranging from 10 to 60% of 80% acetonitrile, 0.1% formic acid at 250 nl/min, and the Q-Exactive instrument (Thermo Fisher Scientific) was run in a DD-MS2 top 10 method. Full MS spectra were collected at a resolution of 70,000, with an automatic gain control (AGC) target of 3×106 or maximum injection time of 20 ms and a scan range of 300–1750 m/z. The MS2 spectra were obtained at a resolution of 17,500, with an AGC target value of 1 × 10E6 or maximum injection time of 60 ms, a normalised collision energy of 25 and an intensity threshold of 1.7e4. Dynamic exclusion was set to 60 s and ions with a charge state < 2 or unknown were excluded. MS performance was verified for consistency by running complex cell lysate quality control standards, and chromatography was monitored to check for reproducibility. The mass spectrometry data have been deposited to the ProteomeXchange Consortium (http://proteomecentral.proteomexchange.org) via the PRIDE partner repository with the dataset identifier PXD 023249. The reviewer account details: Username: [email protected]; Password: xteOmcDd. The raw files were analysed using Proteome Discoverer 2.4. Label-free quantitation (LFQ) was enabled in the processing and consensus

steps, and spectra were matched against the P. rubra S4059 database obtained from Uniprot (UniProt ID: UP000305729). Dynamic modifications were set as Oxidation (M), Deamidation (N, Q) and Acetyl on protein N-termini. Cysteine carbamidomethyl was set as a static modification. The methods of proteomic analyses were modified from Beyene et al [51]. Briefly, all the statistical analyses were performed using Perseus software (version 1.6.14.0, https://maxquant.net/perseus/), all results were filtered to a 1% false discovery rate (FDR). The normalized abundance values for each protein were log2 transformed and at least two valid values were required in the bio-triplicates for quantitation. When the original signals were zero, they were imputed with random numbers from a normal distribution, which the mean and standard deviation were chosen from low abundance values below the noise level (Width= 0.3; shift=1.8) [51,52]. To identify proteins with significantly different abundance when S4059 where grown on chitin as compared to on mannose, the FDR were estimated using Benjamini-hochberg method and a two tailed unpaired t-test was used with a combination of p- value ≤ 0.05 and Log2 fold-change ≥ 1.5 [53]. The resulting significant proteins were exported from Perseus and visualized in GraphPad Prism 8 (https://www.graphpad.com/scientific-software/prism/) using volcano plots.

4.6. DNA manipulation Genomic DNA of P. rubra S4059 was extracted using the Genomic DNA buffer set (QIAGEN, 19060) as mentioned above. All primers used in this study are listed in Table S6. All purified DNA fragments were amplified using PrimeSTAR® Max Premix (TaKaRa, catalog number: R045A). Blue TEMPase Hot Start Master Mix K (catalog number: 733-2584) was used for homologous recombination event checking by PCR. All primers and plasmids were designed in A Plasmid Editor- ApE. The specificity of primers was checked by BLAST against the P. rubra S4059 genome. All primers were ordered from Integrated DNA technologies (Leuven, Belgium).

4.7. Construction of suicide plasmids for in-frame deletion of GH19 chitinase in P. rubra S4059. The suicide plasmid was constructed by direct cloning method using pDM4 as the backbone [42,44]. The pDM4 plasmid contains a R6K replicon origin, mob genes and oirT for conjugation, and a chloramphenicol resistance gene cat and a sacB gene for counter selection [44]. An approximately 1-kb upstream and downstream region flanking of GH19 gene were amplified with primer pairs GH19-L- F/GH19-L-R, GH19-R-F/GH19-R-R (Table S6). The amplified recombining arms were fused with overlap extension PCR to form the recombining arm segments, which were cloned into pJET1.2 subcloning vector using CloneJET PCR Cloning Kit (Thermo Fisher Scientific, K1231) for sequencing. Subsequently, the homologous segment was amplified from sequencing-confirmed pJET1.2-dGH19 arms using primers GH19-pJET1.2-F and GH19-pJET1.2-R. The linear backbone was amplified from pDM4 suicide vector using primers (GH19-pDM4-F/GH19-pDM4-R). After gel purification, the linear vector and the homologous fragment were co-electroporated into E. coli GB dir-pir116 and ligated by the RecET direct cloning system [42]. Restriction cloning method was also attempted several times to construct this plasmid, however, it was unsuccessful. All plasmids were extracted using QIAprep Spin Miniprep Kit (QIAGEN, 27106).

4.8. Conjugation of P. rubra S4059. The conjugation protocol was modified from Yu et al. [54]. E. coli WM3064 harboring the suicide plasmid was used as the donor and P. rubra S4059 as recipient. Overnight cultures of donor and recipient were prepared as pre-cultures one day before the conjugation. During conjugation, both strains were diluted 100 times and grown to OD600≈ 0.6. One-mL donor cells were harvested at 6,000xg for 1 min. The pellets were resuspended and washed once using 1 ml LB+DAP. Then, 1 mL recipient was added to the E. coli WM3064 pellet and centrifuged at 6,000 x g for 1 min. One-mL MB+DAP was added to wash the mixture by pipetting and centrifuged at 6,000 x g for 1 min. The supernatant was

removed until 20-30 µl liquid remained. The mixture of cells was resuspended and placed on a 0.2-μm pore-size membrane (MF-Millipore, GSWP02500) that was placed on an APY+DAP agar plate. The mating plates were incubated at 20°C for 24h. The cells were suspended in 1mL MB and incubated with shaking at 750 rpm in an Eppendorf® thermomixer comfort, 25 °C for 1 h. After recovery, cells were spread on plates with antibiotics and incubated at 25 °C for 24-48 h.

4.9. Confirmation of the first crossing over mutants and deletion mutants. Colonies from the first crossover selective plates were picked and cultured in 5 mL MB containing 30 µg/mL chloramphenicol overnight. Genomic DNA extraction from pre-cultures using NucleoSpin® Tissue kit (Macherey-Nagel, 740952.250). To determine whether the plasmid integrated into target regions, the genomic DNA was used as the template for PCR checking with primers (Cmr-F/Cmr-R, GH19-p 1/GH19-p 4, and GH19-p 2/GH19-p 3). Colonies carrying the integrated plasmid were cultured in MB with 30 µg/mL chloramphenicol at 25°C overnight as pre-culture. The pre-culture was diluted 100 times and inoculated in 5 mL fresh MB without antibiotics until OD600≈0.6. The culture was 10-fold diluted and spread on the counter selection plates (half nutrients of MA) containing 10% sucrose. These counter selection plates were incubated at 20°C until colonies were visible. Confirmation of the in-frame deletion mutants were carried out by PCR application and sequencing. Primers (GH19-p1/GH19-p2) were designed to amplify mutation region and the PCR produced was purified and sent for DNA sequencing.

4.10. Growth curves of wild type and mutants. Growth kinetics of P. rubra S4059 WT (wild type) and ΔGH19 (GH19 chitinase mutant) were established in MMM with or without 0.3% casamino acids, and supplemented with 0.2% colloidal chitin, 0.2% crab chitin (C9752, Sigma) 0.2% mannose (63580, Sigma), or 0.2% NAG (A3286, Sigma), respectively, as carbon source. Pre-cultures of P. rubra WT and mutants were grown in MB overnight at 25°C. The cultures were diluted to a starting concentration of 103 CFU/mL in MMM. Samples were taken every 4 hours for estimation of cell density by plate counting. All experiments were done in biological duplicate.

4.11. Growth and attachment on shrimp shells. 6 mm diameter circular disks were prepared from the exoskeleton of Vannamei shrimps (description in supplementary material). Bacterial cultures (WT and mutants) were grown in 5 mL MB in 50 mL flasks at 25 °C, 200 rpm overnight. The cultures were diluted to 104 CFU/mL in artificial sea water (3% sea salt, Sigma, S9883) and incubated with or without shrimp shells in 1.5 mL Eppendorf tubes. Samples of the suspension were taken after 0, 2, 5, 7, 10 days incubation and plated after serial dilution. The tubes were sonicated five minutes at 50/60 Hz in a ultrasonic bath (Emmi D20 Q, EMAG, Germany) and vortexed 10 s to remove bacteria attached to the shrimp shells [55]. Suspension of cells with known cell counts were subjected to the same sonication and cell counts done after sonication, demonstrating that this procedure did not reduce cell counts. Then serial dilutions and colony counts were done to determine cell densities. The experiment was done in biological duplicate.

4.12. Extraction of metabolites for chemical analysis. Bacterial cultures were grown in MMM with different carbon sources and samples taken for chemical analyses after 72 h incubation. Ten mL sample was extracted with an equal volume of high- performance liquid chromatography (HPLC)-grade Ethyl acetate in 50 mL Falcon tubes. The organic phase was transferred to a new vial and evaporated under nitrogen. The dried extract was re-dissolved in 500 μL methanol and stored at -20°C. Chemical analysis was performed in biological triplicate.

4.13. Chemical analysis by UHPLC-HRMS. The chemical analysis approach was modified from Holm et al [56]. Chemical analysis was performed on a Bruker maXis 3G orthogonal acceleration quadrupole time-of-flight mass spectrometer (Bruker Daltonics) equipped with an electrospray ionization (ESI) source and connected to an Ultimate 3000 UHPLC system (Dionex). The column used was a reverse-phase Kinetex 1.7 μm C18, 100 mm × 2.1 mm (Phenomenex). The column temperature was kept at 40°C throughout the analysis. A linear gradient of LC-MS grade water and acetonitrile both buffered with formic acid was used, starting at

10% (v/v) acetonitrile and increased to 100% in 10 min, maintaining this rate for 3 min before returning to the starting conditions in 0.1 min and staying there for 2.4 min before the following run. A flow rate of 0.40 mL/min was used. TOFMS was performed in ESI+ with a data acquisition range of 10 scans per second at m/z 75–1250. The TOFMS was calibrated using Bruker Daltonics high precision calibration algorithm by means of the internal standard sodium formate, which was automatically infused before each run. This provided a mass accuracy of better than 1.5 ppm in MS mode. UV-visible spectra were collected at wavelengths from 200 to 700 nm. Data processing was performed using DataAnalysis 4.0 and Target Analysis 1.2 software (Bruker Daltonics). Tandem MS spectra were acquired on an Agilent 6545 QTOF-MS using the method described in Isbrandt et al. (2020) [57].

Supplementary Materials: The following are available online at www.mdpi.com/xxx/s1,

Table S1. The predicted biosynthetic gene clusters (BGCs) of Pseudoalteromonas rubra S4059 by antiSMASH 6.0.

Table S2. The fold change of six significantly upregulated chitinolytic enzymes in cell or supernatant samples.

Table S3. Significantly up-and down-regulated proteins in Pseudoalteromonas rubra S4059 proteome when grown on chitin as compared to on mannose.

Table S4. Antibiotic sensitivity of Pseudoalteromonas rubra S4059 growth on MB agar plate.

Table S5. Bacteria and plasmids used in this study.

Table S6. Primers used in this study.

Table S7. The normalized protein abundance of chitinolytic enzymes in S4059.

Figure S1. (A) Base peak chromatograms of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 mutant when cultivated on marine minimal medium using mannose, crystalline chitin, colloidal chitin or N-acetyl glucosamine (NAG) as the carbon source. The red pigment prodigiosin and two of its analogues (hexyl prodigiosin and heptyl prodigiosin) could be identified in the extract and confirmed based on MS/MS experiments and the acquired absorption spectra. Experiments were done in biological triplicates, and a reference chromatogram of the sterile growth medium is included to show media components also present in the experiments. (B) Tandem MS spectra recorded for 1) prodigiosin, 2) hexyl-prodigiosin, and 3) heptyl-prodigiosin. The prodigiosin MS/MS spectra matches our in-house MS/MS library, and the characteristic fragment m/z 252.11, and loss of CH3 (m/z 15.02) additionally matches those previously reported for prodigiosin and analogues [58]. (C) Recorded UV-Vis absorption spectrum of prodigiosin. All proposed prodigiosins share identical absorption spectra. The spectrum is in agreement with previously reported literature [59].

Figure S2. (A-D) Growth kinetics of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 mutant in mannose (A), colloidal chitin (B), crystalline chitin (crab chitin) (C) and NAG (chitin monomer) (D) containing MMM with casamino acids. (E-F) WT and ΔGH19 growth in crystalline chitin (E) or colloidal chitin containing MMM without casamino acid. Square: WT; Triangle: ΔGH19. The points are bio-replicates and error bars are standard deviation. (G-H) Chitin degradation activities of wild type and the mutant on colloidal chitin plate (G) and crystalline chitin plate (H).

Figure S3. Biofilm formation on the microtiter-well plastic surface of Pseudoalteromonas rubra S4059 wild type and ΔGH19 mutant in four different sole carbon contained medium as determined by the O’Toole & Kolter crystal violet assay. (A) in mannose; (B) in NAG (chitin monomer); (C) in colloidal chitin; (D) in crab chitin. All experiments were in bio-triplicates and error bars are standard deviation.

Author Contributions: XW, SSP, SDZ and LG conceived and design experiments; XW performed the experiments; TI and TOL performed chemical analysis; EMS and MWN performed the proteomic analyses. MILST assembled the genome of P. rubra S4059 and supervised statistical analysis. YB assisted to approve that GH19 is expressed in P. rubra S4059 when grown on chitin. XW wrote the first manuscript draft; SDZ and LG supervised edited and finalized the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding: This research was supported by Chinese Scholarship Council (CSC scholarship No. 201706170066), the Danish National Research Foundation (DNRF137) for the Center for Microbial Secondary Metabolites, the Independent Research Fund Denmark (grant DFF – 7017-00003), and the European Union's Horizon 2020 research

and Innovation Programme under the Marie Sklodowska-Curie grant agreement no. 713683 (COFUND fellows DTU) via the H. C. Ørsted fellowship programme.

Acknowledgments: We gratefully acknowledge Dr. Professor Youming Zhang and Dr. Zhen Li from Shandong University for kindly providing the RecET direct cloning strains and vectors.

Conflicts of Interest: The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

References 1. Romano, G.; Costantini, M.; Sansone, C.; Lauritano, C.; Ruocco, N.; Ianora, A. Marine microorganisms as a promising and sustainable source of bioactive molecules. Mar. Environ. Res. 2017, 128, 58–69, doi:10.1016/j.marenvres.2016.05.002.

2. Parte, S.; Sirisha, V.L.; D’Souza, J.S. Biotechnological Applications of Marine Enzymes From Algae, Bacteria, Fungi, and Sponges. Adv. Food Nutr. Res. 2017, 80, 75–106, doi:10.1016/bs.afnr.2016.10.005.

3. Bowman, J.P. Bioactive compound synthetic capacity and ecological significance of marine bacterial genus Pseudoalteromonas. Mar. Drugs 2007, 5, 220–241, doi:10.3390/md504220.

4. Offret, C.; Desriac, F.; Le Chevalier, P.; Mounier, J.; Jégou, C.; Fleury, Y. Spotlight on antimicrobial metabolites from the marine bacteria Pseudoalteromonas: Chemodiversity and ecological significance. Mar. Drugs 2016, 14, 1–26, doi:10.3390/md14070129.

5. Tang, B.L.; Yang, J.; Chen, X.L.; Wang, P.; Zhao, H.L.; Su, H.N.; Li, C.Y.; Yu, Y.; Zhong, S.; Wang, L.; et al. A predator-prey interaction between a marine Pseudoalteromonas sp. and Gram-positive bacteria. Nat. Commun. 2020, 11, 285, doi:10.1038/s41467-019-14133-x.

6. Zhao, H.L.; Chen, X.L.; Xie, B. Bin; Zhou, M.Y.; Gao, X.; Zhang, X.Y.; Zhou, B.C.; Weiss, A.S.; Zhang, Y.Z. Elastolytic mechanism of a novel M23 metalloprotease pseudoalterin from deep-sea Pseudoalteromonas sp. CF6-2: Cleaving not only glycyl bonds in the hydrophobic regions but also peptide bonds in the hydrophilic regions involved in cross-linking. J. Biol. Chem. 2012, 287, 39710–39720, doi:10.1074/jbc.M112.405076.

7. Paulsen, S.S.; Strube, M.L.; Bech, P.K.; Gram, L.; Sonnenschein, E.C. Marine Chitinolytic Pseudoalteromonas Represents an Untapped Reservoir of Bioactive Potential . mSystems 2019, 4, 1–12, doi:10.1128/msystems.00060-19.

8. Rinaudo, M. Chitin and chitosan: Properties and applications. Prog. Polym. Sci. 2006, 31, 603–632, doi:10.1016/J.PROGPOLYMSCI.2006.06.001.

9. RUDALL, B.K.. M..; KENCHINGTON, W.. THE CHITIN SYSTEM. BioL Rev . 1973, 49, 597–636.

10. Beier, S.; Bertilsson, S. Bacterial chitin degradation-mechanisms and ecophysiological strategies. Front. Microbiol. 2013, 4, 1–12, doi:10.3389/fmicb.2013.00149.

11. Vaaje-Kolstad, G.; Horn, S.J.; Sørlie, M.; Eijsink, V.G.H. The chitinolytic machinery of Serratia marcescens - A model system for enzymatic degradation of recalcitrant polysaccharides. FEBS J. 2013, 280, 3028–3049, doi:10.1111/febs.12181.

12. Oyeleye, A.; Normi, Y.M. Chitinase: Diversity, limitations, and trends in Engineering for suitable applications. Biosci. Rep. 2018, 38, 1–21, doi:10.1042/BSR20180323.

13. Gooday, G.W. The Ecology of Chitin Degradation. 1990, 387–430, doi:10.1007/978-1-4684-7612-5_10.

14. Bai, Y.; Eijsink, V.G.H.; Kielak, A.M.; van Veen, J.A.; de Boer, W. Genomic comparison of chitinolytic enzyme systems from terrestrial and aquatic bacteria. Environ. Microbiol. 2016, 18, 38–49, doi:10.1111/1462- 2920.12545.

15. Oyeleye, A.; Normi, Y.M. Chitinase: Diversity, limitations, and trends in Engineering for suitable applications. Biosci. Rep. 2018, 38.

16. Udaya Prakash, N.A.; Jayanthi, M.; Sabarinathan, R.; Kangueane, P.; Mathew, L.; Sekar, K. Evolution, homology conservation, and identification of unique sequence signatures in GH19 family chitinases. J. Mol.

Evol. 2010, 70, 466–478, doi:10.1007/s00239-010-9345-z.

17. Konno, N.; Takahashi, H.; Nakajima, M.; Takeda, T.; Sakamoto, Y. Characterization of β-N- acetylhexosaminidase (LeHex20A), a member of glycoside hydrolase family 20, from Lentinula edodes (shiitake mushroom). AMB Express 2012, 2, 1–7, doi:10.1186/2191-0855-2-29.

18. Vaaje-Kolstad, G.; Westereng, B.; Horn, S.J.; Liu, Z.; Zhai, H.; Sørlie, M.; Eijsink, V.G.H. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science (80-. ). 2010, 330, 219–222, doi:10.1126/science.1192231.

19. García-Fraga, B.; da Silva, A.F.; López-Seijas, J.; Sieiro, C. A novel family 19 chitinase from the marine- derived Pseudoalteromonas tunicata CCUG 44952T: Heterologous expression, characterization and antifungal activity. Biochem. Eng. J. 2015, 93, 84–93, doi:10.1016/j.bej.2014.09.014.

20. Giubergia, S.; Phippen, C.; Gotfredsen, C.H.; Nielsen, K.F.; Gram, L. Influence of niche-specific nutrients on secondary metabolism in Vibrionaceae. Appl. Environ. Microbiol. 2016, 82, 4035–4044, doi:10.1128/AEM.00730-16.

21. Giubergia, S.; Phippen, C.; Nielsen, K.F.; Gram, L. Growth on Chitin Impacts the Transcriptome and Metabolite Profiles of Antibiotic-Producing Vibrio coralliilyticus S2052 and Photobacterium galatheae S2753. mSystems 2017, 2, 1–12, doi:10.1128/msystems.00141-16.

22. Wietz, M.; Duncan, K.; Patin, N. V.; Jensen, P.R. Antagonistic Interactions Mediated by Marine Bacteria: The Role of Small Molecules. J. Chem. Ecol. 2013, 39, 879–891, doi:10.1007/s10886-013-0316-x.

23. Nazari, B.; Kobayashi, M.; Saito, A.; Hassaninasab, A.; Miyashita, K.; Fujiia, T. Chitin-induced gene expression in secondary metabolic pathways of Streptomyces coelicolor A3(2) grown in soil. Appl. Environ. Microbiol. 2013, 79, 707–713, doi:10.1128/AEM.02217-12.

24. Blin, K.; Shaw, S.; Steinke, K.; Villebro, R.; Ziemert, N.; Lee, S.Y.; Medema, M.H.; Weber, T. AntiSMASH 5.0: Updates to the secondary metabolite genome mining pipeline. Nucleic Acids Res. 2019, 47, W81–W87, doi:10.1093/nar/gkz310.

25. Saravanan, P.; Nancharaiah, Y.V.; Venugopalan, V.P.; Rao, T.S.; Jayachandran, S. Biofilm formation by Pseudoalteromonas ruthenica and its removal by chlorine. Biofouling 2006, 22, 371–381, doi:10.1080/08927010601029103.

26. Margolis, J.J.; El-Etr, S.; Joubert, L.M.; Moore, E.; Robison, R.; Rasley, A.; Spormann, A.M.; Monack, D.M. Contributions of francisella tularensis subsp. novicida chitinases and sec secretion system to biofilm formation on chitin. Appl. Environ. Microbiol. 2010, 76, 596–608, doi:10.1128/AEM.02037-09.

27. A. Veliz, E.; Martínez-Hidalgo, P.; M. Hirsch, A. Chitinase-producing bacteria and their role in biocontrol. AIMS Microbiol. 2017, 3, 689–705, doi:10.3934/microbiol.2017.3.689.

28. Reguera, G.; Kolter, R. Virulence and the environment: A novel role for Vibrio cholerae toxin-coregulated pili in biofilm formation on chitin. J. Bacteriol. 2005, 187, 3551–3555, doi:10.1128/JB.187.10.3551-3555.2005.

29. Sun, S.; Tay, Q.X.M.; Kjelleberg, S.; Rice, S.A.; McDougald, D. Quorum sensing-regulated chitin metabolism provides grazing resistance to Vibrio cholerae biofilms. ISME J. 2015, 9, 1812–1820, doi:10.1038/ismej.2014.265.

30. Pruzzo, C.; Vezzulli, L.; Colwell, R.R. Global impact of Vibrio cholerae interactions with chitin. Environ. Microbiol. 2008, 10, 1400–1410, doi:10.1111/j.1462-2920.2007.01559.x.

31. Tuveng, T.R.; Arntzen, M.Ø.; Bengtsson, O.; Gardner, J.G.; Vaaje-Kolstad, G.; Eijsink, V.G.H. Proteomic investigation of the secretome of Cellvibrio japonicus during growth on chitin. Proteomics 2016, 16, 1904– 1914, doi:10.1002/pmic.201500419.

32. Monge, E.C.; Tuveng, T.R.; Vaaje-Kolstad, G.; Eijsink, V.G.H.; Gardner, J.G. Systems analysis of the glycoside hydrolase family 18 enzymes from Cellvibrio japonicus characterizes essential chitin degradation functions. J. Biol. Chem. 2018, 293, 3849–3859, doi:10.1074/jbc.RA117.000849.

33. Hayes, C.A.; Dalia, T.N.; Dalia, A.B. Systematic genetic dissection of chitin degradation and uptake in Vibrio cholerae. Environ. Microbiol. 2017, 19, 4154–4163, doi:10.1111/1462-2920.13866.

34. Bengtsson, O.; Arntzen, M.; Mathiesen, G.; Skaugen, M.; Eijsink, V.G.H. A novel proteomics sample preparation method for secretome analysis of Hypocrea jecorina growing on insoluble substrates. J. Proteomics 2016, 131, 104–112, doi:10.1016/j.jprot.2015.10.017.

35. Peterson, B.W.; Sharma, P.K.; van der Mei, H.C.; Busscher, H.J. Bacterial cell surface damage due to centrifugal compaction. Appl. Environ. Microbiol. 2012, 78, 120–125, doi:10.1128/AEM.06780-11.

36. Martínez-Caballero, S.; Cano-Sánchez, P.; Mares-Mejía, I.; Díaz-Sánchez, A.G.; Macías-Rubalcava, M.L.; Hermoso, J.A.; Rodríguez-Romero, A. Comparative study of two GH19 chitinase-like proteins from Hevea brasiliensis, one exhibiting a novel carbohydrate-binding domain. FEBS J. 2014, 281, 4535–4554, doi:10.1111/febs.12962.

37. Ohno, T.; Armand, S.; Hata, T.; Nikaidou, N.; Henrissat, B.; Mitsutomi, M.; Watanabe, T. A modular family 19 chitinase found in the prokaryotic organism Streptomyces griseus HUT 6037. J. Bacteriol. 1996, 178, 5065– 5070, doi:10.1128/jb.178.17.5065-5070.1996.

38. Gram, L.; Melchiorsen, J.; Bruhn, J.B. Antibacterial activity of marine culturable bacteria collected from a global sampling of ocean surface waters and surface swabs of marine organisms. Mar. Biotechnol. 2010, 12, 439–451, doi:10.1007/s10126-009-9233-y.

39. Zhang, S. Da; Santini, C.L.; Zhang, W.J.; Barbe, V.; Mangenot, S.; Guyomar, C.; Garel, M.; Chen, H.T.; Li, X.G.; Yin, Q.J.; et al. Genomic and physiological analysis reveals versatile metabolic capacity of deep-sea Photobacterium phosphoreum ANT-2200. Extremophiles 2016, 20, 301–310, doi:10.1007/s00792-016-0822-1.

40. Östling, J.; Goodman, A.; Kjelleberg, S. Behaviour of IncP‐1 plasmids and a miniMu transposon in a marine Vibrio sp.: isolation of starvation inducible lac operon fusions. FEMS Microbiol. Lett. 1991, 86, 83–94, doi:10.1111/j.1574-6968.1991.tb04797.x.

41. Paulsen, S.S.; Andersen, B.; Gram, L.; MacHado, H. Biological potential of chitinolytic marine bacteria. Mar. Drugs 2016, 14, doi:10.3390/md14120230.

42. Wang, H.; Li, Z.; Jia, R.; Hou, Y.; Yin, J.; Bian, X.; Li, A.; Müller, R.; Stewart, A.F.; Fu, J.; et al. RecET direct cloning and Redαβ recombineering of biosynthetic gene clusters, large operons or single genes for heterologous expression. Nat. Protoc. 2016, 11, 1175–1190, doi:10.1038/nprot.2016.054.

43. Dehio, C.; Meyer, M. Maintenance of broad-host-range incompatibility group P and group Q plasmids and transposition of Tn5 in Bartonella henselae following conjugal plasmid transfer from Eescherichia coli. J. Bacteriol. 1997, 179, 538–540, doi:10.1128/jb.179.2.538-540.1997.

44. Milton, D.L.; O’Toole, R.; Hörstedt, P.; Wolf-Watz, H. Flagellin A is essential for the virulence of Vibrio anguillarum. J. Bacteriol. 1996, 178, 1310–1319, doi:10.1128/jb.178.5.1310-1319.1996.

45. Wick, R.R.; Judd, L.M.; Gorrie, C.L.; Holt, K.E. Unicycler: Resolving bacterial genome assemblies from short and long sequencing reads. PLoS Comput. Biol. 2017, 13, 1–22, doi:10.1371/journal.pcbi.1005595.

46. Seemann, T. Prokka: Rapid prokaryotic genome annotation. Bioinformatics 2014, 30, 2068–2069, doi:10.1093/bioinformatics/btu153.

47. Vallenet, D.; Calteau, A.; Dubois, M.; Amours, P.; Bazin, A.; Beuvin, M.; Burlot, L.; Bussell, X.; Fouteau, S.; Gautreau, G.; et al. MicroScope: an integrated platform for the annotation and exploration of microbial gene functions through genomic, pangenomic and metabolic comparative analysis. Nucleic Acids Res. 2019, doi:10.1093/nar/gkz926.

48. Almagro Armenteros, J.J.; Tsirigos, K.D.; Sønderby, C.K.; Petersen, T.N.; Winther, O.; Brunak, S.; von Heijne, G.; Nielsen, H. SignalP 5.0 improves signal peptide predictions using deep neural networks. Nat. Biotechnol. 2019, 37, 420–423, doi:10.1038/s41587-019-0036-z.

49. Chevallier, V.; Schoof, E.M.; Malphettes, L.; Andersen, M.R.; Workman, C.T. Characterization of glutathione proteome in CHO cells and its relationship with productivity and cholesterol synthesis. Biotechnol. Bioeng. 2020, 117, 3448–3458, doi:10.1002/bit.27495.

50. Haddad Momeni, M.; Leth, M.L.; Sternberg, C.; Schoof, E.; Nielsen, M.W.; Holck, J.; Workman, C.T.; Hoof, J.B.; Abou Hachem, M. Loss of AA13 LPMOs impairs degradation of resistant starch and reduces the growth of Aspergillus nidulans. Biotechnol. Biofuels 2020, 13, 1–13, doi:10.1186/s13068-020-01775-z.

51. Beyene, G.T.; Kalayou, S.; Riaz, T.; Tonjum, T. Comparative proteomic analysis of Neisseria meningitidis wildtype and dprA null mutant strains links DNA processing to pilus biogenesis. BMC Microbiol. 2017, 17, 1–18, doi:10.1186/s12866-017-1004-8.

52. Yimer, S.A.; Birhanu, A.G.; Kalayou, S.; Riaz, T.; Zegeye, E.D.; Beyene, G.T.; Holm-Hansen, C.; Norheim, G.; Abebe, M.; Aseffa, A.; et al. Comparative proteomic analysis of Mycobacterium tuberculosis lineage 7 and lineage 4 strains reveals differentially abundant proteins linked to slow growth and virulence. Front. Microbiol. 2017, 8, 1–14, doi:10.3389/fmicb.2017.00795.

53. Dunn, J.; Ferluga, S.; Sharma, V.; Futschik, M.; Hilton, D.A.; Adams, C.L.; Lasonder, E.; Hanemann, C.O. Proteomic analysis discovers the differential expression of novel proteins and phosphoproteins in meningioma including NEK9, HK2 and SET and deregulation of RNA metabolism. EBioMedicine 2019, 40, 77–91, doi:10.1016/j.ebiom.2018.12.048.

54. Yu, Z.C.; Zhao, D.L.; Ran, L.Y.; Mi, Z.H.; Wu, Z.Y.; Pang, X.; Zhang, X.Y.; Su, H.N.; Shi, M.; Song, X.Y.; et al. Development of a genetic system for the deep-sea psychrophilic bacterium Pseudoalteromonas sp. SM9913. Microb. Cell Fact. 2014, 13, 1–9, doi:10.1186/1475-2859-13-13.

55. Kim, E.; Kinney, W.H.; Ovrutsky, A.R.; Vo, D.; Bai, X.; Honda, J.R.; Marx, G.; Peck, E.; Lindberg, L.; Falkinham, J.O.; et al. A surface with a biomimetic micropattern reduces colonization of Mycobacterium abscessus. FEMS Microbiol. Lett. 2014, 360, 17–22, doi:10.1111/1574-6968.12587.

56. Holm, D.K.; Petersen, L.M.; Klitgaard, A.; Knudsen, P.B.; Jarczynska, Z.D.; Nielsen, K.F.; Gotfredsen, C.H.; Larsen, T.O.; Mortensen, U.H. Molecular and chemical characterization of the biosynthesis of the 6-MSA- derived meroterpenoid yanuthone D in Aspergillus niger. Chem. Biol. 2014, 21, 519–529, doi:10.1016/j.chembiol.2014.01.013.

57. Isbrandt, T.; Tolborg, G.; Ødum, A.; Workman, M.; Larsen, T.O. Atrorosins: a new subgroup of Monascus pigments from Talaromyces atroroseus. Appl. Microbiol. Biotechnol. 2020, 104, 615–622, doi:10.1007/s00253- 019-10216-3.

58. Lee, J.S.; Kim, Y.S.; Park, S.; Kim, J.; Kang, S.J.; Lee, M.H.; Ryu, S.; Choi, J.M.; Oh, T.K.; Yoon, J.H. Exceptional production of both prodigiosin and cycloprodigiosin as major metabolic constituents by a novel marine bacterium, Zooshikella rubidus S1-1. Appl. Environ. Microbiol. 2011, 77, 4967–4973, doi:10.1128/AEM.01986-10.

59. Couturier, M.; Bhalara, H.D.; Chawrai, S.R.; Monson, R.; Williamson, N.R.; Salmond, G.P.C.; Leeper, F.J. Substrate Flexibility of the Flavin-Dependent Dihydropyrrole Oxidases PigB and HapB Involved in Antibiotic Prodigiosin Biosynthesis. ChemBioChem 2020, 21, 523–530, doi:10.1002/cbic.201900424.

1 Supplementary Materials

2 Chitin degradation machinery and secondary metabolite 3 profiles in the marine bacterium Pseudoalteromonas rubra 4 S4059

6 Xiyan Wang 1, Thomas Isbrandt 1, Mikael Lenz Strube 1, Sara Skøtt Paulsen 1, Maike Wennekers 7 Nielsen 1, Yannick Buijs 1, Erwin M. Schoof 1, Thomas Ostenfeld Larsen 1, Lone Gram 1 and 8 Shengda Zhang 1 *

9 1 Department of Bioengineering, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark; 10 [email protected] (X.W.); [email protected] (T.I.); [email protected] (M.L.S.); [email protected] (S.S.P.); [email protected] 11 (M.W.N.); [email protected] (E.M.S.); [email protected] (Y.B.); [email protected] (T.O.L); [email protected] (L.G.). 12 * Correspondence: [email protected] (S.Z.); Tel.: +45-5011-7765. 13 14 * Corresponding author: 15 Sheng-Da Zhang 16 Department of Biotechnology and Bioengineering, Technical University of Denmark, Søltofts Plads 17 bldg. 221, DK-2800 Kgs. Lyngby, Denmark 18 E-mail: [email protected] Telephone: +45-50117765 19 20 Running title: Bioactive potential of Pseudoalteromonas rubra S4059

22 Supplementary Results 23 Anti-fungal activity 24 P. rubra S4059 WT and ΔGH19 were tested against Aspergillus niger in an antifungal assay. Both strains 25 were antifungal against A. niger. However, there is no signficant difference between WT and ΔGH19 26 and neither of them could retain the antifungal activity over time. 27 28 Table and Figure legends 29 30 Table S1. The predicted biosynthetic gene clusters (BGCs) of Pseudoalteromonas rubra S4059 by 31 antiSMASH 6.0.

32 Table S2. The fold change of six significantly upregulated chitinolytic enzymes in cell or supernatant 33 samples.

34 Table S3. Significantly up-and down-regulated proteins in Pseudoalteromonas rubra S4059 proteome 35 when grown on chitin as compared to on mannose. 36 Table S4. Antibiotic sensitivity of Pseudoalteromonas rubra S4059 growth on MB agar plate. 37 Table S5. Bacteria and plasmids used in this study. 38 Table S6. Primers used in this study. 39 Table S7. The normalized protein abundance of chitinolytic enzymes in S4059.

40 Figure S1. (A) Base peak chromatograms of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 41 mutant when cultivated on marine minimal medium using mannose, crystalline chitin, colloidal 42 chitin or N-acetyl glucosamine (NAG) as the carbon source. The red pigment prodigiosin and two of 43 its analogues (hexyl prodigiosin and heptyl prodigiosin) could be identified in the extract and 44 confirmed based on MS/MS experiments and the acquired absorption spectra. Experiments were done 45 in biological triplicates, and a reference chromatogram of the sterile growth medium is included to 46 show media components also present in the experiments. (B) Tandem MS spectra recorded for 1) 47 prodigiosin, 2) hexyl-prodigiosin, and 3) heptyl-prodigiosin. The prodigiosin MS/MS spectra matches 48 our in-house MS/MS library, and the characteristic fragment m/z 252.11, and loss of CH3 (m/z 15.02) 49 additionally matches those previously reported for prodigiosin and analogues [5]. (C) Recorded UV- 50 Vis absorption spectrum of prodigiosin. All proposed prodigiosins share identical absorption spectra. 51 The spetrum is in agreement with previously reported literature [6].

52 Figure S2. (A-D) Growth kinetics of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 mutant 53 in mannose (A), colloidal chitin (B), crystalline chitin (crab chitin) (C) and NAG (chitin monomer) (D) 54 containing MMM with casamino acids. (E-F) WT and ΔGH19 growth in crystalline chitin (E) or 55 colloidal chitin containing MMM without casamino acid. Square: WT; Triangle: ΔGH19. The points 56 are bio-replicates and error bars are standard deviation. (G-H) Chitin degradation activities of wild 57 type and the mutant on colloidal chitin plate (G) and crystalline chitin plate (H). 58 Figure S3. Biofilm formation on the microtiter-well plastic surface of Pseudoalteromonas rubra S4059 59 wild type and ΔGH19 mutant in four different sole carbon contained medium as determined by the 60 O’Toole & Kolter crystal violet assay. (A) in mannose; (B) in NAG (chitin monomer); (C) in colloidal 61 chitin; (D) in crab chitin. Each experiments are repeated in bio-triplicates and error bars are standard 62 deviation. 63

64 SUPPLEMENTARY RESULTS

65 Table S1. The predicted biosynthetic gene clusters (BGCs) of Pseudoalteromonas rubra S4059 by antiSMASH 6.0.

Type From To Most similar known cluster Similarity chrI BGC1 RRE-containing 484,852 504,385 BGC2 NRPS-like 665,623 707,167 BGC3 NRPS 1,025,623 1,124,309 BGC4 NRPS-like 1,152,132 1,194,273 indigoidine NRP 40% BGC5 NRPS 1,204,311 1,261,426 BGC6 T3PKS 1,684,665 1,725,708 BGC7 hserlactone 2,486,553 2,507,302 BGC8 RiPP-like 2,814,767 2,824,604 BGC9 prodigiosin 3,203,980 3,239,003 prodigiosin 70% NRP + Polyketide: Modular BGC10 NRPS,T1PKS,transAT-PKS 3,627,019 3,764,918 kalimantacin A type I + Polyketide: Trans-AT 10% type I

NRPS,T1PKS,betalactone,thioamide- amonabactin P BGC11 4,235,481 4,358,990 NRP 42% NRP 750 BGC12 lanthipeptide-class-i 4,533,397 4,557,768 BGC13 RiPP-like 4,561,329 4,572,171 chrII BGC14 lanthipeptide-class-iv 165,032 187,755 BGC15 NRPS,T1PKS 209,321 279,019 BGC16 RiPP-like 629,147 639,983 BGC17 NRPS 864,063 1,000,602 BGC18 RRE-containing 1,025,566 1,047,998

BGC19 NRPS 1,052,424 1,095,471 66

67 Table S2. The fold change of six significantly upregulated chitinolytic enzymes in cell or supernatant 68 samples.

Glycoside hydrolase type Log2(Fold change) cells samples GH18-1 13.5 GH18-2 9.76 GH18-3 6.89 GH18-4 14.36 GH20-1 8.86 GH20-2 2.74 supernatant samples GH18-1 13.27 GH18-2 7.71 GH18-3 9.02 GH18-4 15.97 GH20-1 6.49 GH20-2 7.46 70

72 Table S3. Significantly up-and down-regulated proteins in Pseudoalteromonas rubra S4059 proteome 73 when grown on chitin as compared to on mannose. 74

Function Log2(Fold change) Accession Protein Pilus +2.80 A0A5S3V0R4 Pilin biogenesis protein assembly +1.97 A0A0U2P9L9 Pilus assembly protein PilM +2.68 A0A5S3UZ53 Pilus assembly protein PilN +1.65 A0A0L0EQN7 Pilus assembly protein PilP +2.65 A0A5S3UZ07 Pilus assembly protein PilP +2.47 A0A5S3V1T9 Pilus assembly protein PilW +3.10 A0A5S3V0T8 Pilus assembly protein PilX +2.62 A0A5S3UQS3 Pilus assembly protein PilZ +2.01 A0A5S3UWH8 PilZ domain-containing protein +1.86 A0A5S3URP5 PilZ domain-containing protein +5.68 A0A5S3URS7 PilZ domain-containing protein Chemotaxis +3.09 A0A0L0EQP9 Chemotaxis protein CheX and Flagellar assembly +4.43 A0A0U3HPG4 Flagellar motor switch protein FliG +2.82 A0A0L0ERK8 Flagellar motor switch protein FliN +4.19 A0A0L0EU84 Chemotaxis protein CheY +3.27 A0A0F4QWI4 Chemotaxis protein CheY +2.80 A0A0L0EV12 Chemotaxis protein CheY +3.25 A0A0U3I1F8 Chemotaxis protein CheV +2.73 A0A0L0EW69 Fis family transcriptional regulator +1.81 A0A0F4QWB6 Chemotaxis protein CheW +1.26 A0A0F4QWN8 Flagellar basal-body rod protein FlgG +4.90 A0A0L0ERQ9 Flagellar motor switch protein FliM +4.17 A0A5S3UYM4 Flagellar assembly protein FliH +3.85 A0A5S3UTL8 Flagellar biogenesis protein +3.62 A0A5S3UXB9 Flagellar M-ring protein +3.52 A0A5S3UX71 Flagellar protein FliL +2.84 A0A5S3UQH6 Flagellar motor protein MotA +2.06 A0A5S3UTK5 Flagellar biosynthesis protein FlgP +2.05 A0A0L0EPQ6 Flagellar motor protein PomA +1.89 A0A5S3UX07 Flagellar hook protein FlgE +1.66 A0A5S3UWZ7 Flagellar L-ring protein FlgH +1.48 A0A0L0ERW2 Anti-sigma-28 factor FlgM +2.91 A0A0L0EUK7 Twitching motility protein PilT +3.04 A0A5S3V634 Methyl-accepting chemotaxis protein +3.11 A0A5S3V3M4 Methyl-accepting chemotaxis protein +4.18 A0A5S3V3J7 Methyl-accepting chemotaxis protein +2.46 A0A5S3V0Y1 Methyl-accepting chemotaxis protein +2.75 A0A5S3V0M8 Methyl-accepting chemotaxis protein

+2.12 A0A5S3V0H3 Methyl-accepting chemotaxis protein +2.34 A0A5S3UZH1 Methyl-accepting chemotaxis protein +3.91 A0A5S3UXP0 Methyl-accepting chemotaxis protein +3.08 A0A5S3UXF8 Methyl-accepting chemotaxis protein +2.82 A0A5S3UWG2 Methyl-accepting chemotaxis protein +4.11 A0A5S3UVB3 Methyl-accepting chemotaxis protein +5.61 A0A5S3UU56 Methyl-accepting chemotaxis protein +1.92 A0A5S3UTY6 Methyl-accepting chemotaxis protein +1.91 A0A5S3UTH7 Methyl-accepting chemotaxis protein +2.51 A0A5S3UTB3 Methyl-accepting chemotaxis protein +2.45 A0A5S3UR67 Methyl-accepting chemotaxis protein +3.76 A0A5S3URP8 Methyl-accepting chemotaxis protein Cell division +1.93 A0A5S3V4C6 Cell division ATP-binding protein FtsE +2.68 A0A5S3V008 Cell division coordinator CpoB +2.31 A0A0L0EW35 Cell division inhibitor MinD +3.17 A0A5S3UZ24 Cell division protein DamX +3.18 A0A0L0EVX6 Cell division protein FtsA +3.41 A0A5S3V4C7 Cell division protein FtsX +1.38 A0A5S3UW61 Cell division protein FtsX +2.71 A0A5S3V1E1 Cell division protein FtsZ +2.78 A0A0L0EWQ7 Cell division protein ZapB +2.80 A0A5S3UQ38 Cell division protein ZipA +2.40 A0A0F4QML1 Cell division topological specificity factor Core +1.70 A0A0L0EMS8 Methylmalonate-semialdehyde metabolism dehydrogenase +2.45 A0A0F4QGS7 Ribose-phosphate pyrophosphokinase +2.90 A0A0U2Y0N0 Shikimate kinase +2.18 A0A0U3IDX6 Tryptophan synthase beta chain +2.80 A0A0L0ESU1 S-adenosylmethionine synthase +2.81 A0A0L0EN60 Succinate--CoA ligase [ADP-forming] subunit beta +3.39 A0A0L0EX71 Phosphoribosylformylglycinamidine cyclo- ligase +2.04 A0A0L0ETQ3 CoA transferase subunit A +2.46 A0A0L0ENB5 Dihydrolipoyllysine-residue succinyltransferase component of 2- oxoglutarate dehydrogenase complex +3.28 A0A0L0EP99 3-ketoacyl-ACP reductase Type II +2.41 A0A5S3UQN2 Type II secretion system core protein G secretion +4.51 A0A5S3UT29 Type II secretion system protein F +2.69 A0A5S3US13 Type II secretion system protein GspC +2.11 A0A5S3UR30 Type II secretion system protein GspD +3.58 A0A5S3UQK8 Type II secretion system protein J +3.00 A0A5S3URY9 Type II secretion system protein K +1.88 A0A5S3UQK7 Type II secretion system protein L

+1.90 A0A5S3UQZ6 Type II secretion system protein M +3.32 A0A5S3V4K1 Type IV pili twitching motility protein PilT OS mannosidase -5.55 A0A5S3URX4 Sugar hydrolase

75 -: downregulate; +; upregulate. 76

77 Table S4. Antibiotic sensitivity of Pseudoalteromonas rubra S4059 growth on MB agar plate.

+: sensitivity. -: insensitivity. NT: not tested

Antibiotics 10μg/mL 30μg/mL 50μg/mL 100μg/mL

Ampicillin NT NT - +

Kanamycin NT NT - +

Chloramphenicol - + + NT

Erythromycin + + + NT

Gentamycin NT NT - +

79 Strains, plasmids and primers

80 Table S5. Strains and plasmids used in this study.

Reference Strains/plasmids Genotype or relevant characteristics or source Bacterial strains Escherichia coli WM3064 thrB1004 pro thi rpsL hsdS lacZΔM15 RP4-1360 Δ(araBAD)567 ΔdapA1341::[erm pir], 37 ℃. Donner strains in conjuation. [1] GB dir pir116 An arabinose-inducible ETγA operon (full- length recE, recT, redγ, and recA), a copy-up pir116 gene. Host strain for constructing suicide plasmids with R6K replication origin. [2] TOP 10 F- mcrA Δ(mrr-hsdRMS-mcrBC) Φ80lacZ ΔM15 Δ lacX74 recA1 araD139 Δ(araleu)7697 galU galK rpsL (StrR) endA1 nupG. InvitrogenTM Pseudoalteromonas rubra S4059 Wild type strain, Isolated from seaweed. [3] ΔGH19 GH19 chitinase gene in-frame deletion mutant of P. rubra S4059 ΔGH19 This study Plasmids pDM4 Suicide vector for targeted mutagenesis; sacB; oriR6K; CmR [4]

pDM4 –del-GH19 The left arm and right arm DNA region of the GH19 chitinase gene were PCR amplified from P. rubra S4059 genome and cloned in the pDM4 This study plasmid by direct cloning

81 KanR: Kanamycin resistance; CmR: Chloramphenicol resistance 82 83 84

85 Table S6. Primers used in this study.

Primer Sequence Description Expected size (bp) GH19-L-F GCtctagaACTCAATAATCCACTAAA Left arm of GH19 1032 GCC GH19-L-R ATTAAAGCCTAAAAAAGGACATC CTTTACATGTTG GH19-R-F TGTAAAGGATGTCCTTTTTTAGGCT Right arm of GH19 1014 TTAATTTTACTTCG GH19-R-R CCGctcgagAGGGTTCCTTTGTACTTA AAC CmR F GGCATTTCAGTCAGTTGCTC Detection of CmR gene 525

CmR R CCATCACAAACGGCATGATG

GH19-pJet1.2-F CCACATGTGGAATTGTGAGCTAAA GH19 homologous 1914 GCCGCCACCAATGATG fragments GH19-pJet1.2-R CTTATCGATACCGTCGACCCGGCA GACCCAGAATAAGCG GH19-pDM4-F CATCATTGGTGGCGGCTTTAGCTC pDM4 plasmid (GH19) 7054 ACAATTCCACATGTGG GH19-pDM4-R CGCTTATTCTGGGTCTGCCGGGTCG ACGGTATCGATAAG GH19-p 1 GGGTTCCTTTGTACTTAAAC Confirmation of GH19 1st: P1, P4 mutants 2452a; 1012b GH19-p 2 CTTCAAGGACACATACCTTC P2, P3 2577a bp; 1137b bp GH19-p 3 CTATTCAGGCTACCAAAGC 2nd: P1, P2 3487a bp; 2047b bp GH19-p 4 CAACAACATGTAAAGGATGTC

86 a the size of WT; b the size of mutant.

88 Table S7. The normalized protein abundance of chitinolytic enzymes in S4059.

chi-WT chi-WT cells chi-WT chi WT chi WT chi WT man-WT man-WT man-WT man-WT man-WT man-WT Accession Description cells 1 2 cells 3 sup 1 sup 2 sup 3 cells 1 cells 2 cells 3 sup 1 sup 2 sup 3 A0A5S3USE2 GH18 32.06327 32.61660578 33.4121 37.44231 37.65211 37.49193 20.48135 20.43851 22.67027 28.92823 24.77495 25.07364 A0A5S3V351 GH18 20.6461 20.96568668 20.22428 23.46404 23.16674 23.25328 NA NA NA NA NA NA A0A5S3V0U4 GH18 28.15098 27.67043822 29.01734 34.34522 34.25175 33.45857 NA NA NA NA NA NA A0A5S3V6T3 GH18 26.33148 26.75637076 28.43113 32.76361 32.03789 31.64236 18.73021 NA 21.37777 26.46891 26.48377 26.36123 A0A5S3V3K3 GH18 27.80207 28.00121752 30.02482 36.11999 35.99521 35.4816 23.64874 24.07021 23.43214 28.80977 29.00245 28.72435 A0A5S3USH6 GH18 30.77192 30.92668926 32.04919 37.43495 37.65743 37.04639 NA 17.87567 NA 23.19466 23.32522 23.6994 A0A5S3V378 GH18 21.42957 21.86235625 21.93963 27.15255 26.57559 26.68415 NA NA NA NA NA NA A0A5S3UX38 GH19 19.85311 19.99490452 20.16807 26.66788 26.37509 22.88288 NA NA NA NA NA NA A0A5S3V1X9 GH20 28.35271 27.9787318 28.45681 24.3699 23.91617 25.1039 21.347 21.34887 21.51012 21.78837 NA 18.85425 A0A5S3UX95 GH20 28.62493 28.18178963 29.02698 27.56267 27.05506 26.8132 27.99074 27.8446 27.78883 18.63304 21.84458 24.56435 A0A5S3UV09 GH20 21.22181 21.59476677 22.67991 NA NA NA NA NA NA NA NA NA A0A5S3UTD1 AA10 25.19116 25.3838232 27.22621 31.44592 31.29765 30.12035 NA NA 21.18536 21.92399 21.03666 NA 89

93 94 Figure S1. (A) Base peak chromatograms of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 95 mutant when cultivated on marine minimal medium using mannose, crystalline chitin, colloidal 96 chitin or N-acetyl glucosamine (NAG) as the carbon source. The red pigment prodigiosin and two of 97 its analogues (hexyl prodigiosin and heptyl prodigiosin) could be identified in the extract and 98 confirmed based on MS/MS experiments and the acquired absorption spectra. Experiments were done 99 in biological triplicates, and a reference chromatogram of the sterile growth medium is included to 100 show media components also present in the experiments. (B) Tandem MS spectra recorded for 1) 101 prodigiosin, 2) hexyl-prodigiosin, and 3) heptyl-prodigiosin. The prodigiosin MS/MS spectra matches

102 our in-house MS/MS library, and the characteristic fragment m/z 252.11, and loss of CH3 (m/z 15.02) 103 additionally matches those previously reported for prodigiosin and analogues [5]. (C) Recorded UV- 104 Vis absorption spectrum of prodigiosin. All proposed prodigiosins share identical absorption spectra. 105 The spetrum is in agreement with previously reported literature [6].

106

107 108 Figure S2. (A-D) Growth kinetics of Pseudoalteromonas rubra S4059 wild type (WT) and ΔGH19 mutant 109 in mannose (A), colloidal chitin (B), crystalline chitin (crab chitin) (C) and NAG (chitin monomer) (D) 110 containing MMM with casamino acids. (E-F) WT and ΔGH19 growth in crystalline chitin (E) or 111 colloidal chitin containing MMM without casamino acid. Square: WT; Triangle: ΔGH19. The points 112 are bio-replicates and error bars are standard deviation. (G-H) Chitin degradation activities of wild 113 type and the mutant on colloidal chitin plate (G) and crystalline chitin plate (H). 114 115 116

117

118 119 Figure S3. Biofilm formation on the microtiter-well plastic surface of Pseudoalteromonas rubra S4059 120 wild type and mutants in four different sole carbon contained medium as determined by the O’Toole 121 & Kolter crystal violet assay. (A) in mannose; (B) in NAG (chitin monomer); (C) in colloidal chitin; (D) 122 in crab chitin. Positive control is Phaeobacter inhibens; negative control is sterile media. Each 123 experiments are repeated in bio-triplicates and error bars are standard deviation. 124

SUPPLEMENTARY EXPERIMENTAL PROCEDUR

Antibiotic sensitivity assay for selection marker. The antibiotic sensitivity of P. rubra S4059 to five antibiotics was tested: Ampicillin (Sigma, A9518), Kanamycin (Sigma, K4000), Chloramphenicol (Sigma, C0378), Erythromycin (Sigma, E6376), Gentamycin (Sigma, G3632). P. rubra S4059 was grown in MB at 25 °C to late exponential phase and a 10-fold dilution series plated on MA plates with antibiotics in the final concentration of 10 µg/mL, 30 µg/mL, 50 µg/mL, 100 µg/mL. The plates were incubated at 25 °C for 48 h and inspected for colonies. The assay was performed in triplicate.

Electroporation of E. coli strains. The electroporation procedure was modified from Wang et al [2]. Briefly, a pre-culture was grown and diluted 100 times in 5 mL fresh LB and grown to OD600≈0.4. 1.5 mL cells were harvested at 6,000xg for 1 min. The cells were washed twice in 1 mL MilliQ H20 and harvested at 6,000xg for 1 min. Around 200 ng DNA was added to the harvested cells and MilliQ H20 was added to a final volume of 30 µL. The mixture was placed in a 1-mm electroporation cuvette and electroporated at 1,200 V. Cells were resuspended in 1 mL LB Broth without antibiotics immediately after electroporation and incubated in a 1.5-mL Eppendorf® thermomixer comfort at 37 oC, 750 rpm for 1 h. The recovered cells were spread on selection plates with antibiotics and incubated at 37 °C, overnight. Chitin degradation. The protocol was modified from Paulsen et al. [7]. The medium consisted of 2% sea salt (Sigma, S9883), 1.5% agar, 0.3% casamino acid and 0.2% chitin (crystalline chitin from crab and colloidal chitin). Briefly, all strains (wild type and ΔGH19 mutant) were grown in MB at 25 °C, overnight. The pre-cultures were diluted 100 times in 5 mL fresh MB and grown to OD600=0.1. Twenty-µL culture was spotted on chitin plates. The plates were incubated at 25 °C for 7 days. A qualitative grading of chitinase activity was measured as previous description [7]. Biofilm formation assay. Biofilm formation was evaluated in 96-well polystyrene plates (Thermo scientific, 163320) with crystal violet staining as previous reported [8]. Cells were incubated with 5 mL MB in 50 mL tubes at 25 °C, 200 rpm overnight. The overnight cultures were diluted to OD600=0.01 in MMM with different carbon sources (the same medium used for growth kinetics) in a volume of 0.5 mL. To establish the biofilm, 100 µL liquid from the diluted culture were added into each well in triplicate. The plates were incubated at 25 °C for 72 hours in a humidity chamber. The biofilm was visualized by crystal violet staining and was dissolved in 95% ethanol to measure the absorbance at 590 nm. Phaeobacter inbibens DSM17395 was used as positive control and fresh medium was used as negative control. Anti-fungal activity assay. Anti-fungal activity assay was done with a modified protocol from Paulsen et al. 2016 [7]. In brief, Aspergillus niger (IBT 32191) was precultured from IBT Culture Collection at Department of Biotechnology and Biomedicine, Technical University of Denmark. A volume of 20 µL spore suspension of A. niger was added to a puncture well in the center of a MA plate. Then, 20 µL of overnight culture of P. rubra S4059 in MB was spotted 2 cm away from the center of well. Due to slow growth, we also tested that the spore suspension of A. niger was added 4 days prior to the S4059 culture. Plates from both settings were incubated and checked the size of inhibition zone every day for 14 days. Preparation of desks from shrimp shells. Strip the shells off the shrimps carefully by hand and put it into 1L beaker with fresh dH20. After striping all of shells, a teeth brush was used for cleaning the shrimp shells under flowing water to make sure no tissues (or meat) left on the shell. Then put the cleaned shrimp shells into a new 1L beaker with fresh dH20 for washing. Tear the shrimp shells and use the puncher machine to punch the shrimp shells out. Remember each piece of shrimp shell should only be used for punching once and only pick the perfect circular shrimp shells (do not use the gaped shells or broken shells). Put the punched shrimp shells into 100 mL bottles with 30 mL, 3% Sea salts (Sigma, catalog number S9883). Do not let them dry. Otherwise, they will break easily. The shells in 3% Sea salts solution were autoclaved at 121 ℃ for 15 min and stored at 4 ℃.

References

1. Dehio, C.; Meyer, M. Maintenance of broad-host-range incompatibility group P and group Q plasmids and transposition of Tn5 in Bartonella henselae following conjugal plasmid transfer from Eescherichia coli. J. Bacteriol. 1997, 179, 538–540, doi:10.1128/jb.179.2.538-540.1997.

2. Wang, H.; Li, Z.; Jia, R.; Hou, Y.; Yin, J.; Bian, X.; Li, A.; Müller, R.; Stewart, A.F.; Fu, J.; et al. RecET direct cloning and Redαβ recombineering of biosynthetic gene clusters, large operons or single genes for heterologous expression. Nat. Protoc. 2016, 11, 1175–1190, doi:10.1038/nprot.2016.054.

3. Gram, L.; Melchiorsen, J.; Bruhn, J.B. Antibacterial activity of marine culturable bacteria collected from a global sampling of ocean surface waters and surface swabs of marine organisms. Mar. Biotechnol. 2010, 12, 439–451, doi:10.1007/s10126-009-9233-y.

4. Milton, D.L.; O’Toole, R.; Hörstedt, P.; Wolf-Watz, H. Flagellin A is essential for the virulence of Vibrio anguillarum. J. Bacteriol. 1996, 178, 1310–1319, doi:10.1128/jb.178.5.1310-1319.1996.

5. Lee, J.S.; Kim, Y.S.; Park, S.; Kim, J.; Kang, S.J.; Lee, M.H.; Ryu, S.; Choi, J.M.; Oh, T.K.; Yoon, J.H. Exceptional production of both prodigiosin and cycloprodigiosin as major metabolic constituents by a novel marine bacterium, Zooshikella rubidus S1-1. Appl. Environ. Microbiol. 2011, 77, 4967–4973, doi:10.1128/AEM.01986-10.

6. Couturier, M.; Bhalara, H.D.; Chawrai, S.R.; Monson, R.; Williamson, N.R.; Salmond, G.P.C.; Leeper, F.J. Substrate Flexibility of the Flavin-Dependent Dihydropyrrole Oxidases PigB and HapB Involved in Antibiotic Prodigiosin Biosynthesis. ChemBioChem 2020, 21, 523–530, doi:10.1002/cbic.201900424.

7. Paulsen, S.S.; Andersen, B.; Gram, L.; MacHado, H. Biological potential of chitinolytic marine bacteria. Mar. Drugs 2016, 14, doi:10.3390/md14120230.

8. O’Toole, G.; Kolter, R. Initiation of biofilm formation in Pseudomonas fluorescens WCS365. Mol. Microbiol. 1998, 28, 449–461.

Paper 2

Identification and verification of the prodigiosin biosynthetic gene cluster (BGC) in Pseudoalteromonas rubra S4059.

Identification and verification of the prodigiosin biosynthetic gene cluster (BGC) in Pseudoalteromonas rubra S4059

Xiyan Wang 1, Thomas Isbrandt 1, Emil Ørsted Christensen 1, Thomas Ostenfeld Larsen 1, Shengda Zhang 1 and Lone Gram 1 *

1 Department of Bioengineering, Technical University of Denmark, DK-2800 Kgs. Lyngby,

Denmark; [email protected] (X.W.); [email protected] (T.I.); [email protected] (E. Ø. C); [email protected] (T.O.L); [email protected] (S.Z.) * Correspondence: [email protected] (L.G.); Tel: +45-4525-2586

Abstract: Pseudoalteromonas rubra S4059 devotes up to 15% of its genome to biosynthetic gene clusters (BGCs) and until now, only prodigiosin has been chemically identified from this strain. Prodigiosin has attracted much attention due to its pharmaceutical and industrial potential. The purpose of this study is to identify the prodigiosin BGC in Pseudoalteromonas rubra S4059 and further to explore whether this strain can produce other bioactive compounds that may be masked by prodigiosin. A putative prodigiosin synthesizing transferase PigC was targeted and a pigC in-frame deletion mutant was generated in S4059. Deletion of the pigC gene resulted in the prodigiosin deficiency in S4059. Further, extractions from the culture of the pigC mutant retained antibacterial activity. Bioassay-guided fractionation demonstrated that two blue fractions were obtained from the mutant cultures and both had antibacterial activity. A precursor of prodigiosin, 4-methoxy-2,2’- bipyrrole-5- carbaldehyde (MBC), was the dominant compound in both the fractions and a stable blue pigment, dipyrrolyldipyrromethene prodigiosin, was identified from the two fractions. This is the first report of such a blue pigment produced by Pseudoalteromonas rubra. In conclusion, a prodigiosin BGC was identified and verified genetically and chemically in P. rubra S4059 and a stable blue pigment was isolated from the pigC mutant of S4059, suggesting that this strain has a large potential on pharmaceutical and industrial potential. Keywords: Pseudoalteromonas rurba, prodigiosin, BGC, dipyrrolyldipyrromethene prodigiosin.

Introduction: Many secondary metabolites from microorganisms and plants have bioactivities of pharmaceutical and agricultural interest. For instance, more than 60% of the antibiotics used in the clinic have been derived from microorganisms, notably soil-dwelling Actinobacteria (Barka et al., 2016). The current antibiotic crisis is driven both by the rapid development and spread of antibiotic resistance in pathogenic bacteria and by the lack of discovery of novel antibiotic compounds (C. Lee Ventola, 2015). We are thus in urgent need of discovering novel antibiotics, and exploring microorganisms from new environments is one of the promising strategies to uncover novel antibiotics. Although many bioactive compounds have been discovered from marine (micro)organisms, such as antibiotics from the marine genus Pseudoalteromonas, the marine environment remains a less explored drug discovery reservoir as compared to soil.

Pseudoalteromonas species are strictly marine bacteria and can be divided into two groups: pigmented and non-pigmented strains (Bowman, 2007) and especially the pigmented group are producers of bioactive compounds (Bowman, 2007; Vynne et al., 2011). Recently, genome mining on several Pseudoalteromonas species indicated that the genetic potential for secondary metabolite production of pigmented species is equal to that of bacteria from the Actinobacteria. For instance, a red pigmented P. rubra strain S4059 dedicates as much as 15% of its genome to genes likely involved in secondary metabolite production and contains 17 biosynthetic gene clusters (BGCs). However, only one compound class, namely prodigiosins has been chemically identified in this strain, and the biosynthetic gene cluster responsible for the production of this red pigment in S4059 is not known (Vynne et al., 2011).

Prodigiosins are members of the prodiginine family, which is characterized by a tripyrrole structure. Prodigiosin consists of two moieties, 2-methyl-3-n-amyl-pyrrole (MAP) and 4- methoxy-2,2’-bipyrrole-5-carbaldehyde (MBC), which are linked via a condensation reaction to from the final product. Several prodigiosin analogues have been described, mostly varying in the length of the aliphatic side chain on the MAP moiety (Kim et al., 2007). The two moieties are coupled by a synthesizing transferase pigC in a final condensation step to form prodigiosin (Williams, 1973). Prodigiosin was first discovered from a terrestrial bacterium Serratia marcescens (Kalesperis et al., 1975), and has since been found in several other bacterial species, such as Hahella chejuensis (Kim et al., 2007), Streptomyces coelicolor (Kawasaki et al.,

2009), and Pseudoalterononas rubra (Vynne et al., 2011), and has attracted attention due to its multiple activities, such as antibacterial (OKAMOTO et al., 1998), anticancer (Kawauchi et al., 2008), algicidal (Zhang et al., 2016), and immunosuppressive activity (Kawauchi et al., 2007). The prodigiosin derivatives are also of interest, for instance, cycloprodigiosin which was isolated from P. rubra and is more active against Staphylococcus aureus than prodigiosin (Setiyono et al., 2020). Interestingly, most of the prodigionsins are red pigments, however, dipyrrolyldipyrromethene prodigiosin (Dip-PDG), which has only been reported in few bacterial species, such as Hahella chejuensis (Kim et al., 2007), and a mutant Serratia marcescens strain (Wasserman et al., 1968) is a blue pigment.

Although prodigiosin was reported to exhibit cytotoxicity on eukaryotic cells (chick embryos) (Kalesperis et al., 1975) and was presumed not suitable as a clinical drug, more recently, prodigiosin has received renewed attention because prodigiosin and its derivatives are effective proapoptotic agents against cancer cell lines with no or little toxicity on normal cell lines (Darshan and Manonmani, 2015). This finding indicated that prodigiosin is a promising drug candidate and further identification of the BGC is the prerequisite to do heterologous expression for efficient production of this compound. Therefore, a pigmented P. rubra S4059 which can produce prodigiosin was selected to identify the prodigiosin BGC and further to explore whether this strain could produce other bioactive compounds.

Results A prodigiosin BGC was identified and verified in Pseudoalteromonas rubra S4059. Vynne et al. (2011) have demonstrated that P. rubra S4059 can produce the red pigment prodigiosin (Vynne et al., 2011). However, the BGC responsible for prodigiosin production in S4059 remains unknown. A putative prodigiosin synthesizing transferase PigC was annotated in S4059 by Prokka (Seemann, 2014) and its surroundings genes are putative prodigiosin biosynthesis associated genes. Further, antiSMASH 6.0 was used to tentatively predict the prodigiosin BGC in S4059 and the predicted prodigiosin BGC is the same as annotated by Prokka. To further identify the prodigiosin BGC in S4059, a comparison was performed between the putative prodigiosin associated genes in S4059 with known prodigiosin BGC in Serratia sp. ATCC 39006 (Figure 1, Table S1). The prodigiosin BGC in ATCC 39006 contains 14

genes that are arranged pigA through to pigO (Figure 1, A) (Williamson et al., 2006), while the putative prodigiosin BGC in S4059 consists of 13 genes that are arranged pigA through to pigM (Figure 1, A). The pig genes are putatively assigned similar to the genes in the ATCC 39006 pig gene cluster, except that no homologs of pigN and pigO were present (Figure 1, A). The relatedness between each of the 13 prodigiosin proteins found in S4059 and their representative homologs in the prodigiosin BGC of ATCC 39006 were compared (Table S1). Proteins encoded by the genes in the prodigiosin BGC show a wide variation in the level of identity. For example, the ATCC 39006 PigE homologs in S4059 were on average 79.3% identical whilst the PigL homologs on average only were 37.63% identical (Table S1). To verify the prodigiosin BGC in S4059, the putative prodigiosin synthesizing transferase pigC was selected as a target gene to generate an in-frame deletion mutant (Figure 1, B). The phenotype of ΔpigC colonies was slightly brownish, but it did not produce a red pigment (Figure 1 C). The chemical profile of ΔPigC was analysed by LC-MS and prodigiosins were not produced in ΔpigC mutant (Figure 1, D). In conclusion, a prodigiosin BGC was identified and verified in P. rubra S4059 by genomic, genetic, and metabolomic evidence.

Deletion of a pigC gene does not influence the growth and biofilm formation ability of S4059. To explore whether prodigiosin deficiency influences the growth of S4059, wild type and the ΔpigC mutant were cultured in 100 mL marine minimal medium (MMM) with mannose. The deletion of the pigC gene did not influence the growth rate or maximum cell density of S4059 under the condition studied (Figure 2). The biofilm formation of wild type S4059 and the mutant were compared in mannose contained MMM in 96 microtiter plates after 24h, 48h, and 72h growth. Both strains adhered at the same level to the plastic surfaces in the testing media (Figure S1).

Antibacterial activity of wild type S4059 and prodigiosin deficient mutant ΔpigC. To investigate whether S4059 can produce other bioactive compounds than prodigiosin, extracts and culture supernatants of wild type S4059 and ΔpigC (sampled after 24h, 48h, and 72h) were tested for antibacterial activity. Culture extractions from wild type growth after 24h and 48h had antibacterial activity being more inhibitory against S. aures than against V. anguillarum (Figure 3, A, B, C, D). Interestingly, the extractions from the prodigiosin deficient mutant ΔpigC retained a slight antimicrobial activity (Figure 3).

A blue pigment, dipyrrolyldipyrromethene prodigiosin (Dip-PDG), was produced from the prodigiosin deficient mutant ΔPigC. To further explore which compound(s) was/were responsible for the bioactivity in the prodigiosin deficient mutant ΔpigC, a bioassay-guided fractionation was performed. A total of 29 fractions were obtained from the ethyl acetate extract of 500mL of the prodigiosin deficient mutant and used for disc diffusion assays. Two fractions, fraction 11 and 12, had a clear blue color and were the only fractions showing antibacterial activity, this being more pronounced against S. aureus compared to V. anguillarum (Figure 4).

The blue fractions 11 and 12 were analysed by ultra-high performance liquid chromatography coupled to diodearray detection and high resolution mass spectrometry (UHPLC-DAD-HRMS), resulting in similar profiles, although significantly higher concentration in fraction 12. Both fractions contained MBC (meas. m/z = 191.0822, C10H10N2O2, mass accuracy = 3.66 ppm) as the major component. However, MBC has an absorption maximum around 360 nm, which is expected to result in a yellow color (Figure S2). Further inspection of the obtained data from fraction 12 led to identification of two peaks corresponding to the previously reported blue tetrapyrrole pigment, dipyrrolyldipyrromethene prodigiosin (Dip- PDG), formed via condensation of two MBC molecules (Wasserman et al., 1968; Gerber, 1975; Kim et al., 2007), as well as a tentative non-methylated analogue (Dip-PDG: meas. m/z =

335.1509, C19H18N4O2, mass accuracy = 1.79 ppm. Demethyl-Dip-PDG: meas. m/z = 307.1199,

C17H14N4O2, mass accuracy = 2.93 ppm) (Figure S2). Dip-PDG was previously isolated from an MAP pathway blocked mutant of the bacterium Serratia marcescens 9-3-3 and later reported from the wild type of marine bacterium Hahella chejuensis KCTC 2396 (Wasserman et al., 1968; Kim et al., 2007). This strengthens our hypothesis that Dip-PDG and the tentative dimethyl-Dip-PDG are likely responsible for the color of the two blue fractions. Neither Dip- PDG nor the proposed analogue were produced by the wildtype, and only in extremely low amounts in the mutant, thus not allowing for purification and further studies of the compounds.

The filtered supernatants from wild type S4059 and prodigiosin deficient mutant ΔPigC have antibacterial activity. To explore whether some compounds were not extracted with the current extraction method, sterile filtered supernatants from cultures grown in mannose were tested. The cultures were also extracted using ethyl acetate with or without 1% formic

acid and the water phase after extraction was also tested antibacterial activity. Similar antibacterial activity was seen from the extractions with and without formic acid (Figure 5), however, prodigiosin was more efficiently extracted using acidified ethyl acetate (Figure S3). The samples taken from the water phase of the extractions had antibacterial activities (Figure 5), however this activity was lost upon heat treatment of the supernatant (Figure 5).

To further investigate whether this bioactivity was due to proteins or peptides in the spent culture medium, the filtered supernatants were treated with proteinase K or pepsin, neither of which had any effect on the antibacterial activity (Figure S3).

Discussion Prodigiosin and its analogs have medicinal potential due to their anticancer and immunosuppressive properties. Here we identified the prodigiosin BGC including a likely biosynthesis pathway in P. rubra S4059 and this can be useful for future efficient production of this compound for industrial applications. We based the identification of the prodigiosin BGC in Pseudoalteromonas rubra S4059 on genomic, genetic and chemical evidences, and further for the first time reported the production of a stable blue antibacterial pigment, Dip- PGD, in the prodigiosin deficient mutant of Pseudoalteromonas rubra S4059.

Recently, a phylogenetic study on the evolution of the prodigiosin BGC in several bacteria such as the genera Serratia and Pseudoalteromonas was examined by Ravindran et al (Ravindran et al., 2019) and the authors suggested that the these bacteria contain similar proteins in the prodigiosin BGCs originating from a common ancestor. In our study, in silico comparison between putative prodigiosin BGC in S4059 and Serratia sp. ATCC 39006 indicated that a high homology of the pig operon in the genomes of the two strains. This could suggest that the prodigiosin BGC in the two strains was derived from a common ancestor.

The biosynthesis of prodigiosin has been studied in Serratia marcescens and the production involves the coupling step of two precursors: MBC and MAP. The prodigiosin synthesizing transferase PigC is the condensing enzyme in Serratia sp ATCC 39006 (Williamson et al., 2005). In our study, the pigC gene was selected as a target for identifying prodigiosin BGC in S4059 and a pigC in-frame deletion mutant generated in P. rubra S4059. Both of the two precursors can be detected in the ATCC39006 pigC mutant indicating that the deletion of pigC does not disrupt the biosynthesis of the two precursors and only disrupts the formation of prodigiosins (Williamson et al., 2005). Prodigiosins were not detected in extracts from the mutant culture and a high amount of accumulated MBC was detected in the mutant culture extracts. However, MAP was not be detected in the pigC mutant of S4059. That could be due to the volatility of MAP, as reported in another marine bacterium, Hahella chejuensis KCTC 2396 (Kim et al., 2007). Furthermore, two blue fractions obtained by fractionation of the culture of the pigC mutant of S4059 retained antibacterial activity. Chemical analysis showed that MBC was the dominant compound in the two fractions and that both of the two fractions contained low amounts of Dip-PDG as well as a tentative non-methylated analogue. Both MBC and Dip-PDG contain methoxy groups that has been shown to contribute to the bioactivity of

prodigiosin, suggesting that MBC and Dip-PDG are indeed the bioactive compounds (Boger and Patel, 1988; Kim et al., 2007), however, further work is needed to determine the exact nature of their antibacterial activity. Furthermore, the authors have not been able to find any work on the bioactivity of non-methylated prodigiosins.

Antibacterial activity is not only caused by “small molecules” but can also be caused by proteolytic enzymes (Culp and Wright, 2017). For instance, Pseudoalteromonas piscicida can kill competing bacteria by secretion of proteolytic enzymes, such as a trypsin-like serine protease (Richards et al., 2017a) and similar mechanisms of antibacterial activity has also been found in other Pseudoalteromonas species (Longeon et al., 2004). In this study, the filtered aqueous supernatants from both the wild type and the pigC mutant exhibited high antibacterial activity that was lost after heat treatment, suggesting that unknown proteins/peptides or unextractable components with the currently used methods may be responsible for the activities. Three putative RiPP-like proteins were predicted in the genome of S4059 by antiSMASH 6.0 and they all contain a DUF692 domain which always be reported in bacteriocin BGCs (Jackson et al., 2018; Skinnider et al., 2020), suggesting that bacteriocins could be responsible for the antibacterial activity in the supernatants. However, the antibacterial activity of the supernatants was not abolished by treatment with proteinase K or pepsin (Figure S3). This could be because bacteriocins produced by S4059 were not inactivated by treatment with proteinase K and pepsin as reported in several lactic acid bacteria (Lim, 2016). Furthermore, previous proteomic analysis on P. rubra S4059 grown in mannose contained MMM (data are available via ProteomeXchange with identifier PXD023249) indicated that a range of proteases were produced, suggesting that proteases may have the same function as described in P. piscicida and could be responsible for the antibacterial activity in the filtered aqueous supernatants. Further experimental work, e.g. using fluorogenic peptide substrates, will be needed to identify proteases in this strain (Richards et al., 2017b).

In conclusion, we succeeded in identifying the prodigiosin BGC in Pseudoalteromonas rubra S4059 and a stable blue pigment was for the first time reported in the pigC mutant of S4059. These findings provide essential information for further exploring the industrial potential of prodigiosin and its analogs.

Experimental procedures

Bacterial strains, plasmids and cultural conditions. The bacterial strains and plasmids used in this study are listed in Table S2. P. rubra S4059 was isolated during the Galathea 3 expedition (Gram et al., 2010) and cultured on marine agar (MA, BD Difco 2216) or in marine both (MB, BD Difco 2216) at 25 °C. Liquid cultures were incubated with shaking at 25 °C, 200 rpm. P. rubra S4059 wild type and prodigiosin deficient mutant ΔpigC were cultured in marine minimal medium (MMM) (Östling et al., 1991) supplemented with mannose at 25 °C, 200 rpm for chemical extraction. The composition of medium for antibacterial activity assay against Gram-negative bacteria Vibrio anguillarum 90-11-287 and Gram-positive bacteria Staphylococcus aureus 8325 was 1.2 g agar (A0949, ITW reagents), 3.6 g instant ocean salts

(Aquarium systems), 0.4 g casamino acid (BD, 223050) in 120mL distilled H20. The medium was supplemented with 1.2 g peptone (BD, 211677) in assay with Staphylococcus aureus as a target organism.

All Escherichia coli strains were cultured in Luria Bertani broth (BD Difco. 244520) at 37 °C, 200 rpm. E. coli GB dir- pir 116 was used for constructing suicide plasmid (Wang et al., 2016). E. coli WM 3064 is a dapA mutant that requires exogenously supplied diaminopimelic acid (DAP, sigma, D1377) with a final concentration of 0.3μM for growth (Dehio and Meyer, 1997) and was used as a donor strain for intergeneric conjugation. pDM4 was used as the backbone of suicide vector (Milton et al., 1996). E. coli strains harboring pDM4 or its derivatives were grown in LB broth with 10 µg/mL chloramphenicol or on LB agar with 15 µg/mL chloramphenicol.

DNA manipulation. Genomic DNA of P. rubra S4059 was extracted using Genomic DNA buffer set (QIAGEN, 19060) following the supplier instructions. All purified DNA fragments were amplified using PrimeSTAR® Max Premix (TaKaRa, catalog number: R045A). Blue TEMPase Hot Start Master Mix K (catalog number: 733-2584) was used for homologous recombination event checking by PCR. All primers and plasmids were designed in A Plasmid Editor- ApE. The specificity of primers was checked by BLAST against the P. rubra S4059 genome. All primers were synthesized by Integrated DNA Technologies (Leuven, Belgium).

In silico analysis of prodigiosin biosynthesis gene cluster. The genome of P. rubra S4059 is available at the National Center for Biotechnology Information (NCBI) under the accession number CP045429. The genome was annotated by Prokka (Seemann, 2014). Annotated homologous genes (pigA-pigM) of prodigiosin biosynthesis gene cluster (BGC) were identified using CLC Main Workbench 8.0.1 (CLC bio, Aarhus, Denmark) and antiSMASH 6.0 bacterial version.

In-frame deletion of pigC gene in P. rubra S4059. The gene knock-out strategy is followed from Wang et al 2021 (submitted). Briefly, an approximately 1-Kb upstream and downstream region flanking of pigC gene were amplified and fused by overlap extension PCR and the recombining segment was inserted into pDM4 vector to construct suicide plasmid by direct cloning method (Milton et al., 1996; Wang et al., 2016). The suicide plasmid was transferred from E. coli WM 3064 to P. rubra S4059 by intergeneric conjugation. With two times homologous recombination and counter selection, the mutant was identified by PCR and DNA fragments sequencing.

Growth kinetics of P. rubra S4059 wild type and prodigiosin deficient ΔpigC mutant. Pre- culture of S4059 wild type and ΔpigC were incubated in MB overnight at 25°C, 200 rpm. The cultures were diluted to a starting cell density of 103 CFU/mL in 20 mL MMM or 3% sigma sea salt containing 0.2% crystalline chitin. Samples were taken every 2 or 4 hours for estimation of cell density by colony counting on MA plate when S4059 grown in MMM. In chitin medium, samples were taken every day. All experiments were done in biological triplicate.

Biofilm formations of P. rubra S4059 wild type and prodigiosin deficient ΔpigC mutant. The protocol was modified from O’Toole & Kolter crystal violet assay (O’Toole and Kolter, 1998). Briefly, P. rubra S4059 and ΔpigC mutant strain were cultured with MB in 50mL Falcon tubes at 25 °C, 200 rpm overnight. The overnight cultures were diluted to OD600=0.01 using MMM containing mannose in a final volume of 500 µL. Biofilm formation was evaluated in 96 well polystyrene plates (Thermo scientific, 163320) and 100 µL liquid of the diluted cultures were added into each well in bio-triplicates. The plates were incubated at 25 °C for 24, 48 or 72 hours in a humidity chamber. The biofilm was visualized by crystal violet staining and dissolved in 95% ethanol to measure the absorbance at 590 nm. Fresh medium was used as a negative control.

Extraction of metabolites for chemical analysis. The protocol was modified from Wang et al 2020 (Gram et al., 2010). Briefly, P. rubra S4059 and the prodigiosin deficient mutant ΔpigC were cultured in 100 mL MMM or 3% sigma sea salt containing 0.2% crystalline chitin at 25 °C, 200 rpm and samples were taken in 1, 2, 3 days. 20 mL of the culture were extracted with an equal volume of high-performance liquid chromatography (HPLC) grade ethyl acetate (EtOAc) with or without 1% formic acid in 50mL Falcon tubes. The organic phase was transferred to a new Falcon tube and evaporated under nitrogen. The dried extracts were re-dissolved in 200µL methanol and extractions were used for antibacterial activity assay and chemical analysis by UHPLC-DAD-HRMS.

Antibacterial activity assay. The protocol was modified from Gram et al 2010 (Gram et al., 2010). Briefly, the extractions of metabolites were tested for antibacterial activity against Vibrio anguillarum 90-11-287 and Staphylcococcus aureus 8325. The composition of agar medium was described above, and 6 mm wells were punched in the agar plates with a least 2-centimeter distances between the wells. 20 µL of extractions or 40 µL of filtered supernatants were pipetted into the well of the plates for bioassay. To verify whether the antibacterial activity of the supernatants is due to unextracted compounds or proteins/ peptides, the filtered supernatants were also treated with heating at 95°C, 10mins, proteinase K (P6556, Sigma) or pepsin (P6887, Sigma). The plates were incubated at 25 °C for 24-48 h. All experiments were done in biological duplicate.

Fractionation of extract from prodigiosin deficient P. rubra mutant. The extract used for fractionation was made using a modified protocol similar to the one used for chemical analysis. Five 100 mL cultures were combined and extracted using and equal amount EtOAc. The aqueous and organic phases were separated, and the organic phase was evaporated to dryness, yielding 30 mg of crude extract. The extract was redissolved in methanol and fractionated in three 10 mg portions on a Kinetex Core-Shell C18 column (250mm x 10mm, 5µm, Phenomenex), equipped to an Agilent Infinity II 1260 preparative HPLC system. The mobile phase was made up of HPLC grade water (solvent A) and acetonitrile (solvent B) both buffered with 20mM formic acid (FA). Elution was achieved using a flow rate of 5 mL/min and linear gradient starting with 10% B increasing to 100% in 30 min, collecting time slices of one minute each.

Chemical analysis. Chemical analysis was performed using a maXis 3G orthogonal acceleration quadrupole time-of-flight mass spectrometer (QTOF-MS) (Bruker Daltonics) connected to an Ultimate 3000 UHPLC system (Dionex). Separation was achieved using a Kinetex 1.7 μm C18, 100 mm × 2.1 mm column (Phenomenex). The column temperature was maintained at 40°C throughout the analysis, and a linear gradint using LC-MS grade water and acetonitrile both buffered with 20 mM LC-MS grade formic acid, starting from 10% (v/v) acetonitrile and increased to 100% in 10 min, maintaining this rate for 3 min before returning to the starting conditions in 0.1 min and staying there for 2.4 min before the following run. A flow rate of 0.4 ml⋅min−1 was used. Mass spectrometric detection was performed in ESI+ with a data acquisition frequency of 10 scans per second in the m/z range 75-1250. Mass calibration was done calibrated using Bruker Daltonics high precision calibration algorithm by automatically infusing the internal standard sodium formate before each run. UV-Vis absorption spectra were collected at wavelengths from 190 to 800 nm. Data processing and analysis was performed using DataAnalysis 4.0.

Author contributions: XW, SZ and LG conceived and designed experiments; TI performed the chemical analyses supported by TOL; EØC performed in silico comparison. XW wrote the first manuscript draft; SDZ and LG supervised edited and finalized the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding: This research was supported by Chinese Scholarship Council (CSC scholarship No. 201706170066) and the Danish National Research Foundation (DNRF137) for the Center for Microbial Secondary Metabolites, the Independent Research Fund Denmark (grant DFF – 7017-00003).

Acknowledgment: We gratefully acknowledge Dr. Professor Youming Zhang and Dr. Zhen Li from Shandong University for kindly providing the RecET direct cloning strains and vectors.

Reference Barka, E. A., Vatsa, P., Sanchez, L., Nathalie Gaveau-Vaillant, C. J., Klenk, H.-P., Clément, C., et al. (2016). Taxonomy, Physiology .... Am. Soc. Microbiol. 80, 1–43. doi:10.1128/MMBR.00019-15.Address. Boger, D. L., and Patel, M. (1988). Total Synthesis of Prodigiosin, Prodigiosene, and Desmethoxyprodigiosin: Diels-Alder Reactions of Heterocyclic Azadienes and Development of an Effective Palladium(II)-Promoted 2,2’-Bipyrrole Coupling Procedure. J. Org. Chem. 53, 1405–1415. doi:10.1021/jo00242a013. Bowman, J. P. (2007). Bioactive compound synthetic capacity and ecological significance of marine bacterial genus Pseudoalteromonas. Mar. Drugs 5, 220–241. doi:10.3390/md504220. C. Lee Ventola (2015). The Antibiotic Resistance Crisis Part 1: Causes and Threats. P&T 40, 277–283. doi:10.5796/electrochemistry.82.749. Culp, E., and Wright, G. D. (2017). Bacterial proteases, untapped antimicrobial drug targets. J. Antibiot. (Tokyo). 70, 366–377. doi:10.1038/ja.2016.138. Darshan, N., and Manonmani, H. K. (2015). Prodigiosin and its potential applications. J. Food Sci. Technol. 52, 5393–5407. doi:10.1007/s13197-015-1740-4. Dehio, C., and Meyer, M. (1997). Maintenance of broad-host-range incompatibility group P and group Q plasmids and transposition of Tn5 in Bartonella henselae following conjugal plasmid transfer from Eescherichia coli. J. Bacteriol. 179, 538–540. doi:10.1128/jb.179.2.538-540.1997. Gerber, N. N. (1975). Prodigiosin-like pigments. Crit. Rev. Microbiol. 3, 469–485. doi:10.3109/10408417509108758. Gram, L., Melchiorsen, J., and Bruhn, J. B. (2010). Antibacterial activity of marine culturable bacteria collected from a global sampling of ocean surface waters and surface swabs of marine organisms. Mar. Biotechnol. 12, 439–451. doi:10.1007/s10126-009-9233-y. Jackson, S. A., Crossman, L., Almeida, E. L., Margassery, L. M., Kennedy, J., and Dobson, A. D. W. (2018). Diverse and abundant secondary metabolism biosynthetic gene clusters in the genomes of marine sponge derived Streptomyces spp. Isolates. Mar. Drugs 16, 1–18. doi:10.3390/md16020067. Kalesperis, G. S., Prahlad, K. V., and Lynch, D. L. (1975). Toxigenic studies with the antibiotic pigments from Serratia marcescens. Can. J. Microbiol. 21, 213–220. doi:10.1139/m75- 030. Kawasaki, T., Sakurai, F., Nagatsuka, S. Y., and Hayakawa, Y. (2009). Prodigiosin biosynthesis gene cluster in the roseophilin producer Streptomyces griseoviridis. J. Antibiot. (Tokyo). 62, 271–276. doi:10.1038/ja.2009.27. Kawauchi, K., Tobiume, K., Iwashita, K., Inagaki, H., Morikawa, T., Shibukawa, Y., et al. (2008). Cycloprodigiosin hydrochloride activates the Ras-PI3K-Akt pathway and suppresses protein synthesis inhibition-induced apoptosis in PC12 cells. Biosci. Biotechnol. Biochem. 72, 1564–1570. doi:10.1271/bbb.80064.

Kawauchi, K., Tobiume, K., Kaneko, S., Kaneshiro, K., Okamoto, S., Ueda, E., et al. (2007). Suppression of AP-1 activity by cycloprodigiosin hydrochloride. Biol. Pharm. Bull. 30, 1792–1795. doi:10.1248/bpb.30.1792. Kim, D., Lee, J. S., Park, Y. K., Kim, J. F., Jeong, H., Oh, T. K., et al. (2007). Biosynthesis of antibiotic prodiginines in the marine bacterium Hahella chejuensis KCTC 2396. J. Appl. Microbiol. 102, 937–944. doi:10.1111/j.1365-2672.2006.03172.x. Lim, E. S. (2016). Inhibitory effect of bacteriocin-producing lactic acid bacteria against histamine-forming bacteria isolated from Myeolchi-jeot. Fish. Aquat. Sci. 19, 1–10. doi:10.1186/s41240-016-0040-x. Longeon, A., Peduzzi, J., Barthélemy, M., Corre, S., Nicolas, J. L., and Guyot, M. (2004). Purification and partial identification of novel antimicrobial protein from marine bacterium Pseudoalteromonas species strain X153. Mar. Biotechnol. 6, 633–641. doi:10.1007/s10126-004-3009-1. Milton, D. L., O’Toole, R., Hörstedt, P., and Wolf-Watz, H. (1996). Flagellin A is essential for the virulence of Vibrio anguillarum. J. Bacteriol. 178, 1310–1319. doi:10.1128/jb.178.5.1310-1319.1996. O’Toole, G. A., and Kolter, R. (1998). Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways: A genetic analysis. Mol. Microbiol. 28, 449–461. doi:10.1046/j.1365-2958.1998.00797.x. OKAMOTO, H., SATO, Z., SATO, M., KOISO, Y., IWASAKI, S., and ISAKA, M. (1998). Identification of Antibiotic Red Pigments of Serratia marcescens F-1-1, a Biocontrol Agent of Damping- off of Cucumber, and Antimicrobial Activity against Other Plant Pathogens. Japanese J. Phytopathol. 64, 294–298. doi:10.3186/jjphytopath.64.294. Östling, J., Goodman, A., and Kjelleberg, S. (1991). Behaviour of IncP‐1 plasmids and a miniMu transposon in a marine Vibrio sp.: isolation of starvation inducible lac operon fusions. FEMS Microbiol. Lett. 86, 83–94. doi:10.1111/j.1574-6968.1991.tb04797.x. Ravindran, A., Sunderrajan, S., and Pennathur, G. (2019). Phylogenetic Studies on the Prodigiosin Biosynthetic Operon. Curr. Microbiol. 76, 597–606. doi:10.1007/s00284-019- 01665-0. Richards, G. P., Watson, M. A., Needleman, D. S., Uknalis, J., Boyd, E. F., and Fay, J. P. (2017a). Mechanisms for Pseudoalteromonas piscicidainduced killing of vibrios and other bacterial pathogens. Appl. Environ. Microbiol. 83, 1–17. doi:10.1128/AEM.00175-17. Richards, G. P., Watson, M. A., Needleman, D. S., Uknalis, J., Boyd, E. F., and Fay, P. (2017b). Mechanisms for Pseudoalteromonas piscicida-Induced Killing of Vibrios and Other Bacterial Pathogens. 83, 1–17. Seemann, T. (2014). Prokka: Rapid prokaryotic genome annotation. Bioinformatics 30, 2068– 2069. doi:10.1093/bioinformatics/btu153. Setiyono, E., Adhiwibawa, M. A. S., Indrawati, R., Prihastyanti, M. N. U., Shioi, Y., and Brotosudarmo, T. H. P. (2020). An Indonesian Marine Bacterium, Pseudoalteromonas rubra, Produces Antimicrobial Prodiginine Pigments. ACS Omega 5, 4626–4635. doi:10.1021/acsomega.9b04322.

Skinnider, M. A., Johnston, C. W., Gunabalasingam, M., Merwin, N. J., Kieliszek, A. M., MacLellan, R. J., et al. (2020). Comprehensive prediction of secondary metabolite structure and biological activity from microbial genome sequences. Nat. Commun. 11, 1–9. doi:10.1038/s41467-020-19986-1. Vynne, N. G., Månsson, M., Nielsen, K. F., and Gram, L. (2011). Bioactivity, Chemical Profiling, and 16S rRNA-Based Phylogeny of Pseudoalteromonas Strains Collected on a Global Research Cruise. Mar. Biotechnol. 13, 1062–1073. doi:10.1007/s10126-011-9369-4. Wang, H., Li, Z., Jia, R., Hou, Y., Yin, J., Bian, X., et al. (2016). RecET direct cloning and Redαβ recombineering of biosynthetic gene clusters, large operons or single genes for heterologous expression. Nat. Protoc. 11, 1175–1190. doi:10.1038/nprot.2016.054. Wasserman, H. H., Friedland, D. J., and Morrison, D. A. (1968). A novel dipyrrolyldipyrromethene prodigiosin analog from Serratia marcescens. Tetrahedron Lett. 9, 641–644. doi:10.1016/S0040-4039(00)75602-4. Williams, R. P. (1973). Biosynthesis of prodigiosin, a secondary metabolite of Serratia marcescens. Appl. Microbiol. 25, 396–402. doi:10.1128/aem.25.3.396-402.1973. Williamson, N. R., Fineran, P. C., Leeper, F. J., and Salmond, G. P. C. (2006). The biosynthesis and regulation of bacterial prodiginines. Nat. Rev. Microbiol. 4, 887–899. doi:10.1038/nrmicro1531. Williamson, N. R., Simonsen, H. T., Ahmed, R. A. A., Goldet, G., Slater, H., Woodley, L., et al. (2005). Biosynthesis of the red antibiotic, prodigiosin, in Serratia: Identification of a novel 2-methyl-3-n-amyl-pyrroie (MAP) assembly pathway, definition of the terminal condensing enzyme, and implications for undecylprodigiosin biosynthesis in Streptomyces. Mol. Microbiol. 56, 971–989. doi:10.1111/j.1365-2958.2005.04602.x. Zhang, H., Peng, Y., Zhang, S., Cai, G., Li, Y., Yang, X., et al. (2016). Algicidal effects of prodigiosin on the harmful algae Phaeocystis globosa. Front. Microbiol. 7, 1–10. doi:10.3389/fmicb.2016.00602.

FIGURE AND TABLE LEGENDS

Figure 1. The genetic organization of prodigiosin biosynthetic gene clusters of Serratia sp. ATCC 39006 and Pseudoalteromonas rubra S4059. Block arrows are color coded to show which genes encode proteins involved in the biosynthesis of prodigiosin (A). The pigC gene is prodigiosin synthesizing transferase. A prodigiosin deficient mutant was generated by knocking out pigC gene (B) and the phenotype of ΔpigC is brownish (C). The chemical profiles (D) of P. rubra S4059 and prodigiosin deficient mutant ΔpigC. The red peaks are prodigiosin and its analogs.

Figure 2. Growth kinetics of Pseudoalteromonas rubra S4059 wild type and prodigiosin deficient mutant ΔpigC when grown in marine minimal medium containing mannose for 4 days. Grey lane: wild type S4059; black lane: ΔpigC. Each experiment is repeated in bio- triplicates and error bars are standard deviation.

Figure 3. Screening Pseudoalteromonas rubra S4059 wild type (1, 2) and prodigiosin deficient mutant ΔpigC (3, 4) for antibacterial activity against Staphylococcus aureus 8325 (A, C, E) and Vibrio anguillarum 90-11-287 (B, D, F). Wild type S4059 and the ΔpigC were cultured in marine minimal medium for 24h, 48h, or 72h. A-B, 24h; C-D, 48h; E-F,72h; 5: medium extractions, negative control. 20 mL cultures were taken and extracted from each time point for bioassay.

Figure 4. A total of 29 fractions were obtained from the ethyl acetate extract of 500mL of the prodigiosin deficient mutant ΔpigC and used for disc diffusion assays. Only two fractions, Fraction 11 (F11), Fraction 12 (F12), which are two blue fractions, showed antibacterial activity against both Staphylococcus aureus 8325 (A) and Vibrio anguillarum 90-11-287 (B). C: extracts from marine minimal medium containing mannose as a negative control.

extraction using ethyl acetate with 1% formic acid. (B) Pseudoalteromonas rubra wild type S4059 and prodigiosin deficient mutant ΔpigC were cultured in marine minimal medium containing mannose. 1mL culture supernatants were filtered and then treated without or with heating at 95 °C for 10mins. 40 µL supernatants were tested against Staphylococcus aureus 8325. 1,2: filtered wild type supernatants; 3, 4: filtered wild type supernatants after heating; 5, 6: filtered ΔpigC supernatants; 7, 8: filtered ΔpigC supernatants after heating.

4 Log 10(CFU/mL) Log

0 1 2 3 4 5 Time (days)

Figure 5. (A) Pseudoalteromonas rubra wild type S4059 was cultured in marine minimal medium containing mannose. 5mL of the cultures were extracted with the equal volume of ethyl acetate with or without 1% formic acid and 20 µL of each extract was tested antibacterial activity against Staphylococcus aureus 8325. 20 µL liquid from water phase after the extraction also were tested against the two bacteria. 1,2: wild type culture extracts using ethyl acetate with 1% formic acid; 3, 4: wild type culture extracts using ethyl acetate; 5,6: liquid from water phase after extraction using ethyl acetate; 7, 8; liquid from water phase after extraction using ethyl acetate with 1% formic acid. (B) Pseudoalteromonas rubra wild type S4059 and prodigiosin deficient mutant ΔpigC were cultured in marine minimal medium containing mannose. 1mL culture supernatants were filtered and then treated without or with heating at 95 °C for 10mins. 40 µL supernatants were tested against Staphylococcus aureus 8325. 1,2: filtered wild type supernatants; 3, 4: filtered wild type supernatants after heating; 5, 6: filtered ΔpigC supernatants; 7, 8: filtered ΔpigC supernatants after heating.

SUPPLEMENTARY MATERIALS TABLE AND FIGURE LEGENDS

Figure S1. Biofilm formation of Pseudoalteromonas rubra S4059 wild type and prodigiosin deficient mutant ΔpigC in marine minimal medium (MMM) or marine both (MB) for 24 h (A and B), 48 h (C and D), and 72 h (E and F) as determined by the O’Toole & Kolter crystal violet assay. Each experiment is repeated in bio-triplicates and error bars are standard error. Wt: wild type S4059; pigC: prodigiosin deficient mutant ΔpigC.

Figure S2. (A) UV-Vis absorption spectrum for MBC (m/z 191.0822) was measured during the chromatographic analysis. (B) Base peak chromatogram (BPC) of fraction 12, indicating proposed identity of tentatively identified metabolites. (C) the chemical structure of Prodigiosin, (4-methoxy-2,2’- bipyyrrole-5-carbaldehyde (MBC) and Dipyrrolyldipyrromethene prodigiosin (Dip-PDG).

Figure S3. Pseudoalteromonas rubra S4059 was cultured in marine minimal medium containing mannose and the cultures were extracted using ethyl acetate with (A) or without 1% formic acid (B). The filtered supernatants (FSNs) of wild type P. rubra S4059 (1, 2) and ΔpigC mutant (3, 4) were treated with proteinase K (Pk) or pepsin (Pn) and still retain anti- Staphylococcus aureus 8325 activities (C).

Table S1. Comparison of proteins from the prodigiosin gene clusters in Serratia sp. ATCC 39006 and Pseudoalteromonas rubra S4059. Putative functions were assigned based on the results of BLASTp searches.

Table S2. The bacterial strains, primers and plasmids used in this study.

Table S3. Primers used in this study.

Figure S1. Biofilm formation of Pseudoalteromonas rubra S4059 wild type and prodigiosin deficient mutant ΔpigC in marine minimal medium (MMM) or marine both (MB) for 24 h (A and B), 48 h (C and D), and 72 h (E and F) as determined by the O’Toole & Kolter crystal violet assay. Each experiment is repeated in bio-triplicates and error bars are standard deviation. Wt: wild type S4059; pigC: prodigiosin deficient mutant ΔpigC.

Figure S3. Pseudoalteromonas rubra S4059 was cultured in marine minimal medium containing mannose and the cultures were extracted using ethyl acetate with (A) or without 1% formic acid (B). The filtered supernatants (FSNs) of wild type P. rubra S4059 (1, 2) and pigC mutant (3, 4) were treated with proteinase K (Pk) or pepsin (Pn) and still remain anti- Staphylococcus aureus 8325 activities (C).

Serratia sp. Pseudoalteromonas Biosynthetic ATCC 39006 rubra S4059 Homologs Predicted protein function pathway Protein (identity %) PigA 69.19 MBC L-prolyl-PCP dehydrogenase PigB 62.29 MAP Oxidoreductase Terminal PigC 73.3 Terminal Condensation enzyme condensation Thiamine diphosphate dependent-3- PigD 74.46 MAP acetyloctanal synthase PigE 79.46 MAP/Enamine Aminotransferase S-adenosyl-L-methionine-dependent PigF 78.99 MBC methyltransferase PigG 74.36 MBC putative acyl carrier protein PigH 71.82 MBC HBM synthase/aminotransferase PigI 60.25 MBC L-prolyl-AMP ligase PigJ 60.76 MBC β-ketoacylsynthase PigK 66.67 MBC Hypothetical protein (RedY) PigL 36.9 MBC Phosphopantetheinyl transferase PigM 53.54 MBC Putative NAD(P)H nitroreductase

Table S2. The bacterial strains and plasmids used in this study

Reference Strains/plasmids/primers Genotype or relevant characteristics or source Bacterial strains Escherichia coli WM3064 thrB1004 pro thi rpsL hsdS lacZΔM15 RP4- (Dehio and 1360 Δ(araBAD)567 ΔdapA1341::[erm pir], Meyer, 37 ℃. Donner strains in conjuation. 1997) GB dir pir116 An arabinose-inducible ETγA operon (full- length recE, recT, redγ, and recA), a copy-up pir116 gene. Host strain for constructing (Wang et suicide plasmids with R6K replication origin. al., 2016) Pseudoalteromonas rubra S4059 Wild type strain, Isolated from seaweed. (Gram et al., 2010) ΔpigC PigC gene in-frame deletion mutant of P. rubra S4059 Δ PigC This study Plasmids pDM4 Suicide vector for targeted mutagenesis; (Milton et sacB; oriR6K; CmR al., 1996)

pDM4 –del-pigC The left arm and right arm DNA region of the pigC gene were PCR amplified from P. rubra S4059 genome and cloned in the This study pDM4 plasmid by direct cloning

Table S3. Primers used in this study.

Primer Sequence Description Expected size (bp) pigC-L-F ATCTGTTCGAGCCGGAAA Left arm of pigC 1000 pigC -L-R GGATTAGATATACCGTCTCCTTATTAC GCG pigC -R-F AGGAGACGGTATATCTAATCCATCGC Right arm of pigC 1017 GGGC pigC -R-R TTTGGACCACGTAGCACC CmR F GGCATTTCAGTCAGTTGCTC Detection of CmR gene 525 CmR R CCATCACAAACGGCATGATG pigC-pDM4-F GGTGCTACGTGGTCCAAAGGTCGAC Linear pDM4 plasmid 7106 GGTATCGATAAG (pigC) pigC-pDM4-R TTTCCGGCTCGAACAGATTAGATCTT GCATGCGGGT pigC -p 1 GGTTAACAGGCCAGCTACTG Confirmation of pigC 1st: P1, P2 mutant 3879a; 1230b pigC -p 2 GCCCGCGATGGATTAGA P3, P4 3878a; 1229b pigC -p 3 CGCGTAATAAGGAGACGGTATA 2nd: P1, P2 3879a; 1230b pigC -p 4 GGTGCCCTGAAAGCTATACC a the size of WT; b the size of mutant.

Paper 3

Attachment to and growth on chitin surfaces results in loss of prodigiosin production in a pigmented Pseudoalteromonas rubra S4059

Xiyan Wang 1, Thomas Isbrandt 1, Thomas Ostenfeld Larsen 1, Shengda Zhang 1 and Lone Gram 1 *

1 Department of Bioengineering, Technical University of Denmark, DK-2800 Kgs. Lyngby,

Denmark; [email protected] (X.W.); [email protected] (T.I.); [email protected] (T.O.L); [email protected] (S.Z.) * Correspondence: [email protected] (L.G.); Tel: +45-4525-2586

Abstract Bacteria attach to surfaces and form biofilms that can protect them against several adverse conditions. Spontaneous mutations occur in cells in bacterial biofilms and this can provide genetic variations driving the bacterial evolution. In this study, we observed that attachment to and growth on natural chitin surfaces or chitin flakes resulted in pigment deficiency in an otherwise red-pigmented Pseudoalteromonas rubra. The frequency of non-pigmented cells (colonies) was 24 ± 2.1%. The purpose of this study is to explore the relationship between this assumed spontaneous mutation and chitin surface associated biofilm formation in pigmented Pseudolateromonas rubra S4059. The appearance of non-pigmented colonies (cells with spontaneous mutations) only occurred on natural chitin surfaces or chitin flakes and not on seagrass or stainless-steel surfaces. Resequencing of nine randomly picked non-pigment colonies of S4059 indicated that mutations had not occurred in or around the prodigiosin biosynthetic gene cluster (BGC). Six of nine mutants had a short insertion in a GGDEF domain contained diguanylate cyclase (DGC) and this is the most likely candidate that led to phenotypic changes in S4059. The spontaneous mutation of S4059 leads to consistent prodigiosin deficiency and a lower maximum cell density in stationary phase as compared to the wild type S4059. However, cell numbers of the wild type declined after entering stationary phase whereas the mutant retained high cell numbers. We tested another pigmented Pseudoalteromonas piscicida S2724, and non-pigmented mutants occurred when grown on natural chitin surfaces. In conclusion, chitin surface associated biofilm formation in pigmented Pseudoalteromonas species results in spontaneous mutation. The mutation can provide genetic variation in these species and provide a growth (survival) advantage for long- time incubation. Keywords: Pigment, Pseudoalteromonas, GGDEF domain, Chitin surface, Spontaneous mutation.

Introduction:

Bacteria use many mechanisms to adapt to new environment and genetic changes that confer a fitness advantage are likely to drive evolution in bacterial population. Genetic changes can be caused by several genetic mechanisms, such as DNA duplication, insertion or deletion, that subsequently results in phenotypic changes [1,2]. Genetic variation is a prerequisite for bacterial evolution and spontaneous mutations provide a major force in generating genetic variations. Spontaneous mutations are often present within an individual cell, and will either increase in frequency in the bacterial population, or vanish during natural selection [3]. The mutation that provides beneficial effects to bacteria to adapt to new environment will be fixed in the population.

Bacteria are often associated with surfaces and form communities known as biofilms, that form either on biotic or abiotic surfaces [4]. Biofilms are the most widely distributed matrix that can protect bacteria against a wide range of environmental challenges [5]. Interestingly, bacterial biofilms often exhibit phenotypic variation [6–8], suggesting that mutation and genetic change are important traits during biofilm formation. For example, an insertion sequence (IS) element was inserted in the capsular polysaccharide operon during biofilm formation of Pseudoalteromonas lipolytica and this resulted in an increasing attachment ability of this strain on abiotic and biotic surface [9].

In marine environments, the genus Pseudoalteromonas is widely associated with biotic or abiotic surface and many species of the genus are excellent biofilm formers [10]. This genus can be divided into two groups: pigmented and non-pigmented [10] of which the pigmented typically are capable of producing many secondary metabolites with bioactivity. Also, these bacteria are capable of degrading different polymers and chitin degradation is a key trait in the pigmented Pseudoalteromonas species [11] that carry a multitude of genes encoding enzymes involved in chitin degradation [11]. Therefore, natural chitin surfaces are likely to be one of ecological niches for these pigmented species in marine environment. We have found that the red-pigmented bacterium Pseudoalteromonas rubra S4059 can produce a chitinolytic enzyme cocktail and a red pigment, prodigiosin. In this study, we observed that growth of P. rubra S4059 on either shrimp-shells (natural chitin) or chitin flakes as the sole carbon-source resulted in appearance of spontaneous non-pigmented colonies. Such non-pigmented colonies were never observed when S4059 was grown in colloidal or crystalline chitin or on

abiotic surfaces but only on large chitin surfaces. It is the first report that chitin surface is likely to be an important ecological niche for providing genetic variations to pigmented Pseudoalteromonas. The purpose of this study is to investigate the relationship between the loss of pigmentation of P. rubra S4059 and chitin surfaces and determine the genetic background. Further, we also explored other phenotypic variations between P. rubra S4059 wild type and the non-pigmented (spontaneous mutants) strains.

Results:

Non-pigmented P. rubra S4059 accumulate when grown on chitin surface. P. rubra S4059 was incubated in 3% sea salt water with natural shrimp shells to determine attachment and biofilm formation. Samples were taken regularly for eight days at 25 ◦C. Surprisingly, colonies devoid of red pigmentation appeared on the MB-plates (Figure 1). A total of five independent experiments were conducted growing P. rubra 4059 on natural shrimp shells and the appearance of non-pigmented colonies was consistently observed after eight days incubation.

Prodigiosin deficient mutants were associated with chitin surface. To investigate whether the appearance of non-pigmented colonies (mutants) was associated with the substrate (chitin) or the surface-association (biofilm) or both, natural shrimp shells, chitin flakes, chitin power (crystalline and colloidal chitin), metal coupons or seagrass (Zostera marina) were used for culturing S4059. Cells were plated and counted after eight days incubation. Non- pigmented colonies (mutants) only appeared after incubation with shrimp shells or chitin flakes and were not observed when S4059 was grown on other surfaces or in chitin power. The frequency of mutation was 24 ± 2.1% in the entire bacterial population.

Non-pigmented S4059 have altered chemical profiles and antibacterial activities as compared to wild type P. rubra S4059. P. rubra S4059 wild type and three randomly picked mutants, S1, F1 and Fi1, were grown in MMM containing mannose for two days. The cultures were extracted and analyzed by high-performance liquid chromatography coupled to high- resolution mass spectrometry (HPLC-HRMS). Neither prodigiosin, nor the precursor of prodigiosin, 4-methoxy-2, 2’- bipyrrole-5-carbaldehyde (MBC), could be detected in the non-

pigmented mutants (Figure 2). Further, antibacterial activity of the culture extractions of the three mutants was tested against Staphylococus aureus 8325 and Vibrio anguillarum 90-11- 287. The extracts did not have antibacterial activity against the two bacteria (Figure 2).

Non-pigmented mutants of S4059 exhibits a growth advantage after stationary phase in marine minimal medium containing mannose. To investigate whether the mutation influenced growth of S4059 mutants, growth kinetics of wild type S4059 and the three mutants, S1, F1 and Fi1, were established by measuring optical density. The mutants grew slower than wild type S4059. However, after reaching the stationary phase, the density of wild type S4059 decreased significantly, while this was not seen for the mutants.

Most spontaneous mutants have a short insertion in a diguanylate cyclase. Nine randomly picked prodigiosin deficient mutants (S1, S2, S3, F1, F2, F3, Fi1, Fi2, Fi3) were re-sequenced. Three different variant types were detected in the nine strains. These were single nucleotide polymorphism (SNP), insertion or deletion of ≤ 50 bp sequences (Indel) and copy-number variation (CNV). 113 SNPs, 2 Indels and 2 CNVs were detected from S1; 116 SNPs, 4 Indels and no CNVs were detected from S2; 95 SNPs, 4 Indels and 164 CNVs were detected from S3; 87 SNPs, 5 Indels and 144 CNVs were detected from F1; 98 SNPs, 5 Indels and 118 CNVs were detected from F2; 94 SNPs, 4 Indels and 137 CNVs were detected from F3; 95 SNPs, 3 Indels and 109 CNVs were detected from Fi1; 97 SNPs, 4 Indels and 150 CNVs were detected from Fi2; 95 SNPs, 1 Indel and 140 CNVs were detected from Fi3 (Table 1).

No mutation was found in or around the prodigiosin biosynthesis gene cluster (BGC). Of the nine colonies, six have a short insertion in the same GGDEF domain of a diguanylate cyclase (DGC) (Table 2). The mutated diguanylate cyclase is located in the operon containing genes encoding putative riboflavin synthesis associated enzymes.

Loss of pigment in a yellow pigmented Pseudoalteromonas piscicida S2724. Chitin degradation machinery is a common trait in pigment Pseudoalteromonas [11], that led us to ask if the loss of pigment production is observed in other pigmented Pseudoalteromonas when grown on chitin surface? A yellow pigmented bacterium P. piscicida S2724 was selected and grown on shrimp shells for eight days at 25 ◦C. Indeed, pigment deficient mutants of S2724 were observed and 16sRNA sequencing indicated that the pigment deficient mutants were indeed P. piscicida.

Discussion:

Chitin degradation capacity is a key trait for pigmented Pseudoalteromonas [12] and natural chitin surfaces are likely to be ecological niches for these species. Studies of spontaneous mutations associated with this ecological niche are crucial to understanding bacterial evolution in natural environment. In this study, spontaneous mutation led to pigment deficient strains of Pseudoalteromonas rubra when associated with chitin surface. Genome resequencing of nine randomly picked mutants indicated that no mutation took place in prodigiosin BGC of S4059 and the mutation was detected in the GGDEF domain of a diguanylate cyclase (DGC) in six of the nine re-sequenced mutants. The mutations in Pseudoalteromonas rubra S4059 led to consistent pigment deficiency and a slightly lower cell density before stationary phase as compared to the wild type, however, the mutation likely provided a growth advantage to Pseudoalteromonas rubra S4059 as cell density did not decline in the mutants as it did in wild type thus providing a beneficial trait for long-time incubation under the tested conditions. However, this study did not reveal the mechanism behind the mutations and altered phenotype and more experimental work is needed as described below.

Of the nine re-sequenced colonies, a mutation in the GGDEF domain containing DGC was detected in six strains. The GGDEF domain is sufficient to encode DGC activity, which is involved in cyclic-di-GMP synthesis [13]. Cyclic-di-GMP is a bacterial second messenger that can control multiple cellular processes in bacteria, such as biofilm formation, virulence and motility [14]. Thus, experiments in which cyclic-di-GMP is manipulated should be performed and the possible effect on biofilm and/or motility assessed. Similarly, the cyclic-di-GMP production by the mutants should be compared to that of the wild type. Recently, Yin et al. have reported a novel regulatory mechanism of prodigiosin biosynthesis in Pseudoalteromona sp. R3, which a stringent starvation protein A (SspA) can positively regulate prodigiosin production by influencing the siderophore-dependent iron uptake pathway [15], however, no mutation was detected in SspA or siderophore associated genes in the spontaneous mutants of S4059. Considering that GGDEF domain contained protein could involve c-di-GMP synthesis and that may involve siderophore production as reported in other bacteria [16]. Thus, we can measure the level of siderophore production in both wild

type S4059 and the mutants to explore whether loss of prodigiosin in the mutants is associated with siderophore-dependent iron uptake.

However, many proteins that contain the GGDEF domain have no c-di-GMP synthesis activity [17,18] and the enzymatic activity of the GGDEF is highly regulated by surrounding sensory protein domains [13], suggesting that the surrounding sensory protein domains will significantly influence the activity of the GGDEF domain. To analyse which sensory domains are adjacent the mutated GGDEF domain, the domain structure of the mutated diguanylate cyclase was predicted by Pfam. The mutated DGC contains one two-component regulator propeller at N-terminal, one YYY motif and one GGDEF domain at C-terminal. Therefore, the DGC can be a hybrid two-component regulator according to the architecture of domain structure as reported by Menke et al. [19]. In this study, the prodigiosin production was changed as were the growth and antibacterial activities in S4059. In a targeted mutant in which we deleted the pigC of the prodigiosin BGC, growth was not affected, suggesting that the changes in the chitin-induced mutants were not caused by the lack of prodigiosin but a function of the mutation and parallel effect. This suggested that the mutated gene may play a regulatory role in S4059. Therefore, the mutated DGC has the potential to be a global regulator in S4059. The DGC is located in the operon with a putative riboflavin biosynthetic gene cluster. Riboflavin is known as vitamin B2. Vitamin B2 is the precursor of two coenzymes known as flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD). The mutation of diguanylate cyclase could potentially influence riboflavin production in S4059. Simultaneously, FMN and FAD are involved in the biosynthesis pathway of prodigiosin, such as pigM is a probably FMN dependent oxidoreductase and pigB is a probably FAD dependent dehydrogenase [20,21]. Therefore, the lack of FMN and FAD may shut down the biosynthesis of MBC and MAP that results in prodigiosin deficiency of S4059. To verify these hypotheses, a DGC in-frame deletion mutant needs to be further generated.

Mutation of Pseudomonas protegens in the global GacS- GacA regulatory system has been widely reported when grown in nutrient rich medium [22,23]. Yan et al. [23] have indicated that accumulation of spontaneous mutants is associated with the expression of pyoluteorin biosynthetic genes by overexpressing a transcriptional regulator of pyoluteorin. When P. protegens was co-cultured with a competitor, the numbers of spontaneous mutants were significantly decreased as compared to in pure culture. The authors suggested that the

expression of unnecessary secondary metabolites impose a metabolic burden and the mutation in the GacS-GacA regulatory system can shut down the cost and thus provide a fitness advantage as compared to wild type. In our study, growth on chitin surface led to spontaneous mutation of the tested two pigmented Pseudolateromonas species. This phenomenon may due to the expression of unnecessary pigment associated biosynthetic genes. To investigate this hypothesis, further co-cultivation-based approach needs to be set up with a competitor together with S4059 on chitin surface to investigate the mutation frequency as compared to pure culture.

Pigment deficiency was observed in the two tested pigmented Pseudoalteromoans species. To further address the question “are mutations leading to pigment deficiency commonly observed in pigmented Pseudoalteromonas when grown on chitin surface?” We need to test a broader range of pigmented Pseudoalteromonas species. Further, pigment deficiency in pigmented Pseudoalteromonas species is easily visible, however, pigment deficiency is not the only phenotypic change. Furthermore, we also did bacterial antagonism assay by spotting S4059 wild type and the mutants on agar plates against Staphylococcus aureus 8325 or Vibrio anguillarum 90-11-287. The antibacterial activities of the mutants were decreased as compared to the wild type, however, we also observed that the wild type colonies autolyzed which were not seen in the mutants (data not shown). These phenotypic differences can be due to the difference in cell density between wild type and the mutants and thus this experiment needs to be repeated and optimized. Altogether the influence of the mutation on growth, autolysis, and bioactive activities in S4059 suggested that the mutation may affect the expression of multiple genes, and transcriptome and proteome will tell us how the mutation affects genes expression in S4059.

In summary, growth on chitin surface results in loss of prodigiosin production of P. rubra S4059 and resequencing of non-pigment strains indicated that the mutation is likely in a GGDEF domain contained DGC. Prodigiosin production correlates negatively with cellular levels ATP in Serratia marcescens [24]. Therefore, loss of prodigiosin production may save energy costs in S4059. In addition, Price et al. have reported that genes with no detectable benefits are responsible for 31% of proteins productions in E. coli K-12 by using a combination of a library of transposon mutants and individual gene information from knockout strains and that will lead to remarkable cost during evolution [25]. Further, previous proteomic analysis

indicated that 66% of detected proteins were upregulated when S4059 grown on chitin as compared to growth on mannose (In paper 1), suggesting that growth on chitin may result in more energy costs that contribute to proteins expression in S4059. In summary, loss of prodigiosin production may save energy cost in S4059 cells when growth on chitin surface and such mutations may be decreased when the prodigiosin is necessary for bacterial cells, such as under competitors challenging. Spontaneous mutations of P. rubra S4059 provide genetic diversity driving evolution of bacteria and a potential advantage to adapt to various environmental conditions in natural environment.

Experimental procedures

Bacterial strains and cultural conditions. Two species of Pseudoalteromonas, P. piscicida S2724 producing a yellow pigment and P. rubra S4059 producing a red pigment, were used to determine phenotypic (pigment) changes when grown on natural shrimp shells. The two Pseudoalteromonas species were isolated during the Galathea 3 expedition [26] and cultured on marine agar (MA, BD Difco 2216) or in marine both (MB, BD Difco 2216) at 25 °C unless specifically indicated. Liquid cultures were grown with shaking at 200 rpm or under stagnant condition in 96 wells microplates. The two Pseudoalteromonas species were cultured in marine minimal medium (MMM) [27] supplemented with mannose at 25 °C, 200 rpm for chemical extraction and growth curves. Antibacterial activity against Gram-negative bacteria Vibrio anguillarum 90-11-287 and Gram-positive bacteria Staphylococcus aureus 8325 was tested according to [26].

Growth on nutrient and non-nutrient surfaces. To investigate the relationship between loss of pigmentation and surface-growth, natural shrimp shells, chitin flakes, metal coupons or seagrass (Zostera marina) were used for the experiment setup. Chitin power (crystalline or colloidal chitin) was used as carbon-source control. The protocol of shrimp shell preparation was from Wang et al 2021 (revision). Pre-cultures of S4059 were grown in 5 mL MB in 50 mL flasks at 25°C, 200 rpm overnight and diluted to 104 CFU/mL in sea water (3% sea salt, sigma, S9883) with shrimp shells, chitin flakes, coupons, chitin power or seagrass in 1.5 mL Eppendorf tubes. Metal coupons do not provide nutrients and bacteria were therefore grown in marine minimal medium with or without mannose. The tubes were sonicated five minutes at 50/60 Hz in an ultrasonic bath (Emmi D20 Q, EMAG, Germany) and vortexed 10s to remove bacteria from the surface after 8 days incubation[28], and samples of the liquid were plated on MA plates after serial dilution for colonies counting. The numbers of colonies with red pigment (or yellow pigment) or without the pigment were counted, respectively. All experiments were done in biological duplicate.

DNA manipulation and resequencing analysis. Genomic DNA of nine randomly picked non- pigmented colonies from S4059 were extracted using Genomic DNA buffer set (QIAGEN, 19060) following the supplier instructions. A Qubit (v2.0) dsDNA BR assay kit (Lot: 2159622, Invitrogen) were used for gDNA.quantification. The colonies were resequenced at the Novogene (United Kingdom) using 150-bp paired- end sequencing on an IIIumina® platform.

The chitin shell attachment experiment was carried out at total of 5 times and the nine random picked colonies were S1, S2 and S3 from 2nd experiment, F1, F2, and F3 from 4th experiment and Fi1, Fi2 and Fi3 from 5th experiment, respectively. Variations calling from the nine colonies were performed by the Novogene as following:

Original data of sequencing data (e.g. Illumina HiSeqTM) were first transformed to sequence reads by base calling with the CASAVA software. The sequences and corresponding sequencing quality information are stored in a FASTQ file [29]. Raw data in FASTQ file were filtered using CASAVA software to get clean data. Clean data were aligned with the reference sequence through BWA [30]. The duplicates were removed by using SAMTOOLS [31]. Single nucleotide polymorphisms (SNPs) were detected using SAMtools [31] with the following parameter: 'mpileup -m 2 -F 0.002 -d 1000'. To reduce the error rate in SNP detection, the results were filtered with the criterion as follows: (1) The number of support reads for each SNP should be more than 4; (2) The mapping quality (MQ) of each SNP should be higher than 20. InDels (insertions and deletions) were detected using SAMTOOLs [31] with the following parameter 'mpileup -m 2 -F 0.002 -d 1000' to detect InDels and followed by annotation using ANNOVAR [30]. The filter conditions are same with SNPs. Structural variants (SVs) were detected using BreakDancer [30]. The detected SVs were filtered by removing those with less than two supporting PE reads. The INS, DEL and INV were further annotated by ANNOVAR [32]. CNVnator [33] were used to detect copy-number variations (CNVs) of potential deletions and duplications with the following parameter '-call 100'. The detected CNVs were further annotated by ANNOVAR.

Growth kinetics of P. rubra S4059 wild type and three randomly picked non-pigmented mutants. The growth kinetics of three mutants, S1, F1 and Fi1, and the wild type S4059 were compared. Pre-cultures of S4059 wild type and the three mutants were incubated in MB overnight at 25°C, 200 rpm. The cultures were diluted to OD600≈ 0.01 in the final volume of 100µL MMM in 96 wells microplates. The cell density in 96 wells microplates was estimated at 600 nm by Multi-mode readers for every 10 minutes at 25°C (BioTek Synergy, S1LFA). The 96 wells microplates were shacked for 10 seconds for each measurement. All experiments were done in biological triplicate.

Extraction of metabolites for chemical analysis The protocol was modified from Wang et al. 2020 [26]. Briefly, P. rubra S4059 and the three mutants (S1, F1 and Fi1) were cultured in 100 mL MMM at 25 °C, 200 rpm for two days. 20 mL of the culture were extracted with an equal volume of high-performance liquid chromatography (HPLC) grade Ethyl acetate (EtoAc) in 50mL Falcon tubes. The organic phase was transferred to a new Falcon tube and evaporated under nitrogen. The dried extract was re-dissolved in 200 µL methanol and extractions were used for antibacterial activity assay and chemical analysis by UHPLC-HRMS.

Antibacterial activity assay. The protocol was modified from Gram et al. [26]. Briefly, the extractions of metabolites were tested for antibacterial activity against Staphylcococcus aureus 8325. The composition of agar medium was used according to Gram et al. [26], and 6 mm wells were punched in the plates with a least 2-centimeter distance are between the wells. 40 µL of each extraction of metabolites was dropped into the well of the plate for bioassay. The plates were incubated at 25 °C for 24-48 h. All experiments were done in biological duplicate.

References:

1. Dybvig, K. DNA rearrangements and phenotypic switching in prokaryotes. Mol. Microbiol. 1993, 10, 465–471, doi:10.1111/j.1365-2958.1993.tb00919.x.

2. Sousa, A.M.; Machado, I.; Pereira, M.O. Phenotypic switching: an opportunity to bacteria thrive. Sci. against Microb. Pathog. Commun. Curr. Res. Technol. Adv. 2011, 252–262.

3. Hershberg, R. Mutation—the engine of evolution: Studying mutation and its role in the evolution of bacteria. Cold Spring Harb. Perspect. Biol. 2015, 7, 1–13, doi:10.1101/cshperspect.a018077.

4. Hall-Stoodley, L.; Costerton, J.W.; Stoodley, P. Bacterial biofilms: From the natural environment to infectious diseases. Nat. Rev. Microbiol. 2004, 2, 95–108, doi:10.1038/nrmicro821.

5. Flemming, H.C.; Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 2010, 8, 623–633, doi:10.1038/nrmicro2415.

6. Häußler, S.; Ziegler, I.; Löttel, A.; Götz, F. V.; Rohde, M.; Wehmhöhner, D.; Saravanamuthu, S.; Tümmler, B.; Steinmetz, I. Highly adherent small-colony variants of Pseudomonas aeruginosa in cystic fibrosis lung infection. J. Med. Microbiol. 2003, 52, 295–301, doi:10.1099/jmm.0.05069-0.

7. D’Argenio, D.A.; Calfee, M.W.; Rainey, P.B.; Pesci, E.C. Autolysis and autoaggregation in Pseudomonas aeruginosa colony morphology mutants. J. Bacteriol. 2002, 184, 6481–6489, doi:10.1128/JB.184.23.6481-6489.2002.

8. Drenkard, E.; Ausubel, F.M. Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation. Nature 2002, 416, 740–743, doi:10.1038/416740a.

9. Zeng, Z.; Zhan, W.; Wang, W.; Wang, P.; Tang, K.; Wang, X. Biofilm formation in Pseudoalteromonas lipolytica is related to IS5-like insertions in the capsular polysaccharide operon. FEMS Microbiol. Ecol. 2019, 95, 1–11, doi:10.1093/femsec/fiz065.

10. Bowman, J.P. Bioactive compound synthetic capacity and ecological significance of marine bacterial genus Pseudoalteromonas. Mar. Drugs 2007, 5, 220–241, doi:10.3390/md504220.

11. Paulsen, S.S.; Strube, M.L.; Bech, P.K.; Gram, L.; Sonnenschein, E.C. Marine Chitinolytic Pseudoalteromonas Represents an Untapped Reservoir of Bioactive Potential . mSystems 2019, 4, 1–12, doi:10.1128/msystems.00060-19.

12. Giubergia, S.; Phippen, C.; Nielsen, K.F.; Gram, L. Growth on Chitin Impacts the Transcriptome

and Metabolite Profiles of Antibiotic-Producing Vibrio coralliilyticus S2052 and Photobacterium galatheae S2753. mSystems 2017, 2, 1–12, doi:10.1128/msystems.00141-16.

13. Ryjenkov, D.A.; Tarutina, M.; Moskvin, O. V.; Gomelsky, M. Cyclic diguanylate is a ubiquitous signaling molecule in bacteria: Insights into biochemistry of the GGDEF protein domain. J. Bacteriol. 2005, 187, 1792–1798, doi:10.1128/JB.187.5.1792-1798.2005.

14. Jenal, U.; Reinders, A.; Lori, C. Cyclic di-GMP: Second messenger extraordinaire. Nat. Rev. Microbiol. 2017, 15, 271–284, doi:10.1038/nrmicro.2016.190.

15. Yin, J.; Ding, M.; Zha, F.; Zhang, J.; Meng, Q.; Yu, Z.; Province, Z.; Yu, Z. Stringent starvation protein regulates prodigioine biosynthesis via affecting siderophore production in Pseudoalteromonas sp. R3. 2021, doi:10.1128/AEM.02949-20.

16. Little, R.H.; Woodcock, S.D.; Campilongo, R.; Fung, R.K.Y.; Heal, R.; Humphries, L.; Pacheco- Moreno, A.; Paulusch, S.; Stigliano, E.; Vikeli, E.; et al. Differential regulation of genes for cyclic- di-GMP metabolism orchestrates adaptive changes during rhizosphere colonization by Pseudomonas fluorescens. Front. Microbiol. 2019, 10, 1–14, doi:10.3389/fmicb.2019.01089.

17. Matilla, M.A.; Travieso, M.L.; Ramos, J.L.; Ramos-González, M.I. Cyclic diguanylate turnover mediated by the sole GGDEF/EAL response regulator in Pseudomonas putida: Its role in the rhizosphere and an analysis of its target processes. Environ. Microbiol. 2011, 13, 1745–1766, doi:10.1111/j.1462-2920.2011.02499.x.

18. Christen, M.; Christen, B.; Folcher, M.; Schauerte, A.; Jenal, U. Identification and characterization of a cyclic di-GMP-specific phosphodiesterase and its allosteric control by GTP. J. Biol. Chem. 2005, 280, 30829–30837, doi:10.1074/jbc.M504429200.

19. Menke, M.; Berger, B.; Cowen, L. Markov random fields reveal an N-terminal double beta- propeller motif as part of a bacterial hybrid two-component sensor system. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 4069–4074, doi:10.1073/pnas.0909950107.

20. Williamson, N.R.; Simonsen, H.T.; Ahmed, R.A.A.; Goldet, G.; Slater, H.; Woodley, L.; Leeper, F.J.; Salmond, G.P.C. Biosynthesis of the red antibiotic, prodigiosin, in Serratia: Identification of a novel 2-methyl-3-n-amyl-pyrroie (MAP) assembly pathway, definition of the terminal condensing enzyme, and implications for undecylprodigiosin biosynthesis in Streptomyces. Mol. Microbiol. 2005, 56, 971–989, doi:10.1111/j.1365-2958.2005.04602.x.

21. Sakai-Kawada, F.E.; Ip, C.G.; Hagiwara, K.A.; Awaya, J.D. Biosynthesis and bioactivity of prodiginine analogs in marine bacteria, Pseudoalteromonas: A mini review. Front. Microbiol.

2019, 10, 1–9, doi:10.3389/fmicb.2019.01715.

22. Song, C.; Kidarsa, T.A.; van de Mortel, J.E.; Loper, J.E.; Raaijmakers, J.M. Living on the edge: emergence of spontaneous gac mutations in Pseudomonas protegens during swarming motility. Environ. Microbiol. 2016, 18, 3453–3465, doi:10.1111/1462-2920.13288.

23. Yan, Q.; Lopes, L.D.; Shaffer, B.T.; Kidarsa, T.A.; Vining, O.; Philmus, B.; Song, C.; Stockwell, V.O.; Raaijmakers, J.M.; McPhail, K.L.; et al. Secondary metabolism and interspecific competition affect accumulation of spontaneous mutants in the GacS-GacA regulatory system in Pseudomonas protegens. MBio 2018, 9, 1–18, doi:10.1128/mBio.01845-17.

24. Haddix, P.L.; Shanks, R.M.Q. Production of prodigiosin pigment by serratia marcescens is negatively associated with cellular ATP levels during high-rate, low-cell-density growth. Can. J. Microbiol. 2020, 66, 243–255, doi:10.1139/cjm-2019-0548.

25. Price, M.N.; Wetmore, K.M.; Deutschbauer, A.M.; Arkin, A.P. A comparison of the costs and benefits of bacterial gene expression. PLoS One 2016, 11, 1–22, doi:10.1371/journal.pone.0164314.

26. Gram, L.; Melchiorsen, J.; Bruhn, J.B. Antibacterial activity of marine culturable bacteria collected from a global sampling of ocean surface waters and surface swabs of marine organisms. Mar. Biotechnol. 2010, 12, 439–451, doi:10.1007/s10126-009-9233-y.

27. Östling, J.; Goodman, A.; Kjelleberg, S. Behaviour of IncP‐1 plasmids and a miniMu transposon in a marine Vibrio sp.: isolation of starvation inducible lac operon fusions. FEMS Microbiol. Lett. 1991, 86, 83–94, doi:10.1111/j.1574-6968.1991.tb04797.x.

28. Kim, E.; Kinney, W.H.; Ovrutsky, A.R.; Vo, D.; Bai, X.; Honda, J.R.; Marx, G.; Peck, E.; Lindberg, L.; Falkinham, J.O.; et al. A surface with a biomimetic micropattern reduces colonization of Mycobacterium abscessus. FEMS Microbiol. Lett. 2014, 360, 17–22, doi:10.1111/1574- 6968.12587.

29. Cock, P.J.A.; Fields, C.J.; Goto, N.; Heuer, M.L.; Rice, P.M. The Sanger FASTQ file format for sequences with quality scores, and the Solexa/Illumina FASTQ variants. Nucleic Acids Res. 2009, 38, 1767–1771, doi:10.1093/nar/gkp1137.

30. Li, H.; Durbin, R. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics 2009, 25, 1754–1760, doi:10.1093/bioinformatics/btp324.

31. Li, H.; Handsaker, B.; Wysoker, A.; Fennell, T.; Ruan, J.; Homer, N.; Marth, G.; Abecasis, G.; Durbin, R. The Sequence Alignment/Map format and SAMtools. Bioinformatics 2009, 25, 2078–

2079, doi:10.1093/bioinformatics/btp352.

32. Wang, K.; Li, M.; Hakonarson, H. ANNOVAR: Functional annotation of genetic variants from high-throughput sequencing data. Nucleic Acids Res. 2010, 38, 1–7, doi:10.1093/nar/gkq603.

33. Abyzov, A.; Urban, A.E.; Snyder, M.; Gerstein, M. CNVnator: An approach to discover, genotype, and characterize typical and atypical CNVs from family and population genome sequencing. Genome Res. 2011, 21, 974–984, doi:10.1101/gr.114876.110.

FIGURE AND TABLE LEGENDS

Figure 1. Wild type (red) and Spontaneous mutants (non-pigment) of Pseudoaleromonas rubra S4059 after 8 days growth on natural shrimp shells.

Figure 2. Chemical profiles (A) and anti-Staphylococcus aureus 8325 activities (B) of Pseudoalteromonas rubra S4059 wild type (WT) and the three randomly picked spontaneous mutants S1 (spont_1), F1 (spont_2) and Fi1 (spont_3). Mannose: medium control, medium extractions, pigC: a pigC gene in-frame deletion mutant as a standard to show the retention time of the prodigiosin precursor, 4-methoxy-2, 2’- bipyrrole-5-carbaldehyde (MBC). 1, 2: the culture extractions of wild type; 3, 4: the culture extractions of spont_1; 5, 6: the culture extractions spont_2; 7, 8: the culture extractions of spont_3. All experiments were performed in bio-replicates.

Figure 3. Growth kinetics of Pseudoalteromonas rubra S4059 wild type and three randomly picked mutants (S1, F1 and Fi1) when culturing in marine minimal medium containing mannose. Red lane: S4059 wild type; blue lane: S1; grey lane: F1; yellow lane: Fi1; error bar: standard error.

Table 1. The numbers of SNPs, Indels and CNVs were detected in the nine strains.

Table 2. The mutation type and position of the six strains in a diguanylate cyclase gene.

Figure 1. Wild type (red) and spontaneous mutants (non-pigment) of Pseudoaleromonas rubra S4059 after eight days growth on natural shrimp shells or chitin flakes.

Figure 2. Chemical profiles (A) and antibacterial activities (B) of Pseudoalteromonas rubra S4059 wild type (WT) and the three randomly picked spontaneous mutants S1 (spont_1), F1 (spont_2) and Fi1 (spont_3). Mannose: medium control, medium extractions; pigC: a pigC gene in-frame deletion mutant as a standard to show the retention time of the prodigiosin precursor, 4-methoxy-2, 2’- bipyrrole-5-carbaldehyde (MBC). 1, 2: the culture extractions of wild type; C: medium extraction negative control. All experiments were performed in bio- replicates.

0.4

0.2

-0.2

-0.4

-0.6 Log10(OD600) -0.8

-1

-1.2 0 1000 2000 3000 4000 5000 Time (mins)

Figure 3. Growth kinetics of Pseudoalteromonas rubra S4059 wild type and three randomly picked mutants (S1, F1 and Fi1) when culturing in marine minimal medium containing mannose. Red lane: S4059 wild type; blue lane: S1; black lane: F1; green lane: Fi1; error bar: standard error.

Table 1. The numbers of SNPs, Indels and CNVs were detected in the nine strains.

Samples SNPs Indels CNVs S1 113 2 2 S2 116 4 0 S3 95 4 164 F1 87 5 144 F2 98 5 118 F3 94 4 137 Fi1 95 3 109 Fi2 97 4 150 Fi3 95 1 140

Table 2. The mutation type and position of the six strains in a diguanylate cyclase gene.

mutation insertion Samples position type sequences S1 3395305 insertion AC S2 3394536 insertion GTCAGGT S3 3395305 insertion AC Fi1 3393782 insertion C Fi2 3393782 insertion C Fi3 3393782 insertion C