Diversity, abundance and ecological function of fungi and viruses in marine sponges.

A thesis in fulfillment of the requirements for the degree of Doctor of Philosophy Thi Hong Duc (Mary) Nguyen

Supervisor: Torsten Thomas

School of Biotechnology and Biomolecular Sciences

Faculty of Science

December 2018 i ii iii iv v Abstract The sponge holobiont can be considered as a complex ecosystem of the host, the microbiota and the interactions between them. Sponges are known to harbour diverse microorganisms, including bacteria, archaea, eukaryotes (such as fungi) and recently viruses. However, most studies have focused on the abundance and diversity of prokaryotes, while very little is known about sponge-associated fungi and viruses. Therefore, this thesis aimed to elucidate the gaps in our current knowledge of sponge symbiosis and provide insights to their potential functions.

Sponge fungal diversity was investigated using traditional cultivation and ITS amplicon sequencing over two time periods. Our results indicated that there was relatively low fungal diversity and low host-specificity in the sponges studied here, which broadly reflected the fungal community in seawater.

The diversity, abundance and functional analysis of viruses associated with the microbial cell fraction were investigated through metagenomic and (meta)transcriptomic analysis. Viral metagenomes were dominated by bacteriophages from the order Caudovirales (Myoviridae, Siphoviridae and Podoviridae), while eukaryotic viruses such as Phycodnaviridae and Pandoraviruses were highly expressed. Sponges appeared to contain both variable environment-derived and sponge-specific viral assemblages. Functional gene analysis revealed common features of viruses in all host environments, such as DNA replication and virion production. Interestingly, sponge viromes also contained diverse auxiliary metabolic genes, which vary with different host environments.

To elucidate virus-host dynamics, a virus-host model system was established using Ruegeria phage 67, isolated from the sponge-associated bacteria Ruegeria sp. AU67. Infection growth dynamics and phage genomic analysis suggested that the phage was pseudolysogenic. Phage infection induced drastic morphological changes in the bacterial host, which is postulated to enhance the bacteria’s resistance to phagocytosis. Thus, the virus-host interaction may be mutualistic, increasing the survival of both bacteria and phage. Chemical extracts from the host sponge were found to inhibited bacterial growth and subsequently virion production, which suggests that sponges may contain specific mechanisms to maintain a stable holobiont.

Lastly, fungal pathogens and viruses infecting pathogens were detected, indicating that sponges may act as reservoirs for the survival and spread of pathogens. This has global ecological implications on the health and disease of reef ecosystems.

vi This thesis is dedicated to my mother,

Thi Kim Phuong Phan

whose courage, perseverance and optimism has been my inspiration.

vii Acknowledgements

Firstly, I would like to thank my supervisors, Torsten Thomas (UNSW) and Cristina Moraru (University of Oldenburg), for your support, guidance and knowledge. Without your continued patience and passion, this thesis would not have been possible. You have not only been amazing supervisors and mentors but also a friend.

I would like to thank all my friends, fellow students, colleagues and collaborators at UNSW and University of Oldenburg, for your guidance and assistance in not only the scientific field but also in my personal development. Most of all, thank you for your friendship and compassion throughout the years and I will cherish our shared experiences.

Lastly, I would like to thank my family, Maria, An and especially my mother, Phuong to which I owe everything. To my sister, thank you for your generosity, love and lessons in perseverance, compassion and strength. To my brother, thank you for the lessons in patience, understanding and empathy. And to my mother, thank you for your courage and sacrifice, your unwavering support and dedication. Who has taught me to work with diligence and to endure hard work and labour with little complaint or show.

viii Table of Contents Acknowledgements………………………………………………………………………………………………….v List of Figures ...... Error! Bookmark not defined. List of Tables ...... Error! Bookmark not defined. 1 Introduction ...... 1 1.1 The importance of sponges in reef ecosystems ...... 1

1.2 The sponge host ...... 3

1.3 Sponge symbionts and their significance to sponge heath and function...... 5

1.4 Under-explored sponge symbionts: fungi and viruses ...... 7

1.5 Future perspective ...... 9

2 Host-specificity and stability of sponge-associated fungal communities of co- occurring sponges ...... 11 2.1 Co-authors and collaborators ...... 11

2.2 Introduction ...... 11

2.3 Materials and Methods ...... 14

2.3.1 Sample collection ...... 14

2.3.2 Cultivation of fungi ...... 14

2.3.3 Identification of fungal isolates...... 15

2.3.4 Sequence analysis of ITS amplicon community profiling ...... 16

2.3.5 Sequence analysis of 16S rRNA community profiling ...... 17

2.3.6 Statistical analysis ...... 17

2.4 Results ...... 18

2.4.1 Diversity of fungal communities through culture-dependent and independent methods ...... 18

2.4.2 Analysis of the temporal stability and host-specificity of sponge- associated fungal communities ...... 20

2.5 Discussion ...... 26

2.5.1 Assessment of fungal diversity in sponges ...... 26

ix 2.5.2 Host specificity of sponge-associated fungi ...... 28

2.5.3 Potential roles of sponge-associated fungi ...... 30

2.6 Conclusion ...... 33

3 Taxonomic, functional and transcriptomic analysis of viral communities associated with marine sponges...... 34 3.1 Co-authors and collaborators ...... 34

3.2 Introduction ...... 34

3.3 Materials and Methods ...... 38

3.3.1 Metagenomic datasets of microbial cells associated with sponges ...... 38

3.3.2 Transcriptome and metatranscriptome ...... 39

3.3.3 Identification and normalisation of viral contigs ...... 40

3.3.4 Taxonomic and phylogenetic classification of viral contigs ...... 41

3.3.5 Functional annotation of viral contigs ...... 42

3.4 Results ...... 42

3.4.1 Viral diversity associated with microbial cells in sponges ...... 42

3.4.2 Viral communities in the metatranscriptome and transcriptome datasets 46

3.4.3 Functional gene analysis of viral contigs associated with microbial cell metagenomes ...... 48

3.4.4 Functional gene analysis of viral contigs found in (meta)transcriptomes 51

3.5 Discussion ...... 53

3.5.1 Viral assemblages associated with microbial cells in sponges ...... 53

3.5.2 Diversity of active viral consortia in sponges ...... 54

3.5.3 Virus-host dynamics within the sponge environment ...... 56

3.5.4 Functional contribution of the virome to the sponge holobiont ...... 57

3.6 Conclusion ...... 61

x 4 Interactions of a novel phage and the sponge-associated bacteria Ruegeria sp. AU67 ...... 64 4.1 Co-authors and collaborators ...... 64

4.2 Introduction ...... 64

4.3 Materials and Methods ...... 68

4.3.1 Sponge sampling and viral extraction ...... 68

4.3.2 Sponge extract preparation ...... 69

4.3.3 Isolation of sponge-associated bacteria ...... 69

4.3.4 Viral isolation and purification ...... 70

4.3.5 Transmission electron microscopy ...... 71

4.3.6 Bacterial genome sequencing ...... 71

4.3.7 Viral genome sequencing and phylogenetic analysis ...... 71

4.3.8 Bacterial growth curves ...... 72

4.3.9 Plaque assays and one-step infection growth curves ...... 73

4.3.10 Bacterial cell count via flowcytometry ...... 73

4.3.11 Microscopy of phage-host infections ...... 74

4.4 Results ...... 74

4.4.1 Virus isolation ...... 74

4.4.2 Phage-host dynamics ...... 75

4.4.3 Morphological changes in the bacterial host induced by viral infection .. 76

4.4.4 Genomic analysis of Ruegeria sp. AU67 and Ruegeria phage 67 ...... 77

4.4.5 Phage genome comparison ...... 80

4.5 Discussion ...... 82

4.5.1 Isolation of lytic viruses from sponge-associated bacteria ...... 82

4.5.2 Phylogenetic and genomic analysis of Ruegeria phage 67 ...... 84

4.5.3 The life-cycle of Ruegeria phage 67 ...... 85

4.5.4 Morphological changes in host cells induced by viral infection ...... 88

xi 4.5.5 Development of phage resistance ...... 90

4.5.6 Impact of sponge extracts on the phage-host dynamics ...... 90

4.6 Conclusion ...... 91

5 Conclusions and future directions ...... 93 5.1 Summary ...... 93

5.2 The sponge holobiont ...... 96

5.3 Molecular mechanisms of sponge-symbiosis interactions ...... 97

5.4 Implications of sponges as reservoirs of pathogens in global epidemiology ...... 99

5.5 Future directions ...... 101

References ...... 102 6 Appendix ...... 125 6.1 Chapter Two ...... 125

6.2 Chapter Three ...... 131

6.3 Chapter Four ...... 158

xii List of Figures

Chapter One

Figure 1.1 Schematic diagram of the structure of a sponge ...... 4 Figure 1.2 Schematic diagram of symbiotic relationships between sponges and microorganisms adapted from Lee et al. (2001)...... 5

Chapter Two

Figure 2.1 Presence (black)-absence (white) map of the fungal OTUs obtained from ITS-amplicon sequencing (brown) and cultivation (green) for samples collected in 2014...... 20 Figure 2.2 Heatmap of “variable/core” sponge fungal communities, with OTUs only found in seawater removed. ...25 Figure 2.3 Heatmap of the bacterial community composition based on 16S rRNA gene sequences for the sponge samples collected in 2014...... 26

Chapter Three

Figure 3.1 Viral communities of sponges and seawater...... 45 Figure 3.2 (Meta)transcriptome analysis of viral community in sponges...... 48 Figure 3.5 Functional gene analysis of sponges and seawater viral (pink) and whole (viral and prokaryotic) (brown) metagenomes...... 49 Figure 3.6 Genes associated with KEGG pathways of sponges and seawater viral (pink) and whole (viral and bacterial) (brown) metagenome...... 50 Figure 3.7 Functional auxiliary proteins in virus associated with the microbial cell fraction...... 50 Figure 3.8 Functional gene analysis of sponge viral (pink) and whole (viral and prokaryotic) (brown) (meta)transcriptomes...... 52 Figure 3.9 Genes associated with KEGG pathways of sponge viral (pink) and whole (viral and prokaryotic) (brown) (meta)transcriptomes...... 52

Chapter Four

Figure 4.1 TEM images of Ruegeria phage 67...... 75 Figure 4.2 One-step infection of Ruegeria phage 67 in marine broth and marine broth amended with 10% w/v of sponge extracts from T. anhelans, Scopalina sp. and C. concentrica...... 76 Figure 4.3 Morphological changes observed in phage infections of Ruegeria sp. AU67...... 78 Figure 4.4 Ruegeria phage 67 genome annotated via IMG and visualised with Snap gene ...... 79 Figure 4.5 Phylogenetic analysis of Ruegeria phage 67 (highlighted in bold) using the DNA polymerase protein sequence...... 80 Figure 4.6 Multiple genome comparison using the Mauve program ...... 82

xiii List of Tables

Chapter Two

Table 2.1 PERMANOVA of fungal communities based on presence-absences of OTUs obtained by cultivation and ITS- amplicon sequencing of sponges and seawater...... 19 Table 2.2 Summary of ITS amplicon sequencing results of 2014 and 2016 datasets...... 22 Table 2.3 Summary of the percentages of different fungal categories in sponge and seawater...... 23 Table 2.4 Alpha diversity of “variable/core” fungal community as measured by the mean Shannon’s index ± standard deviation (sd)...... 23 Table 2.5 PERMANOVA analysis (based on relative abundance and presence-absence values) of “variable/core” fungal communities (normalised) at an OTU-level of sponges and seawater...... 24

Chapter Three

Table 3.1 Summary of the number of viral contigs found in each sample metagenome...... 43

Chapter Four

Table 4.1 Sponge-associated bacterial cultures used to isolate viruses...... 70

xiv Chapter One

1 Introduction

1.1 The importance of sponges in reef ecosystems

Sponges (phylum Porifera) are among the oldest multicellular organisms (metazoan) and can be divided into four classes; Hexactinellida, Calcarea,Demospongiae and Homoscleromorpha (Gazave et al., 2011). There are more than 6000 sponge species described inhabiting a wide variety of aquatic environments from shallow freshwater to deep oceans, tropical waters, temperate and even in polar regions (Hooper and Van Soest, 2002). Sponges make up a large portion of the benthic community and play crucial roles in the reef ecosystems including; 1) nutrient cycling, such as of carbon, silicon, oxygen and nitrogen, 2) facilitating primary and secondary production and 3) provision of microhabitats for a range of fauna and flora (Bell, 2008).

Sponges filter large quantities of water, from which they remove food particles (carbon source) and other nutrients (e.g. oxygen, silicon and nitrogen), thereby potentially significantly impacting pelagic ecosystems. This interaction between the benthic and pelagic environments has been termed ‘bentho-pelagic coupling’ (Reiswig, 1974). Feeding of sponges on plankton (which includes bacteria, archaea, algae, protozoa and drifting microscopic animals) enables the carbon flow to higher trophic levels through predation of the sponge itself (Wulff, 2006). Sponges are a food source for a wide range of organisms, including fish, opisthobranchs, crustaceans, molluscs (nudibranchs) and echinoderms, including star fish and sea urchins (Wulff, 2006).

Sponges play an important role in nutrient cycles by taking up dissolved organic matter (DOM) and transferring the DOM to higher trophic levels by rapidly expelling filter cells as detritus, which is subsequently consumed by reef fauna (Yahel et al., 2003, De Goeij et al., 2013). The deposition of silicon (Si) is a fundamental process in the production of sponge skeletons and thus are considered to have an important impact on global Si cycling, particularly as a Si sink (Maldonado et al., 2005a). Nitrogen cycling in sponges

1 and its release to the water column for use by other organisms is thought to be undertaken primarily by associated cyanobacteria, which fix atmospheric nitrogen (Wilkinson and Fay, 1979). Ammonia-oxidizing and nitrite-oxidizing microorganisms have also been reported in sponges (Bayer et al., 2008, Mohamed et al., 2010). Sponges and their associated photosynthetic organisms, such as cyanobacteria and to a lesser extent dinoflagellates, have been reported to contribute to the primary production of coral reefs in tropical waters (Wilkinson, 1983, 1987). For example, Wilkinson (1983) found that 6 out of the 10 most common sponge species in Davies Reef (Great Barrier Reef) were net primary producers. Sponge populations have the potential to cause cascading ecosystem-level effects due to their high abundance in benthic ecosystems and their important role in nutrient cycling (Butler IV et al., 1995). For example, decreases in sponge populations have been correlated with increased occurrences of phytoplankton blooms (Peterson et al., 2006).

Sponges support diverse macrofaunal and to a lesser extent macrofloral communities (Villamizar and Laughlin, 1991, Ribeiro et al., 2003, Wulff, 2006, Taylor et al., 2007). For example, Ribeiro et al. (2003) reported over 2000 individual organisms of 75 invertebrate species (belonging to 9 phyla) associated with the encrusting sponge Mycale microsigmatosa. Likewise, Villamizar and Laughlin (1991) reported 139 and 53 species inhabiting the two Caribbean vase-shaped sponges Aplysina lacunosa and Aplysina archeri, respectively. The most commonly represented macrofauna are crustaceans, polychaetes, ophiuroids, cnidarians, molluscs, and fish (Wulff, 2006). Sponges have also been reported to associate with macroalgae and flowering plants, such as coralline red algae Jania adherens (Wulff, 2006), vascular plants (seagrass) in estuaries (Fell and Lewandrowski, 1981) and water hyacinth roots (Tavares et al., 2005).

Abundant and extremely diverse microbial communities, including bacteria, archaea, eukaryotes (such as fungi, protists and diatoms) and viruses, have also been reported to associate with marine sponges (Taylor et al., 2007, Thomas et al., 2016, Pita et al., 2018). Interestingly, apparently healthy individuals of the sponges Agelas tubulata and Amphimedon compressa from Florida reefs were found to harbour microbes associated with coral disease (Negandhi et al., 2010). Additionally, the pathogenic

2 fungi Aspergillus sydowii, which is the causative agent of sea fan disease in the Caribbean, was isolated from healthy Spongia obscura in the Bahamas (Ein-Gil et al., 2009). This suggest that sponges may act as reservoirs for putative marine pathogens, which has significant ecological implications on global reef health and disease (Webster, 2007, Webster and Taylor, 2012).

1.2 The sponge host

Sponges are sessile filter feeders, which pump large volumes of water (up to 24 m3 of seawater day-1 kg sponge-1) through specialised aquiferous systems, comprising of chambers, ostia, osculum and canals, in which organic particulates and microbes are trapped and consumed by ameobocytes in the mesohyl (Hentschel et al., 2012). The outer layer and interior canals (picoderm) are formed by epithelial cells called pinacocytes. Inside the sponge, flagellated cells (choanocytes) beat to draw water through the ostia into a series of chambers, where feeding takes place and water is expelled through the osculum (Figure 1a) (Hentschel et al., 2012). The choanocytes filter food particles from the water and transferred them to the mesohyl, where the food particles are ingested by cells called amoebocytes (also known as archaeocytes). The omnipotent amoebocytes are responsible for nutrition uptake, excretion, gas exchange and also forms part of sexual bodies (oocytes and germocytes) during reproduction (Taylor et al., 2007). The mesohyl is an extensive layer of tissue, that can contain collagenous and amorphous material, such as spongin, as well as spicules that provide structural support (Figure 1b) (Hentschel et al., 2012). Despite the simple body plan of sponges, sponge morphology is extremely diverse with a colourful array ranging from encrusting, branching, cup-shaped and amorphous types, and range in size from a few millimetres to more than a meter in width (Hooper and Van Soest, 2002, Hentschel et al., 2012).

3

Figure 1.1 Schematic diagram of the structure of a sponge, a) Aquiferous body plan of a sponge, b) Close up diagram of the structure of different layers of cells in the sponge (Hentschel et al., 2012).

Marine sponges are known to harbour dense microbial communities from all three domains of life; archaea, bacteria and eukaryotes (fungi and microalgae), comprising of up to 40% of the sponges’ biomass and play an important role in host functions (Taylor et al., 2007, Webster and Taylor, 2012). Symbionts can be found everywhere in the sponge, both intra- and extracellularly (Figure 1.2), and each symbiont seems to have a specific habitat inside the sponge (Lee et al., 2001a). Exosymbiont are present on the outer layers of sponges, endosymbionts are present in the mesohyl and intracellular or inter-nuclear symbionts reside in sponge cells or nuclei (Figure 1.2) (Lee et al., 2001a). These microorganisms have been reported to benefit the host via diverse mechanisms, such as chemical defence through the production of secondary metabolites, contribution to the mechanical structure, transportation of waste, and provision of nutrients and nitrogen fixation (Taylor et al., 2007). While the benefits for the symbionts inside the sponge host may include a more nutrient-richer environment compared to seawater and sediments as well as a ‘safe’ environment from predation.

Metagenomic analyses have uncovered exceptional bacterial diversity in sponges (Webster et al., 2010, Thomas et al., 2016) and also an increased understanding of molecular interactions between bacterial symbionts and sponges (Thomas et al., 2010,

4 Hentschel et al., 2012, Nguyen et al., 2014, Reynolds and Thomas, 2016, Webster and Thomas, 2016). Additionally, proteomic and metatranscriptomic data has provided insights into possible symbiont functions (Liu et al., 2011, Fan et al., 2012, Díez‐Vives et al., 2017, Díez‐Vives et al., 2018).

Figure 1.2 Schematic diagram of symbiotic relationships between sponges and microorganisms adapted from Lee et al. (2001).

1.3 Sponge symbionts and their significance to sponge heath and function.

Sponges form symbiotic relationships with complex microbial communities and have been reported to play an important role in the health and function of sponges including metabolic exchange, provision of nutrients and host defense against predation and pathogens. Symbiotic cyanobacteria may benefit host sponges through nitrogen fixation and supplemented nutrition (Wilkinson and Fay, 1979). For example, cyanobacteria have been report to provide > 50% of the energy requirements in tropical (phototrophic) sponges (Wilkinson, 1983). Additionally, symbiotic microorganisms have been thought to be a possible food source for sponges (Ilan and Abelson, 1995). This has been postulated for the symbiotic interaction between methanotrophic bacteria and deep-sea cladorhizid sponges, which possess no aquiferous system and thus obtain a significant portion of their nutrition from the consumption of their associated methanotrophs (Vacelet et al., 1995, Vacelet and Boury-Esnault, 2002). Sponge symbionts have also been reported to be enriched in

5 genes related to the synthesis of vitamins, such as vitamin B1 (thiamine), vitamin B2 (riboflavin), vitamin B7 (biotin) and B12 (cobalamin), which may benefit the sponge host, which is unable to synthesis these essential vitamins (Thomas et al., 2010, Fan et al., 2012, Fiore et al., 2015, Webster and Thomas, 2016, Lackner et al., 2017).

Sponge-associated microorganisms have also been suggested to play a role in host defense through the production of a wide range of biologically active metabolites (Unson et al., 1994, Schmidt et al., 2000, Sipkema et al., 2005). Various sponge and symbiont-derived compounds have been reported, such as terpenoids, alkaloids, peptides and polyketides with a wide range of properties including anticancer, antibacterial, antifungal, antiviral, anti-inflammatory and antifouling (Sipkema et al., 2005). Some bioactive metabolites (tentatively ascribed to be produced by bacterial symbionts) have been reported to protect the sponge from pathogens, predation and fouling (Unson et al., 1994, Schmidt et al., 2000, Hentschel et al., 2001). For example, Aplysina sponges contained high concentration of brominated metabolites with antimicrobial activity, which are believed to serve as a chemical defense against predators and biofouling (Thompson et al., 1983, Weiss et al., 1996).

Occurrences of sponge disease have increased dramatically over the last two decades with reports of decimated sponge populations in the Mediterranean and Caribbean (Webster, 2007). Despite these epidemics, which can have severe impacts on the ecology of the reef, little is known about the causative agents of sponge disease. Only one reported case identified an Alphaproteobacteria strain as the primary pathogen of sponge disease in Rhopaloeides odorabile (confirming Koch’s postulates) (Webster et al., 2002). Other potential disease agents have been suggested including fungi, viruses and bacteria, such as cyanobacteria, Bacillus and Pseudomonas (Webster, 2007). The correlation of sponge disease and environmental factors such as climate change and urban/agricultural runoff have also been reported (Webster, 2007). Recently, changes in the microbial community structure have been correlated with the development of ‘disease-like’ symptoms in sponges (Webster, 2007, Webster et al., 2008, Fan et al., 2013). Temperature-stress experiments revealed an immediate stress response from both the host and symbiont community and disruptions to functions that mediate symbiosis (Fan et al., 2013). Fan et al. (2013) postulates that the loss of symbionts and

6 consequently the introduction of new microorganisms with scavenging lifestyles (such as fungi) and high growth rates (potential opportunistic pathogens) is a major determinant for mortality of marine sponges (Webster, 2007, Fan et al., 2013). Therefore, the balance of the sponge microbial community is essential to health and function of sponges and subsequently the reef ecosystem (Pita et al., 2018). Environmental disturbance, such as climate change, can drastically impact the microbial communities of sponges, which could result in mortality or disease. Thus, more research is needed to understand the diversity, stability and dynamic of the sponge microbial communities.

1.4 Under-explored sponge symbionts: fungi and viruses

Bacterial symbionts and their potential functions in sponges have been extensively studied, however very little is known about sponge-associated fungi and viruses. Despite the prevalence of fungi in sponges (hundreds of fungal species have been identified from marine sponges) (Holler et al., 2000, Taylor et al., 2007), very few studies have investigated the symbiotic interaction or ecological function of sponge- associated fungi.

Identification of sponge-specific fungi are difficult due to the sponges’ filter feeding activity, which can contain fungal spores from terrestrial environments. Additionally, marine fungi are an ecological group (not a taxonomic group) and therefore can be difficult to determine whether the fungal species are obligate or facultative marine species or terrestrial contaminants (Bugni and Ireland, 2004, Taylor et al., 2007). Nevertheless, fungi may reside in sponges temporarily from terrestrial wash off and form symbiotic relationships over time (Baker et al., 2009). Fungal isolates from sponges have been found to differ from those in the surrounding seawater (Holler et al., 2000) and sponge-associated fungi contained different metabolic activities, such as production of novel secondary metabolites, not found in their terrestrial counterparts (Holler et al., 2000, Raghukumar, 2008). One exceptional case has been reported indicating a true symbiotic relationship by showing the vertical transmission of an unidentified yeast in the sponge genus Chondrillia (Maldonado et al., 2005b). Other

7 indirect evidence of symbiosis was inferred from the mitochondrial intron in the sponge Tetilla sp., which was believed to originate from fungi through cross-kingdom horizontal gene transfer (Rot et al., 2006). Furthermore, Rozas et al. (2011) obtained fungal species directly from in vitro cell cultures and single cells of the sponge Hymeniacidon sp. which were not detected in whole tissue samples and thus the fungal isolates were postulated to be intracellular symbionts. Molecular techniques have been used to analyse fungal communities in marine sponges, and suggested that there were host-specific between sponge species and the surrounding seawater (Gao et al., 2008). In contrast, a recent study by Naim et al. (2017) employed pyrosequencing of 18S ribosomal RNA gene amplicon to investigate the diversity of fungi communities in sponges and surrounding seawater and found that the presence of fungi in these sponges may be accidental and not sponge species-specific. Therefore, greater research efforts are needed to assess the relationship between fungi and sponges.

Considering the high abundance of viruses in seawater (10-7 viral particle/mL) (Wommack and Colwell, 2000) and the large filter feeding capacity of sponges (Vogel, 2008), they would be expect to contain high viral loads. Indeed, recent metagenomic analysis of sponge microbiomes have reported high abundances of viral defense mechanisms (Fan et al., 2012, Horn et al., 2016) and diverse viral communities (Laffy et al., 2018). For example, high abundances of cyanophages (detected via the viral capsid protein G20) were found in the Great Barrier Reef sponge Stylissa sp. 445 (Fan et al., 2012). Bacteriophages (order Caudovirales) including Myoviridae, Siphoviridae and Podoviridae were found to dominate the viral community in four tropical sponges (Amphimedon queenslandica, Xestospongia testudinaria, Ianthella basta and Rhopaloeides odorabile) (Laffy et al., 2018). Eukaryotic viruses (Mimiviridae and Phycodnaviridae) and ssDNA (Microviridae, and Circoviridae) were also detected at variable abundances in these sponges. Interestingly, diverse auxiliary genes were also identified in the sponge viromes, including genes involved in herbicide resistance, viral pathogenesis pathways and cobalamin (vitamin B12) biosynthesis (Laffy et al., 2018).

Viruses are known to influence microbial abundance and community compositions in the ocean via bacterial infection and lysis (Breitbart, 2012). Environmental factors,

8 such as temperature stress and chemical/physiological cues, can induce temperate phages to switch from a lysogenic to a lytic life cycle causing lysis of host cells, which can directly affect microbial populations and structure (Chu et al., 2011). With the high abundance of potential hosts for viral infections in sponges, viruses may play an important role in the microbial ecology and evolution through phage mediated horizontal gene transfer (Wommack and Colwell, 2000, Weinbauer and Rassoulzadegan, 2004, Weitz et al., 2015).

The recognition that macroorganisms live in symbiotic association with microbial communities has opened a new approach to the study of animals, plants and macroalgae, termed the “holobiont” (Margulis, 1991). The sponge with their microbial communities is considered as one of the most diverse and complex holobionts in the marine environment (Pita et al., 2018). This approach considers the host, the microbiota and the interactions among them and thus the first important approach to defining the holobiont is to characterise the core microbiota including the much understudied fungi and viruses (Qin et al., 2010, Tanca et al., 2017).

1.5 Future perspective

Over the last decade we have seen the field of sponge symbiosis rapidly expanding, especially with the application of next-generation sequencing technologies, which has provided insights into the rare biosphere of sponge microbes and revealed incredible diverse microbial communities (Thomas et al., 2016, Moitinho-Silva et al., 2017). However, there are still areas in sponge symbiosis that have largely been overlooked and require additional research effort. This thesis aims to address some of the gaps in our current knowledge of sponge symbiosis focusing on the abundance, diversity and potential ecological functions of sponge-associated fungi and viruses.

Firstly, diversity of sponge associated fungal communities will be assessed using both cultivation-dependent and independent methods. Fungal-host stability and specificity will be investigated through comparison of fungal communities in sponges to seawater over two time periods and to elucidate the potential functional roles of fungi in sponges. Secondly, the abundance, diversity and expression of sponge-associated

9 viruses will be investigated through metagenome and (meta)transcriptome analysis. Taxonomic and functional analysis will be applied to gather insights into the diversity of viral communities of temperate and tropical sponge species and seawater. Thirdly, to elucidate insights into virus-host dynamics in sponges by establishing virus-host model systems relevant to the sponge microbiome.

10 Chapter Two

2 Host-specificity and stability of sponge-associated fungal communities of co-occurring sponges

2.1 Co-authors and collaborators

This chapter is based on the following publication:

NGUYEN, M. T. & THOMAS, T. 2018. Diversity, host-specificity and stability of sponge- associated fungal communities of co-occurring sponges. PeerJ, 6, 4965.

MN conceived and designed the experiments, performed the experiments, analysed the data, prepared figures and/or tables, authored or reviewed drafts of the paper, approved the final draft. TT conceived and designed the experiments, analysed the data, authored or reviewed drafts of the paper, approved the final draft.

2.2 Introduction

Fungi are ecologically important in terrestrial environments performing vital functions as free-living decomposers, nutrient cyclers, parasites and symbionts. The global fungal richness has been estimated to be between 1.5 and 1.6 million species (Hawksworth, 1991). Most of our current understanding of the ecology and function of fungi are derived from studies of cultivated fungal isolates, mainly from terrestrial environments (Richards et al., 2012). In comparison, little is known about the diversity and ecology of fungi in the marine environment, it is estimated that only ~0.6% of cultured fungi are marine derived (Kis-Papo, 2005, Burgaud et al., 2009). Marine fungi

11 form an ecological, but not a taxonomic group and can be classified, according to Kohlymeyer and Volkmann-Kohlmeyer (1990), as “obligate marine” (i.e. those that grow and sporulate exclusively in marine habitats) and “facultative marine” (i.e. those that are from freshwater and terrestrial milieus, that are able to grow and possibly sporulate in marine environments). As more studies emerge, marine fungi are increasingly considered to play an important ecological role as saprotrophs, parasites or symbionts in the marine ecosystem (Hyde et al., 1998). Fungi have been reported to be associated with various marine organisms, such as algae, coral and sponges (Richards et al., 2012).

To date, 22 orders of Ascomycota and eight orders of Basidiomycota have been found in sponges (Yu et al., 2013), largely through cultivation-dependent approaches (Morrison-Gardiner, 2002, Wang et al., 2008, Li and Wang, 2009), which have mainly focussed on the discovery of biologically active, secondary metabolites (Holler et al., 2000, Taylor et al., 2007, Baker et al., 2009, Liu et al., 2010, He et al., 2014). Ubiquitous fungal genera (e.g. Aspergillus, Penicillium, Trichoderma and Acremonium) have been isolated from numerous sponge species worldwide suggesting that these fungi may be either “generalist” and/or are preferentially detected due to their ease of cultivation (Richards et al., 2012). In fact, cultivation conditions as well as sample preparation methods have likely biased the comparison between studies (Anderson et al., 2003). To overcome these issues, molecular techniques that amplify and sequence the 18S rRNA gene or the internal transcribed spacer (ITS) region directly from DNA extracted from samples have been applied to examine fungal diversity in a range of environments (Gardes and Bruns, 1993, Döring et al., 2000, Anderson et al., 2003, De Beeck et al., 2014). PCR-primer bias towards certain fungal groups or non-targeted DNA, however, are drawbacks of this approach (Wang et al., 2014). Cultivation- dependent and -independent approaches have thus been shown to capture different assemblages of the fungal community in deep-sea sediments (Jebaraj et al., 2010, Singh et al., 2012) and soil (Jeewon and Hyde, 2007).

Few studies have investigated the diversity, ecological function and/ or the nature of sponge-fungi interactions. There are conflicting reports on the host-specificity of

12 fungal communities in sponges. For example, Gao et al. (2008) reported distinct fungal communities between two co-occurring Hawaiian sponges which differ to the surrounding seawater using denaturing gradient gel electrophoresis (DGGE)-based ITS analysis. He et al. (2014) found that the fungal communities differed on the order level between some Antarctic sponge species and Rodriguez-Marconi et al. (2015) further suggested that there was a high degree of host-specificity of fungi in Antarctic sponges. However, it should be noted that both studies mentioned above ((He et al., 2014, Rodríguez-Marconi et al., 2015) lacked biological replications. In contrast, Naim et al. (2017) recently reported low host-specificity and suggested the presence of fungi in sponges to be rather “accidental”.

Aside from these community-wide studies, there has been some observational evidence for sponge-fungi interactions. This includes the vertical transmission of an endosymbiotic yeast in the marine sponge Chondrilla sp. (Maldonado et al., 2005b) and the putative horizontal gene transfer of a fungal mitochondrial intron into the genome of the sponge Tetilla sp. (Rot et al., 2006). In addition, the sponge Suberites domuncula has been suggested to be able to recognise fungi via the ᴅ-glucans on their surfaces (Perović-Ottstadt et al. 2004). Ascomycetes of the genus Koralionastes have been reported to have a unique physical association with crustaceous sponges (Kohlmeyer and Volkmann-Kohlmeyer, 1990). Finally, the ability of many sponge- derived fungi to produce novel bioactive compounds and have been suggested to contribute to host defense (Holler et al., 2000, Proksch et al., 2003, Proksch et al., 2008, Wiese et al., 2011, Yu et al., 2013, Imhoff, 2016). These studies indicate that certain close symbiotic interactions between fungi and sponges exist, however this conclusion seems to be not necessarily supported by cultivation-dependent and – independent community-wide analyses.

The aim of this chapter is to assess the diversity of the fungal community associated with sponges using traditional cultivation methods and ITS community profiling to examine their suitability to describe sponge-associated fungal diversity. Fungal communities of three co-occurring sponges and the surrounding seawater were assessed over two time periods to help elucidate host-specificity, stability and

13 potential core members, which may shed light into the ecological function of fungi in sponges.

2.3 Materials and Methods

2.3.1 Sample collection Sponges were sampled at Bare Island in Botany Bay, NSW, Australia (33° 59’S, 151° 14’E) on two separate occasions, on the 13 November 2014 and on the 4 May 2016. At each sampling event, seawater samples and three specimens of C. concentrica, Scopalina sp. and T. anhelans sponges were collected by SCUBA diving at a depth of 7- 10 m and within an area of about 20 x 20 m. Sampling of sponges was performed under the scientific collection permit P13/0007-1.1 issued by the New South Wales Department of Primary Industries. Sponge specimens were identified by the morphological characteristics and their locality as per a previous study (Fan et al., 2012). Samples were placed individually into Ziploc® bags with seawater and then transported in buckets filled with seawater to the laboratory at ambient temperature (travel time approximately 30 min). Sponge samples were processed immediately for cultivation or frozen at -80°C for DNA extraction. Seawater (200 mL) were vacuum filtered onto 0.22 µm filters (Whatman, Sigma-Aldrich, St. Louis, USA) in replicates and used immediately for cultivation or frozen at -80°C for DNA extraction.

2.3.2 Cultivation of fungi Sponge tissues were processed as previously described by Wang et al. (2018). Briefly, sponge samples were washed three times in sterile calcium/magnesium-free seawater

(CMFSW; 25 g NaCl, 0.8 g KCl, 1 g Na2SO4 and 0.04 g NaHCO3 per 1 L) to remove natural seawater from the sponge and the outer surfaces, samples were sterilized with 70% ethanol. Two different cultivation methods were applied; a) sponges were sliced into thin sections (approximately 1 cm2 and 1-2mm thick) and placed directly onto agar plates (listed below) and b) sponge tissue (1 mg/ mL) was homogenized at maximum speed for 50-60s with a dispersing homogenizer (Ulta-Turrax TR50, IKA, Selangor, Malaysia). The homogenate was diluted with sterile seawater (100, 10-1, 10-2) and 100 µl of each dilution was plated. For seawater samples two methods were applied a)

14 200ml of seawater were vacuumed filtered onto 0.22 µm Whatman filters (Sigma- Aldrich, St. Louis, MO, USA) and filters were directly placed onto agar plates and b) 1 mL of seawater was directly plated onto agar plates. Triplicates of each sample and preparation method were plated onto three different media: 1) peptone yeast glucose agar (PYG; 1.0 g glucose, 0.1 g yeast extract, 0.5 g peptone and 15 g Difco-bacto agar (Fisher scientific, Pittsburgh, PA, USA) per 1 L, pH 8.0); 2) Dextrose potato agar (BD

TM Difco , Melbourne, Australia) and 3) Gause I (20 g starch, 1.0 g KNO3, 0.5 g K2HPO4,

0.5 g MgSO4·7H2O, 0.5 g NaCl, 0.01 g FeSO4 and 15 g Difco-bacto agar per 1 L). All media was made up with 0.22 µm filtered and autoclaved seawater from Bare Island. The plates were incubated at 18-20°C for 1-2 months until fungi growth was visible. Every isolate was picked and transferred onto new PYG, Potato dextrose and Gause I agar plates. The resulting pure cultures were stored in sterile artificial seawater (ASW;

23.38 g NaCl, 2.41 g MgSO4, 1.19 g MgCl2, 1.47 g CaCl2.2H2O, 0.75 g FCl and 0.17 g

NaHCO3 per 1 L) in 2 mL cryogenic vials (Sigma-Aldrich, St. Louis, MO, USA) at 4°C.

2.3.3 Identification of fungal isolates

DNA extraction from fungal isolates were conducted using the CTAB method (Lee, 1988) with modifications. Briefly, fungal mycelia were added to 1 mL of CTAB buffer, 3- 6 1 mm glass beads (Sigma-Aldrich, St. Louis, MO, USA) and 10 µL of mercaptoethanol and bead beaten (TissueLyser II, Qiagen, Hilden, Germany) at maximum speed for 8 minutes. Samples were heated at 65°C for 10 min, then extracted with phenol:chloroform:isoamyl alcohol (25:24:1) and DNA was precipitated with isopropanol. DNA was dissolved in 50 µL of pure water and used for PCR amplification. PCR was conducted in 25 µL reactions consisting of 12.5 µL of Econotaq master mix, 9.5 µL of water, 1 µL of each forward primer ITS1f-F (10µM) (5’ TTGGTCATTTAGAGGAAGTAA 3’) and reverse ITS4 (5’ TCCTCCGCTTATTGATATGC 3’) (White et al., 1990) and 1 µL of template DNA. PCR conditions were 95°C/ 2 min, then 94°C/ 30s, 53°C/ 30s, 72°C/ 45s for 35 cycles and 72°C/ 5 min. Amplicon products were assessed by gel electrophoresis, cleaned with Exosap-IT (ThermoFisher Scientific, Waltham, MA, USA) and sequenced with the BigDye Terminator v3.1 chemistry (Applied Biosystems, Austin, TX, USA) and an Applied Biosystems 3730 DNA Analyzer at the Ramaciotti Centre for Genomics (University of New South Wales, NSW, Australia).

15 Sequences were manually quality trimmed using Sequences Scanner v1.0 software and forward and reverse sequencing reads were assembled (when applicable) using BioEdit v7.2.5 (Ibis Biosciences, Carlsbad, CA, USA).

2.3.4 Sequence analysis of ITS amplicon community profiling Total DNA were extracted from sponge samples and seawater filters from the two sampling time points using the Power Soil DNA Isolation kit (Qiagen, Hilden, Germany) following the manufacturer’s instructions. Fungal ITS amplicon sequencing was conducted by Molecular Research LP (Mr. DNA, Houston, TX, USA) using the IST1f-ITS4 primers. PCRs were conducted with a HotStarTaq Plus Master Mix kit (Qiagen, Hilden, Germany) under the following conditions: 94 °C/ 3 min, followed by 28 cycles of 94 °C/ 30 s, 53 °C/ 40 s and 72 °C/ 1 min, and a final elongation step at 72 °C/ 5 min. After amplification, PCR products were checked in 2% agarose gel to determine the success of amplification and the relative intensity of bands. PCR samples were pooled together in equal proportions based on their molecular weight and DNA concentrations and purified using calibrated Ampure XP beads. The pooled and purified PCR products were then used to prepare an Illumina DNA library. Sequencing was performed on the Illumina MiSeq sequencing platform and 2  300 bp chemistry following the manufacturer’s guidelines. Because of the variable length of the ITS region (400–800 bp), forward and reverse sequences could often not be assembled into contigs. In addition, reverse reads had generally lower quality than the forward reads and therefore only the forward ITS amplicon sequence reads (300 bp) were quality filtered and analysed together with the isolate sequences from above. All sequences were quality filtered with a maximum expected error threshold of 1 and minimum length of 250 bp and then clustered into operational taxonomic units (OTUs) at 97% similarity using the UPARSE pipeline (Usearch v9.2) (Edgar, 2013). Chimeras were removed using UCHIME (Edgar et al., 2011) and taxon classification of OTUs were conducted using the UNITE ITS reference database (Abarenkov et al., 2010). Lowest taxon classification at a confidence level of 70% or above were considered and checked with the Basic Local Alignment Search tool (BLASTn) against the non-redundant nucleotide database from the National Centre for Biotechnology Information (NCBI) (Appendix Table S2.5). Non- fungal sequences were removed from further analysis. Raw ITS-amplicon sequences

16 are available through the NCBI Sequence Read Archive under Bioproject ID: PRJNA419577, accession number SRP125576.

2.3.5 Sequence analysis of 16S rRNA community profiling Given the high variability of fungal communities observed between biological replicates (see below), we wanted to understand if this is a peculiar aspect of the samples we took and how we processed them or if this is due to real biological variation. We therefore also analysed all samples for the bacterial community composition, which has been shown to be very consistent between replicates of the three sponges analysed here (Fan et al., 2012, Esteves et al., 2016). Bacterial 16S rRNA gene amplicon sequencing of sponge and seawater samples were conducted using primers 515F (5’-GTG CCA GCM GCC GCG GTA A-3’) and 806R (5’-GGA CTA CHV GGG TWT CTA AT-3’) with 2  250 bp chemistry at the Ramaciotti Centre for Genomics (University of New South Wales, Sydney, NSW, Australia), according to the methodology described by Caporaso et al. (2012). Bacterial 16S rRNA sequences were processed using the MiSeq SOP pipeline (Mothur v1.37.3) (Schloss et al., 2009). Briefly, raw forward and reverse sequence reads were assembled into contigs, quality filtered and aligned to the SILVA 16S rRNA gene reference alignment v102 (Quast et al., 2012). Sequences were filtered to only include overlapping regions, pre-clustered to merge all sequences within three mismatches (difference = 3) and checked for chimeras using the UCHIME algorithm (Edgar et al., 2011). To separate chloroplasts from cyanobacteria, sequences were first classified using the SILVA reference v119 (Quast et al., 2012) with a 60% confidence threshold and sequences classified as chloroplasts were removed. The rest of the sequences were re-classified using the RDP training set release 9 (Cole et al., 2014), sequences classified as “unknown” or “mitochondria” were removed before clustering into OTUs at 97% similarity and sub- sampled to the size of the smallest sample (4193 sequences). 2.3.6 Statistical analysis

The fungal OTU matrix of sponge and seawater samples were used to calculate the species richness estimate (Chao1) and Shannon’s index using the summary.single command in Mothur v1.39.0. Fungal community coverage was estimated using Good’s coverage (= 1- (number of singleton OTUs/number of reads)). Comparison of beta- 17 diversity was conducted with permutational multivariate analysis of variance (PERMANOVA) (Anderson, 2001) of Bray-Curtis dissimilarities of relative abundance and presence-absence values at the OTU level. Variability of communities was analysed using Multivariate Homogeneity of Group Dispersions (PERMDISP) (Anderson et al., 2006, Anderson, 2006) based on Bray-Curtis dissimilarities. Heatmaps and statistics were performed in the R statistical program language (R core Team, 2014) using the vegan package (Oksanen et al., 2013).

2.4 Results

2.4.1 Diversity of fungal communities through culture-dependent and independent methods Culture-dependent and -independent approaches were applied to sponge and seawater samples collected in 2014. Cultivation yielded a total of 108 isolates and after redundant sequences were removed, resulted in 42 unique isolate sequences (see Appendix Table S2.1 for details), which clustered into 8 OTUs at 97% similarity. In contrast, 26 OTUs were obtained via ITS-amplicon sequencing. Cultivation obtained one fungal division, Ascomycota, and four fungal orders (Eurotiales, Hypocreales, Diaporthales and Capnodiales) compared to ITS-amplicon sequencing which obtained three divisions (Ascomycota, Basidiomycota and Glomeromycota) and 12 orders (Pleosporale, Capnodiales, Eurotiales, Leotiales, Malasseziales, Xylariales, Helotiales, Lecanorales, Teloschistales, Hypocreales, Agaricales and Diaporthales) (Figure 2.1). Significant differences (PERMANOVA P-value <0.008) in the fungal community compositions were observed overall and for each sponge species and seawater between the cultivation-dependent and -independent approaches (Table 2.1), with the cultivation method producing a lower fungal diversity (Shannon’s index of 0.77 ± 0.32) compared to the ITS amplicon sequencing (1.17 ± 0.54; one-way ANOVA, P- value=0.04). Although a higher fungal diversity was obtained by ITS-amplicon-based community profiling, the cultivation method yielded five unique OTUs not present in the sequencing data. OTU_266 (Aspergillus sp.), OTU_286 (Acrostalagmus sp.) and

18 OTU_269 (Cladosporium sp.) were only cultivated from T. anhelans, OTU_265 (Togniniaceae unclassified) was cultivated from Scopalina sp., and OTU_264 (Trichoderma sp.) was cultivated from C. concentrica, Scopalina sp. and seawater. Three Penicillium spp. were found to overlap between the two methods (OTU_119, OTU_165 and OTU_9). OTU_119 was commonly cultivated from all sample types and detected in the T. anhelans ITS-amplicon sequencing data. The most common OTUs detected in the sequencing data were OTU_4 (Epicoccum sp.) and OTU_6 (Cladosporium sp.) and were observed in all sample types (i.e. three sponge species and seawater) (Figure 2.1).

Table 2.1 PERMANOVA of fungal communities based on presence-absences of OTUs obtained by cultivation and ITS-amplicon sequencing of sponges and seawater.

ITS-amplicon sequencing vs. P-value Cultivation All samples 0.001 C. concentrica 0.001 Scopalina sp. 0.001 T. anhelans 0.001 Seawater 0.008

19

Figure 2.1 Presence (black)-absence (white) map of the fungal OTUs obtained from ITS- amplicon sequencing (brown) and cultivation (green) for samples collected in 2014. Columns were clustered based on Bray-Curtis dissimilarity using hierarchical clustering with the ‘average’ method (scale depicts the percentage of dissimilarity). Samples are indicated in purple: C. concentrica, orange: Scopalina sp. , red: T. anhelans and blue: seawater. Numbers 1, 2 and 3 indicate sample replicates.

2.4.2 Analysis of the temporal stability and host-specificity of sponge- associated fungal communities Since cultivation recovered only a small proportion of the total fungal diversity found, compared to ITS-amplicon sequencing, the latter approach was used to analyse temporal changes and specificity of OTUs to sponge species. A larger number of quality-filtered sequences were obtained from samples collected on the 4 May 2016 (average 13,136 sequences per samples, range of 6,578 - 30,977) compared to the collection effort on the 13 November 2014 (4,062 sequences per samples 64 - 16,199 range). Sequences were clustered into OTUs at a 97% similarity and non-fungal sequences were removed leaving a total of 155,298 sequences and 148 OTUs.

20 Estimation of the Chao1, Good’s coverage and Shannon’s index were conducted on normalised data (sub-sampled to 250 sequences), resulting in the removal of samples S_1_14, S_2_14, S_2_16, S_3_14, SW_1_14 and T_3_14 (sample_type replicate_number year) (Table 2.2). Good’s coverage estimates were greater than 94% for most remaining samples, showing that the majority of OTUs were captured through the ITS-amplicon sequencing effort. Generally, Chao1 estimates positively correlated with the Shannon’s diversity index.

Due to the filter feeding capacities of sponges, we expect the presence of ‘incidental’ environmental fungi in our sponge samples. Fungal OTUs were therefore grouped into three categories; ‘occasional’ (OTUs occurring only once in the six replicates per sample type), ‘variable’ (OTUs occurring in two to five of the six replicates per sample type) and core OTUs (OTUs occurring in all six replicates per sample type). No core OTUs were observed in C. concentrica, Scopalina sp. and seawater, and only one core OTU was observed in T. anhelans (Table 2.3). Fungal communities of all three sponges were predominantly comprised of ‘variable’ OTUs (>60% mean relative abundance).

Occasional OTUs were removed to create the ‘resident/core’ fungal community dataset and was sub-sampled to a size of 250 reads, resulting in a total of 38 OTUs. Similar to the whole community analysis, Shannon’s diversity index of the ‘variable/core’ communities were significantly lower in 2014 compared to 2016 (Table 2.4 and Appendix Table S2.1). ‘Variable/core’ communities of C. concentrica and seawater were significantly different, but fungal diversity of T. anhelans was comparable between the two time points. Temporal samples were combined to analyse the community diversity of each sponge species and seawater. ‘Variable/core’ community of Scopalina sp. had the highest Shannon’s index followed by T. anhelans, C. concentrica and seawater (Table 2.4), in contrast to the whole community analysis where the Shannon’s index was highest for Scopalina sp., followed by seawater, C. concentrica and T. anhelans (Appendix Table S2.1). However, Shannon’s indices for both whole and ‘variable/core’ communities for all samples were comparable to each other (Table 2.4 and Appendix Table S2.1).

21 Table 2.2 Summary of ITS amplicon sequencing results of 2014 and 2016 datasets. Number of reads, observed fungal OTUs, expected fungal OTUs (Chao1), Coverage (Good’s coverage) and Shannon’s index in seawater and sponge samples.

Sample name Abbreviat Total no. No. of Observed Expected Good’s Shannon’s ion of fungal fungal OTUs coverage index (sub- quality- reads OTUs (Chao1) ± (sub- sampled) filtered SE sampled) reads (sub- sampled)

C. concentrica 1 C_1_14 16199 14849 8 3 ± 0 1.00 0.09 2014

C. concentrica 2 C_2_14 3026 1120 4 3 ± 0 1.00 0.90 2014

C. concentrica 3 C_3_14 6198 4808 4 1 ± 0 1.00 0.00 2014

C. concentrica 1 C_1_16 6578 269 24 28 ± 18 0.97 1.94 2016

C. concentrica 2 C_2_16 21700 3437 27 10 ± 3 0.99 0.63 2016

C. concentrica 3 C_3_16 9483 320 30 44 ± 35 0.94 2.07 2016

Scopalina sp. 1 2014 S_1_14 1532 49 2 NA NA NA

Scopalina sp. 2 2014 S_2_14 7071 12 3 NA NA NA

Scopalina sp. 3 2014 S_3_14 1992 107 4 NA NA NA

Scopalina sp. 1 2016 S_1_16 4859 444 29 27 ±15 0.97 1.33

Scopalina sp. 2 2016 S_2_16 3281 206 32 NA NA NA

Scopalina sp. 3 2016 S_3_16 24565 665 36 39 ± 50 0.96 1.38

T. anhelans 1 2014 T_1_14 681 681 2 2 ± 1 1.00 0.03

T. anhelans 2 2014 T_2_14 6947 6703 9 5 ± 4 1.00 0.96

T. anhelans 3 2014 T_3_14 64 8 3 NA NA NA

T. anhelans 1 2016 T_1_16 19481 18300 21 5 ± 4 1.00 1.21

T. anhelans 2 2016 T_2_16 6947 16379 27 9 ± 17 0.99 0.77

T. anhelans 3 2016 T_3_16 30977 30956 19 3 ± 2 1.00 0.72

Seawater 1 2014 SW_1_14 1066 2 2 NA NA NA

Seawater 2 2014 SW_2_14 3379 3314 2 2 ± 1 1.00 0.03

Seawater 3 2014 SW_3_14 2647 1588 3 1 ± 0 1.00 0.00

Seawater 1 2016 SW_1_16 3379 6139 33 12 ± 8 1.00 2.05

Seawater 2 2016 SW_2_16 12439 5576 45 21 ± 9 0.98 2.37

Seawater 3 2016 SW_3_16 25069 19496 40 14 ± 18 0.99 1.24

NA: Non-applicable as samples contained sequences below the 250 reads cut-off.

22 Table 2.3 Summary of the percentages of different fungal categories in sponge and seawater. Numbers in brackets indicate the sum of the mean relative abundances (percentages ± standard deviations).

Sample type Total OTUs Occasional OTUs Variable OTUs Core OTUs C. concentrica 52 24 (11.0% ± 5.2) 28 (88.9% ± 13) 0 (0%) Scopalina sp. 56 27 (35.7% ± 9.3) 29 (64.2% ± 10.7) 0 (0%) T. anhelans 49 32 (11.9% ± 5.4) 16 (73.2% ± 10.2) 1 (19.5% ± 35) Seawater 76 44 (5% ± 3.7) 32 (94% ± 13.7) 0

Table 2.4 Alpha diversity of “variable/core” fungal community as measured by the mean Shannon’s index ± standard deviation (sd). Comparison of diversity between samples were calculated with ANOVA and multiple comparison with Tukey’s test of subsampled ‘resident/core’ fungal communities at an OTU-level. P-values are shown in brackets and those smaller than 0.05 are highlighted in bold.

Temporal samples Shannon’s Index (P-values) 2014 vs. 2016 0.011 ± 0.014 vs. 1.35 ± 0.66 (6.96e-05) C. concentrica 2014 vs. 2016 0.008 ± 0.015 vs. 1.41 ± 0.9 (0.04) Scopalina sp. 2014 vs. 2016 NA vs. 1.38 ± 0.0001 (NA) T. anhelans 2014 vs. 2016 0.013 ± 0.018 vs. 0.75 ± 0.45 (0.62) Seawater 2014 vs. 2016 0.013 ± 0.018 vs. 1.89 ± 0.39 (0.01) Sample type Seawater vs. Scopalina sp. 0.45 ± 0.52 vs. 1.38 ± 0.0001 (0.98) T. anhelans vs. Scopalina sp. 1.14 ± 1.06 vs. 1.38 ± 0.0001 (0.58) T. anhelans vs. Seawater 1.14 ± 1.06 vs. 0.45 ± 0.52 (0.59) Scopalina sp. vs. C. concentrica 1.38 ± 0.0001 vs. 0.71 ± 0.96 (0.77) Seawater vs. C. concentrica 0.45 ± 0.52 vs. 0.71 ± 0.96 (0.84) T. anhelans vs. C. concentrica 1.14 ± 1.06 vs. 0.71 ± 0.96 (0.96)

Fungal communities overall were variable in structure with the average distances from the centroid calculated with PERMDISP of 0.57, 0.46, 0.44, and 0.58 for C. concentrica, Scopalina sp., T. anhelans and seawater, respectively. In contrast, the average distance to centroid for bacterial communities of C. concentrica, Scopalina sp., T. anhelans and seawater were relatively low with values of 0.21, 0.17, 0.11, and 0.22. A clear distinction in the composition and structure of the bacterial communities in the samples were observed (Figure 2.3). Despite the relatively high variability in the fungal community, beta-diversity analysis of the whole and ‘variable/core’ fungal communities showed significant overall differences between samples in 2014 and 2016 (Table 2.3 and Appendix Table S2.2). C. concentrica and seawater community compositions were also different between the two time points, in contrast to T. anhelans. However, temporal communities of sponge and seawater did not differ

23 between the two time points based on relative abundances. Beta-diversity analysis further showed that the fungal communities of all three sponge species were comparable to the surrounding seawater. The only difference was observed between the fungal community of T. anhelans and Scopalina sp., where the whole community analysis showed only differences in the community compositions, in contrast to the ‘variable/core’ community analysis, where significant differences were seen both in terms of the structure and composition (Table 2.3 and Appendix Table S2.2).

Table 2.5 PERMANOVA analysis (based on relative abundance and presence-absence values) of “variable/core” fungal communities (normalised) at an OTU-level of sponges and seawater. Significant P-values ≤ 0.05 are highlighted in bold.

Temporal samples P-value base on P-value based on relative abundance presence-absence 2014 vs. 2016 0.023 0.001 C. concentrica 2014 vs. 2016 0.1 0.001 Scopalina sp. 2014 vs. 2016 NA NA T. anhelans 2014 vs. 2016 0.40 0.40 Seawater 2014 vs. 2016 0.1 0.008 Sample type C. concentrica vs. Scopalina sp. 0.4 0.386 C. concentrica vs. T. anhelans 0.24 0.154 C. concentrica vs. seawater 0.54 0.632 Scopalina sp. vs. T. anhelans 0.048 0.048 Scopalina sp. vs. seawater 0.5 0.338 T. anhelans vs. seawater 0.143 0.139

A total of 30 out of 32 OTUs observed in sponges were also present in the seawater samples, however 12 of these OTUs were enriched in sponge (based on mean relative abundances) (Figure 2.2 and Appendix Table S2.4). Majority of enriched OTUs were found in more than one sponge species. For example, OTU_6, OTU_5, OTU_10, OTU_7, OTU_8, OTU_11, OTU_12 and OTU_13 were found in all three sponges, OTU_63 and

24 OTU_45 were observed in T. anhelans and Scopalina sp., and OTU_14 were observed in C. concentrica and Scopalina sp. (Figure 2.2). OTU_41 (Alatospora sp.) was only observed in C. concentrica) and OTU_112 (belonging to the class Agaricomycetes) was unique to Scopalina sp.. Both OTUs were highly variable in abundance and sometimes absent in one or more replicate samples. Fungal communities in sponges were generally dominated by a few OTUs. The top seven, eight, and four most abundant OTUs made up ∼90% of the total relative abundance of C. concentrica, Scopalina sp. and T. anhelans, respectively. The most abundant OTU observed across all samples was OTU_6 (Cladosporium sp.), which was most abundantly found in T. anhelans with a relative abundance range of 0–99% across six individual sponge samples. Only one core OTU was found in the sponge T. anhelans (OTU_63) identified as belonging to the class Leotiomycetes.

Figure 2.2 Heatmap of “variable/core” sponge fungal communities, with OTUs only found in seawater removed. Values are fourth root transformation of the relative abundance values. Columns are clustered based on Bray-Curtis dissimilarity using hierarchical clustering with the ‘average’ method (scale depicts percentage of dissimilarities). OTUs in teal indicate sponge enriched OTUs (measured by relative abundances) compared to seawater and OTUs in maroon indicate OTUs present in sponges which were absent in seawater. Samples are indicated by the colour purple: C. concentrica, orange: Scopalina sp., red: T. anhelans and blue: seawater. Temporal samples are indicated by the colour pink: 2014 and green: 2016. Numbers 1, 2 and 3 refer to individual replicates of each sample type.

25

Figure 2.3 Heatmap of the bacterial community composition based on 16S rRNA gene sequences for the sponge samples collected in 2014. OTUs were clustered at 97% similarity and the lowest taxonomy classification are given. Columns are clustered based on Bray–Curtis dissimilarity using hierarchical clustering with the ‘average’ method (scale depicts percentage of dissimilarities). Samples are indicated in purple: C. concentrica, orange: Scopalina sp., blue: seawater, and red: T. anhelans. Numbers 1, 2, and 3 indicate sample replicates.

2.5 Discussion

2.5.1 Assessment of fungal diversity in sponges Sponge-associated fungal communities have been studied through culture-dependent (Holler et al., 2000, Wang et al., 2008, Proksch et al., 2008, Li and Wang, 2009, Liu et al., 2010, Wiese et al., 2011, Yu et al., 2013, Henríquez et al., 2014) and culture- independent methods (Gao et al., 2008, He et al., 2014, Wang et al., 2014, Passarini et

26 al., 2015, Rodríguez-Marconi et al., 2015, Naim et al., 2017). However, no study so far, has combined both approaches to provide insights into the extent to which the two community assessments capture sponge-associated diversity.

In this study, 42 fungal isolates were classified into 8 different OTUs (Penicillium spp. (3), Togniniaceae, Acostalagmus sp., Trichoderma sp., Cladosporium sp., and Aspergillus sp.). Three isolates of the genus Penicillium (Eurotiales) were also found in the ITS amplicon sequencing and one isolate (OTU_119) was cultivated from all sample types. Penicillium can be considered as a ‘generalist’ (or core organism) and is an ubiquitous fungal genus commonly cultured from various environments, including sponges from around the world (Holler et al., 2000, Pivkin et al., 2006, Gao et al., 2008, Proksch et al., 2008, Wang et al., 2008, Li and Wang, 2009, Paz et al., 2010, Liu et al., 2010, Passarini et al., 2013, Passarini et al., 2015). Fungi from the class Togniniaceae (order Diaporthales) are likely of terrestrial origins as a common genus (Togninia) within is often associated with wilt and disease of woody plants (Crous et al., 1996, Rossman et al., 2007). Acrostalagmus (order Hypocreales) contains pathogenic fungi against animals, plants and humans (Kubicek et al., 2008). Some species of this genus have been isolated from deep-sea sediments and have been reported to produce anti- tumour (Wang et al., 2012) and antifungal compounds (Ellestad et al., 1969, Sato and Kakisawa, 1976). Togniniaceae and Acrostalagmus sp. so far have not been reported to be cultured from marine sponges, while Trichoderma sp., Aspergillus sp. and Cladosporium sp. have previously been cultivated from various marine sponges (Holler et al., 2000, Proksch et al., 2008, Wang et al., 2008, Paz et al., 2010, Passarini et al., 2013).

ITS-amplicon based community profiling captured more fungal diversity (26 OTUs) compared to cultivation (eight OTUs), which is consistent with previous studies examining fungal communities in marine sediments, decaying leaves and soil (Borneman and Hartin, 2000, Nikolcheva et al., 2003, Jebaraj et al., 2010, Singh et al., 2012). However, similar to the studies of Jabaraj et al. (2010) and Singh et al. (2012), the cultivation method yielded distinct isolates (five) that were not found in the molecular analysis. Three of the unique isolates were cultivated from T. anhelans (OTU_266, OTU_268 and OTU_269), OTU_265 was cultivated from Scopalina sp. and

27 OTU_ 264, which was cultivated from C. concentrica, Scopalina sp. and seawater. This occurrence could be explained by the overall low number of fungal reads obtained for T. anhelans and Scopalina samples in 2014, when the cultivation was also performed (Table 2.2). The low number of fungal reads due to the amplification of non-targeted DNA (e.g. sponge DNA) was a challenged reported in previous fungal diversity studies (He et al., 2014, Naim et al., 2017, Gao et al., 2008). Another explanation for the discrepancy between the two methods is that the isolates could belong to the rare biosphere, which is supported by the fact that three of the isolates were unique (i.e. only cultivated once in all biological and technical replicates) (Appendix Table S2.1). Similarly, the three Penicillium spp., which were found by both methods, were only found in low relative abundances in the ITS-amplicon sequencing data. This indicates that fungal cultivation, even under the various conditions tried here, does not capture abundant community members in sponges, but rather those that are relatively rare. This situation is analogous to what has been seen in many studies examining bacterial communities in sponges (Webster and Hill, 2001, Hentschel et al., 2001, Li et al., 2007, Muscholl-Silberhorn et al., 2008, Esteves et al., 2016). Our results indicate that ITS amplicon community profiling likely provides a more realistic assessment of fungal diversity in sponges and thus should be seen as the “gold standard” for community assessment, similar to what has been proposed for fungal diversity research in other environments (Pang and Mitchell, 2005, Jeewon and Hyde, 2007).

2.5.2 Host specificity of sponge-associated fungi Li and Wang (2009) have previously classified fungi found in sponges into three groups: ‘sponge-generalists’ (i.e. found in all sponge species), ‘sponge-associates’ (i.e. found in more than one sponge species), and ‘sponge-specialists’ (i.e. found in only one sponge species). This classification is very limited and dependent on a) the number of sponge species studied, b) the methods used (culture-dependent or -independent approach (see above), and c) experimental design (use (or lack) of biological replicates or seawater reference communities). These limitations have been highlighted by Yu et al. (2013) in their summary of sponge-associated fungi reported since 1996. The summary illustrates differences in classification of the same fungal genus in different studies. For example, the fungal genera Fusarium and Trichoderma were classified as ‘sponge

28 generalists’ according to Menezasa et al. (2010) but were classified as ‘sponge- associates’ by Hӧller et al. (2000) and Li and Wang (2009) and were completely absent in various sponges investigated by Gao et al. (2008) and Pivkin et al. (2006). Therefore, one should be careful when using these terminologies for host-specificity of fungal genera. Another challenge in determining the specificity of sponge-associated fungi is the filter-feeding capacity of sponges that brings large amounts of seawater along with all its constituents into the sponge tissue. Studies that are based on the once-off, presence-absence measurement of isolates (Li and Wang, 2009, Paz et al., 2010, Liu et al., 2010, Thirunavukkarasu et al., 2012) or sequences (Gao et al., 2008, He et al., 2014, Rodríguez-Marconi et al., 2015, Zhang et al., 2016) to support claims of specificity, could therefore be misleading as fungi found inside the sponge might simply just be accidentally trapped, rather than having any meaningful interaction with the sponge. Indeed, studies on bacterial communities in sponges have found that ecologically important organisms are consistently present in the sponge and usually at high relative abundances (Wilkinson and Fay, 1979, Hentschel et al., 2002, Hentschel et al., 2003, Lee et al., 2011, Fan et al., 2012, Thomas et al., 2016). Therefore, we propose and recommend implementing a different guideline to the above classifications, where “occasional” and “variable/core” fungi can be defined by examining biological replicate samples over different time periods. “Variable/core” fungi that are found in combination with high relative abundances compared to the surrounding environment (i.e. seawater) should then be considered “sponge-enriched”. Implementing these guidelines, our study revealed that the “variable/core” fungal communities in sponges have low diversity (≤28 fungal OTUs were observed in each sponge species), high variability and that the majority of fungal OTUs were not specifically enriched in any of the sponge species investigated here. These findings are consistent with a recent study by Naim et. al. (2017). In contrast, the bacterial communities of the sponges studied here showed diverse and host-specific sponge enriched bacterial phyla, which was consistent over time and space (Fan et al., 2012, Esteves et al., 2016). Furthermore, bacterial communities had low variability and were distinct from the bacterial community in the surrounding seawater (Fan et al., 2012, Thomas et al., 2016). Indeed, we could reproduce this low variability and distinctive composition of bacterial communities (see Figure 2.3) using the same technical approaches and DNA extracts

29 used for the fungal analysis, indicating that the high variability in the fungal communities truly reflects fundamental differences in the ecology of these two microbial groups.

Fungal communities in the three sponges studied here appear to be largely influenced by the community of the surrounding seawater. This is broadly reflected in the higher fungal diversity observed in 2016 compared to 2014, in both the seawater and in all sponges. However, the degree of influence by the fungal community in seawater appear to differ for different sponge species. For example, fungal communities in T. anhelans did not significantly differ between the two time periods, in contrast to C. concentrica and seawater. Additionally, fungal communities of T. anhelans and Scopalina sp. were found to be significantly different, suggesting that the two sponge species may exert different selective pressures on their fungal community. Differences in the relative abundances of the same OTUs in the sponges further support the occurrence of preferential interactions of fungi with different sponge species. For example, OTU_10 was found in T. anhelans, Scopalina sp. and C. concentrica with mean relative abundance of 0.09% ± 0.21 (range of 0-0.43%), 1.8% ± 3.6 (range of 0- 9,2%) and 16.7% ± 38 (range of 0-94.7%), respectively.

2.5.3 Potential roles of sponge-associated fungi Seven out of the 12 “sponge-enriched” OTUs could not be classified to a genus level, which indicates that they are novel, yet-to-be-studied organisms or that their ITS sequences are not available in the current UNIT ITS database (Abarenkov et al., 2010) (Appendix Table S2.4). The seven OTUs were identified to the lowest taxonomic classification as: kingdom fungi (OTU_11), class Lecanoromycetes (OTU_45 and OTU_87), Leotiomycetes (OTU_63), Agaricomycetes (OTU_14) and within class Agaricomycetes orders Pleosporales (OTU_10) and Agaricales (OTU_12). The other five sponge-enriched OTUs were classified as Cladosporium sp. (OTU_6), Fusarium sp. (OTU_8), Aureobasidium sp. (OTU_5), Curvularia sp. (OTU_7), and Beauveria sp. (OTU_13) (Figure 2.2). OTU_112 and OTU_41 were found exclusively in Scopalina sp. and C. concentrica, respectively. OTU_112 was classified to belong to class Agaricomycetes, which contains saprotrophs, pathogens and mutualists (Hibbett et al., 2014). Agaricomycetes were found frequently in the sponge samples studied here, in

30 fact eight out of 10 Basidiomycota fungi observed belong to this class. Sponge- enriched order Agaricales and Polyporales (within class Agaricomycetes) have also been detected in other marine sponges (He et al., 2014, Naim et al., 2017) and corals (Amend et al., 2012). OTU_41 (Alatospora sp., order Leotiales) has not been previously reported in marine sponges. Alatospora spp. are commonly associated with decaying wood/leaves in fresh water streams (Sridhar and Kaveriappa, 1989, Hosoya and Tanaka, 2007, Das et al., 2008). Together this shows that fungi with potential saprophytic properties are widespread in sponges and indicates that they may exploit the nutrient rich environment of the sponge host as parasites. Alternatively, they could potentially contribute to nutrient uptake via the breakdown of plant-base detritus or plankton filtered from the surrounding seawater (Zhang et al., 2006, Hyde et al., 2013, Hibbett et al., 2014).

The core OTU (OTU_63) detected in T. anhelans was classified to belong to the class Leotiomycetes. Interestingly, Leotiomycetes were prevalently cultivated (75% of total isolates) from Antarctic sponges, where a third of the fungal isolated were classified into the genus Geomyces and had strong antimicrobial activity. Another third of the isolates could not be identified to a genus level (Henríquez et al., 2014), indicating that this class has much unexplored genus-level diversity. More studies on Leotiomycetes in the future are essential to help elucidate its potential ecological function in sponges.

The class Lecanoromycetes (mostly enriched in T. anhelans) contains most of the lichen-forming fungal species (Miadlikowska et al., 2006). An OTU belonging to the order Telochistales (class Lecanoromycetes) was also found in sponge the Halichondrida panicea (an encrusting sponge) and was closely related to a lichen- forming fungi isolate (Naim et al., 2017). The lichen-forming genus Koralionastes has been reported to have a unique physical association with crustaceous sponge and was postulated to be nutritionally dependent on the sponges (Kohlmeyer and Volkmann- Kohlmeyer, 1992). Lichen have been reported to facilitate the destruction of rocks and provide the adjacent water layers with nutrients and trace elements that can benefit hydrobionts, including sponges (Kulikova et al., 2013). This raised the possibility that the presence of Lecanoromycetes in T. anhelans (relative abundance range of 0-0.33%) and in sponge H. panicea may arise from a close co-existence of crustose lichen and

31 encrusting sponges on rock fragments (Suturin et al., 2003). However, further detailed surveys and sample analysis of the surrounding habitats of encrusting sponges are needed to prove this.

OTU_6 (Cladosporium sp.) and OTU_8 (Fusarium sp.) belong to genera that are known to be saprotrophs and have been previously isolated from various marine sponges (Proksch et al., 2008, Wang et al., 2008, Li and Wang, 2009, Menezes et al., 2010, Paz et al., 2010, Liu et al., 2010, Zhou et al., 2011, Ding et al., 2011, Thirunavukkarasu et al., 2012) and were reported to produce bioactive compounds (Jadulco et al., 2002, Gesner et al., 2005, Wiese et al., 2011). For example, Cladosporium herbarum isolated from sponges Callyspongia aerizusa, Aplysina aerophoba and Callyspongia aerizusa produced various different antimicrobial compounds such as acetyl Sumiki’s acid (Gesner et al., 2005), pyrone derivatives and macrocyclic lactones (Jadulco et al., 2002). The marine-derived Cladosporium sp. F14 has also been found to produce compounds which inhibited larval settlement of the bryozoan Bugular neritina and the barnacle Balanus amphitrite (Qi et al., 2009). Fusarium spp. isolated from sponge Tethya aurantium was found to produce antibacterial and insecticidal compounds such as equistins and enniatine (Wiese et al., 2011). Cladosporium sp. and Fusarium sp. may similarly produce secondary metabolites in the sponges investigated here, which may potentially contribute to the sponge host defense.

OTU_5 (Aureobasidium sp., order Dothideales), OTU_7 (Curvularia sp., order Pleosporales) and OTU_13 (Beauveria sp., order Hypocreales) had closest sequence similarity to common terrestrial species ( Appendix Table S2.4), which can be facultative pathogens of plants, humans and insects (Rinaldi et al., 1987, Zalar et al., 2008, Liu et al., 2009, Rehner et al., 2011). All three genera have been previously isolated from various sponges around the globe (Holler et al., 2000, Wang et al., 2008, Li and Wang, 2009, Gao et al., 2008, Wiese et al., 2011, Henríquez et al., 2014, Passarini et al., 2015). Sponges have been reported to harbour various other fungal species related to terrestrial plant, human and animal pathogens, such as Aspergillus terreus and Cladosporium tenuissimum (Liu et al., 2010, Zhou et al., 2011). Additionally, the sponge species Spongia obsucra and Ircina strobilina have been reported to host the fungus Aspergillus sydowii, which is a causative agent of a disease

32 in the Caribbean sea fan corals (Gorgonia spp.) but had no notable affect to the sponge health (Ein-Gil et al., 2009). This suggests that sponges could be reservoirs of potential marine and terrestrial pathogens and may provide an environment for fungal propagule survival and dispersal.

2.6 Conclusion

Our study showed that the sponge samples analysed here contained phylogenetically diverse fungi (eight fungal classes were observed) that have low host-specificity and broadly reflected communities within seawater. The same or highly similar fungi species have been detected in sponges around the world, which suggests a prevalence of horizontal transmission where selectivity and enrichment of some fungi occur for those that can survive and/or exploit the sponge environment. Our current sparse knowledge about sponge-associated fungi indicates that fungal communities may perhaps not play as an important ecological role (beside the exceptional case reported of vertically transmitted endosymbiotic yeast (Maldonado et al., 2005b)) in the sponge holobiont compared to bacteria or archaea. However, the interaction between sponges and fungi provides another layer to the already complex sponge environment, which may drive the evolution of not only bacteria and archaea (Hentschel et al., 2002, Thomas et al., 2010, Fan et al., 2012, Thomas et al., 2016), but possibly also fungi. The ecology and function of sponge-associated fungi represents a frontier of microbial diversity research awaiting further studies.

Chapter Three

33 3 Taxonomic, functional and transcriptomic analysis of viral communities associated with marine sponges.

3.1 Co-authors and collaborators

This chapter is a collaboration between Mary T. H. D Nguyen, Bernd Wemheuer, Fan Lu, Cristina Diez-Vives, Nicole Webster and Torsten Thomas.

MN conceived and designed the experiments, performed the experiments, analysed the data, prepared figures and/or tables, and authored the chapter. BW assisted in the functional gene analysis. FL, NW and TT produced the sponge metagenome dataset. CD-V and TT produced the (meta)transcriptome dataset. TT conceived and designed the experiments, analysed the data and edited the chapter.

3.2 Introduction

Viruses, and predominately bacteriophages, are thought to be the most abundant biological entities in the ocean and are estimated to be present at approximately 107- 108 particles per millilitre of seawater, outnumbering their bacterial hosts of up to 15- fold (Fuhrman, 1999, Wommack and Colwell, 2000, Weinbauer, 2004). Viruses can infect all forms of life; animals, plants and microorganisms, including protists, bacteria and archaea. Marine viruses are considered to play an important role in a number of processes including: 1) global biogeochemical cycles through lysis of microorganisms (Middelboe and Lyck, 2002, Weinbauer et al., 2011), 2) microbial community structures and diversity through what is known as the “kill the winner”, where dominant taxa are killed through infection, allowing the growth of less-competitive

34 microorganisms (Angly et al., 2006, Winter et al., 2010), and 3) the evolution of microorganisms through virus-mediated genetic exchange (Riemann and Middelboe, 2002, Fuhrman and Schwalbach, 2003, Rodriguez-Brito et al., 2010, Breitbart, 2012, Thurber et al., 2017).

Metagenomics approaches have become the benchmark for research on viral communities, as they circumvent the need for cultivation and bypasses the drawback that there are no universal marker genes to assess viral diversity (Rohwer and Edwards, 2002). In addition, metagenomic analysis provides not only information on taxonomic diversity, but also functional genes within viral communities (Hurwitz and Sullivan, 2013). Viromes (i.e. the collections of viral genomes in a system) have been shown to contain genes encoding for extensive metabolic capacities, suggesting that they serve as a repository for storing and potentially transferring genes among different hosts (Hendrix et al., 2002, Dinsdale et al., 2008, Laffy et al., 2018). For example, viruses have been reported to carry ‘specialisation’ genes, including phosphate metabolism (Rohwer et al., 2000), cyanobacterial photosystems (Mann et al., 2003, Weynberg et al., 2017, Laffy et al., 2018), exotoxins and antibiotic resistance genes, which can confer benefits to the infected host (Młynarczyk et al., 1997, Davis and Waldor, 2002).

The influence of viruses on microbial evolution and community structure are particularly interesting for sponges, given the capacity of sponges to filter large amounts of seawater and the existence of dense microbial communities available for viral attack. Thus, sponges would be expected to contain high viral loads and may be a potential reservoir for new viruses. However, the area of sponge-associated viruses has received little attention, so far, only a few studies have reported the presence of viruses within sponges. Vacelet and Gallissian, (1978) noted an uncommon observation of virus-like particles in disturbed areas devoid of choanocytes chambers from one in >500 specimens of the sponge Verongia cavernicola and Claverie et al. (2009) upon reviewing this work predicted that sponge phagocytes were undergoing infection by a giant virus, possibly related to the Mimiviridae.

Metagenomic studies have revealed indirect evidence for viruses in which contained high abundances of restriction-modification (R-M) systems, clustered regularly 35 interspaced short palindromic repeats (CRISPRs) and CRISPR-associated (Cas) proteins (Thomas et al., 2010, Fan et al., 2012, Horn et al., 2016, Karimi et al., 2017). R-M systems are involved in recognising and tagging invading foreign DNA (e.g. phage genomes), which can then be cleaved by restriction endonucleases (Vasu and Nagaraja, 2013). CRISPRs are sites in the bacterial genome that provide resistance against the integration of extrachromosomal DNA (Barrangou et al., 2007). CRISPRs can acquire short DNA fragments from phages and incorporate them into an array of spacer sequences. Upon reinfection by the same phage, these spacer sequences can then get transcribed and hybridized to the phage DNA, which then get degraded by Cas proteins (Barrangou et al., 2007). High abundances of these defence systems suggest that microbial communities in sponges may be exposed to high viral loads (Thomas et al. 2010). The presence of mechanisms to prevent phage attack and control excessive genetic transfer may further contribute to the stability of microbial communities of sponges over time and space, despite the constant influx of viruses (Fan et al., 2012, Thomas et al., 2016).

Furthermore, Karimi et al. (2017) recently observed a negative correlation between CRISPR/Cas proteins as well as restriction endonucleases and viral abundances in the microbial community of Spongia officinalis. Similarly, Fan et al. (2012) previously observed an absence of CRISPR/Cas proteins accompanied with high abundances of cyanophages in the sponge Stylissa sp. 445. Interestingly, Stylissa sp. 445 also contained high abundances of cyanobacteria (Fan et al., 2012), which suggests that the cyanophages may have a lysogenic relationship with their host (Paul, 2008). Lysogeny involves the integration of the phage genome into the host’s chromosome, which can be passed on to daughter cells and can be “activated” to cause host lysis in response to environmental or physiological cues (Paul, 2008). Evidence suggests that lysogeny is a highly sophisticated relationship as a result of coevolution of phage and host genomes (Chen et al., 2005). However, very little is known about factors that trigger the conversion from a lysogenic to a lytic state (Paul, 2008, Thurber et al., 2017). The advantage of lysogeny for the phage includes survival in times of low host abundance as well as protection from proteolytic digestion and UV inactivation. Lysogeny can also confer benefits to the host, such as immunity to infection by other related viruses

36 (Waterbury and Valois, 1993) and can also boost host metabolism and survival through expression of auxiliary metabolic genes or encode for genes (‘fitness factors’) that enhance the fitness of the host (Brussow et al., 2004).

A recent study by Laffy et al. (2018) directly investigated viromes of corals and sponges using shotgun sequencing of amplified DNA derived from size and density-fractionated viruses. This work found diverse viral assemblages that exhibited significant host species specificity (Laffy et al., 2018). Functional gene analyses revealed that certain genes were conserved across all sample types (i.e. coral, sponge and seawater), such as genes associated with viral structure (capsid head and tail formation) and infection mechanisms (DNA replication), while genes involved in herbicide resistance and viral pathogenesis pathways were differentially enriched in host environments (Laffy et al., 2018). The study also showed quite variable relative abundances of eukaryotic viruses, such as Mimiviridae and Phycodnaviridae, among individuals of the same sponge species, while bacteriophages (e.g. Myoviridae, Podoviridae and Siphoviridae) were much more consistent in their relative abundance.

This recent study focussed on the free fraction of DNA viruses and it is currently unknown what viral diversity is associated with the microbial cell fraction and what part of viral assemblages are active (in the sense of showing gene expression). These aspects are important to further understand the role of viruses in the sponge holobiont. This chapter therefore aims to investigate the abundance, diversity and potential ecological function of viral assemblages associated with the microbial cells of three tropical sponges (Cymbastela coralliophila, Rhopaloeides odorabile and Stylissa sp. 445) and three temperate sponges (Cymbastela concentrica, Scopalina sp., and Tedania anhelans) as well as seawater. These tropical and temperate sponges were selected, because they are abundant in the Great Barrier Reef and Sydney waters, respectively, as well as to cover a wide selection of sponge morphologies, incorporating taxonomically diverse species and to encompass a broad geographical range. The bacterial communities of these sponges have been previously well studied using metagenomic approaches Scopalina sp. and T. anhelans contained a high relative abundances of bacteria belonging to the family Nitrosomonadaceae, while C. concentrica had high relative abundances of Phyllobacteriaceae and R. odorabile

37 contained high relative abundances of Poribacteria, Acidobacteria and Gemmatimonadetes (Fan et al., 2012). Of interest are Stylissa sp. 455 and C. coralliophila, which were reported to contain high relative abundances of Synechococcus and cyanobacteria, respectively (Fan et al., 2012). Because of the sponges’ filter feeding rates, metagenomic data will capture not only sponge- associated viruses, but also environmental viruses. To gather insights into potentially more ecologically relevant viruses, the expression of “active” (transcribed) viruses in C. concentrica, Scopalina sp., and T. anhelans sponges was also investigated using meta- transcriptomic data.

3.3 Materials and Methods

3.3.1 Metagenomic datasets of microbial cells associated with sponges The presence of viruses associated with microbial cells was investigated in metagenomic data of six sponges (Cymbastela coralliophila, Rhopaloeides odorabile, Stylissa sp. 445, Cymbastela concentrica, Scopalina sp., and Tedania anhelans) and seawater samples from a previous study by Fan et al. (2012). Briefly, in that study, tropical and temperate sponge species were collected in Palm Island, Great Barrier Reef and Bare Island, Sydney, respectively. on the multiple expeditions from 29.07.2009 to 23.09.2009 and seawater samples were collected on the 18.10.2006 in Bare Island, Sydney. Sponge samples were stored at room temperature in natural seawater and transported directly to the laboratory for processing. Homogenised sponge tissue and seawater was size-fractionated to enrich for microbial cells. This involved a series of low-speed centrifugation and filtration steps to 1) remove as many eukaryotic cells and organelles as possible (checked by microscopy and 16S/ 18S rRNA gene semi-quantitative PCR), and 2) no preferential removal of cellular microorganisms (checked by microscopy and community fingerprinting). DNA was extracted from the microbial cell fraction and shotgun sequenced with the Roche 454 Titanium pyrosequencing platform. Dereplicated reads of each sample were assembled using the GS De Novo Assembler with “genomic” default settings (for more details see Fan et al. (2012)). Taxonomic and functional analyses of the bacterial and archaeal

38 communities showed highly consistent profiles between individuals of the same sponge species (Fan et al. 2012).

3.3.2 Transcriptome and metatranscriptome Viral gene expression and the presence of RNA viruses was investigated for the transcriptome and metatranscriptome dataset of the sponges C. concentrica, Scopalina sp. and T. anhelans from a recent study by Diez-Vives et al. (2017). Total RNA extracted from snap frozen sponge samples were separated into eukaryotic RNA and prokaryotic RNA using the poly(A) RNA selection step. It is expected that the majority of viral genes would be found in the metatranscriptome, especially those from bacteriophages. Some eukaryotic viruses have been reported to have poly(A) tails (Poon et al., 1999) and thus would be in the transcriptome. Transcriptome libraries were generated from total RNA of three pooled sponge individuals using a poly(A) RNA selection step to enrich for poly-adenylated (eukaryotic) mRNA fragments. Meta-transcriptomic libraries were produced by multiple rounds of removal of the poly-adenylated mRNA using the

Dynabeads Oligo(dT)25 (ThermoFisher Scientific, Melbourne, Australia) followed by rRNA depletion using subtractive hybridization with bacteria-specific probes attached to magnetic beads (Ribo-Zero Bacteria Kit, Epicentre, Madison, WI, USA). This library should therefore be enriched in non-eukaryotic mRNA. RNA for the two types of libraries were further processed with the TruSeq RNA Sample Prep Kit protocol and sequenced on the Illumina HiSeq 2000 platform with 100 bp paired-end reads at the Ramaciotti Centre for Genomics (University of New South Wales, NSW, Australia).

Transcriptomic and meta-transcriptomic reads were de novo assembled with the TRINITY software v2.0.6 (Grabherr et al., 2011) with default settings. To analyse the eukaryotic transcriptome, CD-HIT-EST v4.6.4 (Li and Godzik, 2006) was used to remove of identical sequences in the assembly (parameter “c” = 1) based on a word size of 8. Transcripts were searched with BlastX against the non-redundant NCBI database and transcripts with hit E-values < 1.0E-5 were taxonomically classified with MEGAN v5.11.3 (Huson et al., 2007) using the lowest common ancestor algorithm (LCA) with settings min score = 50, max expected = 1.0E-5, top per cent = 10 and LCA per cent = 100. Transcripts classified as bacterial or archaeal were removed from the assembly before further analysis (with E-value < 1.0E-3). The microbiome mRNA reads from

39 treatment RZ (RZ-reads) were mapped against the eukaryotic transcriptome assembly using BOWTIE2 v2.2.4 (Langmead and Salzberg, 2012). Any read mapping the eukaryotic transcriptome with a minimum default score of -60 was removed. No viral sequences were analysed or removed from the transcriptome and metatranscriptome

(for more details see Diez-Vives et al. (2017)).

3.3.3 Identification and normalisation of viral contigs The VirSorter program (Roux et al., 2015) was used to identify viral sequence from the contigs of the sponge microbial cell-associated metagenome the sponge transcriptome and metatranscriptome. VirSorter uses an inbuilt ‘Viromes’ reference database, in which 826 proteins clusters and single genes were identified as “viral hallmark genes”. Briefly, for each contig provided, VirSorter first detects circular sequences (Roux et al., 2014), then predicts genes with MetaGeneAnnotator (Noguchi et al., 2008) and selects all contigs with more than two genes predicted. Predicted protein sequences are then compared to the PFAM v27 database and to the viral database (Viromes) with HMMsearch (Eddy, 2011) and BLASTp (Altschul et al., 1997). Each gene is assigned to the most significant hit based on an alignment score with a minimum score of 40 and maximum E-value of 10-5 for the HMMsearch, and a minimum score of 50 and maximum E-value of 10-3 for the BLASTp search. Contigs are then characterised through a computation of multiple criteria as follows: 1) presence of viral hallmark genes (Koonin et al., 2006, Roux et al., 2014), 2) enrichment in viral-like genes 3) depletion in PFAM affiliated genes, 4) enrichment in uncharacterized genes (i.e. predicted genes with no hits either in PFAM or the viral reference database), 5) enrichment in short genes (genes with a size within the 10% of the shortest genes of the genome or contig), and 6) depletion in strand switching (i.e. change of coding strand between two consecutive genes) (Roux et al., 2015). Contigs are then classified into three categories (i) category 1 (“most confident” viral predictions) for contigs fulfilling criteria 1 and 2; (ii) category 2 (“likely” viral predictions) for contigs fulfilling either criteria 1 or 2 and at least one of the other criteria (3, 4, 5 or 6); and (iii) category 3 (“possible” viral predictions) for contigs not fulfilling criteria 1 or 2, but at least two of the other criteria with at least one significance score greater than 4 (for more details see Roux et al. (2015)). All contigs from the metagenome, transcriptome

40 and metatranscriptome that met category 1 and 2 were used for further taxonomic analysis as these were considered as most confident and most likely to be of viral origins. Viral (meta)transcriptome contigs were normalised using transcripts per million (TPM) counts (Diez-Vives et al. 2017). Viral contigs from the cellular metagenomes were normalised to the contig coverage and the estimated number of prokaryotic genomes (Fan et al. 2012). The number of prokaryotic genomes per sample was inferred using the average abundance of 82 single-copy genes, which were present in ≥ 90% of all complete NCBI archaeal and bacterial RefSeq genomes (archaea = 261, bacteria = 7315; July 2017) and, when present, being single-copy in ≥95% of all genomes. The PFAM and TIGRFAM hmm profiles (version 31.0 and 14.0, respectively) were used for protein identification in the genomes using HMMsearch v3.1b2. The number of prokaryotic genomes per sample for each sponge individual replicate are indicated in Table 3.1.

3.3.4 Taxonomic and phylogenetic classification of viral contigs Taxonomic classifications of viral contigs were conducted using the MetaVir tool (Roux et al., 2014). Open reading frames (ORFs) were first predicted for each contig through MetaGeneAnnotator (Noguchi et al., 2008) and all translated ORFs were compared to the RefseqVirus protein database from NCBI using BLASTp (Altschul et al., 1990) with an E-value threshold of 10-3, and the PFAM database (version 2.0) (Punta et al., 2011) using HMMScan with a score threshold of 30 (Eddy, 2011). Taxonomic classification was based on the lowest common ancestor (LCA) affiliation of each contig. The LCA affiliation considers the multiple hits on a single contig (up to five, if available) and the affiliation is made at the highest common taxonomic level of the best BLAST hits from the selected genes against the RefSeqVirus database.

Heatmaps, community diversity and statistical analysis were conducted in the R statistical program language (R core Team 2014) using the vegan package (Oksanen et al., 2013). Heatmaps were transformed with cube root or fourth root transformation to allow for better visualisation and comparison between samples.

41 3.3.5 Functional annotation of viral contigs For functional annotation of viral sequences, ORFs were predicted using Prodigal v.2.6.2 (Hyatt et al., 2010). Predicted protein sequences were functionally classified using Diamond searches (Buchfink et al., 2015) against the COG (version 2014) (Galperin et al., 2014) and KEGG (October 2017) databases (Kanehisa et al., 2002) with an E value of <10-10. In addition, predicted proteins were compared to the PFAM v31 (Finn et al., 2015) and TIGRFAM v15 (Haft et al., 2003) databases using HMMsearch with an E-value of <10-10. Functional profiles for each sample were calculated by summing the abundance of each functional categories. Proteins with more than one assigned functional category or domain contributed to each function/functional domain equally. The abundance of KEGG and COG pathways in each sample was calculated from the abundance of all affiliated specific functional categories. The abundance of functional categories that belong to more than one pathway was equally divided between the pathways.

3.4 Results

3.4.1 Viral diversity associated with microbial cells in sponges The majority of detected viral contigs using the VirSorter program (Roux et al., 2015) fell into categories 1 or 2 and only contigs that fell into these two categories (predicted to be of “most confident” or “likely” viral sources), were used in the analysis. Viral contigs made up a small percentage of the total contigs ranging from 0-0.47% for different samples analysed here (Table 3.1). A total of 451 viral contigs were identified and of these 107 viral contigs could not be taxonomically classified to a species level using the lowest common ancestor (LCA) approach in the MetaVir program (Roux et al., 2014) (Figure 3.1, Appendix Figure S3.1 and S3.2). Metagenome of seawater contained an estimated average sum of 16 viral contigs per bacterial genome followed by the sponges Scopalina sp., C. coralliophila, Stylissa sp. 445, C. concentrica, T. anhelans and R. odorabile with an average sum of 3.6, 2.7, 2.5, 1.2, 1.1 and 0.68, respectively.

42 Viromes in all samples were dominated by Caudovirales. Myoviridae was most abundant in seawater followed by Siphoviridae and Podoviridae, and the relative abundance of the three viral families differed among each sponge species (see Figure 3.1 and Appendix Figure S3.1). Alpha diversity (measured by Shannon’s index) on the family level showed that seawater contained the highest viral diversity, followed by Scopalina sp., C. coralliophila, T. anhelans, R. odorabile, C. concentrica and Stylissa sp. 445 (Appendix Table S3.1 and S3.2). This was generally consistent with viral diversity observed on a species-level (Appendix Figure S3.2). Viral assemblages in sponges generally did not correlate with the profile and abundance of the bacterial communities at a species level conducted in the previous study by Fan et al. (2012). The exception was the high abundances of Synechococcus phages in the sponges C. coralliophila and Stylissa sp. 445, which is consistent with a relative high abundance of cyanobacteria in these sponges reported by Fan et al. (2012).

Table 3.1 Summary of the number of viral contigs found in each sample metagenome. A, B and C indicate sample replicates for sponges and 01, 02 and 08 indicate seawater replicates.

Sample No. of total contigs No. of viral contigs No. of prokaryotic % of viral contigs in genome per sample metagenome

Seawater 01 18797 88 53.00 0.468

43 Seawater 02 24581 109 63.78 0.443

Seawater 08 42686 11 62.21 0.026

C. coralliophila A 62543 3 39.94 0.005

C. coralliophila B 45264 46 45.82 0.102

C. coralliophila C 58846 17 8.04 0.029

R. odorabile A 76275 14 38.64 0.018

R. odorabile B 31373 9 15.54 0.029

R. odorabile C 34669 3 16.7 0.009

C. concentrica A 27990 6 32.65 0.021

C. concentrica B 26734 8 94.60 0.030

C. concentrica C 68642 13 73.42 0.019

Scopalina sp. A 25395 21 52.62 0.083

Scopalina sp. B 30982 21 21.62 0.068

Scopalina sp. C 28414 27 26.76 0.095

T. anhelans A 15972 11 69.84 0.069

T. anhelans B 15949 10 62.02 0.063

T. anhelans C 11789 7 29.04 0.059

Stylissa sp. 445 A 32276 25 7.8 0.077

Stylissa sp. 445 B 21243 2 0.55 0.009

Stylissa sp. 445 C 24377 0 1.16 0.0

44

Figure 3.1 Viral communities of sponges and seawater. Samples were clustered based on Bray-Curtis dissimilarities of taxonomic profiles (tree scale indicates dissimilarity percentages). Values are normalised to viral contigs per prokaryotic genome transformed with fourth root. Numbers indicate the number of viral species found within each taxon. Taxa highlighted in bold indicate unclassified taxa above species- level and corresponding numbers in brackets indicate the number of contigs. Sample types are indicated by colour in the figure legend. A, B and C indicate sample replicates for sponges and number 01, 02 and 08 indicate replicates for seawater samples.

Viral assemblages had high variability among individual replicates and between different host environments on both taxonomic families and species-level classification (Figure 3.1 and Appendix Figure S3.1 and Figure S3.3). Viral assemblages associated with microbial cells from the three tropical sponge species (C. coralliophila, R. odorabile, Stylissa sp. 445) did not differ to those of the temperate sponge species (C. concentrica, Scopalina sp., and T. anhelans) (Appendix Figure S3.3). Significant

45 differences were seen in the viral community composition across all sponge species at a family-level and species level (PERMANOVA, P=0.011 and P=0.001, respectively), however no significant differences were detected in pairwise comparison of host species (Appendix Table S3.2). Additionally, no significant differences were observed between environments (i.e. sponge vs seawater) at the family and species-level (PERMANOVA, P-value=0.11 and P-value=0.208, respectively). The overlap of viral species observed in sponge and seawater ranged between 11-60% (Appendix Table S3.3).

3.4.2 Viral communities in the metatranscriptome and transcriptome datasets The percentage of viral contigs in meta-transcriptomes of C. concentrica and Scopalina sp. were 0.003% and 0.002%, respectively, and no viral contigs were identified in the meta-transcriptome of T. anhelans. Similarly, low percentages of viral contigs were identified in the transcriptome of C. concentrica and T. anhelans (0.002% and 0.0005%, respectively) and no viral contigs were identified in transcriptomic data of Scopalina sp.. Additionally, no RNA viruses were identified in the (meta)transcriptome data, which may be due to the scarcity of RNA virus genomes in the “Virome” database used in the VirSorter program (Roux et al., 2015).

A total of 14 viral contigs were identified from the meta-transcriptome of the sponges C. concentrica (12) and Scopalina sp. (two) and 5 viral contigs were identified from the transcriptome of the sponges C. concentrica (four) and T. anhelans (one). In contrast to the compositional data from the microbial cell metagenomes (above), expression of viral contigs in the meta-transcriptomes were relatively consistent between individual replicates, showing strong clustering based on Bray-Curtis similarity on the lowest taxonomical level assigned (Figure 3.2).

C. concentrica showed a more diverse “active” viral consortia compared to Scopalina sp., which included bacteriophages belonging to the families Myoviridae and Siphoviridae (within order Caudovirales), and Phycodnaviridae (Figure 3.2). All viral species observed in the meta-transcriptome of C. concentrica were distinct from those found in the microbial cell metagenome (see above). Ostreoccoccus lucimarinus virus 7

46 (belonging to family Phycodnaviridae) were observed at high abundances with an average expression of greater than three-fold the average total TPM in the C. concentrica meta-transcriptome (Figure 3.2). Meta-transcriptomes of Scopalina sp. contained two eukaryotic viruses, with closest similarity to Pandoravirus dulcis and Ostreoccoccus lucimarinus virus 7, which were expressed approximately equal to the total average TPM. (Figure 3.2). Phycodnaviridae was also observed in the metagenome data of microbial cells in Scopalina sp..

Four contigs found in the C. concentrica transcriptome were assigned to the bacteriophages Cronobacter phage vB_CsaM_GAP32, Klebsiella phage K64-1, Bacillus phage G and Synechococcus phage S-CBS3. The contigs assigned to the Cronobacter phage vB_CsaM_GAP32 was expressed at high levels (approximately twice the average TPM in C. concentrica transcriptomes) and was also observed in the metatranscriptome and metagenome data of C. concentrica (see above), while the contigs assigned to the latter three viral species were not found in corresponding metatranscriptome or metagenome datasets. The T. anhelans transcriptome also contained a viral contig assigned to the family Phycodnaviridae, which was absent in the microbial cell metagenome of T. anhelans.

47

Figure 3.2 (Meta)transcriptome analysis of viral community in sponges. Viral assemblages were clustered using Bray-Curtis dissimilarities of taxonomic profiles (scales indicate dissimilarity percentages). 1, 2, and 3 indicate individual sponge replicates. For transcriptome data, the three replicates were pooled together. Values are natural logs of normalised data transcripts per million (TPM), coloured markers on the scale represent average TPM of all transcripts in the corresponding samples coded by colours in the legend, orange: Scopalina sp. meta- transcriptome, red: T. anhelans transcriptome, purple: C. concentrica meta-transcriptome and navy: C. concentrica transcriptome.

3.4.3 Functional gene analysis of viral contigs associated with microbial cell metagenomes Functional gene annotation showed that 16%, 20%, 21.4% and 13% of all predicted genes could be mapped to the KEGG, COG, PFAM and TIGRFAM database, respectively. Functional profiles showed high variability between individual replicates and host environments (similarly to the taxonomic analysis) (Figure 3.1, Figure 3.5, Figure 3.6, and Appendix Figure S3.4, Figure S3.5, Figure S3.6 and Figure S3.7). Functional categories of the viral assemblage were distinct from the whole (viral and prokaryotic) metagenomes (Figure 3.5). Generally, COG annotation associated with “mobilome: prophages, transposon” and “DNA replication, recombination and repair” were enriched in the viral metagenomes compared to the whole metagenomes (Figure 3.6). This is consistent with abundant proteins, such as “Mu-like prophage proteins”, “DNA/RNA polymerase” and “DNA primase and helicases” proteins detected in the PFAM, KEGG, COG and TIGRFAM annotations (Appendix Figure S3.4, Figure S3.5 and

48 Figure S3.6). Interestingly, genes involved in “energy production and conversion”, such as genes associated with photosystem II and ATP synthase, were only observed in viral metagenomes of Scopalina sp. (average of 5.1 copy/ prokaryotic genome) and Stylissa sp. 445 (average of 3.2 copy/ prokaryotic genome), which contained high abundances of Synechococcus phages (Appendix Figure S3.4). “pantothenate and CoA biosynthesis” pathways were enriched in viral metagenomes of C. concentrica (replicate A and B) and “purine and pyrimidine metabolism” pathways were enriched in Stylissa sp. 445 (replicate B). Several specific genes with potential auxiliary functions for bacterial or eukaryotic hosts were detected in the viral assemblages, including pathways for the biosynthesis of antibiotics, genes for aegerolysin and bacterial toxin-antitoxin systems (ydaT and HicA) (Figure 3.7 and Appendix Figure S3.4).

Figure 3.3 Functional gene analysis of sponges and seawater viral (pink) and whole (viral and prokaryotic) (brown) metagenomes. Functional enrichment based on COG annotations of assembled predicted genes normalised to proportion of gene copies per prokaryotic genome and transformed with cube root. Samples are clustered with Bray-Curtis dissimilarity using the ‘average’ method, tree scale is based on dissimilarity percentages. A, B and C indicate individual sponge replicates and 01, 02 and 08 indicate seawater replicates.

49

Figure 3.4 Genes associated with KEGG pathways of sponges and seawater viral (pink) and whole (viral and bacterial) (brown) metagenome. Values are normalised to proportion of gene copies per prokaryotic genome and transformed with fourth root. Samples are clustered with Bray-Curtis dissimilarity using the ‘average’ method, tree scale is based on dissimilarity percentages. A, B and C indicate individual sponge replicates and 01, 02 and 08 indicate seawater replicates.

Figure 3.5 Functional auxiliary proteins in virus associated with the microbial cell fraction. Proteins results are hits against the PFAM database (PF) and TIGRFAM database (TIGR). Values are normalised to copies per bacteria genomes with fourth root transformation. Columns are clustered using the Bray-Curtis dissimilarities using the ‘average method’ and scale indicates dissimilar percentages. A, B and C indicate sample replicates and 02 indicates seawater replicate sample.

50 3.4.4 Functional gene analysis of viral contigs found in (meta)transcriptomes Functional gene analysis based on COGs showed distinct metatranscriptomes profiles between Scopalina sp. and C. concentrica and similar to taxonomic analysis, annotation profiles were conserved between biological replicates (Figure 3.8, and Appendix Figure S3.8, Figure S3.9 and Figure S3.10). Generally, enrichment of genes associated with “posttranslational modification, protein turnover, chaperones” was observed in the viral meta-transcriptomes compared to the whole meta-transcriptome of both sponges (Figure 3.8). C. concentrica metatranscriptomes had an enrichment of genes associated with “amino acid transport and metabolism” and “transcription” and the viral metatranscriptomes of Scopalina sp. had genes associated with “replication, recombination and repair” enriched compared to the whole meta-transcriptomes. COG and KEGG profiles of the viral (meta)transcriptomes were distinct from the whole (viral and prokaryotic) (meta)transcriptomes (Figure 3.8 and Figure 3.9). No genes associated with any known KEGG pathways were observed for viral meta- transcriptomes of Scopalina sp. (Figure 3.9). Interestingly, genes associated with “thiamine metabolism”, “sulfur relay systems” and “antibiotic biosynthesis” pathways were enriched in the viral compared to the whole metatranscriptomes of C. concentrica (Figure 3.9).

51 Figure 3.6 Functional gene analysis of sponge viral (pink) and whole (viral and prokaryotic) (brown) (meta)transcriptomes. Functional enrichment based on COG annotations of assembled predicted genes were normalised to transcripts per million (TPM) and transformed with square root. Samples are clustered with Bray-Curtis dissimilarity using the ‘average’ method, tree scale is based on dissimilarity percentages. 1, 2 and 3 indicate sponge replicates for Scopalina sp. and C. concentrica metatranscriptomes. For transcriptomic data, the three replicate datasets were pooled together for T. anhelans and C. concentrica samples.

Figure 3.7 Genes associated with KEGG pathways of sponge viral (pink) and whole (viral and prokaryotic) (brown) (meta)transcriptomes. Genes associated with KEGG pathways were normalised to transcripts per million (TPM) and transformed with cube root. Samples are clustered with Bray-Curtis dissimilarity using the ‘average’ method, tree scale is based on dissimilarity percentages. 1, 2 and 3 indicate individual sponge replicates for Scopalina sp. and C. concentrica metatranscriptomes. For transcriptomic data, the three replicate datasets were pooled together for T. anhelans and C. concentrica samples.

52 3.5 Discussion

3.5.1 Viral assemblages associated with microbial cells in sponges Metagenome analysis was conducted on the fractionated microbial cell from six sponge species and seawater (Fan et al., 2012), and therefore the viral data obtained here are likely derived mostly from either virus attached to prokaryotic cells, virions present inside cells or prophages, rather than free viruses within the sponge environment. Viral assemblages in all hosts were dominated by bacteriophages with the majority being predicted as dsDNA viruses and less frequently ssDNA viruses (such as Inoviridae). The most abundant bacteriophage order was Caudovirales, which consists of three viral families, Myoviridae (viruses with contractile tails), Siphoviridae (viruses with long, non-contractile tails) and Podoviridae (viruses with short, non- contractile tails) (Maniloff and Ackermann, 1998). Caudovirales have also been reported to dominate the fraction of free viral particles in sponges and corals (Williamson et al., 2012, Laffy et al., 2016, Weynberg et al., 2017, Laffy et al., 2018) and in other environments, including seawater and soil (Clokie et al., 2011, Zablocki et al., 2014).

Viral assemblages associated with the microbial cell fraction of sponges showed high variability within and across sponge species, both in terms of community structure and composition (Appendix Figure S3.3). The viral communities of some sponge species analysed here overlapped with up to 60% (ranging from 11-60%) with viral species found in seawater (Appendix Table S3.4), indicating that a substantial fraction of the viruses found here may originate from the surrounding seawater. However, some sponge species contained high percentages of specific viral assemblages. For example, R. odorabile, C. concentrica and T. anhelans, which contained 75%, 68% and 67% unique viral species, respectively. However, PERMANOVA analysis of viral assemblage on a family and species-level did not show any significant pairwise differences between the sponge species and seawater (Appendix S3.2). This contrasts to the only other study by Laffy et al. (2018) on sponge-associated viromes, which showed high consistency between replicates and clear distinctions between the viromes of sponge species and seawater. One explanation for this discrepancy is that Laffy et al. (2018) investigated the metagenome of the free viral fraction, while our study focused on the

53 metagenome of the microbial cellular fraction. Given the scarcity of data currently available, it is clear that further work is required to fully understand the viral specificity in sponges.

Based on our sequencing data, seawater contained an estimation of 16 viral contigs per prokaryotic genome, which is consistent with a previous estimate of up to 15 viruses per bacterium in various ocean waters (Fuhrman, 1999, Wommack and Colwell, 2000, Weinbauer, 2004, Mokili et al., 2012). The sponges investigated here, contained much lower average values ranging of 4 - 0.7 viral contigs per prokaryotic genome. This indicates that the viruses from seawater that enter the sponge may not effectively infect microbial symbionts and therefore reproduce to a lesser extent. This would be consistent with the high abundances of restriction modification (R-M) systems, CRISPR/Cas proteins and genes associated with phage growth limitations found in previous studies of sponge metagenomes (Thomas et al., 2010, Fan et al., 2012, Horn et al., 2016, Karimi et al., 2017).

3.5.2 Diversity of active viral consortia in sponges In contrast to the variability of the cell-associated viral metagenomes, the viral metatranscriptome of sponges C. concentrica and Scopalina sp. showed consistent expression of viruses in the biological replicates (Figure 3.2). No viruses were found in the T. anhelans metatranscriptome, indicating that could be low or no expressions of active viruses. This is consistent with the relatively low number of viral species identified in the T. anhelans metagenome. Interestingly, active viruses in C. concentrica and Scopalina sp. were generally observed at high expression levels, equal to or above the total average TPM. For example, Phycodnaviridae and Pandoraviridae were observed in both sponge meta-transcriptomes at 2-3 folds more than the total average TPM in C. concentrica and comparable to the total average TPM in Scopalina sp..

Phycodnaviridae belong to the superfamily of Nucleocytoplasmic Large DNA viruses (NCLDV), which infect marine and freshwater algae (Koonin and Yutin, 2010) and were also reported to be prevalent in the free viral fraction of the marine sponges Amphimedon queenslandica and Iantella basta (Laffy et al., 2018). C. concentrica contains a high abundance of diatoms (Taylor et al., 2004), which could potentially be

54 hosts to Phycodnaviridae viruses. Sponges could also filter and concentrate phototrophic organisms that are host to Phycodnaviridae from the seawater (Laffy et al., 2018). This later scenario would be consistent with the observation made here that some Phycodnaviridae sequences are related to Ostreoccoccus lucimarinus virus 7, which infects Ostreoccoccus lucimarinus, a microalga that is abundant in coastal and mesotrophic systems in the Atlantic and Pacific oceans (Derelle et al., 2015).

Pandoraviruses have been reported to infect amoeba, which have been observed to exist in sponges (He et al., 2014), but abundances of amoeba in the sponges studied here have not been investigated. Another potential host for amoeba-infecting viruses (such as Pandoraviruses) may be sponge amoebocytes (phagocytic sponge cells) which have many physiological and structural similarities to free-living amoeba (Reade, 1968, Hahn-Keser and Stockem, 1997). The presence of amoeba-infecting viruses seen here through sequence-based analysis is consistent with microscopic observation, which shows “parasitic infections” of sponge amoebocytes that resembled micrographs of Acanthamoeba infected with Mimivirus (Claverie et al., 2009).

RNA sequences for bacterial viruses were only observed in C. concentrica and were related to Cronobacter phage vB_CsaM_GAP32, Burkholderia phage ST79, Aeromonas phage 31, Enterobacteria phage (phi92 and mEp237) and Shigella phage SfII (Figure 3.2). Different species of Burkholderia and Enterobacteria phages were also observed in the C. concentrica metagenomic data, while Cronobacter phages were observed in both C. concentrica and seawater metagenomes and Aeromonas phages were observed in the seawater metagenome (Figure 3.1 and Appendix Figure S3.2). Interestingly, these bacteriophages are known to infect pathogenic bacteria. Burkholderia phage ST97 was reported to infect Burkholderia pseudomallie, a gram- negative pathogen found in soil that causes a severe septic infectious disease called meliodiosis (Khakhum et al., 2016). Enterobacteria and Shigella phages infect bacteria belonging to the Enterobacteriaceae family (Casjens, 2008), which are common intestinal bacteria with many pathogenic strains causing infection in animals and humans (Paterson, 2006). Cronobacter phage vB_CsaM_GAP32 infects Cronobacter sakazaki, which is an environmental pathogen that cause opportunistic infections in immunocompromised individuals (Abbasifar et al., 2014). Aeromonas phage 31

55 commonly infects Aeromonas salmonicida, which is a common pathogen of fish belonging to the salmonid family (Comeau et al., 2007). While sequences related to these bacteriophages are clearly expressed and hence suggest “active” viruses, sequences corresponding to their bacterial hosts have so far not been found through extensive metagenomic and 16S rRNA amplicon sequencing (Fan et al., 2012, Esteves et al., 2016). One potential explanation for this observation is that these viruses are able to infect bacteria other than their reported hosts in the sponge microbial community, that is, these viruses may have a large host range yet to be elucidated. Alternatively, these “active” bacterial viruses and their bacterial hosts may be derived from human/terrestrial contaminants that are accidentally filtered into the sponge. Within the sponge, the pathogenic hosts may become abundant enough to support viral infection and lysis resulting in a high abundance of “active” viruses and low (or not detectable) abundances of the reported corresponding bacterial hosts. However further work is required to explore if and how sponges act as a reservoir for pathogenic bacteria and their viruses.

3.5.3 Virus-host dynamics within the sponge environment Viral abundance is thought to be generally positively correlated with microbial abundances (Wommack and Colwell, 2000, Breitbart and Rohwer, 2005). On a phylum level, Proteobacteria, Bacteroidetes, and Cyanobacteria hosts were previously found to be in high abundances in the samples analysed here (Fan et al., 2012), this positively correlated to phages that infect these bacterial phyla found in this study. Particularly, high abundances of Synechococcus phages seen here corresponded with high abundances of Synechococcus bacteria observed previously in Stylissa sp. 445 (Fan et al., 2012). This was also consistent with high abundances of the T4-like phage capsid assembly protein G20 previously observed in Stylissa sp. 445 (Fan et al., 2012). Phylogenetic analysis of cyanophages contigs from Stylissa sp. 445 were closely related to the Synechococcus phage S-SM2 (T4-like phage) (Sullivan et al., 2010) and Synechococcus phage Bellamy (Hua et al., 2017) isolated from free-living cyanobacteria. The cyanophage contig from Scopalina sp. were divergent to those found in Stylissa sp. 445 and was more closely related to Synechococcus phage S-SSM7, a T4-like virus isolated from the Atlantic Ocean (Sullivan et al., 2010). This indicates

56 that Synechococcus phages found in Stylissa sp. and Scopalina sp. may have been filtered into the sponge from the surrounding seawater. However, their high abundance found in only these two sponges (and not in the other four sponge species) may also suggest that they perhaps interact with specific sponge-associated Synechococcus. Positive correlation of abundances of viruses and their hosts also indicates that the virus-host dynamic may be lysogenic (Jiang and Paul, 1998, McDaniel et al., 2002, Brussow et al., 2004). Evidence for lysogeny in the current study is given by the prevalence of viruses related to Myoviridae and Siphoviridae, which contains many lysogenic viruses and prophages (Maniloff and Ackermann, 1998, King et al., 2011) (Appendix Figure S3.1). Additionally, high abundances of lysogenic Mu-like prophage proteins were observed (Appendix Figure S3.7). Furthermore, high abundances of viral latency genes were observed in coral and sponge viromes, providing further evidence of lysogeny (Weynberg et al., 2017, Laffy et al., 2018).

The majority of viruses observed here however, did not have an apparent correlation with the abundances of putative host, which further indicates that the viruses may originate from the surrounding environment (as discussed earlier). This was also observed in other metagenome studies that investigated both viral and bacterial communities in corals and soil (Zablocki et al., 2014, Adriaenssens et al., 2015, Weynberg et al., 2017). The disparity in the correlation of viruses and associated hosts in the sponge environment could also be due to viral misclassification caused by the bias towards cultivated viruses in the reference database (Hendrix et al., 2000, Hendrix et al., 2002) or by a potentially uncharacterised wide host range of the classified viruses (Hendrix et al., 2002, Sano et al., 2004).

3.5.4 Functional contribution of the virome to the sponge holobiont In this chapter, approximately >80% of ORFs could not be assigned to any known functions using various annotation systems, which is consistent with the high percentage of “unknown” proteins (generally >70%) commonly observed in other viral metagenome studies (Williamson et al., 2012, Hurwitz and Sullivan, 2013, Zablocki et al., 2014, Adriaenssens et al., 2015, Weynberg et al., 2017). Generally, genes associated with “replication, recombination and repair”, “protein metabolism” and “carbohydrate metabolism” were abundant in all viral (meta)transcriptomes. High 57 abundances of genes associated with these three functions and viral structural proteins are consistent with various virome studies from different environments, including sponges (Laffy et al., 2018), corals (Weynberg et al., 2017) seawater (Williamson et al., 2012), soil (Zablocki et al., 2014, Adriaenssens et al., 2015) and animals, such as fish, mosquito and terrestrial animals (Dinsdale et al., 2008). This indicates the viromes from various environments share common features that reflects the focus of the viruses on energy production for DNA replication and virion production (Modrow et al., 2013).

However, several specific genes with potential auxiliary functions for bacterial or eukaryotic hosts were detected in the viral assemblages, such as genes associated with the biosynthesis of antibiotics, photosystem II (PSII), aegerolysin and bacterial toxin- antitoxin systems (Figure 3.9 and Appendix Figure S3.4). An enrichment in expressions of genes associated with thiamine metabolism was also observed in the C. concentrica metatranscriptomes (Figure 3.8).

Genes associated with the biosynthesis of antibiotics were observed in both viral metagenomes and metatranscriptomes of C. concentrica and were assigned to the synthesis of streptomycin and the vancomycin group of antibiotics. Cationic antimicrobial peptide (CAMP) resistance pathway were also identified in the viral metatranscriptome of C. concentrica. Antibiotic resistance genes (ARGs) have previously been reported in the viromes of aquaculture facilities (Colombo et al., 2016), where transcriptional regulators (associated with modulating antibiotic efflux) as well as glycopeptide, tetracycline and β-lactam resistance genes were observed. Here, CAMP resistant genes were observed in both the viral and the whole metagenomes (and enriched in the viral metatranscriptomes), suggesting that gene mobilisation events occurred between viruses and hosts. Viruses encoding for AGRs may confer benefits to the bacterial host by enhancing the host’s survival in the presence of antibiotics. This may be advantageous in the sponge environment, which has been reported to contain various antibiotics (Faulkner, 1978, Kim et al., 2006, Laport et al., 2009, Kelman et al., 2009). Similarly, Laffy et. al (2018) reported the presence of herbicide resistant genes assigned to Synechococcus phages were enriched in the sponge Xestospongia testudinaria. Herbicides are highly effective in controlling

58 cyanobacterial populations and it was postulated that the Synechococcus phages confer herbicide resistance to their cyanobacterial hosts and enhanced their host’s survival (Laffy et al., 2018).

Genes for the photosystem associated proteins PsbA and PsbD were detected in a viral contig most closely related to Synechococcus phage S-SM2 (in the Stylissa sp. 445 metagenome). Interestingly, psbA and psbD have also been recently found in coral viromes (Weynberg et al., 2017, Laffy et al., 2018) and in cyanophages infecting marine free-living Synechococcus spp. and Prochlorococcus spp. (Mann et al., 2003, Lindell et al., 2004, Weigele et al., 2007). Viral PSII genes are thought to supplement photosynthesis of their host and boost energy production, presumably for virion production (Lindell et al., 2005, Clokie et al., 2006). Additionally, genes encoding the carboxysome shell carbonic anhydrase were detected in the viral metagenome of Stlyissa sp. 445 (average of 2.2 copies/prokaryotic genome). Carbonic anhydrase is responsible for catalysing the conversion of carbonic acid to CO2 and is utilised by cyanobacteria and marine phytoplankton to support photosynthetic carbon fixation (Badger et al., 2005, Reinfelder, 2010). Viral carbonic anhydrase could thus help to increase carbon fixation, which could contribute to the production of virions.

Proteins of the aegerolysin family are widely distributed in fungi and bacteria, and to a lesser extent in plants, protozoa and insects (Berne et al., 2009, Butala et al., 2017). The functions of aegerolysins are not well understood, but they are thought to have a wide range of biological properties, including antitumor, antiproliferation and antibacterial activities (Berne et al., 2009, Butala et al., 2017). High abundance of aegerolysins were detected in the Scopalina sp. metagenomes (average abundance of 7.8 copies per prokaryotic genome) (Figure 3.7). Interestingly, the ascovirus Trichoplusia ni 2c, an obligate viral pathogen of Pseudoplusia includens larvae and other insects from the family Noctuidae, was reported to encode for a hypothetical aegerolysin-like protein (Wang et al., 2006). This hypothetical aegerolysin-like protein was found to be most similar to the aegerolysin protein P23 identified in the expression library of the host Pseudoplusia includens (Zhang et al., 2011). This indicates that the aegerolysin genes can indeed be transferred between the virus and host and may provide benefits to the host, such as antibacterial activity. Sponge

59 extracts and associated microorganisms have been reported to contain abundances of antitumor, antiproliferation and antibacterial activities (Lee et al., 2001b, Monks et al., 2002, Thakur et al., 2003, Laport et al., 2009, Yung et al., 2011). Our results suggest that viruses may carry aegerolysin-like genes which can potentially be passed onto other microorganisms via horizontal gene transfer.

Toxin-antitoxin systems are widely distributed in bacterial and are associated with the formation of antibiotic-tolerant (persister) cells (Butt et al., 2014). A putative bacterial toxin gene ydaT was found in the viral metagenome of Scopalina sp. at an average of 6.7 copies per prokaryotic genome (Figure 3.9). HicA toxin was identified to be actively expressed in the viral metatranscriptome of C. concentrica. YdaT belongs to the type II toxin-antitoxin systems (TA) (Sevin and Barloy-Hubler, 2007), where ydaS expressed the toxin and YdaT constitutes the antitoxin (Yamaguchi and Inouye, 2011). It was recently reported that YdaS and YdaT may both act as toxins by inhibiting the regular cell division of E. coli, which may provide resistance against β-lactamase antibiotics (Bindal et al., 2017). Interestingly, HicA toxins from Burkholderia pseudomallei has also been reported to cause cell growth arrest and increased the number of persister cells tolerant to ciprofloxacin and ceftazidime (Butt et al., 2014). Some sponges can contain high amounts of antibiotics and antimicrobial compounds (Faulkner, 1978, Hentschel et al., 2001, Torres et al., 2002, Sipkema et al., 2005, Kim et al., 2006, Kelman et al., 2009). The HicA/YdaT-type proteins might play a role in providing resistance against them. Also filament cells, which are generated as a result of the inhibition of cell division, are considered to be less likely to be consumed by phagocytotic cells, such as sponge amoebocytes or protozoans (Butt et al., 2014). Therefore, viral YdaT and HicA could potentially also provide bacterial symbionts with resistance to the feeding activity of the sponge host.

Thiamine (vitamin B1) functions as an enzyme cofactor involved in many essential metabolic pathways and is only produced by bacteria, fungi and plants (Jurgenson et al., 2009). Expressed genes associated with thiamine metabolism were enriched in the viral metatranscriptome of C. concentrica (Appendix Table S3.4). Specifically, tyrosine decarboxylase and many cysteine desulfurase (IscS) enzymes were detected (Appendix Table S3.4). The thiazole moiety of thiamine is derived from tyrosine, cysteine and 60 deoxy-D-zylylose-5-phosphate, which is linked with pyrimidine followed by a final phosphorylation step to yield thiamine pyrophosphate and subsequently thiamine (Begley et al., 1999). Tyrosine decarboxylase is part of the tyrosine synthesis pathway and catalyses the reversible chemical reaction of tyramine and carbon dioxide to L- tyrosine. The L-tyrosine can then enter the thiamine synthesis pathway as a substrate for the synthesis of thiazole moiety of thiamine (Jurgenson et al., 2009) (Appendix Figure S3.11). Cysteine desulfurase is involved in the initial stages of sulfur trafficking, which catalyses the conversion of L-cysteine to L-alanine and sulfane via the formation of an enzyme-bound persulfide intermediate (IscS-SH and IscS-SSH). The IscS-SSH persulfide transfers the sulfur to the ThiS enzyme (forming ThiS-COSH), which catalyses the incorporation of sulfur into the thiazole ring of thiamine (Begley et al., 1999, Mihara and Esaki, 2002) (Appendix Figure S3.11). As far as we know, this is the first time that viral-associated genes involved in thiamine metabolism have been detected and may be acquired from prokaryotic symbionts of sponges (Thurber, 2009, Fan et al., 2012). These phage-encoded genes to produce important precursors of thiamine could help to increase metabolic flow and boost bacterial growth during viral infection. Genes associated with other vitamin synthesis pathways, such as cobalamin (vitamin B12), have also been reported in sponge viromes (Laffy et al., 2018) and in myoviruses that infect Prochlorococcus (Sullivan et al., 2005).

3.6 Conclusion

The sponge environment is home to complex virus-host interactions, including lytic or lysogenic infection dynamics and contains variable environmental-derived and sponge- species specific viral assemblages. Environmentally-derived viruses observed in the sponge might be present by chance and thus may have no or little biological or ecological interactions with the microbial symbionts and/or the sponge host. Therefore, although the metagenomic approach is helpful to gather insights into the overall abundance and diversity of the viral community present, (meta)transcriptomic and/or viral cultivation should also be conducted to understand more ecologically and biological relevant viruses. However, metagenomes and (meta)transcriptome can help

61 us detect viruses that are enriched in sponges (compared to the environment), which may play an important role in the evolution of the sponge holobiont. Recent studies have highlighted that marine viruses can encode for auxiliary metabolic genes previously thought to be restricted to host genomes (Hurwitz et al., 2013, Enav et al., 2014, Weynberg et al., 2017, Laffy et al., 2018). Consistent with these studies, we also detected various auxiliary metabolic genes encoded by viruses associated with microorganisms found in sponges. These genes appear to expand the metabolic capacities of their hosts and potentially enhance the survival of infected cells for viral replication and virion production. Functional analysis of viral metagenomes indicate that viromes from diverse environments, including oceans, soils, corals, and animals (fish, mosquito and terrestrial associated) encode for similar metabolic functions, such as DNA replication and repair, and carbohydrate and amino acid metabolism (Dinsdale et al., 2008, Williamson et al., 2012, Zablocki et al., 2014, Adriaenssens et al., 2015, Weynberg et al., 2017, Laffy et al., 2018). This emphasises the common function of viruses to manipulate host cellular processes for viral survival and reproduction (Hargreaves et al., 2014). Some viral-associated functions appear to be specific to different host species. For example, genes associated with thiamine metabolism and biosynthesis of antibiotics were only observed in C. concentrica, and genes associated with photosystems were only detected in Stylissa sp. 445. Our study highlights that sponges contain taxonomically and functionally diverse viral communities, which may play an important role in the evolution of the sponge microbial communities, such as the spread of ‘fitness factors’ including toxins, antibiotics and vitamin biosynthesis.

Currently very little is known about the natural reservoirs of viruses that are carried by, or cause disease in marine animals, and even less is known about their potential to spread to terrestrial systems (Smith et al., 1998, Suttle, 2005). Some viruses, for example calici and distemper viruses, are thought to cycle between marine and terrestrial mammals (Smith et al., 2002, Philippa et al., 2004). Our findings indicate that sponges may act as a reservoir for the survival of viruses infecting marine and terrestrial (pathogenic) bacteria and may potentially influence the spread and/or growth of pathogens in the environment. Our study also suggests that not all sponges may act as reservoirs and that those sponges that do, may only interact with specific

62 bacterial pathogens. This has global ecological implications on the health of reef ecosystems. Thus, further studies into temporal population dynamics of viral communities and their respective hosts in marine sponges are much needed.

Chapter Four

63 4 Interactions of a novel phage and the sponge-associated bacteria Ruegeria sp. AU67

4.1 Co-authors and collaborators

This chapter is a collaboration between Mary T. H. D Nguyen, Cristina Moraru and Torsten Thomas.

MN conceived and designed the experiments, performed the experiments, analysed the data, prepared figures and/or tables, and authored the chapter. CM and TT conceived and designed the experiments. TT edited the chapter.

4.2 Introduction

Viruses are arguably the most abundant entity in marine environments with an estimate of 107-108 viral particles per millilitre of seawater. They play an important ecological role in geochemical nutrient cycling, controlling prokaryotic community composition and structure, and the evolution of microorganisms (Fuhrman and Suttle, 1993, Fuhrman, 1999, Breitbart, 2012). Virus-induced mortality of prokaryotes in oceans have been estimated to be as high as 10-50% (Wommack and Colwell, 2000, Weinbauer, 2004) with an estimated ratio of up to 15 viruses per bacteria (Fuhrman, 1999, Mokili et al., 2012). Our previous study (Chapter three) revealed that microbial metagenomes of sponges contained a much lower average ratio range of 4-0.7 viral contigs per prokaryotic genome. This is surprising given the large amount of water filtered through sponges (Vogel, 1977) and the large abundance of microorganisms as potential hosts (Hentschel et al., 2006). The lower rate of viral infection in the sponge environment may be attributed to the high abundances of defense mechanisms, such as CRISPRs/Cas proteins and restriction-modification systems (Fan et al., 2012, Horn et al., 2016).

64 Currently very little is known about the diversity and ecological function of viruses in marine sponges. So far only two studies have investigated viral assemblages associated with marine sponges, our previous study (Chapter three) and a recent study by Laffy et. al (2018). Both studies revealed diverse and abundant viral assemblages, including bacteriophages and eukaryotic-viruses. Functional gene analysis indicated that viral assemblages could be involved in the evolution of microorganism through virus- mediated genetic transfer such as the spread of ‘fitness factors’ (Chapter three, Laffy et al., 2018). Evidence for dynamic virus-host interactions such as lytic and lysogenic infections can be inferred from the viral metagenomic and transcriptomic analysis (Laffy et al., 2018, Chapter three). However, isolation of viruses and its host are needed to investigate the details of virus-host dynamics. Cultivation of bacterial hosts are thus required for the isolation of phages and this represent one of the bottlenecks of studying virus-host interactions in the sponge environment (Esteves et al., 2016). So far, only one study has reported the isolation of phage φJL001 from a sponge associated α-proteobacterial strain JL001 (Lohr et al., 2005). Infection characteristics and genomic analysis of phage φJL001 indicated that the phage may be temperate with pseudolysogenic characteristics (Chen and Lu, 2002, Lohr et al., 2005).

Lytic and lysogenic interactions of viruses are well understood. During the lytic cycle, viruses take advantage of the host cell to extensively replicated their DNA and package it into virions, which are released through host lysis and spread to infect other neighbouring cells (Fuhrman, 1999, Cenens et al., 2013). Lysogeny is the ability of viruses to integrate their genome as a prophage into the host chromosome allowing the viral genome to stably replicate as part of the host’s replication cycle (Zeng et al., 2010). Prophages can be activated into the lytic cycle by environmental factors or host cues, such as DNA damage or through low rates of spontaneous induction (Jiang and Paul, 1998, Quinones et al., 2005, Knowles et al., 2016). A less well-defined type of virus-host interaction is termed pseudolysogeny. The current consensus definition of pseudolysogeny is a virus-host interaction that neither establishes a long-term, stable integration (lysogeny) nor elicits a lytic response (Ripp and Miller, 1997, Jiang and Paul, 1998, Fuhrman, 1999, Williamson et al., 2001, Łoś and Węgrzyn, 2012, Cenens et al., 2013). Pseudolysogeny has been related to host starvation, where virion production

65 and cell lysis proceed when nutrients were added to the starved hosts (Ripp and Miller, 1997, Ripp and Miller, 1998). Alternatively, pseudolysogeny may be regarded as a transient state of host immunity, possibly induced by the release of immunizing agents from lysed infected cells, promoting coexistence of virus and host cells (Moebus, 1996).

Pseudolysogeny has been postulated to confer a number of advantages over obligate lytic viruses, such as protection of viral DNA from harsh conditions outside the host (Ripp and Miller, 1997, Ripp and Miller, 1998). This might be particularly advantageous in the sponge environment, which contains abundant anti-viral compounds (Perry et al., 1988, Jares-Erijman et al., 1991, Perry et al., 1994, Sipkema et al., 2005, Sagar et al., 2010, Horn et al., 2016). Additionally, physiochemical factors, such as UV-light, pH and temperature, can affect the survival of virions (Chibani-Chennoufi et al., 2004). The potential advantage of pseudolysogeny over true lysogeny may be to allow the virus to escape the host (i.e. change from the lysogeny into a lytic life cycle ) without being dependent on host physiochemical cues, such as DNA damage or through spontaneous induction (which is estimated to occur once every 105-108 cells) (Czyz et al., 2001). The virus thus can potentially leave the dying/compromised host cells/population to infect neighbouring cells and decreased the risk of the virus possibly “dying” with the host cells/population as a prophage.

The transition from lysogeny to lytic life cycles has been correlated to a rise in host density in pelagic environments (Jiang and Paul, 1998, Paul, 2008, Payet and Suttle, 2013, Maurice et al., 2013). However, in environments with high microbial densities, such as coral reefs, a “piggyback-the-winner” model has been proposed, whereby an increase in host density is thought to be accompanied by a transition from lytic to temperate (lysogenic) dynamics (Knowles et al., 2016). This model reflects the increased contribution of temperate viruses in ecosystems with high host abundance, such as sponges, yielding “more microbes” and “fewer viruses”. The model suggests that in environments with high host density, viruses exploit their hosts through lysogeny, rather than killing them. This is consistent with the observation of high abundances of the host bacterium Synechococcus and cyanophages in the sponge Stylissa sp. 445 (Chapter Three, Fan et al., 2012). Lysogeny may also be important in

66 conferring resistant to superinfection by related viruses (Thingstad et al., 2014). Furthermore, lysogeny can decouple microbial taxonomic and functional composition through horizontal gene transfer (Kelly et al., 2014). Virulence genes carried by viruses could allow host bacteria to evade protistan/phagocytic predation (Knowles et al., 2016). These viral-host dynamics therefore play an important role in the sponge microbial community diversity, function and health (Barr et al., 2013, Silveira et al., 2015, Weitz et al., 2015).

Viral activities regulate biodiversity and food web efficiency through the conversion of biomass to dissolved and particulate organic matter via host cell lysis (termed “the viral shunt”) (Suttle, 2007). The extent and efficiency of viruses to drive microbial processes are impacted by biotic and abiotic environmental factors, such as host physiology, temperature, salinity, UV and nutrients (Mojica and Brussaard, 2014). Other factors that influence viral activity are grazing. For example, the coral reef sponge Negombata magnifica was reported to have an average removal efficiency rate for virus of 23% (from the seawater), which may affect the virus-to-host ratios in the surrounding waters (Hadas et al., 2006). However, effects of the sponge environment on virus activity or virus-host interactions have so far not been investigated. There has been indirect evidence to suggest that sponges may act as reservoirs for viruses of marine and terrestrial pathogens (Chapter three), which has global ecological implications on the health and disease of reef ecosystems. Therefore, research on how sponge-derived factors may influence virus-host dynamics are much needed. To gain a clearer understanding of viral-host interactions within sponges, we aim here to isolate viruses from sponge-associated bacteria to provide a virus-host model system and additionally, to investigate how sponge-derived factors may influence virus-host dynamics.

The isolated sponge-associated bacterial strains used in this chapter were chosen due to their potential ecological or biological importance in the sponge microbial community in the well-studied model sponges Cymbastela concentrica, Tedania anhelans and Scopalina sp.. This was accessed based on their high relative abundance in the cultivated fraction and sequencing studies and/or potential important function within the sponge microbiome such as cyanobacteria (Fan et al., 2012, Esteves et al.,

67 2016). Bacteria from different phyla that dominated the cultured fraction in sponges C. concentrica, T. anhelans and Scopalina sp. were chosen as hosts to isolate viruses such as Proteobacteria (order Rhodobacteracae, Pseudovibrio sp., Ruegeria sp. and Loktanella sp.), Firmicutes (order Bacillales, Bacilllus sp.), Bacteroidetes (order Flavobacteriales, Tenacibaculum sp.) and cyanobacteria (Leptolygnbya sp.) (Esteves et al., 2016, Unpublished data from A. Esteves).

4.3 Materials and Methods

4.3.1 Sponge sampling and viral extraction The sponges Cymbastela concentrica, Tedania anhelans and Scopalina sp. were collected on the 13-09-2016 at Bare Island in Botany Bay, NSW, Australia (33° 59’S, 151° 14’E) by SCUBA diving at depths of 7-10 m and placed individually with surrounding seawater into Ziploc ® bags. Samples were transported to the UNSW laboratory (approximately 30 min) in a large container containing natural seawater (NSW). All following work was conducted in a laminar flow cabinet under sterile conditions.

Viruses were extracted from sponge tissue using the centrifugation-filtration method adapted from Thurber et al. (2009). Briefly, sponges were washed three times with sterile calcium magnesium-free seawater (CMFSW; 25 g NaCl, 0.8 g KCl, 1 g Na2SO4 and

0.04 g NaHCO3 per 1 L) and place in a 50 mL Falcon tube (Fisher Scientific, Melbourne, Australia) at a concentration of 1 g/mL of sterilised NSW. Sponge tissue were homogenized for 1 min using a dispersing homogeniser (Ultra-Turrax TR50, IKA, Selangor, Malaysia) at maximum speed. To remove cellular tissue and microorganisms, the homogenate was first centrifuged at 2500 x g for 10 min and the supernatant was vacuum filtered through a 0.45 µm filter (Whatman, Sigma-Aldrich, St. Louis, USA). The flowthrough was then centrifuged at 2500 x g for 5 min and the supernatant was filtered through a 0.22 µm filter (Whatman, Sigma-Aldrich, St. Louis, USA). Viral particles were precipitated using 6% v/v final concentration of polyethylene glycol (PEG) 8000 (Sigma-Aldrich, St. Louis, USA) and incubated overnight at 4°C. The viral- PEG solution was centrifuged at 76,000 g at 4°C for 30 min. The supernatant was

68 discarded, and viral pellets were resuspended in 200 µL of 0.22 µm filtered and autoclaved NSW. Samples were stored at 4°C in the dark until further processing.

4.3.2 Sponge extract preparation Extract of the sponges C. concentrica, T. anhelans and Scopalina sp. were generated to understand how compounds from the sponge can influence the virus-host dynamics. Sponge specimens were collected as above on the 13-09-2016. 10% w/v sponge extract were prepared according to the methods described by Esteves et al. (2016) with minor modifications. Briefly, sponge tissue was homogenised in 10 mL of sterile NSW per gram of sponge sample using a dispersing homogeniser (Ultra-Turrax TR50, IKA, Selangor, Malaysia) at maximum speed on ice until complete tissue disruption. The homogenate was shaken at 100 rpm for 30 min at 4°C and filtered through a 125 µm metal sieve to remove large sponge debris. The supernatant was then filtered through a series of vacuum filtration steps, using a decreasing pore size of 12 µm, 0.45 µm and 0.22 µm filters (Whatman, Sigma-Aldrich, St. Louis, USA). Sponge extracts were freeze dried with an Alpha 1-2 Routine Freeze Dryer (Betatek, North York, ON, Canada) and stored at -80°C. Since all sponge extracts were treated with the same ratio of sponge tissue to seawater (1g of sponge per 10 mL of natural seawater), the resulting salt (from seawater) to sponge compounds would be the same in all samples. Sponge extracts were then used at a final concentration of 10% w/v in one-step infection growth curves (see below).

4.3.3 Isolation of sponge-associated bacteria Bacterial strains used in this studied were provided by Esteves et al. (2016) or cultivated in this study as indicated in Table 4.1. Bacillus sp. 673, Tenacibaculum sp. 330, Pseudovibrio sp. 243, Ruegeria sp. AU67 and Loktanella sp. 529 were grown in marine broth agar (37.4 g/L DifcoTM (BD Biosciences, New Jersey, USA) 15 g/L Bacteriological agar (Sigma-Aldrich, St. Louis, USA)). The cyanobacterium Leptolyngbya sp. (cultivated in this study) was isolated using the methods described in Esteves et al.

(2016) for cyanobacteria, and grown in liquid SNAX media (25.5 mg K2HPO4, 90 mg

KNO3, 1 mg Na2CO3, 0.5 mg Na2EDTA.2H2O, 13 mg (NH4)2SO4, 750 mL NSW, 250 mL MilliQ water and 100 µL micronutrients solutions (6 mg/mL iron citrate, 1.4 mg/mL

MnCl2.4H2O, 20 µg/mL CoCl2.6H2O and 0.22 mg/mL ZnSO4.7H2O)) at room

69 temperature, shaking at 100 rpm and under continuous illumination with cool white light (~20 µE m-2 s-1).

Table 4.1 Sponge-associated bacterial cultures used to isolate viruses.

Strain Sponge source Genome sequenced; Reference GenBank ID Bacillus sp. AU673 C. concentrica No; N/A (Esteves et al., 2016) Tenacibaculum sp. AU330 Tedania sp. Yes; N/A Unpublished data from Esteves, A. Pseudovibrio sp. AU243 C. concentrica Yes; 6511168 (Esteves et al., 2016) Ruegeria sp. AU67 Tedania sp. Yes; 6511288 Unpublished data from Esteves, A. Loktanella sp. AU529 C. concentrica No; N/A (Esteves et al., 2016) Leptolyngbya sp. CCY C. concentrica Yes; N/A This study

4.3.4 Viral isolation and purification Bacterial cultures were grown at room temperature, shaking at 100 rpm to exponential phase (2-3 days, Loktanella sp. 4-5 days). Dilutions of 100 µl of viral extracts (100, 10-1 and 10-2) from each sponge species were added to 1 mL of bacterial culture in triplicates and incubated at room temperature and shaking at 100 rpm for 1 hr to facilitate viral absorption. Mixtures were added to 4 mL of 0.7% marine soft agar (37.4 g/L Marine broth, DifcoTM (BD Biosciences, New Jersey, USA) 7 g/L Bacteriological agar (Sigma-Aldrich, St. Louis, USA) (maintained at 50°C), briefly vortexed and poured onto marine agar plates (37.4 g/L Marine broth, DifcoTM (BD Biosciences, New Jersey, USA) 15 g/L bacteriological agar (Sigma-Aldrich, St. Louis, USA). Plates were incubated at room temperature in the dark under sterile conditions until viral plaques were observed (up to ~6 months).

Viral isolates were purified by picking viral plaques with a pipette tip and transferred into an Eppendorf tube with 500 µL of marine broth (37.4 g/L Marine broth, DifcoTM (BD Biosciences, New Jersey, USA)). The viral plaques were incubated at room temperature for 1 hr to facilitate the diffusion of viruses into the solution and the soft agar was removed. Viruses were concentrated with 6% v/v PEG 8000 at 4°C overnight. Samples were then centrifuged at 76,000 g for 30 mins at 4°C and resuspended in 200 µL of marine broth. Dilutions (10-2, 10-4, 10-6 and 10-8) of the viral lysate were used to 70 re-infected new bacterial hosts (grown to logarithmic phase) and plated to get single plaque forming units (PFUs). Plaques were picked, purified and plaque assays were repeated three times to get pure viral isolates, which were stored at either 4°C or - 20°C until further processing.

4.3.5 Transmission electron microscopy Pure viral isolates were visualised using transmission electron microscopy (TEM) using a JOEL TEM-140 at the Mark Wainwright Analytical Centre (University of New South Wales, NSW, Australia). One drop of viral sample was placed onto a formvar-coated copper grid and left for 2 min before excess sample was wicked off with filter paper. The grids were then placed twice on to a drop of water and wicked off with filter paper to get rid of the salts. Samples were then stained with 2% uranyl acetate for 2 min and excess stain was wicked off with filter paper. Grids were viewed with transmission electron microscopy immediately and images were captured using a Gatan digital camera and Digital Micrograph software (Mark Wainwright Analytical Centre, University of New South Wales, NSW, Australia).

4.3.6 Bacterial genome sequencing Bacterial DNA from Ruegeria sp. AU67, Pseudovibrio sp. 243, Tenacibaculum sp. 330 and Leptolyngbya sp. were isolated using the DNeasy Blood and Tissue extraction kit (Qiagen), according to the manufacturer’s instructions. Bacterial genome sequencing was conducted with Illumina sequencing using MiSeq 2 x 250 bp chemistry at the Ramaciotti Centre for Genomics (University of New South Wales, NSW, Australia). Bacterial sequences were assembled using SPAdes v3.7.0 (Bankevich et al., 2012) with default settings and assembled contigs were annotated via the Integrated Microbial Genomes database (IMG) (Markowitz et al., 2011).

4.3.7 Viral genome sequencing and phylogenetic analysis One viral plaque plate containing high numbers (>300) of PFUs/mL were used to extract viral DNA. Ten mL of marine broth was added to the soft agar plate and incubated at room temperature for 1 hour to facilitate diffusion of viruses into the liquid media. The supernatant was filtered through a 0.22 µm filter (Millipore, Sigma- Aldrich, St. Louis, USA) and precipitated with sterile 6% v/v PEG 8000 (Sigma-Aldrich,

71 St. Louis, USA) overnight at 4 °C. Viral PEG solutions were centrifuged at 76,000 g at 4°C for 30 mins. The supernatant was discard and viral particles were resuspended in 100 µL molecular grade water. Free bacterial DNA was degraded with 10 U DNase I (RNase-free) (New England Biolabs, Massachusetts, USA) and viral DNA extraction was then conducted using the DNeasy Blood and Tissue kit (Qiagen, Hilden, Germany) according to manufacturer’s instructions. Sequencing was conducted using Illumina Miseq 2 x 250 bp chemistry at the Ramaciotti Centre for Genomics (University of New South Wales, NSW, Australia).

Viral sequences were assembled using SPAdes v3.7.0 with default settings and assembled contigs were annotated via the Integrated Microbial Genomes database (IMG) (Markowitz et al., 2011). The Ruegeria phage 67 genome has been submitted to Genbank with submission number SUB:2136728. Viral hallmark gene encoding the DNA polymerase (obtained from the genome annotation) were used for phylogenetic analysis. DNA polymerase protein sequences were aligned using MUSCLE and phylogenetic trees were constructed using the maximum likelihood tree with complete deletion and bootstrap of 100 using the MEGA v6 program (Tamura et al., 2013). Phage genome protein sequences were search with BLASTp against the non-redundant protein database from the National Centre of Biotechnology Information (NCBI). Multiple genome comparison was conducted with Mauve (Darling et al., 2004) using default progressive alignment parameters (http://darlinglab.org/mauve/user- guide/introduction.html).

4.3.8 Bacterial growth curves Growth curves of the bacterium Ruegeria sp. AU67 were determined in biological replicates. Bacterial growth was measured via spectrophotometry (OD= 650 nm) using a Beckman DU 530 spectrophotometer (Beckman Coulter, California, USA) and flow cytometry (BD AccuriTM C6 Plus Flow Cytometer, BD Biosciences, New Jersey, USA; see below). The growth rate of Ruegeria sp. AU67 was determined in marine broth and marine broth amended with three sponge extracts (C. concentrica, Scopalina sp. and T. anhelans) with a final concentration of 10 % w/v and filtered through a 0.22 µm filter (Merck Millipore, Massachusetts, USA).

72 4.3.9 Plaque assays and one-step infection growth curves All infection experiments were conducted with bacterial cells in log phase (OD= 0.4, ~2 x 108 cells/mL). Infection experiments were conducted in marine broth (DifcoTM, BD Biosciences, New Jersey, USA), and marine broth supplemented with a final concentration of 10% w/v sponge extracts of C. concentrica, Scopalina sp., and T. anhelans. One-step growth curves were adapted from Dang et al. (2015) with slight modifications. Briefly, phage-host infection experiments were conducted in three biological replicates and a negative control (host bacterial culture with no phages) were analysed over time. Bacteria were infected with phages at a multiplicity of infection (MOI) of 2 and incubated at 4°C for 15 min to facilitate phage absorption. Infection cultures were then diluted 100-fold in respective media (above) to synchronise the infection, this timepoint was considered as T0 (0 hr). Samples of infected cultures were collected at different timepoints for analysis of: 1) virion production 2) bacterial cell counts and 3) microscopy of phage infection. Virion production was enumerated via plaque forming units (PFUs/mL). Collected samples were filtered through 0.22 µm filters (Merch Millipore, Massachusetts, USA) to remove bacterial cells and free (extracellular) phages were enumerated by averaging the PFUs/mL from three technical replicates of two consecutive dilution series. Bacterial cell counts were conducted via flow cytometry (see below) and microscopy of phage infections were visualised with DAPI stain (see below). The latent period was estimated from the time point where free virus abundances did not increase significantly from the previous time point. The burst size was the average number of phages released per infected host cell and calculated as the ratio between the number of phages before and after the burst (Middelboe et al., 2010).

4.3.10 Bacterial cell count via flowcytometry At each assayed time point, infected cultures were sampled, diluted with 1 x phosphate buffered saline (PBS: 8 g Na Cl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4, pH 7.4 per 1 L) and fixed with 25 % glutaraldehyde solution (Sigma-Aldrich, St. Louis, USA) at a final concentration of 2%. Fixation was conducted at 4°C for 30 min and stored at - 80°C until processing. For flow cytometry, samples were diluted with 1 x PBS, 20 µL of 1 x rainbow fluorescent particles (3.0-3.4 µm) solution (SpheroTM, BD Biosciences, BD

73 Biosciences, New Jersey, USA) and 20 µL of 1 x SYBR green solution (Sigma-Aldrich, St. Louis, USA), and incubated at room temperature, in the dark for 30 min. Bacterial cell counts were measured using the BD AccuriTM C6 Plus Flow Cytometer (BD Biosciences, New Jersey, USA) with parameters of 60 µL limit and slow flow rate.

4.3.11 Microscopy of phage-host infections At each assayed time points during the one-step infection, 40 mL of infected cultures were extracted and fixed with 25% paraformaldehyde (Sigma-Aldrich, St. Louis, USA) at a final concentration of 4% and incubated at room temperate for 1 hr. Fixed cultures were vacuum filtered onto 0.22 µm polytetrafluoroethylene (PTFE) filters (Merck Millipore, Massachusetts, USA) and washed with 10 mL of sterile MilliQ water. Filters were air dried and stored at -20°C until further processing. Cells were fixed onto filters via successive ethanol washed at 50%, 80% and 100% for 1 min and then air dried. Filters were covered with permeabilization and RNase buffer (0.05 M EDTA (pH 8.0), 0.1 M Tris-HCl (pH 8.0), 0.5 mg/ mL lysozyme, 100 µg/ mL RNaseA) and incubated for 1 hr at 37°C in a humid chamber to prevent evaporation. Filters were then washes in succession of 1 x PBS solution for 5 min, then MilliQ water for 1 min and lastly in 100% ethanol for 1 min. Filters were then stained with 4 µg/mL DAPI (4’, 6’ -diamindino-2- phenylindole) solution and embedded in a 3:1 mix of Citifluor TM (glycerol-PBS solution, Citifluor Ltd, United Kingdom) and Vecasheild ® antifade mounting media onto microscopy slides. Bacteria cells were visualised with an epifluorescence microscope (Zeiss Imager.Z2m) and image processing was performed with the AxioVision v4.8.2 program.

4.4 Results

4.4.1 Virus isolation Six bacterial strains were used as potential bacterial hosts to isolate viruses, Leptolyngbya sp., Tenacibaculum sp. 330, Pseudovibrio sp. 243, Ruegeria sp. AU67, Loktanella sp. 529 and Bacillus sp. 673. However, viral plaques were only obtained for Ruegeria sp. AU67, which was isolated from the sponge T. anhelans, using viral extracts from the same sponge. This virus was purified and termed Ruegeria phage 67. TEM of this virus revealed a morphology comprising of a non-enveloped icosahedron

74 head (40-50 nm in width) and a tail of 150-200 nm (length) x 10 nm (width) with short terminal fibres, consistent with viruses belonging to the Siphoviridae family (Figure 4.1).

Figure 4.1 TEM images of Ruegeria phage 67. Image A depicts phages at possibly different viral maturation stages (capsid head development and with and without tail formation). Image B shows details of the tail and short terminal fibres

4.4.2 Phage-host dynamics The one step infection growth curves were performed to elucidate phage-host dynamics under standard laboratory conditions (marine broth) and with sponge extracts to assess if or how sponge-derived factors may influence the virus-host interaction. The growth rate of Ruegeria sp. AU67 host (non-infection experiments) in marine broth (MB), MB plus Scopalina sp. and C. concentrica extracts were comparable at 49.4, 45.7 and 47.4 min, respectively (Figure 4.2). Interestingly, no bacterial growth was observed in marine broth supplemented with 10% w/v T. anhelans sponge extracts, and subsequently no increase in extra-cellular PFUs counts were observed (burst size = 0.2 PFUs/cell, no significant difference was seen in the two-tailed t-test between time points T0 and T12, P-value = 0.9). In contrast, the burst size of viruses in marine broth, and marine broth supplemented with Scopalina sp. and C. concentrica sponge extracts were 22.3, 33.9 and 27.7 PFUs/cell, respectively. The estimated latent period was approximately 6 hours in marine broth, and marine broth supplemented with C. concentrica and Scopalina sp. sponge extracts (Figure 4.2). The rate of bacterial lysis appeared to equal the rate of bacterial cell growth in infection cultures as no changes in the bacterial population was observed over the 12 hours.

75

Figure 4.2 One-step infection of Ruegeria phage 67 in marine broth and marine broth amended with 10% w/v of sponge extracts from T. anhelans, Scopalina sp. and C. concentrica. The y-axis shows counts/mL (bacterial cells and PFUs) and x-axis indicate time after dilution in hours. The blue lines indicate average bacterial cell counts/mL in control experiments (no phage) for each medium. The red lines indicate average bacterial cell counts/mL in infection cultures (bacteria and viruses) and the green lines indicate average viral PFUs/mL. Error bars depict standard deviations from the mean.

4.4.3 Morphological changes in the bacterial host induced by viral infection Microscopy analysis of the phage-host infection in marine broth showed a drastic morphological change in Ruegeria sp. AU67 after phage infection (Figure 4.3). Uninfected bacteria appear rod-shaped with a size of 2-5 µm (length) x 1 µm (width). After only 15 minutes of phage infection (T0), host cells appear rounded (oval shaped) with an increase in cellular diameter of 2-3 µm (length remained approximately the same), till two hours post infection (T2). After four hours post infection (T4), large cellular debris could be observed indicating cell lysis. At T6 “infected” cells appear to increase in size 10-20 µm (length) x 1-3 µm (width), but rod-shaped cells similar to “non-infected” cells (in the negative control) was still present. The pear-shaped, infected cell morphology was more pronounced in T8, with an increase in average cell size of 5-10 µm (length) x 2-5 µm (width). The majority of cells appear to be infected in T8, possibly indicating a second wave of infection. Rounded pear-shaped and

76 elongated cells are observed at T10, T12 and T14. At these time points there was also an increased number of smaller rod-shaped cells (similar to non-infected cells). The ratio of “non-infected” to “infected” cell morphology continued to increase in T16, T18 and T20, suggesting possibly the death of infected cells and growth of resistant cells. Some cells appear dramatically elongated at T16 and T18 (20-45 µm in length) (Figure 4.3).

4.4.4 Genomic analysis of Ruegeria sp. AU67 and Ruegeria phage 67 The bacterium Ruegeria sp. AU67 had a genome size of 4.6 Mbps, a G+C content of 56.5% and a total of 4557 ORFs, of which 98.82% are protein coding. Interestingly, no CRISPRs were found in Ruegeria sp. AU67, but were found in the other bacterial hosts used to isolate phages (i.e. Tenacibaculum sp. AU330 (two CRISPRs), Leptolyngbya sp. CCY (five CRISPRs), and Pseudovibrio sp. AU243 (one CRISPR) (Appendix Table S4.1). However, Ruegeria sp. AU67 contained 65 proteins annotated by Cluster of Orthologous Genes (COG) to “defense mechanisms”, including against phage infection, such as two enzymes belonging to the type I restriction-modification system. The COG analysis further revealed 19 proteins associated with “mobilome, prophages and transposons”. The majority of the proteins found were putative transposases, however phage proteins were also observed, such as a phage uncharacterized protein (putative large terminase) C-terminal domain containing protein and an integrase. The viral genome (see below) was searched against the host genome and one protein was found to have 86% identity to a thymidylate synthase (ThyX). ThyX catalyses the reaction of dUMP to dTMP (thymidylate), which is an essential DNA precursor (Graziani et al., 2004) and which could be used by the virus to generate higher concentrations of deoxynucleotides (dNTPs) for DNA replication (Graziani et al., 2004, Myllykallio et al., 2002).

Rugeria 67 phage is a linear double-stranded DNA virus with a genome size of 61 680 bp and G+C content of 58.87% (Figure 4.4). A total of 64 ORFs were identified with 28% of the proteins resulted in a functional prediciton using KEGG, COG, PFAM and TIGRFAM annotation systems while 72% had no known functions (hypothetical proteins) (Figure 4.4). The genome contained genes encoding for stuctural proteins, DNA replication and potentail proteins assocaited with host interactions.

77

Figure 4.3 Morphological changes observed in phage infections of Ruegeria sp. AU67. NEG indicates non-infection experiment at T0 (after dilution). T0-T20 indicate hours post infection of samples in marine broth. Scale bar indicates 5 µm.

78

Figure 4.4 Ruegeria phage 67 genome annotated via IMG and visualised with Snap gene (GSL Biotech LLC, http://www.snapgene.com/). ORFs are indicated with arrows in the forward and reverse directions. Arrows in white indicate hypothetical proteins, green indicate proteins annotated with KEGG functions, blue arrows indicate proteins with COG functions, red are protein predicted with PFAM, and yellow are proteins predicted with TIGRFAM databases. Proteins that multiple hits to different annotation systems, the highest functional group colour was assigned.

79 4.4.5 Phage genome comparison Phylogenetic analysis of Ruegeria phage 67 was conducted using the DNA polymerase protein sequence. Ruegeria phage 67 was most closely related to two roseophages, Silicibacter phage DSS3-P1 (family unclassified) and Ruegeria phage DSS3-P1 (family Siphoviridae) (Figure 4.5). The phage clustered with other Siphoviridae viruses that infects phylogenetically different bacterial hosts and was distantly related to Podoviridae N4 roseophages (Figure 4.5). The Alphaproteobacteria virus φJI001 (isolated from a sponge-associated α-proteobacteria (Lohr et al., 2005), also belonged to the Siphoviridae family, but was distantly related to Ruegeria phage 67 (Figure 4.5).

Figure 4.5 Phylogenetic analysis of Ruegeria phage 67 (highlighted in bold) using the DNA polymerase protein sequence. Viral families Siphoviridae, Podoviridae and outgroup are indicated. Protein sequences were aligned with MUSLCE and maximum likelihood tree with percentage bootstrapping value of 100 were constructed using the MEGA v6 program (Tamura et al., 2013). Number scale bar indicates number of base substitutions per site.

80 Multiple genome comparison showed similarities between the genomes of Ruegeria phage 67, Ruegeria phage DSS3-P1, Silicibacter phage DSS3-P1 and Synechococcus virus S-ESS1 (Figure 4.6). Similarities were observed in genes associated with structural proteins, such as the tail and tape-measure proteins (highlighted in the purple block), capsid protein (highlighted in the red block). Genes encoding for the portal protein, the gpW protein (head to tail joining protein), the terminase large subunit and the DNA polymerase (highlighted in the orange block) were similar in all four genomes. Virulence associated protein E was homologous to a virulence-associated protein in Ruegeria phage DSS3-P1, but to hypothetical proteins in Silicibacter phage DSS3-P1 and Synechococcus virus S-ESS1 (Figure 4.3). Genes highlighted in the yellow block encoded for hypothetical proteins, ribonucleoside-diphosphate reductase alpha chain and chromosomal segregation protein had sequence similarities to hypothetical proteins in Ruegeria phage DSS3-P1 and Silicibacter phage DSS3-P1, and to three proteins in Synechococcus virus S-ESS1 encoding for FAD-dependent thymidylate synthase, DUF 932 domain-containing protein and ribonucleoside-diphosphate reductase adenosyl, cobalamin-dependent protein (Figure 4.6). The overall genome arrangement of Ruegeria phage 67 most closely resembled Ruegeria phage DSS3-P1 genome (Figure 4.6).

Approximately 34% of the Ruegeria phage 67 genome did not have any homology to the three other genomes. These unique genes (locus tags Ga0138520_111- Ga0138520_116, Ga0138520_1120-Ga0138520_1134, and Ga0138520_1160-1161 and Ga0138520_1164) encoded for hypothetical proteins, uncharacterized conserved protein, UPF0335 family, thymidylate synthase, putative peptidoglycan binding domain-containing protein and subtilase family protein, respectively (Appendix Table S4.2). Thymidylate synthase was detected to have high sequence similarity to the same protein from the host Ruegeria sp. AU67. The putative peptidoglycan binding domain- containing protein was found to have sequence similarity to a DUF3380 domain- containing protein from Pararhizobium sp. XC0140 and subtilase had closest similarity to a hypothetical protein from Roseobacter phage RD-1410W1-01 (Appendix Table S4.2).

81

Figure 4.6 Multiple genome comparison using the Mauve program (Darling et al., 2004). The number on top of each genome indicate the base pair position. The colour blocks indicate sequence regions that are homologous to the other genomes (connected by same coloured lines). Blocks above the centre black line indicate regions in the forward orientation relative to the first genome. Blocks below the centre line indicate regions that align in the reverse complement orientation. Regions outside the blocks indicate unique sequences. Inside each block, depicts the similarity profiles of the genome sequence, the height of the similarity profiles corresponds to the average level of conservation in that region. The areas in white in the blocks were not aligned and contains sequence elements specific to that genome. The white band below the black centre lines indicates ORFs in each phage genome.

4.5 Discussion

4.5.1 Isolation of lytic viruses from sponge-associated bacteria Six different sponge-associated bacteria strains were used as bacterial hosts to isolate viruses, however only one viral isolate was obtained from Ruegeria sp. AU67 (cultivated from the sponge T. anhelans) using T. anhelans viral extracts. Genome sequences were obtained for four bacteria (Ruegeria sp. AU67, Tenacibaculum sp. AU330, Pseudovibrio sp. AU243, Leptolyngbya sp. CCY). Interestingly, genome analysis of the last three bacteria detected CRISPRs and all bacterial hosts contained genes associated with phage-defense mechanisms, such as enzymes involved in restriction modification systems (Appendix Table S4.1). Another form of viral resistance is the presence of prophage/s, which can confer immunity to viral infection from related phages (Stern and Sorek, 2011). A putative prophage was found in the genome of Pseudovibrio sp. AU243 and BLASTp of the terminase protein sequence showed

82 similarities to terminase proteins from Rhodobacter phages RCSaxon, RcCronus and RcRhea (~47% identity) (Bollivar et al., 2016) (Appendix Table S4.1). This is consistent with data from culture-independent studies that showed a lower number of viral contigs in sponges compared to seawater (Chapter three) and high abundances of CRISPRs and defense-related mechanisms in sponge metagenomes (Fan et al., 2012, Horn et al., 2016). Due to sponges’ large filter feeding rate (Vogel, 2008), and the high abundance of viruses in seawater, (Breitbart, 2012), sponges are likely to be exposed to extremely high viral loads (up to 2.4 x 1014-15 viruses/L per kg of sponge per day) (Vogel, 2008). Thus, the sponge microbial communities would need effective defence mechanisms against foreign DNA and viral infections (Fan et al., 2012, Horn et al., 2016, Webster and Thomas, 2016).

So far, only one lytic bacteriophage pT24 has been reported to infect Tenacibaculum mesophilum and Tenacibaculum discolour (Khoa et al., 2017). Currently, no studies have been published on phages that infect the genus Pseudovibrio and no phage genomes for this genus are present in the NCBI database. Two phages, cyanophage LPP-1 and Phormidium phage Pf-WMP3, have been reported to infect Leptolyngbya boryana and Leptolyngbya foveolarum, respectively (Simis et al., 2007, NCBI database). Interestingly, the genome of Leptolyngbya boryana NIES-2135 contains phage proteins, such as an integrase, a phage terminase large subunit, and a DNA polymerase, which suggest that the bacteria may potentially be a lysogen with previous acquisitions of prophages (accession number: BAY58527.1, NCBI). The low number of reported phages and the presence of phage defense mechanisms such as CRISPRs suggests that lytic infections may be rare for these three genera.

Ruegeria belong to the family Rhodobacteraceae was obtained in the cultured fraction of C. cymbastela, Scopalina sp. and T. anhelans (Unpublished data from A. Esteves, Esteves et al., 2016). Genome analysis of Ruegeria was shown to be enriched in a putative hemolysin (COG3042), an adhesin (COG3468), a predicted extracellular nuclease (COG2374), a WD40 repeat protein (COG2319) and a tetratricopeptide repeat (TPR) domain (pfam13429) (Díez‐Vives et al., 2018), however the function of Ruegeria in these sponges are yet to be elucidated. Interestingly, Ruegeria have been previously cultivated from other sponge species such as Suberites domuncula (Mitova et al.,

83 2004a, Mitova et al., 2004b), Mycale laxissima (Zan et al., 2011) and various sponges from the coast of South East India (Anand et al., 2006). Ruegeria sp. SDC-1 (Mitova et al., 2004a, Mitova et al., 2004b) and Ruegeria sp. SC15 (Anand et al., 2006) was found to exhibited moderate antimicrobial activity against Bacillus subtilis, this suggests that the bacterium could potentially play a role in chemical defense.

4.5.2 Phylogenetic and genomic analysis of Ruegeria phage 67 The CG content of Ruegeria phage 67 was 58.87%, which is consistent with other roseophages (range of 55-64%) and the high CG content of marine roseobacter hosts (Rohwer et al., 2000, Chan et al., 2014). A total of 32 roseophages have been isolated so far, however only 11 of these phages contained the DNA polymerase gene and could be included in a phylogenetic analysis. Ruegeria phage 67 was most closely related to Ruegeria phage DSS3-P1 (Siphoviridae, host Ruegeria pomeroyi DSS-3), Silicibacteria phage DSS3-P1 (unclassified dsDNA phage, host Ruegeria pomeroyi DSS- 3) and Synechococcus virus S-ESS1 (host Synechococcus sp. SJ01) (Han et al., 2017). Currently, only genome information for Ruegeria phage DSS3-P1 and Silicibacteria phage DSS3-P1 are available through NCBI. So far, no studies have investigated the phage genome or phage-host dynamic of these two phages. Synechococcus virus S- ESS1 virus was recently sequenced and was found to be distantly related to the eight known cyanosiphoviruses and was closely related to E. coli bacteriophage T7 and Ruegeria phage DSS3-P1, and thus was suggested to be a new subtype of cyanosiphovirus (Han et al., 2017). However, the phage-host dynamics of Synechococcus virus S-ESS1 are yet to be determined.

The majority of protein sequences of Ruegeria phage 67 were most closely related to those from viruses in the Siphoviridae family, but few proteins also matched those from the viral families Myoviridae and Podoviridae (Appendix Table S4.2). This is consistent with several studies that revealed phages often containing proteins similar to those from multiple viral families (Hendrix et al., 2002, Allison et al., 2002, Recktenwald and Schmidt, 2002, Zhan et al., 2016, Han et al., 2017). For example, Roseobacter phage SIO67 had genes with high sequence similarities to podoviruses and siphoviruses (Zhan et al., 2016). Hendrix et al. (2002) proposed that all dsDNA phage genomes are a “mosaic” with access, by horizontal exchange, to a large genetic

84 pool with varying or selective capacities. This suggests that phages can acquire genes from phylogenetically distant viruses (Hendrix et al., 2000). Interestingly, Ruegeria phage 67 contained proteins with high sequence similarity to bacterial proteins, including those belonging to the Rhodobacteraceae and α-proteobacteria (Appendix Table S4.2). High sequence similarity was observed for the protein thymidylate synthase ThyX between the Ruegeria phage 67 and its host, indicating horizontal gene transfer (Appendix Table S4.2). ThyX catalyses the reaction of dUMP to dTMP (thymidylate), which is an essential DNA precursor (Graziani et al., 2004), and acquisition of the enzyme by the virus could aid its DNA replication by providing higher concentrations of deoxynucleotides (dNTPs) (Myllykallio et al., 2002, Graziani et al., 2004).

4.5.3 The life-cycle of Ruegeria phage 67 Putative structural proteins of the phage were identified, such as those with similarity to the major capsid protein E and bacteriophage lamda head decoration protein, which are involved in the assemly of the icosahedral capsids, conistent with the transmission electron microscopy observations. A putative phage tail protein and tape measure domain-containing protein found in the phage likely dictate the tail length and facilitate DNA transport to the bacterial cell cytoplasm during infection (Mahony et al., 2016). The phage genome also encoded a ortholog to the phage protal protein, lambda family, which functions to form a hole (or portal) that enables the passage of DNA during packaging and ejection (Lebedev et al., 2007). A pgW protein was detected, which is thought to be involved in the attachment of the tail to the capsid head and is suggested to interact with the DNA, forming a plug at the connector (site of head and tail attachment) to prevent ejection of the DNA (Maxwell et al., 2001).

Proteins associated with viral genome replication were detected including DNA polymerase (involved in DNA replication) and phage terminase (involved in viral DNA packaging) (Feiss and Catalano, 2005). The majority of the phage’s structural proteins and the proteins involved in DNA replicaiton had high sequence similarities to Ruegeria phage DSS3-P1 and Silicibacteria phage DSS3-P1 (Figure 4.5 and Figure 4.6). A chromosome segregation protein SpoOJ (stage 0 sporulation protein J, a member of the ParB superfamily) was also detected and is thought to be used by viruses to

85 disentangle and resolve multiple phage genomes during phage DNA replication (Chelikani et al., 2014, Chen et al., 2015). The genome also encodes for a ribonucleotide-diphosphate reductase that belongs to the pyrimidine metabolism pathway and is involved in the conversion of the ribose sugar of RNA into the deoxyribose sugar of DNA and is essential for DNA biosynthesis and repair (Uhlin and Eklund, 1994). Those proteins may be involved in increasing DNA biosynthesis for virion production (Uhlin and Eklund, 1994).

The phage genome also encoded proteins that may be involved in modulating cellular host functions, such as a SNF2 N-terminal domain-containing protein, von Willebrand factor type A domain and subtilase. SNF2 N-terminal domain is found in proteins involved in a variety of processes, including transcriptional regulation, DNA repair, DNA recombination and chromatin unwinding (Eisen et al., 1995). Von Willebrand factor type A domain are found in various plasma proteins and other extracellular proteins (in all eukaryotes) and are involved in multiple functions such as transcription, DNA repair, ribosomal and membrane transport and the proteasome (Ruggeri and Ware, 1993). Subtilase (belongs to the family of subtilisin-like serine proteases) are enzymes that cleave peptide bonds in proteins (Siezen and Leunissen, 1997) and thus may play a role in energy production via the breakdown of amino acids or modulation of the host cell functions (Vincent et al., 2003, Paton and Paton, 2010).

Proteins involved in host infection and virulence were also observed, such as virulence- associated protein E, P-loop domain and putative peptidoglycan binding domain, which is found in a variety of enzymes involved in bacterial cell wall degradation (Krogh et al., 1998). This protein may potential play a role in phage abosorption (i.e. infection of host cells) or lysis of bacterial cells to realease virions (Krogh et al., 1998).

The long incubation period of the initial viral plaque assay (approximately six months) suggests that the phage-host dynamic may be lysogenic and the production of virions were induced by host starvation (Cenens et al., 2013). However, Ruegeria phage 67 did not contain an integrase gene, which is consistent with the phylogenetically related Synechococcus virus S-ESS1 and other lytic roseophages, including Roseosiphophage R5C (Yang et al., 2017), Roseophage SI01 (Angly et al., 2009) and Roseophage vB_DshP- R1 (Ji et al., 2015). In contrast, closely related Ruegeria phage DSS3-P1 and 86 Silicibacteria phage DSS3-P1 both contained an integrase gene suggesting a possible lysogenic lifestyle. No phage genes were observed in the host genome indicating that phage DNA did not integrated into the host chromosome. However, a phage integrase gene was found in the bacterial host, which had low sequence similarity (28% identity) to an integrase protein from the Clostridium phage phiCD211 (a prophage of Clostridioides difficile) (Garneau et al., 2017). This suggests that the phage could potentially utilise the host integrase protein to integrate its DNA into the bacterial chromosome (Grainge and Jayaram, 1999). Together with the initial delayed plaque formation and phage-host dynamics (discussed below), this indicates that the phage may have pseudolysogenic characteristics as a the lack of chromosomal integration is consistent with pseudolysogeny (Williamson et al., 2001). Interestingly, Lohr et. al (2005), isolated a phage from a sponge-associated α-proteobacteria, which also was postulated to have pseudolysogenic characteristics, such as positive lysogeny induction, and the presence of both DNA replication genes and integrase in the phage genome. Further studies such as prophage induction should be carried out to test whether the phage can utilise the host integrase to integrate its genome into the host chromosome.

Ruegeria phage 67 had a latent period of 4-6 hours, which was longer compared to other isolated phages infecting the Roseobacter clade (average latent period of 2-3 hours) (Zhao et al., 2009, Zhang and Jiao, 2009, Yang et al., 2017). Long latent periods are thought to be a useful strategy in a non- or slow-growing population to ensure that the host population is dense enough for phage progeny to infect neighbouring cells. This is also accompanied with an increase in burst size to ensure viral spread when more favourable conditions become available (Wommack and Colwell, 2000). Thus, temperate phages have been suggested to have long latent periods and a high burst size, whereas lytic phages exhibit a short latent period and low burst size (Stewart and Levin, 1984, Parada et al., 2006). However, the bust size of Ruegeria phage 67 (range of 22-34 PFUs/cell in different media) was lower compared to other lytic roseobacter phages. For example, phage DSS3φ2 (host Ruegeria pomeroyi DSS-3) and EE36φ1 (host Sulfitobacter sp. EE-36) had a burst size of 350 and 1500 PFUs/cell, respectively (Zhao et al., 2009), and phage RDJLφ1 (host Roseobacter denitrificans OCh114) had an

87 estimated burst size of 203 PFUs/cell (Zhang and Jiao, 2009). This suggests that the phage-host system is atypical to “regular” lysogenic or lytic infection dynamics and may again point towards a pseudolysogeny-like interaction (Williamson et al., 2001, Parada et al., 2006).

4.5.4 Morphological changes in host cells induced by viral infection Swelling in the host cells were observe after 15 min to 2 hours post infection (T0-T2) (Figure 4.5). This was also observed in a phage-FISH (fluorescent in situ hybridisation) study of Pseudoaltermonas sp. H100 and phage PSA-HP1, and is thought to be caused by the accumulation of virions within infected cells (Allers et al., 2013). Cellular debris can be seen after 4 hours post infection (T4), indicating bacterial cell lysis, which is earlier than the observed latent period via PFU counts (at T6). A delay in the observed increased in virion production via PFU counts compared to microscopy has also been reported in another phage-host infection study by Dang et al. (2015) (studying Cellulophage baltica strains and podovirus φ38:1). This observation could be explained by a long bacterial lysis period, where low numbers of virions are being released over time (Dang et al., 2015).

The drastic changes in the host cell size and shape observed here at 6-12 hours post infection have however not been reported in the literature before. The large increase in bacterial cell size was observed after cell lysis (first round of cell lysis observed in the viral infection microscopy was at T4) and indicates that this morphology could be induced by lysis of infected cells or a second round of viral infection. This morphological change could potential be a defense mechanism against the second wave of viral infection. Studies have reported that differences in cell morphology can affect viral infection (Mojica and Brussaard, 2014). For example, haploid cells of Emiliania huxleyi, which are flagellated, are more resistant to viral infection compare to diploid cells, which are non-motile and coccolith-bearing cells (Frada et al., 2008). Alternatively, the dramatic changes seen here might be a result of selecting for certain morphological variants/mutants that are resistant to viral infection.

The morphological changes seen here could potentially be a result of inhibition of cell segregation and continued cellular growth resulting in the filamentous formation and

88 increased in cellular size. Bacterial cell division is orchestrated by FtsZ, which polymerises to form a ring-like structure and act as a scaffold for the assembly of the bacterial cytokinetic machinery (Adams and Errington, 2009). FtsZ assembly is tightly regulated and a diverse repertoire of accessory proteins contributes to the formation of a functional division machinery (Adams and Errington, 2009). The phage-encoded subtilase could potential play a role in cleaving proteins essential for the formation of the Z ring assembly and thus inhibit cell division. Clearly, further work is needed to understand the underlying mechanisms for the morphological changes observed. The induced morphological changes could potential confer benefits to the phage, such as a continued resource for DNA replication and virion production in a prolonged “protected” cellular environment. Likewise, it could also confer benefits to the infected bacterial host by inducing cellular changes that provide resistance to phagocytosis by the sponge host. This suggests that the phage-host system may be a mutualistic interaction, enhancing the survival of both phage and bacterial host in the sponge environment.

Numerous studies have reported the role of phages in enhancing bacterial resistance to serum and phagocytes (van der Vijver et al., 1972, Kaneko et al., 1997, De Groote et al., 1997, Farrant et al., 1997). For example, phages have been reported to encode for CHIPS (a phagocytotoxin) (Wagner and Waldor, 2002), Panton-Valentine leucocidin (PVL, a cytotoxin with direct activity against human phagocytes) (Kaneko et al., 1997) and superoxide dismutase (De Groote et al., 1997, Farrant et al., 1997), which catalyses the conversion of superoxide ion into hydrogen peroxide and molecular oxygen and is thus thought to aid bacteria to survive the oxidative stress inside phagocytes. The morphological increase in bacterial cell size (filamentous) and pear shaped (average of 10-20 µm in length and 2-5µm in width) observed here may enhance bacterial resistance to sponge amoebocytes and/or protozoans (Pernthaler, 2005, Justice et al., 2008). Filamentous bacterial symbionts have been reported in sponges, which appeared to be resistance to phagocytosis (Schmidt et al., 2000, Hentschel et al., 2002). Similarly, filamentous prey have been reported to be resistant to protist predation in marine environments (Pernthaler, 2005). Generally, filamentous bacteria that are larger than 7µm in length are considered inedible by marine protists

89 (Jürgens and Matz, 2002). This feeding size restriction is thought to be related to the kinetics of phagocytosis or due to the size of the phagocytic vesicles (Pernthaler, 2005). Further investigations such as phagocytosis assays are needed to prove that infected cells with the morphological changes are indeed resistance to phagocytosis.

4.5.5 Development of phage resistance In bacterial populations there are usually a small number of genetic variants that are naturally resistant to phage attack. Fully or partially phage-resistant bacterial strain/s can often evolve and increase in frequency within experimental microcosms and thus increase the complexity of phage-host dynamics (Bohannan and Lenski, 1999). Similarly, we observed an increased frequency of cells resembling non-infected cells after 6 hours post infection (>T6), suggesting they may be fully resistant to phage infection. Phage resistance in bacteria are thought to be commonly caused by alterations/mutations in cell membrane proteins (phage receptors) that prevent the absorption of phages (Bohannan and Lenski, 2000). This has also been reported as a resistance mechanism against phage in roseobacter (Zhang et al., 2009, Huang et al., 2010). For example, a phage-resistant mutant strain of Roseobacter denitificans OCh114 (M1) was found to have five membrane proteins (hypothesised to be potential phage receptors) were downregulated compared to the sensitive strain (Huang et al., 2010). Similarly, a phage-resistant mutant of roseobacter Ruegeria pomeroyi DSS-3 was found to have protein modifications in four highly expressed proteins of unknown functions, (Zhang et al., 2009).

4.5.6 Impact of sponge extracts on the phage-host dynamics Viruses are reliant on their host to provide the cellular machinery and the necessary energy and resources required for viral replication and assembly. Consequently, factors that regulate the physiology of the host, as well as its proliferation are also important in governing virus-host dynamics (Mojica and Brussaard, 2014). Virion production was linked to bacterial growth, observed in the infection culture in marine broth (bacterial generation time of 49 min and phage burst size of 22.3 PFUs/cell). Infected cultures supplemented with Scopalina sp. and C. concentrica sponge extracts showed a slight increase in bacterial growth (generation time of 45 min and 47 min, respectively) which corresponded to an increase in viral burst size of 33.9 PFUs/cell

90 and 27.7 PFUs/cell, respectively. This may be due to the increased amount of organic and inorganic nutrients in the added sponge extracts. This is consistent with the study by (Motegi and Nagata, 2007) where the addition of nitrogen or nitrogen plus carbon in subtropical surface waters resulted in an increased in the bacteria growth rates and subsequent increased in virion production.

Intriguingly, sponge extracts from T. anhelans inhibited bacterial growth and thus also virion production which appeared to be host-specific as sponge extracts from the co- occurring sponges C. concentrica and Scopalina sp. had no effect. This is ecologically significant as it indicates that sponges may have specific mechanisms to modulate bacterial growth and thus indirectly prevent the death or lysis of its symbionts. This mechanism may potentially be employed by the sponge host to help maintain a stable microbial community.

4.6 Conclusion

The Red Queen hypothesis attempts to explain that competitive interactions will lead to continuous variation and selection towards adaptation of the host, and counter- adaptation on the side of the parasite (Weinbauer, 2004, Clokie et al., 2011, Hargreaves et al., 2014). This evolutionary trend is arguably most evident in the arms race between phages and bacteria and has implications for the evolution of virulence in pathogens (Brüssow et al., 2004). Viruses and their infection of bacteria and eukaryotes causing cell lysis is believed to have a substantial impact on global nutrient cycling through what has been termed the “viral shunt” (Suttle, 2007), and on the evolution of the biosphere through phage-mediated horizontal transfer (Comeau and Krisch, 2005).

The phage-host interaction studied here displayed characteristics of pseudolysogeny, which is consistent with the only other report of a sponge-associated phage φJL001 of an α-proteobacterium (Lohr et al., 2005). There is growing evidence that pseudolysogeny is widespread among divergent environments and has been suggested that it affords “phage populations with a means of quickly reacting to environmental changes” (Wommack and Colwell, 2000). This may be an ecological adaptation to

91 survive in the dynamic sponge environment (Maldonado et al., 2012). Lysogeny and pseudolysogeny have important environmental and ecological consequences, increasing the likelihood of both phage and host survival. As far as we are aware, this is the first report of extreme bacterial morphological changes induced by viral infection. This may be caused by the bacterial host’s response against further infection i.e. alterations in cell morphology to prevent viral absorption. Alternatively, filamentous and increased cellular size may enhance the bacterial resistance to phagocytosis in the sponge environment. This last potential explanation would indicate that the phage- host system may be mutualistic where both the phage and bacterial host benefit. If so, this indicates an intriguing complex co-evolution of phage-bacteria-sponge interaction that warrants further research.

Environmental factors have been reported to impact virus-host dynamics, such as temperature, pH, salinity and the presence/absence of nutrients, such as carbon, nitrogen and phosphate (Mojica and Brussaard, 2014). Here, we demonstrated that the ecology of the host environment can impact virus-host dynamics, which differs greatly from standard laboratory conditions. This highlights the increasing intricacy and complexity of interactions within the sponge holobiont. This unique phage-host system may provide an interesting model to study factors that help to maintain the balance of diverse sponge microbial communities.

Chapter Five

92 5 Conclusions and future directions

5.1 Summary

Marine sponges play a crucial role in the benthic ecosystem through nutrient cycling and the provision of habitats for a range of fauna and flora (Bell, 2008). The health and function of sponges are closely tied to their diverse community of symbiotic microorganisms and environmental factors, such as climate change, which can influence the symbiotic community composition and structure potential resulting in sponge mortality or disease (Webster, 2007, Fan et al., 2013, Webster and Thomas, 2016). Throughout the past decade, there has been many advances in our understanding of the diversity and to a lesser extent the function of sponge-associated microbes (Webster and Taylor, 2012, Thomas et al., 2016). However, these studies have mainly focused on bacterial symbionts, while very few studies have investigated sponge-associated fungi and viruses (He et al., 2014, Naim et al., 2017, Laffy et al., 2018). This thesis aimed to provide novel insights and to build on the knowledge in these largely understudied areas in sponge ecology. Additionally, we aimed to elucidate some of the functions that may be carried out by fungi and viruses and their potential ecological roles in sponges.

Chapter two explored fungal diversity, host-stability and host-specificity of three co- occurring sponge species. To comprehensively assess fungal diversity, both traditional cultivation and molecular methods were employed. Our results showed that the cultivation method captured only approximately 10% of fungal diversity compared to ITS amplicon sequencing. Community-wide profiling using ITS amplicon sequencing revealed that sponges, C. concentrica, Scopalina sp. and T. anhelans had relatively low fungal diversity compared to the bacterial and archaeal communities reported in these sponges previously (Fan et al., 2012, Esteves et al., 2016). The fungal communities in the sponges studied here, had low host-specificity and broadly reflected the seawater

93 community. Highly similar fungal species have been detected in sponges around the world, which suggests a prevalence of horizontal transmission, where selection and enrichment of some fungi may occur for those that can survive and/or exploit the sponge environment.

There is evidence to suggest that fungi have various interactions with sponges, including parasitic, mutualistic and/ or commensal. The presences of many saprophytic fungi in C. concentrica, Scopalina sp. and T. anhelans indicate that they may be able to exploit the nutrient rich environment of the sponge host. Alternatively, they may play a role in enhancing nutrient uptake by aiding in the breakdown of plant-derived detritus or plankton filtered from seawater. Some fungi detected in our study were related to previously isolated fungal species, which have been reported to produce a range of secondary metabolites (Holler et al., 2000, Henríquez et al., 2014). This suggest that some sponge-associated fungi may play a role in host defense. Furthermore, common plant fungal pathogens were detected in (apparently healthy) sponge samples studied here. Similarly, other studies have also detected fungal and microbial pathogens of corals in sponges (Ein-Gil et al., 2009, Negandhi et al., 2010). This indicates that sponges could potential act as reservoirs of marine and terrestrial pathogens.

Previous studies on microbial communities of sponges have indirectly suggested that there may be high viral loads in sponges due to the presence and high abundances of CRISPRs/Cas proteins and restriction modification systems (Fan et al., 2012, Horn et al., 2016, Karimi et al., 2017). A recent study by Laffy et. al. (2018) directly investigated the diversity, relative abundance and functional gene analysis of viromes associated with the free-viral fraction of coral and sponges (Laffy et al., 2018). Chapter three thus aimed to provide insights into the diversity, abundance and functional analysis of viral assemblages of sponges focusing on the microbial cell fraction. Additionally, the expression of “active” (transcribed) viruses were also investigated. Viral assemblages in sponge metagenomes were dominated by bacteriophages from the order Caudovirales, whereas, eukaryotic viruses (Phycodnaviridae and Pandoraviruses) were highly expressed in the viral metatranscriptomes. Sponges appeared to contain both variable environmentally-derived and sponge-specific viral consortia with some sponge

94 species containing up to 60% overlap of viral species in the seawater community. Functional gene analysis revealed common functions, such as DNA replication and virion assembly in all samples investigated here. Additionally, sponge viral communities contained diverse auxiliary metabolic genes that were specific to different host environments. For example, genes associated with thiamine metabolism and biosynthesis of antibiotics were only found in C. concentrica, while genes associated with photosystem were only observed in Stylissa sp. 445 and aegerolysin genes were only detected in Scopalina sp..

Taxonomical and functional analysis of the sponge viromes indicated that there may be dynamic virus-host interactions associated with the sponge microbial cell fraction, including lytic and lysogenic infections. Metatranscriptome analysis detected “active” expression of viruses infecting marine and terrestrial bacterial pathogens, providing further evidence that sponges may act as reservoirs for viruses of pathogenic bacteria, which may influence the spread and/or growth of pathogens in the environment.

Chapter four aimed to establish a virus-host model system for sponges. A novel phage termed Ruegeria phage 67 was isolated from the sponge-associated Ruegeria sp. AU67. TEM and genome analysis of the phage showed that it belonged to the Siphoviridae family and was most closely related to roseophages Ruegeria phage DSS3- P1 and Silicibacter phage DSS3-P1. Genomic analysis of the phage and host along with infection growth dynamics suggested that the phage was pseudolysogenic. This is consistent with the only other reported sponge-associated phage-host system of an α- proteobacterium and phage φJL001 (Lohr et al., 2005). Pseudolysogeny is thought to allow phage populations to quickly react to environmental change (Łoś and Węgrzyn, 2012). This may be an important ecological adaptation to survive the dynamic sponge environment (Maldonado et al., 2012). Furthermore, phage infection induced extreme morphological changes in the bacterial host, such as filamentation and increased in cell size, which may enhance the bacteria’s resistance to phagocytosis in the sponge environment. Thus, the pseudolysogeny-like interaction may be beneficial to both phage and host, increasing the likelihood of survival of both parties within the sponge environment. Lastly, we demonstrated that the ecology of the sponge can impact the virus-host dynamics. Media supplemented with sponge extracts from T. anhelans,

95 which is the isolation host of Ruegeria sp. AU67 and phage, inhibited bacterial growth and subsequently also virion production, whereas sponge extracts from the co- occurring sponges C. concentrica and Scopalina sp. had no effect. This indicated that there may be host-specific mechanisms that modulate bacterial growth and thus indirectly prevent viral infection and possibly death of its symbionts. This may be an important mechanism by which sponges maintain a stable microbial community.

5.2 The sponge holobiont

Macroorganisms, such as animals, plants and macroalgae, are increasingly recognised as holobionts, that is, a complex ecosystem of the host, the microbiota and the interactions between them (Margulis, 1991, Guerrero et al., 2013). To define the sponge holobiont it is necessary to characterise the core microbial taxa and the core set of functional genes that ensures homeostasis of the holobiont (Qin et al., 2010, Tanca et al., 2017). The holobiont functions by not only the processes carried out by individual members, but also the interactions among them. Thus, the sponge holobiont can be regarded as a complex ecosystem, where the microbiome provides essential functions to the host and together they mediate the interactions of the sponge holobiont with the surrounding community (Sommer et al., 2017, Adair and Douglas, 2017). Consequently, the microbiome can have cascading effects on the health and functioning of sponges and the entire reef ecosystem (Pita et al., 2018).

Global surveys of sponges along different environmental gradients, such as geographical distance, season, depth and habitat have consistently shown that sponges harbour species-specific and stable microbial communities (Erwin et al., 2012, Pita et al., 2013b, Pita et al., 2013a, Zhang et al., 2014, Thomas et al., 2016, Pita et al., 2018), This is remarkable considering the dynamic bacterioplankton communities in the surrounding water, in which the sponges filter feed (Erwin et al., 2015). Despite the high variability of the viral communities in sponge individual replicates (Chapter three), the microbial communities were consistent (Fan et al., 2012). This hints to the importance of host-related factors in shaping and maintain the core microbiome (Pita et al., 2018). Further evidence for this was seen in Chapter four, where sponge-species

96 specific bacterial growth and virion production was inhibited in infection cultures supplemented with sponge extracts from the host.

Many theories have been proposed on the evolution of symbiont-host interactions including 1) an ancient symbiosis maintained by vertical transmission, 2) parental and environmental; symbiont transmission and 3) environmental acquisition (Wilkinson, 1984, Wilkinson et al., 1984, Müller and Müller, 2003, Usher et al., 2005). However, little is known about the mechanisms and/ or factors by which sponges maintain a stable diverse microbial community. One theory is that sponges are “ecosystem engineers” (Coyte et al., 2015) and provide a certain habitat that selects for the presence and persistence of certain microorganisms. This has been suggested in Chapter two, where certain fungal saprotrophs were detected in sponges and may represent those that are able to exploit the nutrient-rich sponge environment. Another theory has been postulated that sponges are able to control their microbial residents by specifically recognising and differentiating between foreign and symbiotic microbes (Wilkinson et al., 1984, Wehrl et al., 2007), likely through the innate immune system (Chu and Mazmanian, 2013) (discussed further below). However, the underlying molecular mechanisms of microbial recognition by sponges are yet to be experimentally determined as establishment of sponge symbiont models remains elusive (Schippers et al., 2012, Esteves et al., 2016, Pita et al., 2016). Our bacterial isolate Ruegeria sp. AU67 may be a good candidate model to investigate the mechanisms that influence the growth of sponge-associated bacteria and thus potentially provide insights into bacterial population control and maintenance in the sponge holobiont.

5.3 Molecular mechanisms of sponge-symbiosis interactions

Two main theories have been proposed to explain how sponges establish and maintain sponge-microbe associations: 1) sponges are able to specifically recognise between symbionts and “food” bacteria through their innate immune system or 2) microbes are able to avoid detection by sponge cells or modulate host cell functions to survive and thrive inside the sponge environment. Evidence to support both theories have been

97 reported and suggests that the establishment and maintenance of the microbial communities may result from a synergistic combination of both mechanisms. That is, some sponge-microbial interactions may have been established through theory one, while others may have arisen from theory two.

Early feeding studies have reported that seawater-derived bacteria were consumed at a much higher rate compared to sponge-derived bacteria suggesting that sponges are able to differentiate between “food” bacteria and symbionts (Wilkinson et al., 1984, Wehrl et al., 2007). Furthermore, sponges have been shown to elicit different immune responses against gram-negative and -positive bacteria (Boehm et al., 2001, Müller and Müller, 2003, Wiens et al., 2005, Thakur et al., 2005), indicating that sponges can specifically recognise different bacteria. For example, the sponge Suberites domuncula has been shown to increase production of compounds with antibacterial activity when exposed to gram-negative bacterial endotoxin lipopolysaccharides (LPS) (Müller et al., 2004), while when exposed to gram-positive LPS, endocytosis followed by the release of lysozymes was activated (Thakur et al., 2005). Recently, high-throughput sequencing data have revealed that sponges harbour a complex genomic repertoire of diverse immune receptors, including Toll- and NOD-like receptors, scavenger receptor cysteine-rich (SRCR) family members (Srivastava et al., 2010, Hentschel et al., 2012, Degnan, 2015, Ryu et al., 2016) as well as receptors for fungi (Perović‐Ottstadt et al., 2004) and viruses (Wiens et al., 1999). The Toll-like receptor pathway, such as MyD88, have been found to be involved in the sponges’ (S. domuncula) response to signals from different bacterial species (Wiens et al., 2005). These studies suggest that sponges can actively recognise and discriminate specific microbes via their immune systems.

Different molecular mechanisms have been speculated to help microbes avoid detection and/ or contain mechanisms that may allow them to persist inside the sponge host. These mechanism include slime layers or sheaths (Wilkinson, 1978, Friedrich et al., 1999), modification of lipopolysaccharides (Wehrl et al., 2007, Burgsdorf et al., 2015) and eukaryotic-like proteins domains (Thomas et al., 2010, Liu et al., 2012, Nguyen et al., 2014, Díez‐Vives et al., 2017). The cyanobacteria symbiont “Candidatus Synechococcus spongiarum” was reported to lack a lipopolysaccharide

98 antigen compared to the free-living Synechococcus (Burgsdorf et al., 2015). This modification was speculated as a mechanism to avoid detection from sponge predation and phage attack. Eukaryotic-like protein domains (ELPs), such as ankyrin repeat proteins, tetratricopeptide repeat proteins, and leucine-rich repeat proteins, were found to be highly enriched and expressed in sponge symbionts (Thomas et al., 2010, Díez‐Vives et al., 2017). Ankyrin repeat proteins from a sponge symbiont have also been demonstrated to retard phagocytosis in amoeba (Nguyen et al., 2014). ELPs mediate protein-protein interactions and are hypothesis to be involved in establishing an intracellular lifestyle in the sponge environment (Webster and Thomas, 2016).

Here (Chapter four), we propose that viruses could potential play a role in the establishment or prolonged persistence of bacteria in the sponge. Viral infection was shown to cause morphological changes in the bacterial host, which may enhance its resistance to phagocytosis by sponge amoebocytes. We speculate that this potentially mutualistic infection strategy may confer survival advantages to both the bacteria and phage. Viral infection may also be an effective mechanism in influencing microbial community population and dynamics (through cell lysis) (Weinbauer and Rassoulzadegan, 2004, Thurber et al., 2017). Additionally, the sponge host was able to specifically inhibit the growth of bacteria and thus subsequently virion production. This provides further evidence that sponges can specifically recognise bacteria and respond appropriately to maintain a healthy core microbiome. However, the factors that underpin this mechanism remains to be determined. The potential role that viruses may play in the establishment and maintenance of microbial communities adds another layer of complexity to the sponge holobiont.

5.4 Implications of sponges as reservoirs of pathogens in global epidemiology

Climate change has been implicated in the recent increased prevalence and severity of disease outbreaks in marine ecosystems (Webster, 2007, Harvell et al., 2009, Angermeier et al., 2012, Gochfeld et al., 2012, Easson et al., 2013, Olson et al., 2014, Lough et al., 2018). These influences likely results in the expansion of pathogen ranges in response to warming, changes to host susceptibility as a result of increasing

99 environmental stresses, and the expansion of potential disease vectors (i.e. a mobile organism that transmits a parasite from one host to another) (Hoegh-Guldberg and Bruno, 2010, Gochfeld et al., 2012). Over the past 30 years, increasing occurrences of disease in marine species and communities have been reported (Ward and Lafferty, 2004). Modular and colonial life forms, such as sponges, corals, bryozoans and ascidians, may facilitate a build-up of more virulent strains due to the genetic homogeneity of the hosts (McCallum et al., 2004). Additionally, the relatively rapid evolution of pathogens compared with hosts may further facilitate the epidemic spread of virulent pathogens (McCallum et al., 2004). Phages can affect pathogenesis by 1) encoding for virulence factors, such as exotoxins, which can be passed onto different bacterial host/s through horizontal gene transfer, and 2) by contributing directly to pathogenesis at the time of infection, such as by encoding for regulatory factors that increase the expression of virulence genes in the host (Spanier and Cleary, 1980, Mavris et al., 1997, Guan et al., 1999). Sponges are host to diverse viral and microbial communities, which could be a hotspot for the spread of virulence factors through phage-mediated horizontal gene transfer across different organisms and this could potentially create new pathogens (Sano et al., 2004, Brabban et al., 2005, Woolhouse et al., 2005, Stewart et al., 2008, Buerger and van Oppen, 2018).

Negandhi et al. (2010) detected coral disease-associated microbes in the Florida reef sponges Agelas tubulata and Amphimedon compressa. Additionally, the causative fungal pathogen Aspergillus sydowii of gorgonian sea fan corals was found in apparently healthy Spongia obscura individuals (Ein-Gil et al., 2009). Sponges were speculated to act as symptomless carriers of marine pathogens and may play a role in the spread of fungal propagules after fungal growth and conidiation (a biological process in which filamentous fungi produce asexually from spores), fragmentation of living sponge tissues or after the sponges’ death (Smith et al., 1998, Philippa et al., 2004, Ein-Gil et al., 2009). Our studies (Chapter two and three), detected both terrestrial and marine fungal pathogens and pathogenic viruses in sponges. The coastal geographic location of many sponge species may provide a potential route between terrestrial habitats and the marine environment. Sponges may retain some of these pathogens by filter feeding, providing a more “sheltered” habitat whereby certain

100 terrestrial pathogens are able to survive and over time adapt to the marine environment (Shearer et al., 2007). Thus “sponge hopping” may be a mechanism in which terrestrial pathogens may survive and disperse in the aquatic environment (Ein- Gil et al., 2009). Analysis of the interaction between host animals and the coastal watershed, as well as the survival, prevalence and proliferation of pathogens in marine sponges are an important area of concern for disease emergence (Stewart et al., 2008).

5.5 Future directions

The healthy sponge holobiont can be considered as an ecosystem that is in a state of dynamic equilibrium (Pita et al., 2018). The interactions among the members of the holobiont may be affected by perturbations causing an imbalance to the healthy equilibrium potentially leading to dysbiosis and disease (Egan and Gardiner, 2016). Future research in the area of sponge ecology should therefore aim to adopt a “holobiont approach”, which involves defining and determining the core microbiome, including the understudied eukaryotic and viral communities. To understand the healthy function of the sponge holobiont, we need to investigate the relationship between the sponge host and its entire microbial community, including eukaryotic, prokaryotic and viral symbionts. This will also incorporate the study of the interaction and function of microbial symbionts in the sponge, how the microbial communities are maintained and to determine the mechanisms that underpin these sponge-microbial interactions.

In this study, we have elucidated some insights into the diversity of lesser studies microbial symbionts associated with sponges, namely fungi and viruses, but future studied should also explore other eukaryotic organisms such as protozoans, algae and/or diatoms and archaea. This can be achieved through deep sequencing of sponges from diverse geographies, morphologies and phylogenetic relationships to gather insights into the holobiont of different sponge species.

To gather insights on the interaction and potential function of symbionts in sponges, this could be tackled through the “omics” or “cultivation” approaches. Metagenome,

101 (meta)transcriptome and proteomics can provide insights on functional genes encoded by organisms and/or certain microbial communities and help elucidate potential metabolic interactions between host and symbionts. Traditional cultivation of symbionts would allow for physiochemical and metabolic analysis, such as the production of secondary metabolites.

The balance of a healthy holobiont in sponges is extremely interesting and little is known about the mechanisms of this interaction. Because of the complex nature of the sponge holobiont, it is very difficult to tease out specific interactions between the host and its microbial community. One approach is to study changes in the sponge holobiont when it undergoes stresses or in disease and compare this with healthy samples. Through these experiments we may elucidate key members or functions that are important for a healthy holobiont.

Lastly, the function of sponges as “ecosystem engineers” and their potential roles as a reservoir for marine and terrestrial pathogens is a new concept that has global ecological implications on the health of reef ecosystems and pathogenesis. Further studies in this area could involve monitoring and sampling sponges over time and when disease outbreaks occur.

References

ABARENKOV, K., HENRIK NILSSON, R., LARSSON, K. H., ALEXANDER, I. J., EBERHARDT, U., ERLAND, S., HØILAND, K., KJØLLER, R., LARSSON, E. & PENNANEN, T. 2010. The UNITE database for molecular identification of fungi–recent updates and future perspectives. New Phytologist, 186, 281-285. ABBASIFAR, R., GRIFFITHS, M. W., SABOUR, P. M., ACKERMANN, H.-W., VANDERSTEEGEN, K., LAVIGNE, R., NOBEN, J.-P., VILLA, A. A., ABBASIFAR, A. & NASH, J. H. 2014. Supersize me: Cronobacter sakazakii phage GAP32. Virology, 460, 138-146. ADAIR, K. L. & DOUGLAS, A. E. 2017. Making a microbiome: the many determinants of host- associated microbial community composition. Current opinion in microbiology, 35, 23- 29. ADAMS, D. W. & ERRINGTON, J. 2009. Bacterial cell division: assembly, maintenance and disassembly of the Z ring. Nature Reviews Microbiology, 7, 642. ADRIAENSSENS, E. M., VAN ZYL, L., DE MAAYER, P., RUBAGOTTI, E., RYBICKI, E., TUFFIN, M. & COWAN, D. A. 2015. Metagenomic analysis of the viral community in Namib Desert hypoliths. Environmental microbiology, 17, 480-495.

102 ALLERS, E., MORARU, C., DUHAIME, M. B., BENEZE, E., SOLONENKO, N., BARRERO‐CANOSA, J., AMANN, R. & SULLIVAN, M. B. 2013. Single‐cell and population level viral infection dynamics revealed by phageFISH, a method to visualize intracellular and free viruses. Environmental microbiology, 15, 2306-2318. ALLISON, G. E., ANGELES, D., TRAN-DINH, N. & VERMA, N. K. 2002. Complete genomic sequence of SfV, a serotype-converting temperate bacteriophage of Shigella flexneri. Journal of bacteriology, 184, 1974-1987. ALTSCHUL, S. F., GISH, W., MILLER, W., MYERS, E. W. & LIPMAN, D. J. 1990. Basic local alignment search tool. Journal of molecular biology, 215, 403-410. ALTSCHUL, S. F., MADDEN, T. L., SCHAFFER, A. A., ZHANG, J., ZHANG, Z., MILLER, W. & LIPMAN, D. J. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res, 25, 3389-3402. AMEND, A. S., BARSHIS, D. J. & OLIVER, T. A. 2012. Coral-associated marine fungi form novel lineages and heterogeneous assemblages. The ISME journal, 6, 1291-1301. ANAND, T. P., BHAT, A. W., SHOUCHE, Y. S., ROY, U., SIDDHARTH, J. & SARMA, S. P. 2006. Antimicrobial activity of marine bacteria associated with sponges from the waters off the coast of South East India. Microbiological research, 161, 252-262. ANDERSON, I. C., CAMPBELL, C. D. & PROSSER, J. I. 2003. Potential bias of fungal 18S rDNA and internal transcribed spacer polymerase chain reaction primers for estimating fungal biodiversity in soil. Environmental Microbiology, 5, 36-47. ANDERSON, M. J. 2001. A new method for non-parametric multivariate analysis of variance. Austral Ecology, 26, 32-46. ANDERSON, M. J. 2006. Distance‐based tests for homogeneity of multivariate dispersions. Biometrics, 62, 245-253. ANDERSON, M. J., ELLINGSEN, K. E. & MCARDLE, B. H. 2006. Multivariate dispersion as a measure of beta diversity. Ecology letters, 9, 683-693. ANGERMEIER, H., GLÖCKNER, V., PAWLIK, J. R., LINDQUIST, N. L. & HENTSCHEL, U. 2012. Sponge white patch disease affecting the Caribbean sponge Amphimedon compressa. Diseases of aquatic organisms, 99, 95-102. ANGLY, F., YOULE, M., NOSRAT, B., SRINAGESH, S., RODRIGUEZ‐BRITO, B., MCNAIRNIE, P., DEYANAT‐YAZDI, G., BREITBART, M. & ROHWER, F. 2009. Genomic analysis of multiple Roseophage SIO1 strains. Environmental microbiology, 11, 2863-2873. ANGLY, F. E., FELTS, B., BREITBART, M., SALAMON, P., EDWARDS, R. A., CARLSON, C., CHAN, A. M., HAYNES, M., KELLEY, S. & LIU, H. 2006. The marine viromes of four oceanic regions. PLoS biology, 4, e368. BADGER, M. R., PRICE, G. D., LONG, B. M. & WOODGER, F. J. 2005. The environmental plasticity and ecological genomics of the cyanobacterial CO2 concentrating mechanism. Journal of experimental botany, 57, 249-265. BAKER, P. W., KENNEDY, J., DOBSON, A. D. & MARCHESI, J. R. 2009. Phylogenetic diversity and antimicrobial activities of fungi associated with Haliclona simulans isolated from Irish coastal waters. Marine biotechnology, 11, 540-547. BANKEVICH, A., NURK, S., ANTIPOV, D., GUREVICH, A. A., DVORKIN, M., KULIKOV, A. S., LESIN, V. M., NIKOLENKO, S. I., PHAM, S. & PRJIBELSKI, A. D. 2012. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. Journal of computational biology, 19, 455-477. BARR, J. J., AURO, R., FURLAN, M., WHITESON, K. L., ERB, M. L., POGLIANO, J., STOTLAND, A., WOLKOWICZ, R., CUTTING, A. S. & DORAN, K. S. 2013. Bacteriophage adhering to mucus provide a non–host-derived immunity. Proceedings of the National Academy of Sciences, 110, 10771-10776. BARRANGOU, R., FREMAUX, C., DEVEAU, H., RICHARDS, M., BOYAVAL, P., MOINEAU, S., ROMERO, D. A. & HORVATH, P. 2007. CRISPR Provides Acquired Resistance Against Viruses in Prokaryotes. Science, 315, 1709-1712. 103 BAYER, K., SCHMITT, S. & HENTSCHEL, U. 2008. Physiology, phylogeny and in situ evidence for bacterial and archaeal nitrifiers in the marine sponge Aplysina aerophoba. Environmental Microbiology, 10, 2942-2955. BEGLEY, T. P., XI, J., KINSLAND, C., TAYLOR, S. & MCLAFFERTY, F. 1999. The enzymology of sulfur activation during thiamin and biotin biosynthesis. Current opinion in chemical biology, 3, 623-629. BELL, J. J. 2008. The functional roles of marine sponges. Estuarine, Coastal and Shelf Science, 79, 341-353. BERNE, S., LAH, L. & SEPČIĆ, K. 2009. Aegerolysins: structure, function, and putative biological role. Protein Science, 18, 694-706. BINDAL, G., KRISHNAMURTHI, R., SESHASAYEE, A. S. N. & RATH, D. 2017. CRISPR-Cas-Mediated Gene Silencing Reveals RacR To Be a Negative Regulator of YdaS and YdaT Toxins in Escherichia coli K-12. mSphere, 2, 483-517. BOEHM, M., HENTSCHEL, U., FRIEDRICH, A., FIESELER, L., STEFFEN, R., GAMULIN, V., MUELLER, I. & MUELLER, W. 2001. Molecular response of the sponge Suberites domuncula to bacterial infection. Marine Biology, 139, 1037-1045. BOHANNAN, B. J. & LENSKI, R. E. 1999. Effect of prey heterogeneity on the response of a model food chain to resource enrichment. The American Naturalist, 153, 73-82. BOHANNAN, B. J. & LENSKI, R. E. 2000. Linking genetic change to community evolution: insights from studies of bacteria and bacteriophage. Ecology letters, 3, 362-377. BOLLIVAR, D. W., BERNARDONI, B., BOCKMAN, M. R., MILLER, B. M., RUSSELL, D. A., DELESALLE, V. A., KRUKONIS, G. P., HATFULL, G. F., CROSS, M. R. & SZEWCZYK, M. M. 2016. Complete genome sequences of five bacteriophages that infect Rhodobacter capsulatus. Genome announcements, 4, 51-56. BORNEMAN, J. & HARTIN, R. J. 2000. PCR primers that amplify fungal rRNA genes from environmental samples. Applied and environmental microbiology, 66, 4356-4360. BRABBAN, A., HITE, E. & CALLAWAY, T. 2005. Evolution of foodborne pathogens via temperate bacteriophage-mediated gene transfer. Foodbourne Pathogens & Disease, 2, 287-303. BREITBART, M. 2012. Marine viruses: truth or dare. Ann Rev Mar Sci, 4, 425-48. BREITBART, M. & ROHWER, F. 2005. Here a virus, there a virus, everywhere the same virus? Trends in microbiology, 13, 278-284. BRÜSSOW, H., CANCHAYA, C. & HARDT, W.-D. 2004. Phages and the evolution of bacterial pathogens: from genomic rearrangements to lysogenic conversion. Microbiology and molecular biology reviews, 68, 560-602. BRUSSOW, H., CANCHAYA, C. & HARDT, W. D. 2004. Phages and the evolution of bacterial pathogens: from genomic rearrangements to lysogenic conversion. Microbiol Mol Biol Rev, 68, 560-602, table of contents. BUCHFINK, B., XIE, C. & HUSON, D. H. 2015. Fast and sensitive protein alignment using DIAMOND. Nature methods, 12, 59. BUERGER, P. & VAN OPPEN, M. J. 2018. Viruses in corals: hidden drivers of coral bleaching and disease? Microbiology Australia, 9-11. BUGNI, T. S. & IRELAND, C. M. 2004. Marine-derived fungi: a chemically and biologically diverse group of microorganisms. Natural Product Reports, 21, 143-163. BURGAUD, G., LE CALVEZ, T., ARZUR, D., VANDENKOORNHUYSE, P. & BARBIER, G. 2009. Diversity of culturable marine filamentous fungi from deep‐sea hydrothermal vents. Environmental Microbiology, 11, 1588-1600. BURGSDORF, I., SLABY, B. M., HANDLEY, K. M., HABER, M., BLOM, J., MARSHALL, C. W., GILBERT, J. A., HENTSCHEL, U. & STEINDLER, L. 2015. Lifestyle evolution in cyanobacterial symbionts of sponges. American Society for Microbiology, 6, 391-415. BURROUGHS, A. M., IYER, L. M. & ARAVIND, L. 2007. Comparative genomics and evolutionary trajectories of viral ATP dependent DNA-packaging systems. Gene and protein evolution. Karger Publishers. 104 BUTALA, M., NOVAK, M., KRAŠEVEC, N., SKOČAJ, M., VERANIČ, P., MAČEK, P. & SEPČIĆ, K. Aegerolysins: lipid-binding proteins with versatile functions. Seminars in Cell & Developmental Biology, 2017. Elsevier, 142-151. BUTLER IV, M. J., HUNT, J. H., HERRNKIND, W. F., CHILDRESS, M. J., BERTELSEN, R., SHARP, W., MATTHEWS, T., FIELD, J. M. & MARSHALL, H. G. 1995. Cascading disturbances in Florida Bay, USA: cyanobacteria blooms, sponge mortality, and implications for juvenile spiny lobsters Panulirus argus. Marine Ecology Progress Series, 129, 119-125. BUTT, A., HIGMAN, V. A., WILLIAMS, C., CRUMP, M. P., HEMSLEY, C. M., HARMER, N. & TITBALL, R. W. 2014. The HicA toxin from Burkholderia pseudomallei has a role in persister cell formation. Biochemical Journal, 459, 333-344. CASJENS, S. R. 2008. Diversity among the tailed-bacteriophages that infect the Enterobacteriaceae. Research in microbiology, 159, 340-348. CENENS, W., MAKUMI, A., MEBRHATU, M. T., LAVIGNE, R. & AERTSEN, A. 2013. Phage–host interactions during pseudolysogeny: Lessons from the Pid/dgo interaction. Bacteriophage, 3, 25029. CHAN, J. Z.-M., MILLARD, A. D., MANN, N. H. & SCHÄFER, H. 2014. Comparative genomics defines the core genome of the growing N4-like phage genus and identifies N4-like Roseophage specific genes. Frontiers in microbiology, 5, 506. CHELIKANI, V., RANJAN, T. & KONDABAGIL, K. 2014. Revisiting the genome packaging in viruses with lessons from the “Giants”. Virology, 466, 15-26. CHEN, B.-W., LIN, M.-H., CHU, C.-H., HSU, C.-E. & SUN, Y.-J. 2015. Insights into ParB spreading from the complex structure of Spo0J and parS. Proceedings of the National Academy of Sciences, 112, 6613-6618. CHEN, F. & LU, J. 2002. Genomic sequence and evolution of marine cyanophage P60: a new insight on lytic and lysogenic phages. Applied and Environmental Microbiology, 68, 2589-2594. CHEN, Y., GOLDING, I., SAWAI, S., GUO, L. & COX, E. C. 2005. Population fitness and the regulation of Escherichia coli genes by bacterial viruses. PLoS biology, 3, e229. CHIBANI-CHENNOUFI, S., BRUTTIN, A., DILLMANN, M.-L. & BRÜSSOW, H. 2004. Phage-host interaction: an ecological perspective. Journal of bacteriology, 186, 3677-3686. CHU, H. & MAZMANIAN, S. K. 2013. Innate immune recognition of the microbiota promotes host-microbial symbiosis. Nature immunology, 14, 668. CHU, T. C., MURRAY, S. R., HSU, S. F., VEGA, Q. & LEE, L. H. 2011. Temperature-induced activation of freshwater Cyanophage AS-1 prophage. Acta Histochem, 113, 294-9. CLAVERIE, J. M., GRZELA, R., LARTIGUE, A., BERNADAC, A., NITSCHE, S., VACELET, J., OGATA, H. & ABERGEL, C. 2009. Mimivirus and Mimiviridae: giant viruses with an increasing number of potential hosts, including corals and sponges. J Invertebr Pathol, 101, 172- 80. CLOKIE, M. R., MILLARD, A. D., LETAROV, A. V. & HEAPHY, S. 2011. Phages in nature. Bacteriophage, 1, 31-45. CLOKIE, M. R., SHAN, J., BAILEY, S., JIA, Y., KRISCH, H. M., WEST, S. & MANN, N. H. 2006. Transcription of a ‘photosynthetic’T4‐type phage during infection of a marine cyanobacterium. Environmental Microbiology, 8, 827-835. COLE, J. R., WANG, Q., FISH, J. A., CHAI, B., MCGARRELL, D. M., SUN, Y., BROWN, C. T., PORRAS- ALFARO, A., KUSKE, C. R. & TIEDJE, J. M. 2014. Ribosomal Database Project: data and tools for high throughput rRNA analysis. Nucleic Acids Res, 42, 633-642. COLOMBO, S., ARIOLI, S., GUGLIELMETTI, S., LUNELLI, F. & MORA, D. 2016. Virome-associated antibiotic-resistance genes in an experimental aquaculture facility. FEMS microbiology ecology, 92, https://doi.org/10.1093/femsec/fiw003. COMEAU, A. M., BERTRAND, C., LETAROV, A., TÉTART, F. & KRISCH, H. 2007. Modular architecture of the T4 phage superfamily: a conserved core genome and a plastic periphery. Virology, 362, 384-396. 105 COMEAU, A. M. & KRISCH, H. M. 2005. War is peace—dispatches from the bacterial and phage killing fields. Current opinion in microbiology, 8, 488-494. COYTE, K. Z., SCHLUTER, J. & FOSTER, K. R. 2015. The ecology of the microbiome: networks, competition, and stability. Science, 350, 663-666. CROUS, P. W., GAMS, W., WINGFIELD, M. J. & VAN WYK, P. 1996. Phaeoacremonium gen. nov. associated with wilt and decline diseases of woody hosts and human infections. Mycologia, 88, 786-796. CZYZ, A., LOS, M., WROBEL, B. & WEGRZYN, G. 2001. Inhibition of spontaneous induction of lambdoid prophages in Escherichia coli cultures: simple procedures with possible biotechnological applications. BMC biotechnology, 1, 1. DANG, V. T., HOWARD-VARONA, C., SCHWENCK, S. & SULLIVAN, M. B. 2015. Variably lytic infection dynamics of large Bacteroidetes podovirus phi38:1 against two Cellulophaga baltica host strains. Environmental Microbiology, 17, 4659-4671. DARLING, A. C., MAU, B., BLATTNER, F. R. & PERNA, N. T. 2004. Mauve: multiple alignment of conserved genomic sequence with rearrangements. Genome research, 14, 1394-1403. DAS, M., ROYER, T. V. & LEFF, L. G. 2008. Fungal communities on decaying leaves in streams: a comparison of two leaf species. Mycological progress, 7, 267-275. DAVIS, B. M. & WALDOR, M. K. 2002. Mobile genetic elements and bacterial pathogenesis. Mobile DNA II. American Society of Microbiology. DE BEECK, M. O., LIEVENS, B., BUSSCHAERT, P., DECLERCK, S., VANGRONSVELD, J. & COLPAERT, J. V. 2014. Comparison and validation of some ITS primer pairs useful for fungal metabarcoding studies. PLoS One, 9, e97629. DE GOEIJ, J. M., VAN OEVELEN, D., VERMEIJ, M. J., OSINGA, R., MIDDELBURG, J. J., DE GOEIJ, A. F. & ADMIRAAL, W. 2013. Surviving in a marine desert: the sponge loop retains resources within coral reefs. Science, 342, 108-110. DE GROOTE, M. A., OCHSNER, U. A., SHILOH, M. U., NATHAN, C., MCCORD, J. M., DINAUER, M. C., LIBBY, S. J., VAZQUEZ-TORRES, A., XU, Y. & FANG, F. C. 1997. Periplasmic superoxide dismutase protects Salmonella from products of phagocyte NADPH-oxidase and nitric oxide synthase. Proceedings of the National Academy of Sciences, 94, 13997-14001. DEGNAN, S. M. 2015. The surprisingly complex immune gene repertoire of a simple sponge, exemplified by the NLR genes: a capacity for specificity? Developmental & Comparative Immunology, 48, 269-274. DERELLE, E., MONIER, A., COOKE, R., WORDEN, A. Z., GRIMSLEY, N. H. & MOREAU, H. 2015. Diversity of viruses infecting the green microalga Ostreococcus lucimarinus. Journal of virology, 89, 5812-5821. DÍEZ‐VIVES, C., ESTEVES, A. I., COSTA, R., NIELSEN, S. & THOMAS, T. 2018. Detecting signatures of a sponge‐associated lifestyle in bacterial genomes. Environmental microbiology reports, 10, 433-443. DÍEZ‐VIVES, C., MOITINHO‐SILVA, L., NIELSEN, S., REYNOLDS, D. & THOMAS, T. 2017. Expression of eukaryotic‐like protein in the microbiome of sponges. Molecular ecology, 26, 1432-1451. DING, B., YIN, Y., ZHANG, F. & LI, Z. 2011. Recovery and phylogenetic diversity of culturable fungi associated with marine sponges Clathrina luteoculcitella and Holoxea sp. in the South China Sea. Marine Biotechnology, 13, 713-721. DINSDALE, E. A., EDWARDS, R. A., HALL, D., ANGLY, F., BREITBART, M., BRULC, J. M., FURLAN, M., DESNUES, C., HAYNES, M. & LI, L. 2008. Functional metagenomic profiling of nine biomes. Nature, 452, 629-632. DÖRING, H., CLERC, P., GRUBE, M. & WEDIN, M. 2000. Mycobiont-Specific PCR Primers for the Amplification of Nuclear its and LSU rDNA from Lichenized Ascomycetes. The Lichenologist, 32, 200-204. EASSON, C. G., SLATTERY, M., MOMM, H. G., OLSON, J. B., THACKER, R. W. & GOCHFELD, D. J. 2013. Exploring Individual-to Population-Level Impacts of Disease on Coral Reef 106 Sponges: Using Spatial Analysis to Assess the Fate, Dynamics, and Transmission of Aplysina Red Band Syndrome (ARBS). PloS one, 8, e79976. EDDY, S. R. 2011. Accelerated profile HMM searches. PLoS computational biology, 7, e1002195. EDGAR, R. C. 2013. UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nature methods, 10, 996-998. EDGAR, R. C., HAAS, B. J., CLEMENTE, J. C., QUINCE, C. & KNIGHT, R. 2011. UCHIME improves sensitivity and speed of chimera detection. Bioinformatics, 27, 2194-2200. EGAN, S. & GARDINER, M. 2016. Microbial dysbiosis: rethinking disease in marine ecosystems. Frontiers in microbiology, 7, 991. EIN-GIL, N., ILAN, M., CARMELI, S., SMITH, G. W., PAWLIK, J. R. & YARDEN, O. 2009. Presence of Aspergillus sydowii, a pathogen of gorgonian sea fans in the marine sponge Spongia obscura. The ISME journal, 3, 752. EISEN, J. A., SWEDER, K. S. & HANAWALT, P. C. 1995. Evolution of the SNF2 family of proteins: subfamilies with distinct sequences and functions. Nucleic acids research, 23, 2715- 2723. ELLESTAD, G. A., EVANS JR, R. H. & KUNSTMANN, M. P. 1969. Structure of a C17 antifungal terpenoid from an unidentified Acrostalagmus species. Journal of the American Chemical Society, 91, 2134-2136. ENAV, H., MANDEL-GUTFREUND, Y. & BÉJÀ, O. 2014. Comparative metagenomic analyses reveal viral-induced shifts of host metabolism towards nucleotide biosynthesis. Microbiome, 2, 9. ERWIN, P. M., COMA, R., LÓPEZ-SENDINO, P., SERRANO, E. & RIBES, M. 2015. Stable symbionts across the HMA-LMA dichotomy: low seasonal and interannual variation in sponge- associated bacteria from taxonomically diverse hosts. FEMS Microbiology Ecology, 91, 115. ERWIN, P. M., PITA, L., LÓPEZ-LEGENTIL, S. & TURON, X. 2012. Stability of Sponge-Associated Bacteria over Large Seasonal Shifts in Temperature and Irradiance. Applied and Environmental Microbiology, 78, 7358-7368. ERWIN, P. M. & THACKER, R. W. 2008. Phototrophic nutrition and symbiont diversity of two Caribbean sponge–cyanobacteria symbioses. Marine Ecology Progress Series, 362, 139- 147. ESTEVES, A. I. S., AMER, N., NGUYEN, M. & THOMAS, T. 2016. Sample processing impacts the viability and cultivability of the sponge microbiome. Frontiers in Microbiology, 7, 499. FAN, L., LIU, M., SIMISTER, R., WEBSTER, N. S. & THOMAS, T. 2013. Marine microbial symbiosis heats up: the phylogenetic and functional response of a sponge holobiont to thermal stress. Isme j, 7, 991-1002. FAN, L., REYNOLDS, D., LIU, M., STARK, M., KJELLEBERG, S., WEBSTER, N. S. & THOMAS, T. 2012. Functional equivalence and evolutionary convergence in complex communities of microbial sponge symbionts. PNAS, 109, 1878-1887. FARRANT, J. L., SANSONE, A., CANVIN, J. R., PALLEN, M. J., LANGFORD, P. R., WALLIS, T. S., DOUGAN, G. & KROLL, J. S. 1997. Bacterial copper‐and zinc‐cofactored superoxide dismutase contributes to the pathogenesis of systemic salmonellosis. Molecular microbiology, 25, 785-796. FAULKNER, D. 1978. Antibiotics from sponges. Topics in antibiotic chemistry, 2, 20-29. FEISS, M. & CATALANO, C. E. 2005. Bacteriophage lambda terminase and the mechanism of viral DNA packaging. Viral Genome Packaging Machines: Genetics, Structure, and Mechanism. Springer. FELL, P. E. & LEWANDROWSKI, K. B. 1981. Population dynamics of the estuarine sponge, Halichondria sp., within a New England eelgrass community. Journal of Experimental Marine Biology and Ecology, 55, 49-63.

107 FINN, R. D., COGGILL, P., EBERHARDT, R. Y., EDDY, S. R., MISTRY, J., MITCHELL, A. L., POTTER, S. C., PUNTA, M., QURESHI, M. & SANGRADOR-VEGAS, A. 2015. The Pfam protein families database: towards a more sustainable future. Nucleic acids research, 44, D279-D285. FIORE, C. L., LABRIE, M., JARETT, J. K. & LESSER, M. P. 2015. Transcriptional activity of the giant barrel sponge, Xestospongia muta Holobiont: molecular evidence for metabolic interchange. Frontiers in microbiology, 6, 364. FRADA, M., PROBERT, I., ALLEN, M. J., WILSON, W. H. & DE VARGAS, C. 2008. The “Cheshire Cat” escape strategy of the coccolithophore Emiliania huxleyi in response to viral infection. Proceedings of the National Academy of Sciences, 105, 15944-15949. FRIEDRICH, A., MERKERT, H., FENDERT, T., HACKER, J., PROKSCH, P. & HENTSCHEL, U. 1999. Microbial diversity in the marine sponge Aplysina cavernicola (formerly Verongia cavernicola) analyzed by fluorescence in situ hybridization (FISH). Marine biology, 134, 461-470. FUHRMAN, J. & SCHWALBACH, M. 2003. Viral influence on aquatic bacterial communities. The Biological Bulletin, 204, 192-195. FUHRMAN, J. & SUTTLE, C. 1993. Viruses in marine planktonic systems. Oceanography, 6, 51- 63. FUHRMAN, J. A. 1999. Marine viruses and their biogeochemical and ecological effects. Nature, 399, 541-548. GALPERIN, M. Y., MAKAROVA, K. S., WOLF, Y. I. & KOONIN, E. V. 2014. Expanded microbial genome coverage and improved protein family annotation in the COG database. Nucleic acids research, 43, D261-D269. GAO, Z., LI, B., ZHENG, C. & WANG, G. 2008. Molecular detection of fungal communities in the Hawaiian marine sponges Suberites zeteki and Mycale armata. Applied and Environmental Microbiology, 74, 6091-6101. GARDES, M. & BRUNS, T. D. 1993. ITS primers with enhanced specificity for basidiomycetes‐ application to the identification of mycorrhizae and rusts. Molecular ecology, 2, 113- 118. GARNEAU, J. R., SEKULOVIC, O., DUPUY, B., SOUTOURINA, O., MONOT, M. & FORTIER, L.-C. 2017. High prevalence and genetic diversity of large phiCD211/phiCDIF1296T-like prophages in Clostridioides difficile. Applied and environmental microbiology, 84, 2164- 2217. GAZAVE, E., LAPÉBIE, P., ERESKOVSKY, A. V., VACELET, J., RENARD, E., CÁRDENAS, P. & BORCHIELLINI, C. 2011. No longer Demospongiae: Homoscleromorpha formal nomination as a fourth class of Porifera. Ancient Animals, New Challenges. Springer. GESNER, S., COHEN, N., ILAN, M., YARDEN, O. & CARMELI, S. 2005. Pandangolide 1a, a metabolite of the sponge-associated fungus Cladosporium sp., and the absolute stereochemistry of pandangolide 1 and iso-cladospolide B. Journal of natural products, 68, 1350-1353. GOCHFELD, D. J., EASSON, C. G., SLATTERY, M., THACKER, R. W. & OLSON, J. B. 2012. Population Dynamics of a Sponge Disease on Caribbean Reefs. Diving For Science 2012, 125. GRABHERR, M. G., HAAS, B. J., YASSOUR, M., LEVIN, J. Z., THOMPSON, D. A., AMIT, I., ADICONIS, X., FAN, L., RAYCHOWDHURY, R. & ZENG, Q. 2011. Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nature biotechnology, 29, 644-652. GRAINGE, I. & JAYARAM, M. 1999. The integrase family of recombinases: organization and function of the active site. Molecular microbiology, 33, 449-456. GRAZIANI, S., XIA, Y., GURNON, J. R., VAN ETTEN, J. L., LEDUC, D., SKOULOUBRIS, S., MYLLYKALLIO, H. & LIEBL, U. 2004. Functional analysis of FAD-dependent thymidylate synthase ThyX from Paramecium bursaria Chlorella virus-1. Journal of Biological Chemistry, 279, 54340-54347. 108 GUAN, S., BASTIN, D. A. & VERMA, N. K. 1999. Functional analysis of the O antigen glucosylation gene cluster of Shigella flexneri bacteriophage SfX. Microbiology, 145, 1263-1273. GUERRERO, R., MARGULIS, L. & BERLANGA, M. 2013. Symbiogenesis: the holobiont as a unit of evolution. Int Microbiol, 16, 133-143. HADAS, E., MARIE, D., SHPIGEL, M. & ILAN, M. 2006. Virus predation by sponges is a new nutrient-flow pathway in coral reef food webs. Limnology and oceanography, 51, 1548-1550. HAFT, D. H., SELENGUT, J. D. & WHITE, O. 2003. The TIGRFAMs database of protein families. Nucleic acids research, 31, 371-373. HAHN-KESER, B. & STOCKEM, W. 1997. Detection of distinct endocytotic and phagocytotic activities in epithelial cells (pinacocytes) of freshwater sponges (Porifera, Spongillidae). Zoomorphology, 117, 121-134. HAN, Y., ZHANG, Y., ZHAO, J. & CHENG, K. 2017. Cyanosiphovirus S-ESS1 Infecting Marine Synechococcus (Chroococcales) Almost Shows No Genetic Relationship to Known Cyanosiphoviruses. Genomics and Applied Biology, 8, 8-15. HARGREAVES, K. R., KROPINSKI, A. M. & CLOKIE, M. R. 2014. Bacteriophage behavioral ecology: how phages alter their bacterial host’s habits. Bacteriophage, 4, 85131. HARVELL, D., ALTIZER, S., CATTADORI, I. M., HARRINGTON, L. & WEIL, E. 2009. Climate change and wildlife diseases: when does the host matter the most? Ecology, 90, 912-920. HAWKSWORTH, D. L. 1991. The fungal dimension of biodiversity: magnitude, significance, and conservation. Mycological research, 95, 641-655. HE, L., LIU, F., KARUPPIAH, V., REN, Y. & LI, Z. 2014. Comparisons of the Fungal and Protistan Communities among Different Marine Sponge Holobionts by Pyrosequencing. Microbial Ecology, 67, 951-961. HENDRIX, R. W., HATFULL, G. F., FORD, M. E., SMITH, M. C. & BURNS, R. N. 2002. Evolutionary relationships among diverse bacteriophages and prophages: all the world's a phage. Horizontal Gene Transfer (Second Edition). Elsevier. HENDRIX, R. W., LAWRENCE, J. G., HATFULL, G. F. & CASJENS, S. 2000. The origins and ongoing evolution of viruses. Trends in microbiology, 8, 504-508. HENRÍQUEZ, M., VERGARA, K., NORAMBUENA, J., BEIZA, A., MAZA, F., UBILLA, P., ARAYA, I., CHÁVEZ, R., SAN-MARTÍN, A. & DARIAS, J. 2014. Diversity of cultivable fungi associated with Antarctic marine sponges and screening for their antimicrobial, antitumoral and antioxidant potential. World Journal of Microbiology and Biotechnology, 30, 65-76. HENTSCHEL, U., FIESELER, L., WEHRL, M., GERNERT, C., STEINERT, M., HACKER, J. & HORN, M. 2003. Microbial diversity of marine sponges. Sponges (Porifera). Springer. HENTSCHEL, U., HOPKE, J., HORN, M., FRIEDRICH, A. B., WAGNER, M., HACKER, J. & MOORE, B. S. 2002. Molecular evidence for a uniform microbial community in sponges from different oceans. Applied and Environmental Microbiology, 68, 4431-4440. HENTSCHEL, U., PIEL, J., DEGNAN, S. M. & TAYLOR, M. W. 2012. Genomic insights into the marine sponge microbiome. Nature Reviews Microbiology, 10, 641-654. HENTSCHEL, U., SCHMID, M., WAGNER, M., FIESELER, L., GERNERT, C. & HACKER, J. 2001. Isolation and phylogenetic analysis of bacteria with antimicrobial activities from the Mediterranean sponges Aplysina aerophoba and Aplysina cavernicola. FEMS Microbiology Ecology, 35, 305-312. HENTSCHEL, U., USHER, K. M. & TAYLOR, M. W. 2006. Marine sponges as microbial fermenters. FEMS microbiology ecology, 55, 167-177. HIBBETT, D., BAUER, R., BINDER, M., GIACHINI, A., HOSAKA, K., JUSTO, A., LARSSON, E., LARSSON, K.-H., LAWREY, J. D. & MIETTINEN, O. 2014. 14 Agaricomycetes. Systematics and evolution. Springer. HOEGH-GULDBERG, O. & BRUNO, J. F. 2010. The impact of climate change on the world’s marine ecosystems. Science, 328, 1523-1528. 109 HOLLER, U., WRIGHT, A. D., MATTHEE, G. F., KONIG, G. M., DRAEGER, S., AUST, H.-J. & SCHULZ, B. 2000. Fungi from marine sponges: diversity, biological activity and secondary metabolites. Mycological Research, 104, 1354-1365. HOOPER, J. N. & VAN SOEST, R. W. 2002. Systema Porifera. A guide to the classification of sponges. Systema Porifera. Springer. HORN, H., SLABY, B. M., JAHN, M. T., BAYER, K., MOITINHO-SILVA, L., FÖRSTER, F., ABDELMOHSEN, U. R. & HENTSCHEL, U. 2016. An enrichment of CRISPR and other defense-related features in marine sponge-associated microbial metagenomes. Frontiers in microbiology, 7, 1751. HOSOYA, T. & TANAKA, K. 2007. Survey of freshwater hyphomycetes in Yakushima Island, southern Japan. Bulletin of the National Museum of Nature and Science, Series B (Botany), 33, 127-132. HUA, J., HUET, A., LOPEZ, C. A., TOROPOVA, K., POPE, W. H., DUDA, R. L., HENDRIX, R. W. & CONWAY, J. F. 2017. Capsids and Genomes of Jumbo-Sized Bacteriophages Reveal the Evolutionary Reach of the HK97 Fold. mBio, 8, 1579-1586. HUANG, C., ZHANG, Y. & JIAO, N. 2010. Phage resistance of a marine bacterium, Roseobacter denitrificans OCh114, as revealed by comparative proteomics. Current microbiology, 61, 141-147. HURWITZ, B. L., HALLAM, S. J. & SULLIVAN, M. B. 2013. Metabolic reprogramming by viruses in the sunlit and dark ocean. Genome biology, 14, 123. HURWITZ, B. L. & SULLIVAN, M. B. 2013. The Pacific Ocean Virome (POV): a marine viral metagenomic dataset and associated protein clusters for quantitative viral ecology. PloS one, 8, 57355. HUSON, D. H., AUCH, A. F., QI, J. & SCHUSTER, S. C. 2007. MEGAN analysis of metagenomic data. Genome research, 17, 377-386. HYATT, D., CHEN, G.-L., LOCASCIO, P. F., LAND, M. L., LARIMER, F. W. & HAUSER, L. J. 2010. Prodigal: prokaryotic gene recognition and translation initiation site identification. BMC bioinformatics, 11, 119. HYDE, K. D., JONES, E. G., LEAÑO, E., POINTING, S. B., POONYTH, A. D. & VRIJMOED, L. L. 1998. Role of fungi in marine ecosystems. Biodiversity and Conservation, 7, 1147-1161. HYDE, K. D., JONES, E. G., LIU, J.-K., ARIYAWANSA, H., BOEHM, E., BOONMEE, S., BRAUN, U., CHOMNUNTI, P., CROUS, P. W. & DAI, D.-Q. 2013. Families of dothideomycetes. Fungal Diversity, 63, 1-313. ILAN, M. & ABELSON, A. 1995. The life of a sponge in a sandy lagoon. The Biological Bulletin, 189, 363-369. IMHOFF, J. F. 2016. Natural products from marine fungi—Still an underrepresented resource. Marine drugs, 14, 19. JADULCO, R., BRAUERS, G., EDRADA, R. A., EBEL, R., WRAY, V., SUDARSONO, A. & PROKSCH, P. 2002. New Metabolites from Sponge-Derived Fungi Curvularia l unata and Cladosporium h erbarum. Journal of natural products, 65, 730-733. JARES-ERIJMAN, E. A., SAKAI, R. & RINEHART, K. L. 1991. Crambescidins: new antiviral and cytotoxic compounds from the sponge Crambe crambe. The Journal of Organic Chemistry, 56, 5712-5715. JEBARAJ, C. S., RAGHUKUMAR, C., BEHNKE, A. & STOECK, T. 2010. Fungal diversity in oxygen- depleted regions of the Arabian Sea revealed by targeted environmental sequencing combined with cultivation. FEMS Microbiology Ecology, 71, 399-412. JEEWON, R. & HYDE, K. 2007. Detection and diversity of fungi from environmental samples: traditional versus molecular approaches. Advanced Techniques in Soil Microbiology. Springer. JI, J., ZHANG, R. & JIAO, N. 2015. Complete genome sequence of Roseophage vB_DshP-R1, which infects Dinoroseobacter shibae DFL12. Standards in genomic sciences, 10, 6.

110 JIANG, S. & PAUL, J. 1998. Significance of lysogeny in the marine environment: studies with isolates and a model of lysogenic phage production. Microbial ecology, 35, 235-243. JÜRGENS, K. & MATZ, C. 2002. Predation as a shaping force for the phenotypic and genotypic composition of planktonic bacteria. Antonie van Leeuwenhoek, 81, 413-434. JURGENSON, C. T., BEGLEY, T. P. & EALICK, S. E. 2009. The structural and biochemical foundations of thiamin biosynthesis. Annual review of biochemistry, 78, 569-603. JUSTICE, S. S., HUNSTAD, D. A., CEGELSKI, L. & HULTGREN, S. J. 2008. Morphological plasticity as a bacterial survival strategy. Nature Reviews Microbiology, 6, 162. KANEHISA, M., GOTO, S., KAWASHIMA, S. & NAKAYA, A. 2002. The KEGG databases at GenomeNet. Nucleic acids research, 30, 42-46. KANEKO, J., KIMURA, T., KAWAKAMI, Y., TOMITA, T. & KAMIO, Y. 1997. Panton-valentine leukocidin genes in a phage-like particle isolated from mitomycin C-treated Staphylococcus aureus V8 (ATCC 49775). Bioscience, biotechnology, and biochemistry, 61, 1960-1962. KARIMI, E., RAMOS, M., GONÇALVES, J., XAVIER, J. R., REIS, M. P. & COSTA, R. C. 2017. Comparative metagenomics reveals the distinctive adaptive features of the Spongia officinalis endosymbiotic consortium. Frontiers in Microbiology, 8, 2499. KELLY, L. W., WILLIAMS, G. J., BAROTT, K. L., CARLSON, C. A., DINSDALE, E. A., EDWARDS, R. A., HAAS, A. F., HAYNES, M., LIM, Y. W. & MCDOLE, T. 2014. Local genomic adaptation of coral reef-associated microbiomes to gradients of natural variability and anthropogenic stressors. Proceedings of the National Academy of Sciences, 111, 10227-10232. KELMAN, D., KASHMAN, Y., HILL, R. T., ROSENBERG, E. & LOYA, Y. 2009. Chemical warfare in the sea: The search for antibiotics from Red Sea corals and sponges. Pure and Applied Chemistry, 81, 1113-1121. KHAKHUM, N., YORDPRATUM, U., BOONMEE, A., TATTAWASART, U., RODRIGUES, J. L. & SERMSWAN, R. W. 2016. Cloning, expression, and characterization of a peptidoglycan hydrolase from the Burkholderia pseudomallei phage ST79. AMB Express, 6, 77. KHAYAT, R., TANG, L., LARSON, E. T., LAWRENCE, C. M., YOUNG, M. & JOHNSON, J. E. 2005. Structure of an archaeal virus capsid protein reveals a common ancestry to eukaryotic and bacterial viruses. Proceedings of the National Academy of Sciences, 102, 18944- 18949. KHOA, H. V., MIDORIKAWA, Y., UCHINO, T., NAKAI, T., KATO, G., KONDO, H., HIRONO, I., LABAIDEN, M., DIREKBUSARAKOM, S. & SANO, M. 2017. Complete Genome Sequence of the Lytic Giant Bacteriophage pT24 Infecting Tenacibaculum spp., Isolated from a Shrimp Culture Pond. Genome announcements, 5, 81-88. KIM, T. K., HEWAVITHARANA, A. K., SHAW, P. N. & FUERST, J. A. 2006. Discovery of a new source of rifamycin antibiotics in marine sponge actinobacteria by phylogenetic prediction. Applied and environmental microbiology, 72, 2118-2125. KING, A. M., LEFKOWITZ, E., ADAMS, M. J. & CARSTENS, E. B. 2011. Virus taxonomy: ninth report of the International Committee on Taxonomy of Viruses, Elsevier. KIS-PAPO, T. 2005. Marine fungal communities. Mycology Series, 23, 61. KNOWLES, B., SILVEIRA, C., BAILEY, B., BAROTT, K., CANTU, V., COBIÁN-GÜEMES, A., COUTINHO, F., DINSDALE, E., FELTS, B. & FURBY, K. 2016. Lytic to temperate switching of viral communities. Nature, 531, 466. KOHLMEYER, J. & VOLKMANN-KOHLMEYER, B. 1990. New species of Koralionastes (Ascomycotina) from the Caribbean and Australia. Canadian Journal of Botany, 68, 1554-1559. KOHLMEYER, J. & VOLKMANN-KOHLMEYER, B. 1992. Two Ascomycotina from coral reefs in the Caribbean and Australia. Cryptogamic Botany, 2, 367-374. KOONIN, E. V., SENKEVICH, T. G. & DOLJA, V. V. 2006. The ancient Virus World and evolution of cells. Biology direct, 1, 29. 111 KOONIN, E. V. & YUTIN, N. 2010. Origin and evolution of eukaryotic large nucleo-cytoplasmic DNA viruses. Intervirology, 53, 284-292. KROGH, S., JØRGENSEN, S. T. & DEVINE, K. M. 1998. Lysis Genes of the Bacillus subtilisDefective Prophage PBSX. Journal of bacteriology, 180, 2110-2117. KUBICEK, C. P., KOMON-ZELAZOWSKA, M. & DRUZHININA, I. S. 2008. Fungal genus Hypocrea/Trichoderma: from barcodes to biodiversity. Journal of Zhejiang University Science B, 9, 753. KULIKOVA, N., SAIBATALOVA, E., BOIKO, S., SEMITURKINA, N., BELOZEROVA, O. Y., MEKHONOSHIN, A., TIMOSHKIN, O. & SUTURIN, A. 2013. Biogeochemistry of encrusting sponges of the family Lubomirskiidae in Southern Lake Baikal. Geochemistry International, 51, 326-337. KUMAR, S., NEI, M., DUDLEY, J. & TAMURA, K. 2008. MEGA: a biologist-centric software for evolutionary analysis of DNA and protein sequences. Briefings in bioinformatics, 9, 299-306. LACKNER, G., PETERS, E. E., HELFRICH, E. J. & PIEL, J. 2017. Insights into the lifestyle of uncultured bacterial natural product factories associated with marine sponges. Proceedings of the National Academy of Sciences, 114, 347-356. LAFFY, P. W., WOOD‐CHARLSON, E. M., TURAEV, D., JUTZ, S., PASCELLI, C., BOTTÉ, E. S., BELL, S. C., PEIRCE, T. E., WEYNBERG, K. D. & VAN OPPEN, M. J. 2018. Reef invertebrate viromics: diversity, host specificity and functional capacity. Environmental microbiology. LAFFY, P. W., WOOD CHARLSON, E., TURAEV, D., WEYNBERG, K. D., BOTTE, E., VAN OPPEN, M. J., WEBSTER, N. & RATTEI, T. 2016. HoloVir: A workflow for investigating the diversity and function of viruses in invertebrate holobionts. Frontiers in Microbiology, 7, 822. LANGMEAD, B. & SALZBERG, S. L. 2012. Fast gapped-read alignment with Bowtie 2. Nature methods, 9, 357. LAPORT, M., SANTOS, O. & MURICY, G. 2009. Marine sponges: potential sources of new antimicrobial drugs. Current pharmaceutical biotechnology, 10, 86-105. LEBEDEV, A. A., KRAUSE, M. H., ISIDRO, A. L., VAGIN, A. A., ORLOVA, E. V., TURNER, J., DODSON, E. J., TAVARES, P. & ANTSON, A. A. 2007. Structural framework for DNA translocation via the viral portal protein. The EMBO journal, 26, 1984-1994. LEE, O. O., WANG, Y., YANG, J., LAFI, F. F., AL-SUWAILEM, A. & QIAN, P.-Y. 2011. Pyrosequencing reveals highly diverse and species-specific microbial communities in sponges from the Red Sea. The ISME journal, 5, 650-664. LEE, S. 1988. A rapid high yield mini-prep method for isolation of total genomic DNA from fungi. Fungal Genet. Newsl., 35, 23-24. LEE, Y. K., LEE, J.-H. & LEE, H. K. 2001a. Microbial symbiosis in marine sponges. Journal of Microbiology, 39, 254-264. LEE, Y. K., LEE, J.-H. & LEE, H. K. 2001b. Microbial symbiosis in marine sponges. JOURNAL OF MICROBIOLOGY-SEOUL-, 39, 254-264. LI, Q. & WANG, G. 2009. Diversity of fungal isolates from three Hawaiian marine sponges. Microbiological research, 164, 233-241. LI, W. & GODZIK, A. 2006. Cd-hit: a fast program for clustering and comparing large sets of protein or nucleotide sequences. Bioinformatics, 22, 1658-1659. LI, Z., HE, L. & MIAO, X. 2007. Cultivable bacterial community from South China Sea sponge as revealed by DGGE fingerprinting and 16S rDNA phylogenetic analysis. Current microbiology, 55, 465-472. LINDELL, D., JAFFE, J. D., JOHNSON, Z. I., CHURCH, G. M. & CHISHOLM, S. W. 2005. Photosynthesis genes in marine viruses yield proteins during host infection. Nature, 438, 86-89. LINDELL, D., SULLIVAN, M. B., JOHNSON, Z. I., TOLONEN, A. C., ROHWER, F. & CHISHOLM, S. W. 2004. Transfer of photosynthesis genes to and from Prochlorococcus viruses. 112 Proceedings of the National Academy of Sciences of the United States of America, 101, 11013-11018. LIU, M., FAN, L., ZHONG, L., KJELLEBERG, S. & THOMAS, T. 2012. Metaproteogenomic analysis of a community of sponge symbionts. The ISME journal, 6, 1515-25. LIU, M. Y., KJELLEBERG, S. & THOMAS, T. 2011. Functional genomic analysis of an uncultured delta-proteobacterium in the sponge Cymbastela concentrica. The ISME journal 5, 427- 35. LIU, T., LIU, L., JIANG, X., HUANG, X. & CHEN, J. 2009. A new furanoid toxin produced by Curvularia lunata, the causal agent of maize Curvularia leaf spot. Canadian Journal of Plant Pathology, 31, 22-27. LIU, W., LI, C., ZHU, P., YANG, J. & CHENG, K. 2010. Phylogenetic diversity of culturable fungi associated with two marine sponges: Haliclona simulans and Gelliodes carnosa, collected from the Hainan Island coastal waters of the South China Sea. Fungal Diversity, 42, 1-15. LOHR, J. E., CHEN, F. & HILL, R. T. 2005. Genomic Analysis of Bacteriophage ΦJL001: Insights into Its Interaction with a Sponge-Associated Alpha-Proteobacterium. Applied and Environmental Microbiology, 71, 1598-1609. ŁOŚ, M. & WĘGRZYN, G. 2012. Pseudolysogeny. Advances in virus research. Elsevier. LOUGH, J., ANDERSON, K. & HUGHES, T. 2018. Increasing thermal stress for tropical coral reefs: 1871–2017. Scientific reports, 8, 6079. MAHONY, J., ALQARNI, M., STOCKDALE, S., SPINELLI, S., FEYEREISEN, M., CAMBILLAU, C. & VAN SINDEREN, D. 2016. Functional and structural dissection of the tape measure protein of lactococcal phage TP901-1. Scientific reports, 6, 36667. MALDONADO, M., CARMONA, M. C., VELÁSQUEZ, Z., PUIG, A., CRUZADO, A., LÓPEZ, A. & YOUNG, C. M. 2005a. Siliceous sponges as a silicon sink: an overlooked aspect of benthopelagic coupling in the marine silicon cycle. Limnology and Oceanography, 50, 799-809. MALDONADO, M., CORTADELLAS, N., TRILLAS, M. I. & RÜTZLER, K. 2005b. Endosymbiotic yeast maternally transmitted in a marine sponge. The Biological Bulletin, 209, 94-106. MALDONADO, M., RIBES, M. & VAN DUYL, F. C. 2012. Nutrient fluxes through sponges: biology, budgets, and ecological implications. Advances in marine biology. Elsevier. MANILOFF, J. & ACKERMANN, H.-W. 1998. Taxonomy of bacterial viruses: establishment of tailed virus genera and the other Caudovirales. Archives of Virology, 143, 2051-2063. MANN, N. H., COOK, A., MILLARD, A., BAILEY, S. & CLOKIE, M. 2003. Marine ecosystems: bacterial photosynthesis genes in a virus. Nature, 424, 741-741. MARGULIS, L. 1991. Symbiogenesis and symbionticism. Symbiosis as a source of evolutionary innovation, 1-14. MARKOWITZ, V. M., CHEN, I.-M. A., PALANIAPPAN, K., CHU, K., SZETO, E., GRECHKIN, Y., RATNER, A., JACOB, B., HUANG, J. & WILLIAMS, P. 2011. IMG: the integrated microbial genomes database and comparative analysis system. Nucleic acids research, 40, 115- 122. MAURICE, C., BOUVIER, C., DE WIT, R. & BOUVIER, T. 2013. Linking the lytic and lysogenic bacteriophage cycles to environmental conditions, host physiology and their variability in coastal lagoons. Environmental microbiology, 15, 2463-2475. MAVRIS, M., MANNING, P. A. & MORONA, R. 1997. Mechanism of bacteriophage SfII‐mediated serotype conversion in Shigella flexneri. Molecular microbiology, 26, 939-950. MAXWELL, K. L., YEE, A. A., BOOTH, V., ARROWSMITH, C. H., GOLD, M. & DAVIDSON, A. R. 2001. The solution structure of bacteriophage λ protein W, a small morphogenetic protein possessing a novel fold1. Journal of molecular biology, 308, 9-14. MCCALLUM, H. I., KURIS, A., HARVELL, C. D., LAFFERTY, K. D., SMITH, G. W. & PORTER, J. 2004. Does terrestrial epidemiology apply to marine systems? Trends in Ecology & Evolution, 19, 585-591. 113 MCDANIEL, L., HOUCHIN, L., WILLIAMSON, S. & PAUL, J. 2002. Plankton blooms: Lysogeny in marine Synechococcus. Nature, 415, 496-496. MENEZES, C. B., BONUGLI-SANTOS, R. C., MIQUELETTO, P. B., PASSARINI, M. R., SILVA, C. H., JUSTO, M. R., LEAL, R. R., FANTINATTI-GARBOGGINI, F., OLIVEIRA, V. M. & BERLINCK, R. G. 2010. Microbial diversity associated with algae, ascidians and sponges from the north coast of São Paulo state, Brazil. Microbiological research, 165, 466-482. MIADLIKOWSKA, J., KAUFF, F., HOFSTETTER, V., FRAKER, E., GRUBE, M., HAFELLNER, J., REEB, V., HODKINSON, B. P., KUKWA, M. & LÜCKING, R. 2006. New insights into classification and evolution of the Lecanoromycetes (Pezizomycotina, Ascomycota) from phylogenetic analyses of three ribosomal RNA-and two protein-coding genes. Mycologia, 98, 1088-1103. MIDDELBOE, M., CHAN, A. & BERTELSEN, S. K. 2010. Isolation and life cycle characterization of lytic viruses infecting heterotrophic bacteria and cyanobacteria. Manual of aquatic viral ecology. American Society of Limnology and Oceanography, Inc. MIDDELBOE, M. & LYCK, P. G. 2002. Regeneration of dissolved organic matter by viral lysis in marine microbial communities. Aquatic Microbial Ecology, 27, 187-194. MIHARA, H. & ESAKI, N. 2002. Bacterial cysteine desulfurases: their function and mechanisms. Applied microbiology and biotechnology, 60, 12-23. MITOVA, M., POPOV, S. & DE ROSA, S. 2004a. Cyclic peptides from a Ruegeria strain of bacteria associated with the sponge Suberites domuncula. Journal of natural products, 67, 1178-1181. MITOVA, M., TOMMONARO, G., HENTSCHEL, U., MÜLLER, W. E. & DE ROSA, S. 2004b. Exocellular cyclic dipeptides from a Ruegeria strain associated with cell cultures of Suberites domuncula. Marine Biotechnology, 6, 95-103. MŁYNARCZYK, G., MŁYNARCZYK, A., ZABICKA, D. & JELJASZEWICZ, J. 1997. Lysogenic conversion as a factor influencing the vancomycin tolerance phenomenon in Staphylococcus aureus. Journal of Antimicrobial Chemotherapy, 40, 136-137. MODROW, S., FALKE, D., TRUYEN, U. & SCHÄTZL, H. 2013. Viruses: Definition, Structure, Classification. Molecular Virology. Springer. MOEBUS, K. 1996. Marine bacteriophage reproduction under nutrient-limited growth of host bacteria. I. Investigations with six phage-host systems. Marine Ecology Progress Series, 144, 1-12. MOHAMED, N. M., SAITO, K., TAL, Y. & HILL, R. T. 2010. Diversity of aerobic and anaerobic ammonia-oxidizing bacteria in marine sponges. The ISME journal, 4, 38. MOITINHO-SILVA, L., STEINERT, G., NIELSEN, S., HARDOIM, C. C., WU, Y.-C., MCCORMACK, G. P., LOPEZ-LEGENTIL, S., MARCHANT, R., WEBSTER, N. & THOMAS, T. 2017. Predicting the HMA-LMA status in marine sponges by machine learning. Frontiers in microbiology, 8, 752. MOJICA, K. D. & BRUSSAARD, C. P. 2014. Factors affecting virus dynamics and microbial host– virus interactions in marine environments. FEMS microbiology ecology, 89, 495-515. MOKILI, J. L., ROHWER, F. & DUTILH, B. E. 2012. Metagenomics and future perspectives in virus discovery. Current opinion in virology, 2, 63-77. MONKS, N. R., LERNER, C., HENRIQUES, A. T., FARIAS, F. M., SCHAPOVAL, E. E., SUYENAGA, E. S., DA ROCHA, A. B., SCHWARTSMANN, G. & MOTHES, B. 2002. Anticancer, antichemotactic and antimicrobial activities of marine sponges collected off the coast of Santa Catarina, southern Brazil. Journal of Experimental Marine Biology and Ecology, 281, 1-12. MORRISON-GARDINER, S. 2002. Dominant fungi from Australian coral reefs. Fungal Divers, 9, 105-121. MOTEGI, C. & NAGATA, T. 2007. Enhancement of viral production by addition of nitrogen or nitrogen plus carbon in subtropical surface waters of the South Pacific. Aquatic microbial ecology, 48, 27-34. 114 MÜLLER, W., KLEMT, M., THAKUR, N., SCHRÖDER, H., AIELLO, A., D'ESPOSITO, M., MENNA, M. & FATTORUSSO, E. 2004. Molecular/chemical ecology in sponges: evidence for an adaptive antibacterial response in Suberites domuncula. Marine biology, 144, 19-29. MÜLLER, W. E. & MÜLLER, I. M. 2003. Origin of the Metazoan Immune System: Identification of the Molecules and Their Functions in Sponges. Integrative and comparative biology, 43, 281-292. MUSCHOLL-SILBERHORN, A., THIEL, V. & IMHOFF, J. F. 2008. Abundance and bioactivity of cultured sponge-associated bacteria from the Mediterranean Sea. Microbial Ecology, 55, 94-106. MYLLYKALLIO, H., LIPOWSKI, G., LEDUC, D., FILEE, J., FORTERRE, P. & LIEBL, U. 2002. An alternative flavin-dependent mechanism for thymidylate synthesis. Science, 297, 105- 107. NAIM, M. A., SMIDT, H. & SIPKEMA, D. 2017. Fungi found in Mediterranean and North Sea sponges: how specific are they? PeerJ, 5, 3722. NEGANDHI, K., BLACKWELDER, P. L., ERESKOVSKY, A. V. & LOPEZ, J. V. 2010. Florida reef sponges harbor coral disease-associated microbes. Symbiosis, 51, 117-129. NGUYEN, M. T. H. D., LIU, M. & THOMAS, T. 2014. Ankyrin-repeat proteins from sponge symbionts modulate amoebal phagocytosis. Molecular Ecology, 23, 1635-1645. NIKOLCHEVA, L. G., COCKSHUTT, A. M. & BÄRLOCHER, F. 2003. Determining diversity of freshwater fungi on decaying leaves: comparison of traditional and molecular approaches. Applied and environmental microbiology, 69, 2548-2554. NOGUCHI, H., TANIGUCHI, T. & ITOH, T. 2008. MetaGeneAnnotator: detecting species-specific patterns of ribosomal binding site for precise gene prediction in anonymous prokaryotic and phage genomes. DNA research, 15, 387-396. OKSANEN, J., BLANCHET, F. G., KINDT, R., LEGENDRE, P., MINCHIN, P. R., O’HARA, R., SIMPSON, G. L., SOLYMOS, P., STEVENS, M. H. H. & WAGNER, H. 2013. Package ‘vegan’. Community ecology package, version, 2, 1-198. OLSON, J. B., THACKER, R. W. & GOCHFELD, D. J. 2014. Molecular community profiling reveals impacts of time, space, and disease status on the bacterial community associated with the Caribbean sponge Aplysina cauliformis. FEMS microbiology ecology, 87, 268-279. PANG, K.-L. & MITCHELL, J. I. 2005. Molecular approaches for assessing fungal diversity in marine substrata. Botanica Marina, 48, 332-347. PARADA, V., HERNDL, G. J. & WEINBAUER, M. G. 2006. Viral burst size of heterotrophic prokaryotes in aquatic systems. Journal of the Marine Biological Association of the United Kingdom, 86, 613-621. PASSARINI, M. R., MIQUELETTO, P. B., DE OLIVEIRA, V. M. & SETTE, L. D. 2015. Molecular diversity of fungal and bacterial communities in the marine sponge Dragmacidon reticulatum. Journal of basic microbiology, 55, 207-220. PASSARINI, M. R., SANTOS, C., LIMA, N., BERLINCK, R. G. & SETTE, L. D. 2013. Filamentous fungi from the Atlantic marine sponge Dragmacidon reticulatum. Archives of microbiology, 195, 99-111. PATERSON, D. L. 2006. Resistance in gram-negative bacteria: Enterobacteriaceae. American journal of infection control, 34, 20-28. PATON, A. W. & PATON, J. C. 2010. Escherichia coli subtilase cytotoxin. Toxins, 2, 215-228. PAUL, J. H. 2008. Prophages in marine bacteria: dangerous molecular time bombs or the key to survival in the seas? Isme j, 2, 579-89. PAYET, J. P. & SUTTLE, C. A. 2013. To kill or not to kill: the balance between lytic and lysogenic viral infection is driven by trophic status. Limnology and Oceanography, 58, 465-474. PAZ, Z., KOMON-ZELAZOWSKA, M., DRUZHININA, I., AVESKAMP, M., SHNAIDERMAN, A., ALUMA, Y., CARMELI, S., ILAN, M. & YARDEN, O. 2010. Diversity and potential antifungal properties of fungi associated with a Mediterranean sponge. Fungal Diversity, 42, 17-26. 115 PERNTHALER, J. 2005. Predation on prokaryotes in the water column and its ecological implications. Nature Reviews Microbiology, 3, 537. PEROVIĆ‐OTTSTADT, S., ADELL, T., PROKSCH, P., WIENS, M., KORZHEV, M., GAMULIN, V., MÜLLER, I. M. & MÜLLER, W. E. 2004. A (1→ 3)‐β‐d‐glucan recognition protein from the sponge Suberites domuncula. European Journal of Biochemistry, 271, 1924-1937. PERRY, N. B., BLUNT, J. W., MUNRO, M. H. & PANNELL, L. K. 1988. Mycalamide A, an antiviral compound from a New Zealand sponge of the genus Mycale. Journal of the American Chemical Society, 110, 4850-4851. PERRY, N. B., ETTOUATI, L., LITAUDON, M., BLUNT, J. W., MUNRO, M. H., PARKIN, S. & HOPE, H. 1994. Alkaloids from the antarctic sponge Kirkpatrickia varialosa.: Part 1: Variolin b, a new antitumour and antiviral compound. Tetrahedron, 50, 3987-3992. PETERSON, B. J., CHESTER, C. M., JOCHEM, F. J. & FOURQUREAN, J. W. 2006. Potential role of sponge communities in controlling phytoplankton blooms in Florida Bay. Marine Ecology Progress Series, 328, 93-103. PHILIPPA, J., LEIGHTON, F., DAOUST, P. Y., NIELSEN, O., PAGLIARULO, M., SCHWANTJE, H., SHURY, T., VAN, R. H., MARTINA, B. & KUIKEN, T. 2004. Antibodies to selected pathogens in free-ranging terrestrial carnivores and marine mammals in Canada. The Veterinary Record, 155, 135-140. PITA, L., ERWIN, P. M., TURON, X. & LÓPEZ-LEGENTIL, S. 2013a. Till death do us part: stable sponge-bacteria associations under thermal and food shortage stresses. PLoS One, 8, 80307. PITA, L., FRAUNE, S. & HENTSCHEL, U. 2016. Emerging sponge models of animal-microbe symbioses. Frontiers in microbiology, 7, 2102. PITA, L., RIX, L., SLABY, B. M., FRANKE, A. & HENTSCHEL, U. 2018. The sponge holobiont in a changing ocean: from microbes to ecosystems. Microbiome, 6, 46. PITA, L., TURON, X., LÓPEZ-LEGENTIL, S. & ERWIN, P. M. 2013b. Host rules: spatial stability of bacterial communities associated with marine sponges (Ircinia spp.) in the Western Mediterranean Sea. FEMS microbiology ecology, 86, 268-276. PIVKIN, M., ALESHKO, S., KRASOKHIN, V. & KHUDYAKOVA, Y. V. 2006. Fungal assemblages associated with sponges of the southern coast of Sakhalin Island. Russian Journal of Marine Biology, 32, 207-213. POON, L. L., PRITLOVE, D. C., FODOR, E. & BROWNLEE, G. G. 1999. Direct evidence that the poly (A) tail of influenza A virus mRNA is synthesized by reiterative copying of a U track in the virion RNA template. Journal of virology, 73, 3473-3476. PROKSCH, P., EBEL, R., EDRADA, R., RIEBE, F., LIU, H., DIESEL, A., BAYER, M., LI, X., HAN LIN, W. & GREBENYUK, V. 2008. Sponge-associated fungi and their bioactive compounds: the Suberites case. Botanica Marina, 51, 209-218. PROKSCH, P., EBEL, R., EDRADA, R., SCHUPP, P., LIN, W., WRAY, V. & STEUBE, K. 2003. Detection of pharmacologically active natural products using ecology. Selected examples from Indopacific marine invertebrates and sponge-derived fungi. Pure and applied chemistry, 75, 343-352. PUNTA, M., COGGILL, P. C., EBERHARDT, R. Y., MISTRY, J., TATE, J., BOURSNELL, C., PANG, N., FORSLUND, K., CERIC, G. & CLEMENTS, J. 2011. The Pfam protein families database. Nucleic acids research, 40, 290-301. QI, S.-H., XU, Y., XIONG, H.-R., QIAN, P.-Y. & ZHANG, S. 2009. Antifouling and antibacterial compounds from a marine fungus Cladosporium sp. F14. World Journal of Microbiology and Biotechnology, 25, 399. QIN, J., LI, R., RAES, J., ARUMUGAM, M., BURGDORF, K. S., MANICHANH, C., NIELSEN, T., PONS, N., LEVENEZ, F. & YAMADA, T. 2010. A human gut microbial gene catalogue established by metagenomic sequencing. nature, 464, 59.

116 QUAST, C., PRUESSE, E., YILMAZ, P., GERKEN, J., SCHWEER, T., YARZA, P., PEPLIES, J. & GLÖCKNER, F. O. 2012. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic acids research, 50, 1219. QUINONES, M., KIMSEY, H. H. & WALDOR, M. K. 2005. LexA cleavage is required for CTX prophage induction. Molecular cell, 17, 291-300. RAGHUKUMAR, C. 2008. Marine fungal biotechnology: an ecological perspective. Fungal Diversity, 31, 19-35. READE, P. C. 1968. Phagocytosis in invertebrates. Immunology and Cell Biology, 46, 219. RECKTENWALD, J. & SCHMIDT, H. 2002. The nucleotide sequence of Shiga toxin (Stx) 2e- encoding phage φP27 is not related to other Stx phage genomes, but the modular genetic structure is conserved. Infection and immunity, 70, 1896-1908. REHNER, S. A., MINNIS, A. M., SUNG, G.-H., LUANGSA-ARD, J. J., DEVOTTO, L. & HUMBER, R. A. 2011. Phylogeny and systematics of the anamorphic, entomopathogenic genus Beauveria. Mycologia, 103, 1055-1073. REINFELDER, J. R. 2010. Carbon concentrating mechanisms in eukaryotic marine phytoplankton. REISWIG, H. M. 1974. Water transport, respiration and energetics of three tropical marine sponges. Journal of Experimental Marine Biology and Ecology, 14, 231-249. REYNOLDS, D. & THOMAS, T. 2016. Evolution and function of eukaryotic‐like proteins from sponge symbionts. Molecular ecology, 25, 5242-5253. RIBEIRO, S. M., OMENA, E. P. & MURICY, G. 2003. Macrofauna associated to Mycale microsigmatosa (Porifera, Demospongiae) in Rio de Janeiro State, SE Brazil. Estuarine, Coastal and Shelf Science, 57, 951-959. RICHARDS, T. A., JONES, M. D., LEONARD, G. & BASS, D. 2012. Marine fungi: their ecology and molecular diversity. Annual review of marine science, 4, 495-522. RIEMANN, L. & MIDDELBOE, M. 2002. Viral lysis of marine bacterioplankton: implications for organic matter cycling and bacterial clonal composition. Ophelia, 56, 57-68. RINALDI, M., PHILLIPS, P., SCHWARTZ, J., WINN, R., HOLT, G., SHAGETS, F., ELROD, J., NISHIOKA, G. & AUFDEMORTE, T. 1987. Human Curvularia infections: report of five cases and review of the literature. Diagnostic microbiology and infectious disease, 6, 27-39. RIPP, S. & MILLER, R. V. 1997. The role of pseudolysogeny in bacteriophage-host interactions in a natural freshwater environment. Microbiology, 143, 2065-2070. RIPP, S. & MILLER, R. V. 1998. Dynamics of the pseudolysogenic response in slowly growing cells of Pseudomonas aeruginosa. Microbiology, 144, 2225-2232. RODRIGUEZ-BRITO, B., LI, L., WEGLEY, L., FURLAN, M., ANGLY, F., BREITBART, M., BUCHANAN, J., DESNUES, C., DINSDALE, E. & EDWARDS, R. 2010. Viral and microbial community dynamics in four aquatic environments. The ISME journal, 4, 739-751. RODRÍGUEZ-MARCONI, S., DE LA IGLESIA, R., DÍEZ, B., FONSECA, C. A., HAJDU, E. & TREFAULT, N. 2015. Characterization of bacterial, archaeal and eukaryote symbionts from Antarctic sponges reveals a high diversity at a three-domain level and a particular signature for this ecosystem. PloS one, 10, 138837. ROHWER, F. & EDWARDS, R. 2002. The Phage Proteomic Tree: a genome-based taxonomy for phage. Journal of bacteriology, 184, 4529-4535. ROHWER, F., SEGALL, A., STEWARD, G., SEGURITAN, V., BREITBART, M., WOLVEN, F. & FAROOQ AZAM, F. 2000. The complete genomic sequence of the marine phage Roseophage SIO1 shares homology with nonmarine phages. Limnology and Oceanography, 45, 408-418. ROSSMAN, A. Y., FARR, D. F. & CASTLEBURY, L. A. 2007. A review of the phylogeny and biology of the Diaporthales. Mycoscience, 48, 135-144. ROT, C., GOLDFARB, I., ILAN, M. & HUCHON, D. 2006. Putative cross-kingdom horizontal gene transfer in sponge (Porifera) mitochondria. BMC Evolutionary Biology, 6, 71. 117 ROUX, S., ENAULT, F., HURWITZ, B. L. & SULLIVAN, M. B. 2015. VirSorter: mining viral signal from microbial genomic data. PeerJ, 3, 985. ROUX, S., TOURNAYRE, J., MAHUL, A., DEBROAS, D. & ENAULT, F. 2014. Metavir 2: new tools for viral metagenome comparison and assembled virome analysis. BMC bioinformatics, 15, 76. ROZAS, E. E., ALBANO, R. M., LÔBO-HAJDU, G., MÜLLER, W. E., SCHRÖDER, H.-C. & CUSTÓDIO, M. R. 2011. Isolation and cultivation of fungal strains from in vitro cell cultures of two marine sponges (Porifera: Halichondrida and Haplosclerida). Brazilian Journal of Microbiology, 42, 1560-1568. RUGGERI, Z. M. & WARE, J. 1993. von Willebrand factor. The FASEB Journal, 7, 308-316. RYU, T., SERIDI, L., MOITINHO-SILVA, L., OATES, M., LIEW, Y. J., MAVROMATIS, C., WANG, X., HAYWOOD, A., LAFI, F. F. & KUPRESANIN, M. 2016. Hologenome analysis of two marine sponges with different microbiomes. BMC genomics, 17, 158. SAGAR, S., KAUR, M. & MINNEMAN, K. P. 2010. Antiviral lead compounds from marine sponges. Marine drugs, 8, 2619-2638. SANO, E., CARLSON, S., WEGLEY, L. & ROHWER, F. 2004. Movement of viruses between biomes. Applied and Environmental Microbiology, 70, 5842-5846. SATO, M. & KAKISAWA, H. 1976. Structures of three new C 16 terpenoids from an Acrostalagmus fungus. Journal of the Chemical Society, Perkin Transactions 1, 2407- 2413. SCHIPPERS, K. J., SIPKEMA, D., OSINGA, R., SMIDT, H., POMPONI, S. A., MARTENS, D. E. & WIJFFELS, R. H. 2012. Cultivation of sponges, sponge cells and symbionts: achievements and future prospects. Advances in marine biology. Elsevier. SCHLOSS, P. D., WESTCOTT, S. L., RYABIN, T., HALL, J. R., HARTMANN, M., HOLLISTER, E. B., LESNIEWSKI, R. A., OAKLEY, B. B., PARKS, D. H. & ROBINSON, C. J. 2009. Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Applied and environmental microbiology, 75, 7537-7541. SCHMIDT, E., OBRAZTSOVA, A., DAVIDSON, S., FAULKNER, D. & HAYGOOD, M. 2000. Identification of the antifungal peptide-containing symbiont of the marine sponge Theonella swinhoei as a novel δ-proteobacterium,“Candidatus Entotheonella palauensis”. Marine Biology, 136, 969-977. SCHMITT, S., TSAI, P., BELL, J., FROMONT, J., ILAN, M., LINDQUIST, N., PEREZ, T., RODRIGO, A., SCHUPP, P. J. & VACELET, J. 2012. Assessing the complex sponge microbiota: core, variable and species-specific bacterial communities in marine sponges. The ISME Journal, 6, 564. SEVIN, E. W. & BARLOY-HUBLER, F. 2007. RASTA-Bacteria: a web-based tool for identifying toxin-antitoxin loci in prokaryotes. Genome biology, 8, 155. SHEARER, C. A., DESCALS, E., KOHLMEYER, B., KOHLMEYER, J., MARVANOVÁ, L., PADGETT, D., PORTER, D., RAJA, H. A., SCHMIT, J. P. & THORTON, H. A. 2007. Fungal biodiversity in aquatic habitats. Biodiversity and Conservation, 16, 49-67. SIEZEN, R. J. & LEUNISSEN, J. A. 1997. Subtilases: the superfamily of subtilisin‐like serine proteases. Protein science, 6, 501-523. SILVEIRA, C. B., SILVA‐LIMA, A. W., FRANCINI‐FILHO, R. B., MARQUES, J. S., ALMEIDA, M. G., THOMPSON, C. C., REZENDE, C. E., PARANHOS, R., MOURA, R. L. & SALOMON, P. S. 2015. Microbial and sponge loops modify fish production in phase‐shifting coral reefs. Environmental microbiology, 17, 3832-3846. SIMIS, S. G., TIJDENS, M., HOOGVELD, H. L. & GONS, H. J. 2007. Optical signatures of the filamentous cyanobacterium Leptolyngbya boryana during mass viral lysis. Limnology and oceanography, 52, 184-197.

118 SINGH, P., RAGHUKUMAR, C., MEENA, R. M., VERMA, P. & SHOUCHE, Y. 2012. Fungal diversity in deep-sea sediments revealed by culture-dependent and culture-independent approaches. Fungal Ecology, 5, 543-553. SIPKEMA, D., FRANSSEN, M. C., OSINGA, R., TRAMPER, J. & WIJFFELS, R. H. 2005. Marine sponges as pharmacy. Marine Biotechnology, 7, 142-162. SMITH, A., MATSON, D., STEIN, D., SKILLING, D., KROEKER, A., BERKE, T. & IVERSEN, P. 2002. Antisense treatment of caliciviridae: an emerging disease agent of animals and humans. Current opinion in molecular therapeutics, 4, 177-184. SMITH, A. W., SKILLING, D. E., CHERRY, N., MEAD, J. H. & MATSON, D. O. 1998. Calicivirus emergence from ocean reservoirs: zoonotic and interspecies movements. Emerging Infectious Diseases, 4, 13. SOMMER, F., ANDERSON, J. M., BHARTI, R., RAES, J. & ROSENSTIEL, P. 2017. The resilience of the intestinal microbiota influences health and disease. Nature Reviews Microbiology, 15, 630. SPANIER, J. G. & CLEARY, P. P. 1980. Bacteriophage control of antiphagocytic determinants in group A streptococci. Journal of Experimental Medicine, 152, 1393-1406. SRIDHAR, K. & KAVERIAPPA, K. 1989. Water-Borne Flyphomycete Flora of Two Freshwater Streams. Environment & Ecology, 7, 771-772. SRIVASTAVA, M., SIMAKOV, O., CHAPMAN, J., FAHEY, B., GAUTHIER, M. E., MITROS, T., RICHARDS, G. S., CONACO, C., DACRE, M. & HELLSTEN, U. 2010. The Amphimedon queenslandica genome and the evolution of animal complexity. Nature, 466, 720. STERN, A. & SOREK, R. 2011. The phage‐host arms race: shaping the evolution of microbes. Bioessays, 33, 43-51. STEWART, F. M. & LEVIN, B. R. 1984. The population biology of bacterial viruses: why be temperate. Theoretical population biology, 26, 93-117. STEWART, J. R., GAST, R. J., FUJIOKA, R. S., SOLO-GABRIELE, H. M., MESCHKE, J. S., AMARAL- ZETTLER, L. A., DEL CASTILLO, E., POLZ, M. F., COLLIER, T. K. & STROM, M. S. The coastal environment and human health: microbial indicators, pathogens, sentinels and reservoirs. Environmental Health, 2008. BioMed Central, 3. SULLIVAN, M. B., COLEMAN, M. L., WEIGELE, P., ROHWER, F. & CHISHOLM, S. W. 2005. Three Prochlorococcus cyanophage genomes: signature features and ecological interpretations. PLoS biology, 3, 144. SULLIVAN, M. B., HUANG, K. H., IGNACIO‐ESPINOZA, J. C., BERLIN, A. M., KELLY, L., WEIGELE, P. R., DEFRANCESCO, A. S., KERN, S. E., THOMPSON, L. R. & YOUNG, S. 2010. Genomic analysis of oceanic cyanobacterial myoviruses compared with T4‐like myoviruses from diverse hosts and environments. Environmental microbiology, 12, 3035-3056. SUTTLE, C. A. 2005. Viruses in the sea. Nature, 437, 356-361. SUTTLE, C. A. 2007. Marine viruses—major players in the global ecosystem. Nature Reviews Microbiology, 5, 801. SUTURIN, A., TIMOSHKIN, O., PARADINA, L., KRAVTSOVA, L., ROZHKOVA, N., KULIKOVA, N., SAIBATALOVA, Y. V. & SEMITURKINA, N. 2003. Biogeochemical Processes on the Stony Littoral-Unlimited Element and Nutrient Source for Baikal Ecosystem. Berl. Palaeobiol. Abh, 4, 129-139. TAMURA, K., STECHER, G., PETERSON, D., FILIPSKI, A. & KUMAR, S. 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Molecular biology and evolution, 30, 2725- 2729. TANCA, A., ABBONDIO, M., PALOMBA, A., FRAUMENE, C., MANGHINA, V., CUCCA, F., FIORILLO, E. & UZZAU, S. 2017. Potential and active functions in the gut microbiota of a healthy human cohort. Microbiome, 5, 79. TAVARES, M. D. C. M., VOLKMER-RIBEIRO, C. & HERMANY, G. 2005. Seasonal abundance in a sponge assembly at a southern neotropical inner delta. Journal of Coastal Research, 335-342. 119 TAYLOR, M. W., RADAX, R., STEGER, D. & WAGNER, M. 2007. Sponge-Associated Microorganisms: Evolution, Ecology, and Biotechnological Potential. Microbiology and Molecular Biology Reviews, 71, 295-347. TAYLOR, M. W., SCHUPP, P. J., DAHLLÖF, I., KJELLEBERG, S. & STEINBERG, P. D. 2004. Host specificity in marine sponge‐associated bacteria, and potential implications for marine microbial diversity. Environmental Microbiology, 6, 121-130. THAKUR, N., PEROVIĆ-OTTSTADT, S., BATEL, R., KORZHEV, M., DIEHL-SEIFERT, B., MÜLLER, I. & MÜLLER, W. 2005. Innate immune defense of the sponge Suberites domuncula against gram-positive bacteria: induction of lysozyme and AdaPTin. Marine biology, 146, 271- 282. THAKUR, N. L., HENTSCHEL, U., KRASKO, A., PABEL, C. T., ANIL, A. C. & MÜLLER, W. E. 2003. Antibacterial activity of the sponge Suberites domuncula and its primmorphs: potential basis for epibacterial chemical defense. Aquatic microbial ecology, 31, 77-83. THINGSTAD, T. F., VÅGE, S., STORESUND, J. E., SANDAA, R.-A. & GISKE, J. 2014. A theoretical analysis of how strain-specific viruses can control microbial species diversity. Proceedings of the National Academy of Sciences, 201400909. THIRUNAVUKKARASU, N., SURYANARAYANAN, T. S., GIRIVASAN, K. P., VENKATACHALAM, A., GEETHA, V., RAVISHANKAR, J. P. & DOBLE, M. 2012. Fungal symbionts of marine sponges from Rameswaram, southern India: species composition and bioactive metabolites. Fungal Diversity, 55, 37-46. THOMAS, T., MOITINHO-SILVA, L., LURGI, M., BJÖRK, J. R., EASSON, C., ASTUDILLO-GARCÍA, C., OLSON, J. B., ERWIN, P. M., LÓPEZ-LEGENTIL, S. & LUTER, H. 2016. Diversity, structure and convergent evolution of the global sponge microbiome. Nature Communications, 7, 11870. THOMAS, T., RUSCH, D., DEMAERE, M. Z., YUNG, P. Y., LEWIS, M., HALPERN, A., HEIDELBERG, K. B., EGAN, S., STEINBERG, P. D. & KJELLEBERG, S. 2010. Functional genomic signatures of sponge bacteria reveal unique and shared features of symbiosis. The ISME journal, 4, 1557-1567. THOMPSON, J. E., BARROW, K. D. & FAULKNER, D. J. 1983. Localization of two brominated metabolites, aerothionin and homoaerothionin, in spherulous cells of the marine sponge Aplysina fistularis (= Verongia thiona). Acta Zoologica, 64, 199-210. THURBER, R. V. 2009. Current insights into phage biodiversity and biogeography. Current opinion in microbiology, 12, 582-587. THURBER, R. V., PAYET, J. P., THURBER, A. R. & CORREA, A. M. 2017. Virus–host interactions and their roles in coral reef health and disease. Nature Reviews Microbiology, 15, 205. TORRES, Y. R., BERLINCK, R. G., NASCIMENTO, G. G., FORTIER, S. C., PESSOA, C. & DE MORAES, M. O. 2002. Antibacterial activity against resistant bacteria and cytotoxicity of four alkaloid toxins isolated from the marine sponge Arenosclera brasiliensis. Toxicon, 40, 885-891. UHLIN, U. & EKLUND, H. 1994. Structure of ribonucleotide reductase protein R1. Nature, 370, 533. UNSON, M., HOLLAND, N. & FAULKNER, D. 1994. A brominated secondary metabolite synthesized by the cyanobacterial symbiont of a marine sponge and accumulation of the crystalline metabolite in the sponge tissue. Marine Biology, 119, 1-11. USHER, K. M., SUTTON, D. C., TOZE, S., KUO, J. & FROMONT, J. 2005. Inter-generational transmission of microbial symbionts in the marine sponge Chondrilla australiensis (Demospongiae). Marine and Freshwater Research, 56, 125-131. VACELET, J. & BOURY-ESNAULT, N. 2002. A new species of carnivorous deep-sea sponge (Demospongiae: Cladorhizidae) associated with methanotrophic bacteria. Cahiers de Biologie Marine, 43, 141-148. VACELET, J., BOURY-ESNAULT, N., FIALA-MEDIONI, A. & FISHER, C. 1995. A methanotrophic carnivorous sponge. Nature, 377, 296-296. 120 VAN DER VIJVER, J. C., VAN ES-BOON, M. & MICHEL, M. F. 1972. Lysogenic conversion in Staphylococcus aureus to leucocidin production. Journal of virology, 10, 318-319. VASU, K. & NAGARAJA, V. 2013. Diverse functions of restriction-modification systems in addition to cellular defense. Microbiology and molecular biology reviews, 77, 53-72. VILLAMIZAR, E. & LAUGHLIN, R. 1991. Fauna associated with the sponges Aplysina archeri and Aplysina lacunosa in a coral reef of the Archipielago de Los Roques, National Park, Venezuela. Fossil and recent sponges. Springer. VINCENT, M. J., SANCHEZ, A. J., ERICKSON, B. R., BASAK, A., CHRETIEN, M., SEIDAH, N. G. & NICHOL, S. T. 2003. Crimean-Congo hemorrhagic fever virus glycoprotein proteolytic processing by subtilase SKI-1. Journal of virology, 77, 8640-8649. VOGEL, G. 2008. The Inner Lives of Sponges. Science, 320, 1028-1030. VOGEL, S. 1977. Current-induced flow through living sponges in nature. Proceedings of the National Academy of Sciences, 74, 2069-2071. WAGNER, P. L. & WALDOR, M. K. 2002. Bacteriophage control of bacterial virulence. Infection and immunity, 70, 3985-3993. WANG, F.-Z., HUANG, Z., SHI, X.-F., CHEN, Y.-C., ZHANG, W.-M., TIAN, X.-P., LI, J. & ZHANG, S. 2012. Cytotoxic indole diketopiperazines from the deep sea-derived fungus Acrostalagmus luteoalbus SCSIO F457. Bioorganic & medicinal chemistry letters, 22, 7265-7267. WANG, G., LI, Q. & ZHU, P. 2008. Phylogenetic diversity of culturable fungi associated with the Hawaiian sponges Suberites zeteki and Gelliodes fibrosa. Antonie Van Leeuwenhoek, 93, 163-174. WANG, L., XUE, J., SEABORN, C. P., ARIF, B. M. & CHENG, X.-W. 2006. Sequence and organization of the Trichoplusia ni ascovirus 2c (Ascoviridae) genome. Virology, 354, 167-177. WANG, Y., TIAN, R. M., GAO, Z. M., BOUGOUFFA, S. & QIAN, P.-Y. 2014. Optimal eukaryotic 18S and universal 16S/18S ribosomal RNA primers and their application in a study of symbiosis. PloS one, 9, 90053. WARD, J. R. & LAFFERTY, K. D. 2004. The elusive baseline of marine disease: are diseases in ocean ecosystems increasing? PLoS biology, 2, 120. WATERBURY, J. B. & VALOIS, F. W. 1993. Resistance to co-occurring phages enables marine synechococcus communities to coexist with cyanophages abundant in seawater. Appl Environ Microbiol, 59, 3393-9. WEBSTER, N. & HILL, R. 2001. The culturable microbial community of the Great Barrier Reef sponge Rhopaloeides odorabile is dominated by an α-Proteobacterium. Marine Biology, 138, 843-851. WEBSTER, N. S. 2007. Sponge disease: a global threat? Environmental Microbiology, 9, 1363- 1375. WEBSTER, N. S., COBB, R. E. & NEGRI, A. P. 2008. Temperature thresholds for bacterial symbiosis with a sponge. The ISME Journal, 2, 830-842. WEBSTER, N. S., NEGRI, A. P., WEBB, R. I. & HILL, R. T. 2002. A spongin-boring α- proteobacterium is the etiological agent of disease in the Great Barrier Reef sponge Rhopaloeides odorabile. Marine Ecology Progress Series, 232, 305-309. WEBSTER, N. S. & TAYLOR, M. W. 2012. Marine sponges and their microbial symbionts: love and other relationships. Environ Microbiol, 14, 335-46. WEBSTER, N. S., TAYLOR, M. W., BEHNAM, F., LÜCKER, S., RATTEI, T., WHALAN, S., HORN, M. & WAGNER, M. 2010. Deep sequencing reveals exceptional diversity and modes of transmission for bacterial sponge symbionts. Environmental Microbiology, 12, 2070- 2082. WEBSTER, N. S. & THOMAS, T. 2016. The sponge hologenome. American Society for Microbiology, 7, 135-141.

121 WEHRL, M., STEINERT, M. & HENTSCHEL, U. 2007. Bacterial uptake by the marine sponge Aplysina aerophoba. Microbial ecology, 53, 355-365. WEIGELE, P. R., POPE, W. H., PEDULLA, M. L., HOUTZ, J. M., SMITH, A. L., CONWAY, J. F., KING, J., HATFULL, G. F., LAWRENCE, J. G. & HENDRIX, R. W. 2007. Genomic and structural analysis of Syn9, a cyanophage infecting marine Prochlorococcus and Synechococcus. Environmental microbiology, 9, 1675-1695. WEINBAUER, M. G. 2004. Ecology of prokaryotic viruses. FEMS microbiology reviews, 28, 127- 181. WEINBAUER, M. G., BONILLA-FINDJI, O., CHAN, A. M., DOLAN, J. R., SHORT, S. M., ŠIMEK, K., WILHELM, S. W. & SUTTLE, C. A. 2011. Synechococcus growth in the ocean may depend on the lysis of heterotrophic bacteria. Journal of plankton research, 33, 1465- 1476. WEINBAUER, M. G. & RASSOULZADEGAN, F. 2004. Are viruses driving microbial diversification and diversity? Environmental microbiology, 6, 1-11. WEISS, B., EBEL, R., ELBRÄCHTER, M., KIRCHNER, M. & PROKSCH, P. 1996. Defense metabolites from the marine sponge Verongia aerophoba. Biochemical Systematics and Ecology, 24, 1-12. WEITZ, J. S., STOCK, C. A., WILHELM, S. W., BOUROUIBA, L., COLEMAN, M. L., BUCHAN, A., FOLLOWS, M. J., FUHRMAN, J. A., JOVER, L. F. & LENNON, J. T. 2015. A multitrophic model to quantify the effects of marine viruses on microbial food webs and ecosystem processes. The ISME journal, 9, 1352. WEYNBERG, K. D., LAFFY, P. W., WOOD-CHARLSON, E. M., TURAEV, D., RATTEI, T., WEBSTER, N. S. & VAN OPPEN, M. J. 2017. Coral-associated viral communities show high levels of diversity and host auxiliary functions. PeerJ, 5, 4054. WHITE, T. J., BRUNS, T., LEE, S. & TAYLOR, J. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR protocols: a guide to methods and applications, 18, 315-322. WIENS, M., KORZHEV, M., KRASKO, A., THAKUR, N. L., PEROVIĆ-OTTSTADT, S., BRETER, H. J., USHIJIMA, H., DIEHL-SEIFERT, B., MÜLLER, I. M. & MÜLLER, W. E. 2005. Innate immune defense of the sponge Suberites domuncula against bacteria involves a MyD88- dependent signaling pathway induction of a perforin-like molecule. Journal of Biological Chemistry, 280, 27949-27959. WIENS, M., KUUSKSALU, A., KELVE, M. & MÜLLER, W. E. 1999. Origin of the interferon‐ inducible (2′‐5′) oligoadenylate synthetases: cloning of the (2′‐5′) oligoadenylate synthetase from the marine sponge Geodia cydonium 1. FEBS letters, 462, 12-18. WIESE, J., OHLENDORF, B., BLÜMEL, M., SCHMALJOHANN, R. & IMHOFF, J. F. 2011. Phylogenetic identification of fungi isolated from the marine sponge Tethya aurantium and identification of their secondary metabolites. Marine drugs, 9, 561-585. WILKINSON, C. 1978. Microbial associations in sponges. III. Ultrastructure of the in situ associations in coral reef sponges. Marine Biology, 49, 177-185. WILKINSON, C. & FAY, P. 1979. Nitrogen fixation in coral reef sponges with symbiotic cyanobacteria. WILKINSON, C., GARRONE, R. & VACELET, J. 1984. Marine sponges discriminate between food bacteria and bacterial symbionts: electron microscope radioautography and in situ evidence. Proc. R. Soc. Lond. B, 220, 519-528. WILKINSON, C. R. 1983. Net primary productivity in coral reef sponges. Science, 219, 410-412. WILKINSON, C. R. 1984. Immunological evidence for the Precambrian origin of bacterial symbioses in marine sponges. Proc. R. Soc. Lond. B, 220, 509-518. WILKINSON, C. R. 1987. Productivity and abundance of large sponge populations on Flinders Reef flats, Coral Sea. Coral Reefs, 5, 183-188.

122 WILLIAMSON, S., MCLAUGHLIN, M. & PAUL, J. 2001. Interaction of the ΦHSIC virus with its host: lysogeny or pseudolysogeny? Applied and environmental microbiology, 67, 1682- 1688. WILLIAMSON, S. J., ALLEN, L. Z., LORENZI, H. A., FADROSH, D. W., BRAMI, D., THIAGARAJAN, M., MCCROW, J. P., TOVCHIGRECHKO, A., YOOSEPH, S. & VENTER, J. C. 2012. Metagenomic exploration of viruses throughout the Indian Ocean. PLoS One, 7, 42047. WINTER, C., BOUVIER, T., WEINBAUER, M. G. & THINGSTAD, T. F. 2010. Trade-offs between competition and defense specialists among unicellular planktonic organisms: the “killing the winner” hypothesis revisited. Microbiology and Molecular Biology Reviews, 74, 42-57. WOMMACK, K. E. & COLWELL, R. R. 2000. Virioplankton: viruses in aquatic ecosystems. Microbiology and molecular biology reviews, 64, 69-114. WOOLHOUSE, M. E., HAYDON, D. T. & ANTIA, R. 2005. Emerging pathogens: the epidemiology and evolution of species jumps. Trends in ecology & evolution, 20, 238-244. WULFF, J. L. 2006. Ecological interactions of marine sponges. Canadian Journal of Zoology, 84, 146-166. YAHEL, G., SHARP, J. H., MARIE, D., HÄSE, C. & GENIN, A. 2003. In situ feeding and element removal in the symbiont‐bearing sponge Theonella swinhoei: Bulk DOC is the major source for carbon. Limnology and Oceanography, 48, 141-149. YAMAGUCHI, Y. & INOUYE, M. 2011. Regulation of growth and death in Escherichia coli by toxin–antitoxin systems. Nature Reviews Microbiology, 9, 779-790. YANG, Y., CAI, L., MA, R., XU, Y., TONG, Y., HUANG, Y., JIAO, N. & ZHANG, R. 2017. A novel roseosiphophage isolated from the oligotrophic South China Sea. Viruses, 9, 109. YU, Z., ZHANG, B., SUN, W., ZHANG, F. & LI, Z. 2013. Phylogenetically diverse endozoic fungi in the South China Sea sponges and their potential in synthesizing bioactive natural products suggested by PKS gene and cytotoxic activity analysis. Fungal Diversity, 58, 127-141. YUNG, P. Y., BURKE, C., LEWIS, M., KJELLEBERG, S. & THOMAS, T. 2011. Novel antibacterial proteins from the microbial communities associated with the sponge Cymbastela concentrica and the green alga Ulva australis. Applied and environmental microbiology, 77, 1512-1515. ZABLOCKI, O., VAN ZYL, L., ADRIAENSSENS, E. M., RUBAGOTTI, E., TUFFIN, M., CARY, S. C. & COWAN, D. 2014. High-level diversity of tailed phages, eukaryote-associated viruses, and virophage-like elements in the metaviromes of antarctic soils. Applied and environmental microbiology, 80, 6888-6897. ZALAR, P., GOSTINČAR, C., DE HOOG, G., URŠIČ, V., SUDHADHAM, M. & GUNDE-CIMERMAN, N. 2008. Redefinition of Aureobasidium pullulans and its varieties. Studies in Mycology, 61, 21-38. ZAN, J., FRICKE, W. F., FUQUA, C., RAVEL, J. & HILL, R. 2011. Genome sequence of Ruegeria sp. KLH11, a N-acylhomoserine lactone-producing bacterium isolated from the marine sponge Mycale laxissima. Journal of bacteriology, JB. 05556-11. ZENG, L., SKINNER, S. O., ZONG, C., SIPPY, J., FEISS, M. & GOLDING, I. 2010. Decision making at a subcellular level determines the outcome of bacteriophage infection. Cell, 141, 682- 691. ZHAN, Y., HUANG, S., VOGET, S., SIMON, M. & CHEN, F. 2016. A novel roseobacter phage possesses features of podoviruses, siphoviruses, prophages and gene transfer agents. Scientific Reports, 6, 30372. ZHANG, F., PITA, L., ERWIN, P. M., ABAID, S., LÓPEZ-LEGENTIL, S. & HILL, R. T. 2014. Symbiotic archaea in marine sponges show stability and host specificity in community structure and ammonia oxidation functionality. FEMS microbiology ecology, 90, 699-707.

123 ZHANG, L., KANG, M., HUANG, Y. & YANG, L. 2016. Fungal communities from the calcareous deep-sea sediments in the Southwest India Ridge revealed by Illumina sequencing technology. World Journal of Microbiology and Biotechnology, 32, 1-11. ZHANG, N., CASTLEBURY, L. A., MILLER, A. N., HUHNDORF, S. M., SCHOCH, C. L., SEIFERT, K. A., ROSSMAN, A. Y., ROGERS, J. D., KOHLMEYER, J. & VOLKMANN-KOHLMEYER, B. 2006. An overview of the systematics of the Sordariomycetes based on a four-gene phylogeny. Mycologia, 98, 1076-1087. ZHANG, S., CLARK, K. D. & STRAND, M. R. 2011. The protein P23 identifies capsule-forming plasmatocytes in the moth Pseudoplusia includens. Developmental & Comparative Immunology, 35, 501-510. ZHANG, Y. & JIAO, N. 2009. Roseophage RDJLΦ1, infecting the aerobic anoxygenic phototrophic bacterium Roseobacter denitrificans OCh114. Applied and environmental microbiology, 75, 1745-1749. ZHANG, Y., JIAO, N., COLQUHOUN, D. R., HALDEN, R. U. & CHEN, F. 2009. Protein modifications related to phage resistance in a marine roseobacter. Aquatic Microbial Ecology, 55, 203-207. ZHAO, Y., WANG, K., JIAO, N. & CHEN, F. 2009. Genome sequences of two novel phages infecting marine roseobacters. Environmental microbiology, 11, 2055-2064. ZHOU, K., ZHANG, X., ZHANG, F. & LI, Z. 2011. Phylogenetically diverse cultivable fungal community and polyketide synthase (PKS), non-ribosomal peptide synthase (NRPS) genes associated with the South China Sea sponges. Microbial ecology, 62, 644-654.

124 6 Appendix

6.1 Chapter Two

Table S2.1 Summary of isolation source and growth media of unique isolates (i.e. after redundant isolate sequences were removed). Media: Peptone yeast glucose agar (PYG), Potato dextrose agar (PD) and Gause 1 agar (G1). Isolate sequence ID code (media source_sample replicate number_isolate number). Isolate sequences are available through the NCBI GenBank, accession numbers are indicated in brackets.

OTU_ID C. concentrica (C) Scopalina sp. (S) T. anhelans (T) Seawater (S) OTU_9 PYG_S_2_05 (MG548542) G1_S_2_03 (MG548546) OTU_119 PYG_C_3_02 (MG548512) PYG_S_2_03 (MG548534) PD_T_3_01 (MG548514) PYG_SW_2_01 PYG_C_3_03 (MG548524) PYG_S_2_01 (MG548535) G1_T_2_02 (MG548532) (MG548513) PD_C_1_01 (MG548518) PYG_S_3_04 (MG548533) G1_T_1_01 (MG548550) G1_SW_3_01 PD_C_2_01 (MG548552) PYG_S_2_04 (MG548521) PD_T_1_01 (MG548519) (MG548544) PYG_S_1_01 (MG548529) PD_S_2_02 (MG548537) PYG_S_2_02 (MG548536) G1_S_2_01 (MG548527) G1_S_2_02 (MG548543) PYG_S_3_02 (MG548541) PD_S_1_01 (MG548523) PYG_S_3_01 (MG548547) G1_S_3_01 (MG548551) PD_S_2_01 (MG548545) OTU_165 G1_C_3_01 (MG548531) PD_S_2_03 (MG548517) PYG_C_2_02 (MG548516) PGY_C_3_01 (MG548539) PYG_C_2_01 (MG548540) G1_C_1_01 (MG548553) OTU_264 PD_C_3_01 (MG548530) G1_S_1_01 (MG548538) G1_SW_3_02 PD_C_2_02 (MG548525) (MG548526) PYG_C_2_03 (MG548522)

OTU_265 PYG_S_3_03 (MG548515) OTU_266 PYG_T_3_01 (MG548528) G1_T_2_01 (MG548520) OTU_268 PYG_T_1_01 (MG548549) OTU_269 PYG_T_1_02 (MG548548)

Table S2.2 Shannon’s index summary of sub-sampled data with ANOVA and Tukey’s multiple comparison between temporal samples and sample types. P-values are shown in brackets and significant difference taken at P-values ≤0.05 are highlighted in bold. 125 Temporal samples Shannon’s index (P-values) 2014 vs. 2016 0.29 ± 0.44 vs. 1.43 ± 0.60 (0.0005) C. concentrica 2014 vs. 2016 0.33 ± 0.49 vs. 1.54 ± 0.79 (0.15) Scopalina sp. 2014 vs. 2016 NA vs. 1.36 ± 0.037 (NA) T. anhelans 2014 vs. 2016 0.49 ± 0.66 vs. 0.90 ± 0.27 (0.97) Seawater 2014 vs. 2016 0.01 ± 0.018 vs. 1.89 ± 0.58 (0.02) Sample type Seawater vs. Scopalina sp. 1.13 ± 1.11 vs. 1.36 ± 0.037 (0.99) T. anhelans vs. Scopalina sp. 0.77 ± 0.44 vs. 1.36 ± 0.037 (0.81) T. anhelans vs. Seawater 0.77 ± 0.44 vs. 1.13 ± 1.11 (0.87) Scopalina sp. vs. C. concentrica 1.36 ± 0.037 vs. 0.94 ± 0.89 (0.92) Seawater vs. C. concentrica 1.13 ± 1.11 vs. 0.94 ± 0.89 (0.98) T. anhelans vs. C. concentrica 0.77 ± 0.44 vs. 0.94 ± 0.89 (0.98)

Table S2.3 Summary of PERMANOVA analysis of the whole fungal community (sub-sampled to 250 reads) of sponges and seawater samples in 2014 and 2016. Significant difference was taken at P-values ≤0.05 and are highlighted in bold.

Temporal samples P-value base on P-value based on

126 relative abundance presence-absence 2014 vs. 2016 0.017 0.001 C. concentrica 2014 vs. 2016 0.1 0.1 Scopalina sp. 2014 vs. 2016 NA NA T. anhelans 2014 vs. 2016 0.5 0.7 Seawater 2014 vs. 2016 0.1 0.008 Sample type C. concentrica vs. Scopalina sp. 0.4 0.51 C. concentrica vs. T. anhelans 0.29 0.24 C. concentrica vs. seawater 0.52 0.58 Scopalina sp. vs. T. anhelans 0.078 0.049 Scopalina sp. vs. seawater 0.39 0.12 T. anhelans vs. seawater 0.14 0.075

Table S2.4 Mean relative abundance percentages of OTUs in sponges (three species combined and seawater with standard deviations (sd). Enrichment ratios are calculated by the OTU mean relative abundance in sponges/ OTU mean relative abundance in seawater. The OTUs were considered as enriched in sponges if the ratio values were >2 and are highlighted in bold.

127 OTU_ID Sponges (Sps) Seawater (SW) Enrichment

mean % ± sd mean % ± sd ratio (Sps/SW)

OTU_3 9.09% ± 7.64 16.08% ± 25.96 0.6

OTU_4 11.1% ± 8.23 8.34% ± 20.41 1.3

OTU_5 8.2% ± 7.09 0.17% ± 0.19 48.2

OTU_6 33.88% ± 10.19 17.28% ± 40.46 2.0

OTU_7 3.01% ± 4.47 0.07% ± 0.09 40.5

OTU_8 0.59% ± 1.83 0.04% ± 0.05 15.6

OTU_10 6.21% ± 6.49 0.03% ± 0.03 246.9

OTU_11 1.82% ± 2.9 0.07% ± 0.1 24.4

OTU_12 1.8% ± 3.37 0.03% ± 0.05 51.6

OTU_13 0.28% ± 1.09 0.05% ± 0.12 5.1

OTU_14 1.77% ± 3.21 0.02% ± 0.04 78.7

OTU_16 1.03% ± 2.45 2.98% ± 7.29 0.3

OTU_18 1.37% ± 3.14 1.01% ± 2.43 1.4

OTU_19 0.12% ± 0.76 1.25% ± 3.04 0.1

OTU_21 0.17% ± 0.82 1.98% ± 3.07 0.1

OTU_23 0.07% ± 0.6 3.33% ± 8.13 0.0

OTU_24 0.08% ± 0.6 2.37% ± 5.81 0.0

OTU_25 1.23% ± 2.87 3.87% ± 4.93 0.3

OTU_27 0.14% ± 0.84 2.33% ± 5.71 0.1

OTU_28 0.16% ± 0.76 4.13% ± 10.11 0.0

OTU_37 0.06% ± 0.58 0.37% ± 0.89 0.2

OTU_38 0.06% ± 0.6 1.17% ± 2.85 0.1

OTU_40 0.09% ± 0.66 0.94% ± 2.31 0.1

OTU_41 5.55% ± 6.38 0% ± 0 -

OTU_45 0.21% ± 1.22 0.01% ± 0.01 24.5

OTU_50 0.17% ± 1.04 1.01% ± 2.46 0.2

OTU_51 0.05% ± 0.62 0.34% ± 0.56 0.2

OTU_59 0.08% ± 0.67 0.09% ± 0.23 0.9

OTU_60 0.09% ± 0.61 0.19% ± 0.46 0.5

OTU_63 5.76% ± 6.39 0.01% ± 0.02 516.4

OTU_87 0.05% ± 0.47 0.01% ± 0.01 7.6

OTU_112 5.7% ± 6.37 0% ± 0 -

OTU_142 0.03% ± 0.43 0.1% ± 0.25 0.3 Table S2.5 BLASTn results of the “variable/core” fungal community using the representative sequences clustered at 97% similarity against the NCBI non-redundant database.

OTU_ID Taxon classification Accession Best match Isolation source E-value Query Max UPARSE program number cover- identit

128 age y (%) (%) OTU_3 (Genus) EF060745 Sporidiolales sp. Seawater 1E- 100 100 Sporidiobolus sp. .1 LM439 134 OTU_4 (Genus) Epicoccum MF97249 Epicoccum sp. Archaeologic 1E- 59 100 sp. 8.1 strain 1a al marble 134 OTU_5 (Genus) KY322539 Aureobasidium Pinu 1E- 100 100 Aureobasidium sp. .1 pullulans heldreichii 134 fine roots OTU_6 (Genus) KY781771 Cladosporium cf. Inland sea 1E- 100 100 Cladosporium sp. .1 cladosporoides water 134 OTU_7 (Genus) Curvularia KY694447 Curvularia lunata Cotton plant 2E- 100 96 sp. .1 strain L7 113 OTU_8 (Genus) Fusarium EF488404 Fusarium sp. XL- Annona 5E- 100 99 sp. .1 D19 squamosa L. 133 OTU_10 (Order) KF800567 Uncultured fungus Indoor air 1E- 58 100 Leosporales .1 clone CMH476 133 OTU_11 (Kingdom) Fungi KP889871 Uncultured fungus Soil 1E-13 23 90 .1 clone SG020_F01 OTU_12 (Order) Agaricales KC753423 Unclutured Environment 6E- 100 98 .1 Calyptlla clone al samples 126 R1_21 OTU_13 (Genus) Beauveria AY261369 Beauveria feline Isolate 4E-90 100 89 sp. .1 strain CBS 250.34 OTU_14 (Class) JQ007418 Uncultured fungus Environment 1E- 100 100 Agaricomycetes .1 clone al samples 134 OTU_16 (Genus) LN898703 Aspergillus jensenii Clinical 1E- 100 100 Aspergillus sp. .1 samples 134 OTU_18 (Genus) JQ732918 Mycosphearella sp. Leaves 1E- 100 100 Mycospaerella sp. .1 AA-2012 134 OTU_19 (Kindom) Fungi KU16381 Uncultured fungus Environment 5E-70 99 84 2.1 clone S12T_70 al sample OTU_21 (Genus) LC133885 Mortierella sp. JCM Soil 3E-16 22 95 Mortierella sp. .1 8527 OTU_23 (Genus) MG23032 Metarhizium sp. (Plant) 1E- 100 100 Metarhizium sp. 8.1 clone A17-S Spiranthes 134 novae- zelandiae OTU_24 (Kingdom) Fungi JX384714. Uncultured fungus Environment 8E-42 100 75 1 clone 109A72941 al samples OTU_27 (Genus) HG93703 Uncultured Soil 3E- 100 99 Cryptococcus sp. 9.1 Cryptococcus 129 OTU_28 (Genus) AY301025 Cryptococcus Vishniacozy 1E- 100 96 Cryptococcus sp. .1 victoriae strain ma victoriae 116 S762

129 Table S2.5 (continued).

OTU_37 (Class) PK954735.1 Uncultured Glomus Soil 7E- 51 87 Lecanoromycetes clone cr-4 64201 24 OTU_38 (Genus) Coriolopsis KJ093492.1 Coriolopsis trogii Sapwood 5E- 100 99 sp. isolate LG1 133 OTU_40 (Class) JX042727.1 Uncultured fungus Environmental 7E- 32 85 Lecanoromycetes clone G1_FF11 samples 17 OTU_41 (Genus) Alatospora HM069492.1 Uncultured fungus Pine forest soil 1E- 100 100 sp. clone 115 Fungi_Clone_89 OTU_45 (Class) KM247690.1 Uncultured fungus Environmental 9E- 22 95 Lecanoromycetes clone OTU51 samples 16 OTU_50 (Class) MF966060.1 Fungal sp. clone Ponds 6E- 25 94 Lecanoromyctes 18 OTU_51 (Division) MF965433.1 Fungal sp. clone Ponds 3E- 21 97 Ascomycota 16 OTU_59 (Genus) Malassezia KM454161.1 Malassezia globose Homo sapiens 1E- 100 100 sp. strain 149.1 134 OTU_60 (Kingdom) Fungi KP889956.1 Uncultured fungus Soil 2E- 23 97 clone SG071_D04 18 OTU_63 (Class) JN396554.1 Uncultured fungus Environmental 1E- 33 89 Leotiomycetes clone samples 20 U_QM_090722_33_ Aa01.b1 OTU_87 (Class) FN397125.1 Uncultured fungus Environmental 7E- 21 97 Lecanoromycetes samples 17 OTU_112 (Class) MF965844.1 Fungal sp. clone Ponds 3E- 22 94 Agaricomycetes 15 OTU_142 (Class) MF965546.1 Fungal sp. clone Ponds 3E- 71 97 Agaricomycetes 21

130 6.2 Chapter Three

Figure S3.1 Viral communities classified to the family-level (where applicable) are clustered based on Bray-Curtis dissimilarities of taxonomic profiles (tree scale indicates dissimilarity percentages). Values are normalised to viral contigs per prokaryotic genome transformed with fourth root. A, B and C indicate sample replicates for sponges and 01, 02 and 08 indicate replicates for seawater samples.

131

Figure S3.2 Viral metagenome communities of sponges and seawater classified to the lowest classification level (species) where applicable. Sample were clustered based on Bray-Curtis dissimilarity (tree scale indicates dissimilarity percentages). Values are normalised to viral contigs per prokaryotic genome transformed with fourth root. A, B and C indicate sample replicates and 01, 02 and 08 indicate replicates for seawater samples.

132 Table S3.1 Shannon’s indices of viral assemblages associated with the sponge microbial cell fraction and the Tukey’s multiple comparison of Shannon’s indices. Significant differences are taken at P ≤ 0.05 and are highlighted in bold.

Shannon’s indices ± standard deviations

Sample C. C. R. Scopalina Stylissa sp. Seawater T. anhelans concentrica coralliophila odorabile sp. 445

Family-level 0.78 ± 0.22 1.34 ± 0.34 1.21 ± 0.46 1.54 ± 0.14 0.25 ± 0.36 1.6 ± 0.15 1.26 ± 0.50

Species-level 1.58 ± 0.28 1.85 ± 0.85 1.60 ± 0.80 2.23 ± 0.41 0.96 ± 0.45 2.63 ± 0.69 1.67 ± 0.69

Tukey multiple comparison of Shannon’s indices means (P-values of family-level, species-level)

C. concentrica 0.43, 0.99 0.69, 1.00 0.15, 0.81 0.61, 0.89 0.11, 0.36 0.60, 0.99

C. coralliophila 0.99, 0.99 0.98, 0.99 0.04, 0.63 0.95, 0.67 0.99, 0.99

R. odorabile 0.88, 0.83 0.08, 0.88 0.78, 0.37 0.99, 0.99

Scopalina sp. 0.01, 0.27 0.99, 0.97 0.93, 0.89

Stylissa sp. 445 0.009, 0.08 0.06, 0.82

Seawater 0.85, 0.45

Table S3.2 Summary of PERMANOVA analysis of pair-wise comparisons of viral assemblages (at a taxonomic family level; species level) of microbial cell associated with sponges. Values indicated P-values and significant differences were taken at P ≤ 0.05.

Samples C. C. R. odorabile Scopalina Stylissa sp. Seawater T. anhelans concentrica coralliophila sp. 445

C. concentrica 0.5, 01 0.1, 01 0.1, 01 0.7, 0.1 0.2, 0.2 0.1, 0.1

C. 0.4, 0.2 0.7, 0.3 0.3, 0.1 0.8, 0.8 0.3, 0.5 coralliophila

R. odorabile 0.1, 0,1 0.1, 0.1 0.5, 0.1 0.2, 0.1

Scopalina sp. 0.2, 0.1 0.3, 0.1 0.1, 0.2

Stylissa sp. 0.2, 0.1 0.1, 0.1 445

Seawater 0.1, 0.1

Table S3.3 Percentage of viral species that overlap with those found in seawater and percentage of unique viral species in each sample.

133 Host environment C. C. R. Scopalina Stylissa T. Seawater concentrica coralliophila odorabile sp. sp. 445 anhelans

No. of assigned viral 19 24 16 27 5 9 72 species

Percentage of viral species 25% 37.5% 12.5% 26% 60% 11% NA in common with seawater

Percentage of unique viral 68% 50% 75% 63% 40% 66% 75% species

Figure S3.3 NMDS plot base on Bray-Curtis similarity of viral community composition on a family-level taxonomic assignment in sponges and seawater. Host and environments are represented in different colours as shown in the legend. Sponges from tropical waters are indicated with a circle symbol and samples from temperate waters are indicated by a plus symbol.

134

Figure S3.4 Heatmap of the top most abundant 35 proteins hits against the PFAM database. Values are normalised to copies per genome with fourth root transformation. Columns are clustered using the Bray-Curtis dissimilarities using the ‘average method’ and scale indicates percentages of dissimilarity. A, B and C indicate sponge replicates of sponges and 01, 02 and 08 indicate seawater replicates.

135

Figure S3.5 Heatmap of the top most abundant 50 proteins hits against the TIGRFAM database. Values are normalised to copies per genome with fourth root transformation. Columns are clustered using the Bray-Curtis dissimilarities using the ‘average method’ and scale indicates dissimilar percentages. A, B and C indicate sponge replicates and 01 and 02 indicate seawater replicates.

136

Figure S3.6 Heatmap of the top most abundant 50 proteins hits against the COG database. Values are normalised to copies per genome with fourth root transformation. Columns are clustered using the Bray-Curtis dissimilarities using the ‘average method’ and scale indicates dissimilar percentages. A, B and C indicate sponge replicates and 01, 02 and 08 indicate seawater replicates.

137

Figure S3.7 Heatmap of proteins hits (E-value of 1-10) against the KEGG database). Values are normalised to copies per genomes with fourth root transformation. Columns are clustered using the Bray-Curtis dissimilarities using the ‘average method’ and scale indicates dissimilar percentages. A, B and C indicate sponge replicates and 01, 02 and 08 indicate seawater replicates.

138

Figure S3.8 Heatmap of the top most abundant 35 proteins hits against the COG database in the A) viral (meta)transcriptome B) and the whole (meta)transcriptome. Values are normalised to transcripts per million (TPM) with natural log transformation. Samples are clustered using the Bray-Curtis dissimilarities using the “average method” and scale indicates dissimilar percentages. 1, 2 and 3 represent sample replicates. For transcriptome data, the three replicates were pooled together.

139

Figure S3.9 Heatmap of the top most abundant 35 proteins hits against the PFAM database in the A) viral (meta)transcriptome and B) the whole (meta)transcriptome. Values are normalised to transcripts per million (TPM) with natural log transformation. Samples are clustered using the Bray-Curtis dissimilarities using the “average method” and scale indicates dissimilar percentages. 1, 2 and 3 represent sample replicates. For transcriptome data, the three replicates were pooled together.

140

Figure S3.10 Heatmap of the top most abundant 35 proteins hits against the TIRGFAM database in A) viral (meta)transcriptome and B) the whole (meta)transcriptome. Values are normalised to transcripts per million (TPM) with natural log transformation. Samples are clustered using the Bray-Curtis dissimilarities using the “average method” and scale indicates dissimilar percentages. 1, 2 and 3 represent sample replicates. For transcriptome data, the three replicates were pooled together.

141

Table S3.4 Summary of proteins identified in the TIGRFAM annotation system associated with thiamine metabolism in the metatranscriptome of C. concentrica.

TIGRFAM ID Protein name

TIGR01976 Am_tr_V_VC1184: cysteine desulfurase family protein

TIGR01977 SufS: cysteine desulfurase

TIGR01979 IscS: cysteine desulfurase IscS

TIGR02006 DNA_S_dndA: cysteine desulfurase DndA

TIGR03235 FeS_syn_CsdA: cysteine desulfurase

TIGR03392 Catalytic subunit CsdA

TIGR03402 FeS_nifS: cysteine desulfurase NifS

TIGR03403 NifS_epsilon: cysteine desulfurase

TIGR03812 Tyr_de_CO2_Arch: tyrosine decarboxylase MnfA

TIGR04343 EgtE_PLP_lyase: ergothioneine biosynthesis PLP-dependent enzyme EgtE

142

Figure S4.2 KEGG pathway map of thiamine metabolism and annotations are sourced from https://www.genome.jp/kegg-bin/show_pathway?map=map00730&show_description=show. Tyrosine decarboxylase belongs to the tyrosine synthesis pathway which catalyses the production of L-tyrosine (highlighted in yellow). Cysteine desulfurase (IscS) is involved in transferring the sulfur from L-cysteine to ThiS (forming ThiS-COSH), which incorporates the sulfur into the thiazole ring of thiamine (highlighted in red).

143 Table S3.5 Identification of functional groups of viral contigs in the metagenome.

Contig ID Sample COG KEGG PFAM TIGRFAM CynA_contig08608 CynA COG0059 K00053 PF01450.18;PF07991.11 TIGR00465 CynA_contig08608 CynA COG0854 K03474 PF03740.12 TIGR00559 CynA_contig08608 CynA COG0507 K01144 PF13538.5 TIGR01447;TIGR01448 CynA_contig08608 CynA COG0507 K01144 NA TIGR01448 CynA_contig08608 CynA COG0507 K01144 NA NA CynA_contig08608 CynA COG0440 K01653 PF01842.24;PF13710.5;PF10369.8 TIGR00119 CynA_contig08608 CynA COG0028 K01652 PF02775.20;PF00205.21;PF02776.17 TIGR00118;TIGR00173;TIGR01504;TIGR02418;TIGR02720;TIGR03 254;TIGR03393;TIGR03394;TIGR03457;TIGR04377 CynA_contig08608 CynA NA NA PF12183.7 NA CynA_contig08608 CynA COG0338 K06223 PF02086.14 NA CynA_contig10340 CynA COG0338 K06223 PF02086.14 TIGR00571 CynA_contig10340 CynA COG2189 K07319 PF01555.17 NA CynA_contig10340 CynA NA NA PF09195.10 NA CynA_contig12772 CynA NA K21367 PF01531.15 NA CynA_contig12772 CynA COG1089 K01711 PF01370.20;PF16363.4 TIGR01179;TIGR01181;TIGR01472;TIGR02622;TIGR04180 CynA_contig13406 CynA COG4799 K01966 PF01039.21 TIGR00515;TIGR01117;TIGR03133 CynA_contig13406 CynA COG0542 K03695 PF00004.28;PF07724.13;PF07728.13;PF10431.8 TIGR02639;TIGR03345;TIGR03346 CynA_contig13406 CynA COG0542 K03695 NA TIGR02639;TIGR03345;TIGR03346 CynA_contig13406 CynA COG0542 K03695 PF02861.19 TIGR02639;TIGR03345;TIGR03346 CynB_contig00172 CynB COG2801 K07497 PF02914.14;PF09039.10;PF09299.10 NA CynB_contig00172 CynB COG2842 K07132 PF13401.5;PF09077.10 NA CynB_contig00172 CynB COG0593 K02313 PF08299.10 TIGR00362 CynB_contig00172 CynB NA NA PF14216.5 NA CynB_contig00172 CynB COG2932 NA NA NA CynB_contig00172 CynB COG4382 NA PF06252.11 NA CynB_contig00172 CynB COG3772 K01185 PF01471.17 TIGR02869 CynB_contig00172 CynB NA NA PF10805.7 NA CynB_contig00172 CynB NA NA PF11985.7 NA CynB_contig00172 CynB COG4373 NA NA NA CynB_contig00172 CynB COG4383 NA PF06074.11 NA CynB_contig00172 CynB NA K07724 PF13693.5 NA CynB_contig00172 CynB COG2369 NA PF04233.13 TIGR01641 CynB_contig00172 CynB COG1475 NA NA NA CynB_contig00172 CynB COG4388 NA PF10123.8 NA CynB_contig00172 CynB COG4397 NA PF10124.8 NA CynB_contig00172 CynB COG4387 NA PF07030.11 NA CynB_contig00172 CynB COG5005 NA PF05069.12 TIGR01635

144 CynB_contig00172 CynB NA NA PF08809.10 NA CynB_contig00172 CynB COG1475 K03497 PF02195.17 TIGR00180;TIGR03454;TIGR03734;TIGR04285 CynB_contig00172 CynB NA NA PF00486.27 NA CynB_contig01795 CynB NA NA PF00462.23 TIGR02196 CynB_contig01795 CynB COG0553 K08282 PF00271.30;PF00176.22 NA CynB_contig02890 CynB COG0338 K06223 PF02086.14 NA CynB_contig04786 CynB COG1702 K06217 PF02562.15 NA CynB_contig10133 CynB NA NA PF06067.10 NA CynB_contig13589 CynB COG1198 K04066 NA TIGR00595 CynB_contig13589 CynB COG0458 K01955 NA TIGR01369 CynB_contig13589 CynB COG0458 K01955 PF15632.5;PF02786.16;PF02787.18 TIGR00514;TIGR00768;TIGR00877;TIGR01142;TIGR01205;TIGR01 235;TIGR01369;TIGR02068;TIGR02144;TIGR02712 CynB_contig13589 CynB COG0505 K01956 PF00117.27 TIGR00566;TIGR00888;TIGR01368 CynB_contig13589 CynB COG0192 K00789 PF02773.15;PF02772.15;PF00438.19 TIGR01034 CynB_contig13589 CynB COG0452 K13038 PF04127.14;PF02441.18 TIGR00521;TIGR02113;TIGR02114 CynB_contig13589 CynB COG1758 K03060 PF01192.21 TIGR00690 CynB_contig13589 CynB COG0194 K00942 PF00625.20 TIGR02322;TIGR03263 CynB_contig13589 CynB COG2759 K01938 PF01268.18 NA CynB_contig13589 CynB COG0458 K01955 NA TIGR01369 CynB_contig17902 CynB NA NA PF03796.14 TIGR03600 CynC_contig03202 CynC COG0542 K03695 PF00004.28;PF13191.5;PF07724.13;PF07728.13;PF02 TIGR02639;TIGR03345;TIGR03346 861.19 CynC_contig03202 CynC COG0542 K03695 PF10431.8 TIGR02639;TIGR03345;TIGR03346 CynC_contig03202 CynC COG0265 K08070 PF13180.5;PF00089.25;PF13365.5 TIGR02037;TIGR02038 CynC_contig03202 CynC COG0432 NA PF01894.16 TIGR00149 CynC_contig03202 CynC COG2206 NA NA NA CynC_contig04879 CynC COG0745 K02483 PF00072.23;PF00486.27 TIGR01387;TIGR01818;TIGR02154;TIGR02875;TIGR03787 CynC_contig04879 CynC COG5002 K02484 PF00672.24;PF02518.25;PF00512.24 TIGR01386;TIGR02916;TIGR02938;TIGR02956;TIGR02966;TIGR03 785 CynC_contig04879 CynC COG0265 K04771 PF00595.23;PF13180.5;PF00089.25;PF13365.5 TIGR02037;TIGR02038 CynC_contig04879 CynC COG0568 K03086 PF04542.13;PF04539.15;PF04545.15 TIGR02392;TIGR02393;TIGR02394;TIGR02479;TIGR02835;TIGR02 846;TIGR02850;TIGR02885;TIGR02937;TIGR02941;TIGR02980;TI GR02997 CynC_contig04879 CynC COG1770 K01354 PF00326.20 NA CynC_contig04879 CynC COG1770 K01354 NA NA CynC_contig18440 CynC COG5323 NA NA NA CynC_contig18440 CynC COG1783 K06909 NA NA CynC_contig24800 CynC COG0629 K03111 NA TIGR00621 CynC_contig24800 CynC COG0629 K03111 PF00436.24 TIGR00621 CynC_contig24800 CynC COG0583 NA NA TIGR03418 CynC_contig24800 CynC COG0583 K03566 PF00126.26;PF03466.19 TIGR02036;TIGR02424;TIGR03298;TIGR03339;TIGR03418

145 CynC_contig35303 CynC COG5412 NA NA NA CynC_contig35303 CynC COG3499 K06906 PF06995.10 NA CynC_contig35303 CynC COG5004 NA PF05489.11 NA CynC_contig35303 CynC COG3500 K06905 NA NA CynC_contig35303 CynC COG3500 K06905 PF05954.10 NA CynC_contig35429 CynC COG2369 NA PF04233.13 NA CynC_contig35429 CynC COG4383 NA PF06074.11 NA CynC_contig35429 CynC NA NA PF06074.11 NA CynC_contig36463 CynC COG0468 K04483 PF08423.10;PF00154.20 TIGR02012;TIGR02236;TIGR02237;TIGR02238;TIGR02239 CynC_contig38160 CynC NA NA PF05135.12 TIGR02215 CynC_contig38160 CynC NA NA PF11367.7 NA CynC_contig38160 CynC COG1403 K07451 NA NA CynC_contig38160 CynC COG4626 NA PF03354.14 NA CynC_contig38160 CynC COG4695 NA PF04860.11 TIGR01537 CynC_contig38160 CynC COG3740 K06904 PF04586.16 TIGR01543 CynC_contig38160 CynC COG4653 K06919 PF05065.12 TIGR01554 CynC_contig45324 CynC NA NA PF00521.19 NA CynC_contig45324 CynC COG0188 K02469 PF00521.19 TIGR01061;TIGR01062;TIGR01063 CynC_contig45332 CynC COG2171 K00674 PF00132.23;PF14805.5 TIGR00965;TIGR03532;TIGR03570 CynC_contig45332 CynC COG0624 K01439 PF07687.13;PF01546.27;PF04389.16 TIGR01246;TIGR01892;TIGR01900;TIGR01902;TIGR01910;TIGR03 526 CynC_contig45332 CynC NA NA PF09195.10 NA CynC_contig45332 CynC COG2189 K07319 PF01555.17 NA CynC_contig45332 CynC COG0338 K06223 PF02086.14 TIGR00571 CynC_contig45332 CynC COG1974 K01356 PF00717.22 TIGR00498 CynC_contig58819 CynC NA NA PF05869.10 NA CynC_contig58819 CynC COG1074 K19465 NA NA CynC_contig58819 CynC COG1783 NA NA NA CyrA_contig00230 CyrA COG4805 K01322 PF05960.10 NA CyrA_contig13025 CyrA NA NA PF05658.13 NA CyrA_contig13025 CyrA COG3023 K01447 PF01510.24 NA CyrA_contig13025 CyrA NA NA PF00166.20 NA CyrA_contig13025 CyrA COG0459 K04077 PF00118.23 TIGR02339;TIGR02340;TIGR02341;TIGR02342;TIGR02343;TIGR02 344;TIGR02345;TIGR02346;TIGR02347;TIGR02348 CyrA_contig13025 CyrA COG2105 NA PF06094.11 NA CyrA_contig13025 CyrA NA NA PF06378.10 NA CyrA_contig13025 CyrA NA NA PF12684.6 NA CyrA_contig13025 CyrA NA NA PF08800.9 NA CyrA_contig13025 CyrA COG3378 NA NA NA CyrA_contig13025 CyrA COG0553 NA NA NA CyrB_contig00102 CyrB COG3108 NA PF08291.10 NA

146 CyrB_contig00102 CyrB NA NA PF11351.7 NA CyrB_contig00102 CyrB NA NA PF16684.4 NA CyrB_contig00102 CyrB COG5545 K06919 PF05272.10 NA CyrB_contig00102 CyrB NA NA NA TIGR01547 CyrB_contig02187 CyrB COG1783 NA NA NA CyrB_contig02187 CyrB COG3772 K01185 NA NA CyrB_contig02187 CyrB NA NA PF07087.10 NA CyrB_contig02187 CyrB NA NA PF05866.10 NA CyrB_contig02187 CyrB COG1061 K19789 PF04851.14 TIGR04095 CyrB_contig02187 CyrB NA NA PF13481.5 NA CyrB_contig02973 CyrB COG3299 NA NA NA CyrB_contig02973 CyrB COG1783 K06909 PF04466.12 TIGR01547 CyrB_contig02973 CyrB COG0338 K06223 NA NA CyrB_contig02973 CyrB COG3567 K09961 PF06381.10 TIGR01555 CyrB_contig02973 CyrB COG2369 NA PF04233.13 TIGR01641 CyrB_contig02973 CyrB NA NA PF09950.8 NA CyrB_contig02973 CyrB NA NA PF11041.7 NA CyrB_contig02973 CyrB NA NA NA TIGR01760 CyrB_contig05153 CyrB COG1074 K19465 NA NA CyrB_contig05867 CyrB COG3772 K01185 NA NA CyrB_contig09182 CyrB COG1475 K03497 PF02195.17 TIGR00180;TIGR03454;TIGR03734;TIGR04285 CyrB_contig09182 CyrB COG1195 K03629 PF13175.5;PF13476.5;PF02463.18 TIGR00611 CyrB_contig09182 CyrB COG0187 K02470 PF02518.25 TIGR01055;TIGR01058;TIGR01059 CyrB_contig09182 CyrB COG0187 K02470 PF00204.24;PF00986.20;PF01751.21 TIGR01055;TIGR01058;TIGR01059 CyrB_contig09182 CyrB COG0188 K02469 NA TIGR01062;TIGR01063 CyrB_contig09182 CyrB COG1192 K03496 PF13614.5;PF02374.14;PF01656.22;PF06564.11;PF00 TIGR01007;TIGR01287;TIGR01968;TIGR01969;TIGR03371;TIGR03 142.17;PF10609.8 453 CyrB_contig09182 CyrB NA NA PF02527.14 TIGR00138 CyrB_contig09182 CyrB COG1847 K06346 PF01424.21 NA CyrB_contig09182 CyrB COG0706 K03217 PF02096.19 TIGR03592 CyrB_contig09182 CyrB COG0759 K08998 PF01809.17 TIGR00278 CyrB_contig09182 CyrB COG0593 K02313 PF00308.17;PF08299.10;PF01695.16 TIGR00362;TIGR03420 CyrB_contig09182 CyrB COG0592 K02338 PF00712.18;PF02767.15;PF02768.14 TIGR00663 CyrB_contig09386 CyrB COG0207 K00560 PF00303.18 TIGR03283;TIGR03284 CyrB_contig09386 CyrB COG0358 K17680 PF13155.5 TIGR01391 CyrB_contig09386 CyrB NA NA PF12684.6 NA CyrB_contig09386 CyrB NA NA PF04404.11 NA CyrB_contig09386 CyrB NA NA NA TIGR02393;TIGR02937;TIGR02980;TIGR02997 CyrB_contig10565 CyrB NA NA PF00166.20 NA CyrB_contig10565 CyrB COG0459 K04077 PF00118.23 TIGR02348 CyrB_contig10565 CyrB COG0459 K04077 PF00118.23 TIGR02339;TIGR02348

147 CyrB_contig12817 CyrB NA NA PF08800.9 NA CyrB_contig12817 CyrB NA NA PF12684.6 NA CyrB_contig12817 CyrB NA NA PF06378.10 NA CyrB_contig22402 CyrB NA NA NA TIGR01547 CyrB_contig22402 CyrB COG4805 NA PF05960.10 NA CyrB_contig24754 CyrB COG1694 NA NA NA CyrB_contig24754 CyrB NA NA PF06356.10 NA CyrB_contig24756 CyrB NA NA PF00303.18 NA CyrB_contig24756 CyrB COG0305 K17680 NA NA CyrB_contig25493 CyrB COG0208 K00526 PF00268.20 TIGR04171 CyrB_contig26167 CyrB COG1793 K10747 PF01068.20 TIGR00574;TIGR02779 CyrB_contig30369 CyrB COG0208 K00526 PF00268.20 TIGR04171 CyrB_contig37240 CyrB COG0459 K04077 PF00118.23 TIGR02339;TIGR02340;TIGR02341;TIGR02342;TIGR02343;TIGR02 345;TIGR02348 CyrB_contig37240 CyrB COG0459 K04077 PF00118.23 TIGR02348 CyrB_contig40757 CyrB NA NA PF04404.11 NA CyrB_contig40757 CyrB NA NA PF12684.6 NA CyrB_contig40757 CyrB NA K17680 NA NA CyrC_contig00238 CyrC NA NA PF11351.7 NA CyrC_contig00238 CyrC COG3108 NA PF08291.10 NA CyrC_contig02028 CyrC NA NA PF04404.11 NA CyrC_contig02028 CyrC NA NA PF04404.11 NA CyrC_contig02036 CyrC NA NA PF02037.26 NA CyrC_contig04265 CyrC COG0338 K06223 PF02086.14 TIGR00571 RhoA_F5ZQ6II01EHCV4 RhoA COG0745 K07658 NA TIGR01387;TIGR02154;TIGR03787 RhoA_F5ZQ6II01EHCV4 RhoA COG0745 K07668 PF00072.23 TIGR02154;TIGR02875 RhoA_F5ZQ6II01EROAC RhoA NA NA PF07230.10 NA RhoA_contig02659 RhoA COG0006 NA PF00557.23 NA RhoA_contig02659 RhoA COG1011 K01560 PF13419.5 TIGR01428;TIGR01493 RhoA_contig02659 RhoA COG2801 K07497 PF13333.5;PF13683.5 NA RhoA_contig02659 RhoA COG2826 K07482 PF13936.5 NA RhoA_contig02659 RhoA COG2801 K07497 PF00665.25 NA RhoA_contig25098 RhoA COG1783 K06909 NA TIGR01547 RhoA_contig25098 RhoA NA NA PF07460.10 TIGR01453 RhoA_contig27050 RhoA COG3666 K07487 PF05598.10 NA RhoA_contig27050 RhoA COG3666 NA PF01609.20;PF13751.5 NA RhoA_contig35769 RhoA COG3588 K01623 NA NA RhoA_contig37159 RhoA COG0582 NA NA NA RhoA_contig37159 RhoA COG0582 NA NA NA RhoA_contig37598 RhoA COG4974 K04763 PF00589.21 TIGR02224;TIGR02225;TIGR02249 RhoA_contig37598 RhoA COG0582 NA PF00589.21 TIGR02224;TIGR02225

148 RhoA_contig37598 RhoA COG4974 K03733 PF00589.21;PF02899.16 TIGR02224;TIGR02225;TIGR02249 RhoA_contig37705 RhoA COG3436 K07484 PF01527.19;PF05717.12 NA RhoA_contig37705 RhoA COG2963 K07484 PF03050.13;PF13007.6;PF13005.6 NA RhoA_contig37705 RhoA NA K07484 PF03050.13 NA RhoA_contig37705 RhoA COG4372 K07484 PF03050.13;PF13817.5 NA RhoA_contig43551 RhoA COG0582 NA PF00589.21 NA RhoA_contig43551 RhoA COG0582 NA NA NA RhoA_contig55073 RhoA COG2801 K07497 PF13276.5;PF00665.25;PF13683.5 NA RhoA_contig55073 RhoA COG2963 K07483 PF01527.19 NA RhoA_contig55943 RhoA NA K03581 PF13604.5 TIGR01447;TIGR01448 RhoA_contig55943 RhoA COG0507 K03581 PF13538.5 TIGR01447;TIGR01448;TIGR02768 RhoA_contig60442 RhoA COG0582 NA NA NA RhoA_contig60442 RhoA COG0582 NA PF13356.5 NA RhoB_contig02701 RhoB NA K01155 PF09019.10 NA RhoB_contig02701 RhoB COG3727 K07458 PF03852.14 TIGR00632 RhoB_contig02753 RhoB COG0410 K01996 PF13304.5;PF00005.26 TIGR00955;TIGR00956;TIGR00958;TIGR00968;TIGR00972;TIGR01 166;TIGR01184;TIGR01186;TIGR01187;TIGR01188;TIGR01189;TI GR01192;TIGR01193;TIGR01257;TIGR01271;TIGR01277;TIGR012 88;TIGR01842;TIGR01846;TIGR01978;TIGR02142;TIGR02203;TIG R02204;TIGR02211;TIGR02314;TIGR02315;TIGR02323;TIGR0232 4;TIGR02633;TIGR02673;TIGR02769;TIGR02770;TIGR02857;TIGR 02868;TIGR02982;TIGR03005;TIGR03258;TIGR03265;TIGR03269; TIGR03375;TIGR03410;TIGR03411;TIGR03415;TIGR03522;TIGR03 608;TIGR03719;TIGR03740;TIGR03771;TIGR03796;TIGR03797;TI GR03864;TIGR03873;TIGR04406;TIGR04520;TIGR04521 RhoB_contig02753 RhoB COG0411 K01995 PF00005.26;PF12399.7 TIGR00955;TIGR00958;TIGR00968;TIGR00972;TIGR01166;TIGR01 184;TIGR01186;TIGR01187;TIGR01188;TIGR01189;TIGR01192;TI GR01193;TIGR01194;TIGR01257;TIGR01277;TIGR01288;TIGR018 42;TIGR01846;TIGR01978;TIGR02142;TIGR02203;TIGR02204;TIG R02211;TIGR02314;TIGR02315;TIGR02323;TIGR02324;TIGR0263 3;TIGR02673;TIGR02769;TIGR02770;TIGR02857;TIGR02868;TIGR 02982;TIGR03005;TIGR03258;TIGR03265;TIGR03269;TIGR03375; TIGR03410;TIGR03411;TIGR03415;TIGR03522;TIGR03608;TIGR03 719;TIGR03740;TIGR03771;TIGR03796;TIGR03797;TIGR03864;TI GR03873;TIGR04406;TIGR04520;TIGR04521 RhoB_contig24396 RhoB COG0488 K06158 PF00005.26;PF12848.6 TIGR00958;TIGR00968;TIGR00972;TIGR01166;TIGR01184;TIGR01 186;TIGR01188;TIGR01189;TIGR01193;TIGR01277;TIGR01288;TI GR01842;TIGR01846;TIGR01978;TIGR02142;TIGR02204;TIGR022 11;TIGR02315;TIGR02324;TIGR02633;TIGR02673;TIGR02769;TIG R02770;TIGR02857;TIGR02868;TIGR02982;TIGR03005;TIGR0325 8;TIGR03265;TIGR03269;TIGR03375;TIGR03410;TIGR03411;TIGR 03522;TIGR03608;TIGR03719;TIGR03740;TIGR03771;TIGR03796;

149 TIGR03797;TIGR03864;TIGR03873;TIGR04406;TIGR04520;TIGR04 521 RhoB_contig24396 RhoB NA NA PF01047.21;PF12802.6 NA RhoB_contig24396 RhoB COG1595 K03088 PF07638.10;PF04542.13;PF04545.15;PF08281.11 TIGR02937;TIGR02939;TIGR02943;TIGR02947;TIGR02948;TIGR02 950;TIGR02952;TIGR02954;TIGR02957;TIGR02959;TIGR02960;TI GR02983;TIGR02984;TIGR02985;TIGR02989 RhoB_contig24396 RhoB COG0568 K03086 PF04542.13;PF04545.15 TIGR02392;TIGR02393;TIGR02394;TIGR02479;TIGR02835;TIGR02 846;TIGR02850;TIGR02885;TIGR02937;TIGR02941;TIGR02980;TI GR02997 RhoC_contig02058 RhoC COG2963 K07483 PF01527.19 NA RhoC_contig02058 RhoC COG2801 K07497 PF13276.5;PF00665.25;PF13683.5 NA RhoC_contig05466 RhoC COG2801 K07497 PF13276.5;PF00665.25;PF13333.5;PF13683.5 NA RhoC_contig05466 RhoC COG2963 K07483 PF01527.19 NA SW01_F4H3OAQ02HCRW0 SW01 COG1209 NA NA NA SW01_F4H3OAQ02HRXCC SW01 COG0542 K03696 NA TIGR02639;TIGR03345;TIGR03346 SW01_F4H3OAQ02HRXCC SW01 COG0542 K03696 PF07724.13 TIGR02639;TIGR03345;TIGR03346 SW01_F6S3EYI01A7DHP SW01 NA NA PF00574.22 NA SW01_F6S3EYI01A91CY SW01 NA NA PF08291.10 NA SW01_F6S3EYI01AQJXC SW01 COG1327 K07738 NA TIGR00244 SW01_F6S3EYI01AQJXC SW01 COG1327 K07738 NA TIGR00244 SW01_F6S3EYI01AS4XJ SW01 COG0112 K00600 PF00464.18 NA SW01_F6S3EYI01AS4XJ SW01 COG0112 K00600 PF00464.18 NA SW01_F6S3EYI01AT9HV SW01 NA NA PF02562.15 NA SW01_F6S3EYI01AT9HV SW01 COG0678 K03386 NA NA SW01_F6S3EYI01BBP1X SW01 COG0187 K02622 NA TIGR01055;TIGR01058;TIGR01059 SW01_F6S3EYI01BBP1X SW01 COG0187 K02622 NA TIGR01055;TIGR01058;TIGR01059 SW01_F6S3EYI01CET6D SW01 COG0162 K01866 PF00579.24 TIGR00234 SW01_F6S3EYI01CNVHB SW01 COG1192 K03496 PF13614.5;PF01656.22;PF10609.8 TIGR03453 SW01_contig00096 SW01 COG1403 NA PF01844.22;PF14279.5 NA SW01_contig01716 SW01 COG3023 K01447 PF01510.24 NA SW01_contig01811 SW01 COG4254 NA NA NA SW01_contig02031 SW01 COG0272 K01972 PF00533.25;PF12826.6 TIGR00575 SW01_contig02031 SW01 COG1131 K01990 PF00005.26 TIGR00968;TIGR00972;TIGR01166;TIGR01184;TIGR01186;TIGR01 188;TIGR01189;TIGR01288;TIGR02203;TIGR02204;TIGR02211;TI GR02315;TIGR02673;TIGR02857;TIGR03265;TIGR03375;TIGR034 10;TIGR03411;TIGR03522;TIGR03608;TIGR03740;TIGR03771;TIG R03864;TIGR03873;TIGR04406;TIGR04520;TIGR04521 SW01_contig02031 SW01 COG1131 K01990 NA TIGR00955;TIGR00958;TIGR00968;TIGR00972;TIGR01166;TIGR01 184;TIGR01186;TIGR01187;TIGR01188;TIGR01189;TIGR01193;TI GR01257;TIGR01277;TIGR01288;TIGR01842;TIGR01846;TIGR019 78;TIGR02142;TIGR02203;TIGR02204;TIGR02211;TIGR02314;TIG R02315;TIGR02323;TIGR02633;TIGR02673;TIGR02769;TIGR0277 150 0;TIGR02857;TIGR02868;TIGR02982;TIGR03005;TIGR03258;TIGR 03265;TIGR03269;TIGR03375;TIGR03410;TIGR03411;TIGR03415; TIGR03522;TIGR03608;TIGR03740;TIGR03771;TIGR03796;TIGR03 864;TIGR03873;TIGR04406;TIGR04520;TIGR04521 SW01_contig02661 SW01 COG0389 K03502 PF00817.19 NA SW01_contig02661 SW01 COG0389 K03502 PF00817.19 NA SW01_contig02661 SW01 COG1974 K03503 PF00717.22 NA SW01_contig03517 SW01 COG1694 NA NA NA SW01_contig04365 SW01 NA K17680 NA NA SW01_contig04407 SW01 NA NA NA TIGR00138 SW01_contig04407 SW01 COG1192 K03496 PF13614.5;PF01656.22;PF10609.8 TIGR03371;TIGR03453 SW01_contig04407 SW01 COG1192 K03496 PF13614.5 TIGR01969 SW01_contig04407 SW01 COG1475 K03497 PF02195.17 TIGR00180;TIGR03454;TIGR03734;TIGR04285 SW01_contig08769 SW01 COG0629 K03111 PF00436.24 TIGR00621 SW01_contig08769 SW01 COG0629 K03111 PF00436.24 TIGR00621 SW01_contig09770 SW01 COG0389 K03502 PF00817.19 NA SW01_contig09770 SW01 COG0389 K03502 PF00817.19 NA SW01_contig09770 SW01 COG1974 K03503 NA NA SW01_contig11526 SW01 COG0459 K04077 NA TIGR02348 SW02_F51BN4L01AKP2A SW02 COG0188 K02621 PF00521.19 TIGR01061;TIGR01062;TIGR01063 SW02_F51BN4L01AKP2A SW02 NA NA PF00521.19 TIGR01061;TIGR01062;TIGR01063 SW02_F51BN4L01BH2R8 SW02 NA K19465 NA NA SW02_F51BN4L01C538O SW02 COG0112 K00600 PF00464.18 NA SW02_F51BN4L01C7CO5 SW02 NA NA PF06568.10 NA SW02_F51BN4L01C88W1 SW02 COG2089 K01654 NA TIGR03569;TIGR03586 SW02_F51BN4L01CH9S4 SW02 NA NA PF08291.10 NA SW02_F51BN4L01CJRKG SW02 NA K17733 NA NA SW02_F51BN4L01DGDIE SW02 NA NA PF13884.5 NA SW02_F51BN4L01DKERX SW02 NA NA PF04545.15 NA SW02_F51BN4L01DQTPK SW02 NA NA PF08722.10 NA SW02_F51BN4L01DSXJG SW02 COG0756 K01520 PF00692.18 TIGR00576 SW02_F51BN4L01EUBTW SW02 NA NA NA TIGR02348 SW02_contig00280 SW02 NA NA PF16473.4 NA SW02_contig00651 SW02 COG0192 K00789 PF02772.15 TIGR01034 SW02_contig00651 SW02 COG0192 K00789 PF02773.15 TIGR01034 SW02_contig00651 SW02 COG2236 K07101 PF00156.26 NA SW02_contig00832 SW02 COG4675 NA PF07484.11 NA SW02_contig01747 SW02 COG1525 K01174 PF00565.16 NA SW02_contig02741 SW02 COG1442 NA NA NA SW02_contig03479 SW02 COG1738 K09125 PF02592.14 TIGR00697 SW02_contig03479 SW02 COG1087 K01784 PF01370.20;PF16363.4 TIGR01179;TIGR01181

151 SW02_contig03529 SW02 NA NA PF07230.10 NA SW02_contig03550 SW02 COG4502 NA PF06941.11 NA SW02_contig04517 SW02 COG1372 K00525 NA TIGR02505 SW02_contig05763 SW02 NA NA PF05065.12 TIGR01554 SW02_contig05975 SW02 COG1212 K00979 PF02348.18 TIGR00466 SW02_contig05975 SW02 COG2877 K01627 PF00793.19 TIGR01361;TIGR01362 SW02_contig05975 SW02 COG2877 K01627 PF00793.19 TIGR01362 SW02_contig06151 SW02 COG2192 K00612 PF02543.14 NA SW02_contig06151 SW02 COG2192 K00612 PF16861.4 NA SW02_contig06396 SW02 COG0542 K03694 PF10431.8 TIGR02639;TIGR03346 SW02_contig06396 SW02 COG0542 K03694 PF00004.28;PF07724.13;PF07728.13;PF00158.25 TIGR02639;TIGR03345;TIGR03346 SW02_contig06396 SW02 COG0242 K01462 PF01327.20 TIGR00079 SW02_contig07851 SW02 COG1080 K01006 NA TIGR01828 SW02_contig07851 SW02 NA NA NA TIGR01828 SW02_contig07851 SW02 COG1080 K01006 PF01326.18 TIGR01828 SW02_contig07851 SW02 COG1080 K01006 PF01326.18 TIGR01828 SW02_contig07998 SW02 COG0208 K00526 PF00268.20 TIGR04171 SW02_contig08061 SW02 COG0389 K03502 NA NA SW02_contig08061 SW02 COG0389 K03502 PF00817.19 NA SW02_contig08061 SW02 COG1974 K03503 PF00717.22 TIGR00498 SW02_contig08384 SW02 NA NA PF13884.5 NA SW02_contig08786 SW02 NA NA PF00940.18 NA SW02_contig08786 SW02 NA NA PF00940.18 NA SW02_contig08786 SW02 COG5108 NA PF00940.18 NA SW02_contig09585 SW02 COG2925 K01141 NA NA SW02_contig09585 SW02 NA K19465 NA NA SW02_contig09585 SW02 NA K19465 NA NA SW02_contig09960 SW02 NA NA PF13385.5 NA SW02_contig10294 SW02 COG3179 K03791 PF01471.17 TIGR02869 SW02_contig10868 SW02 COG0171 K01916 NA NA SW02_contig10868 SW02 COG0171 K01916 PF02540.16 TIGR00552 SW02_contig12242 SW02 NA NA PF00462.23 TIGR02180;TIGR02181;TIGR02183;TIGR02190;TIGR02196 SW02_contig12242 SW02 COG0208 K00526 NA NA SW02_contig13327 SW02 NA NA PF11351.7 NA SW02_contig13710 SW02 COG0071 K04080 NA NA SW02_contig16270 SW02 COG1003 K00281 PF02347.15 TIGR00461 SW02_contig16270 SW02 NA K00281 NA TIGR00461 SW02_contig16270 SW02 COG1003 K00281 NA TIGR00461 SW02_contig16270 SW02 COG1003 K00281 NA TIGR00461 SW02_contig16270 SW02 COG1003 K00281 NA TIGR00461 SW02_contig16270 SW02 COG1003 K00281 NA TIGR00461

152 SW02_contig22738 SW02 NA K02469 NA TIGR01063 SW02_contig22738 SW02 COG0188 K02469 PF00521.19 TIGR01061;TIGR01062;TIGR01063 SW02_contig22738 SW02 COG0188 K02469 PF00521.19 TIGR01061;TIGR01062;TIGR01063 SW02_contig22738 SW02 COG0188 K02469 NA TIGR01061;TIGR01062;TIGR01063 SW02_contig22738 SW02 COG0629 K03111 PF00436.24 TIGR00621 SW08_F7T5LKM01BTD2S SW08 NA K06214 PF03783.13 NA SW08_F7T5LKM02HSKKT SW08 COG0451 K02377 PF01370.20 NA ScoA_contig00057 ScoA NA NA PF17236.1 NA ScoA_contig00309 ScoA NA NA PF17236.1 NA ScoA_contig00309 ScoA COG3064 NA NA NA ScoA_contig00589 ScoA COG5565 NA PF03237.14 NA ScoA_contig01418 ScoA COG2369 NA NA TIGR01641 ScoA_contig01418 ScoA COG1783 NA NA NA ScoA_contig02215 ScoA COG1351 NA PF02511.14 TIGR02170 ScoA_contig02215 ScoA COG0208 K00526 PF00268.20 NA ScoA_contig02841 ScoA COG0464 NA PF00004.28 TIGR01242;TIGR01243 ScoA_contig02841 ScoA COG0464 NA NA NA ScoA_contig02841 ScoA COG1404 NA PF00082.21 NA ScoA_contig04573 ScoA COG5405 K01419 PF00227.25 TIGR03692 ScoA_contig04573 ScoA COG1220 K03667 PF00004.28;PF07724.13 TIGR00382;TIGR00390 ScoA_contig04573 ScoA COG0265 K04771 PF13365.5 TIGR02037;TIGR02038 ScoA_contig04573 ScoA COG0265 K04771 PF13180.5 TIGR02037;TIGR02038 ScoA_contig04573 ScoA COG0210 K03657 PF00580.20;PF13361.5;PF13538.5 TIGR00609;TIGR01073;TIGR01074;TIGR01075;TIGR02784;TIGR02 785 ScoA_contig06915 ScoA COG2521 K00571 PF01555.17 NA ScoA_contig06915 ScoA NA NA PF02945.14 NA ScoA_contig07755 ScoA COG0749 K02335 PF00476.19 NA ScoA_contig07755 ScoA COG0258 K02335 PF02739.15 TIGR00593 ScoA_contig07755 ScoA COG0358 K17680 NA NA ScoA_contig07755 ScoA NA K17680 NA NA ScoA_contig07755 ScoA COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoA_contig08813 ScoA COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoA_contig08813 ScoA COG0358 K17680 NA NA ScoA_contig08813 ScoA COG0358 K17680 PF13155.5 NA ScoA_contig08813 ScoA COG0749 K02335 PF02739.15 TIGR00593 ScoA_contig08813 ScoA COG0749 NA PF00476.19 NA ScoA_contig08916 ScoA COG3772 K01185 PF00959.18 NA ScoA_contig08916 ScoA NA NA PF11351.7 NA ScoA_contig09224 ScoA COG0358 K17680 PF13155.5 NA ScoA_contig09224 ScoA COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoA_contig09224 ScoA NA NA PF00940.18 NA

153 ScoA_contig09580 ScoA COG0749 K02335 PF00476.19 NA ScoA_contig09580 ScoA COG0258 K02335 PF02739.15 TIGR00593 ScoA_contig10158 ScoA COG0749 K02335 PF00476.19 NA ScoA_contig10158 ScoA COG0258 K02335 PF02739.15 TIGR00593 ScoA_contig10158 ScoA COG0358 K17680 PF13155.5 TIGR01391 ScoA_contig10158 ScoA NA NA PF00940.18 NA ScoA_contig11045 ScoA NA NA PF05838.11 NA ScoA_contig11045 ScoA NA NA PF11195.7 NA ScoB_contig00065 ScoB COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoB_contig00065 ScoB COG0358 K17680 PF13155.5 NA ScoB_contig00065 ScoB COG0749 K02335 PF02739.15 TIGR00593 ScoB_contig01350 ScoB NA NA PF17236.1 NA ScoB_contig02357 ScoB COG2131 K01493 PF00383.22 NA ScoB_contig02357 ScoB NA K17680 PF13155.5;PF13362.5;PF13662.5 NA ScoB_contig02357 ScoB COG0258 K02335 PF02739.15 TIGR00593 ScoB_contig02357 ScoB COG0749 K02335 PF00476.19 NA ScoB_contig04566 ScoB COG0791 NA NA TIGR02219 ScoB_contig04566 ScoB COG4695 NA PF04860.11 TIGR01537 ScoB_contig04566 ScoB COG5323 NA PF03237.14;PF17289.1 NA ScoB_contig04566 ScoB COG2983 K09160 NA NA ScoB_contig04566 ScoB COG5449 NA PF09931.8;PF09356.9 TIGR02218 ScoB_contig04566 ScoB COG5448 NA PF09343.9 TIGR02217 ScoB_contig04566 ScoB NA NA PF05521.10 NA ScoB_contig04566 ScoB NA NA NA TIGR02215 ScoB_contig04566 ScoB COG4653 NA PF05065.12 TIGR01554 ScoB_contig04566 ScoB COG3740 K06904 PF04586.16 TIGR01543 ScoB_contig04628 ScoB COG2189 K07316 PF01555.17 NA ScoB_contig04628 ScoB COG3587 K01156 PF04851.14 NA ScoB_contig05778 ScoB COG3772 K01185 PF00959.18 NA ScoB_contig08270 ScoB NA NA PF01391.17 NA ScoB_contig08270 ScoB NA NA PF17236.1 NA ScoB_contig10406 ScoB NA NA PF08291.10 NA ScoB_contig11792 ScoB COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoB_contig11792 ScoB NA K17680 NA NA ScoB_contig11792 ScoB COG0358 K17680 NA NA ScoB_contig11792 ScoB COG0258 K02335 PF02739.15 TIGR00593 ScoB_contig11792 ScoB COG0749 K02335 PF00476.19 NA ScoB_contig11915 ScoB COG0358 K17680 NA NA ScoB_contig11915 ScoB COG0358 K17680 PF13155.5;PF13662.5 TIGR01391 ScoB_contig13026 ScoB NA NA PF04404.11 NA ScoB_contig14796 ScoB COG0358 K17680 PF13155.5 TIGR01391

154 ScoB_contig14796 ScoB COG0258 K02335 PF02739.15 TIGR00593 ScoB_contig14796 ScoB COG0749 K02335 PF00476.19 NA ScoB_contig15285 ScoB NA NA PF02925.15 NA ScoB_contig15285 ScoB NA NA PF02305.16 NA ScoB_contig16413 ScoB NA NA PF01391.17 NA ScoB_contig16413 ScoB NA NA PF17236.1 NA ScoB_contig18743 ScoB COG0358 K17680 NA NA ScoB_contig18743 ScoB COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoB_contig18743 ScoB NA NA PF00940.18 NA ScoB_contig18837 ScoB COG0451 K02377 PF01370.20;PF16363.4 TIGR01179;TIGR01181;TIGR04180 ScoB_contig18837 ScoB COG1089 K01711 PF01370.20;PF16363.4 TIGR01179;TIGR01181;TIGR01472;TIGR02622;TIGR04180 ScoB_contig18837 ScoB COG1089 K01711 PF01370.20;PF16363.4 TIGR01472;TIGR02622 ScoC_contig00741 ScoC NA K06909 NA NA ScoC_contig00741 ScoC COG4373 NA NA NA ScoC_contig01271 ScoC NA NA PF03420.12 NA ScoC_contig04323 ScoC NA NA PF07068.10 NA ScoC_contig04646 ScoC COG0592 K02338 PF02768.14 TIGR00663 ScoC_contig04646 ScoC NA NA PF00712.18 TIGR00663 ScoC_contig04740 ScoC COG4626 NA PF03354.14 NA ScoC_contig04740 ScoC NA NA PF02675.14 TIGR03330 ScoC_contig04740 ScoC COG1403 K07451 PF01844.22 NA ScoC_contig07180 ScoC COG0358 K17680 PF13155.5 NA ScoC_contig07180 ScoC COG0749 K02335 PF02739.15 TIGR00593 ScoC_contig12503 ScoC COG0372 K01659 PF00285.20 TIGR01793;TIGR01798;TIGR01800 ScoC_contig12503 ScoC COG2513 K03417 PF00463.20;PF13714.5 TIGR01346;TIGR02317;TIGR02319;TIGR02320;TIGR02321 ScoC_contig12503 ScoC COG0542 K03694 PF00004.28;PF02861.19 TIGR02639;TIGR03345;TIGR03346 ScoC_contig12503 ScoC COG0542 K03694 PF00004.28;PF07724.13;PF07728.13;PF10431.8 TIGR02639;TIGR03345;TIGR03346 ScoC_contig12503 ScoC COG0451 K01709 PF01370.20;PF16363.4;PF02719.14 TIGR01179;TIGR01181;TIGR01472;TIGR02622;TIGR04180 ScoC_contig12503 ScoC COG0399 K12452 PF01041.16 TIGR02379;TIGR03588;TIGR04181;TIGR04427 ScoC_contig12503 ScoC COG0399 K12452 PF01041.16 NA ScoC_contig12503 ScoC COG1898 K01790 PF00908.16 TIGR01221 ScoC_contig12762 ScoC COG0208 K00526 NA NA ScoC_contig12762 ScoC COG0208 K00526 PF00268.20 TIGR04171 ScoC_contig12762 ScoC COG0209 K00525 PF02867.14 TIGR02504;TIGR02506;TIGR02510;TIGR04170 ScoC_contig12762 ScoC COG0209 K00525 PF00317.20 TIGR02506;TIGR02510 ScoC_contig13426 ScoC NA NA NA TIGR02348 ScoC_contig13426 ScoC COG0459 K04077 PF00118.23 TIGR02345;TIGR02348 ScoC_contig13519 ScoC COG0749 K02335 PF01367.19;PF02739.15 TIGR00593 ScoC_contig13519 ScoC COG0358 K17680 PF13155.5 NA ScoC_contig13519 ScoC COG2131 K01493 PF00383.22;PF14437.5 TIGR02571 ScoC_contig13582 ScoC COG1004 K00012 PF00984.18 TIGR03026

155 ScoC_contig16980 ScoC NA NA PF11056.7 NA ScoC_contig20525 ScoC COG0358 K17680 PF13155.5;PF13662.5 TIGR01391 ScoC_contig21402 ScoC NA NA PF13640.5 NA ScoC_contig24315 ScoC NA NA PF01391.17 NA ScoC_contig24315 ScoC NA NA PF17236.1 NA StyA_contig00172 StyA COG4653 NA PF05065.12 TIGR01554 StyA_contig00172 StyA NA NA PF04586.16 TIGR01543 StyA_contig00172 StyA NA NA PF04860.11 TIGR01537 StyA_contig05421 StyA NA NA PF04965.13 NA StyA_contig09107 StyA NA NA NA TIGR02656 StyA_contig09107 StyA COG0176 K00616 PF00923.18 TIGR00874;TIGR00875;TIGR02134 StyA_contig09717 StyA NA K01955 NA TIGR01689 StyA_contig09717 StyA NA NA PF00294.23 TIGR02152;TIGR02198 StyA_contig09717 StyA COG2870 NA NA TIGR02198 StyA_contig09717 StyA COG2870 K03272 PF01467.25 TIGR00125;TIGR01518;TIGR02199 StyA_contig09909 StyA COG4695 NA PF04860.11 TIGR01537 StyA_contig09909 StyA NA NA PF04586.16 TIGR01543 StyA_contig09909 StyA NA NA NA TIGR01554 StyA_contig09909 StyA NA NA PF05065.12 TIGR01554 StyA_contig09909 StyA NA NA PF05135.12 TIGR02215 StyA_contig11879 StyA NA NA PF13476.5 TIGR00618 StyA_contig11879 StyA NA NA PF11360.7 NA StyA_contig12383 StyA COG0171 K01916 PF02540.16 TIGR00552 StyA_contig12383 StyA NA K01955 NA NA StyA_contig12383 StyA NA K03272 PF00294.23 TIGR02152;TIGR02198 StyA_contig12383 StyA NA NA NA TIGR02198 StyA_contig12383 StyA COG2870 K03272 PF01467.25 TIGR00125;TIGR01518;TIGR02199 StyA_contig12564 StyA NA K21313 PF00940.18 NA StyA_contig12564 StyA COG0358 K17680 NA NA StyA_contig13411 StyA COG0176 K00616 PF00923.18 TIGR00874;TIGR00875;TIGR02134 StyA_contig13411 StyA NA K02706 PF00124.18 TIGR01151;TIGR01152 StyA_contig14203 StyA NA NA PF08994.9 NA StyA_contig14203 StyA NA NA PF08855.9 NA StyA_contig15606 StyA NA NA PF07230.10 NA StyA_contig16084 StyA COG0171 K01916 PF02540.16 TIGR00552 StyA_contig16084 StyA NA NA PF03721.13 TIGR03026 StyA_contig16084 StyA COG1004 K00012 PF00984.18 TIGR03026 StyA_contig16260 StyA NA NA PF04577.13 NA StyA_contig16307 StyA NA K01955 NA NA StyA_contig16307 StyA COG3347 NA NA NA StyA_contig17511 StyA NA NA NA TIGR03182

156 StyA_contig17511 StyA COG0022 K21417 PF02779.23 NA StyA_contig17522 StyA NA K00012 PF03721.13 TIGR03026 StyA_contig17522 StyA COG1004 K00012 PF00984.18 TIGR03026 StyB_contig07188 StyB COG0208 K00526 PF00268.20 NA StyB_contig07188 StyB COG0208 K00526 NA NA TedA_contig00635 TedA COG0553 NA PF04851.14;PF00176.22 NA TedA_contig00635 TedA COG5545 NA PF08800.9 NA TedA_contig05443 TedA NA NA PF12684.6 NA TedA_contig05443 TedA NA NA PF06378.10 NA TedA_contig05443 TedA COG1475 NA NA NA TedA_contig05650 TedA COG3108 NA PF08291.10 NA TedA_contig05650 TedA NA NA PF03382.13 NA TedA_contig05817 TedA COG0863 K00571 PF01555.17 NA TedB_contig00086 TedB COG1475 NA NA NA TedB_contig00086 TedB NA NA PF06378.10 NA TedB_contig00086 TedB NA NA PF12684.6 NA TedB_contig00195 TedB NA NA PF03382.13 NA TedB_contig00195 TedB COG3108 NA PF08291.10 NA TedB_contig04928 TedB COG0553 NA NA NA TedB_contig04928 TedB COG5545 NA PF08800.9 NA TedB_contig05488 TedB COG0749 K02335 PF00476.19 TIGR00593 TedB_contig06483 TedB COG0459 K04077 PF00118.23 TIGR02339;TIGR02340;TIGR02341;TIGR02342;TIGR02343;TIGR02 345;TIGR02348

157

6.3 Chapter Four

To check if the viral extraction methods resulted in the extraction of viruses, transmission electron microscopy (TEM) of sponge viral extracts were conducted and check for the presence of viral-like particles (Appendix Figure S4.1). Figure S4.1 A and E resemble icosahedron enveloped viruses with a width of 40-50 nm (similar in structure to myoviruses (Modrow et al., 2013)). Figure S4.1 B and C are non-tailed virus with an icosahedron capsid head size of approximately 120 nm and 40 nm, respectively. Figure 4.1 D has a helix structure, resembling an RNA virus (Modrow et al., 2013) of approximately 125 nm in length and Figure S4.1 F appear to be an enveloped virus with a width of 60-70 nm.

Figure S4.1 TEM images of viral extracts containing potential viral-like particles of different morphologies.

158 Table S4.1 Genes associated with defense mechanism in sponge-associated bacteria genomes used to isolate viruses.

Bacterial culture Phage genes CRISPR Restriction sequences enzymes

Ruegeria sp. Phage uncharacterized protein (putative large terminase), C- 0 2 AU67 terminal domain-containing protein

Pseudovibrio sp. Putative phage tail protein 1 3 AU243 Putative phage cell wall peptidase, NlpC/P60 family

Phage conserved hypothetical protein BR0599

Phage prohead protease, HK97 family/phage major capsid protein, HK97 family, TIGR01554

Phage portal protein, lambda family

Phage terminase, large subunit GpA

Phage DNA packaging protein, Nu1 subunit of terminase

Phage integrase family protein

Leptolyngbya Phage integrase family protein 5 3 spCCY

Tenacibaculum Conserved hypothetical phage tail region protein 2 6 sp. AU330 Phage baseplate assembly protein W

159 Table S4.2 Summary of COG, KEGG, PFAM and TIGRFAM protein classifications of Ruegeria sp. 67 phage ORFs. The gene highlighted in bold indicate sequence homology to proteins from the host Ruegeria sp. AU67.

Locus Tag Gene Product Name Functional annotation % query % E-value Accession number Organism Protein predicted via IMG via IMG coverage identity function match Ga0138520_111 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_112 Hypothetical protein 98 34.72 6.00E-20 gi|1172574560|ref|WP_0 Pseudaminobacter manganicus Hypothetical protein 80920472.1| (Alphproteobacteria) Ga0138520_113 Hypothetical protein 100 44.53 4.00E-28 gi|640277052|ref|WP_02 Aminobacter sp. J41 Hypothetical protein 4847820.1| (Alphproteobacteria) Ga0138520_114 Hypothetical protein 23 59.35 1.00E-32 gi|1168015087|gb|ARB05 Synechococcus virus S-ESS1 Hypothetical protein 742.1| (Siphoviridae) Ga0138520_115 Hypothetical protein 19 66.67 8.00E-05 gi|1184849988|gb|ARK07 Sphingobium phage Lacusarx Hypothetical protein 527.1| Ga0138520_116 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_117 Hypothetical protein 76 44.34 9.00E-83 gi|1119678217|ref|WP_0 Sphingomonas sp. JJ-A5 Hypothetical protein 72597331.1| (Alphproteobacteria) Ga0138520_118 Hypothetical protein 98 54.45 7.00E-145 gi|1168015072|gb|ARB05 Synechococcus virus S-ESS1 DUF932 domain- 727.1| (Siphoviridae) containing protein Ga0138520_119 Hypothetical protein N/A

Ga0138520_1110 Hypothetical protein 100 46.59 2.00E-16 gi|1338671248|gb|AUX83 Microbacterium phage Raccoon Hypothetical protein 266.1| (Siphoviridae) Ga0138520_1111 Hypothetical protein 98 49.67 2.00E-99 gi|1039941200|ref|WP_0 Mesorhizobium sp. AA22 Hypothetical protein 65011803.1| (Alphproteobacteria) Ga0138520_1112 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_1113 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_1114 Hypothetical protein N/A N/A N/A N/A N/A N/A

160 Table S4.2 (continued). Ga0138520_1115 Ribonucleoside- K00525, Purine and 98 57.26 0 gi|705244755|gb|AIW567 uncultured virus Ribonucleotide diphosphate reductase Pyrimidine 21.1| reductase alpha chain metabolism

Ga0138520_1116 Hypothetical protein 94 42.16 1.00E-39 gi|472341437|ref|YP_007 Loktanella phage pCB2051-A Hypothetical protein 674955.1| (Unclassified dsDNA phage)

Ga0138520_1117 Chromosome COG1475, pfam02195- 83 41.63 2.00E-51 gi|435844368|ref|YP_007 Agrobacterium phage 7-7-1 Hypothetical protein segregation protein ParBc 006484.1| (Myoviridae) Spo0J, contains ParB- like nuclease domain

Ga0138520_1118 Hypothetical protein 85 50 7.00E-23 gi|1176003540|gb|OQX10 Thiothrix lacustris Hypothetical protein 524.1| (gammaproteobacterial)

Ga0138520_1119 Hypothetical protein 54 32.94 6.00E-05 gi|1113389848|gb|OJW52 Sphingobacteriales bacterium 50- Hypothetical protein 869.1| 39

Ga0138520_1120 Uncharacterized COG3750, pfam10073- 89 59.3 4.00E-25 gi|1056825338|ref|WP_0 Rhodobacteraceae bacterium DUF2312 domain- conserved protein, DUF2312 68244832.1| O3.65 containing protein UPF0335 family

Ga0138520_1121 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_1122 Hypothetical protein N/A N/A N/A N/A N/A N/A

Ga0138520_1123 Hypothetical protein 87 38.12 2.00E-27 gi|499857403|ref|WP_01 Ruegeria sp. TM1040 Hypothetical protein 1538137.1|

Ga0138520_1124 Thymidylate synthase COG1352, 60 86 3.00E-17 gi|6511288|ref|653418| Ruegeria sp. AU67 Thymidylate (FAD) pfam02511-ThyX, synthase (FAD) K03465 (One carbon pool by folate, Pyrimidine metabolism)

161 Table S4.2 (continued). Ga0138520_1125 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1126 Hypothetical protein 78 54.1 9.00E-07 gi|746112960|ref|WP_03 Leisingera sp. ANG-M7 Hypothetical protein 9178784.1| Ga0138520_1127 Hypothetical protein 97 66.67 2.00E-66 gi|1035522406|gb|ANJ20 Roseobacter phage RD-1410W1-01 Thymidylate 765.1| (Podoviridae) synthase

Ga0138520_1128 Hypothetical protein 99 37.04 4.00E-15 gi|472341425|ref|YP_007 Loktanella phage pCB2051-A Hypothetical protein 674943.1| (Unclassified dsDNA phage)

Ga0138520_1129 Putative peptidoglycan Pfam01471 75 61 5.00E-117 gi|1275800693|ref|WP_0 Pararhizobium sp. XC0140 DUF3380 domain-

binding domain- 99868241.1| containing protein containing protein Ga0138520_1130 Subtilase family protein Pfam00082-Peptidase 34 40.85 2.00E-82 gi|1035522415|gb|ANJ20 Roseobacter phage RD-1410W1-01 Hypothetical protein S8 774.1| (Podoviridae)

Ga0138520_1131 Hypothetical protein 90 38.79 4.00E-10 gi|1247476833|gb|PCJ336 Alphaproteobacteria bacterium Hypothetical protein 16.1| Ga0138520_1132 Protein of unknown Pfam10983-DUF2793 12 49.19 6.00E-27 gi|1357959230|ref|WP_1 Rhodobacteraceae bacterium DUF2793 domain- function (DUF2793) 05321483.1| WD3A24 containing protein Ga0138520_1133 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1134 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1135 Putative phage tail Pfam13550-Phage 90 32.21 6.00E-103 gi|1114543965|gb|APL99 Bordetella phage FP1 (Siphoviridae) Virion structural protein tail_3 377.1| protein

Ga0138520_1136 Hypothetical protein 88 34.25 2.00E-06 gi|738010744|ref|WP_03 Bradyrhizobium sp. WSM3983 hypothetical protein 5972126.1|

162

Table S4.2 (continued). Ga0138520_1137 Hypothetical protein 82 52.38 3.00E-13 gi|712912847|ref|YP_009 Silicibacter phage DSS3-P1; Hypothetical protein 099702.1| Ruegeria phage DSS3-P1 (Siphoviridae)

Ga0138520_1138 Phage conserved Pfam09356-Phage 99 32.19 1.00E-49 gi|712912848|ref|YP_009 Silicibacter phage DSS3-P1; Hypothetical protein hypothetical protein BR0599 099703.1| Ruegeria phage DSS3-P1 BR0599 (Siphoviridae)

Ga0138520_1139 Von Willebrand factor Pfam13519-VWA_2 59 25.56 1.00E-25 gi|712912849|ref|YP_009 Silicibacter phage DSS3-P1; Hypothetical protein type A domain- 099704.1| Ruegeria phage DSS3-P1 containing protein (Siphoviridae)

Ga0138520_1140 Tape measure domain- TIGR02675-tape 100 29.28 2.00E-145 gi|552541458|ref|WP_02 Labrenzia sp. C1B10; Labrenzia sp. MULTISPECIES: tape containing protein measure domain 2999923.1| C1B70 measure domain- containing protein Ga0138520_1141 Hypothetical protein 82 29.55 3.00E-05 gi|559198868|gb|AHB120 Phage Sano (Siphoviridae) pre-tape measure 75.1| frameshift protein G Ga0138520_1142 Hypothetical protein 88 46.33 4.00E-71 gi|472341409|ref|YP_007 Loktanella phage pCB2051-A Hypothetical protein 674927.1| (Unclassified dsDNA phage)

Ga0138520_1143 Hypothetical protein 90 46.15 2.00E-34 gi|712912854|ref|YP_009 Ruegeria phage DSS3-P1 Hypothetical protein 099709.1| (Siphoviridae)

Ga0138520_1144 Hypothetical protein 98 50.75 3.00E-55 gi|1168015059|gb|ARB05 Synechococcus virus S-ESS1 Hypothetical protein 714.1| (Siphoviridae)

163 Table S4.2 (continued). Ga0138520_1145 Hypothetical protein 92 42.74 2.00E-17 gi|712912856|ref|YP_009 Silicibacter phage DSS3-P1; Hypothetical protein 099711.1| Ruegeria phage DSS3-P1 (Siphoviridae)

Ga0138520_1146 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1147 Phage major capsid Pfam03864 97 44.32 4.00E-80 gi|712912858|ref|YP_009 Ruegeria phage DSS3-P1 Major capsid protein protein E 099713.1| (Siphoviridae)

Ga0138520_1148 Bacteriophage lambda Pfam02924 76 44.23 1.00E-15 gi|712912859|ref|YP_009 Silicibacter phage DSS3-P1; Head decoration head decoration 099714.1| Ruegeria phage DSS3-P1 protein protein D (Siphoviridae)

Ga0138520_1149 Protein C Serine COG0616-Perplasmic 78 45.56 1.00E-71 gi|1168015058|gb|ARB05 Synechococcus virus S-ESS1 Peptidase S49 peptidase. MEROPS serine protease, ClpP 713.1| (Siphoviridae) family S49 class, pfam01343 Ga0138520_1150 Phage portal protein, TIGR01539, 93 55.97 0 gi|712912861|ref|YP_009 Ruegeria phage DSS3-P1 Portal protein lambda family pfam05136 099716.1| (Siphoviridae)

Ga0138520_1151 GpW protein Pfam02831 96 48.57 9.00E-14 gi|712912862|ref|YP_009 Silicibacter phage DSS3-P1; Head-to-tail joining 099717.1| Ruegeria phage DSS3-P1 protein (Siphoviridae)

Ga0138520_1152 Phage terminase, large COG5525-Phage 99 52.73 1863 gi|712912863|ref|YP_009 Silicibacter phage DSS3-P1; Terminase large subunit GpA terminase, large 099718.1| Ruegeria phage DSS3-P1 subunit subunit GpA, (Siphoviridae) pfam05876

164 Table S4.2 (continued). Ga0138520_1153 Protein of unknown Pfam07278 97 37.5 3.00E-36 gi|712912864|ref|YP_009 Silicibacter phage DSS3-P1; Putative terminase function (DUF1441) 099719.1| Ruegeria phage DSS3-P1 small subunit (Siphoviridae) Ga0138520_1154 SNF2 family N-terminal Pfam00176 93 42.28 1.00E-110 gi|712912868|ref|YP_009 Ruegeria phage DSS3-P1 Superfamily II domain-containing 099723.1| (Siphoviridae) DNA/RNA helicase protein Ga0138520_1155 Hypothetical protein 87 41.11 2.00E-11 gi|472341396|ref|YP_007 Loktanella phage pCB2051-A Hypothetical protein 674914.1| (Unclassified dsDNA phage) Ga0138520_1156 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1157 DNA polymerase K02334 99 48 0 gi|356927494|gb|AET422 Silicibacter phage DSS3-P1 DNA polymerase I 85.1| (Unclassified dsDNA) Ga0138520_1158 Protein of unknown Pfam10991 87 37.19 8.00E-26 gi|491346218|ref|WP_00 Acinetobacter; Acinetobacter sp. MULTISPECIES: function (DUF2815) 5204155.1| CIP 70.18 DUF2815 domain- containing protein Ga0138520_1159 Protein of unknown Pfam10926 88 30.99 4.00E-12 gi|712912879|ref|YP_009 Ruegeria phage DSS3-P1 Hypothetical protein function (DUF2800) 099734.1| (Siphoviridae) Ga0138520_1160 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1161 Hypothetical protein N/A N/A N/A N/A N/A N/A Ga0138520_1162 Hypothetical protein 82 41.43 8.00E-08 gi|1168015076|gb|ARB05 Synechococcus virus S-ESS1 N/A 731.1| (Siphoviridae) Ga0138520_1163 Virulence-associated COG5545, pfam05272- 96 30 7E-109 YP_009216663.1 Clostridium phage phiCT453A Virulence-associated protein E (predicted P- VirE E domain-containing loop ATPase and protein inactivated derivatives) Ga0138520_1164 Hypothetical protein N/A N/A N/A N/A N/A

165

166