Amphimedon Queenslandica

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Analysis of silicatein gene expression and spicule formation in the demosponge Amphimedon queenslandica

Aude Gauthier BMarSt (Hons)

A thesis submitted for the degree of Master of Philosophy at The University of Queensland in 2014 School of Biological Sciences

Abstract

The skeletal elements in most sponges are siliceous spicules. These are fabricated into species-specific sizes and shapes. Demosponges, in particular, have specialised cells called sclerocytes that possess the unique ability to synthesise biosilica and these spicules. Underlying the diversity of demosponge spicules morphology is a conserved protein, called silicatein. This thesis aims to investigate the process of spiculogenesis in the different developmental stages of the demosponge Amphimedon queenslandica, and the evolution and developmental expression of the silicatein gene family in relation to spicule formation.

A. queenslandica is the only sponge species to have its genome fully sequenced, assembled and annotated, and currently is one of the best models to study sponge development (Srivastava et al. 2010). This species broods embryos year-round, facilitating the access to embryological and larval material (Leys and Degnan 2001). This combination of logistical advantages means that I was able to trace the expression of silicatein genes through A. queenslandica embryonic, larval and postlarval development. Spicule formation starts in the early embryogenesis in A. queenslandica during, gastrulation or the brown stage. Spicule number increases throughout embryonic development until the pre-hatching larval stage, with the emerging larvae having about 1000 spicules. No detectable increase in spicule number was recorded during larval and postlarval development. Spicule number varied remarkably between different individual embryos and larvae of the same stage of development.

I initially identified six silicatein β like genes in the genome of A. queenslandica, among which four can be categorised as non-conventional by the absence of the serine in the catalytic triad of the protein. These genes do not have direct orthologues in other sponge species and appear to have evolved by a lineage-specific gene duplication. A comprehensive phylogenetic analysis of this gene family in sponge indicated that silicatein α arose from silicatein β by gene duplication and that silicatein β gene share traits with both cathepsin L and silicatein. Conservation of gene structure and exon length in silicatein and cathepsin L genes suggests that these genes have preserved an ancestral gene structure common to both gene families in both marine and freshwater sponges.

Using in situ hybridisation, I demonstrated that silicatein genes are expressed during A. queenslandica early embryonic development, with genes being expressed exclusively in sclerocytes. Analysis of gene expression levels through embryogenesis and metamorphosis, using RNA-Seq performed on a pool of same stage individuals, revealed that all silicatein-like genes are differentially expressed throughout development, and the expression of silicatein genes occurs prior to spicule formation. However, some silicatein-like gene expression levels and spicule number do not appear to be tightly correlated.

Declaration by author

This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my research higher degree candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the policy and procedures of The University of Queensland, the thesis be made available for research and study in accordance with the Copyright Act 1968 unless a period of embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis.

Publication during candidature

No publications.

Publication included in this thesis

No publications included.

Contributions by others to the thesis

My supervisor, Professor Bernard Degnan, contributed to the conception and design of the research and critically revised and proofread all sections of this thesis. My co-supervisor, Dr Nagayasu Nakanishi performed technical work required for the RNA-Seq data.

Statement of parts of the thesis submitted to quality for the award of another degree

None.

Acknowledgements

I am extremely grateful to my supervisor Bernie Degnan for accepting me into his group and introducing me to the world of sponges and research, and for his continuous support and invaluable advice through this project. Additionally, I would like to thank my committee members, Dr Nagayasu Nakanishi and Dr David Merritt, for their encouragement and interest in my work.

I would also like to thank everyone in the Degnan lab, both past and present members, for their help, support, coffee breaks, stimulating discussions, and fun times in and out of the office throughout the years. This includes in no particular order, Carmel McDougall, Andrew Calcino, Laura Grice, Kerry Roper, Maely Gauthier, Jo Bayes, Jaret Bilewitch, Nobuo Ueda, Felipe Aguilera, Jabin Watson, Simone Higgie, Federico Gaiti, Katia Jindrich, Kevin Kocot, Sunsuke Sogabe, Selene Fernandez Valverde, Ben Yuen, Tahsha Say, Rebecca Fieth and William Hatleberg. I am most particularly thankful to Carmel McDougall and Kerry Roper for their invaluable help and guidance in the laboratory. A special thanks to Sandie Degnan for her support and encouragement through the different milestones of this project, Nagayasu Nakanishi for providing help and suggestions with my in situ hybridization approach, and Katia Jindrich and Ben Yuen for going through some troubleshooting with me in the lab. I also wish to thank Maely Gauthier, Kevin Kocot, Laura Grice, Simone Higgie and William Hatleberg for taking the time to proofread parts of my thesis.

During the course of this project, I was lucky to participate in field trips to Heron Island. So, I would like to acknowledge staff members of Heron Island Research Station, which are always available and understanding of our last minute experimental set-up, for their technical help and great working environment atmosphere. Thanks to the sponge people who made those trips worth remembering for their assistance in the field, the great times and sunset drinks, as well as for going through those endless night with me fixing material and dissecting brood chambers.

I would like to thank friends and family who supported me during my time here. Above all, my parents for supporting my choice and giving me the opportunity to undertake my studies in Australia. A special thanks ‘au Nain’ (Arnault Gauthier) for listening to my complaints when things were not working to plan and checking for the numerous grammatical mistakes.

Keywords

Porifera, silicatein, silicatein gene expression, spicules

Australian and New Zealand Standard Research Classifications (ANZSRC)

ANZSRC code: 060309 Phylogeny and comparative analysis 50% ANZSRC code: 060808 Invertebrate biology 50%

Fields of Research (FoR) Classification

FoR code: 0603 Evolutionary biology 50% FoR code: 0608 Zoology 50%

Table of Contents

Chapter 1: General Introduction...... 1 1.1 Sponge biosilica structure…………………………………………………………………………………………….2 1.1.1 Spicule morphology………………………………………………………………………………………………3 1.1.2 Spicule formation…………………………………………………………………………………………………4 1.2 Silicatein……………………………………………………………………………………………………………………….7 1.2.1 Catalytic mechanism of silicatein………………………………………………………………………….7 1.3 Amphimedon queenslandica as a study model………………………………………………………………8 1.4 Aims of this study………………………………………………………………………………………………………….9

Chapter 2: Identification of silicatein genes in the demosponge Amphimedon queenslandica and their evolutionary relationship to other sponge silicatein and cathepsin genes………………………………………………..11 2.1 Abstract……………………………………………………………………………………………………………………..11 2.2 Introduction……………………………………………………………………………………………………………….12 2.3 Materials and Methods………………………………………………………………………………………………14 2.3.1 Identification of silicatein genes in Amphimedon queenslandica………………………..14 2.3.2 Gene architecture analyses………………………………………………………………………………..15 2.3.3 Molecular phylogenetic analyses………………………………………………………………………..15 2.4 Results………………………………………………………………………………………………………………………. 16 2.4.1 Identification and genomic organisation of A. queenslandica silicatein genes……..16 2.4.2 Conservation in the genomic structure of A. queenslandica silicatein genes………..20 2.4.3 Phylogenetic relationship of sponge silicateins……………………………………………………23 2.4.4 More detailed analysis of silicatein protein sequences………………………………………..27 2.4.5 Presence of the CY motif in other metazoan cathepsin L sequences……………………28 2.5 Discussion…………………………………………………………………………………………………………………..33

Chapter 3: Analysis of spicule formation and silicatein gene expression in the demosponge Amphimedon queenslandica……………………………………………38 3.1 Abstract……………………………………………………………………………………………………………………..38 3.2 Introduction……………………………………………………………………………………………………………….38 3.3 Materials and Methods………………………………………………………………………………………………41 3.3.1 Sponge collection and fixation of biological materials…………………………………………41 3.3.1.1 Brooded embryos………………………………………………………………………………………41 3.3.1.2 Larval and postlarval stages……………………………………………………………………….42 3.3.2 Spicule preparation…………………………………………………………………………………………….43 3.3.3 Gene expression analysis – RNA-Seq protocol…………………………………………………….43 3.3.3.1 RNA extraction and cDNA synthesis…………………………………………………………..44 3.3.3.2 Samples processing………………………………………………………………………………….. 44 3.3.4 Gene expression analysis – qRT-PCR…………………………………………………………………..44 3.3.4.1 Primer design for genes of interest…………………………………………………………….45 3.3.4.4 qRT-PCR analyses………………………………………………………………………………………46 3.3.5 Whole-mount in situ hybridisation……………………………………………………………………..47 3.3.5.1 Probe synthesis…………………………………………………………………………………………48 3.3.6 Statistical analyses……………………………………………………………………………………………..48 3.4 Results………………………………………………………………………………………………………………………. 49 3.4.1 Spicule number increases during embryogenesis but not in larvae and early postlarvae………………………………………………………………………………………………….49 3.4.2 Temporal expression of A. queenslandica silicatein genes during development…50 3.4.3 Expression pattern of A. queenslandica silicatein genes, Aqu2.41046, Aqu2.41047 and Aqu2.42494……………………………………………………………………………..54 3.4.4 Localised expression of Aqu2.42494 during development…………………………………..54 3.4.5 Individual variation in gene expression and spicule number……………………………….57 3.5 Discussion…………………………………………………………………………………………………………………..60

Chapter 4: General Discussion……………………………………………………………………. 64 4.1 Does silicatein expression correlate with spicule number? …………………………………………65 4.2 Why does variation in silicatein expression and spicule number occur?...... 66 4.3 Conclusions and future directions……………………………………………………………………………….67

References………………………………………………………………………………………………….69

Appendices…………………………………………………………………………………………………81 …A1. GeneBank (NCBI) accession numbers of protein sequences of demosponges and hexactinellid (asterix) used in this study……………………………………………………………….81

…A2. Protein sequences alignment of the conserved domain, Inhibitor I29 and Peptidase C1A, of Amphimedon queenslandica silicatein and cathepsin L genes…………………………83

…A3. Protein sequences alignment of the conserved domain, Inhibitor I29 and Peptidase C1A, of silicatein-like sequences in different animals ………………………………………………..84

List of Figures

Figure 1.1 Evolutionary tree showing poriferan relationship to each other and to other metazoans……………………………………………………………………………………………………………3

Figure 1.2 Schematic of spicule formation……………………………………………………………………………..5

Figure 2.1 Detailed comparison of conserved amino acids of demosponge silicatein…………..14

Figure 2.2 Comparison of the protein sequence of Amphimedon queenslandica (Aqu2.) silicatein genes with homologous sequence from the sponge S. domuncula …….18

Figure 2.3 Phylogenetic relationships and gene structure of Amphimedon queenslandica silicatein and cathepsin L genes………………………………………………………………………….19

Figure 2.4 Comparison of exon structure of silicatein and cathepsin L genes of A. queenslandica with S. domuncula and L. baicalensis…………………………………………..22

Figure 2.5 Phylogenetic relationships of sponge silicatein and cathepsin L genes family……..26

Figure 2.6 Sequence motifs of the two main regions defining silicatein genes…………………….28

Figure 2.8 Predicted emergence of silicatein like genes in metazoan derived from sequences encoding the characteristic SY/CY (Serine-Tyrosine/Cysteine-Tyr)…….31

Figure 3.1 Embryonic development of Amphimedon queenslandica……………………………………42

Figure 3.2 Rate of spicule accumulation during Amphimedon queenslandica embryonic, larval and postlarval development……………………………………………………………………..50

Figure 3.3 Normalised transcript levels (transcripts per million) of Amphimedon queenslandica silicatein genes throughout development. …………………………………..52

Figure 3.4 Comparison of mean spicule number with mean silicatein gene expression per stages…………………………………………………………………………………………………………..53

Figure 3.5 Silicatein expression pattern of type I and II genes in juvenile Amphimedon queenslandica using whole mount in situ hybridisation………………..55

Figure 3.6 Expression of Aqu2.42494 during A. queenslandica development………………………56

Figure 3.7 Individual variations in spicule number and silicatein gene expression throughout A. queenslandica development…………………………………………………….....59

List of Tables

Table 2.1 Intron phases for A. queenslandica silicatein and cathepsin L genes, and S. domuncula and L. baicalensis orthologues…………………………………………………………..23

Table 2.2 EnsemblMetazoa accession numbers of silicatein like sequences in different animals……………………………………………………………………………………………………………….32

Table 3.1 Gene specific primer sequences used in this study for quantitative real tine PCR analysis……………………………………………………………………………………………………………….45

Table 3.2 Total RNA (ng/µl) used for each sample stages to prepare cDNA…………………………46

Table 3.3 List of references genes used in this study for each developmental stages…………..47

Table 3.4 Oligonucleotide primer used in this study for whole mount in situ hybridisation analysis……………………………………………………………………………………………………………….48

List of Abbreviations General Abbreviations 3D: three dimensional AP: anterior-posterior AP buffer: Alkaline phosphatase buffer ANOVA: Analysis of variance BCIP: 5-bromo-4-chlore-3-indolyl-phosphate BI: Bayesian Intereference BLASTP: search protein database using protein query bp: base pairs cDNA: complementary DNA DIG: dioxigenin DNA: deoxyribonucleic acid DNase: deoxyribonuclease dNTP: deoxyribonucleotide triphosphate FSW: filtered sea water GBR: Great Barrier Reed GOI: Gene of interest h: hour HB buffer: Hybridization buffer hpe: hour post emergence hpi: hour post induction Kb: Kilo-base pairs MgCl2: Magnesium chloride min: minute ML: Maximum likelihood NBT: nitro blue tetrazolium NJ: Neighbor-Joining PCR: polymerase chain reaction PP: Posterior probabilities qRT-PCR: real time quantitative PCR RNA: ribonucleic acid RNA-Seq: RNA Sequencing RNase: Ribonuclease SDS PAGE: sodium dodecyl sulfate polyacrylamide gel electrophores s: second TBE: Tris-borate EDTA TEOS: tetraethoxysilane UTR: Untranslated region WMISH: Whole mount in situ hybridization

Abbreviations of gene names (Reference genes) EIF: Eukaryotic initiation factor FKBP: FKBP-type peptidyl-prolyl cis-transisomerase HPRT: Hypoxanthine guanine phosphoribosyl transferase NFkB: Nuclear factor kappaB PERO5: Peroxiredoxin 5 RPL13: Ribosomal protein L13 SDHA: Succinate dehydrogenase complex subunit A TETRA: Tetraspanin YWAHZ: Tyrosine-3-monooxygenase/tryptophan5-monooxygenase activation protein

Abbreviations of protein names Ala/A: alanine Asn/N: Asparagine Cys/C: Cysteine Gly/G: Glycine His/H: Histidine Phe/F: Phenylalanine Ser/S: Serine Trp/W: Tryptophan Tyr/Y: Tyrosine

Chapter 1: General Introduction

Biomineralisation processes are associated with the formation of mineral structures common to both eukaryotes and prokaryotes. These structures often serve protective, feeding or supportive functions (Bengtson 1994; Cusack and Freer 2008; Hamm et al. 2003; Palumbi 1986; Swift 2012; Wealthall et al. 2005). Common biominerals include 1) calcium carbonate, which for instance is used to build mollusc shells or sea urchin spines, 2) calcium phosphate, which is found in vertebrate bones and teeth and brachiopode shells, 3) and silica, which forms the skeleton of most sponge, diatom, silicoflagellates, some choanoflagellates, chrysophytes, synurophytes and radiolarians, and small number of dinoflagellates (Beniash et al. 1997; DeMaster 1981; Elliott 2002; Leadbeater and Jones 1984; Pohnert 2002; Preisig 1994; Simpson 1984). Although there are still gaps in our understanding of biomineralisation mechanisms, proteins have been found to govern the formation of many of these diverse structures (Huang et al. 2007; Kröger et al. 1999; Kugimiya et al. 2005; Shimizu et al. 1998). One remarkable example of biomineralised structures is the skeletal framework of sponges, which is composed of fascinating and complex glassy amorphous silica structures. The biomineralisation of these structures, termed spicules, is controlled by the silicatein protein family (Cha et al. 1999; Shimizu et al. 1998).

Biosilicification is the process by which organisms absorb and accumulate silicic acid from their environment and deposit it as amorphous silica (Exley 2009; Tréguer et al. 1995). Each year, gigatonnes of silicon are processed by marine organisms to build and harden their silica skeletons (DeMaster 1981; Epstein 1994; Schröder et al. 2008; Simpson and Volcani 1981; Tréguer et al. 1995; Wang et al. 2007). For instance, the estimate consumption of silica per year by marine sponges is 8.6 X 1010 to 7.3 X 1012 mol Si year-1, while the accumulation of silica in sponge’s skeletons is estimated at 19 X 1012 mol Si (Maldonado et al. 2011; Maldonado et al. 2012). Among the organisms that possess siliceous skeletons, diatoms and sponges are the best characterised (Hildebrand 2008; Simpson and Volcani 1981; Uriz et al. 2003). A large variety of shapes and sizes of biosilica structures can be observed in diatom skeletons (termed frustules) and sponges (Boury-Esnault and Rützler 1997; Chanas and Pawlik 1995; Picket et al.

1990; Simpson 1984; Uriz 2006). Skeletal structures are typically species-specific and have been used as a taxonomic character for both sponges and diatoms (Bavestrello et al. 1993b; Picket et al. 1990; Round et al. 1990; Simpson 1984).

Biosilica formation is of high interest for the industrial and medical sectors. Silica in its different forms is widely used in everyday products such as glasses, paints, adhesives, stabilizers, food additives and high-tech products (Schröder et al. 2007a; Schröder et al. 2007b). Industrially produced silica-based materials require extremely high pressures and temperatures, and yield significant amounts of harmful chemical waste (Morse 1999; Müller et al. 2008a). In contrast, sponges have this unique ability to form these highly ordered and hierarchical structures under physiological conditions, at ambient temperature and pressure. Sponge are therefore of increasing scientific and economic value as the mechanism responsible for the formation of their biosilica structures is of potential importance for the fabrication of novel silica material under unique properties.

1.1 Sponge biosilica structure

Sponges (phylum Porifera) evolved approximately 700 million years ago (Erwin et al. 2011) and today inhabit both freshwater and marine water worldwide (Van Soest et al. 2012). They are phylogenetically considered the oldest extant metazoan phylum that use silica for the formation of their mineral skeleton (Bavestrello et al. 1993b; Cha et al. 1999; Müller 1995; Müller 1998) (Fig. 1.1). The mineral skeleton of all hexactinellids and most demosponges is composed of amorphous silica; Calcarea build spicules made of calcium carbonate (Bergquist and Sinclair 1973; Imsiecke et al. 1995; Simpson 1984). Most Homoscleromorpha, recently considered as the fourth class, also produce silica skeleton (Gazave et al. 2012; Maldonado and Riesgo 2007).

Figure 1.1 Evolutionary tree showing poriferan relationship to each other and to other metazoans (Wörheide et al. 2012).

1.1.1 Spicule morphology Sponge siliceous spicules display a large variety of shapes and sizes, and they have a characteristic species-specific morphology (Boury-Esnault and Rützler 1997; Hartman 1981; Lévi 1973). They are classified into two classes according to their size, shape and function. Megascleres are larger with lengths ranging from a micrometre to a centimetre, have a simple needle-like shape, and form the skeletal framework of the sponge (Berquist and Sinclair 1973;

Boury-Esnault and Rützler 1997; Uriz et al. 2003). Spicules can be joined by collagenous material like sponging depending on the sponge specie and group (Uriz et al. 2003; Uriz 2006). Exceptionally, the spicule of the hexactinellids sponge Monorhaphis chuni can measure up to 1 to 2 m due to its extracellular formation (Müller et al. 2007a). The total length of spicules in demosponge typically does not exceed a few millimetres due to the mechanisms behind its formation. Indeed, dimensions are limited by the size of the cells involved in the secretion of the axial filament (Uriz et al. 2000; Weaver et al. 2010). Demosponges and hexactinellids can be differentiated based on their meglasclere terminations and number of axes of symmetry, with demosponges having monaxons and tetraxons, and hexactinellids having monaxons and triaxons (Uriz et al. 2003; Uriz 2006). In contrast, microscleres are smaller, more complex in shape and are widely distributed within the sponge body. These may assist to reinforce the sponge skeleton (Berquist and Sinclair 1973; Boury-Esnault and Rützler 1997; Uriz et al. 2003).

1.1.2 Spicule formation Siliceous spicules of demosponges are known to contain an axial filament composed of protein (Shore 1972). However, the mechanism of spicule formation had remained unclear until studies in two demosponges, Tethya aurantium (Cha et al. 1999; Cha et al. 2000; Shimizu et al. 1998) and Suberites domuncula (Krasko et al. 2000), revealed that the axial filament of siliceous spicule was enzymatically fabricated via a protein called silicatein. From these studies and subsequent analyses, spiculogenesis has been divided into three phases. There is an initial intracellular phase where the axial filament made of silicatein is produced, initiating spicule formation and two extracellular phases where subsequent spicule growth occurs (Fig. 1.2) (Müller et al. 2005a; Shimizu et al. 1998; Wilkinson and Garrone 1980).

Figure 1.2. Schematic of spicule formation. (A) Formation of spicule starts in the intracellular space of the sclerocyte. Silicasomes (sis) absorb silica (Si) from the surrounding environment in order to form an axial filament that develops by apposition of silica layer to an immature spicule. This spicule is then extruded to the extracellular space (mesohyl) where subsequent growth occurs. Silicatein released from silicasomes present in the mesohyl bind with galectin molecules in presence of Ca2+ to form a biosilica lamellae (cy). This newly formed structure is layered onto pre-existing ones. A mature spicule is formed when superposed lamellae fuse to create a solid rod. (B) Axial growth of the spicule occurs when silicasomes (sis) present within the axial canal (ac) release silicatein to form a biosilica lamellae (a-si) on the inter-wall of the spicule (Wang et al. 2011b).

Initial phase in the intracellular space Spicule formation appears to start intracellularly within specialised cells called sclerocytes (Simpson 1984; Wilkinson and Garrone 1980). However, this perspective has been debated by Uriz et al. (2000) who proposed that the production of large spicule types (e.g. large megasclere) is initiated in the extracellular mesohyl. Because spicule formation is a rapid process, it remained unclear for a long time if spiculogenesis begins intra- or extracellularly. Progress in cell biology with the development of the primmorph system from Suberites domuncula sponge allowed for a more detailed analysis of spicule formation (Müller et al. 2005a; Müller et al. 2006). Primmorphs are three-dimensional (3D) aggregates containing

proliferating cells, and are generated from cell dissociation. These aggregates have a tissue- like appearance and can be cultured for more than 5 months (Müller et al. 1999; Müller et al. 2005a). Studies on primmorphs have confirmed by electron microscopic imaging that the axial filament is formed in the sclerocyte within a silicasome (Müller et al. 2005a; Müller et al. 2006). Silicasomes actively absorb and accumulate silica from the surrounding environment via Na+/HCO3-[Si(OHO4] cotransporter (Schröder et al. 2004; Schröder et al. 2007a; Wang et al. 2012). Silicatein accumulates in the silicasome along with the silintapin-1 protein (Wiens et al. 2009). Thereafter, the first silica layer is deposited around the filament to form an immature spicule which is released into the mesohyl where its final shape and size are determined (Fig. 1.2A) (Müller et al. 2005a; Schloβmacher et al. 2011; Wang et al. 2011b).

Spicule growth in the extracellular space While spicule formation in the intracellular space is well understood, the extracellular growth of the spicule remains unclear. Spicules growth in the extracellular space might not be observed in every sponge species, and may not be necessary in embryo, larvae and early juveniles as the mechanism of spicule formation in these early developmental stages might slightly differ from the adult. However, several studies have been done to elucidate the different processes associated with the growth of the spicule in the extracellular space.

Spicule growth is mediated by the release of silica and silicatein from silicasomes. These vesicles are present within the axial canal as well as around the spicule in the mesohyl (Müller et al. 2005a; Schröder et al. 2006; Wang et al. 2011a), leading to axial and radial growth of the spicule. The axial growth corresponds to the deposition of silica lamellae on the inner surface of the spicule within the axial canal (Wang et al. 2011a) (Fig. 1.2B). Radial growth corresponds to the deposition of silica lamellae from the mesohyl onto the surface of the spicule. Silicatein appears to bind with galectin molecule in the presence of Ca2+ and silintaphin-2 proteins to form a lamellar structure orientated by collagen around the growing spicule (Müller et al. 2005a; Schröder et al. 2006; Wang et al. 2010; Wiens et al. 2011). A mature spicule is formed when the apposition of those lamellae fuse to form a solid siliceous layer around the filament (Müller et al. 2009).

Spicule morphology in the extracellular space

The mechanisms regulating spicule morphology remain unknown (Boury-Esnault and Rützler 1997; Chanas and Pawlik 1995; Uriz 2006). However, it has been suggested that the final shape of the spicule is driven by the shape of the filament (Ecker et al. 2006; Piserra 2003). The orientation and arrangement of the biosilica lamellae by collagen fibres may also provide a platform in determining the final shape of the spicule (Ecker et al. 2006; Müller et al. 2007a; Müller et al. 2009).

1.2 Silicatein

The axial filaments of siliceous spicules found in demosponges consist predominantly of silicatein protein. Three silicatein isoforms have been identified, silicatein α, silicatein β, and silicatein γ. Their molecular masses have being determined by SDS PAGE to be 29, 28 and 27 kDa respectively, in Tethya aurantium (Shimizu et al. 1998), while the dominant silicatein protein in Suberites domuncula is 24 kDa (Müller et al. 2005a). Silicatein is a member of the papain-like cysteine protease superfamily, belonging more specifically to the cathepsin family with highest similarity to cathepsin L (Krasko et al 2000; Shimizu et al. 1998). In fact, silicateins are reported to differ from cathepsin L by one amino acid in the catalytic triad. In silicatein the catalytic triad is composed of Serine (Ser/S) - Histidine (His/H) - Asparagine (Asn/N); while the Ser is replaced by Cysteine (Cys/C) in cathepsin L (Shimizu et al. 1998). Another specific feature is the presence of a serine rich cluster in silicatein α (Shimizu et al. 1998).

1.2.1 Catalytic mechanism of silicatein The Ser residue present in the catalytic centre of the silicatein molecule instead of the Cys in cathepsin L appears to be important for the catalytic function of the enzyme (Krasko et al 2000; Shimizu et al. 1998). The proposed 3D structure of silicatein shows that these three amino acids (the SHN triad) are part of the active site of the enzyme, while a serine cluster is located at the surface of the molecule (Schröder et al. 2007b). Using silicatein purified from Tethya aurantium spicules, it was demonstrated that the protein catalyses the synthesis of silica from tetraethoxysilane (TEOS) monomers, which is a silica precursor, at neutral pH in vitro (Cha et al. 1999). Although this observation in vitro has not been observed in living sponges, as TEOS is not a natural precursor, it still has biotechnological interest (Cha et al. 1999). Finally, site-directed mutagenesis experiments revealed that the interaction of the

hydroxyl group of the Ser residue with the His residue in the active site is essential for the catalytic mechanism, as the enzymatic activity is prevented when Ser or His was replaced (Zhou et al. 1999).

The protein silicatein not only has a catalytic activity but also has been shown in situ to have a proteolytic activity similar to cathepsin L (Müller et al. 2003; Müller et al. 2007b). Müller et al. (2007b) showed that the axial filament of Lubomirskia baicalensis can cleave a cathepsin L substrate that was labelled with Rhodamine 110+bis-[CBZ-L-Phe-L-Arg] as strong filament staining reflecting biosilica synthesis was detected by fluorescent microscopy after 30 min of incubation. The specificity of the silicatein activity was established using the substrate for another proteinase, elastase (Rhodamine 110+bis-[CBZ-L-Ala–L-Ala]), that could not be cleaved by the filaments (Müller et al 2007b).

1.3 Amphimedon queenslandica as a study model

The demosponge Amphimedon queenslandica (Porifera, Demospongiae, Haploscleridae, Niphatidae) was first discovered on Heron Island reefs in the Capricorn-Bunker Group, southern Great Barrier Reef (GBR) (Hooper and Van Soest 2006), but its distribution extends to One Tree Island and Magnetic Island reefs (Degnan et al. 2008a), related populations have been found in Japan and the Red Sea (Degnan et al. 2008a). A. queenslandica is the first sponge species to have its genome sequenced, assembled and annotated and currently is one of the best models to study sponge development; it broods embryos and larvae year-round, facilitating the access to embryological and larval material (Leys and Degnan 2001; Srivastava et al. 2010).

Amphimedon queenslandica has a biphasic life cycle with a short and well-defined planktonic phase. Larvae are released from brood chamber that may contain embryos from different developmental stages at any one time (Leys and Degnan 2001). The planktonic larval stage may last several hours but larvae are competent to metamorphose after six to eight hours (Degnan and Degnan 2010). Settlement and metamorphosis are induced when the competent larva comes in contact with an inductive environmental cue (Degnan and Degnan 2010).

Numerous molecular techniques have been developed for this animal (Adamska and Degnan 2008; Degnan et al. 2008b; Larroux et al. 2008; Leys et al. 2008) and its development and behaviour are well understood (Adamska et al. 2007; Degnan and Degnan 2010; Leys and Degnan 2001).

Many sponge larvae are known to possess spicules (Leys 2003), including the demosponge Amphimedon queenslandica which start to fabricate spicule early in development (Degnan et al. 2008). The spicules present in larvae usually differ in size and in shape from the adult sponge; larval spicules tend to be smaller. Indeed, the different types of spicule present in the adult are usually absent in larvae, especially the microscleres (Leys 2003). Amphimedon queenslandica larvae and adult both lack microsclere and only possess megascleres (Hooper and Van Soest 2006). Different type of spicule are observed in the adult, the most common being the oxea megasclere, approximately 80-100 µm long and 1-2.3 µm wide (Hooper and Van Soest 2006; Leys and Degnan 2001). Embryo, larvae and post-larvae possess only one type of spicule, the oxea, being approximately 25 and 40 µm long, 0.5 and 0.6 µm wide respectively for the mature spicule. In A. queenslandica, other spicule types present in the adult probably appear later in the development of post-larval life as only one type of has been observed in early juveniles, up to 88h old.

1.4. Aims of this study

In this thesis I aim to provide a better understanding of how spiculogenesis is genetically regulated. By using A. queenslandica I can trace the expression and role of silicatein genes through embryonic, larval and postlarval development. Although recent studies have revealed the role of silica in biosilicification, its developmental expression through embryogenesis, larval development and postlarval development are currently unknown for any sponges. Outcomes of this research will provide insight into how silicatein genes are expressed over the different developmental stages of the sponge and how this expression correlates with the rate of spicule formation.

I first identified all the silicatein genes in the A. queenslandica genome, completing the first genome-wide study of this gene family. I undertook a comprehensive phylogenetic analysis of this gene family to determine how it evolved in A. queenslandica and other sponges. This analysis is presented in Chapter 2 and provides the foundation for further molecular research of this gene family in A. queenslandica. In Chapter 3, I gain insight into the expression of silicatein genes throughout the different developmental stages and relate these expression profiles and patterns to spiculogenesis. This process has not been investigated in natural sponge development. Finally, in the general discussion (Chapter 4), I summarise these results and relate them to prior work on silicatein in other sponges.

Chapter 2: Identification of silicatein genes in the demosponge Amphimedon queenslandica and their evolutionary relationship to other sponge silicatein and cathepsin genes

2.1 Abstract

Silicatein genes are involved in the process of spiculogenesis in sponges (Porifera), and act as a template for the siliceous spicule formation. In this study, six putative silicatein genes were identified and isolated in the genome of the demosponge Amphimedon queenslandica. Protein sequence analysis reveals high sequence identity with previously identified sponge silicateins despite the absence of the conventional catalytic serine (Ser/S). Gene models Aqu2.08677, Aqu2.29399, Aqu2.41047 and Aqu2.41046 contain the conserved catalytic residues Cysteine (Cys/C) - Histidine (His/H) - Asparagine (Asn/N) characteristic of cathepsin L. Gene trees generated using silicatein and cathepsin L sequences, using Bayesian and maximum likelihood approaches, are similar and suggest the presence of four silicatein subclades. Subclades I to III represent marine demosponge species and are comprised of silicatein β sequences (subclade I and II), and of silicatein α and β sequences (subclade III). Subclade IV only has members of the freshwater sponge families. Phylogenetic analysis confirms that the isolated A. queenslandica genes belong to the silicatein family and more precisely to the silicatein β subfamily. Further analyses suggest that the presence of the Cys residue in some silicatein genes represents an intermediate form between cathepsin L and silicatein genes; and that the Tyrosine (Tyr, Y) residue following this catalytic Ser/Cys might be as important. Additionally, the presence of the CY residues have been observed in cathepsin L sequence of other metazoan species suggesting a common ancestor and an unrelated function to spicule formation in these species. Characterisation of the exon-intron structure of genes encoding silicatein and cathepsin L reveals that exon length and exon- intron border phase are highly conserved within most sponge silicatein genes and subset of cathepsin L genes.

2.2 Introduction

Siliceous sponges (phylum Porifera) have the ability to synthesise silica spicules enzymatically via the protein silicatein to form their skeletal framework (Cha et al. 1999; Müller 1995; Müller 1998; Shimizu et al. 1998). Demospongiae, Hexactinellida and Homoscleromorpha are the three classes of Porifera that display this biosynthetic capability, producing morphologically diverse siliceous spicules (Boury-Esnault and Rützler 1997; Imsiecke et al. 1995; Simpson 1984).

While biosilica structures have been identified in sponges since the 19th century (Borojevic et al. 1968; Delage 1892; Donati 1753; Grant 1826; Haeckel 1972; Schulze 1904), it is only with the recent progress in molecular biology that the mechanism of spiculogenesis has been understood, especially with regards to its developmental and genetic regulation. The protein primarily responsible for the biosynthesis of silica spicules in demosponges and hexactinellids is called silicatein and was originally isolated from the axial filament in the spicules of the marine demosponges Tethya aurantia (Shimizu et al. 1998). Subsequently, three closely related silicatein subunits, α, β and γ were identified, all having similar amino acid compositions (Shimizu et al. 1998). Molecular data have also revealed the importance of silicatein in the biosilicification process of siliceous sponges.

Protein sequence comparisons and phylogenetic analyses of silicatein, the α subunit specifically, indicate that it is a member of the cathepsin family, being most closely related to cathepsin L. It is thought that silicatein arose in the phylum Porifera from a cathepsin L-like ancestor, which is a member of the papain-like cysteine protease superfamily (C1 family) (Krasko et al 2000; Shimizu et al. 1998).

In T. aurantia, two silicatein protein isoforms have been identified, silicatein α, and silicatein β (Shimizu et al. 1998). In the following years, gene encoding silicatein α and silicatein β proteins have been found in Suberites domuncula (Krasko et al. 2000; Schröder et al. 2005). Afterward, silicatein has been isolated from other marine sponges, including Petrosia ficiformis (Pozzolini et al. 2004), Hymeniacidon perlevis (Cao et al. 2007), Cratomorpha meyeri (Müller et al. 2008b), Latrunculia oparinae (Kozhemyako et al. 2009) and Geodia cydonium

(Müller et al. 2007c), as well as freshwater sponges including Lubomirskia baicalensis (Kaluzhnaya et al. 2005; Kaluzhnaya et al. 2011), Ephydatia fluviatilis (Funayama et al. 2005; Mohri et al. 2008) and Ephydatia muelleri (Kaluzhnaya et al. 2011).

Silicatein α and silicatein β have been detected at the gene level in several marine sponges species. However, freshwater sponges seem to lack silicatein β but often code for four isoforms of silicatein α (e.g. E. fluviatilis and L. baicalensis; Kaluzhnaya et al. 2005; Kaluzhnaya et al. 2011; Mohri et al. 2008). In E. fluviatilis primmorph the different silicatein α isoforms are expressed at different times during the formation of spicules (Mohri et al. 2008), suggesting that each silicatein gene plays a different role in spicule growth and morphology.

Comparison of the protein sequence between silicatein and cathepsin L reveals a characteristic difference in one residue of the cathepsin catalytic triad, which in the latter sequence consists of three amino acids: Cys-His-Asn. In silicatein Cys is replaced with Ser (Shimizu et al. 1998). This Ser residue is thought to be involved in the catalytic mechanism of the enzyme (Krasko et al 2000; Shimizu et al. 1998). Recently, however, by comparing the silicatein sequences of multiple demosponge species, three amino acids, tyrosine (Tyr, Y), alanine (Ala, A) and phenylalanine (Phe, F), seem to always follow the Ser of the catalytic triad and Cys, Glycine (Gly/G) and Ala are before this residue (Veremeichik et al. 2011) (Fig. 2.1). Thus, silicateins may not only be defined by the presence of a Ser in the catalytic triad but by the following sequence, CGASYAF. Another distinctive component of silicatein, specifically silicatein α is the presence of specific hydroxyl amino acid clusters represented by either Ser- Ser-Arg-Cys-Ser-Ser-Ser-Ser (SSRCSSSS), Ser-Ser-Cys-Thr-Tyr (SSCTY), or two Ser-Xaa-Ser-Xaa- Ser (SXSXS) sequences. The most common serine cluster is SSRCSSSS, which is thought to serve as template for silica deposition (Shimizu et al. 1998).

Silicatein and cathepsin L have six conserved Cys residues that are involved in the formation of disulphide bridges in both, suggesting that the three dimensional structure of silicatein and cathepsin is similar (Shimizu et al. 1998). Additionally, similarity between the propeptide of silicatein and cathepsin L from sponge and human further support a common evolutionary origin (Shimizu et al. 1998).

Figure 2.1 Detailed comparison of conserved amino acids of demosponge silicatein Sequence alignment of freshwater (E. fluviatilis (BAG74346.1) and L. baicalensis (CAI43319.1)) and marine sponges (P. ficiformis (AAO23671.1), T. aurantium (a- AAC23951.1; b- AAF21819.1) and S. domuncula (a- CAI46305.1; b- CAI46304.1)). The catalytic serine (Ser/S) is marked by an arrow. The conserved residues of all demosponge silicatein are underlined. The amino acids following the catalytic Ser, tyrosine (Tyr, Y), alanine (Ala, A) and phenylalanine (Phe, F) are indicated by a black line. The preceding amino acids, Cysteine (Cys/C), Glycine (Gly, G), and Ala, are indicated by a dashed line.

The aim of the present study is to identify and analyse silicatein genes in the demosponge Amphimedon queenslandica, whose genome has been fully sequenced (Srivastava et al., 2010); and show their evolutionary relationship to other silicatein and cathepsin genes. My analysis shows the presence of both conventional and non-conventional silicatein genes in A. queenslandica, with four out of six putative genes lacking the serine residue in the catalytic triad. A comprehensive phylogenetic analysis of this gene family has demonstrated a high level of sequence similarity between A. queenslandica and other demosponges silicateins despite the absence of the catalytic Ser residue. Further amino acid analyses suggest that silicatein genes can be defined by the presence of either SY or CY; and that the serine cluster representing silicatein α, as previously described, is present only in very few demosponges sequences. Additionally, the CY motif has been observed in cathepsin L genes in diverse species suggesting a shared evolutionary function in poriferan and the common ancestor of all metazoans. The investigation of the genomic organisation of silicatein genes shows a well- conserved genomic architecture such as exon length and intron/exon border phases.

2.3 Materials and Methods

2.3.1 Identification of silicatein genes in Amphimedon queenslandica The sequenced genome of A. queenslandica was screened for sequences encoding the conserved domains of silicatein, peptidase C1A and inhibitor I29. A BLASTP search against the

NCBI non-redundant protein sequences was performed on each of the identified sequences to assess their similarity. Additional silicatein and cathepsin L protein sequence from several sponge species were retrieved from the NCBI database and their accession numbers are listed in Appendix A1. Each protein sequences were scanned with the InterProScan plugin implemented in Geneious (R6) Pro software (Biomatters Ltd.) to identify and verify the conserved region position of domains in proteins and genes.

2.3.2 Gene architecture analyses Silicatein and cathepsin L genes in A. queenslandica, S. domuncula (silicatein α and β) and L. baicalensis (Silicatein α1, α2, α3, α4, and cathepsin L), were obtained in order to determine their gene structure. As exon length conservation often correlates with intron phase (Long et al. 1995), lengths of exons and introns were calculated, and intron/exon border phase were manually determined. Intron phase refers to the position of an intron splice site within or between a codon with respect to the open reading frame of the mature mRNA (Sharp 1981). Three different intron phases can be observed, phase 0, 1 and 2. Phase 0 corresponds to the placement of an intron between two codons. Phase 1 corresponds to the location of intron between the first and second nucleotides of a codon, while phase 2 intron is between the second and third nucleotides of a codon (Long et al. 1995; Sharp 1981). Gene structure and domain organisation were designed using Fancy Gene v1.4 (http://bio.ieo.eu/fancygene/) (Rambaldi and Ciccarelli 2009).

2.3.3 Molecular phylogenetic analyses Molecular phylogenetic analyses were carried out using the bioinformatics tools provided by the Geneious (R6) Pro software (Biomatters Ltd.). Alignments used for the multiple phylogenetic analyses were based on the two conserved domains only, Inhibitor I29 and Peptidase C1, discarding any redundant sequences between domains. All multiple sequence alignments were performed using the MAFFT program (v7.017) with default settings (Katoh et al. 2002). They were then edited visually to remove redundant sequences. Phylogenetic trees were generated using Bayesian Interference (BI), Maximum likelihood (ML) and Distance (Neighbor-Joining, NJ) methods. Before performing ML and BI analyses, the most appropriate substitution model that would best fit the data was determined using the ProtTest program (v2.4) (Abascal et al. 2005). The best model was WAG, which was applied

for the BI and ML analyses, after further trial tests using different substitution models (Blosum, LG and Dayhoff) showed similar tree topologies. Maximum likelihood based phylogenetic analyses were performed using the PhyML program (Guindon and Gascuel 2003). One thousand bootstraps replicates were performed to obtain statistical support for the phylogenetic trees generated. Distance Neighbor-Joining (NJ) trees were generated with 1000 bootstraps using the Jukes-Cantor genetic distance model with default settings. Bayesian analyses were performed using Mr Bayes program (v.3.2) (Huelsenbeck and Ronquist 2001). Analyses were run for 1,100,000 generations, with 4 chains, sampling every 200 generations. Trees from the first 100,000 generations (200 trees) were discarded to ensure that burn-in trees were excluded, leaving 2000 trees to construct the 50% majority- rule consensus. Confidence at each node was assessed using the posterior probability values (PP). In every analysis, cathepsin B from A. queenslandica was used as the outgroup.

2.4 Results

2.4.1 Identification and genomic organisation of A. queenslandica silicatein genes Analysis of the genome of A. queenslandica revealed six putative silicatein genes: Aqu2.08677, Aqu2.29399, Aqu2.41047, Aqu2.41046, Aqu2.42494, and Aqu2.42495. Alignment of the A. queenslandica silicatein sequences with representative silicatein and cathepsin L genes of S. domuncula is presented in Fig. 2.2. Only two A. queenslandica genes, Aqu2.42494 and Aqu2.42495, have the characteristic silicatein serine (Ser/S) residue in the catalytic triad (Ser-His-Asn). Analysis of the amino acid sequences encoded by the remaining four A. queenslandica genes (Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047) reveals they share characters of both silicatein and cathepsin L. These sequences contain a Cys residue instead of a Ser in the catalytic triad of the protein, which is characteristic of cathepsin L. However, in all A. queenslandica sequences this Cys is followed by Tyr-Ala-Phe (YAF) residues, which are specific to silicatein (Fig. 2.2). Interestingly, an analysis of amino acid sequence similarity between the novel A. queenslandica sequences and S. domuncula silicatein and cathepsin L indicates that all A. queenslandica genes are most related to silicatein than to cathepsin sequences. The putative proteins encoded by the cathepsin-like silicatein genes Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047 show highest amino

acid identity to the S. domuncula silicatein α (47.6%, 47.3%, 57.7% and 58.2% respectively), while Aqu2.42494 and Aqu2.42495 have a high identity with S. domuncula silicatein β (60.4% and 52.2% respectively).

A. queenslandica silicatein sequences differ from those in other demosponge species by the absence of the characteristic and highly conserved G residue in the motif preceding the catalytic Ser/Cys. It is replaced by a Cys in Aqu2.42495, and His in Aqu2.08677 and Aqu2.29399 (Fig. 2.2). The serine cluster CSSSS representing silicatein α, which is thought to act as template for silica deposition (Shimizu et al. 1998), is absent in all A. queenslandica silicatein genes. Finally, the last residue of the six conserved cysteines, which form a disulphide bridge and play an important role in protein folding and stability, is missing from the silicatein gene Aqu2.42495 (Fig. 2.2).

All three phylogenetic models group the six identified A. queenslandica genes into the silicatein clade with strong to moderate support (PP: 0.97, ML: 74.2 %, NJ: 66%) (Fig. 2.3A) even though they lack the catalytic serine. Two silicatein genes, Aqu2.08677 and Aqu2.29399 form a separate clade, with strong support values (PP: 1, ML: 100, NJ: 100) within the silicatein clade (Fig. 2.3A). Five A. queenslandica cathepsin L genes, Aqu2.41049a, Aqu2.41049b, Aqu2.41050, Aqu2.43732, and Aqu2.43733, confidently form one clade (PP: 0.97, ML: 85.5%, NJ: 100%), within the cathepsin L gene family (Fig. 2.3A) (Appendix A2).

Figure 2.2 Comparison of the protein sequence of Amphimedon queenslandica (Aqu2.) silicatein genes with homologous sequence from the sponge S. domuncula. Conserved amino acid are highlighted in black (100% similarity) and similar amino acid (>60%) are highlighted in grey. Arrows indicate the catalytic amino acids, Cysteine (Cys/C) or Serine (Ser/S), Histidine (His/H) and Asparagine (Asn/N). Key conserved silicateins residues are underlined. The Cys residues involved in the formation of disulphide bonds are represented by dots and the serine cluster by a box.

Figure 1.3 Phylogenetic relationships and gene structure of Amphimedon queenslandica silicatein and cathepsin L genes family. (A) The substitution model WAG was used. Posterior probabilities (PP) and bootstrap proportion percentage (1000 replicates for maximum likelihood (ML) and Neighbor-joining (NJ)) are given (PP/ML/NJ). The outgroup represents cathepsin B from A. queenslandica. (B) Gene architectures of each silicatein/cathepsin L gene. Exons are represented by rectangles where the color scheme is red, UTR region; blue, inhibitor I29 conserved domains; yellow, peptidase C1 conserved domain. Introns are represented by thin lines.

Figure 2.3 Phylogenetic relationships and gene structure of Amphimedon queenslandica silicatein and cathepsin L genes family.A The substitution model WAG was used. Posterior probabilities (PP) and bootstrap proportion percentage (1000 replicates for maximum likelihood (ML) and Neighbor-joining (NJ)) are given (PP/ML/NJ). The outgroup represents cathepsin B from A. queenslandica. B. Gene architectures of each silicatein/cathepsin L gene. Exons are represented by rectangles where the color scheme is red, UTR region; blue, inhibitor I29 conserved domains; yellow, peptidase C1 conserved domain. Introns are represented by thin lines. 19

2.4.2 Conservation in the genomic structure of A. queenslandica silicatein genes Analysis of the genomic contigs bearing the six A. queenslandica silicatein genes reveals that these genes range in size from 1.11 to 1.80 kb and are composed of 5 to 8 exons. The genes each encode two highly conserved domains, Inhibitor I29 and Peptidase C1A (Fig. 2.3B). The total exon/intron lengths of the silicatein genes are 822/288 (Aqu2.08677), 1048/393 (Aqu2.29399), 1385/418 (Aqu2.41046), 1014/318 (Aqu2.41047), 1122/475 (Aqu2.42494) and 756/382 (Aqu2.42495). The length of individual introns range in size between and within genes from 46 to 156 bp. The lengths and organisation of the exons encoding the peptidase C1 domain appears to be very conserved in A. queenslandica (Fig. 2.4). This region is also conserved in S. domuncula and L. baicalensis silicatein genes, as well as some cathepsin L genes in A. queenslandica and L. baicalensis (Fig. 1.4). The exon structure comparison of silicatein and cathepsin L genes showed conservation in exon length across exons 3 to 7. With regards to their length, exons 3 to 5 appear to be the most conserved within the silicatein and cathepsin L family, the length being 115, 110 and 153 bp respectively (Fig. 1.4 red box). One exception is Aqu2.42494, which shares the conserved exon lengths in exon 5 to 7 instead, suggesting this gene has gain 2 exons. The length of exon 6 (119 bp) is also highly conserved across sponge silicatein and cathepsin L genes (Fig. 2.4), while the 100 bp length of exon 7 exists in L. baicalensis and S. domuncula silicatein α but only present in two of the A. queenslandica silicatein genes (Aqu2.41046 and Aqu2.41047) (Fig. 2.4 green box). However, the combined length of exon 6 and 7 in these two genes corresponds to the length of exon 8 in Aqu2.42494 suggesting loss of an intron. Six A. queenslandica genes, three cathepsin L (Aqu2.41049a, Aqu2.41049b and Aqu2.43732) and three silicatein (Aqu2.42495, Aqu2.41046 and Aqu2.41047), have identical genomic structure from exon 3 to exon 6 (Fig. 2.4 blue box). A. queenslandica silicatein genes Aqu2.08677 and Aqu2.29399 do not display this conserved gene structure, although both have similar exon structures (exon 2 to exon 5) (Fig 2.4 purple box). The absence of this conserved exon structure in these genes is supported in the phylogenetic tree as they form an independent subclade, with strong support values (PP: 1, ML: 100, NJ: 100), within the silicatein clade (Fig. 2.3 A and Fig. 2.5, subclade I).

Intron phase analysis showed that each intron phase is also highly conserved in A. queenslandica silicatein genes (Table 2.1). Introns 1, 2, 4 and 5 are in phase 0, intron 3 is in phase 1 and intron 6 in phase 2. This intron phase sequence is also observed in S. domuncula

and L. baicalensis silicatein genes. The intron phase sequence corresponding to these silicatein genes is 5’-0-0-1-0-0-2-3’. This sequence can also be observed in 5 cathepsin L genes (Aqu2.41049a, Aqu2.41049b, Aqu2.43732, Aqu2.43733 and Aqu2.43734). These same genes have the conserved exon structure (Fig. 2.4).

Figure 2.4 Comparison of exon structure of silicatein and cathepsin L genes of A. queenslandica with S. domuncula and L. baicalensis. Exons are represented by a box, each containing the individual exon length (bp), while the untranslated regions are illustrated by dashed boxes. The conserved inhibitor I29 domain is blue, peptidase C1 domain is yellow. Introns are represented by the ‘V’ shape line and are not drawn to scale. Coloured boxes highlight exons of identical length between genes. Red boxes corresponds to genes with identical exons 3 to 5, blue boxes correspond to exons 3-6, and green boxes correspond to exons 3-7. The purple box corresponds to the uniquely identical exon length in two A. queenslandica silicatein genes. The purple box corresponds to identical exon structure of cathepsin L within A. queenslandica.

Table 2.1 Intron phases for A. queenslandica silicatein and cathepsin L genes, and S. domuncula and L. baicalensis orthologues. A generalised 7-exon silicatein gene model is shown for reference

2.4.3 Phylogenetic relationship of sponge silicateins The Bayesian tree generated from 95 unique silicatein and cathepsin L protein sequences from 26 sponge species (19 demosponges and 7 hexactinellids; Appendix A1) shows a number of distinct clades, especially within the silicatein family (Fig. 2.5). Cathepsin L genes are divided within the tree into two distinct groups depending on the sponge class, hexactinellid or demosponges. The phylogeny revealed a well-defined and supported silicatein clade (PP: 1, ML: 75.2, NJ 53.6), that can be divided into four subclades. The first three subclades (I, II and III) correspond to marine sponge species and subclade IV represents freshwater sponge species (PP: 1, ML: 95.5, NJ: 53). The branching within each of these clades might differ slightly as UTR and redundant regions were discarded due to gaps throughout available sequences. Indeed, these regions may contain sequence elements important for gene expression and regulation.

These analyses indicate that silicatein α and β isoforms do not form two distinct clades, as they are distributed among subclades I, II and III. Instead they form individual clusters within

these subclades. For instance, subclade II represents silicatein β only from five sponge species including A. queenslandica (PP: 0.79, ML: 24.5), while subclade III can be separated into two distinct clusters, silicatein α/β (PP: 0.53) and silicatein α (PP: 0.99, ML: 50.7). An unexpected grouping of hexactinellid sequences within demosponge group in the silicatein cluster α/β is seen, but they do not form a monophyletic group. This branching topology might be a consequence of the lack of information for the sequence of the hexactinellid species as only a partial sequence was used. The clear lineage-specific gene duplication that can be observed between freshwater and marine species suggests a monophyletic freshwater silicatein protein family. This clear cluster distinction between marine and freshwater sponges has been previously noted (Mohri et al. 2008; Veremeichik et al. 2011). However, a well- supported sequence from the freshwater sponge E. fluviatilis can be observed within the silicatein β clade and appears to be most similar with silicatein from S. domuncula. (PP: 1, ML: 87.5). The phylogenetic analysis of Veremeichik et al. (2011) also showed this improbable grouping. From this analysis I can infer that the silicatein evolved from a cathepsin L ancestor and that silicatein α evolved from a silicatein β ancestor.

Three A. queenslandica silicatein genes, Aqu2.41046, Aqu2.42494 and Aqu2.42495 are placed within a specific clade (subclade II), although its node support is weak (PP: 0.79, ML: 24.5). These three genes appear to be orthologous to silicatein β from four other demosponges: S. domuncula, Petrosia ficiformis, Latrunculia oparinae and the freshwater sponge Ephydatia fluviatilis. The analysis confidently places Aqu2.42494 within that silicatein β cluster (PP: 1, ML: 89.8, NJ: 53.9), being more closely related to P. ficiformis (PP: 1, ML: 99.8, NJ: 99.9). A cathepsin sequence from L. oparinae also belongs to this β clade, but the analysis of its protein sequence alignment reveals the CYAF cluster residues. Aqu2.08677 and Aqu2.29399 are forming a monophyletic clade (subclade I) and represent the basis branch from which silicatein originates. Other marine sponge species’ silicateins are principally placed in the well- supported multispecies subclade III.

Homologous sequences corresponding to four freshwater silicatein isoforms (α1, α2, α3 and α4) are grouped into four clusters with strong support (α1- PP: 1, ML: 96.7, NJ: 97.8); α2- PP: 1, ML: 98.6, NJ: 99.9; α3- PP: 1, ML: 99.8, NJ 100; α4- PP: 1, ML 76.1, NJ 62.9). The freshwater silicatein α3 cluster forms an independent branch located closer to the common ancestor of

freshwater sponge’ silicatein genes. Phylogenetic analysis suggests that these four silicatein- α isoforms have evolved from gene duplication events from a common ancestor.

One distinction that can be noted between marine and freshwater sponges is the number of silicatein genes present within one species. Freshwater sponges usually have multiple copies of silicatein genes, six for E. fluviatilis and four for B. fungiformis and L. baicalensis, while marine sponges only have one or two copies with the exception of L. oparinae (five gene sequences) and A. queenslandica (six genes). In A. queenslandica, the different numbers of cathepsin L genes are probably a consequence of species-specific gene duplication and divergence as 7 of the 10 sequences form a monophyletic clade (Aqu2.41049a, Aqu2.41049b, Aqu2.41050, Aqu2.43732, Aqu2.43733, Aqu2.33917 and Aqu2.27307 (and Sil 2 Hymeniacidon perlevis). This grouping is arguable as its node support significantly differs between the two phylogenetic analyses (PP: 0.88, ML: 3.7). The presence of the H. perlevis gene within this grouping may affect this result as the alignment show that the relatively short sequence it encodes is missing part of the catalytic domain. Five of those genes (Aqu2.41049a, Aqu2.41049b, Aqu2.41050, Aqu2.43732, Aqu2.43733) form a well-supported independent branch (PP: 0.99, ML: 75, NJ: 74.4). The remaining three cathepsin L genes (Aqu2.22535, Aqu2.39162 and Aqu2.43734) group together in the Bayesian analysis and are more closely related to the common ancestor of cathepsin L. In the ML analysis, the gene Aqu2.39162 groups weakly with C. meyerini, P. raphanus, A. vastus, Aulosaccus sp. and Bathydorus sp. proteins (ML: 24.6 %).

2.4.4 More detailed analysis of silicatein protein sequences Silicatein and cathepsin L have high sequence identity and similarity but one main difference is the replacement of the catalytic Cys in cathepsin L by Ser in silicatein (Shimizu et al. 1998). Previous studies showed that cathepsin L sequences from distantly-related vertebrates could confer a silica condensing activity by the replacement of not only the catalytic Cys by a Ser but also the adjacent amino acid (CW to SY) (Fairhead et al. 2008; Ki et al. 2013). These studies suggest therefore a more similar function between silicatein and cathepsin than previously thought.

Analysis of the protein sequences of each of the sponge silicateins in this study reveals that the Tyr (Y) following the catalytic Cys/Ser is the most highly conserved amino acid within the silicatein family; in Cathepsin L it is replaced with tryptophan (Trp/W) (Fig. 2.6A). The Ser alone cannot define silicatein genes since a number of proteins within silicatein group I/II and III have the Cys instead of the Ser. I conclude from these data that silicatein can either be defined by SY or CY, while cathepsin L is defined by CW. It also noted that the presence of the CY is only observed in the silicatein β genes (group I/II and III), which supports silicatein β being an intermediate form between cathepsin L and silicatein α (c.f. Fig.2.5). Regarding the amino acid motif preceding the catalytic Cys/Ser, silicatein β once again shares some residues with cathepsin L. Cathepsin L is characterised by CGS. Similarly, silicatein β from group I/II often possess CGS (Fig. 2.6A), while silicatein α from group III and IV typically have a CGA motif with the exception of four sequences from sil-α3 in group IV, which form an independent branch (Fig. 2.5). Therefore, silicatein β in marine sponges and silicatein α-3 in freshwater sponges share trait of both cathepsin and silicatein, suggesting that they form the basal clade of their specific grouping. The AF residues downstream of the catalytic S/CY is also highly conserved within the silicateins and less so in cathepsin L (Fig. 1.6A). The serine rich motif (SSRCSSSS) that has been used to define silicatein α does not appear to be as conserved within silicatein sequences as previously thought (Fig. 2.6B). This motif is mainly observed in sequences from silicatein group III, which show high conservation on the SSR cluster. Only four sequences from group III (sil-α/β and sil-α) show the CSSSS serine cluster, namely sil-α S. domuncula, sil-α T. aurantium, sil M. chuni and sil-α3 L. oparinae, consistent with the motif not being necessary for silicification.

Figure 2.6 Sequence motifs of the two main regions defining silicatein genes. A. Region containing the Cys/Ser in the catalytic triad. B. Serine rich cluster region. The sequences were searched for common motif using the MEME suite. The consensus logos for these two regions in each cathepsin L and silicatein group are shown.

2.4.5 Presence of the CY motif in other metazoan cathepsin L sequences. Biosilica is formed by different marine organisms including diatoms, crustaceans, ascidians and sponges (Boury-Esnault and Rützler 1997; Lowenstam 1971; Miller et al. 1990; Monniot et al. 1992; Round et al. 1990; Simpson and Volcani 1981). Sponges are phylogenetically the oldest extant phyletic lineage of metazoans that use silica for the formation of their skeletal framework (Müller 1995; Müller 1998). As described above silicatein and cathepsin L share high sequence similarity but one major distinction between them is the presence of the CY/SY in silicateins, CY being proposed to be an intermediate form between cathepsin L and silicatein α. Currently, however, there is no evidence that CY-containing silicateins catalyse silica spicule formation.

With the new insight that CYAF motif can define silicatein, I sought to determine if silicatein- like sequences exist elsewhere in the animal kingdom. Recent phylogenetic analyses proposed that ctenophore are the most basal animal lineage; although their phylogenetic relationship to the other early branching metazoan phyletic lineages (Cnidaria, Placozoa and Porifera) remains uncertain (Dunn et al. 2008; Philippe et al. 2009; Ryan et al 2013; Schierwater et al. 2009). For instance, it has been proposed based on the presence or absence of homeodomains genes that ctenophore may be more closely related to sponges (Ryan et al. 2010). A BLASTP search of predicted transcripts from the genome of the ctenophore Mnemiopsis leidyi. (http://research.nhgri.nih.gov/mnemiopsis/pfam/pfam_domain.cgi) and Pleurobrachia bachei (http://moroz.hpc.ufl.edu/slimebase2/browse.php) with silicatein sequences of A. queenslandica showed the presence of the CY/SY amino acid residues within cathepsin L sequences (Fig. 2.7). The phylogeny suggests that five genes from M. leidyi (ML007332a, ML012013a, ML071151a, ML098612a and ML10109a) and 3 genes from P. bachei (sb3473063, sb3461712 and sb3468590) are silicatein-like genes as they branch off with silicatein from A. queenslandica (P:0.87, ML:6.3 %) (Fig. 2.7A). Interestingly, five sequences from P. bachei having the CY residues form a distinct clade within the cathepsin L family sister to a clade containing mostly Amphimedon and Mnemiopsis cathepsin L sequences. This branching pattern might be the result of the absence of the AF residues following the CY (Fig. 2.7B), but also of the characteristic amino acids of the catalytic triad His and Asn in some proteins (sb3479662, sb3471699 and sb3468548). Further analysis reveals the presence of the CY residues within cathepsin L sequences of different taxa (Fig. 2.8). This suggests that this particular gene is possibly ancient and has been maintained in different groups, but its function in biosilicification has probably exclusively evolved in poriferans.

Figure 2.7 Phylogenetic relationship between silicatein and silicatein like genes of the sponge Amphimedon queenslandica and ctenophores Mnemiopsis leidyi and Pleurobrachia bachei. A. Bayesian phylogenetic tree. The substitution model WAG was used. Posterior probabilities (PP) and bootstrap proportion percentage (maximum likelihood (ML) with 1000 replicates) are given (PP/ML). The outgroup represents cathepsin B from A. queenslandica. The sequences name ML correspond to M. leidyi and sb to P. bachei ctenophore. B. Protein allignment of the catalytic Cys/Ser region.

A B

X Predicted emergence of silicatein-like sequences Presence of silicatein or silicatein like sequence (SY/CY) in organisms Absence

Figure 2.8. Predicted emergence of silicatein like genes in metazoan derived from sequences encoding the characteristic SY/CY (Serine- Tyrosine/Cysteine-Tyr). Presence (star) or absence (square) of silicatein/ silicatein-like sequences is indicated for each metazoan phylum (A) and for the four classes of poriferan (B). Name and sequence number for each analyse species are indicated in Table 2 .2.

Table 2.2 EnsemblMetazoa accession numbers of silicatein-like sequences in different animals.

2.5 Discussion

In this study, I characterised the silicatein genes in the demosponge Amphimedon queenslandica, as this remains the only sponge with a predicted published genome (Srivastava et al. 2010). I then explored the evolution of silicatein genes first in sponges and then in metazoans. I identified six silicatein β genes in A. queenslandica of which four can be characterised as non-conventional. I will categorise these genes as either type I (Aqu2.42494 and Aqu2.42495) or type II (Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047). Type I correspond to conventional silicatein characterised by the serine (Ser, S) in the catalytic triad, while type II is described as non-conventional silicatein by the catalytic cysteine (Cys, C).

Sequences analysis revealed that Amphimedon queenslandica type II silicatein genes (Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047) potentially represent a new type of silicatein as they contain both silicatein and cathepsin L characteristics (i.e. CYAF) (Fig. 2.2). The catalytic region of the protein contains a Cys in the catalytic triad (Cys-His-Asn), which is characteristic of cathepsin L (Berti and Storer 1995; Shimizu et al. 1998), instead of Ser, which is thought to be essential for the catalytic mechanism of the silicatein (Krasko et al 2000; Shimizu et al. 1998; Zhou et la. 1999). This type II silicatein has a Tyr, which is specific to silicatein. Using a mutated cathepsin L from human it was demonstrated that the surrounding residues of the catalytic Ser/Cys played an important role in silica condensing activity, since the mutation of the cysteine alone did not result in a significant activity (Fairhead et al. 2008). The A. queenslandica genes have either a SYAF or CYAF motif, the latter also being present in the demosponge Latrunculia oparinae cathepsin sequence (ACH48003.1) (Kozhemyako et al. 2009) and in the hexactinellid Aulosaccus schulzei (ACU86976.1 and ACU86974.1) (Veremeichik et al. 2011). This putative catalytic difference within the silicatein gene family may represent an intermediate position of these type II genes between the sponge cathepsin L and type I silicatein. At this time, it is unclear if type II silicateins can promote biosilicification in sponges.

Silicatein encoding genes of A. queenslandica contain between 5 to 8 exons (Fig. 2.3). Three of these genes have the conserved 7 exons observed in the silicatein genes of L. baicalensis and S. domuncula (Fig. 2.4) (Kaluzhnaya et al. 2011; Müller et al. 2003; Schröder et al. 2005).

The addition or loss of one or more exons has also been reported in S. domuncula silicatein β genes (Schröder et al. 2005). This gene structure modification may result in the loss or gain of particular functions of the silicatein protein (Kaluzhnaya et al. 2011). Silicatein proteins have a dual function, a structure-guiding function which provide biosilica a structural support, and a structure-forming with an enzymatic function (Müller et al. 2014). Structure of silicatein genes such as exon length and intron phases are generally conserved. Length of exon 3 to 5 in A. queenslandica is highly conserved in silicatein genes, but also present in a number of cathepsin L genes. This observation has previously been reported between eight freshwater sponge species (L. baicalensis, B. intermedia, L. incrustans, B. fungiformis, S. lacustris, E. muelleri, E. fluviatilis and S. papyracea) by Kayluzhnaya et al. (2011). Conserved Intron phases has also been noticed in both freshwater and marine sponges, where intron phase in silicatein α of S. domuncula and L. baicalensis follows the same phase sequence as in A. queenslandica silicatein genes (intron phase sequence: 5’-0- 0-1-0-0-2-3’) (Table 2.2) (Belikov et al. 2007; Müller et al. 2007b). While the intron sequence phase of S. domuncula silicatein β is 5’-0-0-1- 0-2-3’ (Müller et al. 2007b). Thus, my analysis of A. queenslandica silicatein genes on exon length and intron phases suggests that marine and freshwater sponges share genomic features which are likely to reflect a common ancestral state.

The phylogenetic tree obtained in this study for silicatein and cathepsin L genes from several sponges species (Fig. 2.5), suggests the presence of four silicatein clades. The result also shows that all A. queenslandica silicateins are assigned with confidence to the silicatein clade, suggesting that type II proteins are true silicateins despite not having a Ser in the catalytic triad. These type II (CY) gene sequences do not form a monophyletic group at the base of the silicatein clade implying that they do not form an intermediate group between cathepsin L and type I silicatein, but that they originated from a common ancestor of all silicatein gene.

The silicateins present in Amphimedon are only present in Clade I and II (Fig. 2.5 ). Clade I, is the smallest silicatein clade and is uniquely composed of two type II A. queenslandica sequences (Aqu2. 08677 and Aqu2. 29399). The sequences from this clade have been described as being silicatein β genes but can possibly be silicatein γ, which would be the first identification of this silicatein isoform at the gene level. Initially, three closely related silicatein protein subunits: α, β and γ have been dissociated in T. aurantian axial filament, all having

similar amino acid compositions and present in the relative proportion α:β:γ = 12:6:1 (Shimizu et al. 1998). Currently two out of three silicatein protein isoforms found in the axial filament have been identified at the gene level in demosponges, silicatein α and silicatein β. Aqu2. 08677 and Aqu2. 29399 are more closely related to silicatein β (Clade II) than to silicatein α (Clade III), suggesting that they are silicatein β genes. However, these two genes are distinctly separated from their homologues and may represent the third silicatein isoform, silicatein γ.

Clade II is composed of silicatein β genes belonging to four species, including the remaining four A. queenslandica silicatein sequences (Fig. 2.5). Two species, S. domuncula and L. oparinae also appear in clade III. Clade III differs from clade I and II in that it is the largest and most diverse. This clade can be divided into three groups which seem to represent different isoforms of silicatein, since sequences from the same species do not necessarily branch together (S. domuncula, L. oparinae, T. aurantium and H. perlevis) and thus, are not grouped depending on species nor family. The first and second group appear to be a combination of two sponge classes, demosponge and hexactinellid, and represent Sil. α/ β cluster, while group two (Sil. α/ β) and three (Sil. α) form sister groups. One can wonder if silicatein β sequences from group two (Acanthodendrilla sp., T. aurantium and L. oparinae) are not misnamed, as they appear to be more closely related to silicatein α than to Clade II silicatein β genes. If not, the two sister group might represent two different isoform of silicatein α as seen in freshwater sponge (Clade IV) (Fig. 2.5).

The first two silicatein clades which regroup the A. queenslandica genes correspond to silicatein β, and are more closely related to the silicatein common ancestor, suggesting that silicatein α arose from the β isoform by gene duplication. Both silicatein and cathepsin L genes in A. queenslandica are phylogenetically very close and located at the basal branch of their respective clades (Fig. 2.5), suggesting the emergence of silicatein and cathepsin L of other demosponges from this group. This result implies that A. queenslandica silicatein genes have evolved separately from the other demosponges species used in this phylogenetic study. The phylogeny of demosponges remains unresolved to some extent. However, recent studies using mitochondrial genes have provided insights into demosponges relationship and shown that the class Demospongiae is divided into five main clades, G0 (Homoscleromorpha), G1 (Keratosa), G2 (Myxospongiae), G3 (marine Haplosclerida) and G4 (Others demosponges)

(Lavrov et al. 2008; Wörheide et al. 2012). The distinction seen between A. queenslandica and other demosponges might be the result of being part of two different clades. Indeed, A. queenslandica and P. ficiformis are part of the G3 clade while the other species used in this phylogeny are in the G4 clade (Lavrov et al. 2008). Interestingly, only one silicatein gene is expressed in P. ficiformis, silicatein β gene, which correspond to the one type of spicules found in the adult sponge (Armirotti et al. 2009; Pozzolini et al. 2004). As A. queenslandica appears to only have silicatein β gene, this finding suggests that the silicatein α isoform arose in the G4 clade.

I detected for the first time the occurrence of silicatein-like genes in other metazoan taxa (Fig 2.7; Fig. 2.8 and Table 2.2). The sequences analysis (Appendix A3) revealed that these genes do not only show the CY/SY characteristic of silicatein genes, but are also highly similar in their protein composition to sponge silicatein. Moreover, most of these sequences have the CGS/CGA and AF residues on either side of the catalytic C/S (Table 2.2). This indicated that silicatein-like genes evolved earlier than previously thought and have an ancestor common to all metazoans. However, no evidence of siliceous biomineralization products in these species can be found in the literature. Within the Porifera, we can notice a possible loss of the CY in the calcareous class (Sycon ciliatum). On the other hand, presence of silicatein genes with SY motif (type I silicatein) can be observed within non-spicule forming demosponge (G1: Keratosa, Acanthodendrilla.sp Vietnam) (Kozhemyako et al. 2009) (Fig. 2.5). The genomic structure of these silicatein-like sequences in the different taxa do not appear to show the exon length conservation seen in sponge silicatein gene. Until now, the serine (S) in the catalytic triad of silicatein genes was thought to be the main characteristic and specific to poriferans. We have shown in this study, that silicatein can also be defined by the cysteine (C) and that the presence of the tyrosine (Y) is specific to silicatein. One possible explanation is that the role of the protein has evolved in poriferans and relate to spiculogenesis in demosponge and hexactinellid, while it may perform a cathepsin L like function in other species.

In this study, I identified two types of silicatein genes in A. queenslandica. I found that type II silicatein gene has only been observed in silicatein β genes. Phylogenetic and protein sequence analyses show that silicatein β is the intermediate form between cathepsin L and

silicatein α, and that silicatein α arose from silicatein β by gene duplication in the G4 clade of demosponges. The result also shows that the Ser cluster used to identify silicatein α is not as conserved as initially though (Fig. 2.6). The redefinition of silicatein genes (type I and II) and the presence of silicatein-like genes in other metazoans show that further investigation of the protein gene family might shed new light.

Chapter 3: Analysis of spicule formation and silicatein gene expression in the demosponge Amphimedon queenslandica

3.1 Abstract

Demosponges, hexactinellids and homoscleromorphs have a skeleton composed of siliceous spicules with species-specific morphologies. Silicatein, a protein localised in the axial filament of the spicule, is the major component of these highly structured skeletal elements and is thought to contribute in determining their structure and shape. Spicules, which are either widely distributed in the sponge body or fuse to form the skeletal framework, are fabricated and secreted by specialised cells called sclerocytes. Using the demosponge Amphimedon queenslandica, I provide the first data set of temporal and spatial expression of silicatein genes on a sexual reproductive sponge through embryonic, larval and post-larval development. Additionally, I provide insight on the rate of spicule formation in this demosponge. Quantitative analyses indicate that all A. queenslandica silicatein genes are differentially expressed throughout development, and four of these six genes (Aqu2.41046, Aqu2.41047, Aqu2.42494 and Aqu2.42495) display similar expression profiles. These data also show that A. queenslandica silicatein genes are first expressed in early embryos prior to the formation of mineralised spicules. Although spicule number does not appear to increase in larvae and juveniles, Aqu2.42494 and other silicatein genes are expressed during these stages. Whole mount in situ hybridisation data analysis demonstrates that Aqu2.42494 is expressed in sclerocyte cells from embryonic to post-larval stages. Localised expression across development shows a progressive migration of sclerocytes cells from the outer cell layer of cleavage embryos to the inner cell mass of larvae.

3.2 Introduction

Marine organisms use a variety of biominerals to build and harden their skeletons. For instance, sponges, one of the oldest metazoans phyla, convert silicate absorbed from the surrounding environment into silica, which then serves as structural material (Cha et al. 1999; DeMaster 1981; Epstein 1994). The most diverse and widespread class among the four sponge

classes is the Demospongiae, and these fabricate fascinating and complex silica structures, termed spicules. These spicules often form the skeletal framework of adult sponges, which confers strength and protection from the surrounding environment (Bond 1992; Imsiecke et al. 1995; Palumbi 1986; Simpson 1984). Understanding the developmental basis of spiculogenesis provides a means to understand how these spicules and skeletons form in this ecologically and evolutionarily important group of animals.

Spicule development and morphology have been characterised since the early 1900s (Bergquist and Sinclair 1973; Borojevic et al. 1968; Schulze 1904; Shore 1972; Simpson 1984). However, the more recent discovery that the formation of siliceous spicule is enzymatically mediated via silicatein was a major breakthrough in the understanding of spiculogenesis (Cha et al. 1999; Shimizu et al. 1998). Further studies based on immunofluorescence and electron microscopy have improved our understanding of the different phases of spicule formation. Spiculogenesis is characterised by an initial intracellular phase within specialised cells called sclerocytes, where dissolved silicon is absorbed and immature spicules are formed. This is followed by an extracellular phase where spicule growth is driven by the concentration of soluble silica present in the surrounding environment. The final morphogenesis of spicules is poorly characterised (Chombard et al. 1998; Mohri et al. 2008; Uriz 2006). It has been suggested that the final shape of the spicule is driven by the shape of the filament (Ecker et al. 2006; Piserra 2003). However, despite spicule size and shape being highly diverse and species-specific, the basic structure remains the same. All spicules have an axial canal that is a few micrometres widths that contains a proteinaceous filament structure (axial filament) around which a silica layer is deposited (Shore 1972; Müller et al. 2005a).

The high diversity in spicule morphology has been used as a taxonomic character to identify genera and species (Boury-Esnault and Rützler 1997; Chanas and Pawlik 1995; Simpson 1984; Uriz 2006), as their morphogenesis is genetically controlled (Bavestrello et al. 1993a; Ecker et al. 2006). However, environmental conditions appear to influence both spicule shape and size. It was showed that the expression of a particular spicule type is dependent on the concentration of silicic acid in the surrounding water, which may vary with season (Bavestrello et al. 1993b; Maldonado et al. 1999). Regardless of shape and size, silicatein appears to be the main component of all siliceous spicules studied, apparently contributing to structure and

shape (Cha et al. 1999; Piserra 2003; Shimizu et al. 1998). Silicatein is present specifically in siliceous sponges (demonsponges, hexactinellids and homoscleromorphs) and was first discovered in the axial filament of the demosponge Tethya aurantia where two genes have been isolated, silicatein α and silicatein β (Shimizu et al. 1998). Silicatein genes have since been identified in other marine and freshwater sponges (Krasko et al. 2000; Pozzolini et al. 2004; Cao et al. 2007; Kaluzhnaya et al. 2005; Funayama et al. 2005; Mohri et al. 2008).

Although recent studies have revealed the role of silicatein in biosilicification, its developmental expression from embryogenesis to post-larval development is not known in any sponges species. It is also unknown how silicatein expression correlates with spicule number. Fabrication and secretion of spicules can occur at different developmental stages of the sponge, either during embryogenesis, larval phase and metamorphosis, through to the development of the juveniles and growth of the adult. However, specific details on the ontogenetic basis of spicule formation are scant (Berquist and Sinclair 1968; Leys 2003; Maldonado et al. 1997; Woollacott 1990). Therefore, to better understand the process of spiculogenesis during natural sponge development- embryonic, larval and post-larval stages – using a sexual reproductive system, spicule formation and silicatein expression were analysed in the demosponge A. queenslandica.

In this chapter, I analysed the expression of the six silicatein β genes identified and characterised in the previous chapter (Aqu2.08677, Aqu2.29399, Aqu2.41046, Aqu2.41047, Aqu2.42494 and Aqu2.42495), among which four (Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047) are type II genes described as non-conventional by the presence of the cysteine (Cys, C) instead of the serine (Ser, S) in the catalytic triad of the protein. The expression profile of these genes was characterised through development and related to the accumulation of spicules in embryos, larvae, postlarvae and juveniles. Using whole mount in situ hybridisation (WMISH), I characterised the expression patterns of type I silicatein gene, Aqu2.42494. These analyses demonstrate that silicatein expression is restricted to sclerocytes and precedes the formation of the mineralised spicules, consistent with the hypothesis that this protein is required in spiculogenesis.

3.3 Materials and Methods

3.3.1 Sponge collection and fixation of biological materials.

Adult A. queenslandica were collected from Shark Bay reef flat of Heron Island Reef GBR at low tide as described in Leys et al. (2008) and transferred to outdoor aquaria that were supplied with flow-through lagoonal seawater. Embryos, larvae and postlarvae were procured following procedures in Leys et al. (2008). For whole mount in situ hybridisation (WMISH), individual embryos, larvae and postlarvae were fixed in 4% paraformaldehyde, 0.05% glutaraldehyde and 5 X MOPs buffer for 1 h on a nutator, following the procedure of Larroux et al. (2008). After fixation, samples were transferred into and stored in 70% ethanol at -20oC. For RNA-Seq and qRT-PCR methods, individuals were stored in RNALater at 4oC overnight and then at -20oC.

3.3.1.1 Brooded embryos Brood embryos are found in chambers, containing as many as 100 embryos of different developmental stages (Leys and Degnan 2001). Stage differentiation is made based on embryo pigmentation (Adamska et al. 2007). The cleavage stage embryo is the earliest stage of development, and is mainly found around the edges of the brood chambers (Fig. 3.1 A-B). The brown stage corresponds to the localisation of pigment cells on the embryo surface, giving the embryo its beige colour (Fig. 3.1 C). Cloud and spot stage embryos are when the pigment cells migrate to form a cloud over the posterior pole and then aggregate to form a spot (Fig. 3.1 D-F), respectively. The final stage of embryonic development is when embryos have a bi-layered appearance with a well-defined pigmented ring (Fig. 3.1 G-I) (Adamska et al. 2007; Degnan et al. 2005).

Removal of embryos from the brood chamber was performed following the procedure of Leys et al. (2008). Slices of adult sponges containing brood chambers were transferred and pinned into a petri dish containing a thin base of agarose and filtered sea water (FSW). Under a dissecting microscope, using fine forceps, sharpened pin and a Pasteur pipette, embryos were detached from the connective tissue of the brood chamber and fixed for WMISH, RNA-seq and qRT-PCR as described above.

Figure. 3.1 Embryonic development of Amphimedon queenslandica. (A&B) Early cleavage and cleavage embryo; (C) brown embryo; (D) cloud embryo: pigment cells starting to migrate toward the posterior pole, forming a cloud; spot embryo: cells coming together (E) to form a well-defined spot (F); ring embryo: pigment cells start to migrate outward (G) to form a well-defined pigmented ring (H). Mature larva defined by a bi-layered appearance ready to be released from the brood chamber (I). Adapted from Richards (2010).

3.3.1.2 Larval and post-larval stages Approximately 10 sponges were placed in a large black nally bin for spawning. Release of larvae was triggered by heating the surrounding seawater by up to 3oC above ambient water temperature between 12 pm and 2 pm, as described in Leys et al. (2008). Larvae were collected over a 2 h period, washed in 0.2 µm filtered sea water (FSW) and then stored in FSW

at 25oC with regular changes of water (Leys et al. 2008) until the appropriate fixing time. Precompetent larvae correspond to free swimming larvae up to 6-8 hours post emergence (hpe). Competent larvae were either kept as free swimming or transferred at 8 hpe into six well plates (10 larvae/well), filled with 5 ml FSW, containing pieces of the coralline algae Amphiroa sp(Degnan and Degnan 2010). Plates were kept in the dark in order to induce settlement. To ensure simultaneously development, at 1 hour post induction (hpi), free swimming larvae were removed from wells. At approximately 4-6 hpi, newly settled larvae were transferred into 24 well plates (1 larvae/well), each well containing a small round coverslip onto which they continue to metamorphose. Plates were left in the dark to allow resettlement, with a daily change of water.

3.3.2 Spicule preparation Spicule count preparation and results were obtained from WMISH fixed embryos, larvae and postlarvae collected in Sept 2010 and Jan 2011. Larvae and postlarvae were fixed at 0, 2, 4, 6, 9, 12, 18, 24, 36, 48, 60, 72 and 96 hpe. Juveniles were therefore between 1 and 88 h old from the time of induction. 0 hpe corresponds to 0 to 1 h after larvae have been released from the adult sponges; the same time window can be applied to the other time periods.

The spicule preparation protocol was performed as described by Cao et al. (2007), with some modifications. To maintain the original number of spicules, fixed embryos, larvae and postlarvae were transferred individually on a grid glass slide, in 1-2 µl of 70% ethanol. Samples were then washed twice in distilled water and air dried for 5 min. To obtain spicule preparation, sponge tissue was dissolve in concentrated (~11.1 M) nitric over a hot plate set a 90oC, and the slide was carefully shaken in order for the spicules to spread apart. A few drops of acid were added before complete evaporation, thereby preventing tissue to stick to the slide, which would result in clump of spicules. The process was repeated at least 5 times for complete tissue digestion. Spicules were counted under a compound microscope. Spicules in the earliest phase of formation were discarded in the count as they clumped at the edge of the grid and could not be differentiated from broken spicule tips.

3.3.3 Gene expression analysis – RNA-Seq protocol

Developmental RNA-seq data were generated by Dr Nagayasu Nakanishi and analyse to compare silicatein expression profile across development. RNA was extracted from the cleavage, brown, spot and ring stages embryo, precompetent (0-2 hpe) and competent (9- 11 hpe) larvae, and postlarvae aged 0-1, 23-24 and 71-72 hpi.

3.3.3.1 RNA extraction and cDNA synthesis Total RNA was extracted from pooled samples, composed of 3 to 10 individuals with three replicates per stage, using Trizol reagent (Invitrogen). RNA was quantified using fluorometric quantitation (Qubit 2.0 Fluorometer) and purified by DNase I (Invitrogen) treatment according to manufacturer’s protocol. Complementary DNA (cDNA) was synthesised with Random pentadecamers using dNTP mix and 5X First Strand buffer (Invitrogen), DTT, RNAsOUT (Invitrogen) and Superscript III Reverse Transcriptase (Invitrogen), according to the manufacturer’s instruction. Quality of cDNA was assessed using agarose gel electrophoresis (1.5% TBE) with No-RT control for each sample to detect any genomic DNA contamination.

3.3.3.2 Samples processing Approximately 100 ng of total RNA was used for strand specific library construction and single-end sequencing on the Illumina Hiseq-2000 platform. The sequence read was generated using Illumina consensus assessment of sequence and variation (CASAVA1.8.2) and analysed according to AGRF quality control measures. The clean reads were then aligned against the A. queenslandica genome available at http://metazoa.ensembl.org/Amphimedon_queenslandica/Info/Index and map to the genomic sequence using the Tophat aligner. Transcripts were then assembled using the reference annotation based transcript assembly option (RABT).

3.3.4 Gene expression analysis – qRT-PCR In order to compare gene expression amongst same developmental stage individual, quantitative real time polymerase chain reaction (qRT-PCR) was performed on the selected genes of interest (GOI) (Table 3.1) using a Roche Lightcycler 480 Real-Time PCR System (Roche Applied Science). These three silicatein genes were chosen based on their expression level and because amplification and optimisation of the lower expressed genes were unsuccessful and required more time. This analysis was generated from 1) cleavage, brown, cloud, spot

and ring embryos, 2) newly emerge precompetent larvae (0-2 hpe), larvae reaching competency (6-8 hpe) and competent larvae (24 hpe) and 3) 24, 48 and 72 hpi postlarvae.

3.3.4.1 Primer design for genes of interest Oligonucleotide primers for the target genes Aqu2.41046, Aqu2.41047 and Aqu2.42494 were designed using Primer3Plus software (Rozen and Skaletsky 2000) and synthesised by Sigma. Optimal parameters for all qRT-PCR primers were to generate a product size of 100-250 bp, have a GC content of approximately 50% and an average melting temperature of 60oC. Gene- specific primer sequences used in this experiment are listed in Table 3.1. When possible and in order to detect any genomic DNA contamination, primer pairs were designed to span exon- exon junctions. Primers were tested for specificity and optimized for concentration and melt temperature to avoid primer dimers.

Table 3.1 Gene specific primer sequences used in this study for quantitative real-time PCR analysis. Primer efficiency (Eff.) and standard error of efficiency (SE Eff.) are given and were calculated by the second derivative method using Light Cycler 480 program (Roche). Reference genes name are abbreviated.

Nucleotide sequence (5' - 3') Product Tm Conc. Primer size Gene name Forward primer Reverse primer (bp) (oC) µM Eff. SE Eff. Genes of interest Aqu2.41046 TGGAATCAACACCGAACAGA TGATGAGTGCATGTGGTCAA 187 58 5 1.82 0.0368 Aqu2.41047 TGCTTTAGAAGGCGCTCAAG TCCACCGTTGCATCCTTTGT 109 58 5 1.805 0.029 Aqu2.42494 GTTGCTCTTGCGGTGATGTA TTGAACTTGCAGCCATCTTG 112 58 10 1.835 0.0808 Reference genes YWAHZ AGCGGAGGGCAAAAAGAA CCTCGGCAAGATAGCGATAGTAGT 185 61 10 1.953 0.0131 SDHA AGGGGAGTGGTAGCTATGAA TGAAACTGTACAAACTCCATGTCT 194 58 5 1.889 0.00903 NFkB TCTCTTACAGCAAACAACTCCTC CTTACCACAGAGAGATTCATTGAC 156 58 5 1.914 0.0113 HPRT CAGACGATGAAAACAAGACTG TAGTAATGAGCAGGGACACAG 127 58 10 1.971 0.00574 PERO5 GCATTGAGAGGAAAGCCACT TGCATCAGGCTCAATAGCAA 179 63 5 1.974 0.00248 FKBP GCTGTGTCAAGGCAAATCCT AGGACACCACTGCTTGTGAA 225 63 10 1.878 0.0136 EIF GCTCTGTCTCATCAGCCACTT CATTGGCTTGATTCCCATTT 164 59 2.5 1.723 0.0288 TETRA GCCACTATCATGACTCTTGCTG CAGCCAACGCTACAATCACT 139 63 2.5 1.845 0.012 RPL13 CTTGGAAGGTTATCGCATGAA TCTTTTTCTCATAATAAATTGCTTCC 103 59 2.5 1.737 0.0992

Table 3.2 Total RNA (ng/µl) used for each sample stages to prepare cDNA

Developmental stages Total RNA ng/µl Cleavage 300-500 Brown 300-500 Embryological Cloud 100-300 Spot 130-150 Late ring 70-100 0-2 hpe 60-100 Larval 6-8 hpe 70-100 24 hpe 70-100 24 hpi 60-100 Postlarval 48 hpi 90-120 72 hpi 60-100

3.3.4.4 qRT-PCR analyses Each 15 µl qRT-PCR reaction contained 3 µl of diluted (1:10) cDNA from an individual sample, 7.5 µl of Lightcycler 480 SYBR green master mix (Roche), 1 µl of primers at specific concentration (Table 3.1) and 3.5 µl of RNAse/DNAse free water. Reactions were performed under the following conditions: An initial denaturing step at 95oC for 10 min; followed by 40 to 50 cycles of denaturation at 95oC for 10 s, annealing temperature (Table 3.1) for 5 s and an extension at 72oC for 10 s. A melt curve analysis cycle of amplified products was also performed at the end of the run to verify the specificity of the reactions. Each assay included in addition to the cDNAs, a no template control reaction where cDNA was replaced with 3 µl of RNAse/DNAse free water and calibrator sample where 2 µl of each sample’s cDNA was pooled, mixed, and 3 µl was used in the reaction. The no template control was used to detect any cross-contamination and primer dimer formation while the calibrator sample was used to account for inter-run variation. Each qRT-PCR reaction was performed in triplicate to account for technical variation, on five individuals per stages.

The relative expression data of the GOI was normalised against the appropriate reference genes (Tables 3.1 and 3.3) to compensate for differences in RNA quality and efficiency between samples. The conversion of quantification cycle values (Cq) into calibrated

normalised relative quantities (CNRQs) was performed using the qBase+ software and a modified delta Ct method (Hellemans et al. 2007; Vandesompele et al. 2002).

Candidate reference genes were chosen from a list of previously optimised genes generated by Dr Kerry Roper and assayed for their suitability in normalizing the data of this experiment. A geNorm analysis of these candidate genes identified two or three stably expressed genes for each embryonic, larval and post-larval stages independently (Table 3.1 and Table 3.3) (Vandesompele et al. 2002).

Table 3.3 List of references genes used in this study for each developmental stages

Developmental stages Reference genes Cleavage HPRT - NFkB - YWAHZ Brown HPRT - EIF Embryological Cloud EIF - PERO5 Spot EIF - FKBP- RPL13 Late ring FKBP - SDHA - TETRA Larval 0-2, 6-8 & 24 hpe EIF - YWAHZ Postlarval 24, 48 & 72 hpi EIF - YWAHZ - PERO5

3.3.5 Whole-mount in situ hybridisation WMISH were performed on fixed embryos, larvae and post-larvae as per Larroux et al. (2008) with minor changes. Three genes were chosen, representing type I (Aqu2.42494 – SYAF) and type II (Aqu2.41046, Aqu2.41047 – CYAF) silicatein genes.

In brief, after rehydratation with PBTS embryos, larvae and postlarvae were treated with proteinase K at 37oC for 10 min, post fixed for 1 h at room temperature in WMISH fixative, and incubated for 4 h at 51oC in HB buffer. Hybridisation with RNA probe was performed at 51oC for 60 h. After blocking, samples were incubated in an anti-DIG antibody solution (1:5000) at 4oC overnight. Signals were developed by using NBT/BCIP staining in AP buffer

solution containing PVA and 1 M MgCl2. Larvae and embryo were cleared in benzyl benzoate: benzyl alcohol (2:1) solution. Whole mount individual samples were observed on compound or inverted microscope, and photographed using a Nikon digital camera.

3.3.5.1 Probe synthesis RNA probes were synthesised from purified PCR product of Aqu2.41046, Aqu2.41047 and Aqu2.42494 silicatein genes, using a 10x DIG RNA labelling mix (Roche) following the manufacturer instructions. Probes were transcribed from either cloned Aqu2.42494 silicatein or specifically designed oligonucleotide primers containing the T7 RNA promoter at the 5’- ends of the reverse primers (David and Wedlich, 2001), Aqu2.41046 and Aqu2.41047. Probe lengths were 849, 811 and 872 bp respectively (Table 3.4). Probes concentration was 1 ng/µl.

Table 3.4 Oligonucleotide primer used in this study for whole mount in situ hybridisation analysis. T7 promoter sequences are underlined.

Nucleotide sequence (5' - 3') Product Gene name Forward primer Reverse primer size (bp) Aqu2.41046 CTCACAGCGTTGACTTTCCA AATTAACCCTCACTAAAGGGGGCTCATCAAAAATGCCACT 811 Aqu2.41047 CACGGAGAAGAGGAAAGCAA AATTAACCCTCACTAAAGGGGTAGCAATCCCGCACTGATT 872 Aqu2.42494 CACAGTTGGAAGGCAACTCATGGGAAG TCCCCAGCTGTTCTTCAAAAGCCAGT 849

3.3.6 Statistical analyses The variance in spicule number within each developmental stage was tested with an one-way analysis of variance (ANOVA). Two-way ANOVAs tested the difference in spicule number between larvae and post-larvae of same developmental stages overtime. For the RNA-seq experiments, a one-way ANOVA in expression over time was performed for each gene, while correlation between mean spicule number and mean gene expression was analysed by simple regression analysis. Assumptions of normality were checked prior to each analysis using Shapiro test, and data were log transformed were assumptions of normality was not met. Tukey HSD post hoc test was performed when necessary.

3.4 Results

3.4.1 Spicule number increases during embryogenesis but not in larvae and early postlarvae This experiment sought to determine the rate of spicule formation over the different developmental stages of Amphimedon queenslandica. Analysis of the embryonic stages enabled me to show that spicule formation starts at the brown stage with an average of 8 spicules/embryo. Spicule number significantly increases as the embryo develops into the larval form (ANOVA, F= 3.8239, P= 0.03885), with the emerging larvae having about 1000 spicules (Fig. 3.2). The most dramatic increases in spicule number per individual embryo occurred between spot and ring stages, and ring and newly hatched larval stages (Fig. 3.2).

Spicule number did not increased throughout the larval (ANOVA, F= 0.9877, P= 0.4887) and post-larval phases (ANOVA, F= 1.1988, P= 0.3509), but instead appeared to fluctuate with time. Mean spicule number in the larval stage varied from 1200 spicules at 2 hpe to 903 at 24 hpe, 1103 spicules at 48 hpe and then down to 806 spicules at 96 hpe. The same level of variation occurred in postlarvae with a mean spicule number of 942 at 9 hpe to 1148 at 10 hpe, compared to 881 spicules at 36 hpe and to 1495 spicules at 72 hpe (Fig. 3.2). Comparison made between free swimming larvae and postlarvae, from time point 12 hpe to 96 hpe, showed that there is no significant difference in spicule number between planktonic and benthic forms (ANOVA, F= 0.1910, P= 0.6652) nor with time (ANOVA, F= 1.5853, P= 0.1780) (Fig. 3.2). Based on these results, there appeared to be no significant change in spicule number after the larva emerged from the mother and in the formation of the juvenile body plan. Differences between time points likely reflected individual variation in spicule number. Also, spicule counts did not measure directly rates of spicule formation, except possible during embryogenesis, as it could not be determined if spicules were being turned over (i.e. being fabricated and degraded) during larval and post-larval development.

Figure 3.2 Rate of spicule accumulation during Amphimedon queenslandica embryonic, larval and postlarval development. The graph shows the mean value at each developmental stage with standard error bar. Spicule count was performed in five embryological stages (dark blue): cleavage; brown; cloud; spot; and ring (n= 3 for each stage). Larval stages (light blue), which represent free swimming larvae age between 0 to 120 hpe (n=3 for time point 0 to 72 hpe, and n=2 for 96 hpe) correspond to a 1 h time window. For instance, 0 hpe equal 0 to 1 h from the time larvae have been released from the adult sponges. Post-larval stages (green) include 10 time points, from 9 to 96 hpe (n= 3 for time point 9 to 72 hpe, and n= 2 for 96 hpe). Induction of metamorphosis occurs 8 hpe when free swimming larvae settle on the coralline alga Amphiroa sp. in the dark. Postlarvae and juveniles are therefore age 1 to 88 hpi, which corresponds to 9 to 96 hpe. Errors bars are displayed at each time points. This experiment commenced during my Honours year and data accumulated during that year have been incorporated into this graph.

3.4.2 Temporal expression of A. queenslandica silicatein genes during development RNA-seq was used to determine the expression level of silicatein genes in A. queenslandica throughout development. The data (Fig. 3.3) showed that the six silicatein genes are expressed in all developmental stages: embryos, precompetent larvae (0-2 hpe), competent larvae (6-8 hpe), postlarvae and juveniles. Aqu2.41046 expression does not significantly change (ANOVA, F=1.425, P=0.243) across the sponge development in comparison to Aqu2.08677 (ANOVA, F=3.918, P=0.0053), Aqu2.29399 (ANOVA, F=2.622, P=0.0348), Aqu2.41047 (ANOVA, F=43.83, P=3.2e-11), Aqu2.42494 (ANOVA, F=20.98, P=2.58e-08) and Aqu2.42495 (ANOVA, F=15.78, P=2.97e-07) (Fig. 3.3). The data also indicated that they are no relevant association between silicatein expression and spicule number during A. queenslandica development (Fig. 3.4); Aqu2.41046 (X2=0.05481, df=1, P=0.266), Aqu2.41047

(X2=0.1481, df=1, p=0.166), Aqu2.42494 (X2=-0.02535, df=1, P=0.4002) and Aqu2.42495 (X2=-0.1267, df=1, P=0.7608). Aqu2.08677 and Aqu2.29399 were omitted in this analysis due to their low and constant expression across the embryological and larval stages (Fig. 3.3).

These data indicate that there is a differential expression in silicatein genes, with the type I gene, Aqu2.42494, being the most highly expressed throughout development. It is expressed approximately 10 times higher than Aqu2.42495, and 4 times higher than Aqu2.41046 and Aqu2.41047. Although Aqu2.42494 was more highly expresses, these four genes had similar developmental expression profiles with one peak in the spot stage embryo and a second peak in metamorphosing postlarvae (Fig. 3.3). The expression remained constant throughout the larval stages, from the newly emerge larva until acquisition of competence. Both Aqu2.08677 and Aqu2.29399 had a similar profile, with low levels of expression that slightly increased from the post-larval stage during early metamorphosis (Fig. 3.3).

The data also revealed that Aqu2.41046, Aqu2.41047 and Aqu2.42494 are the only silicatein genes expressed in early development, peaking in the cleavage stage embryo. In the case of Aqu2.41047 and Aqu2.42494, expression levels in cleavage embryos were at least twice that of all other stages of sponge development, with a net decrease in expression observed to the next brown stage. Aqu2.42495 is expressed later in embryogenesis and has its highest expression in spot embryo (Fig. 3.3). Only Aqu2.41046 and Aqu2.42494 transcripts were at a detectable level in the adult sponge (Fig. 3.3).

Figure 3.3 Normalised transcript levels (transcripts per million) of Amphimedon queenslandica silicatein genes throughout development. Graphs represent the mean value of three technical replicates (pool of 3-10 individuals per replicate), the standard errors of the mean are shown. Squiggle on the y-axis of Aqu2.41047 gene represent a break in the y-axis (transcript levels). Larval stages represent time from which larvae have emerged from the adult sponge (hpe), while postlarval stages correspond to the time induction occurred (hpi).

Figure 3.4 Comparison of mean spicule number with mean silicatein gene expression per stages. Bars graphs represent the mean value of normalised transcript levels obtained from three technical replicates (pool of 3-10 individuals per replicate). Squiggle on the y-axis of Aqu2.41047 gene represent breaks in transcript levels. The smooth red line represent the mean value of spicule number per stages (n=3) (secondary axis). Standard error bars are displayed for both data. Larval stages represent time from which larvae have emerged from the adult sponge (hpe), while post-larval stages correspond to the time induction occurred (hpi).

3.4.3 Expression pattern of A. queenslandica silicatein genes, Aqu2.41046, Aqu2.41047 and Aqu2.42494 Using whole mount in situ hybridisation , I compared the expression patterns in 3 days old juveniles of type I silicatein gene, characterise by the SY motif in the active site (Aqu2.42494), to those of type II characterised by CY (Aqu2.41046 and Aqu2.41047) (Chapter 2). All three genes were predominantly expressed in long spindle-shaped cells reminiscent of sclerocytes without the mineralised spicule (Fig. 3.5). Staining was detected in other cells types to differing extents, especially for the type II silicateins (Aqu2.41046 and Aqu2.41047). Although some of this may have been background, deep staining of some archeocytes and pinacocytes suggested that these genes are expressed in other cell types in addition to sclerocytes. In the case of Aqu2.41047, the astrocyte-shaped cells observed in juveniles are larval sclerocytes (Fig. 3.5D).

3.4.4 Localised expression of Aqu2.42494 during development Analysis of expression of silicatein genes through development was restricted to type I Aqu2.42494 genes, largely because of time restrictions and lack of WMISH material. Aqu2.42494 transcripts were detected in spindle-shaped cells in cleavage stage embryos (Fig. 3.6). It was previously shown by transmission electron microscopy and freeze-fracture scanning electron microscopy that sclerocytes with a central proteinaceous filament appear during cleavage in A. queenslandica (Leys and Degnan 2001). These cells are the predominant type expressing Aqu2.42494 throughout development. Interestingly, staining was absent or less intense in sclerocytes containing a mineralised spicule.

Aqu2.42494 is initially expressed in many cells of the outer layer of cleavage embryo (Fig. 3.6A). In the cloud and spot embryos cells start to migrate toward the inner cell mass (Fig. 3.6B-C), only to be present internally in late embryonic stage, in early ring embryo (Fig. 3.6D). Aqu2.42494 is expressed throughout the inner cell mass of the larvae but seems to be at highest density in the posterior pole (Fig. 3.6E-F), region where larval spicules aggregate (Leys and Degnan 2001). After metamorphosis, Aqu2.42494 expressed cells are sparsely distributed within the postlarvae (Fig. 3.6G-H).

Figure 3.5. Silicatein expression pattern of type I and II genes in juvenile Amphimedon queenslandica using whole mount in situ hybridisation. Type I silicatein gene, Aqu2.42494, characterise by SY is expressed in sclerocyte cells in 3 day old juveniles (oscula stage), with unique spindle shape pattern (A). Staining pattern of type II genes, Aqu2.41046 (B-C) and Aqu2.41047 (D-E), in 3 day old juveniles is inconclusive. However sclerocyte (arrows) lacking spicules but expressing Aqu2.41046 and Aqu2.41047 can be seen, as well as asterocyte-like cells (arrowheads). Scale bars: 100 µm

Figure 3.6 Expression of Aqu2.42494 during A. queenslandica development. Aqu2.42494 is expressed in sclerocyte cells in cleavage stage embryo, principally on the outer layer (A). In cloud, spot and early ring embryo sclerocyte cells slowly migrate toward the inner cell mass (B- D). In larval stages (E-F) expression is localised in the inner cell mass but more specifically toward the posterior pole, where staining associate with spicule location. Sclerocyte cells are widely distributed within postlarval stage (G-H). The photos presented in this figure were obtained and taken by myself and other members of the Degnan lab. Scale bars: 100 µm

3.4.5 Individual variation in gene expression and spicule number Data indicated that Aqu2.42494 has a high expression level and is expressed in sclerocytes throughout A. queenslandica development (Section 3.3.3 - 3.4.2). WMISH data showed that Aqu2.41046 and Aqu2.41047 may possibly have a role in spiculogenesis, at least in juveniles. Given this, and the variable number of spicules between individuals of the same stage development (Fig.3.2 and Fig. 3.7A), I sought to determine if these genes displayed variable levels of expression in individuals of same developmental stages. To do this, I employed a qRT-PCR approach, analysing expression levels in individual embryos, larvae and postlarvae.

Spicule number tended to be variable within individuals in the same developmentally staged (Fig. 3.2 and Fig. 3.7A). Thus to determine if variation in gene expression levels correlated with variation in spicule number, I measured the expression levels of Aqu2.41046, Aqu2.41047 and Aqu2.42494 in five individuals at 11 different developmental stages- 5 embryonic, 3 larval and 3 postlarval. Spicule counts were made on three individuals at for each of these stages. Because of the incompatibility with spicule preparations and RNA extractions, I was unable to obtain expression levels and spicule counts from the same individual.

Individual differences in spicule number were observed from late embryonic stages onwards. For instance, the number of spicules in pigmented ring stage embryos varied between 172 and 917 (Fig. 3.7A). Although this variation could have been explained by a subtle developmental difference within a stage (e.g. early ring versus late ring embryo), variation was also observed in similar aged larvae and postlarvae. For instance, number of spicules in 6-8 hpe larvae varied from 831 to 1473, and in 48 hpi postlarvae from 660 to 1884 (Fig. 3.7A).

Similarly, expression levels of the three silicatein genes were variable in the five unrelated individuals at same developmental stage. Differential expression was observed in embryos, larvae and postlarvae, and variation seemed independent of ages and genes (Fig. 3.7B). The expression levels in 0-2 hpe (Aqu2.41046) and 6-8 hpe (Aqu2.42494) larvae, and 72 hpi postlarvae (Aqu2.41046) were less variable that the other developmental stages (Fig. 3.7B). Although there was no correlation between expression levels and spicule number, it was noted that gene expression variation in embryonic development was markedly higher than

spicule number variation. This observation was consistent with the in situ hybridisation results, which demonstrated that gene expression occurred predominantly in sclerocytes before spicule formation.

Figure 3.7 Individual variations in spicule number and silicatein gene expression throughout A. queenslandica development. Spicule count and normalised relative expression (CNRQ) were obtained for 5 embryological stages (cleavage, brown, cloud, spot and ring), 3 larval stages (precompetent (0-2 hpe), reaching competency (6-8 hpe) and competent (24 hpe) larvae), and 3 postlarval stages (24, 48 and 72 hpi). For instance 0-2 hpe correspond to 0 to 2 h from the time larvae have been released from the adult sponge. A Rate of spicule formation during embryonic (dark blue), larval (light blue) and postlarval (green) stages. Each spot represents the spicule count in one individual (n=3/stage). B Comparing gene expression (CNRQ) amongst unrelated individuals of same age (n=5/stage, light grey: embryo, grey: larvae and dark grey: postlarvae) for the 3 genes of interest. References gene were used for the 11 development stages independently (Table 3.3).

3.5 Discussion

In this Chapter, spatial and temporal expression of silicatein genes, from embryonic to postlarval developmental stages, were characterised in order to correlate expression with spicule number. Six silicatein genes were identified in A. queenslandica genome, but three genes of interest, Aqu2.41046, Aqu2.41047 (both type II gene) and Aqu2.42494 (type I) were selected based on their expression level (Fig. 3.3) for further analyses. Indeed, all six silicatein genes were expressed throughout A. queenslandica development, but Aqu2.41046, Aqu2.41047 and Aqu2.42494 have higher transcript levels than Aqu2.08677, Aqu2.29399 and Aqu2.42495, making them better candidate genes to study expression and role of silicatein gene in this demosponge species.

The function of type I and II silicateins in A. queenslandica remains to be verified by functional analyses, however based on in situ hybridisation data I speculate that both types may perform a spicule forming function in A. queenslandica. Aqu2.42494 has approximately a four-fold increase in expression level compare to Aqu2.41046 and Aqu2.41047, suggesting that Aqu2.42494 is the main gene regulating spicule formation in A. queenslandica. Moreover, in the present study, WMISH analyses demonstrate that Aqu2.42494 expression is restricted to sclerocytes prior to the formation of mineralised spicules. Despite the differences in expression levels, the three genes displayed similar developmental expression profiles and pattern (although Aqu2.41046 and Aqu2.41047 embryonic and larval expression patterns are yet to be verified), suggesting that they may performed a similar function in spiculogenesis. Experiments on E. fluviatilis primmorph showed that different silicatein isoforms are used at different times during spicule formation, suggesting that each silicatein gene plays a different role in the growth and morphology of the spicule (Mohri et al. 2008). This study found that the expression of certain types of isoform ceased when spicule reach a size of 100 µm to 150 µm. A similar role might be seen among A. queenslandica silicatein genes, although currently RNA-Seq, qRT-PCR and WMISH data are consistent with these genes being activated before commencement of the mineralisation process and spicule growth.

Aqu2.42494 expression is restricted to sclerocytes throughout development, suggesting that this gene is involved in spicule formation from embryonic to post-larval developmental

stages (Fig. 3.6). Within the larval stage, sclerocytes are mainly localised at the posterior pole along the AP axis, region where larval spicules are distributed and which may allow larvae to orientate themselves (Leys 2003; Maldonado et al. 1997; Woollacott 1993).

If silicatein is the major enzyme responsible for the formation of spicules (Cha et al. 1999; Shimizu et al. 1998), then spicule number should correlate with silicatein gene expression. To trace the temporal expression of silicatein through sponge development, normalised transcript levels were generated and analysed. The data showed that Aqu2.41046, Aqu2.41047 and Aqu2.42494 are expressed before and during spicule formation (Fig. 3.3), but that their expression do not relate to spicule number across development (Fig. 3.4). This finding suggests that silicatein expression is not directly connected with number of spicules, although it should be recognised that these silicatein genes are expressed at high levels before the start of spicule formation. This delay seen in the early stages between silicatein expression and spicule number can be expected as spiculogenesis requires multiple steps before the initial silica deposition, including translation, phosphorylation and self-assembly (Cao et al. 2007). This will confound the correlation of expression with spicule number, in addition to the variability observed in both data throughout development. Supporting this postulation is the variable nature of gene expression during the early embryonic developmental stages, prior to spicule detection and during spicule formation as in embryos with a relatively consistent small number of spicules.

Spicules contain silicatein in the axial filament as well as on their surface layers (Cha et al. 1999; Müller et al. 2005a). Indeed, silicatein also controls spicule growth, which occurs in two different directions, an increase in length due to the elongation of the axial filament, and an increased in width by apposition of silica lamellae (Müller et al. 2007b; Schröder et al. 2006; Wang et al. 2010). Silicatein may therefore be used for growth rather than for the formation of new spicules when a certain stage is reached, such as the larval stage, resulting in an unrelated expression with numbers. However, this hypothesis seems contradictory to the data presented in this study as silicatein expression is at the lowest in the larval stages and does not appear to change significantly through the larval and early post-larval development (Fig. 3.3), as is the case with spicule number. This suggests that silicatein may not be contributing to spicule formation in larval stage and postlarval stages. Another plausible

explanation is that a small amount of silica, with an undetectable change in expression, might be sufficient for the demand of spicule growth, resulting in a small change in length and width of the spicule. Also, given that the sclerocytes expressing silicatein in larvae and juveniles are spindle-shaped and lack biomineralised spicule, it may be that there is a low level of spicule turnover during these stages that essentially results in no change in total spicule number. Thus, silicatein-expressing cells may be responsible for the fabrication of new spicules.

Several studies have been undertaken on sponge spicules in post-metamorphic (i.e. postlarval and juveniles) stages, focussing on the mechanisms behind spicule formation and the role of spicules in the sponge body plan. However, the role of spicules in embryos and larvae is largely unknown, although it has been proposed that they provide protection and confer other advantages during larval and post-larval development (Woollacott 1990; Woollacott 2003). Maldonado et al. (1997) was the first to provide a possible ecological and functional role of larval spicules, and hypothesise that these control larvae buoyancy. Sigmadocia caerulea larvae sank more rapidly with age, implying that larvae density might be affected by the increasing number of spicules as ciliary activity was halted for the experiment (Maldonado et al. 1997). Indeed, length of cilia has been reported to affect the swimming speed of larvae, as larvae possessing a posterior ring with long cilia tend to swim faster (Woollacott 1990; Berquist et al. 1970; Konstantinova 1966). From observation only, I noticed that A. queenslandica larvae sank more rapidly with time and that newly released larvae were unable to reach the bottom of the dish (see Degnan and Degnan 2010 for account of the distribution of larvae in a water column). However, no correlation was found between A. queenslandica larval age and spicule number, which remains constant with time; thereby spicule do not seem to affect larvae density/buoyancy. The result of this study is contradictory with Maldonado et al. (1997) findings, which showed that spicule increased with larval development in the demosponge Sigmadocia caerulea. The study measured the ash content of free swimming larvae, where ash was assumed to correspond to spicule mass, and found an increased in mass between 6 and 24 hour old larvae. Similarly, Bavestrello et al. (1993c) determine the daily rate of spicule production in the adult calcarean sponge Clathrina cerebrum, using a tetracycline marking method, and showed that newly formed spicules represent 10% of the total spicule number after 24 h, resulting in a slight increase of spicule with time. The roles and rates of production of spicules possibly differs between sponge

species and at this time it is unknown the function of spicules in A. queenslandica embryos, larvae, postlarvae and juveniles. It maybe that spicules in the larval stage increase in size rather than in numbers, this in turn may affect both spicule mass and larvae buoyancy. It also has to be noted that newly incompletely formed spicules were discarded in the count experiment of this study.

Variation of spicule number as well as silicatein expression within sponge developmental stages was recorded (Fig. 3.7). In Hymeniacidon perlevis, it was shown that silicatein expression changed depending on the sampling part of the individual, front, middle or back; each part affected by different micro-environment like water current flow capacity, higher flow resulted in higher expression (Cao et al. 2007). Larvae in this study were from a pool of adult sponges collected in different parts of the reef. Sponges were then placed in large aquarium and possibly affected by different water flow depending on their location within the tank. However, variation in expression cannot be attributed to micro-environment alone as sponge and larvae were reared in a common environment for the duration of the experiment. Differential expression may be influenced by the capacity of each sponge to absorb, accumulate and deposit silica. Genetic effect might be a possible explanation for this variation within same developmental stage individuals or environment adult sponges come from. These findings suggest that silicatein expression and spicule number might correlate within a single individual, and that difference may be observed between same age individuals of different maternal sources. Differences within a sponge might also be seen, but perhaps in a less significant way, as multiple brood chambers located in different parts of the sponge can be observed.

Chapter 4: General discussion

The emergence of sponges, one of the oldest extant metazoan phyletic lineages, was probably triggered by changes in the pre-Cambrian environment, and likely to have included changes in temperature, oxygen level and seawater chemistry (Bengtson et al. 1990; Seilacher 1994). The appearance of siliceous sponges corresponds to an increase in dissolve silicon in the aquatic environment due to weathering events between the Sturtian glaciations (710-680 Mya) and the Varanger-Maurinoan ice ages (605-585 Mya) (Hoffman and Schrag 2002). Therefore, it is possible that this influx in soluble silica allowed allow the sponge ancestors to exploit this new resources for novel adaptation such as siliceous spicules (Krasko et al. 2000; Müller et al. 2007a).

Sponge siliceous spicules wrere first described by Donati (1753) and subsequently it has been shown that spicule morphology is highly diverse and species-specific (Simpson 1984). Many studies have investigated spiculogenesis, from the role of silicatein and the different steps involved in spicule formation, to the possible ecological role of larval spicules and effect of environmental factors on spiculogenesis (Berquist and Sinclair 1973; Frøhlich and Barthel 1997; Maldonado et al. 1997; Fell 1976).

To understand the role of silicatein in spicule formation, and its evolution and expression, I undertook a detailed phylogenetic and gene expression analysis of silicatein genes in Amphimedon queenslandica. A. queenslandica remains the only sponge with a publically available genome and is a particularly good model to study developmental processes in demosponges (Leys and Degnan 2001; Srivastava et al. 2010). In this study, I employed bioinformatics, phylogenetic and molecular approaches to provide insight into 1) the presence of multiple silicatein genes in the A. queenslandica genome, some with unique characteristics that are shared between both cathepsin L and silicatein; and 2) how these genes are expressed through development; expression patterns and profiles were related to cell types and rates of spicule formation.

Despite spicules being an integral part of the demosponge body plan, little is known about their ontogenetic origin. As A. queenslandica embryos are easily accessible in brood chambers (Degnan et al. 2008a), I was able to show that spicule production begins in early gastrulation in A. queenslandica, similar to that of another demosponge Haliclona loosanoffi (Fell 1976). Unexpectedly, there is no significant difference in spicule number between different aged larvae and postlarvae, or between larvae and postlarvae of the same age. Moreover, spicule content tends to decrease during early metamorphosis, suggesting that larval spicules are lost during metamorphosis and that this loss does not affect postlarval development. After settlement occurred, ash content of Sigmadocia caerulae postlarvae was found to be lower than in free swimming larvae. Spicules could be seen around the newly settled postlarvae, which is consistent with their extrusion when the larval body flattens at the beginning of metamorphosis (Maldonado et al. 1997). A similar mechanism may be operating in A. queenslandica postlarvae.

4.1 Does silicatein expression correlate with spicule number?

Given the temporal accumulation of spicules during later embryogenesis and the apparent stabilisation of spicule number in larvae and postlarvae, I sought to determine how silicatein gene expression relates to spiculogenesis. First, I characterised the full repertoire of silicatein genes in A. queenslandica. Surprisingly, there are two types of silicatein β in A. queenslandica genome, type I gene (Aqu2.42494 and Aqu2.42495) which is characterised by the conventional serine in the catalytic triad, and type II gene (Aqu2.08677, Aqu2.29399, Aqu2.41046 and Aqu2.41047) which has a cysteine in the position that is characteristic of cathepsin L. Both types of silicatein - and sponge silicatein sequences in general - have a conserved Tyr following the catalytic Ser/Cys, suggesting that this residue is important for silicatein catalytic activity (Chapter 2).

All six silicatein genes are expressed throughout A. queenslandica development from embryonic development to juvenile and adult stages. Aqu2.08677 and Aqu2.29399 have similar expression levels and profiles which differed from the remaining four genes (Aqu2.41046, Aqu2.41047, Aqu2.42494 and Aqu2.42495). This is consistent with the

phylogenetic analyses that place these two genes on a separate branch and in their own subclade (Chapter 2). Aqu2.08677, Aqu2.29399 and Aqu2.42495 were discarded in the spatial and temporal experiments due to their low expression level. The genes of interest in this study (Chapter 3) are Aqu2.42494 type I gene, which is expressed higher than Aqu2.41046 and Aqu2.41047 type II genes. These type I and type II silicatein genes have similar expression profiles, which vary across A. queenslandica development with highest levels of expression in cleavage stage embryos prior to spicule formation, as well as spot embryos and newly settle postlarvae. In general, these genes appear to be expressed in sclerocytes cells prior to the fabrication of the mineralised spicule, although small immature spicules may have been present. This gene expression patterns can be related to that of silicatein isoforms in the freshwater sponge Ephydatia. fluviatilis reported in Mohri et al. (2008). However, it is worth noting that the multiple silicatein genes in E. fluviatilis are not orthologues of the A. queenslandica silicatein genes, since the silicatein gene family expanded independently in the lineages leading to these demosponges. Nonetheless, Ef silicatein-M1 and Ef silicatein-M2 are found to be active in nonspiculous cells as well as sclerocytes containing both short immature and long meglascleres, as in Aqu2.420494. On the other hand, Ef silicatein-M3 and Ef silicatein-M4 are only expressed in sclerocytes containing short immature spicules and in nonspiculous cells, as in Aqu2.41046 and Aqu2.41047. The nonspiculous cells containing silicatein in E. fluviatilis are archeocytes, while Aqu2.41047 seems to be expressed in pinacocyte-like cells. The function of silicatein in these non-spicule forming cells remain unknown, but may explain the reason why Aqu2.41046 and Aqu2.41047 expression does not correlate to spicule number in A. queenslandica.

4.2 Why does variation in silicatein expression and spicule number occur?

A. queenslandica postlarvae grown in the same environmental condition show variation in both silicatein expression and spicule number. Spicule growth is a rapid process, with an average rate of 5 µm/h (Imsiecke et al. 1995). Spicule width is found to be the most affected in embryonic and larval stages (Berquist and Sinclair 1973; Schönberg and Barthel 1997). Since, A. queenslandica mature spicules are on average 83 µm long and 2.3 µm wide in the adult sponge (Hooper & Van Soest 2006), it can be inferred that A. queenslandica spicule

reach maturity within 24 hour. The rapid growth rate of spicules should not have affected the counts data, as no increases in spicule number were recorded from 0 to 96 hour post- emergence (hpe) larvae and postlarvae. However it may have resulted in variation observed between individuals, as some may have been more efficient than others in fabricating spicules. This efficiency may be dependent of maternal sources.

The uptake rate of silica from the surrounding environment seems to be influenced by the health of the sponge (Frøhlich & Barthel 1997; Fell 1976). Other factors, such as reproductive stage and food scarcity also appears to influence sponge silica uptake rate (Frøhlich & Barthel 1997; Fell 1976). Since the A. queenslandica sponges used in this study were kept in aquaria for long period of time, the health of some sponge might have been deteriorating, affecting in turn the ability of their embryos and larvae to efficiently produce spicules. Finally, sponge efficiency in producing spicules may also be dependent on maternal sources as well as genetic variation in the regulatory sequences of the various silicatein genes likely underlining difference in the expression levels between individuals.

4.4 Conclusions and future directions

This analysis of spiculogenesis and silicatein expression through development is the most detailed to date and has yielded a number of unexpected findings. This study identified a new atypical silicatein genes (type II silicatein genes) expressed in sclerocytes cells; these type II silicatein genes are not restricted to A. queenslandica and can be found in other sponges. The role and function of these genes in spiculogenesis remain to be investigated, although it is clear that their expression in sclerocyte cells occurs before the formation of mineralised structures. This is the first study showing detailed spicule counts and silicatein expression through natural sponge development, which highlights the variability in both spicule number and expression patterns in individual of the same developmental stage and age. Previous ideas about the role spicules may play in controlling larval buoyancy may not apply to all sponge species, as spicule content did not increase during the larval and postlarval stages of A. queenslandica. From these results, it appears that role of spicules in sponge early

development, as well as possible correlation between expression and spicule number, might differ between sponge species.

Future studies need to consider possible genetic variation within a population, as a source of variation in spicule number and gene expression. Maternal effects may also contribute to this variation. It will also be interesting to consider the effect of environment, including seasonality and water chemistry, on A. queenslandica spicule formation and gene expression. Reproductive period can affect silica uptake rate and decrease spicule production (Frøhlich & Barthel 1997). A. queenslandica reproduces throughout the year but fecundity appears to be lower in winter, suggesting a higher rates of spicule formation and sponge growth during this period. Spicule morphology has been used as a taxonomic character, but environmental conditions may need to be taken into consideration when identifying sponge genera and species. Expression of a particular spicule types are dependent of the concentration of different elements including silica and iron in the surrounding water (Bavestrello et al. 1993b; Maldonado et al. 1999; Valisano et al. 2012). Additional to water temperature, different concentrations of silica, iron, selenium and germanium can influence spicule formation (Krasko et al. 2002; Le Pennec et al. 2003; Müller et al. 2005b; Simpson et al. 1985). Silica concentration in the surrounding water and water temperature can affect silicatein gene expression, thus affecting spicule production (Krasko et al. 2002; Schröder et al. 2006). Data obtained in this study, specifically for larvae and postlarvae, were based in the laboratory where water quality differs from the natural condition.

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Appendices

A1. GeneBank (NCBI) accession numbers of protein sequences of demosponges and hexactinellid (asterix) used in this study.

Species Silicatein GenBank accession Comments isoforms/cathepsin numbers Acanthodendrilla.sp Vietnam sil-a ACH92669.1 Partial sequence sil-b ACH92668.1 “ cath-L ACJ02498.1 Aphrocallistes vastus cath-L CAI91577.1 Aulosaccus sp. GV-2009 * Sil-like ACU86976.1 cath-L1 ACU86972.1 cath-L2 ACU86974.1 Baikalospongia fungiformis sil-a1 AEO36958.1 Partial sequence sil-a2 AEO36959.1 “ sil-a3 AEO36960.1 “ sil-a4 ADQ74580.1 “ Baikalospongia intermedia sil-a1 ACO51493.1 “ sil-a1’ CAQ54043.1 “ sil-a4 ACO51494.1 “ sil-a4’ CAQ54044.1 “ Bathydorus sp. GV-2009 cath-L1 ACU86973.1 “ cath-L2 ACU86975.1 “ Crateromorpha meyeri * sil CAP49202.2 “ cath-like1 CAP17584 “ cath-like2 CAP17585.1 “ cath-like3 CAP17586.1 “ cath-like4 CAP17587.1 “ cath-like5 CAP17588.1 “ Discodermia japonica sil CBY80151.1 “ Ephydatia fluviatilis sil CAJ44453.1 “ sil-2 CAJ44454.1 “ sil-a4 ACO51487.1 “ sil-G1 BAG74346.1 sil-G2 BAG74347.1 sil-M1 BAE54434.1 sil-M2 BAG74343.1 sil-M3 BAG74344.1 sil-M4 BAG74345.1 Ephydatia muelleri sil-a2 CAQ54046.1 sil-a3 CAQ54047.1 sil-a4 CAQ54048.1 Partial sequence Ephydatia sp. n.1 PW-2008 sil-a4 CAQ54050.1 “ Ephydatia sp. n.2 PW-2008 sil-a2 CAQ54049.1 “ Euplectella aspergillum * sil CBY80150.1 “ Geodia cydonium sil-a CAM57981.1 cath CAA71554.1 Halichondria okadai sil BAB86343.1 Partial sequence Hymeniacidon perlevis sil-a ABC94586.1 sil-1 ABM47424.1 Partial sequence

sil-2 ABM47425.2 “ cath-L ABM47423.1 “ Latrunculia oparinae sil-a1 ACG63793.1 sil-a1’ ACH47999.1 Partial sequence sil-a2 ACH48000.1 sil-a3 ACH48001.1 Partial sequence sil-b ACH48002.1 “ cath ACH48003.1 Lubomirskia baicalensis sil CAH10753.1 Partial sequence sil-a CAI43319.1 sil-a2 ADQ74585.1 Only 2 amino acid differ sil-a2’ CAI91571.1 between a2 and a2’ sil-a3 CAI91572.1 sil-a4 CAI91573.1 cath-L CAI43320.1 cath-L’ CAH10752.1 Partial sequence cath-L2 CAI91575.1 Lubomirskia incrustans sil-a1 ACO51492.1 Partial sequence sil-a4 ACO51491.1 “ Monorhaphis chuni sil CAZ04880.1 “ Petrocia ficiformis sil-b AAO23671.1 Pheromena raphanus cath-L1 ACU82389.1 cath-L2 ACU82390.1 Spongilla lacustris sil-a3 CAQ54051.1 sil-a4 ACO51489.1 Partial sequence sil-a4’ CAQ54052.1 Suberites domuncula sil-a CAI46305.1 sil-b CAI46304.1 Only 2 amino acids differ sil-b’ CAH04635.1 between sil-b and sil-b’ cath-L CAH04632.1 Swartschewskia papyracea sil-a4 CAQ54053.1 Partial sequence Tethya aurantium sil-a AAC23951.1 sil-b AAF21819.1 Tethya aurantium red variant sil CBY80148.1 Tethya aurantium yellow sil CBY80149.1 variant

A2. Protein sequences alignment of the conserved domain, Inhibitor I29 and Peptidase C1A, of Amphimedon queenslandica silicatein and cathepsin L genes. Conserved amino acid are highlighted in black (100% similarity) and similar amino acid (>60%) are highlighted in grey. Arrows indicate the catalytic amino acids, Cysteine (Cys/C) or Serine (Ser/S), Histidine (His/H) and Asparagine (Asn/N). Key conserved silicateins residues are underlined. The Cys residues involved in the formation of disulphide bonds are represented by dots and the serine cluster by a box.

A3. Protein sequences alignment of the conserved domain, Inhibitor I29 and Peptidase C1A, of silicatein-like sequences in different animals