Lessons from The

Home , ABHD18, Bacteriocin, Cassiopea andromeda, Cassiopea xamachana, Recoverin, Urea transporter 1

Pennsylvania State University The Graduate School Eberly College of Science

MICROBES IN LIFE HISTORY TRANSITIONS: LESSONS FROM THE

UPSIDE-DOWN JELLYFISH CASSIOPEA XAMACHANA

A Dissertation in Biology by Aki Ohdera

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2018

The Dissertation of Aki Ohdera was reviewed and approved* by the following:

Mónica Medina Associate Professor of Biology Dissertation Advisor

Todd C. LaJeunesse Associate Professor of Biology Chair or Committee

Timothy Jegla Associate Professor of Biology

Paul Medvedev Assistant Professor of Computer Science and Engineering Assistant Professor of Biochemistry and Molecular Biology

Stephen W. Schaeffer Professor of Biology Associate Department Head of Graduate Education

* Signatures are on file in the graduate school

! ii! Abstract

The ubiquity of symbiotic associations that exist in nature demonstrates the importance mutualism plays in the life history of virtually all metazoans. Even closely related species of hosts can associated with different species or genera of symbionts, with varying degrees of specificity. The co-evolutionary implications of these associations underline the genetic and molecular innovations that are a result of symbiosis. This is highlighted in the symbiosis between insects and Buchnera, as well as the bob-tail squid and Vibrio fischeri, where symbiont specific organs have evolved to facilitate the symbiosis. Genomic evolution can also be observed in bacteria as dramatic reductions in gene content, indicating evolution can act on both host and symbiont. Despite the importance and prevalence of symbiosis in nature, we still lack an understanding of how symbiosis is established and maintained for many of the associations. In particular, the mechanisms behind symbiosis-mediated developmental transitions that are found in some invertebrate symbiosis remain unsolved. Here, I utilize the Cassiopea xamachana model system to understand how symbionts can play a role in host life history transitions, and the mechanisms controlling these developmental events. I use next-generation sequencing techniques to determine how both host and symbiont genetics resulted in the evolution of larval and polyp metamorphosis. I also utilize the newly sequenced Cassiopea genome to explore genes that may have facilitated the evolution of symbiosis with Symbiodinium. The work presented in these chapters also showcase the effort put forth in developing the

Cassiopea system as a model to understand various fields of organismal biology, particular in cnidarian-Symbiodinium symbiosis.

! iii! Table of Contents List of Figures!...... !vi! List of Tables!...... !x! Acknowledgment!...... !xi! ! Chapter!1!8!Introduction:!Symbiosis!as!a!driver!of!evolutionary!novelty!in! organismal!life!history!...... !1! References!...... !5! ! Chapter 2: Upside-down but headed in the right direction: Review of the highly versatile Cassiopea xamachana system!...... !9! Abstract!...... !12! Introduction!...... !13! Evolution and Phylogenetics!...... !14! Life History!...... !16! Bud Morphogenesis!...... !17! Settlement and Metamorphosis!...... !18! Cassiopea-Symbiodinium Symbiosis!...... !20! Nutritional Requirements!...... !22! Behavior!...... !24! Fluid Dynamics!...... !24! Quiescence!...... !26! Cassiopea in the Environment!...... !27! Bioinvasion and Blooms!...... !27! Environmental Monitoring and Ecotoxicology!...... !28! Other Laboratory Applications!...... !30! Toxinology and cnidome!...... !30! Cassiopea Virology!...... !31! Cassiopea as a Laboratory Resource!...... !32! Conclusions!...... !34! References!...... !36! ! Chapter 3: Is larval settlement predictable? Genomic insights of settlement and metamorphosis inducing bacteria of Cassiopea xamachana!...... !70! Abstract!...... !71! Introduction!...... !72! Materials and Methods!...... !74! Larval Collection!...... !74! Bacterial Isolation!...... !75! Identification of bacterial Isolates!...... !75! Settlement Bioassays!...... !76! 16S microbiome extraction and sequencing!...... !77! Genome Sequencing and Analysis!...... !78! Results!...... !79! Bacterial Settlement Bioassay!...... !79! Settlement Substrate Microbiome!...... !79! Bacterial Genome Analysis!...... !80! Discussion!...... !82! References!...... !89!

! iv! Chapter 4: Modulation of gene expression driven by symbiosis: Strobilation mechanism in the upside-down jellyfish Cassiopea xamachana!...... !108! Abstract!...... !109! Introduction!...... !110! Materials and Methods!...... !112! Artificial Induction of Strobilation!...... !112! RNAseq of colonization and strobilation!...... !113! RNAseq Analysis!...... !114! In Situ Hybridization!...... !115! Results!...... !116! Artificial Induction of Strobilation!...... !116! Cassiopea Differential Gene Expression!...... !116! Symbiodinium Differential Gene Expression!...... !118! Discussion!...... !118! References!...... !123! ! Chapter 5: Box, stalked and upside-down? Draft genomes from diverse jellyfish (Cnidaria, Acraspeda) lineages: Alatina alata (Cubozoa), Calvadosia cruxmelitensis (Staurozoa), and Cassiopea xamachana (Scyphozoa)!...... !136! Abstract!...... !137! Introduction!...... !138! Materials and Methods!...... !142! Cassiopea xamachana Sample Collection and DNA extraction!...... !142! Calvadosia cruxmelitensis Sample Collectioin and DNA extraction!...... !142! Alatina alata Sample Collection and DNA extraction!...... !143! Cassiopea xamachana Sequencing and Assembly!...... !143! Calvadosia cruxmelitensis Sequencing and Assembly!...... !144! Alatina alata Sequencing and Assembly!...... !145! Gene Model Prediction!...... !146! Orthologous Gene Analysis!...... !146! Venom Analysis!...... !149! Hox-POU synteny analysis!...... !151! Conclusions!...... !152! Availability of supporting Data!...... !153! References!...... !154! Chapter 6: Conclusion!...... !178! References!...... !182! ! Appendix!A:!!Supplementary!material!for!" Is larval settlement predictable? Genomic insights of settlement and metamorphosis inducing bacteria of Cassiopea xamachana"!...... !188! ! Appendix!B:!Supplementary!material!for!"Modulation!of!gene!expression! driven!by!symbiosis:!Strobilation!mechanism!in!the!upside8down!jellyfish! Cassiopea(xamachana"!...... !196! ! Appendix!C:!Supplementary!Material!for!"Box,!stalked!and!upside8down?! Draft!genomes!from!diverse!jellyfish!(Cnidaria,!Acraspeda)!lineages:! Alatina(alata((Cubozoa),!Calvadosia(cruxmelitensis!(Staurozoa),!and! Cassiopea(xamachana!(Scyphozoa)"!...... !215!

! v!

List of Figures Figure!281.!A!schematic!summarizing!the!tools!and!approaches!available! for!the!Cassiopea(system,!highlighting!the!utility!of!each!life!stage! towards!a!multidisciplinary!approach!to!understanding!the!biology! and!physiology!of!this!early!metazoan.!...... !66! ! Figure!282.!A!working!hypothesis!of!Medusozoa!phylogeny!based!on!several! studies!(Bayha!et!al.,!2010;!Kayal!et!al.,!2013;!Kayal!et!al.,!2017),! including!the!monophyletic!Scyphozoa,!Rhizostomeae,!Kolpophorae,! and!Cassiopeidae.!Principal!morphological!and!life!cycle!characters! are!plotted!on!the!tree!(1!=!cnidae;!2!=!medusa!stage;!3!=!rhopalia;!4!=! strobilation!and!ephyra;!5!=!monodisc!strobilation).!Note:!The! topology!is!not!meant!to!represent!a!consensus.!...... !67! ! Figure!283.!A!Cassiopea(xamachana!poylp!with!a!planuloid!bud!developing! asexually!from!the!aboral!region!of!the!calyx.!...... !68! ! Figure!381.!Mangrove!leaf!sampling!locations!across!the!Florida!Keys! between!2012!and!2014.!Four!sites!were!chosen!for!collection!in!Key! Largo!(hexagon!=!Mangrove!Island,!triangle!=!dock,!square!=!the!cove,! circle!=!North!Bay)!and!a!single!site!on!Crawl!Key!(star!=!quarry).!....!99! ! Figure!382.!Larval!settlement!in!response!to!monoculture!biofilm!grown! from!bacterial!isolated!from!degrading!manrove!leaves.!C!=!0.2!μm! filtered!seawater!negative!control.!C2!=!marine!broth!negative! control.!C3!=!settlement!inducing!artificial!hexapeptide!(GPGGPA)! positive!control.!Statistical!significance!was!calculated!*!=!p8value!

! vi! MEGA!using!MUSCLE!and!phylogenetic!relationship!was!inferred!using! maximum!likelihood!(GTRi)!with!a!bootstrap!of!500.!...... !103! ! Figure!385.!Larval!settlement!frequency!in!response!to!culture!filtrate!from! bacterial!monoculture.!Filtrate!was!added!to!1!ml!of!0.2!μm!filtered! artifical!seawater.!100!μl!of!the!cell!fraction!was!added!to!1!ml!of! artificial!sea!water!and!allowed!to!settle!prior!to!the!start!of!the! experiment.!ASW!=!0.2!μm!filtered!artifical!seawater.!...... !104! ! Figure!386.!Taxonomic!identification!of!OTUs!determined!from!the!V4! region!of!the!16S!rRNA!gene!associated!with!degrading!mangrove! leaves!in!various!states!of!degradation.!OTUs!were!identified!to!the! genus!level!using!Qiime.!...... !105! ! Figure!387.!PCoA!plot!of!micriobial!biofilm!community!associated!with! degrading!mangrove!leaves!in!various!states!of!degradation.!Purple! triangle!=!quarry,!green!traingles!=!North!Bay,!blue!squares!=!the! dock,!red!triangles!=!the!cove,!orange!circles!=!mangrove!island.! Undegraded!leaves!are!outlined!and!encircled.!...... !106! ! Figure!481.!Strobilation!rates!of!Cassiopea(xamachana(polyps!(N=16)!in! response!to!artificial!1!μM!98cis!retinoic!acid,!50!nM!58methoxy828 methylindol,!!and!the!natural!strobilation!trigger!(Symbiodinium).! Control!=!0.2!μM!filtered!artificial!sea!water.!Statistical!significance! (***)!was!calculated!with!Tukey's!honest!significant!difference!post( hoc(test!from!the!ANOVA!(p!≤!0.0005)!...... !128! ! Figure!482.!Heat!map!of!the!top!100!differentially!expressed!genes!of! Cassiopea(xamachana(at!4!different!stages:!aposymbiotic!(0!=!light! blue),!3!and!8!days!post8colonization!(3d!=!light!green!and!8d!=! green),!and!strobila!(strob!=!dark!gold).!Count!data!was!r8log! transformed,!with!hierarchical!clustering!of!samples!labeled!based!on! stage.!Row!z8score!is!represented!with!the!cyan!line.!...... !129! ! Figure!483.!Gene!expression!of!genes!belonging!to!the!retinoic!acid!pathway! (retinoic(x(receptor,!retinol(dehydrogenase(18like,!retinol( dehydrogenase!28like,!CL112)!as!well!as!beta8carotene!9'10'8 dioxygenase!(BCO2).!!Expression!data!was!calculated!from!raw!read! counts,!with!normalization!and!Log2!expression!performed!with!the! DEseq2!R8package.!...... !130! ! Figure!484.!A)!Whole!mount!in!situ!hybridization!visualizing!the!expression! of!retinoic(x(receptor!(RxR),!CL112,!and!miniAcollagen(A!(McolAA;! positive!control)!for!aposymbiotic,!early,!and!late!strobila.!B)!Gene! expression!model!of!strobilation!related!genes!in!Cassiopea! xamachana.!...... !131!

! vii! ! Figure!485.!Venn!diagram!showing!shared!differentially!expressed!genes!at! 3!days!and!8!days!post8colonization!as!well!as!during!strobilation.! Number!of!differentially!expressed!genes!shared!across!the!different! stages!for!A)!Cassiopea(xamachana!and!B)!Symbiodinium( microadriaticum!...... !132! ! Figure!486.!ClusterProfiler!output!of!over8represented!KEGG!pathways!of! Cassiopea(xamachana.!Differentially!expressed!genes!with! annotations!were!mapped!to!KEGG!IDs!and!analyzed!using! ClusterProfiler!with!a!p8adjusted!cutoff!of!0.05.!Size!of!the!circles! indicates!precentage!of!genes!within!the!KEGG!pathway!found!to!be! differentially!expressed!at!3!days!and!8!days!post8colonization!and! during!strobilation.!Numbers!in!brackets!next!to!KEGG!descriptions! denote!the!number!of!genes!found!to!be!differentially!expressed! belonging!to!the!corresponding!pathway.!...... !133! ! Figure!487.!ClusterProfiler!output!of!over8represented!KEGG!pathways!of! Symbiodinium(microadriaticum.!Differentially!expressed!genes!with! annotations!were!mapped!to!KEGG!IDs!and!analyzed!using! ClusterProfiler!with!a!p8adjusted!cutoff!of!0.05.!Size!of!the!circles! indicates!precentage!of!genes!within!the!KEGG!pathway!found!to!be! differentially!expressed!at!3!days!post8colonization!and!during! strobilation.!Numbers!in!brackets!next!to!KEGG!descriptions!denote! the!number!of!genes!found!to!be!differentially!expressed!belonging!to! the!corresponding!pathway.!...... !134! ! Figure!488.!Strobilation!of!aposymbiotic!polyp!of!Cassiopea(xamachana! occurred!in!select!individuals!from!2012.!A)!Aposymbiotic!strobila!B)! 40x!magnification!bright!field!image!of!an!aposymbiotic!C.(xamachana! ephyra!C)!40x!magnficiation!bright!field!image!of!a!symbiotic!C.( xamachana!ephyra.!...... !135! ! Figure!581.!A)!Calvadosia(cruxmelitensis!(Staurozoa),!B)!Alatina(alata! (Cubozoa),!and!C)!Cassiopea(xamachana!(Sycphozoa).!D)!Phylogenetic! relationship!of!major!cnidarian!lineages!after!Kayal!et!al.!(2018),! linking!Cubozoa!and!Scyphozoa!as!sister!groups!and!united!with! Staurozoa!to!form!the!Acraspeda!lineage.!...... !168! ! Figure!582.!Gene!Content!Distribution!in!Cnidarian!Lineages.!Filled!circles! in!the!bottom!panel!indicate!shared!orthogroups!in!these!lineages.! Bar!graphs!indicate!the!number!of!orthogroups!corresponding!to!each! filled8circle!pattern.!Numbers!next!to!each!species!abbreviation! indicate!the!total!number!of!orthogroups!identified!for!that!species.! Hsap!=!Homo(sapiens;!Nvec!=!Nematostella(vectensis;!Hmag!=!Hydra( magnipapillata;!Ccrux!=!Calvadosia(cruxmelitensis;!Aala!=!Alatina( alata;!Cxam!=!Cassiopea(xamachana.!...... !169!

! viii! Figure!583.!Distribution!of!venom8related!genes!in!cnidarian!lineages.! Filled!circles!in!the!bottom!panel!indicate!presence!of!venom8related! gene!in!each!lineage.!Bar!graphs!indicate!the!number!of!venom8 related!orthogroups!corresponding!to!each!filled8circle!pattern.! Numbers!next!to!each!species!abbreviation!indicate!the!total!number! of!venom8related!orthogroups!identified!for!that!species.!Hsap!=! Homo(sapiens;!Nvec!=!Nematostella(vectensis;!Hmag!=!Hydra( magnipapillata;!Ccrux!=!Calvadosia(cruxmelitensis;!Aala!=!Alatina( alata;!Cxam!=!Cassiopea(xamachana.!...... !170! ! Figure!584.!Linkage!of!PPCS8Pou!Genes!with!Hox!Genes!in!Medusozoa! Genomes.!Genomic!scaffolds!from!three!Medusozoa!lineages!(C.( xamachana,(C.(cruxmelitnesis,(and(E.(dichotoma)!show!linkage!of!the! PPCS8POU!gene!linked!to!a!Hox!gene!(dark!green).!This!linkage!is!not! seen!in!Anthozoa!(N.(vectensis).!Scaffold!length!is!shown!to!the!right!of! the!bar.!The!light!green!region!indicates!the!transcribed!portion!of!the! scaffold,!and!exons!are!represented!within!by!curved!rectangles! (PPCS!exons!=!purple,!POU!exons!=!yellow).!Edic!=!Eleutheria( dichotoma;!Ccrux!=!Calvadosia(cruxmelitensis;!Cxam!=!Cassiopea( xamachana;!Nvec!=!Nematostella(vectensis.!...... !171! ! Figure!585.!Gene!ontology!biological!processes!over8enriched!within! Cnidaria!specific!orthogroups!visualized!using!REViGO.!Over8 representation!analysis!was!performed!with!ClusterProfiler,!with!a!p8 adjusted!cutoff!of!0.01.!Color!indicates!Log10!transformed!p8adjusted! value.!Terms!are!plotted!within!a!x8y!semantic!space,!in!which!similar! terms!are!clustered!within!closer!proximities.!Color!indicates!p8value! and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.!...... !172! ! Figure!586.!Gene!ontology!biological!processes!over8enriched!within! Medusozoa!specific!orthogroups!visualized!using!REViGO.!Over8 representation!analysis!was!performed!with!ClusterProfiler,!with!a!p8 adjusted!cutoff!of!0.01.!Color!indicates!Log10!transformed!p8adjusted! value.!Terms!are!plotted!within!a!x8y!semantic!space,!in!which!similar! terms!are!clustered!within!closer!proximities.!Color!indicates!p8value! and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.!...... !173! ! Figure!587.!Gene!ontology!biological!processes!over8enriched!within! Acraspeda!specific!orthogroups!visualized!using!REViGO.!Over8 representation!analysis!was!performed!with!ClusterProfiler,!with!a!p8 adjusted!cutoff!of!0.01.!Color!indicates!Log10!transformed!p8adjusted! value.!Terms!are!plotted!within!a!x8y!semantic!space,!in!which!similar! terms!are!clustered!within!closer!proximities.!Color!indicates!p8value! and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.!...... !174!

! ix!

List of Tables ! Table!281:!Morphological!data!on!valid!Cassiopea!species,!modified!from! Morandini!et(al.,!(2017)!...... !69! ! Table!381.!Presence!absence!of!genomic!features!between!five!bacterial! species!(Pseudoalteromonas!MB481,!Nisaea(MA684,!Thalassospira(MA682,! Pseudoatleromonas(luteoviolacea(HI1,!and!P.(luteoviolacea(6061).!...... !108! ! Table!581.!Statistics!of!the!genomic!assemblies!of!Alatina(alata,(Calvadosia( cruxmelitensis,!and!Cassiopea(xamachana.!...... !176! ! Table!582.!Venom8encoding!gene!repertoire!of!five!cnidarian!genomes! identified!with!the!venomix!database.!Venom!genes!are!categories!by! families!(column!1).!Both!genomic!and!transcriptomic!data!were!used,!with! transcriptomic!isoforms!counted!as!a!single!venom8encoding!gene..!...... !177! ! Table!583.!Orthrogroups!found!to!be!specific!to!cnidarians!that!are! symbiotic!with!Symbiodinium.!Gene!annotations!were!retrieved!from! Swissprot!with!representative!sequences!from!the!Cassiopea(genome.!...!178!

! ! ! ! ! ! ! ! ! ! ! ! ! !

! x! Acknowledgment!

First and foremost, I would like to thank my doctoral dissertation advisor,

Mónica Medina for providing me with the opportunity and mentorship to do the research presented in this dissertation. She allowed me to explore my own interests and ideas with full support, always looking for new avenues to further my education and pushing me to go beyond the boundaries in science and in life. She was instrumental in surrounding me with an amazing group of people, which I would not have been able to complete my PhD without. This includes everyone that was and currently are members of the Medina Lab. I would like to especially thank Bishoy

Kamel for his knowledge and patients in teaching me the skills necessary for much of the bioinformatics presented throughout the chapters, in addition to Erika Diaz-

Almeyda for her assistance in development of the Cassiopea culture. In addition to my lab mates, I would like to thank the group of undergraduate students that assisted me in maintaining the Cassiopea culture and assisting me with experiments. Their commitment to the animals and their friendship has been invaluable and without their help, I would not have been able to accomplish as much as I did.

I addition to the members of the Medina Lab, I would like to thank my committee members Todd LaJeunesse, Tim Jegla, and Paul Medvedev for their expert advice and willingness to provide assistance when needed. Todd LaJeunesse in particular was extremely helpful in providing insight on Symbiodinium, in addition to aspects outside of research. Tim Jegla provided valuable assistance in projects presented here, as well as taking his time to work on side projects that will surely inform future projects. Paul Medvedev provided an outside perspective to my research and project, as well as providing advice on genome assemblies. In addition to members of my committee, I received considerable assistance from those at other

! xi! institutions. I would not have been able to accomplish most of the work without the advice and support provided by William Fitt of the University of Georgia. Not only is he an expert in Cassiopea, he went above and beyond in assisting me with my field research and providing mentorship. Dieter Hofmann, an early researcher of Cassiopea also provided immense knowledge related to the system.

Finally, I'd like to thank my friends and family for their support and friendship for the past 6 years. I would like to thank my parents Ellen and Yasuo for shaping the path for me to begin and complete graduate school, and to my brothers Nao and Kei for providing a respite from my research. I'd like to thank Sheila Kitchen for her scientific advice, friendship, and support in life, particularly towards the final months of PhD. I'd like to thank my roommates and close friends in State College, Isaac,

Arin, Jo, Herschel, and Scott for ensuring that I stay relaxed and live life beyond the laboratory. I'd also like to thank my close friends and former co-members of EGO for their support and affection. I could not have done this without everyone listed above, and I hope this expresses a fraction of the gratitude I have towards everyone.

! xii! Chapter 1

Introduction:!Symbiosis!as!a!driver!of!evolutionary!novelty!in! organismal!life!history!

There is currently no debate in the prevalence of mutualistic symbiosis in metazoans. However, the proposition of symbiosis playing a key role in organismal evolution initially received significant criticism from the scientific community.

Symbiosis as a driver of evolutionary novelty was suggested during the 19th century, notably by Constantin Merezhkowsky, Andrey Famintsyn, and Boris Kozo-

Polyanksy, Paul Portier, and Ivan Wallin (Sapp, 1994;2003). They were early proponents of the endosymbiotic theory of organelles, proposing the bacterial origin of organelles such as mitochondria, plastids, and chloroplasts. This theory did not gain significant attention until its re-introduction by Lynn Margulis in 1967 (Sagan,

1967). The prokaryotic origin of eukaryotic mitochondria became widely accepted with evidence provided from genomic sequencing (Lang et al., 1997;Gray et al.,

1999;Gray et al., 2001;Sapp, 2003).

Since, our understanding of how symbiosis plays a major role in metazoan life history continues to increase, and research in a diverse group of animal systems have provided us with new knowledge and even more questions regarding the importance of symbiosis and how these associations shape both host and symbiont physiology, development, behavior, and genomic architecture.

Humans and other mammals harbor bacterial symbionts within their gut, with humans hosting as many as 100 trillion microbial cells and (Whitman et al., 1998).

These bacteria supplement the metabolic capabilities of the host, thereby expanding the bioavailability of nutrients (Hooper et al., 2002;Chow et al., 2010). Nutritional complimentary is observed across many invertebrate phyla, including the association

! 1! between the pea aphid and the endosymbiotic bacteria Buchnera. Buchnera supplements the low nutritional sap diet of the insect by providing the host with essential amino acids (Sasaki and Ishikawa, 1995;Douglas, 2009). Reef building corals are symbiotic with a photosynthetic dinoflagellate Symbiodinium, which provide the host with photosynthetic products in the form of glucose and glycerol, in addition to amino acids and lipids in exchange for inorganic nitrogen and phosphate, and carbon (Venn et al., 2008;Burriesci et al., 2012;Davy et al., 2012). The reliance of metazoans on microbes as a source of nutrients likely released the hosts' requirements of synthesizing metabolically expensive molecules, and could have allowed some taxa to enter previously unavailable niches due to nutrient limitations (Windoffer and

Giere, 1997;Lopez-Garcia et al., 2002;Douglas, 2014).

While nutritional symbiosis is a hallmark of mutualism, endosymbiosis can also provide the host with non-nutritional benefits. In the case of the bobtail squid,

Euprymna scolopes and its endosymbiont Vibrio fischeri, the bioluminescent bacteria provide countershading for the host, allowing the squid to escape predation. What is striking about this symbiosis is the highly coordinated process by which a single V. fischeri cell colonizes the symbiont housing light organ, which initiates a developmental cascade, allowing the organ to mature (McFall-Ngai, 2002;Kremer et al., 2013;McFall-Ngai, 2014). Symbiont induced development is also observed in the deep-sea hydrothermal vent dwelling tubeworm Riftia and the morphogenesis of the bacteria-housing trophosome (Nussbaumer et al., 2006). Symbiont induced development can also be observed within the mammalian gut, with germ-free rats exhibiting differences in microvilli morphology compared to normal rats (Falk et al.,

1998). These highly coordinated developmental events demonstrate the co- evolutionary outcome of host-microbe symbiosis and the resulting evolutionary

! 2! innovations. Despite recognition of the importance of microbial symbionts in organismal life history, we have yet to understand the molecular and genetic mechanisms that control these developmental transitions. Determining the underpinnings of the association will allow us to better understand the implications of symbiosis in metazoan evolution. With the increasing availability of genomic and molecular tools such as next generation sequencing and the CRISPR-cas9 method, we are better equipped to investigate how these associations are established and maintained, and the mechanism behind the coordinated signaling to drive developmental changes.

The upside-down jellyfish Cassiopea xamachana exhibits symbiotic associations at various points in its lifecycle. The larva to sessile polyp

(scyphistomae) transition, characterized by settlement and subsequent metamorphosis, is triggered by biofilm-associated bacteria. Another association is observed during the polyp stage, the jellyfish acquires their endosymbiont, the dinoflagellate algae

Symbiodinium. While the mutualistic association is nutritionally beneficial for both members of the symbiosis, the host relies on the symbiont in order to transition from the juvenile polyp stage to the adult medusae stage (Colley and Trench,

1985;Hofmann et al., 1996;Ohdera et al., 2018). Unlike other symbiosis-induced development, the upside-down jellyfish is unique in that Symbiodinium triggers metamorphosis of the host, rather than the formation of a symbiosis-specific structure.

The establishment of symbiosis is also marked by the differentiation of the symbiont phagocytosing host cell, which is also a unique feature of the symbiosis (Colley and

Trench, 1985). With the use of next-generation sequencing, I explore the genomic and transcriptomic underpinnings involved in driving the developmental transitions within the Cassiopea life history. Moreover, given their phylogenetic relationship to other

! 3! Symbiodinium-symbiosis forming organisms, I utilize the newly sequenced C. xamachana genome to investigate the presence of novel genes that may have allowed the symbiosis to evolve within the cnidarian lineage, in addition to other genomic features unique to cnidarian lineages. These findings will provide further insight into how symbiosis can drive organismal evolution and not only demonstrate the efficacy of the Cassiopea-Symbiodinium system to explore various aspects organismal biology.

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! 4! References! ! Burriesci,!M.S.,!Raab,!T.K.,!and!Pringle,!J.R.!(2012).!Evidence!that!glucose!is!the!

major!transferred!metabolite!in!dinoflagellateMcnidarian!symbiosis.!

Journal)of)Experimental)Biology!215,!3467M3477.!

Chow,!J.,!Lee,!S.M.,!Shen,!Y.,!Khosravi,!A.,!and!Mazmanian,!S.K.!(2010).!HostM

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! 8! Chapter 2

Upside-down but headed in the right direction: Review of the highly versatile Cassiopea xamachana system ! Aki H. Ohdera1, Michael J. Abrams2, Cheryl Lewis Ames3, David M. Baker4, Luis

Parmenio Suescún-Bolívar5, Allen Collins3,6, Christopher J. Freeman7, Edgar

Gamero-Mora8, Tamar L. Goulet9, Dietrich K. Hofmann10, Adrian Jaimes-Becerra8,

Paul F. Long11, Antonio Carlos Marques8,12, Laura Miller13, Laura D. Mydlarz14,

Andre C. Morandini8,12, Casandra Newkirk15, Sastia Putri16, Julia Samson13, Sérgio N.

Stampar17, Bailey Steinworth15, Michelle Templeman18, Patricia E. Thomé5, Marli

Vlok19, Cheryl M. Woodley20, Jane C. Y. Wong4, Mark Q. Martindale15, William

Fitt21, Mónica Medina1,22

1. Department of Biology, Pennsylvania State University, University Park, PA,

USA

2. Division of Biology and Biological Engineering, California Institute of

Technology, Pasadena, CA, USA

3. Department of Invertebrate Zoology, National Museum of Natural History,

Smithsonian Institution, Washington D.C., USA

4. The Swire Institute of Marine Science, School of Biological Sciences, The

University of Hong Kong, Hong Kong, PR China

5. Unidad Académica de Sistemas Arrecifales Puerto Morelos, Instituto de

Ciencias del Mar y Limnología, Universidad Nacional Autónoma de México,

Quintana Roo, México

6. National Systematics Laboratory of NOAA's Fisheries Service, National

Museum of Natural History, Smithsonian Institution, Washington D.C., USA

7. Smithsonian Marine Station, Fort Pierce, FL, USA

! 9! 8. Departamento de Zoologia, Instituto de Biociências, Universidade de São

Paulo, São Paulo, Brazil

9. Department of Biology, University of Mississippi, University, MS, USA

10. Department of Zoology & Neurobiology, Ruhr-University Bochum, Bochum

Germany

11. School of Cancer and Pharmaceutical Sciences, King's College, London, UK

12. Centro de Biologia Marinha, Universidade de São Paulo, São Sebastião,

Brazil

13. Biology Department, University of North Carolina at Chapel Hill, Chapel

Hill, NC, USA

14. Department of Biology, University of Texas at Arlington, Arlington, TX, USA

15. Whitney Laboratory for Marine Bioscience, University of Florida, St.

Augustine, FL, USA

16. Department of Biotechnology, Graduate School of Engineering, Osaka

University, Osaka, Japan

17. Departamento de Ciências Biológicas, Faculdade de Ciências e Letras de

Assis, Unesp Universidade Estadual Paulista, Assis, Brazil

18. TropWATER and College of Marine & Environmental Sciences, James Cook

University, Townsville, QLD, Australia

19. Department of Botany, University of British Columbia Vancouver,

Vancouver, BC, Canada

20. U.S. National Oceanic & Atmospheric Administration, National Ocean

Service, National Centers for Coastal Ocean Science, Charleston, SC, USA

21. Odum School of Ecology, University of Georgia, Athens, GA, USA

! 10! 22. Smithsonian Tropical Research Institute, Smithsonian Institution, Panama

City PL, Washington D.C. USA

! ! ! ! !

! 11! Abstract ! The upside-down jellyfish Cassiopea xamachana (Scyphozoa: Rhizostomeae) has been predominantly studied to understand its interaction with the endosymbiotic dinoflagellate algae Symbiodinium. As an easily culturable and tractable cnidarian model, it is an attractive alternative to stony corals to understanding the mechanisms driving establishment and maintenance of symbiosis. Cassiopea is also unique in requiring the symbiont in order to complete its transition to the adult stage, thereby providing an excellent model to understand symbiosis-driven development and evolution. Recently, the Cassiopea research system has gained interest beyond symbiosis in fields related to embryology, climate ecology, behavior, and more. With these developments, resources including genomes, transcriptomes, and laboratory protocols are steadily increasing. This review provides an overview of the broad range of interdisciplinary research that has utilized the Cassiopea model and highlights the advantages of using the model for future research.

! 12! Introduction ! The upside-down jellyfish Cassiopea is a benthic scyphozoan (Rhizostomeae) commonly found in tropical and sub-tropical shallow coastal ecosystems, such as mangroves and seagrass beds. Cassiopea spp. are unique among scyphomedusae in that their characteristic flat exumbrella rests on the sea bottom, while their convex subumbrella and oral arms face upwards. High light penetrance is important for species within this genus, as the jellyfish hosts one or more photosynthetic dinoflagellate species of the genus Symbiodinium (Hofmann et al., 1996; Lampert

2016). Similar to their coral relatives, nutrient exchange is a key component supporting this cnidarian-dinoflagellate mutualism (Hofmann and Kremer, 1981;

Welsh et al., 2009; Freeman et al., 2016). In addition, Cassiopea spp. rely on the symbionts as a developmental trigger (Hofmann et al., 1978; Colley and Trench,

1985). A fascination with these requisite traits of the upside-down jellyfish has fueled studies on the Cassiopea-Symbiodinium interaction since the 1980s (see recent review by Lampert 2016). While its efficacy as a system for symbiosis research is well known in the literature, Cassiopea is gaining momentum as a model species in other areas. In mathematics, the symmetric morphology and relatively simple neuromuscular system of the upside-down jellyfish make it an ideal system from which to develop computational fluid dynamic and neuromechanical models (Passano

2004; Hamlet et al., 2011; Santhanakrishnan et al., 2012). Additionally, Cassiopea has been presented as a possible model organism for the study of behavioral biology as highlighted by recent developments with sleep behavior (Nath et al., 2017) (Figure

1). Likewise, this jellyfish genus has been gaining traction as a biomonitor / bioindicator species, with applications for coastal ecosystem management

(Templeman and Kingsford, 2012; Epstein et al., 2016; Klein et al., 2016a; Klein et

! 13! al., 2017). The relative simplicity of culturing Cassiopea polyps and medusae makes this genus a highly amenable laboratory system. With newly established Cassiopea xamachana clonal laboratory lines, and recent advances in affordable genomics tools giving way to draft transcriptome and genome assemblies, the upside-down jellyfish is an appropriate candidate as a model organism. Here, we review Cassiopea research conducted to date and discuss future directions of this highly valuable study system.

Evolution and Phylogenetics

The first species of the genus Cassiopea was described from the Caribbean as

Medusa frondosa by Pallas (1774), and just one year later a second species from the

Red Sea was described as Medusa andromeda (published as a post mortem work; figures published only in Forskål (1776). The generic name Cassiopea was proposed for all jellyfish with foliaceous appendages in 1810 (Péron and Lesueur, 1810).

Members of the genus Cassiopea were treated as a family by Tilesius (1831), and the current spelling appeared ~30 years later (Agassiz, 1862). The species-level relationships of the genus Cassiopea are still unresolved since most of the taxonomic studies are based on overlapping, plastic, and polymorphic features (Table 1) (Mayer,

1910; Kramp, 1961). This is confounded by vague species descriptions with species diagnoses that are descriptive rather than comparative, although morphology has been useful in distinguishing some congeneric species, i.e. Cassiopea frondosa. In total, 24 nominal species, and varieties thereof, have been proposed to date (Table 1), but only

10 species are presently considered to be valid based on molecular data (Holland et al.,

2004; Arai et al., 2017; Morandini et al., 2017). These studies show that populations of C. xamachana are potential introduction of Cassiopea andromeda from the Red

Sea, thereby grouping the species into a single lineage. These species designations,

! 14! which are based on kimura 2-parameter pairwise distance, fall within previous estimations for other scyphozoans, ranging from 0.00-0.034 for conspecifics, and

0.102 - 0.234 for congenerics (Holland et al., 2004; Ortman et al., 2010; Gómez

Daglio and Dawson, 2017). Analyses with expanded taxon sampling (Bayha et al.,

2010; Kayal et al., 2013; Kayal et al., 2017) have placed the monogeneric Cassiopea within the family Cassiopeidae, in an unstable position within the clade Kolpophorae, along with representatives of Mastigiidae, Thysanostomatidae, Versurigidae and

Cepheidae (Figure 2). The Kolpophorae belongs to the order Rhizostomeae, a well- supported clade derived from within the scyphozoan order Semaeostomeae (Bayha et al., 2010; Kayal et al., 2013).

While the polyp stage and strobilation of Cassiopea xamachana, as well as the polyp stage of C. frondosa, were described by Bigelow (Bigelow, 1892, 1893), a complete description of the life cycle of C. xamachana did not appear in the literature until almost a decade later (Bigelow, 1900). Additional details of the life cycle were later published on a population of C. andromeda from Al-Ghardaqa (Red Sea) (Gohar and Eisawy, 1960). Although the morphology of Cassiopea is unique and known only from extant forms, a fossil dated to the Solnhofen formation lagerstätte (Jurassic,

Myogramma speciosum) has a similar arrangement of what appears to be the muscular system (Maas, 1902).

Moving forward, enhanced taxon sampling and character analyses by multi- omics methods on a broad geographical scale will facilitate rigorous testing of phylogenetic hypotheses and assessment of genetic richness related to cryptic species, thereby allowing research on different Cassiopea systems from a comparative evolutionary perspective.

! 15! Life History ! Cassiopea spp. are gonochoristic brooders, with fixed sex determination, although a case of hermaphroditism has been reported from Hawai'i (Hofmann and

Hadfield, 2002). Fertilization has been reported to occur within the gastrovascular cavity of the female (Smith, 1936). The embryos pass from the gastrovascular cavity through the brachial canal out of the secondary mouths, and are subsequently deposited onto the oral disc. Because motile settlement-competent planulae can be found attached to the female’s oral disc at any given point, the duration of attachment

(i.e., brooding) in the absence of outside disturbance is unknown. Cleavage begins approximately one to two hours after the fertilized eggs are visible on the oral disc, and then round, ciliated embryos are present 48 hours post-fertilization. By 96 hours post-fertilization, elongate ovoid planulae, rotating around the longitudinal axis, swim with clear directionality and are capable of settling and becoming polyps. While a detailed characterization of early embryonic development in Cassiopea is yet to be done, developmental timing appears to be temperature dependent and planula morphology has been well described (Martin and Chia, 1982).

Hox genes are present in C. xamachana (Kuhn, et al., 1999; Ferrier and

Holland, 2001), and work on other cnidarians suggests these may be involved in patterning the oral-aboral axis. In both the anthozoan Nematostella vectensis and the hydrozoan Clytia hemisphaerica, planula and polyp stages show differential Hox expression along the oral-aboral axis (Ryan et al., 2007; Chiori et al., 2009).

Patterning of the medusa body form remains a more complicated question, as expression of some Hox genes were localized to sensory structures and the bell

(Chiori, 2009), while others were expressed specifically in oral endoderm. In addition to Hox expression during development, homeobox genes are active during cnidarian

! 16! regeneration (Schummer et al., 1992) and Cassiopea has gained some attention as a model for regeneration (Cary, 1916; Polteva, 1985; Curtis and Cowden, 1974). Of particular note, all Cassiopea life stages show telomerase activity, with potential elongation of the telomere during the medusa stage (Ojimi et al., 2009; Ojimi and

Hidaka, 2010), paving the way for additional research in the genetic mechanisms of regeneration and longevity in non-bilaterian organisms. Future research in Cassiopea development incorporating genomics, transcriptomics, metabolomics, and in situ hybridization will allow further characterization of pre-swimming development, particularly timing and stereotype of gastrulation, and precise identification of morphological characteristics denoting settlement competency. The multi-omics approach will lay the foundations for describing the developmental genes regulating the life history of these scyphozoans and generate testable hypotheses towards understanding the evolution of cnidarian life cycles and bauplans.

Bud Morphogenesis ! Polyps of the genus Cassiopea are capable of producing planuloid buds 3-4 mm in length via evagination from the aboral region of the calyx (Van Leishout and

Martin, 1992) (Figure 3). Bud formation and detachment can occur at a relatively rapid rate, with buds released as frequently as 2-3 per day. Progenitor ectodermal cells are recruited distally and asymmetrically from the site of bud morphogenesis rather than produced by a mitotically active population of cells (Hofmann and Honegger,

1990; Hofmann and Gottlieb, 1991). Planuloid buds form with their future oral pole most proximal to the body. The most distal part of the bud is the future aboral pole, which after detachment will be the leading pole of the swimming bud and the stalk of the future polyp (Curtis and Cowden, 1971). Within 24 hours, settled buds will

! 17! develop a hypostome and up to 16 tentacles, thereby providing the newly developed polyps with the capacity to capture prey. Head (calyx and tentacles) and stalk formation is contingent on settlement to a substrate, with a head-morphogenesis inhibiting signal potentially generated from the aboral pole of the swimming bud

(Neumann, 1980). It has been reported that oral halves of transversely bisected buds properly developed the hypostome and tentacles but lacked a stalk, while the aboral halves developed into full scyphistomae (Curtis and Cowden, 1971; Neumann, 1980).

Further, treatment of the oral fragment with bud homogenate as well as PKC inhibitors (psychosine, chelerythrine) showed inhibitory effects to head development

(Thieme and Hofmann, 2003a, 2003b), suggesting the important involvement of PKC in bud and larval metamorphosis (Fleck and Bischoff, 1992). Conversely, treatments with canathadrin, a serine/threonine protein phosphatase inhibitor, induced development of the head and stalk in scyphistomae, while methionine, homocysteine, trigonelline, nicotinic acid, and cycloleucine treatments all inhibited morphogenesis

(Kehls et al., 1999).

Settlement and Metamorphosis ! Induction of settlement and metamorphosis in cnidarian species, including

Cassiopea spp., is contingent on the presence of certain types of bacteria (Hofmann et al., 1978; Neumann 1979; Leitz 1997; Vermeij et al., 2009). Both larvae and planuloid buds of Cassiopea spp. settle and metamorphose in response to bacterial biofilm, including Vibrio sp. isolated from an Artemia culture (Hofmann et al., 1978).

While the isolated Vibrio sp. showed toxic effects on polyp development, a metabolite

(1 to 10 kDa) that was isolated from the bacteria induced settlement and metamorphosis of Cassiopea planulae (Neumann, 1979). A cholera toxin isolated

! 18! from Vibrio cholerae was also inductive (Wolk et al., 1985). Collagen digestion by

Vibrio alginolyticus also led to the production of inductive fractions, suggesting the involvement of a peptidic cue in polyp development (Hofmann and Brand 1987). The discovery that certain small molecules and peptides are involved in polyp metamorphosis has led to the design of a synthetic peptide (GPGGPA) with settlement and metamorphosis inductive capacities (Fleck, 1998).

Research efforts attempting to isolate a peptidic cue from Cassiopea’s natural substrate, degraded mangrove leaves, resulted in extraction of several bioactive fractions, one of which contained a proline-rich 5.8 kDa peptide capable of inducing settlement and metamorphosis, although the precise amino acid sequence of this peptide or other natural cues have yet to be determined (Fleck and Fitt, 1999; Fleck et al. 1999). While cnidarian species have been shown to settle and metamorphose in response to mono-culture biofilm of multiple bacterial taxa (Tran and Hadfield, 2011), the exact underlying mechanism allowing planulae to react to multiple microbial cues has yet to be fully explored. Variation in induction response to artificial inducers

(inorganic ions, phorbolesters, diacylglycerols, tetradecanoyl-phorbol-13 acetate) has been observed between hydrozoan and scyphozoan species (Müller, 1973; Henning et al.,, 1996; Siefker et al., 2000). The difference in induction response to artificial inducers noted between the moon jellyfish Aurelia aurita and C. xamachana (Fitt et al., 1987; Siefker et al., 2000), for example, are suggestive of potentially differing mechanisms of settlement induction. The evolution of cue specificity and signal transduction mechanisms of cnidarian larvae, and more generally invertebrate larvae, remains a topic requiring further research. The current understanding of Cassiopea species diversity and geographic distribution, coupled with the availability of several

! 19! lab cultures, offers avenues to conduct comparative studies on the induction response of planulae to both natural and synthetic settlement inducers.

Cassiopea-Symbiodinium Symbiosis ! In many scyphozoans, the transition from polyp to medusa, known as strobilation, can be triggered by temperature and other exogenous cues (Spangenberg

1965; Loeb 1973; Holst 2012). Strikingly, strobilation in Cassiopea requires colonization by the endosymbiont Symbiodinium spp. (Gohar and Eisawy, 1960;

Hofmann et al., 1996). Without colonization, polyps will not undergo strobilation, although a single strobilation event in aposymbiotic polyps has been reported by

Rahat and Adar (1980). Cassiopea spp. are monodisc strobilating scyphozoans, producing a single ephyra at each strobilation event. Strobilation generally occurs over several days, with the first few days characterized by a reduction in tentacle length (due to absorption), followed by the development of a disc-like apical structure bearing rhopalia and a medusoid form; shortly following is constriction by the polyp body resulting in release of the ephyra (Ludwig, 1969; Hofmann et al., 1978).

Colonization of the polyp by the symbiont (i.e., dinoflagellate), and subsequent strobilation of an ephyra, happens in several discrete stages. The colonization process begins when symbionts enter the polyp through the mouth and are then phagocytosed by the gastroderm (Colley and Trench, 1983; Fitt and Trench, 1983). Successful colonization of cnidarian cells requires suppression of the host immune response, thereby revealing a link between symbiosis and innate immunity (Miller et al., 2007;

Kvennefors et al., 2008). Along with other cnidarians, jellyfish show evidence of producing antimicrobial peptides that indicate their ability to identify between self and non-self (Rinkevich, 2004; Ovchinnikova et al., 2006; Bosch, 2008). This ability

! 20! would be useful in regulating the host microbial composition as well as symbiont colonization and proliferation, a topic that has garnered significant attention and will benefit from the development of the Cassiopea-Symbiodinium research system

(Detournay et al., 2012; Neubauer et al., 2016; Neubauer et al., 2017).

Events leading to successful colonization post-phagocytosis can be identified by the relative position of the algae within the phagocytosing cell (Colley and Trench,

1985). Initially, symbionts can be found towards the apical end of phagocytic cells and the basal end by day 3 post-entry. Symbiodinium cells that are unable to migrate basally are destroyed by lysosomal fusion. By day 8, host cells containing successful symbionts detach and migrate into the mesoglea where proliferation of symbiont- containing host-cells subsequently occurs (Colley and Trench, 1985). Strobilation takes place approximately 3 weeks later, depending on several factors including the symbiont species, temperature and food intake (Fitt and Costley, 1998). While the precise cue(s) responsible for initiation of strobilation is unknown, inhibition of photosynthesis with 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) results in continuation of the process, albeit at lower rates (Hofmann and Kremer, 1981). In a non-symbiotic scyphozoan, Aurelia aurita, initiation of strobilation was found to involve genes from the retinoic acid pathway (Fuchs et al., 2014; Brekhman et al.,

2015). In contrast to A. aurita, treatment of Cassiopea polyps with retinoic acid does not result in strobilation (Silverstone et al., 1977; Pierce 2005; Cabrales-Arellano et al., 2017). Conversely, indomethacin, a drug with structural resemblance to a peptide involved in strobilation in Aurelia induces metamorphosis of C. xamachana polyps to the mature medusa form (Kuniyoshi et al., 2012; Cabrales-Arellano et al., 2017). This observation suggests that while some aspects of the strobilation mechanism might be shared across Scyphozoa (Helm and Dunn, 2017; Yamamori et al., 2017), the

! 21! evolution of symbiosis within members of the Rhizostomae may have resulted in co- option of other pathways as a developmental trigger. With the availability of transcriptome and genome level sequence data awaiting extensive query, Cassiopea holds promise as an exciting system for the study of symbiosis-mediated development, as well the mechanisms involved in host colonization by its symbionts.

Nutritional Requirements ! For symbiotic cnidarians, the essential nature of the symbiotic relationship strongly suggests that heterotrophic feeding from the surrounding waters may not provide nutrients sufficient to cover the carbohydrate and lipids needed for medusa metamorphosis, growth, and sexual reproduction. A major gap in our knowledge of algal-cnidarian symbiosis has been the understanding of the mechanisms underlying the biosynthesis and regulation of carbon flow from symbiont to host (Yellowlees et al., 2008; Davy et al., 2012), and the uptake of such carbon compounds by the host partner. In this context, identifying the chemical form of photosynthates or so-called

“mobile compounds” (Trench, 1971; Venn et al., 2008) has proven challenging, although it most likely involves complex metabolic modifications. In other well- known examples of algal-cnidarian symbiotic relationships in scleractinian corals, glycerol is the dominant photosynthate produced (Gattuso et al., 1993; Hagedorn et al.,

2010), while glucose is the dominant photosynthate in the sea anemone genus

Exaiptasia (formerly Aiptasia) (Burriesci et al., 2012; Molina et al., 2017). Early 14C

(radiocarbon) labeling studies in Cassiopea andromeda showed that glucose and glycerol were the only two free metabolites detected after short term labeling (3 to 60 min) in medusa tissue for samples incubated in the light (Hofmann and Kremer, 1981).

These experiments showed that labeled glycerol disappeared in less than 30 minutes,

! 22! likely through utilization in lipid synthesis, as suggested by the authors; these findings were later corroborated with findings of a similar study on the sea anemone

Exaiptasia (Davy and Cook, 2001). In vivo carbon fixation by the symbiont is highly efficient in C. xamachana, with rates exceeding daily respiration requirements of the host, supporting a growth rate of 3% body weight per day (Verde and McCloskey,

1998; Welsh et al., 2009; Freeman et al., 2016). Contrary to carbon, nitrogen assimilation levels under both photosynthetic and non-photosynthetic conditions are low, suggesting alternative nitrogen sources are essential for life. Interestingly, nitrogen metabolism appears to be largely driven by non-photosynthetic microbes, highlighting the complexity of the Cassiopea holobiont (Freeman et al., 2016). Lipids are also translocated between host and symbiont (Mortillaro et al., 2009; Imbs, 2013;

Imbs et al., 2014), but no cue has been found that explains the underlying mechanism or degree of translocation, although lipid composition and concentrations are known to shift under low light conditions in Cassiopea (Mortillaro et al., 2009). With the development of metabolomic tools and technologies such as NanoSIMS (nano-scale resolution chemical imaging mass spectrometry) (Kopp et al., 2013; Kopp et al.,

2015), Cassiopea offers an approachable and amenable system for the study of nutritional symbiosis. Given that this host can uptake multiple symbionts, future comparative studies conducted on various colonization regimes and conditions will undoubtedly shed light on cnidarian-Symbiodinium symbiosis.

Similar to the topic of nutrient translocation, little is known about the plasticity in jellyfish nutrient cycling. Jellyfish planulae are all thought to be born symbiont-free, and then following settlement the sessile polyps acquire resources through ingestion of zooplankton. However, less than 10% of essential carbon is provided by heterotrophic means during the adult medusa stage, and decreased

! 23! dependence on heterotrophy as a nitrogen source is seen with increasing medusa size

(Kremer, 2005). In some coral species, a shift from photosynthesis to heterotrophy may be triggered by thermal stress; this trophic switching is believed to enhance host survival in the event of bleaching events (Rodrigues and Grottoli, 2007; Anthony et al., 2009). Under reduced light conditions, a short-term increase in levels of chlorophyll a was measured in Cassiopea, but no evidence of a shift towards heterotrophy was seen that might compensate for the perceived reduction in photosynthetic activity (Mortillaro et al., 2009). In a separate study, providing nutrient supplements to bleached C. xamachana did not prevent a reduction in medusa body mass (McGill and Pomory, 2008), pointing out the limitation of heterotrophic contributions to the overall energy budget.

Behavior

Fluid Dynamics ! Jellyfish use bell contractions while swimming and changing direction in the water and to generate water currents that facilitate feeding and nutrient exchange.

Both fluid dynamics and related biological relevance of these activated currents to

Cassiopea medusae have been studied using empirical data and computational fluid dynamics simulations (Hamlet et al., 2011; Santhanakrishnan et al., 2012). Although the pulsing behavior is cyclic (one cycle comprising a single bell contraction and expansion with pauses in between), it results in continuous flow along the substrate towards the medusa and continuous upward flow above the medusa. During contraction, vortices develop at the bell margins that are forced through the oral arms, where they are broken up. The frilly structure of the oral arms of Cassiopea ensures thorough mixing of the water over their surface and secondary mouths before it is

! 24! pushed into an upward jet propulsion (Hamlet et al., 2011). Recently, a well-resolved three-dimensional computational fluid dynamics (CFD) model of bell contraction in the upside-down jellyfish was developed to better ascertain the flows generated by these medusae (Hamlet et al., 2011; Hamlet and Miller 2012; Santhanakrishnan et al.,

2012). Implementation of these models presents the possibility of further investigations of the effect of jellyfish aggregations on the local fluid and nutrient dynamics of a habitat.

At the ecosystem level, the pulsing behavior of upside-down jellyfish affects nutrient release from the substrate. Cassiopea's pulsing was shown to drive pore- water (interstitial water) up from the underlying sediment into the water column, allowing these derived nutrients to be used by the jellyfish and other reef organisms

(Jantzen et al., 2010). This type of bioturbation (the disturbance of sediment by organisms) is an important mechanism that couples the nutrient-rich sediments with the oligotrophic waters in which coral reefs occur. Cassiopea itself also releases organic matter, which is taken up by zooplankton in the water column (Niggl et al.,

2010). Furthermore, studies have demonstrated that the presence of upside-down jellyfish can have an effect on local nutrient dynamics, greatly affecting local oxygen concentration and ammonium regeneration rates in space (between sediment patches with or without jellyfish) as well as in time (given the diurnal cycle of the photosynthetic symbionts) (Welsh et al., 2009). These findings validate the notion that Cassiopea jellyfish play an important role in benthic-pelagic coupling of nutrient cycles, and some researchers even refer to the upside-down jellyfish as an ecosystem engineer (Jantzen et al., 2010). Further research is necessary to better understand the intricate role that Cassiopea medusae play in coastal ecosystems and nutrient cycling.

! 25! Quiescence ! Animal behavior is the product of numerous pathways, integrating both internal and external information to generate a global behavior. The internal state of an animal greatly affects its behavior – aroused animals act differently than those that are asleep. There is a debate as to how global brain states, like sleep, arise. It has been hypothesized that either specialized regions control the switch between wakefulness and sleep (a top-down mechanism), or neural networks have an emergent bias towards certain global states regulated by local circuits (a bottom-up mechanism). The fundamental question of the cellular mechanism controlling the sleep state is now approachable through detailed analysis of simple behavioral states.

Quiescence in the bilaterian model nematode Caenorhabditis elegans appears to be an emergent property; as the result of an internal network bias, it provides evidence against top-down regulation of a global brain state (Nichols et al., 2017).

Unlike C. elegans, neurons in Cassiopea, and other cnidarians, are organized in non- centralized radially symmetrical nerve nets, and yet there exists deep conservation among cnidarian and bilaterian neural structure and neurotransmitters. Recently, a sleep-like state was identified in Cassiopea, making it the earliest diverging lineage

(as a representative of Cnidaria) in which sleep has been observed (Nath et al., 2017).

Cassiopea medusae display the three behavioral characteristics that define a sleep state: quiescence that is rapidly reversible, homeostatic regulation upon sleep deprivation, and a delayed response to sensory stimuli, called sensory depression.

This recent recognition of a defined sleep-like state in Cassiopea, despite its lack of centralized nervous system, presents a unique opportunity to determine if sleep is, in fact, an emergent property of the earliest nerve networks. As a early diverging metazoan, Cassiopea is an important model for understanding scyphozoan behavior

! 26! and the underlying evolutionary mechanisms that have arisen to control behavior in derived animal lineages.

Cassiopea in the Environment

Bioinvasion and Blooms ! Molecular phylogenetic analyses demonstrated that some individuals of

Cassiopea from the same geographic area fall into different clades, evidencing cryptic species (Arai, 2001; Holland et al., 2004).⁠ At least two non-morphologically differentiated lineages of Cassiopea have been identified as alien species (Table1).

Cassiopea spp. have biological characteristics that make them potentially successful bioinvaders, such as their prolific asexual reproduction (Schiariti et al., 2014) and broad tolerance to variations in temperature and salinity (Morandini et al., 2017). The tenacity of Cassiopea spp. is demonstrated by their often well-established presence in disturbed or human-impacted habitats (Stoner et al., 2011). It is not surprising then, that species of Cassiopea have been documented as invasive in the Mediterranean

(Galil et al., 1990; Çevik, 2006; Schembri et al., 2010; Bayha and Graham, 2013),

Indo-Pacific (Papua New Guinea), Hawai'i (Holland et al., 2004), Australia (Keable and Ahyong, 2016), and Brazil (Morandini et al., 2017). Populations may occasionally reach enormous numbers (Stoner et al., 2011; Morandini et al., 2017) as a function of their environmental condition. With the frequency of these events potentially increasing according to studies on climate change (Richardson et al., 2009;

Brotz et al., 2012), it necessitates development of methods for accurate taxonomic identification of cryptic species. These sudden proliferation events can be useful indicators of shifts in environmental conditions.. Recent work has shown, to the contrary, that projected increased temperatures combined with increased exposure to

! 27! UV-B radiation will lead to a general reduction in Cassiopea population size (Klein et al., 2016b). These findings are perplexing given the perception that jellyfish have an exceptional capacity to adapt to and thrive under conditions associated with ocean warming.

Environmental Monitoring and Ecotoxicology ! As marine systems worldwide continue to decline from local and global physical, chemical and biological threats (De'ath et al., 2012; Epstein et al., 2016), the need to manage marine natural resources and reduce these threats is becoming increasingly more critical. Early detection and quantification tools are needed to assess risks that can have adverse impacts on the biological, physiological, and metabolic conditions at the level of the organism and the ecosystem (Markert et al.,

2003; Downs, 2005). The use of jellyfish as biomonitors and/or bioindicators previously was not explored, possibly due to historical perceptions of their tolerance for poor environmental conditions and ‘boom or bust’ population fluctuations.

Biomonitors are used to derive both qualitative and quantitative information about the environment. There has been recent interest in establishing Cassiopea as a potential biomonitoring tool (i.e., indicator species) for enhancing water quality management.

Previous studies have indicated jellyfish are able to accumulate trace metals in concentrations exceeding those present in ambient seawater, and concern exists that these could have potential for transfer up food webs (Romeo et al., 1987; Romeo and

Gnassia-Barelli, 1992). For example, copper and zinc accumulate in Cassiopea

(Templeman & Kingsford, 2010, 2012, 2015; Epstein et al., 2016) generally within hours to several days and with varying residence times (Fowler et al., 2004), while phosphate concentrations in C. xamachana can reach an order of magnitude greater

! 28! than those of ambient conditions, even when such levels are undetectable in seawater

(Todd et al., 2006). The rapid response of Cassiopea medusae under exposure to metals and nutrients suggests they could serve as a useful indicator tool to detect and quantify short pulses of contaminants/pollutants entering a system, a role achieved by only a few of the current typical biomonitors.

In contrast to biomonitors, bioindicators are defined as organisms that will exhibit a change in structure or function that is linked to the biological effect of a contaminant at the organism, population, community, or ecosystem level (McCarty,

2002; McCarty and Munkittrick, 2008). Bioindicator species are used in laboratory bioassays to establish criteria for ecological sensitivity (e.g. LC50), but there is still a need to identify model organisms for standardization of these assays (Chapman,

1995;Edwards et al., 1996;Van Gestel and Van Brummelen, 1996). There is clear ecological relevance for exploring Cassiopea as a tool for ecotoxicological assessment - briefly defined as the study of effects of toxic chemicals (especially on the population level). As cnidarians, jellyfish occupy a key evolutionary position as early metazoans, serving important ecological roles as predators and prey (Faimali et al., 2014); additionally, they fuel pelagic food webs with dissolved and particulate organic matter (Niggl et al., 2010). Cassiopea offers an additional benefit as a bioindicator, as both the host jellyfish and dinoflagellate symbiont responses to environmental conditions can be assessed. Studies conducted on different life stages of Cassiopea exposed to copper or zinc demonstrated measurable effects at environmentally relevant concentrations (Templeman and Kingsford, 2015). In other studies, agricultural formulations of diuron and hexazinone at ecologically relevant concentrations caused measurable inhibition with respect to Cassiopea growth, photosynthetic efficiency, and symbiont density within seven days of exposure

! 29! (Rowen et al., 2017). A study looking at interactive responses between herbicides and salinity demonstrated that reduced salinity increased herbicide toxicity (Klein et al.,

2016a). Pollutants in sewage and agro-chemicals have also been shown to cause reproductive and recruitment failure across most invertebrate phyla (Depledge and

Billinghurst, 1999). There is an urgent need for new and improved bioassays to detect and assess new chemicals for their potential effects on marine invertebrates.

Cassiopea has the potential to contribute extensively to understanding the effects of contaminants and pollutants on coastal marine ecosystems; the various genomic resources available for Cassiopea provide a tractable means to identify potential in situ molecular biomarkers and cellular diagnostic endpoints.

Other Laboratory Applications

Toxinology and cnidome Despite the ubiquity of cnidarian species in a variety of aquatic environments, toxinological research concerning cnidarian venom has been significantly delayed compared to advances seen in research applications related to venomous terrestrial animals (Turk and Kem, 2009). Early clinical research on cnidarian venom validated a diversity of symptoms presented in human sting victims, including but not limited to anaphylaxis (Togias et al., 1985, Radwan and Burnett, 2001). Studies have also emphasized the potential for cnidarian bioactive proteins in pharmacological applications (Jha and Zi-rong, 2004). Cassiopea medusae have a mild capacity to sting, and only a few cases of human envenomation have been reported to cause symptoms ranging from a rash, swelling, vomiting, and urticaria (Rifkin and Fenner,

1996). Cnidarian venom is delivered to prey and potential predators via the firing of microscopic nematocysts. In Cassiopea andromeda, five different types of nematocysts have been identified: a-isorhizas, O-isorhizas, euryteles, large oval

! 30! birhopaloids, and small lemon-shaped birhopaloid (Heins et al., 2015). Studies including toxinological inferences have focused on several bioactive properties of crude venom extracts, which include cytolysis and hemolysis (Radwan et al., 2001;

Radwan and Burnett, 2001; Torres et al., 2001; Radwan et al., 2005). Neurotoxic effects were also identified in two of Cassiopea’s nematocyst types (birhopaloid and a-isorhiza) (Gülşahin, 2015). The ease at which Cassiopea can be collected globally in mangroves and cultured in the lab, provides a clear advantage to studying this species’ venom, which has antitumor and antiprotozoal properties that may serve as future novel pharmacological and therapeutics agents (Orduña-Novoa et al., 2003;

Morales-Landa et al., 2007; Mirshamsi et al., 2017). A recent study showed that

Cassiopea crude venom induced apoptosis in human breast cancer tissue via reactive oxygen species (ROS) mediated cytotoxicity (Mirshamsi et al., 2017). Cassiopea, therefore, has potential for future bioactive natural product discovery through modern

“venomics” approaches (genome, transcriptome, proteome, metabolome) that have applications to elucidating evolutionary mechanisms driving the complexity and diversity of cnidarian venom (Radwan et al., 2001, Casewell et al., 2013; Jouiaei et al.,

2015).

Cassiopea Virology ! In comparison to bacterial components of the cnidarian holobiont, studies on the virome have received considerably less attention, but are of interest due to the potentially harmful effects that viruses can have on cnidarian health (Thurber and

Correa, 2011). There is increasing recognition for biological and ecological importance of viruses in marine samples (Bosch et al., 2015; Thurber et al., 2017), and their role in evolution (Grasis et al., 2014). Preliminary data from the anthozoan

! 31! Exaiptasia, has identified several RNA viruses, including those from Partitiviridae and Picornaviridae (Brüwer and Voolstra, 2017). The potential medical relevance, complex life strategies, as well as their host range and evolutionary history, the possible presence of RNA viruses associated with the jellyfish is of great interest to the field (Johnson, 2015; Koonin et al., 2015), and future studies should shed light on the yet-to-be explored evolutionary path of virus-cnidarian co-evolution. Furthermore, some cnidarians express genes that have been implicated as potential anti-retroviral agents (Ramessar et al., 2014), and our preliminary findings of several viruses in

Cassiopea validate the necessity of these putative defense mechanisms. Going forward, studies on Cassiopea viromes may provide a unique opportunity to not only expand our understanding of the current host range of Mononega- and retro-like viruses, but also to establish a reliable and robust model system for rigorous investigation of interactions within the holobiont as they relate to possible host immunity, in all stages of the jellyfish’s life history.

Cassiopea as a Laboratory Resource ! The ease at which the asexual polyp stage can be propagated in the laboratory using artificial seawater makes Cassiopea an attractive system for sustained culturing.

Depending on the frequency of feeding, polyps are capable of developing motile planuloid buds within 24 hrs (Van Leishout and Martin, 1992). Swimming buds can settle onto a suitable substrate within 24 hrs and undergo metamorphosis into a polyp.

Therefore, the continuous release of planuloid buds permits an aposymbiotic polyp to propagate a perpetual colony of asexual polyps. Currently, twelve C. xamachana lines

(originally from Florida) have been established by the Cassiopea consortium. A draft

C. xamachana genome has been assembled (ENA Accession: PRJEB23739), as well

! 32! as transcriptomes of the aposymbiotic, symbiotic, and strobila stages (European

Nucleotide Archive Accession: PRJEB21012). Future application of metabolomics approaches in Cassiopea research will also help generate a profile of the underlying metabolic pathways and metabolite dynamics that might be affected by environmental factors during key stages of Cassiopea development and symbiosis.

While aposymbiotic polyps can be propagated as described above, symbiotic polyps can be generated by providing the polyps with Symbiodinium spp. Cassiopea polyps are capable of forming associations with virtually all symbiotic Symbiodinium cultures (Lampert, 2016; Thornhill et al., 2006), making this an ideal system for comparative studies. The colonization characteristics of Cassiopea provide two tractable phenotypes to visually confirm variation in the symbiotic association.

Symbiodinium density within the amoebocytes can vary depending on the colonizing species (Winstead et al., 2018), and strobilation timing also appears to vary accordingly (Lampert, 2016). Newly strobilated ephyrae are available for experimentation; likewise the adult medusae can also be acquired with relative ease, through animal transfer with public aquariums and through the recreational aquarium trade. In addition, only simple setups are required to rear each stage, making

Cassiopea an easily manageable laboratory animal (Widmer, 2008). Fertilized eggs can be obtained from the brooding female adult medusae, confirmed by the initiation of initial cleavage post-collection (Gohar and Eisawy, 1960; Smith, 1936). Cassiopea are gonochoristic, with females identifiable through the presence of specialized central brood tentacles (Hofmann and Hadfield, 2002). Spawning can likely occur year-around under ideal conditions, and fertilized eggs collected directly from the brood tentacles after spawning (Gohar and Eisawy, 1960; Hummelinck, 1968). The planula larvae stage can also be acquired directly from the brooding female and used

! 33! for additional experimentation. Tools for working with Cassiopea are increasingly being made available to researchers and educators, with many such protocols available online. In particular, a customized whole mount in situ hybridization protocol has been developed for Cassiopea, along with other molecular tools and resources, which are accessible through the online portal

(http://sites.psu.edu/cassbase/). Single cell injection for transgenic approaches is currently under development, with a promising outlook for becoming an important resource for mapping jellyfish molecular pathways.

Conclusions ! Cassiopea is a system predominantly studied for the many questions that can be addressed on interdependent organisms given its remarkable Cnidaria-

Symbiodinium association. This review demonstrates that in addition to the effectiveness of the Cassiopea system for elucidating the complex processes underlying symbiosis, Cassiopea has potential to contribute substantially to our knowledge of a wide range of research topics within the field of invertebrate biology.

The recent acquisition of genomics resources and development of several new molecular tools for Cassiopea sets this cnidarian apart as a model for identifying possible genotypic and phenotypic traits to support the hypothesis of co-evolution between Cassiopea and Symbiodinium. Given Cassiopea’s position as a well- established lab organism, the time is right for studies that seek to characterize possible changes in molecular composition (at the gene, metabolite or peptide-level) of one or both symbiotic organisms, phenotypic attributes that may have evolved as a result of this interspecific interaction, or the putative effects of ecological intimacy on each organism’s level of adaptation to the shared environment. These questions, and

! 34! countless others, can be properly interrogated by way of rigorous observations, diverse experimentation, and implementation of phylogenetic analyses in an attempt to infer potential reciprocal evolutionary change.

The Cassiopea system also has obvious applications to the fields of bacterial and viral co-evolution, from the dual perspective of understanding microbial symbiosis between either the cnidarian host or the associated dinoflagellate symbiont.

Furthermore, Cassiopea shows promise as an effective bioindicator for our rapidly changing oceans, and as a biomonitor for further investigations of the medusa’s potential impact on coastal ecology on a broader geographic scale. The relatively straight-forward culturing protocols and approachability of the system make

Cassiopea particularly feasible as a teaching tool, not only at the college level but also from Kindergarten through grade 12. This review is a culmination of many years of multidisciplinary, collaborative research that has helped position the upside-down jellyfish Cassiopea as a valuable resource to researchers in the omiwcs era seeking an effective model system to better understand the natural history and biodiversity of some of the earliest multicellular animals through studies on evolution, co-evolution and beyond.

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! 65!

Figure!1.!A!schematic!summarizing!the!tools!and!approaches!available!for!the! Cassiopea(system,!highlighting!the!utility!of!each!life!stage!towards!a! multidisciplinary!approach!to!understanding!the!biology!and!physiology!of!this! early!metazoan.!

! 66!

Figure!2.!A!working!hypothesis!of!Medusozoa!phylogeny!based!on!several!studies! (Bayha!et!al.,!2010;!Kayal!et!al.,!2013;!Kayal!et!al.,!2017),!including!the! monophyletic!Scyphozoa,!Rhizostomeae,!Kolpophorae,!and!Cassiopeidae.! Principal!morphological!and!life!cycle!characters!are!plotted!on!the!tree!(1!=! cnidae;!2!=!medusa!stage;!3!=!rhopalia;!4!=!strobilation!and!ephyra;!5!=!monodisc! strobilation).!Note:!The!topology!is!not!meant!to!represent!a!consensus.!

! 67!

Figure!3.!A!Cassiopea(xamachana!poylp!with!a!planuloid!bud!developing!asexually! from!the!aboral!region!of!the!calyx.!

! 68!

Lappets in Life Size Exumbrella Type Species each Rhopalia Mouth arms Appendages Distribution cycle Bioinvasion GenBank* (cm) shape locality paramere known n small, 5 or Western Atlantic, C. andromeda more 10- 12-19 North Pacific, (Forskål, flat 1-10 4–6 branches large, club Red Sea Yes+ Yes1 Yes 12 (16) Mediterranean 1775) shaped Sea, Indo-Pacific vesicles Madagascar and C. depressa 10- Madagascar, off coast of Haeckel, flat 9 16 6–8 branches n small clubs No No No 12 Mozambique Mozambique, E. 1880 Africa bifurcated at their ends 30–40 small C. frondosa 12- West Indies, West Indies to flat 5 12 with short, leaf shaped No No Yes (Pallas, 1774) 26 Caribbean Florida pinnate side vesicles branches C. broad, maremetens 4–6 with 1–2 central, 1 Lake shallow, Queensland, Gershwin, 2-20 4 19 distal base Magellan, No No No aboral Australia Zeidler & bifurcation arm, 1 tip arm Australia concavity Davie, 2011 C. medusa n branches (3 Culion Bay, Hawaii & 26 flat 7 17 n small/large No Yes2 No Light, 1914 main distal) Philippines Philippines rounded Kosrae C. mertensi 10- 8-12 main Caroline Islands 3 without a 8 16 n large clubs Island, No Yes No Brandt, 1835 12 branches & Hawaii concavity Micronesia C. ndrosia 6-12 main n small leaf Agassiz & 5 concave 4 18-22 Suva, Fiji Australia & Fiji No No No branches shaped Mayer, 1899 C. ornata Palau, Papua 10- 9- 12 side Malay Haeckel, flat 5 16 n small clubs New No No Yes 12 branches Archipelago 1880 Guinea flatly n small and C. rounded, with pinnately few large Caracas Bay, Caracas Bay, vanderhorsti 17 3-5 14-18 No No No a low central dichotomus vesicles Curaçao Curaçao Stiasny, 1922 dome C. 10–15 large and Kingston xamachana 11-23 West Indies to 15 concave 5 alternate small ribbon- Harbor, Yes- No Yes Bigelow, (16) Florida branches like filaments Jamaica 1892

Holland et al., (2004) 2 Doty (1961) 3Uchida (1970) + Gohar & Eisawy (1960a) - Bigelow (1900)

Table!1:!Morphological!data!on!valid!Cassiopea!species,!modified!from!Morandini! et(al.,!(2017)

! 69! !

Chapter 3

Is larval settlement predictable? Genomic insights of settlement and metamorphosis inducing bacteria of Cassiopea xamachana

Aki H. Ohdera1, Khushboo Attarwala1, Henry Rubain1, Henry Laird2, William Fitt2, Mónica Medina3,4

1. Department of Biology, Pennsylvania State University, University Park, PA, USA

2. University of Wisconsin, Madison, USA

3. Odum School of Ecology, University of Georgia, Athens, GA, USA

4. Smithsonian Tropical Research Institute, Smithsonian Institution, Panama

City PL, Washington D.C. USA

! 70! !

Abstract! ! ! Many!marine!invertebrates!exhibit!a!life!history!transition!whereby!the! planktonic!larvae!undergo!metamorphosis!to!develop!into!the!juvenile!or!adult! form.!The!timing!and!location!of!this!transition!is!informed!by!both!biotic!and! abiotic!factors,!but!larvae!of!many!species!associate!with!bacteria!in!order!to! initiate!the!developmental!transition.!While!the!importance!of!site!selection!is! demonstrated!by!the!high!degree!of!specificity!to!particular!substrates,!we!have! yet!to!understand!the!cue!or!cues!responsible!for!triggering!settlement!and! metamorphosis!in!many!invertebrates.!In!the!scyphozoan!Cassiopea)xamachana,! bacteria!has!been!implicated!in!settlement!and!metamorphosis!but!the!source!of! the!settlement!cue!associated!with!their!preferred!substrate!has!yet!to!be! determined.!Bioassays!indicate!the!marine!specific!genus!Pseudoalteromonas,!as! well!as!bacteria!from!other!phyla,!are!capable!of!inducing!settlement!and! metamorphosis.!Genomic!comparisons!of!select!inducers!highlight!the! difficulties!in!understanding!the!molecular!mechanism!of!the!association.!

However,!the!results!from!this!study!point!to!potential!mechanisms!that!may! facilitate!the!induction!of!settlement!and!metamorphosis!in!Cassiopea) xamachana!and!suggest!an!evolutionary!redundancy!in!cue!detection!is! important!for!site!selection.!

! 71! !

Introduction ! The transition from a planktonic to a benthic environment is a crucial step in the life history of many marine invertebrates (Hadfield and Paul, 2001;Hadfield,

2010). Settlement, defined as the transition in which planktonic larvae enter and make contact with the benthos, is often followed by a developmental transition to form the juvenile/adult body plan or metamorphosis. Abiotic factors, including light, temperature, salinity, and substrate can inform settlement site selection, in addition to biotic factors such as chemical cues (Leitz, 1997;Fleck and Fitt, 1999;Gambill et al.,

2016). These cues can originate from multiple sources, including conspecifics

(Wieczorek and Todd, 1998;Cahill and Koury, 2016), or from substrates such as crustose coralline algae (Williams et al., 2009;Grasso et al., 2011), and bacterial biofilms (Abdul Wahab et al., 2011;Bacchetti De Gregoris et al., 2012). Of these, bacterial biofilm has been found to produce the cue responsible for settlement and metamorphosis in a significant number of marine taxa (Fitt et al., 1990;Unabia and

Hadfield, 1999;Negri et al., 2001;Whalan and Webster, 2014).

Given the prevalence of bacterial-metazoan interactions, this association likely existed early in metazoan evolution (McFall-Ngai et al., 2013). Colony or "rosette" formation in the choanoflagellate Salpingoeca rosetta is regulated by factors released by an environmental bacterium Algoriphagus machipongonensis (Alegado et al.,

2012;Woznica et al., 2016;Woznica and King, 2018). Given the parallels of colony formation in choanoflagellates, and the ubiquity of bacterial interactions related to development in metazoans, bacterial-metazoan signaling likely played an important role early in metazoan evolution. Bacterial signaling driving the plankton to benthos transition may exemplify the importance of bacterial-metazoan interaction (Pawlik,

1992;Huggett et al., 2006;Huang et al., 2012). The biofilm-larval interaction can be

! 72! ! highly specific, with larvae delaying settlement for prolonged periods (Elkin and

Marshall, 2007). Despite the ecological and evolutionary importance of biofilm to marine benthic communities, few chemical cues responsible for settlement and metamorphosis have been identified.

In the polychaete tubeworm Hydroides elegans, a marine bacterial genus

Pseudoalteromonas has been shown to be a strong inducer of settlement and metamorphosis. Thus far, two different inducers, tetrabromopyrrole and a phage tail- like viral particle have been isolated from the genus (Tebben et al., 2011;Shikuma et al., 2014), highlighting the diversity in mode of signaling (chemical vs tactile). The mechanism driving this larval-biofilm interaction has yet to be explored and whether bacterial biofilm can have benefits beyond dictating settlement site. Biofilm found on decomposing leaves has been suggested to have nutritional value to post larval penaid shrimp (Gatune et al. 2012). Bacterial secondary metabolites are also known to have anti-bacterial and anti-fungal properties (Holmstrom and Kjelleberg 1999, Bosch

2013), potentially acting as a "secondary immune system".

The upside-down jellyfish Cassiopea xamachana is found in mangrove forests worldwide. Though capable of motility, the adult stage is primarily benthic and remains in shallow areas with access to sunlight to allow photosynthesis by its intracellular dinoflagellate symbiont (Symbiodinium microadriaticum). The brooded larvae are released into the water column and will often settle preferentially onto the underside of degrading mangrove leaves. Previous studies have shown degraded mangrove leaves to be a source of an inducer for settlement and metamorphosis

(Fleck and Fitt 1999). Fractionation of leaf extracts led to the identification of a water soluble, 5.8 kDa proline-rich peptidic cue (Fleck et al. 1999). While the compound was isolated, the identity of the peptide could not be determined. In addition, it is

! 73! ! unclear whether the sources of the cue is plant or bacterial in origin. In support of

Cassiopea responding to bacterial cues, Neumann (1979) showed larvae of Cassiopea andromeda to metamorphose in response to the biofilm of Vibrio sp. This led to the isolation of a 1 - 10 kDa fraction with inductive capacity, although the identity of the peptide was not determined. A cholera toxin isolated from V. cholerae was also found to induce settlement and metamorphosis (Wolk et al., 1985), and a fraction extracted from collagen digested by V. alginolyticus was also found to be inductive (Hofmann and Brand, 1987). These findings suggest bacteria associated with degrading mangrove leaves may be a likely source of the inductive cue.

In order to determine whether bacteria are responsible for inducing settlement and metamorphosis in C xamachana, we isolated and cultured degrading mangrove leaf associated bacteria and tested the response of C. xamachana larvae to these isolates. We examined the microbiome of degrading mangrove leaves to determine the prevalence of settlement inducing bacteria on the preferred substrate of C. xamachana.

We generated draft genomes of several bacterial isolates and explored the genomes to detect potentially predictive genes for larval settlement and metamorphosis induction and explore the efficacy of using genomic data for cue prediction.

Materials and Methods !

Larval Collection ! Brooding females of C. xamachana were collected from a seagrass bed off of

Key Largo, FL (25.102204, -80.438708, 24.749522, -80.978341)(Fig. 1). Eggs were acquired from the brooding C. xamachana females by gently pipetting seawater at the central brood tentacles and collecting the released egg capsules. The egg capsules

! 74! ! were transferred to glass finger bowls containing antibiotic seawater (100 mg neomycin and 130 mg streptomycin dissolved in 1 L seawater). Embryos were left undisturbed for 48 hours to allow development into motile planulae. The planulae were transferred to fresh antibiotic seawater (ABS) and used within 24 hours for the bioassays.

Bacterial Isolation ! Degraded mangroves were collected from below mangrove stands and rinsed gently three times with 0.2 µm filtered seawater (FSW). 1 cm2 pieces were cut from the leaves and homogenized in 25 ml of FSW using ethanol sterilized mortar and pestle. Serial dilutions of the homogenate were made by transferring 0.5 ml of the diluted homegenate into 9.5 ml of FSW for 5 dilutions. 100 µl of each dilution was plated onto marine agar and incubated at 28°C for 24 hr. Bacterial colonies were picked and preserved in FSW with 15% glycerol at -80°C.

Identification of bacterial Isolates ! The 16S region of the isolates were sequenced using universal primers 27F

(AGAGTTTGATCMTGGTCAG) and 1492R (TACGGYTACCTTGTTACGACT) primer pairs (Weisburg et al., 1991). Bacterial stocks were plated onto marine agar and grown for 48 hours. Colonies were picked and added directly to PCR reactions using the following program: 95 for 5 min, 94 for 1, 55 for 1:30, 72 for 2:00, 72 for

5:00, for 35 cycles. PCR products were submitted for Sanger sequencing at the

Pennsylvania State University Sequencing Core Facility. Chromatograms were manually curated using MEGA and sequences were compared to both RDP and NCBI

! 75! ! databases in order to identify the isolates. Taxonomic identification was further confirmed by generating alignments to sequences retrieved from Genbank using

MUSCLE and phylogenetic inference was determined using MEGA.

Settlement Bioassays ! Biossays were prepared in 12 well cell culture plates by filling individual wells with 1 ml of sterile marine media (Difco Marine Broth 2216). Replicate wells for the FSW negative control were left unfilled with marine media. Wells were inoculated with isolates by transferring a single plate grown colony to a well. Wells were inoculated in triplicate per isolate and were randomly assigned across multiple plates. A second negative control was included in which wells were left un-inoculated and rinsed with FSW. Plates were incubated at 28°C for 24 hours without shaking to allow the bacteria to grow and settle to the bottom of the well. Wells were gently rinsed with 1 ml of FSW without disrupting the biofilm for a total of 2 washes at the end of the 24 hrs. Wells that were still turbid after the two washes were rinsed for a third time, or until the seawater remained clear. Wells were then filled with a total of

2 ml of 0.2 µm FSW. A positive control was included in the assay where the synthetic settlement and metamorphosis inducing peptide (GPGGPA) were added at a final concentration of 1.5 x 10-5 M. Approximately 10 C. xamachana larvae kept in ABS for up to 48 hrs were placed in each of the wells. Plates were kept at 25°C in the dark.

The plates were scored for settlement and metamorphosis every 24 hrs using a dissecting microscope.

In addition to biofilm, larvae were exposed to filtrate produced from select bacterial cultures. Bacteria were grown as described above for 24hrs. The culture was centrifuged at 3000 g for 10 minutes to pellet the cells and supernatant was collected

! 76! ! and filtered through a 0.2 µM filter. Excess supernatant was removed and the pelleted cells were re-suspended in 1 ml of FSW prior to spinning at 3000 g for 10 minutes.

The supernatant was discarded and cells re-suspended in 1 ml of FSW. 50 and 100 µl of the filtrate and 100 µl of re-suspended bacterial cells were added to a total volume of 1 ml FSW in 24 well plates. Plates were allowed to stand for 1 hr prior the start of the experiment to allow cells to settle to the plate bottom. 10 larvae were added to each well and settlement was scored as described above.

16S microbiome extraction and sequencing ! Mangrove leaves were collected during two consecutive years in Key Largo

Florida from five different locations (24.750769, -80.979027, 25.101526, -80.438872.

Leaves were immediately stored at -20 for DNA extraction. DNA was extracted using the Mobio PowerSoil DNA Isolation Kit following the manufacturers instructions.

The V4 region of the 16S rRNA gene PCR amplified using the 515F

(GTGCCAGCMGCCGCGCGGTAA) - 806R (GGACTACHVGGGTWTCTAAT) primer pairs (Caporaso et al., 2011). Sequencing was performed at the Joint Genome

Institute (JGI) for samples collected in 2013 and samples collected in 2015 were sequenced at the University of Illinois - Chicago Core Genomics Facility using the

Illumina HiSeq 2000. Initial processing of the reads was conducted according to the

Earth Microbiome Project (http://www.earthmicrobiome.org/protocols-and- standards/) Qiime pipeline (Caporaso et al., 2010) with the closed reference option using the Greengenes 97% OTUs database. Further downstream analysis we also conducted using Qiime.

! 77! !

Genome Sequencing and Analysis ! Bacterial isolates MB4-1, MA6-2, and MA6-4 were chosen for whole genome sequencing based on their inductive capacities. Colonies were grown in marine media for 24 hrs. Genomic DNA was extracted following the JGI Bacterial DNA Isolation

CTAB-2012 protocol (https://jgi.doe.gov/user-program-info/pmo-overview/).

Illumina sequencing libraries were constructed using the Illumina TruSeq Nano DNA prep Kit with a 550 bp insert size. Libraries were sequenced on the Illumina Miseq, generating 2.8, 3.0, and 2.7 Gb of 300 bp paired-end sequence data for MB4-1, MA6-

4, and MA6-2, respectively. Draft genomes were de novo assembled using the A5- miseq pipeline (Tritt et al., 2012). Genomes of Pseudoalteromonas luteoviolacea strains HI1 (Asahina and Hadfield, 2015) and 6061 were downloaded from NCBI

(accession JWIC01000000, AUYB00000000). Genes were annotated with

Glimmer3.0 option of RAST (Aziz et al., 2008). The presence of mac genes and the tetrabromopyrrole gene cluster were determined using BLAST 2.5.0+ (Camacho et al.,

2009). InterproScan (Zdobnov and Apweiler, 2001) was used to search the genomes for proline-rich proteins, in addition to manually searching for short proline repeats within predicted sequences. Genes identified as homologs of TonB-C were further analyzed to determine whether they were membrane bound or signaling proteins using tmhmm (Krogh et al., 2001) and signalP (Petersen et al., 2011). Predictions for the presence of secretion systems were preformed using T346Hunter (Martinez-Garcia et al., 2015). The genomes were searched for secondary metabolite biosynthetic gene clusters using AntiSmaSH (Weber et al., 2015).

! 78! !

Results

Bacterial Settlement Bioassay ! Bacterial settlement varied between isolates (Fig. 2), with average metamorphic induction rates ranging from 0% to 77.5%. Negative controls (0.22 μm filtered sea water, growth media + FSW) showed less than 1.5% rates of induction, while the artificial inducer treated larvae metamorphosed at an average rate of 58.9%.

Isolate MB4-1 showed greater metamorphic induction than the positive control at

77.5%, while all other settlement inducers ranged from 32.7% to below 5%. Of the isolates assayed, five elicited a significant settlement and metamorphosis response in

Cassiopea larvae (p < 0.05) (Fig. 2). Using the full length16S sequence, isolates were identified to the genus level (Fig. 3). Inductive bacteria were assigned across genera belonging to multiple phyla. The isolate with greatest inductive capacity (MB4-1) was identified as a Pseudoalteromonas sp., most closely related to P. lipolytica (Fig. 4). A subset of the bacteria was cultured to also test the inductive capacity of filtered cells and the media in which the bacteria were grown. Bacterial cells separated from the growth media showed no induction of settlement (Fig. 4), while the culture media filtrate showed high rates of induction, ranging from 30 - 80% at the higher concentration (Fig. 5). The filtrate resulted in higher rates of settlement compared to some biofilm. Interestingly, larvae only settled in response to the filtrate and did not undergo metamorphosis. Settled larvae remained attached, but failed to develop tentacles or a mouth.

Settlement Substrate Microbiome ! We sequenced the 16S microbiome of degrading mangrove in order to determine if inductive bacteria identified from the settlement assays were in high

! 79! ! abundance relative to the rest of the biofilm community. The bacterial community of five sites (Fig. 1) and two different degradation states over two years were compared.

Community composition showed settlement and metamorphosis inducing taxa to be rare, with all isolates comprising less than 0.001 % of total bacterial abundance within the leaf biofilm, including Pseudoalteromonas MB4-1 (Fig 5). PCoA analysis showed samples were largely clustered based on year and location, although leaves collected from the quarry that did not have settled polyps clustered closely with those collected from the North Bay the previous collection year (Fig. 6).

Bacterial Genome Analysis ! Isolates tested for their inductive capacity were classified under three categories (high, moderate, low) based on the settlement and metamorphosis success of the larvae. Isolates that induced metamorphosis in greater than 30 percent of larvae were considered high, between 30 and 5 percent were considered moderate, and less than 5 percent were considered low. A single isolate from the three categories were chosen for whole genome sequencing. The genomes of Pseudoalteromonas MB4-1

(high), Nisaea MA6-4 (moderate), and Thalassospira MA6-2 (low) were sequenced, resulting in draft genomes assemblies with total lengths of 4.85 Mb, 5.5 Mb, and 7.2

Mb, respectively. Gene prediction identified 4454, 5256, and 7261 genes for MB4-1,

MA6-4, and MA6-2, respectively. The newly sequenced genomes were compared to

Pseudoalteromonas luteoviolacea strain HI1 and P. luteoviolacea strain 6061. We utilized available tools to compare the five genomes in order to identify genes potentially related to settlement and metamorphosis of Cassiopea xamachana larvae.

Predictions of secretion systems with T346Hunter (Martinez-Garcia et al. 2015) identified variation in the repertoire of secretion systems found across the five

! 80! ! genomes. All clustered genes associated with the presence of the flagella were confirmed in all the genomes, while the Type VI secretion system (T6SS) gene cluster was only found in MB4-1, MA6-4, and HI1. Interestingly, the type IV secretion system (T4SS) gene cluster was only found in the two genomes (MB4-1 and MA6-4) shown to induce settlement and metamorphosis in C. xamachana.

Phage tail-like viral particles or MAC genes first identified in P. luteoviolacea

HI1 (Huang et al. 2012) could only be identified in the Pseudoalteromonas genome of the same strain. The gene cluster responsible for tetrabromopyrrole production was only identified from the P. luteoviolacea HI1 genome. However, genes with high similarity to those found in the tetrabromopyrrole cluster were found in the other four taxa, but were located at separate loci around the genomes and not found in tandem.

Fleck et al. (1999) identified a 5.8 kDa peptide extracted from degrading mangrove leaves to induce settlement in Cassiopea larvae. As the peptide was rich in proline, we searched the genomes for proline-rich regions using InterProScan and BLAST.

InterProScan did not identify any proline-rich regions within the five genomes, but genes containing the relatively proline-rich TonB-C domain were found in all three

Pseudoalteromonas genomes as well as Nisaea MA6-4. Tmhmm identified many of these genes to contain transmembrane domains. Analysis with the signal peptide prediction software SignalP did not identify TonB-C to produce a secreted signal peptide.

Antibiotic and secondary metabolite prediction using AntiSmaSH identified

14 clusters in Pseudoalteromonas MB41 (Table S1), 22 in Nisaea MA64 (Table S2),

47 in Thalassospira MA62 (Table S3), 33 in Pseudoalteromonas HI1 (Table S4), and

46 in Pseudoalteromonas 6061 (Table S5). While most of the cluster predictions were undetermined in function, bacteriocin clusters were found in 4 of the 5 genomes.

! 81! !

Other common secondary metabolite clusters coded or pathways belonging to polyketide synthases and saccharide biosynthetic pathways. Interestingly, more secondary metabolite gene clusters were predicted in non-inductive bacteria.

Comparisons of the clusters identified the the aryl polyene cluster to be specific to inductive bacteria, although the biosynthesis product of the two clusters are unknown.

Discussion ! Bacteria have been implicated to be the source of larval settlement cues for many invertebrate species (Neumann, 1979;Abdul Wahab et al., 2011;Grasso et al.,

2011;Tebben et al., 2011;Ganesan et al., 2012;Whalan and Webster, 2014;El Gamal et al., 2016). The ubiquity of these bacteria-larvae interactions likely suggests the association has persisted since the Cambrian, playing an important role in metazoan evolution. These cues are diverse, from water-soluble compounds to membrane- bound viral-like particles. Bacteria-larvae interaction is important for benthic recruitment in many invertebrate species, and has garnered attention as it relates to biofouling and commercially important species (Fitt et al., 1990;Satuito et al.,

1997;Huggett et al., 2006;Huang et al., 2007;Ritson-Williams et al., 2009;Williams et al., 2009). Understanding larval settlement provides information on species distribution and can aid in restoration efforts, particularly in the case of ecologically important scleractinian coral species (Vermeij et al., 2009). While several cues have been identified to induce settlement and metamorphosis, the underlying basis for larval-microbe interaction has not been extensively addressed. This is demonstrated in some species exhibiting attachment response to monocultured biofilm, but fail to continue to metamorphosis (Yang et al. 2013). In the case of the carnivorous nudibranch Onchidoris bilamellata, settlement is triggered upon detection of a water- soluble metabolite produced by its barnacle prey, while metamorphosis is triggered

! 82! ! upon contact with shells of dead barnacles (Chia and Koss 1988). On the other hand, both settlement and metamorphosis can be triggered by the same cue in other invertebrates. While research outlining known inducers has been extensively reviewed (Hadfield and Paul 2001, Hadfield 2010), we still lack a thorough understanding of the extent of specificity in marine invertebrates and the basis for the evolution of such interaction. In particular, Pseudoalteromonas species, which induce settlement and metamorphosis in several marine taxa, are capable of producing a suite of anti-bacterial products (Holmstrom and Kjelleberg 1999). Small brominated compounds are a class of compounds found in P. luteoviolacea with bactericidal effects. Tetrabromopyrrole, a similar brominated compound has been shown to induce settlement in Acropora millepora (Tebben et al. 2011). Larval-microbial association may be driven by immune protection afforded by the associating biofilm (Bosch

2013).

Several marine bacterial species are known to be inductive of settlement and metamorphosis in Cassiopea spp. Biofilm formed by a monoculture of the marine bacterium Vibrio sp. was found to induce settlement and metamorphosis in Cassiopea andromeda (Hofmann et al., 1978). Settlement and metamorphosis of C. andromeda larvae was also observed when exposed to cholera toxin isolated from Vibrio cholerae

(Wolk et al., 1985) and Vibrio alginolyticus digested collagen also led to a settlement response in planulae exposed to the resulting peptide (Hofmann and Brand, 1987).

Cassiopea larval response to a peptidic cue was confirmed with the development of an artificial hexapeptide (GPGGPA), which induced high rates of settlement and metamorphosis (Fleck, 1998;Walther and Fleck, 1998). While these studies substantiated the potential involvement of a peptidic cue produced by a marine bacterium in settlement and metamorphosis in Cassiopea spp., the bacterial source of

! 83! ! the natural cue was yet unknown. Fractions extracted from degrading mangrove leaves, the natural substrate of C. xamachana, induced settlement in the planula larvae

(Fleck and Fitt, 1999;Fleck et al., 1999), with one highly inductive fraction containing a 5.8 kDa proline-rich peptide. These finding corroborating the involvement of a natural peptidic cue in C. xamachana settlement and metamorphosis but the identity of the cue is yet to be determined.

In congruence with previous findings, monoculture assays of bacterial isolates from degrading mangrove leaves indicate bacteria are responsible for inducing settlement and metamorphosis in C. xamachana larvae. Divergent bacterial species belonging to multiple phyla were found to induce settlement and metamorphosis to varying degrees, including an isolate belonging to the genus Pseudolateromonas (Fig

3), which has been implicated in settlement and metamorphosis in other invertebrate taxa (Huang et al., 2007;El Gamal et al., 2016). Phylogenetic placement did not seem to preclude inductive capacity of an isolate, as shown for isolate SA412, which was closely related to non-inductive isolates. This was also found for species of the genus

Pseudoalteromonas. This observation has been previously reported for other invertebrate-bacterial interactions (Lau et al., 2003;Freckelton et al., 2017). Analysis of the leaf microbiome did not identify inductive isolates to exist in high abundance, and many were rare, including the highly inductive Pseudoalteromonas MB4-1.

While the genus Pseudoalteromonas is ubiquitous throughout marine environments, they have been shown to be rare in abundance within biofilm (Skovhus et al., 2004).

Despite low OTU abundances of inductive bacteria, larvae may be preferentially settling within close proximity of these taxa on the biofilm (Bacchetti De Gregoris et al., 2012). As we do not know the inductive capacity of the all bacterial members of

! 84! ! the biofilm, it is premature to conclude that inductive taxa are often found in low abundances.

The response of C. xamachana larvae to various bacterial taxa suggests a redundancy in the mechanism controlling larval settlement and metamorphosis.

Larval response to multiple monoculture biofilm suggests A) invertebrate larvae are capable of detecting multiple cues or B) multiple bacterial taxa are capable of producing the settlement and metamorphosis inducer that is specific to the invertebrate taxa. While both scenarios are likely, C. xamachana larvae settled and metamorphosed in response to multiple fractions extracted from degrading mangrove

(Fleck et al., 1999), pointing towards a capacity for C. xamachana to detect multiple cues. Our results suggest a water-soluble cue to be involved in settlement (Fig 5), with larvae responding stronger to the culture filtrate than to the monoculture biofilm.

However, larvae did not undergo metamorphosis when exposed to filtrate, further implicating multiple cues are involved in facilitating the larvae to polyp transition. As larvae did not respond to the cell fraction alone, a two step process is likely required in which settlement is induced by a water soluble cue, followed by a signaling mechanism triggered potentially by direct contact (Leitz, 1997). In order to determine the induction mechanism, we took a bioinformatics approach to identify potential cues for Cassiopea settlement and metamorphosis by comparing the genome of inductive and non-inductive bacteria.

The phage tail-like particles previously identified in Pseudoalteromonas luteoviolacea (Huang et al., 2012;Shikuma et al., 2014) were found in both strains of

P. luteoviolacea but were absent in the other three genomes. Phage tail-like particles

(MAC proteins) have been found in multiple Pseudoalteromonas species, but have only been reported to induce settlement in Hydroides elegans. Presence of the mac

! 85! ! genes cluster is not predictive of settlement induction (Nedved et al., 2017), and C. xamachana larvae do not respond to bacteria carrying the gene cluster. The type VI secretion system cluster (T6SS), homologous to phage-tail proteins (Leiman et al.,

2009), were present in Pseudoalteromonas MB4-1, Pseudoatleromonas HI1, and

Nisaea MA6-4. The type IV secretion system was the only gene cluster specific to settlement and metamorphosis inducing bacteria (Fig. 2, 9). Type IV and VI secretion systems have been well characterized due to their importance to pathogens in exporting virulence factors to the host. These secretion systems function by directly injecting proteins into the host cytosol (Green and Mecsas, 2016), suggesting their potential involvement in inducing settlement of C. xamachana larvae, as larvae may require contact with the biofilm in , addition to a water soluble cue, in order to undergo metamorphosis (Tebben et al. 2001).

We searched the genomes for secreted products in order to identify potential predictors of larval settlement. Searching the genomes for signatures of proline-rich proteins domains using InterProScan produced no hits. However, a Blast search of the genomes produced hits to proteins containing the C-terminal domain of the TonB protein family. TonB proteins contain a transmembrane domain and are embedded within the outer membrane transporters responsible for predominantly trafficking siderophores and vitamin B12 (Chu et al., 2007;Noinaj et al., 2010). TonB-C carrying genes were found in all genomes except for Thalassospira MA6-2, suggesting that it may not be involved in settlement and metamorphosis of Cassiopea larvae. In

Pseudoalteromonas tunicata, a TonB-dependent outer-membrane transporter was simultaneously up-regulated with antifouling compounds and biofilm formation factors under various nutrient conditions (Stelzer et al., 2006;Bowman, 2007).

! 86! !

Therefore, nutrient availability may correlate with metabolite production for some inducers.

We searched for the tetrabromopyrrole biosynthesis cluster (bmp1 - 10) and found most genes present in all genomes. Tetrabromopyrrole is a common moiety found in biologically active bacterial product and has been implicated in settlement of scleractinian coral larvae (Tebben et al., 2011;Sneed et al., 2014;El Gamal et al., 2016).

Although genes belonging to the cluster were found in all genomes, only the

Pseudoalteromonas HI1 genome possessed the cluster organized in tandem. This may suggest tetrabromopyrrole may not be a cue for C. xamachana larvae, but this finding cannot preclude the absence of tetrabromopyrrole related metabolites in settlement inducing larvae.

We predicted secondary metabolite clusters for all genomes, which resulted in identification of 14 clusters for Pseudoalteromonas MB41, 21 for Nisaea MA64, 33 for Pseudoalteromonas HI1, and over 46 for both Pseudoalteromonas 6061 and

Thalassospira MA62 (Table S1-S5). The lack of completeness for our assemblies resulted may have resulted in low power to predict secondary metabolites within the newly assembled genomes, but a greater number of secondary metabolite clusters were detected for non-inductive bacteria. While many of the clusters remain putative in their function, we identified one cluster to be specific in inductive bacteria (Table S1) and annotated as a aryl polyene synthesizing cluster. Aryl polyenes are a class of pigment molecules related to carotenoids and are commonly found in gram-negative bacteria

(Schoner et al., 2016). These molecules antioxidative properties and likely play an important role in the biology of gram-negative bacteria (Cimermancic et al., 2014) but their has not been a link between these compounds and settlement induction of marine larvae. Antimicrobial and antifungal synthesizing gene clusters (bacteriocin, aryl

! 87! ! polyene, and polyketide synthase) were predominant. The Thalassospira MA6-2 genome lacked many of the gene clusters found in the other genomes, but contained three separate terpene-synthesizing clusters. Terpenes produced by the red algae

Sphaerococcus coronopifolius have been shown to inhibit settlement of barnacle larvae (Piazza et al., 2011). P. luteoviolacea genomes also possess the violacein gene cluster, a bacterial pigment with antimicrobial activity and can be toxic to marine invertebrates at high concentrations (Stelzer et al., 2006). Pseudoalteromonas MB4-1 belongs to the non-pigmented group within the genus and loss of pigmentation has been associated with reduction in antifouling activity of mutant strains of pigmented species (Egan et al., 2002;Huang et al., 2011).

Although further research would be necessary, larvae of C. xamachana may associate with bacterial taxa based not only the presence of an inductive cue but also the absence of potentially harmful secondary metabolites. C. xamachana responds to monoculture biofilm but the response to filtrate may suggest multiple cues are required to facilitate the complete process of settlement and metamorphosis. Comparing genomes of non-inductive and inductive bacteria revealed few candidates that may explain the larval response to monoculture biofilm. Both Pseudoalteromonas MB4-1 and Nisaea MA6-4 possessed the gene cluster for secretion system IV, which could be a target of future studies to determine their role in larval metamorphosis, in addition to the aryl polyene synthesizing cluster. The combinatorial effect of an inductive cue and inhibitory compounds likely played an important role in larval settlement preference and animal life history.

! 88! !

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! 98! !

500 m

10 km

500 m

Figure!1.!Mangrove!leaf!sampling!locations!across!the!Florida!Keys!between!2012! and!2014.!Four!sites!were!chosen!for!collection!in!Key!Largo!(hexagon!=!Mangrove! Island,!triangle!=!dock,!square!=!the!cove,!circle!=!North!Bay)!and!a!single!site!on! Crawl!Key!(star!=!quarry).!!

! 99! !

Figure!2.!Larval!settlement!in!response!to!monoculture!biofilm!grown!from! bacterial!isolated!from!degrading!manrove!leaves.!C!=!0.2!μm!filtered!seawater! negative!control.!C2!=!marine!broth!negative!control.!C3!=!settlement!inducing! artificial!hexapeptide!(GPGGPA)!positive!control.!Statistical!significance!was! calculated!*!=!p8value!

! 100! !

MB4-2 98 MB4-2b 93 MB10-1 100 Agarivorans gilvus Agarivorans albus SA4-10

100 Alteromonas macleodii 100 Alteromonas alvinellae

78 Pseudoalteromonas luteoviolace 87 Pseudoalteromonas sp. H1A 99 SA2-2

100 MB8-1 Pseudoalteromonas donghaensis 74 73 MB4-1 81 MB4-7 MB6-2

92 Vibrio fischeri str. ES114 Gammaproteobacteria 99 Vibrio shiloni 55 MB1-2 100 Mangrovibacter sp.

100 MB1-1 100 MB1-1b Pseudomonas syringae SA4-12b

93 Salinicola salarius 59 96 MA8-1 Salinicola sp. Halomonas halmophila 99 Halomonas halocynthiae SA6-4 82 75 MA4-2 SA10-5 100 MB4-9 SA6-6 SA4-1 SA4-12 54 SB10-2

100 Erythrobacter flavus SB10-2b Erythrobacter longus

94 Nisaea denitrificans 100 Nisaea nitritireducens 97 MA6-4 MB4-3 99 Thalassospira lucentensis type strain Thalassospira profundimaris 100 96 MA6-2 87 MA8-2 Alphaproteobacteria

100 MA6-3 53 90 MA8-3 63 Stappia stellulata SB10-3

59 MA4-5 MB6-1 98 MA4-6 100 Paracoccus denitrificans SB10-4 78 81 Paracoccus carotinifaciens 78 SA4-5

99 Exiguobacterium sibiricum 97 Exiguobacterium antarcticum SA4-6

70 SA10-11 100 SA8-1 100 Bacillus sp. Bacillus subtilis lactipan

98 SB4-1 Bacillus cereus

76 Pontibacillus chungwhensis Bacilli 66 99 Pontibacillus litoralis SA4-15

91 Halobacillus trueperi 89 SA8-2 Halobacillus sp. 97 SA10-1 98 SA4-4 SA4-8 Euzebyella saccharophyla Flavobacteria 100 SA4-13 Methanocaldococcus jannaschii Aquifex aeolicus

! 101! !

Figure!3.!Phylogenetic!analysis!of!full!length!16S!rRNA!gene!of!bateria!isolated! from!degrading!mangrove!leaves.!16S!sequences!were!aligned!in!MEGA!using! MUSCLE!and!phylogenetic!relationships!were!inferred!using!maximum!likelihood!! (Tamura8Nei)!with!a!bootstrap!of!500.!Sequences!of!bacteria!that!were!not! isolated!in!the!current!study!were!retrieved!from!genbank.!Bacterial!isolates!are! colored!based!on!inductive!capacity!determined!with!the!monoculture!bioassays.! Blue!=!weakly!or!non8inductive!isolates.!Green!=!moderately!inductive!isoaltes.! Red!=!highly!inductive!isolates.!Black!=!untested.!Scale!bar!=!number!of!nucletoide! substitutions!per!site.!

! 102! !

62 P atlantica IAM 12927T X82134 54 P espejiana NCIMB 2127T X82143

32 P agarovorans KMM 255 AJ417594 P distincta AF043742

39 40 P sp. KMM 3548 AY040229 46 P elyakovii AF082562 P carrageenovora IAM 12662T X82136 19 60 P issachenkonii strain KMM 3549 AF316144 60 P tetraodonis strain IAM 14160 AF214730 P undina NCIMB 2128T X82140 58 20 P nigrifaciens NCIMB 8614T X82146 77 P haloplanktis X67024.1 P marina strain mano4 AY563031

99 P arctica strain A 37-1-2 DQ787199.1 19 P translucida KMM 520 AY040230 63 78 P antartica X98336 16 99 P antartica X98336(2) P aliena AY387858

99 P mariniglutinosa KMM 3635 AJ507251 P prydzensis strain MB8-11 U85855

30 P byunsanensis strain FR1199 DQ011289 23 78 P lipolytica strain LMEB 39 FJ404721 100 MB4-1

69 P spongiae strain UST010723-006 AY769918 P ruthenica strain KMM300 AF316891 P phenolica O-BC30 AF332880 53 100 P luteoviolacea NCIMB 1893T X82144 94 45 P luteoviolacea HI1 P rubra 16S ATCC 29570T X82147 95 76 P flavipulchra AF297958 95 P maricaloris AF144036 70 100 P piscicida ATCC 15057 X82215 57 P peptidysin AF007286 P aurantia ATCC 33046T X82135 100 P citrea NCIMB 1889T X82137.1 P tunicata strain D2 Z25522 P denitrificans ATCC 43337T X82138 Algicola sagamiensis AB063324

Figure!4.!Phylogenetic!analysis!of!full!length!16S!rRNA!gene!of!the!genus! Pseudoalteromonas,!including!isolate!MB481.!Full!length!16S!sequences!were! retrieved!from!genbank.!Sequences!were!aligned!in!MEGA!using!MUSCLE!and! phylogenetic!relationship!was!inferred!using!maximum!likelihood!(GTRi)!with!a! bootstrap!of!500.!!

! 103! !

5% Filtrate

10% Filtrate

Cell Fraction

ASW MB4-8 SA4-12 MB4-1 MA6-4 SA4-6

Figure!5.!Larval!settlement!frequency!in!response!to!culture!filtrate!from!bacterial! monoculture.!Filtrate!was!added!to!1!ml!of!0.2!μm!filtered!artifical!seawater.!100! μl!of!the!cell!fraction!was!added!to!1!ml!of!artificial!sea!water!and!allowed!to!settle! prior!to!the!start!of!the!experiment.!ASW!=!0.2!μm!filtered!artifical!seawater.!!

! 104! !

Quarry Quarry Cove Dock Mangrove Is. N. Bay Mangrove Is. Degraded Undegraded Degraded Degraded Degraded Degraded Undegraded !

Figure!6.!Taxonomic!identification!of!OTUs!determined!from!the!V4!region!of!the! 16S!rRNA!gene!associated!with!degrading!mangrove!leaves!in!various!states!of! degradation.!OTUs!were!identified!to!the!genus!level!using!Qiime.!!

! 105! !

Figure!7.!PCoA!plot!of!micriobial!biofilm!community!associated!with!degrading! mangrove!leaves!in!various!states!of!degradation.!Purple!triangle!=!quarry,!green! traingles!=!North!Bay,!blue!squares!=!the!dock,!red!triangles!=!the!cove,!orange! circles!=!mangrove!island.!Undegraded!leaves!are!outlined!and!encircled.!!!

! 106! !

MB41 MA64 MA62 HI1 6061 Secretion Proteins T4SS + + T6SS + + + + + + + + MAC proteins MacS putative sheath + MacT1 putative tube + MacT2 putative tube + MacB Baseplate + Tetrabromopyrrole + bmp1 + + + + + bmp2 + + + + bmp3 + + + + + bmp4 + + + + bmp5 + + + bmp6 + bmp7 + + bmp8 + bmp9 + + bmp10 + + + + TonB-C + + + +

Table!1.!Presence!absence!of!genomic!features!between!five!bacterial!species! (Pseudoalteromonas!MB481,!Nisaea(MA684,!Thalassospira(MA682,! Pseudoatleromonas(luteoviolacea(HI1,!and!P.(luteoviolacea(6061).!!!

! 107! !

Chapter 4

Modulation of gene expression driven by symbiosis: Strobilation mechanism in the upside-down jellyfish Cassiopea xamachana

Aki H. Ohdera1, Kelly Watson1, Bailey Steinworth2, , Erika M. Diaz-Almeyda3, William Fitt4, Mark Q. Martindale2, Mónica Medina1,5

1. Department of Biology, Pennsylvania State University, University Park, PA, USA

2. Whitney Laboratory for Marine Bioscience, University of Florida, St. Augustine, FL, USA

3. Odum School of Ecology, University of Georgia, Athens, GA, USA

4. Department of Biology, Emory University, Atlanta, GA, USA

5. Smithsonian Tropical Research Institute, Smithsonian Institution, Panama City PL, Washington D.C. USA

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Abstract

Similar to scleractinian corals, the!upside!jellyfish!Cassiopea)xamachana! forms!a!symbiotic!association!with!the!dinoflagellate!algae!Symbiodinium.!Both! host!and!symbiont!exchange!nutrients!in!this!mutualistic!symbiosis,!but!the!host! also!require!Symbiodinium!in!order!to!proceed!through!its!lifecycle.!Strobilation,! the!developmental!transition!by!which!the!sessile!polyp!undergoes! metamorphosis!in!order!to!generate!the!adult!form,!is!typically!triggered!by! environmental!cues!in!nonMsymbiotic!scyphozoans.!However,!in!Cassiopea,)the! transition!is!initiated!approximately!2!weeks!postMcolonization!by!Symbiodinium.!

While!previous!research!implicates!the!retinoic!acid!pathway!to!be!directly! involved!in!mediated!strobilation!in!Aurelia)aurita,!we!have!yet!to!understand! the!mechanisms!involved!in!initiating!strobilation!in!Cassiopea.!Here,!we!use!an!

RNAseq!approach!to!understand!the!genes!involved!in!symbiosisMmediated! development.!We!find!that!genes!in!the!retinoic!acid!pathway!to!be!constitutively! expressed!in!C.)xamachana.!Visualization!of!mRNA!localization!shows!these! genes!to!be!apically!expressed!in!the!polyp,!thereby!potentially!mediating!the! developmental!transition.!Further!examination!of!differentially!expressed!genes! identified!a!gene!responsible!for!conversion!of!betaMcarotene!to!retinoic!acid,! which!has!been!shown!to!induce!strobilation!in!scyphozoans.!This!suggests!

Symbiodinium!triggers!strobilation!in!Cassiopea)by!providing!a!source!of!betaM carotene.!This!highlights!the!evolutionary!implications!of!symbiosis!in!mediating! the!modification!of!host!gene!expression!control!of!existing!pathways.!!

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Introduction

As we continue to understand the intricacies of animal microbe symbiosis, there is increasing evidence of the importance microbes play in basic animal biology

(McFall-Ngai, 2002;McFall-Ngai et al., 2013). Symbiotic microbes can affect host nutrition, development, and reproduction, to name a few. Symbiotic microbes may have played an important role in metazoan evolution. Examples of symbiosis driving development exist across diverse metazoan groups. In the squid-vibrio symbiosis, bacterial infection leads to dynamic changes to the host light-organ, resulting in irreversible changes to organ structure (Montgomery and McFall-Ngai, 1995;McFall-

Ngai, 2014). The development of the symbiont housing trophosome of vestimentiferan tubeworms is initiated by acquisition of environmental bacteria into the midgut (multiple papers from yesterday). While development of structures post- colonization is not typical of symbiosis, differentiation of host tissue can be observed in a diverse array of interactions, including mammals (below). More dramatic changes to host morphology can be observed in induction of settlement and metamorphosis in marine larvae (Hadfield 2011). However, complete metamorphosis induced by endosymbionts is rare. One example is the upside-down jellyfish Cassiopea xamachana, in which strobilation (Fig. 1) is induced by a dinoflagellate

Symbiodinium spp. Symbiodinium is commonly associated with scleractinian coral, which require the dinoflagellate as a nutrient source (Trench, 1971;Tremblay et al.,

2012). While Cassiopea as adults may potentially rely entirely on Symbiodinium as a food source, they also require the symbiont in order to proceed through its lifecycle and reach sexual maturity (Hofmann et al., 1996).

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Species in the class scyphozoa are characterized by strobilation, a process in which sessile polyps begin a metamorphic process marked by regression of tentacles, following by constriction of the calyx (Gohar and Eisawy, 1960). Lappets and rhopalia begin to develop from the margins of the developing ephyra (Berrill 1949,

Arai 1997). Pulsation is observed before fission is complete, releasing the ephyra from the basal polyp. Environmental factors as a trigger appears to be common across scyphozoans (Purcell et al., 2012;Feng et al., 2018). Strobilation in scyphozoans is controlled by environmental factors including temperature, light, and nutrition but it can also be induced artificially by iodinated compounds, and polypeptides (Fuchs et al., 2014;Cabrales-Arellano et al., 2016;Yamamori et al., 2017). Natural strobilation in Aurelia aurita is triggered by a change in the surrounding seawater temperature in a reversible manner; thereby a return of the temperature to normal condition within 6 days prevents strobilation (Spangenberg, 1965;Fuchs et al., 2014;Sukhoputova and

Kraus, 2017). Beyond 6 days, the process of strobilation becomes irreversible. These exogenous cues initiate a signaling cascade, beginning with release of neurosecretory compounds from neurons (Crawford and Webb 1972). In A. aurita, the neuronal signal leads to differential expression of genes involved in the retinoic acid pathway

(RxR, RDH1, RDH2) as well as genes encoding potentially scyphozoan specific proteins (CL390, CL112, CL631)(Fuchs et al. 2014). The involvement of the retinoic acid pathway suggests evolutionary conservation of metamorphic signaling, as the retinoic acid pathway plays an important role in metamorphosis of both insects and amphibians. In insects, the retinoic x receptor (RxR) homologue ultraspiralce (USP) forms a heterodimer with the ecdysone bound ecydosone nuclear receptor (EcR) and initiates downstream transcription to trigger metamorphosis (Buszczak and Segraves,

2000;Kam et al., 2012). A hetermodimer formed with RxR and thyroid hormone

! 111! ! receptor is also involved in amphibian metamorphosis (Tata, 2006), demonstrating the conservation of the retinoic acid pathway in metazoan metamorphosis. While its role in invertebrates is less characterized, it has been suggested to coordinate tissue regeneration and development in Drosophila (Halme A, Cheng M, & Hariharan I

2010).

Unlike most scyphozoans, some members of the Rhizostomae are triggered to strobilate post-colonization by an endosymbiotic dinoflagellate algae Symbiodinium

(Sugiura, 1964;Rahat and Adar, 1980). In Cassiopea, colonization can be divided into a series of stages, beginning with phagocytosis of the dinoflagellate cells.

Phagocytosis is followed by differentiation and entry of the symbiont housing phagocytic cell into the mesoglea, and subsequent migration of the mobile cell to sub- epithelial locations around the animal body. Approximately 2-3 weeks after infection, the scyphistomae will begin strobilation (Colley and Trench 1983, 1985). The scyphistomae will fail to undergo strobilation in an aposymbiotic state (Hofmann et al. 1978), with only one previously reported case of strobilation by uninfected scyphistomae (Rahat and Adar 1980). As the control of strobilation initiation in

Cassiopea involves successful infection by its symbiont, understanding the molecular mechanism can provide insights how symbiosis can drive the evolution of developmental processes.

Materials and Methods

Artificial Induction of Strobilation

Stock solutions of 1 mM retinoic acid and 50 mM 5-methoxy-2-methylindole were made in DMSO. Solutions were added to 2 ml of 0.2 µM filtered artificial seawater in a 24-well plate for a final concentration of 1 µM and 50 µM of 9-cis retinoic acid and 5 methoxy-2-methylindole, respectively. A single polyp (N=16) was

! 112! ! added to each well, and strobilation was monitored over a course of 5 days at 26°C.

The seawater in each well was replaced daily with freshly made treated seawater.

Strobilation rates in response to Symbiodinium were monitored over a course of 19 days. Polyps were exposed to 100,000 cells with simultaneous feeding of Artemia in a single vessel. Polyps were exposed to Symbiodinium for 24 hrs and transferred to individual wells with 0.2 µm filtered artificial seawater. Polyps were maintained at

100 µmol m-2 s-1 of 12:12 light-dark cycle and fed twice per week for the duration of the experiment. Strobilation was determined as complete upon release of the ephyra from the polyp calyx.

RNAseq of colonization and strobilation

Aposymbiotic scyphistomae from 3 clonal lines (T1-A, T1-B, and T2-B) were colonized with Symbiodinium microadriaticum (CladeA1 - strain KB8).

Symbiodinium cells were provided to Artemia at a concentration of 8.025 x 106 cells prior to being fed to polyps. Polyps were allowed to feed for 2 hrs prior to being transferred to clean filtered artificial seawater. Polyps were maintained at 26° C with

100 µmol m-2 s-1 12:12 light-dark cycle, with feeding every other day except on days for sampling. A subset (at least 15 polyps) were collected on each sampling day and immediately preserved in RNAlater for subsequent RNA extraction. For aposymbiotic polyps, a set of 50 polyps was used to isolate enough totalRNA for downstream processing. Polyps were collected 3 days and 8 days post-colonization as well as during mid-strobilation when the tentacles have regressed and the calyx has shown evidence of undergoing constriction. Strobilae with pulsating ephyra were not sampled.

RNA was extracted using CTAB phenol-chloroform protocol. Polyps were homogenized with a bead beater using 250 µl of 0.1 and 0.5 mm zirconia beads in

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Qiazol. Chloroform was added directly to the bead tube prior to proceeding through the extraction protocol. RNA concentration was determined using the Qubit 2.0

(ThermoFisher Scientific) and quality assessed using the Bioanalyzer (Agilent) with the RNA 6000 Nano kit and NanoDrop 1000 spectrophotometer (ThermoFisher

Scientific). Extracted totalRNA was preserved at -80°C until further processing. The mRNA was isolated from the totalRNA using the Dynabeads mRNA purification kit

(Ambion). Purified mRNA was used to construct the sequencing libraries following the JGI's custom Illumina stranded Truseq library protocol with an insert size of 250 bp. 150 bp SE sequencing of the 12 completed libraries was done on the HiSeq2000 at the Pennsylvania State University Genomics Core Facility.

RNAseq Analysis

Trimming and adapter removal was done with Trimmomatic-0.36 (Bolger et al., 2014) and de novo assembly was conducted using Trinity v2.4.0 (Grabherr et al.,

2011) using standard settings. Contaminating Symbiodinium and bacterial reads were removed from the assembled transcriptome using Alien_Index

(https://github.com/josephryan/alien_index) with the standard bacterial database, and the Symbiodinium microadriaticum (KB8) genome (Aranda et al., 2016). The transcriptome was scaffolded to generate non-redundant transcripts using TransPS

(Liu et al., 2014). Reads were mapped to the de novo Cassiopea assembly as well as the S. microadriaticum reference transcriptome (http://smic.reefgenomics.org/) using bowtie2 (Langmead and Salzberg 2012) and format conversion was completed with

SAMtools (Li et al., 2009). Differential expression analysis of the host and symbiont was conducted using DESeq2 (Love et al., 2014). Transcripts with less than 12 reads across all samples were removed and the Wald test was selected for differential expression analysis. Differentially expressed genes for each time point were identified

! 114! ! via pairwise contrasts between the apo-symbiotic state (apo v 3 days, apo v 8 days, apo v strobila). Differentially expressed host genes were annotated using blastx

(Camacho et al., 2009) against the swissprot database (Bairoch and Apweiler, 1999).

Annotations for S. microadriaticum genes were pulled from those accompanying the gene models. KEGG pathway enrichment was performed using clusterProfiler (Yu et al., 2012).

In Situ Hybridization

Probes were constructed using primers listed in Table S3. cDNA was synthesized using RNA isolated from adult gonadal tissue using the Advantage RT for PCR kit (Clontech). Amplified target genes were ligated into plasmids using the pGEM-T cloning kit (Promega) cloned into Dh-5 competent E. coli cells. Successful cloning was confirmed via colony PCR and plasmids were extracted using the Hi-

Speed Mini Plasmid Kit (IBI). Target sequences were amplified using T7 and SP6 primers and purified from a gel using the QiaQuick extraction kit (Qiagen). Final riboprobes were synthesized using the Megascript Sp6 and T7 Transcription kit

(Thermo Fisher Scientific). Polyps were collected immediately preceding the start of the in situ hybridization protocol. Whole mount in situ hybridization was conducted following the methods described in Wolenski et al. (2013). Polyps were incubated in proteinase K for 25 minutes at a final concentration of 0.01 mg/ml. Samples were incubated in pre-hybridization buffer overnight at 60° C. Samples were hybridized in the probe at a concentration of 1 ng/ml for 48 hrs at 60° C. Blocking of samples was carried out at 4° C for 1 hr prior to an overnight incubation in the anti-DIG, AP Fab fragments at 4° C.

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Results

Artificial Induction of Strobilation

Cassiopea showed a strong response to the indole derivative 5-methoxy-2- methylindole, with 100 % of apo-symbiotic polyps strobilating within 4 days of treatment (Figure 1) while approximately 70% of infected polyps strobilated within 3 weeks of colonization by S. microadriaticum. Surprisingly, strobilation by 9-cis retinoic acid only occurred in an average of 10% of treated polyps over a 7-day period.

Cassiopea Differential Gene Expression An average of 25.61% of reads mapped back to the host transcriptome assembly while an average of 13.43% of reads mapped to the S. microadriaticum reference transcriptome. Differential expression analysis with DESeq2 identified 2074 unique genes (7.1%) to be up- or down-regulated across all time points. Expression between aposymbiotic polyps and strobila showed the largest number of differential expression with 971 up-regulated genes and 1231 down-regulated genes.

Aposymbiotic polyps compared to 3 days (791 up, 869 down) and 8 days (651 up,

781 down) post-colonization showed similar numbers of differentially expressed genes. Gene expression profiles between 3 days and 8 days post-colonization did not show any differential expression and clustered together (Figure 2). A blastx search of the transcriptome yielded high similarity hits to likely homologues of genes belonging to the retinoic acid pathway. However, none of the genes belonging to the retinoic acid pathway (RxR, RDH1, RDH2) were differentially expressed throughout each stage. Examination of normalized log2 expression revealed these genes to be constitutively expressed across the four time points (Figure 3). Of the 3 putative scyphozoan/Aurelia specific genes, only CL112 was found in the Cassiopea

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transcriptome with high confidence. CL112 was down-regulated 2.2 fold (log2) between the aposymbiotic and strobila stage, but did not show differential expression

3 days and 8 days post-colonization. Whole mount in situ hybridization with probes against RxR and CL112 revealed transcripts to be localized to the apical half of the polyp in during aposymbiotic and the early/late strobila polyp stages (Figure 4).

Expression was particularly prominent in the tentacles in the aposymbiotic polyps, which remained in the early strobila, with a moderate expansion of expression beyond the tentacle margins. Expression was markedly reduced in the late strobila, with expression localizing to the degenerating tentacle tips and the developing hypostome margins (Figure 4).

We surveyed for patterns in DEGs across the stages and identified genes with increasing expression across the time points (Table S1). Beta, beta-carotene 9'10'- dioxygenase, a gene involved in retinoic acid synthesis, showed constitutive expression during the first three time points but increased 3.71 fold (log2) during strobilation (Figure 4). With the discovery of an up-regulated Beta, beta-carotene

9'10'-dioxygenase, we searched for genes involved in the production of rhodopsin and found calnexin to be down-regulated at 8 days post-colonization (-1.17 fold) and during strobilation (-1.28 fold). Calnexin, in conjunction with 2 other proteins, are responsible for proper folding of rhodopsin (Xiong and Bellen, 2013). Most genes showing steady up-regulation were unannotated. We performed KEGG over- representation analysis in order to determine if any pathways are over-represented throughout the stages. We found stage specific over-representation of KEGG pathways, particularly for those associated with development during strobilation (Wnt signaling pathway, focal adhesion, regulation of actin cytoskeleton, etc) (Figure 5).

Two pathways (ribosome, protein processing in endoplasmic reticulum, pathways in

! 117! ! cancer), in particular were over-represented throughout all three stages. Several of the genes in these pathways were related to canonical developmental signaling, regulation of apoptosis, and notch signaling.

Symbiodinium Differential Gene Expression

4905 genes were found to be up-regulated between the aposymbiotic stage and strobila stage. Interestingly, only 390 S. microadriaticum genes were active at day 3 post-colonization, while less than half were expressed 8 days post-colonization (140 genes). All 140 genes differentially expressed at 8 days-post colonization were found to be expressed at 3 days post-colonization and strobila stage (Figure 5b, Table S2). 3 unique genes were differentially expressed at 3 days post-colonization, while 4518 genes were unique to the strobila stage. Gene annotations suggest most of the genes found to be expressed across all three stages were associated with photosynthetic processes and nutrient exchange (Table S2). Three genes were unique to 3 days post- colonization time point, but annotations were unavailable for these. Summarization of the expressed genes by KEGG over-representation analysis showed no pathways were over-represented across all stages (Figure 7). However, pathways specific to 3 days post-colonization and strobilation suggest potential gene expression responsible for mediating the symbiosis but we were unable to identify genes potentially involved in mediating strobilation in C. xamachana.

Discussion

Metamorphic transitions are a common life history trait among invertebrates, as exemplified by transition from planktonic larvae to the juvenile stage or the dynamic transition between pupa and adult in insects. Metamorphosis can also be observed in vertebrates, including amphibians and fish (Buszczak and Segraves,

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2000;Tata, 2006). Initiation of these transitions is controlled in part by genes involved in the retinoic acid pathway. Homologues of the retinoic X receptor and retinoic acid receptor form heterodimers in order to bind to DNA, controlling downstream gene expression that will initiate the metamorphic signaling cascade. Interestingly, retinoic acid genes appear to be involved in mediating the larvae to juvenile transition in true jellyfish and sea stars (Fuchs et al., 2014;Yamakawa et al., 2018), indicating their involvement in metamorphosis early in evolution. The retinoic acid pathway was found to be involved in initiating strobilation in the non-symbiotic scyphozoan

Aurelia aurita (Fuchs et al., 2014;Brekhman et al., 2015). An investigation into the kinetics of strobilation in Cassiopea revealed some differences between the two species. While Cassiopea does not respond strongly to 9 cis-retinoic acid in the same manner as Aurelia, exposure to 5-methoxy-2-methylindole induces strobilation for both. These findings potentially suggest parallels exist between the two species.

Differential expression analysis of genes expressed between aposymbiotic, 3 days and 8 days post colonization, and strobilation revealed genes involved in the retinoic acid pathway to be constitutively expressed in C. xamachana. Given the importance of RxR signaling in metamorphosis, and its involvement in Aurelia strobilation, it is likely that RxR is involved in strobilation in Cassiopea. RxR and

RAR expression in A. aurita and the sea star Patiria pectinifera shows a clear increase in gene expression as the larvae approaches metamorphosis (Fuchs et al.,

2014;Yamakawa et al., 2018). In addition to RxR, both RDH1 and RDH2 are constitutively expressed through the four stages. Both RxR and CL112 expression overlapped with the apical portion of the polyp, particularly throughout the tentacles.

This expression remains unchanged until the late-strobila stage, in which the signal becomes localized to the degenerating tentacle tips. As in A. aurita and P. pectinifera,

! 119! ! expression of RxR and other metamorphosis-related genes appear to gradually increase, but demonstrating polarity in expression. In A. aurita, the putative Aurelia hormones are often expressed in the apical portion of the metamorphic polyp, similar to what is observed in C. xamachana. These data suggest C. xamachana may have evolved a primed co-expression of strobilation genes as a result of evolving a symbiosis with Symbiodinium. The gene encoding a beta-carotene 9'10'-dioxygenase, an enzyme responsible for catalyzing the reaction to generate retinal, a precursor to retinoic acid, was highly up-regulated during strobilation. This aligns with the involvement of the retinoic acid pathway in C. xamachana strobilation. While C. xamachana did not show a strong response to retinoic acid, this does not preclude the non-involvement of the ligand in Cassiopea strobilation as 9 cis-retinoic acid triggers strobilation in Aurelia, there may be additional requirement in Cassiopea that inhibits activation of strobilation in the presence of the RxR ligand. Alternatively, strobilation initiation in C. xamachana may require a different form of retinoic acid, which may explain the low frequency of strobilation in response to the treatment with exogenous retinoic acid. These data, however, suggest that Symbiodinium is potentially the source of the ligand cue, providing the host with the essential nutrient β-carotene, which is then converted to retinoic acid. We also confirmed the gene responsible for rhodopsin synthesis to be non-differentially expressed. Rhodopsin expression has been found to be up-regulated in the eyes of the cubozoan Alatina alata (Lewis Ames et al., 2016). Chaperone genes implicated in folding of rhodopsin (NinaA, calnexin, and XPORT) to facilitate maturation of the protein were found in the transcriptome excluding XPORT (Xiong and Bellen, 2013). While NinaA (peptidyl-prolyl cis-trans isomerase) was not differentially expressed, calnexin was down regulated at 8 days and during strobilation, potentially suggesting the up-regulation of beta, beta-carotene

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9'10'-dioxygenase to be unrelated to maturation of rhodopsin. Further experiments will be required to confirm the β-carotene sourced from Symbiodinium to be involved in initiating strobilation in Cassiopea.

Unlike other scyphozoans, C. xamachana is dependent on colonization by

Symbiodinium to initiate strobilation. Thus, the upstream signal likely originates from the establishing population of Symbiodinium. Exploration of the Symbiodinium gene expression confirmed active photosynthetic machinery during symbiosis (Figure 7), in addition to genes belonging to pathways likely involved in nutrient transfer. 54 genes were gradually up-regulated post-colonization (Table S2). Genes that were annotated within this set were also involved in photosynthesis, except for three proteins (major basic nuclear protein 2, ABC transporter G family member, N-(5-amino-5- carboxypentanoyl)-L-cysteinyl-D-valine synthase). The pattern likely reflects

Symbiodinium proliferation within the host polyp over the study period. With the exclusion of unannotated genes, these patterns did not identify potential signaling genes that may be involved in triggering strobilation in C. xamachana.

Strobilation in C. xamachana requires colonization by Symbiodinium in order for the developmental process to be initiated. Strobilation by aposymbiotic polyps was reported by Rahat and Adar (1980), whereby polyps 20% of experimental polyps strobilated in response to exposure to temperatures above 25°C after a 5°C increase in temperature. Although this has not been repeated, we'd like to report a similar occurrence of spontaneous strobilation by aposymbiotic polyps in our culture. This report confirms that aposymbiotic strobilation can occur in C. xamachana (Figure 8).

In light of these observations, we hypothesize the existence of dual mechanism of strobilation in C. xamachana, whereby strobilation occurs predominantly via induction by increase in available retinoic acid and the retinoic acid pathway. A

! 121! !

"basal" or alternative mechanism still exists in Cassiopea where strobilation is triggered via a neural trigger, similar to what is found in non-symbiotic scyphozoan.

These neural triggers may perhaps be related to a stress response, as exemplified by an increase in temperature for strobilation in C. xamachana described above

(Sukhoputova and Kraus, 2017). Thus, strobilation in C. xamachana has evolved as a result of the association with Symbiodinium. While expression associated with strobilation has only been determined for A. aurita and C. xamachana, these data may suggest alterations to strobilation related gene expression and changes to developmental mechanisms brought on by the evolution of symbiosis.

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*** ***

Control 9-cis Retnoic Symbiodinium 5-methoxy Acid 2-methylindole !

Figure!1.!Strobilation!rates!of!Cassiopea(xamachana(polyps!(N=16)!in!response!to! artificial!1!μM!98cis!retinoic!acid,!50!nM!58methoxy828methylindol,!!and!the! natural!strobilation!trigger!(Symbiodinium).!Control!=!0.2!μM!filtered!artificial!sea! water.!Statistical!significance!(***)!was!calculated!with!Tukey's!honest!significant! difference!post(hoc(test!from!the!ANOVA!(p!≤!0.0005)!

! 128! !

Figure!2.!Heat!map!of!the!top!100!differentially!expressed!genes!of!Cassiopea( xamachana(at!4!different!stages:!aposymbiotic!(0!=!light!blue),!3!and!8!days!post8 colonization!(3d!=!light!green!and!8d!=!green),!and!strobila!(strob!=!dark!gold).! Count!data!was!r8log!transformed,!with!hierarchical!clustering!of!samples!labeled! based!on!stage.!Row!z8score!is!represented!with!the!cyan!line.!! !

! 129! !

RxR RDH2-like RDH1-like Log2 Expression

CL112 BCO2 Stage Aposymbiotic Scyphistomae

3 Days Post-Colonization

8 Days Post-Colonization

Log2 Expression Strobila ! !

Figure!3.!Gene!expression!of!genes!belonging!to!the!retinoic!acid!pathway! (retinoic(x(receptor,!retinol(dehydrogenase(18like,!retinol(dehydrogenase!28like,! CL112)!as!well!as!beta8carotene!9'10'8dioxygenase!(BCO2).!!Expression!data!was! calculated!from!raw!read!counts,!with!normalization!and!Log2!expression! performed!with!the!DEseq2!R8package.! !

! 130! !

A RxR

B CL112 Mcol-A RxR/CL112

Aposymbiotc Polyp Early Strobila Late Strobila ! !

Figure!4.!A)!Whole!mount!in!situ!hybridization!visualizing!the!expression!of! retinoic(x(receptor!(RxR),!CL112,!and!miniAcollagen(A!(McolAA;!positive!control)!for! aposymbiotic,!early,!and!late!strobila.!B)!Gene!expression!model!of!strobilation! related!genes!in!Cassiopea!xamachana.!! !

! 131! ! A

Figure!5.!Venn!diagram!showing!shared!differentially!expressed!genes!at!3!days! and!8!days!post8colonization!as!well!as!during!strobilation.!Number!of! differentially!expressed!genes!shared!across!the!different!stages!for!A)!Cassiopea( xamachana!and!B)!Symbiodinium(microadriaticum(

! 132! !

Ribosome (58) Protein processing in endoplasmic reticulum (40) Huntington's disease (22) Parkinson's disease (21) Thermogenesis (25) mTOR signaling pathway (17) Hepatocellular carcinoma (16) Alzheimer's disease (21) Percentage Fluid shear stress and atherosclerosis (8) 0.2 Pathways in cancer (57) Oxidative phosphorylation (13) 0.4 Carbon metabolism (16) 0.6 Cardiac muscle contraction (6) 0.8 Basal cell carcinoma (9) 1.0 Non−alcoholic fatty liver disease (NAFLD) (8) Focal adhesion (12) p.adjust Regulation of actin cytoskeleton (11) Wnt signaling pathway (8) 0.04 Signaling pathways regulating pluripotency of stem cells (8) 0.03 Glutathione metabolism (5) 0.02 Leukocyte transendothelial migration (6) Metabolism of xenobiotics by cytochrome P450 (4) 0.01 Protein digestion and absorption (5) Renal cell carcinoma (5) Insulin signaling pathway (6) Chemical carcinogenesis (4) ECM−receptor interaction (5) Axon guidance (7) Pancreatic secretion (5) Phagosome (6) Day3 Day8 Strobila !

Figure!6.!ClusterProfiler!output!of!over8represented!KEGG!pathways!of!Cassiopea( xamachana.!Differentially!expressed!genes!with!annotations!were!mapped!to! KEGG!IDs!and!analyzed!using!ClusterProfiler!with!a!p8adjusted!cutoff!of!0.05.!Size! of!the!circles!indicates!precentage!of!genes!within!the!KEGG!pathway!found!to!be! differentially!expressed!at!3!days!and!8!days!post8colonization!and!during! strobilation.!Numbers!in!brackets!next!to!KEGG!descriptions!denote!the!number! of!genes!found!to!be!differentially!expressed!belonging!to!the!corresponding! pathway.!! !

! 133! !

Ribosome (49) Biosynthesis of amino acids (37) Carbon metabolism (47) Carbon fixation in photosynthetic organisms (13) Glycolysis / Gluconeogenesis (19) Lysosome (19) Protein processing in endoplasmic reticulum (22) Spliceosome (20)

Oocyte meiosis (15) Percentage Valine, leucine and isoleucine degradation (14) 0.25 Glycine, serine and threonine metabolism (15) 0.50 Fatty acid degradation (11) 0.75 Glyoxylate and dicarboxylate metabolism (15) 1.00 Synaptic vesicle cycle (9) RNA degradation (13) p.adjust

Huntington's disease (19) 0.04 Pentose phosphate pathway (13) 0.03 Peroxisome (12) 0.02 Glucagon signaling pathway (10) 0.01 Glutathione metabolism (9) Hedgehog signaling pathway − fly (7) Hedgehog signaling pathway (1) Renal cell carcinoma (1) Citrate cycle (TCA cycle) (1) Insulin signaling pathway (1) Pyruvate metabolism (1) Carbon fixation pathways in prokaryotes (1) Cushing's syndrome (1) scyV3d scyVs !

Figure!7.!ClusterProfiler!output!of!over8represented!KEGG!pathways!of! Symbiodinium(microadriaticum.!Differentially!expressed!genes!with!annotations! were!mapped!to!KEGG!IDs!and!analyzed!using!ClusterProfiler!with!a!p8adjusted! cutoff!of!0.05.!Size!of!the!circles!indicates!precentage!of!genes!within!the!KEGG! pathway!found!to!be!differentially!expressed!at!3!days!post8colonization!and! during!strobilation.!Numbers!in!brackets!next!to!KEGG!descriptions!denote!the! number!of!genes!found!to!be!differentially!expressed!belonging!to!the! corresponding!pathway.!!

! !

! 134! !

A B

Figure!8.!Strobilation!of!aposymbiotic!polyp!of!Cassiopea(xamachana!occurred!in! select!individuals!from!2012.!A)!Aposymbiotic!strobila!B)!40x!magnification! bright!field!image!of!an!aposymbiotic!C.(xamachana!ephyra!C)!40x!magnficiation! bright!field!image!of!a!symbiotic!C.(xamachana!ephyra.!!

! 135! !

Chapter 5

Box, stalked and upside-down? Draft genomes from diverse jellyfish (Cnidaria, Acraspeda) lineages: Alatina alata (Cubozoa), Calvadosia cruxmelitensis (Staurozoa), and Cassiopea xamachana (Scyphozoa)

Aki Ohdera1, Cheryl L. Ames2,3,4, Rebecca B. Dikow2, Ehsan Kayal2, Marta Chiodin3, Ben Busby4, Sean La4,5, Stacy Pirro6, Allen G. Collins2, Mónica Medina1*, Joseph F. Ryan3*

1. Department of Biology, Pennsylvania State University, University Park, PA, USA

2. Department of Invertebrate Zoology, National Museum of Natural History, Smithsonian Institution, Washington D.C., USA

3. Whitney Laboratory for Marine Bioscience, University of Florida, St. Augustine, FL, USA

4. National Center for Biotechnology Information, Bethesda, MD, USA

5. Department of Mathematics, Simon Fraser University, British Columbia, BC, Canada

6. Iridian Genomes, Inc. Bethesda, MD, USA

7. UPMC, CNRS, FR2424, ABiMS, Station Biologique Roscoff, France

8. National Systematics Laboratory of NOAA's Fisheries Service, Washington, DC, USA

* Corresponding authors

! 136! !

Abstract

Anthozoa, Endocnidozoa, and Medusozoa comprise the three major clades of

Cnidaria. Medusozoa is further divided into four clades, Hydrozoa, Staurozoa,

Cubozoa, and Scyphozoa—the latter three lineages make up the clade Acraspeda.

Acraspeda includes some of the most venomous organisms on the planet, numerous nuisance species, some of the most highly developed eyes in the animal kingdom, and it encompasses extraordinary diversity in terms of life history. Currently, no genomes are publicly available for any of these animals. Here we present three new draft genomes of Calvadosia cruxmelitensis (Staurozoa), Alatina alata (Cubozoa), and

Cassiopea xamachana (Scyphozoa) for which we provide preliminary orthology analyses that includes an inventory of their known venom-related genes. To further demonstrate the utility of these datasets, we identify synteny between Pou and Hox genes that had previously been reported in a hydrozoan, suggesting that this linkage is highly conserved, dates back to at least the last common ancestor of Medusozoa, and is likely independent from the Hox-Pou linkages seen in vertebrates. These draft genomes provide a valuable resource for studying the evolutionary history and biology of these extraordinary animals, and for identifying genomic features underlying venom, vision, and life history traits.

Keywords: Staurozoa, Scyphozoa, Cubozoa, Acraspeda, Cnidaria, Medusozoa

! 137! !

Introduction Some of the most fascinating and outstanding mysteries in genome biology are centered around cnidarians. Active areas of research include the basis of venom evolution and diversification (Brinkman!and!Burnell,!2009;Jouiaei!et!al.,!

2015a;Jouiaei!et!al.,!2015b), mechanisms of independent evolution of image- forming vision (lens eyes) (Coates,!2003;Liegertova!et!al.,!2015;Lewis!Ames!et!al.,!

2016), and the emergence of a pelagic adult stage within a biphasic life cycle

(Collins,!2002).!Cnidaria encompasses three major clades: Anthozoa, Endocnidozoa, and Medusozoa. The Endocnidozoa are comprised of the parasitic Myxozoa and

Polypodiozoa were recently accepted as formal members of Cnidaria based on phylogenomic evidence and the evolutionary similarities between the myxozoan polar capsule and nematocysts present in all cnidarians (Chang et al., 2015). Anthozoans are represented by the Hexacorallia, which include scleractinian corals, anemones, and zooanthids. The group is characterized by a six-fold symmetry, with species exhibiting both colonial and solitary forms. The Octocorallia sea fans, gorgonians, and soft corals and are characterized by pinnate tentacles in 8-fold symmtery. While all known anthozoans are sessile, medusozoans are characterized by the emergence of a medusa life history stage within some taxa of the clade, their high diversity in regards to life history and morphology, the presence of a linear mitochondrial genome

(with a variable number of chromosomes), and by the presence of a hinged cap at the apex of the cnidocyst (cnidarian stinging organelles) (Bridge!et!al.,!1992;Collins,!

2002;Reft!and!Daly,!2012)(Bridge!et!al.!1992;!Collins!2002;!Reft!and!Daly!2012).

There are approximately 3900 described species within Medusozoa, classified into four diverse lineages: Hydrozoa (hydroids, hydromedusae, siphonophores),

Staurozoa (stalked jellyfish), Cubozoa (box jellyfish), and Scyphozoa (true jellyfish)

(Figure 1A-C). There exists much debate regarding the phylogenetic relationships

! 138! ! among these lineages (von!SalviniMPlawen,!1978;Bridge!et!al.,!1992;Marques!and!

Collins,!2004;Ortman!et!al.,!2010;Miranda!et!al.,!2016). Recent phylogenomic analyses have placed Staurozoa as the sister to a clade that contains Cubozoa and

Scyphozoa, uniting these lineages in a group called Acraspeda (Figure 1D) (Kayal!et! al.,!2013;Zapata!et!al.,!2015;Kayal!et!al.,!2017). Given the high morphological diversity within Cnidaria, understanding the evolutionary relationships and mechanisms leading to lineage specific innovations has been difficult. In particular, the evolution and subsequent loss of the medusoid form in some lineages hints at a complex evolutionary history within the Medusozoa (Cartwright and Nawrocki,

2010).

The mechanisms of medusa formation are variable amongst medusozoans: often involving two phenotypically distinct life stages - polyp and medusa - that are genotypically identical. Cubozoan polyps undergo partial or complete metamorphosis and develop into the adult medusoid form capable of sexual reproduction, although in some cases a polyp rudiment remains (Toshino!et!al.,!2015).!Scyphozoan polyps

(scyphistomae) undergo a transition known as strobilation, in which the upper calyx proceeds through metamorphosis and transverse fission to produce a medusa (Helm,!

2018).!Unlike other medusozoans, staurozoans lack a free-swimming medusa stage but exhibit medusa-associated characters that are present in other medusozoans. The basal portion of the adult forms a stalk, or peduncle, while coronal muscles and gastric filaments, among other features, characterizes the apical portion (calyx) of the adult (Kikinger!and!von!SalviniMPlawen,!2009;Miranda!et!al.,!2017). Hydrozoans exhibit the greatest variation in life history strategies and often lack a medusa form.

Species that give rise to the medusoid form do so via lateral buds generated by asexual polyps, while others possess sexual polyps without a free-swimming stage

! 139! !

(Boero!et!al.,!1997;Bentlage!et!al.,!2018). Elsewhere within Cnidaria, Anthozoa and the parasitic Endocnidozoa lack the medusa stage or medusoid characters entirely.

Research on medusa development has shown similar gene expression patterns between hydrozoans and scyphozoans, with pre-existing developmental genes co- opted for patterning the medusa body plan (ReberMMuller!et!al.,!2006;Kraus!et!al.,!

2015). Interestingly, strobilation in scyphozoans was recently shown to be under the control of the retinoic acid pathway (Fuchs!et!al.,!2014;Brekhman!et!al.,!2015).

These same genes are involved in metamorphosis of insects and amphibians, hinting that the conservation of metamorphosis regulation might be conserved in metazoans.

The study also found potential lineage specific genes to be involved in controlling strobilation, suggestive of genomic innovations playing a role in medusa morphogenesis within Medusozoa, or perhaps within Scyphozoa.

The genomic resources necessary to understand medusozoan evolution have been lacking, with genomes currently available only for cnidarian models including the anthozoan Nematostella vectensis and the freshwater hydrozoan Hydra (Putnam! et!al.,!2007;Chapman!et!al.,!2010;Shinzato!et!al.,!2011). While the majority of

Medusozoa species are represented by hydrozoans (>90%), both cubozoans and scyphozoans garner significant attention as a result of their impact on economy and tourism (Nastav!et!al.,!2013). Largely due to venom being employed as a mechanisms of defense and prey capture, the inherent risk of jellyfish sting has been exacerbated by uncertainty about how cnidarians will respond to modern-day anthropogenic disturbances along coastal environments (Purcell!and!Arai,!

2001;Purcell!et!al.,!2007). In addition, there has been increased interest in cnidarian venom for their pharmacological applications (Jha!and!ZiMrong,!2004).

! 140! !

Here we present three new genomes, each belonging to a species representing one of the three major Acraspeda lineages: Calvadosia cruxmelitensis (formerly

Lucernariopsis cruxmelitensis)(Staurozoa), Alatina alata (Cubozoa), and Cassiopea xamachana (Scyphozoa). Herein, we perform a range of preliminary gene inventory and synteny analyses to show the utility of these new data, with particular attention to

C. xamachana, which is characterized by the presence of the endosymbiotic dinoflagellate Symbiodinium. Similar to species within Anthozoa, species of the order

Rhizostomeae establish an association in which both host and symbiont coordinate nutritional exchange (Dawson and Hamner, 2008). While previously sequenced genomes of symbiotic cnidarians belong to the Anthozoa, the genome of Cassiopea will provide an additional insight into the evolution of symbiosis. Symbiosis with

Symbiodinium likely evolved multiple times within cnidarians, and while modulation of pre-existing pathways likely played a role in allowing the stable association to occur, key genomic innovations may also have played a role. Several examples of horizontal gene transfers from historical microbial associations been identified in cnidarians, incuding enzymes of the shikimic pathway, mycosporine-like amino acids, and a plant-like peroxidase (Habetha and Bosch, 2005;Starcevic et al., 2008;Shinzato et al., 2011).

We utilize the genomes to understand lineage specific innovation that may have played a role in the evolution of the cnidarian classes, and provide a early evidence for diversification of cnidarian venom. The genomes and corresponding gene annotations from these three lineages will serve as useful resources aimed at sparking investigative research into the evolution and diversification of life history strategies across cnidarians, the evolution of venom within Cnidaria and

! 141! ! phylogeographic patterns of venomous jellyfish, potential jellyfish-derived therapeutic drug development, as well as countless additional research programs.

Materials and Methods ! Genome!Assembly!

Cassiopea xamachana Sample Collection and DNA extraction

We propagated C. xamachana polyps from a single polyp via budding (Line

T1-A). Polyps were maintained symbiont-free at 26 °C, and fed 3 times weekly with

Artemia nauplii. To avoid the possibility of food-source contaminates interfering with downstream bioinformatic analysis, we starved the polyps for seven days in antibiotic-treated seawater prior to preservation in 95% ethanol, with any Artemia cysts still retained within the gut being manually removed before preservation. We extracted genomic DNA using a CTAB (cetyl trimethylammonium bromide) phenol chloroform extraction. We performed an overnight digestion of polyps with proteinase K (20 mg/ml) in CTAB buffer before proceeding with the standard protocol. Extract DNA was stored at -20 °C until further processing.

Calvadosia cruxmelitensis Sample Collectioin and DNA extraction

We collected adult specimens of C. cruxmelitensis in January 2013 at

Chimney Rock, off the coast of Penzance, Cornwall, England. Specimens were immediately preserved in ethanol and stored at -20 °C until further processing. We extracted genomic DNA using a phenol-choloroform protocol in an Autogen mass extractor, and stored the DNA at -20 °C.

! 142! !

Alatina alata Sample Collection and DNA extraction

We collected A. alata material during a spermcasting aggregation in Bonaire,

The Netherlands (April, 2014, 22:00-01:00) according to the methods in Ames et al.

(Lewis Ames et al., 2016). We selected a single live spermcasting male medusa from the same cohort as the medusa used for RNA-Seq studies (Genbank Accession:

GEUJ01000000) (Zapata!et!al.,!2015;Lewis!Ames!et!al.,!2016). We divided the medusa into four longitudinal sections, and one quarter was placed into a 15 ml tube with pure (99%) ethanol. We flash froze the tissue samples at -180 °C (using a dry shipper), and subsequently transported them to the Smithsonian NMNH where they were stored at -20 °C until genomic DNA extraction using a DNeasy Blood & Tissue

Kit (Qiagen) following the manufacturer's protocol.

Cassiopea xamachana Sequencing and Assembly

Library construction and sequencing was performed at HudsonAlpha Institute for Biotechnology. Four 350 bp paired-end linear libraries with insert sizes of 500 bp were made with Illumina TruSeq DNA PCR-Free LT Prep Kits and sequenced on the

Illumina Hiseq2000. Approximately 634 million reads totaling 117.6 Gb of high- quality paired-end sequence data were generated. We performed adaptor trimming and quality filtering using Trimmomatic v0.36 (Bolger!et!al.,!2014) with default settings, followed by error correction with Allpaths-LG version 52488 (Gnerre!et!al.,!

2011). We removed mitochondrial reads using FastqSifter v1.1.1

(https://github.com/josephryan/FastqSifter) with the Cassiopea frondosa mitochondrial genome as a reference (NCBI NC_016466.1). We performed de novo genome assemblies using ABySS 2.0.1 with default settings (Simpson!et!al.,!2009),!

SPAdes genome assembler v3.10.0 (Bankevich!et!al.,!2012), and Platanus version

1.2.1 (with default parameters, k=89) (Kajitani!et!al.,!2014) (Table 1). We used a

! 143! ! custom Perl script, plat.pl, (https://github.com/josephryan/Ohdera_et_al_2018) to invoke the Platanus commands for assembly, scaffolding, and gap closing. We generated a draft assembly with 93,483 scaffolds measuring a total of 393.5 Mb with an N50 of 15,563 bp (Table 1) (ENA Accession OLMO01000000). We recovered

82.6% (53.63 % complete and 29.03 partial) of the core eukaryotic genes and 66.97%

(58.59% complete and 8.38% partial) of the core metazoan genes with CEGMA ver.

2.5 (Parra!et!al.,!2007) and BUSCO ver.2.01 (Simao!et!al.,!2015), respectively, through the gVolante web server (Nishimura!et!al.,!2017) (Table 1).

Calvadosia cruxmelitensis Sequencing and Assembly

Library construction and sequencing for C. cruxmelitensis were performed at the University of Florida Interdisciplinary Center for Biotechnology Research. Four

150 bp paired-end linear libraries and four 150 bp single-end linear libraries with insert size of 300 bp were generated and sequenced on the Illumina NextSeq 500. We performed adaptor trimming and quality filtering using Trimmomatic-0.32 (Bolger!et! al.,!2014) with default settings, followed by error correction using Allpaths-LG version (Gnerre!et!al.,!2011). We removed mitochondrial sequences to improve the final assembly with FastqSifter v1.1.1

(https://github.com/josephryan/FastqSifter,https://doi.org/10.5281/zenodo.1211357), using a de novo assembly of the C. cruxmelitensis mitochondrial genome. We assembled the C. cruxmelitensis mitochondrial genome by capturing contigs from an initial assembly using available staurozoan mitochondrial DNA sequences from NCBI as a reference, following the methods presented in Kayal et al. (2015). We used

Geneious v9.0 to generate the final mitochondrial assembly. We checked completeness of the mitochondrial genome using NCBI BLAST against the nr database in addition to a manually generated set of medusozoan genes. We annotated

! 144! ! tRNA genes separately by using tRNAscan-SE (Lowe!and!Eddy,!1997)!and Arwen

(Laslett!and!Canback,!2008). We checked the integrity of the assembly by aligning the reads to the completed mitochondrial genome. With the mitochondrial sequences removed, we generated two "sub-optimal" assemblies using Platanus v1.2.1 with kmer size of 32 bp and 45 bp and default settings. Subsequently, we used these "sub- optimal" assemblies to construct artificial mate-pair libraries for 9 insert sizes (1000,

2000, 3000, 4000, 5000, 7500, 10000, 15000, 20000) with MateMaker

(https://github.com/josephryan/matemaker). We used the artificial mate-pair libraries to scaffold the optimal assembly (generated using Platanus kmer=45) with SSPACE

Standard v3.0 (Boetzer!et!al.,!2011). This process produced a draft assembly with

50,999 scaffolds measuring a total of 209.4 Mb with an N50 of 16,443 bp (Table 1)

(ENA Accession OFHS01000000). We recovered 91.94 % (61.29% complete and

30.65% partial) of the core eukaryotic genes and 85.07% (70.86% complete and

14.21% partial) of the core metazoan genes with CEGMA and BUSCO, respectively.

Alatina alata Sequencing and Assembly

Illumina library prep and sequencing was conducted at the University of

Kansas Genome Sequencing Core. Libraries were generated with the Illumina

Nextera Library Preparation kit and sequenced twice on the Illumina HiSeq 2500. The two different runs were performed on the same library: one with 100 bp paired-end, and one with 150 bp paired-end sequencing, resulting in 564 million reads totaling

148.6 Gb of paired-end sequence data. PacBio library prep and sequencing were completed at the University of Washington Northwest Genomics Center. Libraries were constructed with unsheared DNA with end-cleanup only, and an average insert size of 6000 bp. Sequencing was completed on the PacBio RS II platform, resulting in

486,000 long-reads totaling 990.2 Mb of data. We performed error correction and

! 145! ! subsequent hybrid assembly using Illumina short-reads and PacBio long-reads using

MaSuRCA 3.2.2 (Zimin!et!al.,!2013),!which!results!in!an!assembly!of!291,445 contigs and an N50 of 7,049 bp (NCBI Accession PUGI00000000). The total length of the assembly was 851 Mbp. We recovered 29.84% (8.06% complete and 21.84% partial) of the core eukaryotic genes and 32.11% (18.30% and 13.81% partial) of the core metazoan genes with CEGMA and BUSCO, respectively.

Gene Model Prediction

We predicted genes for all three genomes using Augustus v3.2.2 (Stanke!and!

Waack,!2003). We used the Nematostella vectensis v1.0 training set

(http://ryanlab.whitney.ufl.edu/downloads/nematostella_training_files_for_augustus.t ar.gz) and hints generated with BLAT (Kent,!2002) alignments of transcriptome data

(C. cruxmelitensis ENA accession= HAHC01000000; C. xamachana ENA accession=

PRJEB21012; A. alata accession= PRJNA312373) to the genome assemblies. We generated 50,725 gene models for A. alatina, 27,276 for C. cruxmelitensis, and 24,718 for C. xamachana.

Orthologous Gene Analysis

We used OrthoFinder v1.1.4 (Emms!and!Kelly,!2015) to construct orthologous groups between gene models of A. alatina, C. cruxmelitensis, C. xamachana, N. vectensis, Hydra magnipapillata, and Homo sapiens. We also included translated transcriptome assemblies for A. alatina, C. cruxmelitensis, and C. xamachana in these ortholog analyses, as well as an additional transcriptome of the apo-symbiotic polyp stage of C. xamachana, which was assembled using Trinity v2.4.0 (Grabherr!et!al.,!2011) with default settings (ENA Project Accession:

PRJEB23739). All transcriptomes were translated using TransDecoder 3.0.0 (Haas!et!

! 146! ! al.,!2013) with minimum protein length (-m) set to 50 and all other settings as default.

Orthogroups were annotated by blasting a representative species against the

Uniprot/Swissprot database. Orthogroups with annotations were further mapped to

Gene Ontology terms and analyzed for enrichment related to biological function using

ClusterProfiler. In order to detect orthogroups associated with dinoflagellate symbiosis, we performed a secondary analysis with four symbiotic (Stylophora pistillata, Exaiptasia pallida, Acropora digitifera, Cassiopea xamachana) and four non-symbiotic cnidarians (N. vectensis, Hydra vulgaris, Aurelia aurita, C. cruxmelitensis).

Our OrthoFinder analysis generated a total of 78,227 orthogroups, with 1,187 species-specific orthogroups (note: orthogroups with single sequences are not counted in these species-specific orthogroups). Using a custom script, we identified 1,052

Acraspeda-specific orthogroups, 480 medusozoan-specific orthogroups, and 799

Cnidaria-specific orthogroups (Figure 2). Of these orthogroups, we were able retrieve

Swissprot annotations for 436 out of 799 Cnidaria-specific orthogroups, 67 of the 480

Medusozoa-specific orthogroups, and 298 of the 1052 Acraspeda-specific orthogroups (Table S1-S3). Unannotated orthogroups may represent taxonomically restricted genes within each lineage and further characterization will be required for those groups. Of those that were annotated, we identified enriched gene ontology terms and identified 46 biological process terms that were enriched within Cnidaria,

25 for medusozoa, and 155 for Acraspeda (p-adjust < 0.01), but failed to identify enrichment of Cnidarian orthogroups. Cnidaria biological process terms were largely clustered into 4 clusters related to G-protein coupled receptor signaling, ion transmembrane transport, detection of stimulus, and synaptic transmission (Figure 5,

Table S1). Medusozoa enriched biological process terms were largely clustered into 2

! 147! ! main groups related to organelle and vesicle localization, and synaptic transmission.

(Figure 6, Table S2). Other enriched terms were related to morphogenesis and neurotransmitter function. Over enriched terms identified for Acraspeda show genes involved in multitude of biological processes, including metabolism, protein processing, response to stress and stimulus, and cell component localization (Figure 7,

Table 3). Enriched biological processes that were most statistically significant were involved in signaling (GO:0023052), cell communication (GO:0007154), developmental process (GO:0032502), single-multicellular process (GO0044707), multicellular organismal process (GO: 0032501), and cell differentiation

(GO:0030154)

The second analysis identified 7 orthogroups that were specific to taxa that are symbiotic with dinoflagellates. Of the 7 identified orthogroups, 5 were annotated, including retrovirus-related pol polyprotein from transposons (Table 3).

Retrotransposon activity have been linked to metazoan genome evolution, with transpositions providing novel promoters leading to differential regulation of proximal genes (Kazazian, 2004;Batut et al., 2013). However, genes sharing the genomic scaffolds with the pol polyprotein predominantly lacked annotations within the Swissprot database or were annotated as additional retroviral transposons. Thus, the implications of the retrovirus-related Pol polyprotein to the evolution of symbosis are unclear. Ankyrin proteins function to provide organizational stability and mechanical support to plasma membranes and membrane spanning proteins, and were likely present in pre-cambrian animals (Bennett and Lorenzo, 2013). While other cnidarian lineages possess ankyrins, a symbiosis-specific ankyrin may suggest its involvement in functioning within the Symbiodinium enveloping symbiosome membrane (Kazandjian et al., 2008). Interestingly, an ankyrin-repeat containing

! 148! ! membrane protein is required for the persistence of the symbiosome in the legume-

Rhizobium symbiosis (Kumagai et al., 2007;Hakoyama et al., 2012). Another striking finding is the presence of the gene coding for L-gulonolactone oxidase, which is involved in the synthesis of ascorbate (Vitamin C), with a transition to vitamin C auxotrophy often resulting from the genomic loss of L-gulonolactone (Helliwell et al.,

2013;Wheeler et al., 2015). Ascorbate plays a role antioxidant activity, reducing metabolically generated reactive oxygen species levels, which can have significant implications for an animal housing a photosynthetically active eurkaryote. During heat stress, reactive oxygen species are generated as the photosynthetic machinery of

Symbiodinium is damaged and leads to disruption of symbiosis (Weis, 2008;Weis et al., 2008;Roth, 2014;Levin et al., 2016). While the gene does not preclude the ability for biosynthesis of ascorbate, the capacity to generate vitamin C can be highly advantageous. In addition ascorbate synthesis, quinone oxidoreductase is an additional enzyme that further supports the presence of a photosynthetic endosymbiotic, as it has been suggested to be involved in induction of melanogenesis

(Choi et al., 2010). While further characterization will be necessary to determine the degree to which the genes belonging to lineage specific orthogroups play in physiology, and morphology of the organisms, all of these orthologous groups potentially play an important role in the evolution of the cnidarian lineages.

Venom Analysis

We identified potential venom-encoding genes from the transcriptomes using the venomix database (a curated set of 6,622 venom-related proteins) and associated pipeline (https://bitbucket.org/JasonMacrander/venomix). Transcripts that were identified as venom-encoding were searched against the protein predictions of the corresponding genomes with BLAST v2.2.31+ (e-value = 10-6) (Camacho!et!al.,!

! 149! !

2009) in order to confirm the presence of these genes within the genome assemblies.

Transcripts identified with the venomix pipeline were treated as a single protein hit, and combined with the BLAST output (Table 2). We identified 119 families of putative venom proteins that were present in at least one of the five cnidarian taxa

(Table 2). We identified 93 families of venom-encoding genes in both C. cruxmelitensis and A.alata, and 98 families in the C. xamachana genome. In addition to the venomix analysis, we analyzed the venom content of the genomes using

OrthoFinder v1.1.4 by adding the venomix database to our initial set of input sequences. Using this process, we identified 123 orthogroups encoding venom genes that occurred in all five cnidarian genomes as well as the human genome (Figure 3).

Of the 123 venom orthogroups, few were found to be specific to any one cnidarian lineage, with only three orthogroups present across all cnidarians and one spanning just medusozoans. This is likely because most venom-related proteins in the venomix database were identified first in bilaterian animals, many of which are model organisms, whereas putative toxins recently identified in non-model cnidarians often lack robust annotations required for accession into this curated database. However, we were successful in identifying two taxon-specific toxin proteins, CrTX and CqTX, the amino acid sequences of which were first characterized in cubozoans, in the A. alata, and C. xamachana genomes (Nagai!et!al.,!2000;Nagai!et!al.,!2002).!We identified seven and six putative proteins of CqTX and CrTX, respectively, from A. alata, while the C. xamachana genome contained only three CqTX genes and one CqTX gene.

However, both toxins were absent from the C. cruxmelitensis genome and transcriptome, potentially indicating these genes may have been lost in the staurozoan lineage.

! 150! !

Hox-POU synteny analysis

In the hydrozoan Eleutheria dichotoma, a POU6 class homeobox gene is fused with a phosphopantothenoylcysteine-synthetase (PPCS); and this fusion gene is linked to a Hox class homeobox gene, Cnox5 (Kamm!and!Schierwater,!2007). The fusion of PPCS and POU6 is not seen outside of Cnidaria, but it is present in the anthozoan

Nematostella vectensis suggesting it was present in the last common cnidarian ancestor. On the other hand the PPCS/POU6 fusion is not linked to a Hox class gene in N. vectensis (cf. Putnam et al. 2007 assembly (Putnam!et!al.,!2007))!suggesting this linkage might be a more recent event. We searched our three Acraspeda genomes for the presence of the PPCS/POU6 fusion gene and for synteny between this gene and the ortholog to the E. dichotoma Cnox5. In the C. cruxmelitensis and C. xamachana genomes we find the PPCS/POU6 fusion gene linked to a Cnox5 ortholog

(Figure 4). This result suggests that these two genes were linked in the last common medusozoan ancestor. Furthermore, considering that the last common medusozoan ancestor may have lived some 630-850 million years ago (Rogers,!2009), it is reasonable to conclude that a functional constraint has led to this conservation in synteny.

Based on the well-established linkage of a POU class homeobox gene to Hox clusters in vertebrates (Kamm and Schierwater, 2007), it had been suggested that a

Pou-Hox linkage may have been present in the last common ancestor of cnidarians and bilaterians. To check this, we searched several additional anthozoan genomes:

Stylophora pistillata, Acropora digitifera, Orbicella faveolata, as well as several invertebrate bilaterian genomes: Capitella teleta (Polychaeta), Strigamia maritima

(Chilopoda), Octopus bimaculoides (Cephalopoda), Mizuhopecten yessoensis

(Bivalvia), and Ciona intestinalis (Ascidiacea) that were not available at the time of

! 151! ! the original study. We found no evidence for ancient Pou-Hox synteny in these anthozoans nor in the invertebrate bilaterian genomes, suggesting that the Hox-Pou linkage in medusozoans was achieved independently from the vertebrate Hox-Pou linkage. These findings demonstrate the utility of the three new medusozoan genomes in addressing questions pertaining to molecular evolution, as well as the synergistic effect that increased genomic-level taxon sampling can provide when testing hypotheses about ancestral state reconstruction.

Conclusions

In this note we provided draft genomes for three species of the medusozoan sub-group Acraspeda (Cnidaria)– Calvadosia cruxmelitensis (Staurozoa), Alatina alata (Cubozoa), and Cassiopea xamachana (Scyphozoa) – and our corresponding bioinformatics workflows for their assemblies and partial annotations. These assemblies represent the first published genomes for acraspedan jellyfish species and have been made available in the NCBI public database. The findings of our preliminary orthology analyses and annotation of Hox-linked and venom-related genes provide a glimpse into genetic components underlying the evolution of certain traits in these early metazoans. In particular, we've identified potential genes involved in facilitating symbiosis within symbiotic cnidarian lineages. Coupled with appropriate bioinformatics tools and data management pipelines, researchers across a broad range of scientific fields can utilize these resources to investigate the genetic basis of defense, reproduction, symbiosis, and communication in this ancient and specious group that encompasses a diversity of life histories, some of which exhibit pelagic life stages. Furthermore, cnidarian genomes offer strategic opportunities to investigate possible genetic links to any number of ecological issues related to jellyfish that are frequently reported in the scientific literature, or in the news media.

! 152! !

These medusozoan genomes will be useful resources in developing functional constructs (e.g. CRISPR/Cas9 guide RNAs) that can be employed to understand the genomic basis for some of the captivating biological features of these animals. Lastly, the availability of these genomic-level sequence data is an important step forward in the pursuit to elucidate evolutionary events that may have shaped Medusozoa, and in reconstructing the last common ancestor of Cnidaria and Bilateria. Therefore, we are confident that these new genomes will prove essential for understanding key evolutionary genomic events that were formative in the early evolution of cnidarians and bilaterians.

Availability of supporting Data

Accession numbers for raw sequencing reads and assemblies are available in

Table 1. Custom scripts and parameters used for the analyses are available at https://github.com/josephryan/Ohdera_et_al_2018

! 153! !

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Figure!1.!A)!Calvadosia(cruxmelitensis!(Staurozoa),!B)!Alatina(alata!(Cubozoa),!and! C)!Cassiopea(xamachana!(Sycphozoa).!D)!Phylogenetic!relationship!of!major! cnidarian!lineages!after!Kayal!et!al.!(2018),!linking!Cubozoa!and!Scyphozoa!as! sister!groups!and!united!with!Staurozoa!to!form!the!Acraspeda!lineage.!!

! 168! !

Figure!2.!Gene!Content!Distribution!in!Cnidarian!Lineages.!Filled!circles!in!the! bottom!panel!indicate!shared!orthogroups!in!these!lineages.!Bar!graphs!indicate! the!number!of!orthogroups!corresponding!to!each!filled8circle!pattern.!Numbers! next!to!each!species!abbreviation!indicate!the!total!number!of!orthogroups! identified!for!that!species.!Hsap!=!Homo(sapiens;!Nvec!=!Nematostella(vectensis;! Hmag!=!Hydra(magnipapillata;!Ccrux!=!Calvadosia(cruxmelitensis;!Aala!=!Alatina( alata;!Cxam!=!Cassiopea(xamachana.!!

! 169! !

Figure!3.!Distribution!of!venom8related!genes!in!cnidarian!lineages.!Filled!circles! in!the!bottom!panel!indicate!presence!of!venom8related!gene!in!each!lineage.!Bar! graphs!indicate!the!number!of!venom8related!orthogroups!corresponding!to!each! filled8circle!pattern.!Numbers!next!to!each!species!abbreviation!indicate!the!total! number!of!venom8related!orthogroups!identified!for!that!species.!Hsap!=!Homo( sapiens;!Nvec!=!Nematostella(vectensis;!Hmag!=!Hydra(magnipapillata;!Ccrux!=! Calvadosia(cruxmelitensis;!Aala!=!Alatina(alata;!Cxam!=!Cassiopea(xamachana.!!

! 170! !

Figure!4.!Linkage!of!PPCS8Pou!Genes!with!Hox!Genes!in!Medusozoa!Genomes.! Genomic!scaffolds!from!three!Medusozoa!lineages!(C.(xamachana,(C.( cruxmelitnesis,(and(E.(dichotoma)!show!linkage!of!the!PPCS8POU!gene!linked!to!a! Hox!gene!(dark!green).!This!linkage!is!not!seen!in!Anthozoa!(N.(vectensis).!Scaffold! length!is!shown!to!the!right!of!the!bar.!The!light!green!region!indicates!the! transcribed!portion!of!the!scaffold,!and!exons!are!represented!within!by!curved! rectangles!(PPCS!exons!=!purple,!POU!exons!=!yellow).!Edic!=!Eleutheria( dichotoma;!Ccrux!=!Calvadosia(cruxmelitensis;!Cxam!=!Cassiopea(xamachana;!Nvec! =!Nematostella(vectensis.!

! 171! !

secondary metabolic process DNA integrationprotein homooligomerization

neurological system process

plot_size

3 cellular polysaccharide biosynthetic process

transmembrane transport

5 detection of mechanical stimulus 0 metal ion transport cationcalcium transport ion transport response to calcium ion potassium ion transport monovalent inorganic cation transport detection of stimulus 6 ion transmembrane transport semantic space x cation transmembrane transport inorganic cation transmembrane transport potassium ion transmembrane transport G−protein coupled receptor signaling pathway inorganic ion transmembrane transport glutamate receptor signaling pathway regulation of ion transmembrane transport log10_p_value 0 adenylate cyclase−modulating G−protein coupled receptor signaling pathway

−5 −5

−10

synaptic transmission, glutamatergic

action potential

−10

−4 0 4 8 semantic space y ! !

Figure!5.!Gene!ontology!biological!processes!over8enriched!within!Cnidaria! specific!orthogroups!visualized!using!REViGO.!Over8representation!analysis!was! performed!with!ClusterProfiler,!with!a!p8adjusted!cutoff!of!0.01.!Color!indicates! Log10!transformed!p8adjusted!value.!Terms!are!plotted!within!a!x8y!semantic! space,!in!which!similar!terms!are!clustered!within!closer!proximities.!Color! indicates!p8value!and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.! ! ! !

! 172! !

neurotransmitter transport

calcium ion regulated exocytosis

chemical synaptic transmission trans−synaptic signaling signal release log10_p_value 0 presynaptic process involved in chemical synaptic transmission synaptic vesicle cycle 4 synaptic signaling −2 neurotransmitter secretion −4 establishment of organelle localization

cell−cell signaling synaptic vesicle localization −6 vesicle localization

plot_size

G−protein coupled receptor signaling pathway 4.0

semantic space x 0 4.5

5.0

regulation of neurotransmitter levels −4

morphogenesis of an epithelium morphogenesis of a branching structure

−4 0 4 semantic space y ! Figure!6.!Gene!ontology!biological!processes!over8enriched!within!Medusozoa! specific!orthogroups!visualized!using!REViGO.!Over8representation!analysis!was! performed!with!ClusterProfiler,!with!a!p8adjusted!cutoff!of!0.01.!Color!indicates! Log10!transformed!p8adjusted!value.!Terms!are!plotted!within!a!x8y!semantic! space,!in!which!similar!terms!are!clustered!within!closer!proximities.!Color! indicates!p8value!and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.! !

! 173! !

single−organism organelle organization protein metabolic process

protein catabolic process log10_p_value 5 0

−3

−6 microtubule−based process movement of cell or subcellular componentmacromolecule modification regulation of cell division regulation of cell cycle −9

cell motility plot_size regulation of transferase activity 4.0 regulation of localization cell differentiation regulation of cell proliferation 4.5

0 vesicle−mediated transport localization of cell immune system process

5.0

semantic space x locomotion

biological adhesion regulation of response to stimulus

5.5 multi−organism process cell proliferation

reproduction 6.0 −5 cell communication single−multicellular organism process signaling

multicellular organismal process cell adhesion

single organism reproductivedevelopmental process process

−5 0 5 semantic space y ! !

Figure!7.!Gene!ontology!biological!processes!over8enriched!within!Acraspeda! specific!orthogroups!visualized!using!REViGO.!Over8representation!analysis!was! performed!with!ClusterProfiler,!with!a!p8adjusted!cutoff!of!0.01.!Color!indicates! Log10!transformed!p8adjusted!value.!Terms!are!plotted!within!a!x8y!semantic! space,!in!which!similar!terms!are!clustered!within!closer!proximities.!Color! indicates!p8value!and!circle!size!indicates!frequency!of!GO!term!in!the!Cassiopea! database.!

)

! 174! !

! Alatina alata Calvadosia cruxmelitensis Cassiopea xamachana NCBI Taxa ID 1193083 1843192 12993 # of Sequences 291,445 50,999 93,483 Total Length (bp) 851,121,747 209,392,379 393,520,168 N50 (bp) 7,049 16,443 15,563 CEGMA (Complete) 8.06 61.29 53.63 CEGMA (Complete + Partial) 29.84 91.94 82.66 BUSCO (Complete) 18.30 70.86 58.59 BUSCO (Complete + Partial) 32.11 85.07 66.97 GC Content (%) 38.07 39.95 37.07 Assembly Accession PUGI00000000 OFHS01000000 OLMO01000000 NCBI Raw Read Accession PRJNA421156 PRJEB23739 PRJEB23739 Specimen Voucher ID USNM 1248604 USNM 1286381 UF Cnidaria 12979

Table!1.!Statistics!of!the!genomic!assemblies!of!Alatina)alata,)Calvadosia)cruxmelitensis,! and!Cassiopea)xamachana.!

! 175! !

Family ID Ccrux Aala Cxam Hmag Nvec Acetylcholinesterase Acetylcholinesterase-1_1 0 6 10 1 1 Acidic_phospholipase Acidic_phospholipase_A2_1_322 1 1 1 2 4 Acidic_phospholipase_A2_2_1 0 0 0 0 3 Acidic_phospholipase_A2_daboiatoxin_B_chain_6 0 0 0 0 6 Alpha-amylase Alpha-amylase_1 0 0 1 0 0 Alpha-latroinsectotoxin Alpha-latroinsectotoxin-Lh1a_1 0 2 6 3 0 Alpha-latroinsectotoxin-Lt1a_2 30 61 45 3 3 Astacin-like_metalloprotease_toxin Astacin-like_metalloprotease_toxin_1_1 24 7 8 5 10 Astacin-like_metalloprotease_toxin_3_2 10 7 8 5 10 Astacin-like_metalloprotease_toxin_4_1 11 7 8 5 10 Astacin-like_metalloprotease_toxin_5_1 11 7 8 5 10 Basic_phospholipase Basic_phospholipase_A2_2_60 1 0 0 2 2 Basic_phospholipase_A2_acanthin-1_3 1 1 1 2 4 Basic_phospholipase_A2_notexin_18 2 1 1 2 8 C-type_lectin C-type_lectin_6 12 6 13 11 27 C-type_lectin_galatrox_1 1 0 2 7 0 C-type_lectin_mannose-binding_isoform_30 9 8 17 11 27 Conodipine-M_alpha_chain_1 Conodipine-M_alpha_chain_1 1 1 1 2 2 Cystatin-1_1 0 0 1 2 0 Cysteine-rich Cysteine-rich_venom_protein_1_2 0 0 0 0 2 Cysteine-rich_venom_protein_bucarin_1 1 1 1 2 0 Cysteine-rich_venom_protein_Mr30_2 2 2 3 2 4 Cytolysin_RTX Cytolysin_RTX-S-2_2 0 0 0 4 0 Delta-latroinsectotoxin Delta-latroinsectotoxin-Lt1a_1 19 53 45 3 3 Disintegrin Disintegrin_acostatin-alpha_26 3 1 1 0 0 Disintegrin_bitistatin_6 4 1 2 4 5 Disintegrin_EMF10B_17 3 0 0 0 0 Disintegrin_lebein-1-beta_8 2 0 0 0 0 Disintegrin_saxatilin_24 3 1 1 4 5 Disintegrin-like Disintegrin-like_leberagin-C_8 5 1 2 4 5 Factor_V Factor_V_activator_RVV-V_gamma_142 3 3 4 5 11 Galactose-specific_lectin Galactose-specific_lectin_nattectin_1 14 8 21 11 20 Hyaluronidase Hyaluronidase_1 1 1 1 2 1 Hyaluronidase_CdtHya1_1 0 0 0 0 1 Hyaluronidase_conohyal-ad1_1 1 1 1 2 1 Hyaluronidase_conohyal-Cn1_1 1 1 2 2 1 Hydralysin Hydralysin-1_6 0 0 0 2 0 KappaPI-theraphotoxin KappaPI-theraphotoxin-Hs1a_80 6 2 2 6 6 Kunitz-type Kunitz-type_conkunitzin-S1_2 3 1 1 4 4 Kunitz-type_proteinase_inhibitor_AEPI-IV_2 1 0 0 2 0 Kunitz-type_serine_protease_inhibitor_2_4 3 4 1 4 5 Kunitz-type_serine_protease_inhibitor_As-fr-19_1 1 2 1 6 4 Kunitz-type_serine_protease_inhibitor_bicolin_1 0 0 0 2 1 Kunitz-type_serine_protease_inhibitor_bitisilin-3_1 5 5 1 6 8 Kunitz-type_serine_protease_inhibitor_BmKTT-2_1 6 4 1 6 7 Kunitz-type_serine_protease_inhibitor_BmKTT-3_1 5 3 1 6 6 Kunitz-type_serine_protease_inhibitor_conotoxin_Cal9 4 4 1 5 7 Kunitz-type_serine_protease_inhibitor_Hg1_1 6 4 1 7 7 Kunitz-type_serine_protease_inhibitor_homolog_alpha-dendrotoxin_3 2 1 1 3 6 Kunitz-type_serine_protease_inhibitor_homolog_beta-bungarotoxin_B5-B_chain_1 1 1 1 6 3 Kunitz-type_serine_protease_inhibitor_homolog_delta-dendrotoxin_2 5 1 2 2 6 Kunitz-type_serine_protease_inhibitor_homolog_dendrotoxin_K_154 7 4 2 6 8 Kunitz-type_serine_protease_inhibitor_Kunitz-1_1 7 4 3 6 8 Kunitz-type_serine_protease_inhibitor_PIVL_13 5 2 1 5 8 Kunitz-type_serine_protease_inhibitor_U1-aranetoxin-Av1a_1 7 4 2 7 7 L-amino-acid_oxidase L-amino-acid_oxidase_31 1 1 1 3 7 Nematocyte Nematocyte_expressed_protein_6_1 22 7 7 5 10 Neprilysin Neprilysin-1_1 4 2 5 2 2 Neutral_phospholipase Neutral_phospholipase_A2_3_21 1 1 1 2 4 Peptide Toxin Peptide_toxins_Am-1_1 0 0 0 0 2 Perivitellin Perivitellin-2_31_kDa_subunit_1 0 0 1 0 0 Perivitellin-2_67_kDa_subunit_1 2 1 2 0 0 Peroxiredoxin Peroxiredoxin-4_1 1 1 1 2 2 Phospholipase Phospholipase_A1_19 1 1 1 2 2 Phospholipase_A2_1 3 1 2 3 8 Phospholipase_A2_2 0 0 1 0 0 Phospholipase_A2_homolog_crotoxin_acid_subunit_CA_21 1 1 1 2 4 Phospholipase_A2_imperatoxin-1_4 1 1 1 2 0 Phospholipase_A2_large_subunit_1 1 1 1 2 0 Phospholipase_A2_phaiodactylipin_6 1 1 0 2 0 Phospholipase_A2_Scol@Pla_1 1 1 1 2 4 Plancitoxin Plancitoxin-1_1 1 1 1 2 0 Potassium Channel Toxin Potassium_channel_toxin_BcsTx3_1 0 0 0 0 2 Protease inhibitor Protease_inhibitor_1_6 7 4 2 9 8 Putative protein-glutamate O-methyltransferase Putative_antimicrobial_peptide_7848_1 0 0 0 0 1 SE-cephalotoxin Putative_protein-glutamate_O-methyltransferase_1 1 1 1 2 2 Serine_proteinase-like SE-cephalotoxin_1 0 0 1 1 0 Snaclec (C-type lectin-like proteins)(CLPs) Serine_proteinase-like_BMK-CBP_1 0 1 1 3 0 Snaclec_aspercetin_subunit_alpha_2 3 1 1 0 6 Snaclec_botrocetin_subunit_beta_1 11 3 6 21 27 Snaclec_coagulation_factor_IX@factor_X-binding_protein_subunit_A_1 9 3 10 5 15 Snaclec_coagulation_factor_X-activating_enzyme_light_chain_2_135 12 3 10 11 24 Snaclec_echicetin_subunit_alpha_1 9 5 4 8 17 Snaclec_echicetin_subunit_beta_6 8 3 9 5 6 Snaclec_lebecin_subunit_alpha_9 8 3 8 3 14 Snaclec_rhodocetin_subunit_alpha_2 3 5 8 1 2 Snaclec_rhodocetin_subunit_delta_1 10 5 11 22 27 Snaclec_VP12_subunit_A_1 3 2 4 0 15 Snake_venom_metalloproteinase Snake_venom_metalloprotease_inhibitor_02A10_1 0 0 0 0 2 Snake_venom_metalloproteinase_Ac1_1 3 3 1 4 6 Snake_venom_metalloproteinase_acutolysin-C_1 3 3 1 4 6 Snake_venom_metalloproteinase_fibrolase_19 3 3 1 4 7 Sticholysin-2_8 0 0 0 4 0 Techylectin-like Techylectin-like_protein_1 24 7 28 0 3 Thrombin-like_enzyme Thrombin-like_enzyme_ancrod_11 4 3 5 5 53 Thrombin-like_enzyme_TLBm_1 3 3 2 5 28 Toxin_AvTX-60A_3 0 0 0 0 1 Toxin_CqTX-A_3 0 6 1 0 0 Toxin_CrTX-A_1 0 7 3 2 2 Toxin_MsePTx1_1 0 0 0 0 0 Turripeptide_Ici9 2 6 5 4 32 Turripeptide Turripeptide_Pal9 1 5 2 4 28 U24-ctenitoxin-Pn1a_1 3 1 1 0 1 Venom_allergen Venom_allergen_3_30 5 4 2 3 23 Venom_allergen_5 3 8 7 3 37 Venom_allergen_5_1 5 5 6 3 36 Venom_allergen_5_15 6 5 4 3 34 Venom peptide isomerase heavy chain Venom_peptide_isomerase_heavy_chain_2 6 4 7 5 58 Venom_peptide_SjAPI-2_1 0 0 0 0 1 Venom_protein_59 0 0 0 2 4 Venom_prothrombin Venom_prothrombin_activator_pseutarin-C_catalytic_subunit_12 5 5 9 5 58 Zinc-metalloproteinase Zinc_metalloproteinase_recombinant_fibrinogenase_II_1 4 3 1 4 105 Zinc_metalloproteinase-disintegrin-like_alternagin_2 3 1 2 4 88 Zinc_metalloproteinase-disintegrin-like_ammodytagin_1 3 1 1 4 56 Zinc_metalloproteinase-disintegrin-like_mikarin_1 3 1 1 4 56 Zinc_metalloproteinase-disintegrin-like_VAP2B_272 8 4 4 4 145 Zinc_metalloproteinase@disintegrin_107 6 4 1 4 139 Zinc_metalloproteinase@disintegrin_4 4 1 1 4 76 Zinc_metalloproteinase@disintegrin_5 4 1 1 4 83

Table!2:!Venom8encoding!gene!repertoire!of!five!cnidarian!genomes!identified! with!the!venomix!database.!Venom!genes!are!categories!by!families!(column!1).! Both!genomic!and!transcriptomic!data!were!used,!with!transcriptomic!isoforms! counted!as!a!single!venom8encoding!gene.!

! 176! !

Orthogroup! ID! Protein!Name!

OG0000584' Cxam.v1_g706.t1' N/A'

OG0003792' Cxam.v1_g4560.t1' Quinone'oxidoreductase'(EC'1.6.5.5)'(NADPH:quinone'reductase)'

RetrovirusMrelated'Pol'polyprotein'from'transposon'297'[Includes:'Protease'(EC' OG0006001' Cxam.v1_g8438.t1' 3.4.23.M);'Reverse'transcriptase'(EC'2.7.7.49);'Endonuclease]'

OG0006787' Cxam.v1_g23465.t1' AnkyrinM3'(ANKM3)'(AnkyrinMG)'

OG0007961' Cxam.v1_g5603.t1' LMgulonolactone'oxidase'(LGO)'(EC'1.1.3.8)'(LMgulonoMgammaMlactone'oxidase)'(GLO)'

OG0010609' Cxam.v1_g21481.t1' N/A' Cxam.v1_g2940.t1,' RetrovirusMrelated'Pol'polyprotein'from'transposon'opus'[Includes:'Protease'(EC' OG0001102' Cxam.v1_g6802.t1,' 3.4.23.M);'Reverse'transcriptase'(EC'2.7.7.49);'Endonuclease]' Cxamv1_g18470.t1' ' ' '

Table!3.!Orthrogroups!found!to!be!specific!to!cnidarians!that!are!symbiotic!with! Symbiodinium.!Gene!annotations!were!retrieved!from!Swissprot!with! representative!sequences!from!the!Cassiopea(genome.!! ! ! ! ! ! ! ! ! ! !

! 177! !

Chapter 6

Conclusion ! The role of symbiosis in the life history and evolution of metazoans is becoming increasingly apparent, to the extent whereby signatures of co-evolution can be detected (Wernegreen, 2012;McFall-Ngai et al., 2013;Gilbert et al., 2015). The evolutionary role of symbiosis can be traced back to a pre-metazoan era, with the now widely accepted theory of endosymbiosis between prokaryotes leading to the evolution of the mitochondria (Sagan, 1967;de Duve, 2007). Examples of symbiosis driving life history changes can be found in a multitude of organisms, and the impacts of symbionts to holobiont physiology can lead to alterations in gene expression, behavior, immune function, etc (Jaenike et al., 2010;Hughes et al., 2011;Kremer et al., 2012;Winkler et al., 2015;Okayama et al., 2016;Tanaka et al., 2018). Symbiotic associations can be found in virtually all metazoans, with a wide range of microbes, from bacteria to single cell eukaryotes, specificity is highly variable form obligate to commensal (Sanchez et al., 2007;Chavez-Dozal et al., 2012;Ainsworth et al., 2015).

Despite the ubiquity of animals associating with microbes, we have yet to fully understand the role microbes play in life history evolution. Changes to genomic architecture are often observed in the microbes, with genome reductions being typical of mutually obligate symbiosis (Bennett et al., 2014;Campbell et al., 2015).

Modifications to host genomes have often been identified as a result of horizontal gene transfer of bacterial origin, often resulting in facilitating the host-symbiont association (Husnik et al., 2013;Nakabachi et al., 2014). Selective pressures driving the evolution of symbiosis related genes have also been observed in plant-microbe symbiosis, highlighting the intricacies of symbiosis in evolution (Delaux et al., 2014).

While the extent of host genomic evolution is not well understood, examples of

! 178! ! phenotypic consequences of symbiosis are abundant. This is particularly apparent through observations of the symbiont housing organs such as the bacteriocyte in insects (Braendle et al., 2003;Baumann, 2005), light organ in bobtail squids (McFall-

Ngai, 2014;Pankey et al., 2014), and the trophosome of hydrothermal vent tube worms (Nussbaumer et al., 2006). These symbiont-housing structures demonstrate the specificity of the association in reshaping the function of available structures to through modulation of gene expression. However, there are few examples in which symbiosis can affect dynamic developmental transitions, such as metamorphosis.

My work presented here addresses the role of microbial symbionts in shaping life history transitions, in particular, the dynamic process of metamorphosis. The upside-down jellyfish Cassiopea xamachana larvae associate with bacterial biofilm on degrading mangrove leaves, resulting in settlement and metamorphosis (Fleck and

Fitt, 1999;Fleck et al., 1999). The cue that induces both settlement and metamorphosis in invertebrates appear to be diverse, with species across multiple bacterial phyla capable of inducing metamorphosis (Tran and Hadfield, 2013). These were corroborated in C. xamachana larvae but searching through the genomes in order to find potential cues has yet to be attempted. Our findings demonstrate the diversity found within bacterial species belonging to the same genera, and the potential specificity larvae can exhibit towards these taxa. Furthermore, settlement- inducing bacteria are potentially rare within the microbiome, further suggesting an yet unknown factor such as nutrient or defense that may be involved in driving the evolution of larval-bacterial associations.

While similar benefits of nutrient translocation to the host have been implicated in cnidarian-dinoflagellate symbiosis, initiating metamorphosis of an animal has only been describe thus far in select members of the Scyphozoa. In C.

! 179! ! xamachana, colonization by Symbiodinium leads to strobilation, whereby the polyp produces the juvenile form of the sexual medusae. Our findings suggest modulation of the expression of key genes involved in strobilation as a result strobilation. While gene expression related to strobilation has only been investigated in Aurelia aurita

(Fuchs et al., 2014;Brekhman et al., 2015), our results highlight the co-option of existing pathway resulting in the obligate association between C. xamachana and

Symbiodinium. In the absence of colonization, C. xamachana will remain in its polyp form indefinitely, preventing sexual reproduction. While the evolutionary benefits of relying on a symbiont to complete the life cycle is unclear, these findings can provide additional avenues of understanding mechanisms of symbiosis driven life history evolution.

The genomic resources that I have generated, including the transcriptome and genome that are now publicly available, will aid the efforts of understanding evolutionary impacts of symbiosis to holobiont life history. In addition to genomic resources, I have generated genetic lines that have been disseminated to labs around the world. These resources provide the foundations for generating the C. xamachana system as a model for understanding symbiosis, with applications towards coral conservation, but towards answering questions in a broad range of fields including neurobiology, toxinology, fluid dynamics, etc. (Ohdera et al., 2018). These efforts have culminated into the establishment of the International Cassiopea Workshop, which brings researchers from multiple countries to share research and facilitate collaborations, allowing attendees to perform experiments and conduct sampling expeditions in the Florida Keys.

These developments indicate the future of the Cassiopea system, and efforts are currently underway to generate additional resources including generating

! 180! ! transgenic lines of C. xamachana using the CRISPR-cas9 system. Expansion and refinement of currently available protocols are also underway, including the development of an RNAi protocol, and genotyping methods for genotype-genotype interaction studies. These and community supplied resources will be made available through the Cassiopea portal, where users will have access to request polyps and download protocols in order to facilitate adoption of the system as easily as possible

(http://sites.psu.edu/cassbase/). Thus, we believe the Cassiopea model has broad applicability for research and will offer a unique model for answering a multitude of questions related to symbiosis and early metazoan biology.

! 181! !

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