Evolution of Developmental Pattern and Larval Form in the Ophiuroids of Oregon and Beyond by Nicole N. Nakata A dissertation accepted and approved in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biology Dissertation Committee: Nadia Singh, Chair Richard Emlet, Advisor Svetlana Maslakova, Core Member Julie Schram, Core Member Samantha Hopkins, Institutional Representative University of Oregon Fall 2023 © 2023 Nicole N. Nakata This work is licensed under a Creative Commons CC-BY. 2 DISSERTATION ABSTRACT Nicole N. Nakata Doctor of Philosophy in Biology Title: Evolution of Developmental Pattern and Larval Form in the Ophiuroids of Oregon and Beyond Despite the ubiquity and known impacts of transitions in developmental pattern in marine invertebrates, many taxa have been insufficiently analyzed. In Chapter II, I report the effects of larval feeding in an undescribed facultative planktotroph, Amphiodia sp. opaque. By culturing larvae with and without food and observing development, I found that larval feeding led to faster development times, greater percent metamorphosis, and larger juveniles able to evade starvation longer than individuals that did not receive food. We compared metrics of early life stages using a series of generalized linear models, and model fitting was determined using Akaike’s Information Criterion (AIC). Scores for each model and test are included as supplementary tables. In Chapter III, I present a summary of developmental diversity in the ophiuroids of the northeast Pacific Ocean. We used DNA barcoding to identify eighteen species from the plankton of the southern Oregon coast. We found four species with reduced plutei, one with vitellaria, three with pelagic direct development, and ten with planktotrophic ophioplutei (including one followed by a vitellaria). This diversity of larval forms suggests multiple transitions from feeding to nonfeeding larvae in the Amphiuridae, which I tested for in Chapter IV using comparative phylogenetic 3 analyses. To do so, I constructed a four-gene phylogenetic hypothesis for species with known development pattern in the family Amphiuridae. Single-gene trees were made to check for congruence between loci and the multi-gene dataset and are included as supplementary figures. This analysis inferred a brooding ancestor, an instance of re-acquisition of feeding, and multiple transitions back to nonfeeding larvae. The analysis inferred multiple transitions from brooding to nonfeeding planktonic development via pelagic direct development. Altogether, this work introduces new examples of the developmental patterns in Ophiuroidea, including an example of facultative planktotrophy, a rare developmental pattern that may represent an evolutionary intermediate between feeding and nonfeeding larvae. We show the effectiveness of DNA barcoding for identifying the early life stages of benthic marine invertebrates. Finally, I found that reconstructed ancestral states of developmental pattern are influenced by tree topology and completeness of the dataset. This dissertation includes previously published and unpublished co-authored material. 4 ACKNOWLEDGMENTS I wish to express sincere appreciation to the many colleagues, family, and friends that have supported this work. First, thank you to my advisor, Dr. Richard Emlet, for planting the seeds of these projects and for always keeping me sharp. Our shared hours at the microscope looking through plankton were some of the best and most educational moments in this program. I am ever thankful to my committee, Drs. Nadia Singh, Svetlana Maslakova, Samantha Hopkins, and Julie Schram, who have provided thoughtful feedback and support throughout this process. Sometimes what a student needs most is a kind word and encouragement; for this, I am especially grateful to Julie. Thank you to the whole OIMB community, especially the graduate students, for making Coos Bay a home. Reyn Yoshioka, Caitlin Plowman, and Kara Robbins were essential guides in getting to know the idiosyncrasies of OIMB. So many others have provided emotional and technical support over the years, including Erin Jezuit, Jessie Masterman, Christina Ellison, Lauren Rice, Ross Whippo, Maureen Heaphy, MacKenna Hainey, Alexa Romersa, Mike Thomas, Jenna Valley, and Ella Lamont. Thank you to the many OIMB staff that keep our institute running and who were always so helpful and kind to me. Especially Trish Mace, Clara Robbins, Maya Watts, Laura Screen, James Johnson, Mike Johnson, Lisa Samuelson, Debbie Seabright, and Jesse Borland. This work would not have been possible without the support of the Raymund Fellowship, William R. Sistrom Memorial Fellowship, and Neil Richmond Memorial Scholarship of the University of Oregon, the Charles Lambert Memorial Fellowship of Friday Harbor Laboratories, University of Washington, and the Melbourne R. Carriker Student Research Grant of the 5 American Malacological Society. This work was also supported in part by NSF grants OCE 1950520 (Emlet and Watts), OCE 1259603 (Emlet, Shanks, and Sutherland). We would like to thank the Smithsonian Tropical Research Institute, particularly Rachel Collin, the Florida Museum of Natural History, the Muséum d’Histoire Naturelle de Marseille, and the National Museum Victoria for lending us specimens to include in this work. Many people contributed to this project by bringing us individual larvae of interest, enumerating and photographing larvae from plankton samples, conducting gene amplification, and confirming adult identifications. They include George von Dassow, MacKenna Hainey, Gordon Hendler, Chris Mah, Jenna Valley, Ella Lamont, and Craig Young. 6 DEDICATION To my parents, Fran and Norm Nakata, and my partner, Tommy Ashcraft. Because of you, I have everything. 7 TABLE OF CONTENTS Chapter Page I. INTRODUCTION .................................................................................................... 14 II. HAVING CAKE AND EATING TOO: THE BENEFITS OF AN INTERMEDIATE LARVAL FORM IN A BRITTLE STAR AMPHIODIA SP. OPAQUE (OPHIUROIDEA ................................................................................. 18 1. Introduction ........................................................................................................ 18 2. Materials and Methods ....................................................................................... 23 2.1. Molecular identification of embryos and larvae ....................................... 23 2.2. Larval cultures and data collection ........................................................... 25 2.3. Feeding experiments ................................................................................. 28 2.4. Analysis..................................................................................................... 29 3. Results ................................................................................................................ 31 3.1. Species identity and occurrence of larvae ................................................. 31 3.2. Larval developmental mode ...................................................................... 33 3.3. Effects of larval food on developmental timing ....................................... 34 3.4. Effects of larval food on percent metamorphosis ..................................... 35 3.5. Effects of larval food on juvenile size ...................................................... 36 3.6. Effects of larval food on juvenile survival ................................................ 37 4. Discussion .......................................................................................................... 39 4.1. Species identity ......................................................................................... 39 4.2. Larval morphology and developmental mode .......................................... 40 4.3. Development time ..................................................................................... 41 4.4. Percent metamorphosis ............................................................................. 43 4.5. Juvenile size .............................................................................................. 43 8 4.6. Juvenile survival ....................................................................................... 44 5. Conclusion ............................................................................................................... 45 III. BRITTLE STAR LARVAE OF THE NORTHEAST PACIFIC ........................... 46 1. Introduction ........................................................................................................ 46 2. Methods.............................................................................................................. 51 2.2. Collection of larvae and adults ................................................................. 51 2.3. Molecular identification of embryos and larvae ....................................... 53 3. Results and Discussion ...................................................................................... 54 3.1. Taxonomic diversity ................................................................................. 54 3.2. Developmental diversity ........................................................................... 55 3.3. Patterns in spawning phenology ............................................................... 59 3.4. Species accounts ....................................................................................... 61 3.4.1. Ophiotrichidae ........................................................................... 61 3.4.2. Ophiopholidae ........................................................................... 67 3.4.3. Amphiuridae ............................................................................. 73 3.4.4. Ophiacanthina ........................................................................... 100 3.4.5. Ophiuridae ................................................................................. 110 3.4.6. Gorgonocephalidae ................................................................... 121 4. Conclusion .................................................................................................... 125 5. Keys to the ophiuroid planktonic forms of the northeast Pacific ....................... 126 IV. EVOLUTION OF LARVAL DEVELOPMENT IN THE BRITTLE STAR FAMILY AMPHIURIDAE ................................................................................... 131 1. Introduction ........................................................................................................ 131 2. Methods.............................................................................................................. 134 9 2.1. Sequence dataset and phylogenetic analyses ............................................ 134 2.2. Phylogenetic comparative analyses .......................................................... 139 3. Results ................................................................................................................ 141 3.1. Egg size and developmental mode ............................................................ 141 3.2. Sequence dataset and phylogenetic analyses ............................................ 142 3.3. Comparative phylogenetic analyses .......................................................... 144 4. Discussion .......................................................................................................... 150 4.1. Inference of development mode from egg size ......................................... 149 4.2. Phylogenetic analyses ............................................................................... 149 4.3. Ancestral state estimations ........................................................................ 150 4.4. Repeated gains and losses of planktonic development ............................. 150 REFERENCES CITED ................................................................................................ 159 SUPPLEMENTAL FILES ........................................................................................... 176 1 0 LIST OF FIGURES Figure Page 2.1. Amphiodia sp. opaque egg, embryo, pluteus, and juvenile ................................... 27 2.2. Maximum likelihood tree of COI sequences from adult and larval amphiurid spp. in the northeastern Pacific, constructed using the PhyML plug-in in Geneious. .............................................................................................................. 33 2.3. Cumulative sum of juveniles over time by treatment and year ............................ 35 2.4. Bar plots of percent metamorphosis averaged across experimental bowls for years 2020 and 2021 ............................................................................................. 36 2.5. Boxplot of juvenile aboral surface area at metamorphosis by food treatment, pooled across years ............................................................................................... 37 2.6. Scatterplots of planktonic duration by juvenile size, and juvenile aboral surface area by time to juvenile starvation ................................................ 38 2.7. Kaplan-Meier survival curves for Amphiodia juveniles according to larval food treatment and year ........................................................................................ 39 3.1. Developmental stages of the Ophiuroidea ............................................................ 50 3.2. Ophioplutei of the northeast Pacific, viewed in transmitted and cross-polarized light ....................................................................................................................... 59 3.3. Spawning phenology for 14 planktonic ophiuroid spp. of the southern Oregon coast ...................................................................................................................... 61 3.4. Spawning phenology for 4 additional ophiuroid spp. that were each observed on a single occasion .............................................................................................. 61 3.5. Maximum likelihood tree of 16S and COI of Ophiothrix spp. from North America ................................................................................................................ 63 3.6. Larvae and juveniles of Ophiopholis kennerlyi, O. bakeri, and Ophiothrix spiculata ............................................................................................................... 67 3.7. Maximum likelihood tree of COI sequences for Ophiopholis spp. ...................... 69 3.8. Maximum likelihood tree of 16S and COI sequences for Amphiuridae species from the northeast Pacific ..................................................................................... 75 3.9. Planktotrophic plutei of Amphiuridae: Amphiodia urtica and Amphipholis pugetana ............................................................................................................... 81 1 1 3.10. Reduced plutei of three species of Amphiodia: A. sp. opaque, A. sp. orange belly, and A. sp. tan ............................................................................................. 90 3.11. Three nonfeeding planktonic forms of Amphiuridae: Amphiodia periercta?, Amphioplus sp. vitellaria, and Amphiura arcystata ............................................ 95 3.12. Maximum likelihood tree for a concatenated dataset of 16S and COI sequences for superfamily Ophiacanthina from the Pacific Ocean .................... 101 3.13. Ophioplutei and juveniles of Ophiacantha diplasia ........................................... 106 3.14. Ophioplutei, vitellaria, and juvenile of Ophiopteris papillosa ........................... 109 3.15. Maximum likelihood tree for a concatenated dataset of 16S and COI sequences for Ophiuridae spp. ............................................................................ 112 3.16. Ophioplutei of four species of Ophiuridae: Ophiura leptoctenia, O. luetkenii, O. sarsii, and Ophiocten hastatum. ..................................................................... 115 3.17. Maximum likelihood tree of COI for 27 spp. of Gorgonocephalus from the Pacific ................................................................................................................. 123 3.18. Planktonic stages of Gorgonocephalus eucnemis ............................................... 125 4.1. Histogram of egg diameters by developmental pattern for 37 spp. of Amphiuridae ......................................................................................................... 142 4.2. Phylogenetic hypothesis for 39 spp. of Amphiuridae, constructed by maximum likelihood .............................................................................................................. 143 4.3. Phylogenetic hypothesis for 30 spp. of Amphiuridae, constructed by Bayesian inference ............................................................................................................... 144 4.4. Estimated ancestral character states for three-state model of developmental patterns mapped on maximum likelihood phylogeny .......................................... 147 4.5. Estimated ancestral character states for five-state model of developmental patterns mapped on maximum likelihood phylogeny .......................................... 149 1 2 LIST OF TABLES Table Page 2.1. Effects of larval feeding on development of facultative planktotrophs ................ 20 2.2. Collection and accession data for specimens included in Fig. 2 ........................... 24 2.3. Summary of traits for larval and juvenile Amphiodia sp. opaque ......................... 34 3.1. PCR primers for molecular identification of ophiuroid larvae ............................. 54 3.2. Ophiuroid species of southern Oregon with planktonic development .................. 56 3.3. Specimen information for Ophiothrix spp. included in Figure 3.4 ....................... 64 3.4. Specimen information for Ophiopholis spp. included in Fig. 3.7 ......................... 69 3.5. Specimen information for Amphiuridae spp. included in Fig. 3.8 ....................... 76 3.6. Specimen information for species of superfamily Ophiacanthina from the Pacific included in Figure 3.12 ............................................................................. 101 3.7. Specimen information for Ophiuridae spp. included in Fig. 3.15 ........................ 112 3.8. Specimen information for Gorgonocephalus spp. included in Fig. 3.17 .............. 123 4.1. Egg size, developmental mode, and larval form for species included in phylogenetic hypothesis for the family Amphiuridae .......................................... 135 4.2. PCR primers used to amplify four loci for phylogenetic analysis ........................ 139 4.3. Summary of the three-state transition models tested using ML .......................... 148 4.4. Summary of the five-state transition models tested using ML ............................. 150 1 3 CHAPTER I INTRODUCTION The evolutionary history of developmental patterns in marine invertebrates is an important system for examining the loss and possible regain of complex and important traits. In most marine taxa, benthic adults spawn or release planktonic larvae that vary in size, complexity, and developmental duration based on the level of parental investment. Many marine invertebrates have elaborate feeding larvae that develop from very many small eggs; others, sometimes even closely related species, have simple nonfeeding larvae that develop from moderately sized eggs. In some taxa, both planktonic and benthic brooding strategies exist (Hendler, 1991; Strathmann & Strathmann, 1982). Evolutionary transitions from planktonic feeding to nonfeeding larvae are persistent and widespread across marine invertebrates, and have consequences for species distributions and population connectivity (Strathmann, 1985). Many groups of marine invertebrates contain a diversity of developmental strategies, sometimes even within families or genera, suggesting multiple transitions in mode of development (Allen & Podolsky, 2007; Collin, 2004; Krug et al., 2015). Some groups, such as the brittle stars (Echinodermata: Ophiuroidea) have great potential to study evolutionary transitions in development but have historically lacked sufficient data for analysis. In the following chapters I investigate the evolution of development in brittle stars. Three categories are accepted (Hendler, 1975): obligate feeding (planktotrophy) in ophioplutei, abbreviated development in vitellaria, reduced plutei, and pelagic direct developers, and direct 1 4 development by brooding. These developmental patterns occur throughout ophiuroid taxa, suggesting transitions between divergent patterns. However, developmental pattern of many ophiuroid species have yet to be characterized. Unlike their echinoid and asteroid relatives, ophiuroids lack reliable means to obtain mature gametes in the lab (Selvakumaraswamy & Byrne, 2000). Instead, I collected wild larvae from the plankton to study development patterns and taxonomic diversity of ophiuroids in the northeast Pacific. First, in Chapter II, I describe a facultatively feeding larva with a reduced pluteus morphology that may feed but can also form a juvenile in the absence of food. This is a rarely documented developmental strategy that has been described in just a handful of marine invertebrate groups, including two urchins and another brittle star (Allen & Pernet, 2007; Emlet, 1986; Hart, 1996). I hypothesized that larval feeding would provide benefits to the larva and carry over post metamorphosis. Chapter II was published in the journal Ecology and Evolution with co-author Dr. Richard Emlet, who provided editorial and manuscript preparation assistance (Nakata & Emlet, 2023). The species from Chapter II is only known in its larval form. We collected all experimental material from the plankton as part of a decade-long project to identify brittle star larvae by DNA barcoding. DNA barcoding has been used to identify marine invertebrate larvae in many taxa, can reveal the presence of rare or difficult to collect species, and greatly increases estimates of regional species diversity (Collin et al., 2020a, 2021; Heimeier et al., 2010; Maslakova et al., 2022; Shanks et al., 2020). In Chapter III, I use DNA barcoding and records collected by R. Emlet and students to investigate the developmental diversity of brittle stars from the northeast Pacific Ocean. I ask if development mode and species identity can be determined 1 5 from larval form. Chapter III is unpublished but will include Dr. Richard Emlet as a co-author, as he contributed all data collected prior to 2018 and will assist with the final manuscript preparation. Brittle stars display a diversity of developmental strategies and larval forms. As such, they have great potential as a system in which to study the evolutionary loss and gain of complex larval morphologies and life history patterns. However, analyses have been conducted in few taxa (Allen & Podolsky, 2007; O’Hara et al., 2019a). Brittle stars display many developmental patterns including obligately feeding (planktotrophic) ophioplutei, facultatively feeding reduced plutei, nonfeeding vitellaria, pelagic direct developers, and brooding (Allen & Podolsky, 2007; Byrne et al., 2008; Hendler, 1991; O’Hara et al., 2019a). In Chapter IV, I built a dataset of developmental pattern and larval form based on data from Chapter III, unpublished data from R. Emlet, and from accounts from the literature. I built a 39-spp. phylogenetic hypothesis to ask if the family Amphiuridae had an ancestor with a feeding larva. Feeding larvae have complex morphologies that aid in swimming and feeding. It may be difficult to re-evolve complex larval structures once lost (Strathmann, 1978b). In Chapter III we observed three planktonic forms from the family Amphiuridae that are classified as abbreviated developers: nonfeeding vitellaria, pelagic direct developers, and reduced plutei. I hypothesized that abbreviated development evolved multiples times in the family and arose as distinct larval forms. To investigate this and other hypotheses on the evolution of development in Amphiuridae I estimated ancestral states for developmental pattern and larval form. I estimated transition rates between states by fitting a series of models to my 1 6 tree. Chapter IV is unpublished but will include Dr. Richard Emlet as a co-author, and he will assist with the final manuscript preparation. Despite their potential use for the study of evolutionary patterns in development, ophiuroids have thus far received little attention due to the lack of molecular and developmental data across taxa. We present our own data for 18 species from the northeast Pacific, with additional data for 13 species from Panama and Australia from co-author R. Emlet. We use this data to characterize the developmental diversity of ophiuroids with planktonic development, and investigate hypotheses of evolution of development in the family Amphiuridae. 1 7 CHAPTER II HAVING CAKE AND EATING TOO: THE BENEFITS OF AN INTERMEDIATE LARVAL FORM IN A BRITTLE STAR AMPHIODIA SP. OPAQUE (OPHIUROIDEA) From Nakata, N. N., & Emlet, R. B. (2023). Having cake and eating too: The benefits of an intermediate larval form in a brittle star Amphiodia sp. opaque (Ophiuroidea). Ecology and Evolution, 13(7), e10298. https://doi.org/10.1002/ece3.10298. Published open access under Creative Commons by Attribution License. Minor editorial changes have been made for dissertation consistency. 1. INTRODUCTION The evolutionary transition in developmental mode from feeding (planktotrophic) to non- feeding (lecithotrophic) larvae is widespread across marine invertebrate taxa, and significantly impacts number and size of offspring (Collin & Moran, 2018; Marshall et al., 2012; Strathmann, 1978b, 1985). Lineages with feeding larvae have given rise to those with nonfeeding larvae in many marine taxa, including in closely related species (e.g., Byrne, 2006; Collin, 2004; Jeffery et al., 2003; Keever & Hart, 2008; Krug et al., 2015; Pappalardo et al., 2014; Waeschenbach et al., 2012), sometimes with great frequency, e.g., at least 15 times in living echinoids (Emlet, 1990). These contrasting patterns signify an evolutionary trade-off between parental investment per offspring and fecundity: feeding larvae develop from smaller eggs that are produced in far 1 8 greater numbers than those of related species with nonfeeding development (Strathmann, 1985). The limited provisions of small eggs require planktotrophic larvae to acquire materials through exogenous food to complete metamorphosis. The larger, more lipid-rich eggs of lecithotrophic larvae contain sufficient material for larvae to create a juvenile without feeding. An intermediate pattern known as facultative planktotrophy (Chia, 1974; Emlet, 1986; Vance, 1973a), where larvae are capable of feeding in the plankton but can also complete metamorphosis without food, is a rare but persistent mode of development that has been observed at least eight times across several marine invertebrate taxa (Table 1; see also Allen & Pernet 2007). Facultative planktotrophy is often considered an intermediate mode of development in the evolutionary transition between feeding and nonfeeding larvae, but whether or not intermediate modes can maximize reproductive success over evolutionary timescales is still up for debate (Christiansen & Fenchel, 1979; Levitan, 2000; McEdward, 1997; Vance, 1973a, 1973b). Larvae with facultative planktotrophy provide an opportunity to test the effects of larval feeding on several aspects of early life history, including planktonic duration, percent metamorphosis, juvenile size, and energetic reserves in juveniles. 1 9 Table 2.1. Effects of larval feeding on development of facultative planktotrophs. Larval and juvenile metrics are represented as increasing (+), decreasing (-), equal (=), or not measured (NM) because of larval feeding. Taxa Develop- Larval Juvenile References ment time survivor- size ship Echinodermata: Echinoidea Clypeaster rosaceus = = + Allen et al., 2006; Emlet, 1986 Brisaster latifrons NM NM + Hart, 1996 Echinodermata: Ophiuroidea Amphiodia sp. opaque - + + This study Macrophiothrix rhabdota - + + Allen & Podolsky, 2007 Mollusca: Gastropoda Adalaria proxima - = NM Kempf & Todd, 1989 Conus pennaceus = = NM Perron, 1981 Phestilla sibogae = + + Kempf & Hadfield, 1985; Miller, 1993 Annelida: Polychaeta Streblospio benedicti NM NM NM Pernet & McArthur, 2006 Arthropoda: Copepoda Tisbe sp. - + + Gangur & Marshall, 2020 Planktonic duration, the time interval between introduction of embryos or larvae to the water column and metamorphosis in the benthos, varies widely across developmental modes of planktonic larvae and affects the composition and distribution of benthic adult populations (Becker et al., 2007; Shanks, 2009; Strathmann, 1985). For feeding larvae, the accumulation of materials necessary for metamorphosis can take weeks to months, whereas nonfeeding larvae can settle in the benthos in hours to days after release (Strathmann, 1987). The larval phase is the dominant dispersal stage for benthic marine invertebrates, and larvae that spend weeks in the plankton tend to disperse further than those that spend only hours in the water column before 2 0 settlement (Shanks, 2009). Realized dispersal distances are modulated by a complex set of temporal, physical, and behavioral factors, but greater interchange of genetic propagules via feeding larvae tends to lead to greater genetic connectivity between benthic adult populations (reviewed in Cowen & Sponaugle, 2009; Shanks, 2009). The increased dispersal potential of feeding larvae influences biogeography: in cone snails, echinoids and cowries, species with feeding planktonic larvae have larger geographic ranges when compared to species with nonfeeding development (Kohn & Perron, 1994; Emlet, 1995; Paulay & Meyer, 2006); and in fossil gastropods, there is evidence that planktotrophic development can positively influence geographical distribution and species longevity (Hansen, 1978, 1980; Jablonski, 1986). Planktonic duration can be mediated by factors both environmental and taxon-specific, but extended planktonic duration generally increases the likelihood of larval mortality prior to metamorphosis (Rumrill, 1990). Low food availability (Miner et al., 2005; Rendleman et al., 2018; Sewell et al., 2004; Strathmann et al., 1992) and low temperature (O’Connor et al., 2007) can slow larval development in echinoids, as well as facultative planktotrophs from some taxa (Allen & Podolsky, 2007; Paulay et al., 1985). Larvae with longer development times relative to conspecifics experienced lower survivorship in two fishes (Hare & Cowen, 1997; Meekan & Fortier, 1996) and increased mortality from predation in a number of invertebrate species (Cowen & Sponaugle, 2009; Rumrill, 1990). Long planktonic durations also expose larvae to the risk of advection away from suitable habitat for settlement (Pineda et al., 2010), although long- lived larvae in coastal habitats may not always disperse widely (Shanks, 2009). In these ways, planktonic duration can be tied to the percentage of larvae in a cohort that survive through metamorphosis, hereafter referred to as percent metamorphosis. 2 1 Metamorphosis does not promise a new beginning: embryonic and larval experiences can be expressed latently in juvenile and adult quality (Emlet & Sadro, 2006; Pechenik, 2006). Food limitation and prolonged planktonic duration have been shown to negatively influence juvenile size, growth, and survival (reviewed in Pechenik, 2018). In facultative planktotrophs, larval feeding resulted in larger juveniles in echinoderms (Allen & Podolsky, 2007; Emlet, 1986; Hart, 1996) and gastropods (Kempf & Hadfield, 1985; Miller, 1993). Juvenile size is an important life history characteristic because larger juveniles tend to exhibit higher survival, growth, reproduction and longevity across several species (Marshall et al. 2018). Here we present evidence for a facultatively planktotrophic larva of a brittle star, Amphiodia sp. opaque (Echinodermata: Ophiuroidea), a previously unknown species that occurs in the northeastern Pacific from Oregon to British Columbia. There is limited documentation of developmental mode within the Ophiuroidea (ca. 12% of ophiuroid species, N. Nakata unpubl. data), but the following developmental modes are known: planktotrophy via an eight-armed ophiopluteus, facultative planktotrophy, lecithotrophy via a reduced pluteus or vitellaria larva that can be pelagic or demersal, and brooding (Hendler 1991; Byrne and Selvakumaraswamy 2002). There are many unresolved relationships within families, but widespread occurrence of developmental diversity suggests frequent transitions from feeding to nonfeeding larvae (Allen & Podolsky, 2007; Lessios & Hendler, 2022; O’Hara et al., 2019a). This study utilizes larvae of intermediate mode of development to assess the effect of larval feeding on planktonic duration, percent metamorphosis, juvenile size, and juvenile energetic reserves. Because our experimental individuals were collected as embryos from the plankton and the adults remain unknown (in 2 2 Oregon), we used DNA barcoding to identify our animals and to reveal their relationship to other ophiuroids in the northeast Pacific. 2. MATERIALS AND METHODS 2.1. Molecular identification of embryos and larvae To investigate species identity of the embryonic morphotypes, we froze one embryo from each year’s cohort of larvae collected for experimentation (2019-2021, see section 2.2) at -20°C in a small volume of seawater. We compared the resulting sequences with those of approximately a dozen larvae of the same morphotype collected over the last decade, as well as with sequences for adults and larvae of other amphiurids from the northeast Pacific. Genomic DNA was extracted with the chelex-based InstaGene™ Matrix (Bio-Rad) and fragments of cytochrome c oxidase subunit I (COI) were amplified and sequenced using the primers jgLCO1490 and jgHCO2198 (Geller et al., 2013). PCR amplification reactions were performed in a 20 µL total reaction volume that included 11.4 µL nuclease-free water, 4 µL 5X Green Buffer, 0.4 µL dNTP 10 mM, 0.2 µL GoTaq Polymerase (Promega), and 1 µL each of forward and reverse primers. PCR conditions were as follows: initial step 95°C for 2 min, followed by 34 cycles of denaturation at 95°C for 40 sec, annealing at 45 or 48°C for 40 sec, and extension at 72°C for 1 min, followed by a final extension at 72°C for 2 min. PCR products were cleaned up using the Wizard SV Gel and PCR Clean up System (Promega) prior to Sanger sequencing (Sequetech, Mountain View, CA). Barcode sequences were compared to GenBank (www.ncbi.nlm.nih.gov/genbank/), BOLD (Ratnasingham & Hebert, 2007), and our unpublished 2 3 dataset of ophiuroid sequences using the BLAST function in Geneious Prime (https://www.geneious.com). We aligned COI sequences of amphiurid spp. of the northeast Pacific (Table 2.2) using the Geneious MAFFT plug-in (Katoh & Standley, 2013) created a maximum likelihood tree using the PhyML plug-in with 100 bootstraps and based on the HKY85 model (Guindon et al., 2010; Hasegawa et al., 1985). Table 2.2. Collection and accession data for specimens included in tree (Fig. 2). Accession numbers refer to records in BOLD (Ratnasingham & Hebert, 2007), unless marked by an asterisk (*), indicating that a record is from GenBank (Benson et al., 2017). Species ID Specimen code Life Collection location, Accession stage State/Province Number Amphichondrius granulatus Agran1 Adult Catalina Is., CA OOPH005-18 Agran2 Adult Catalina Is., CA OOPH006-18 Amphipholis pugetana Ampu1 Adult Cape Arago, OR OOPH003-18 Ampu2 Adult Cape Arago, OR OOPH004-18 Or43-12913 Larva Charleston, OR OLAB015-22 Or811-12115 Larva Charleston, OR OLAB016-22 Amphiodia occidentalis Amoc2 Adult Charleston, OR OOPH002-18 AoMc1a Adult Charleston, OR OOPH020-22 Amoc1 Adult Charleston, OR OOPH001-18 AoMc2a Adult Charleston, OR OOPH022-22 Amphiodia sp. FHLAmphioegg1 Egg Orcas Island, WA OLAB045-22 sensu Emlet 2006 OrR7 Larva Charleston, OR OLAB031-22 Or8P-121812 Larva Charleston, OR OLAB033-22 MMB17 Adult Charleston, OR OOPH032-22 Amphiodia sp. opaque BFHL 4167 Adult Boundary Bay, WA BBPS564-19 Orop3a Larva Charleston, OR OLAB001-22 Opq4 Larva Charleston, OR OLAB006-22 Oop1 Larva Charleston, OR OLAB007-22 2 4 QHAK-00565 Larva Quadra Island, BC QHAK711-21 Opq2 Larva Charleston, OR OLAB004-22 opaque 11-7-18 Larva Charleston, OR OOPH029-22 Opq1 FHL Larva Friday Harbor, WA OLAB047-22 Amphiodia urtica Or988 Juvenile Charleston, OR OLAB028-22 A.urt adt 4-23-19 Adult Cape Arago, OR OOPH026-22 DISA835-19 Adult Los Angeles, CA DISA835-19 HM542069 Adult Bamfield, BC *HM542069 urtica 11-5-18 Larva Charleston, OR OOPH027-22 CCDB-31778 B01 Adult Dana Point, CA ECHCA108-18 KU495782 Adult Queen Charlotte Is., BC *KU495782 2.2. Larval cultures & data collection Adults of Amphiodia sp. opaque have not yet been found in Oregon. We obtained late blastulae, hatched gastrulae, and larvae of Amphiodia sp. from plankton tows collected with a 130 µm mesh net in the Coos Bay estuary (Oregon) approximately 3 km from the entrance to the Pacific Ocean (Charleston Marina: 43°21.2’N, 124°20’W). We collected plankton daily, approximately one hour before high tide and examined our catch within two hours using a stereomicroscope. On several occasions the bright orange embryos characteristic of Amphiodia sp. opaque (Fig. 2.1A, B) were abundant enough to obtain sufficient material for experimentation. In 2020, we collected approximately 120 blastulae or gastrulae, ~1 day post spawn (dps), on February 23 and 100 more of similar stage on February 24. Because of their similar stages of development, embryos from each day represented separate spawning events one day apart. In 2021, we collected 240 embryos of similar stage on March 7. In 2019, no embryos 2 5 were found, but we collected approximately 80 ophioplutei on January 28 and 29. These larvae had either formed juvenile rudiments or did so within a week of capture and were analyzed for juvenile size and starvation time as a ‘wild’ treatment. The ophiuroid hydrocoel rudiment wraps around the larval esophagus and marks the cessation of larval feeding; therefore, we considered larvae with rudiments to have completed or nearly completed larval feeding in their natural environment prior to their collection and monitoring in the lab. Wild ophioplutei and rudiment- stage larvae were kept in filtered sea water (FSW) for the remainder of their development. We did not manipulate microalgal food or estimate planktonic duration for larvae collected in 2019 because we could not determine their prior history in the plankton. 2 6 Figure 2.1. Amphiodia sp. opaque (A) fertilized egg, (B) unhatched blastula, (C) gastrula, (D) early pluteus 3 days post spawn (dps), (E) reduced pluteus 6 dps from no-food treatment, with mouth (m) and empty stomach (st), (F) reduced pluteus 6 dps from food treatment, with algal food visible in the stomach (arrowheads), (G) pluteus at 10 dps from no-food treatment, with three pairs of arms: anterolateral (al), postoral (po), and posterolateral (pl), (H) pluteus 10 dps from food treatment, with juvenile rudiment (r), (I) juvenile from no-food treatment with disc diameter (dd), and (J) juvenile from food treatment. Scale bars are 100 µm; same scale for A-F and for G-J. 2 7 2.3. Feeding experiments For experimental manipulations of microalgal food, we kept embryos (collected in 2020 or 2021) in FSW until just before the formation of the mouth, approximately two days after collection. At this point we haphazardly divided early plutei into replicate finger bowls each with 10 larvae per 30 ml (2020) or 17 larvae per 51 ml (2021) for a standard density of 0.3 larva ml-1. We randomly assigned bowls to ‘food’ or ‘no-food’ treatments and kept them in an incubator at 15°C. We fed larvae in the food treatment a tripartite microalgal diet composed of 2 parts by volume of Rhodomonas lens, to one part each of Dunaliella tertiolecta and Isochrysis galbana at a combined concentration of 5,000 cells ml-1. Larvae in the no-food treatment were kept in FSW alone. Small flakes of cetyl alcohol were added to all cultures to prevent larvae from perishing in the air-water interface. We collected data on stages of larval development every two or three days when FSW and microalgal food were refreshed. We categorized larvae visually as (1) ophiopluteus, (2) rudiment-stage larva, or (3) juvenile (Fig. 2.1). Developmental stages were defined as follows: ophioplutei had up to six larval arms and an open mouth; rudiment-stage larvae had a pair of posterolateral arms, the right anterolateral arm and a well-developed rudiment that occluded the larval mouth; and juveniles lacked larval arms including the larval skeleton and ciliary band, and all locomotion was by their podia. Planktonic duration was defined as the number of days from hatching – estimated as one day prior to the blastula stage in which they were collected – until they were scored as juveniles. We defined percent metamorphosis as the total number of juveniles produced relative to the initial number of larvae from a given bowl. When we found new juveniles, we removed them from the larval culture bowl and put each individual in its own 2 8 35 mm petri dish with FSW. We kept juveniles in FSW without food at 15°C, observing them every two to three days when FSW was changed. A juvenile was considered dead when podia did not move, even following water agitation and the juvenile did not hold onto the dish bottom. 2.4. Analysis We conducted all statistical analyses in the R environment v3.6.0 (R Core Team, 2022), and visualized plots with the package ‘ggplot2’ (Wickham, 2016). For each statistical test to follow, apart from the survival analysis of juveniles, we considered the finger bowl to be the experimental unit. We calculated the percent of larvae that successfully metamorphosed into juveniles (hereafter percent metamorphosis), and mean values for planktonic duration, juvenile size, and juvenile starvation time for each bowl. We calculated survival statistics in two ways: by bowl (Fig. 2.6B) or pooled within treatments and years in survival analysis (Fig. 2.7). To quantify the effect of larval food on percent metamorphosis, we compared between treatments, keeping years separate. As values for percent metamorphosis were not normally distributed and did not have equal variances (Shapiro-Wilk, p = 0.015; Levene’s test, p = 0.004), we used a non-parametric Kruskal-Wallis test, followed by a Dunn’s test for pairwise comparisons. We used aboral surface area of the disc as our value for juvenile size. We measured disc diameter (dd, Fig. 2.1I), the length from one arm tip through the center of the disc to the opposite interradius, using the ocular micrometer on the dissecting microscope (dh, Fig. 2.1I), measured to the nearest 20 microns. Using the disc diameter as the height of a regular pentagon (h, 2 9 Equation 1), we solved for the length of a single side (a) in order to calculate the area of the pentagonal juvenile disc (A, Equation 2). (1) ℎ = 𝑎 ∗ &5 + 2√5 / 2 (2) 𝐴 = 𝑎! ∗ &25 + 10√5/4 The data for juvenile aboral surface area did not satisfy the assumption of normality (Shapiro-Wilk, 2020 p = 0.001, 2021 p = 0.006) but variances were not significantly different (Levene’s test, 2020 p = 0.97, 2021 p = 0.09). To determine if the presence of microalgal food had an effect on juvenile size, we compared aboral surface area across treatments using a non- parametric Kruskal-Wallis test, followed by a post hoc Dunn’s test. To determine whether larvae that developed more quickly also produced larger juveniles, we modeled the association between planktonic duration and juvenile aboral surface area using a set of generalized linear models (GLMs) with a Gaussian family, including combinations of potential covariates including treatment and year. To examine the relationship between juvenile size and survival, we modeled the association between juvenile aboral surface area and time to starvation using a set of GLMs and combinations of covariates treatment, year, and their interaction. As we predict time to juvenile starvation represents a waiting time, we used GLMs with a gamma distribution. We assessed model fit using Akaike’s and Bayesian Information Criteria (Table S4), with the lowest value indicating the best fit. To assess differences in juvenile survival time between treatments and years, we used survival analysis. The survival time data violated the assumptions of parametric statistics, rendering traditional tests inappropriate for their analysis. Such data can be analyzed using non- 3 0 parametric survival analysis (Kleinbaum & Klein, 2012; Moore, 2016), a collection of statistical procedures for which the outcome variable of interest is time until an event occurs. Juvenile starvation times were analyzed using survival analysis in the R package ‘survival’ (Therneau, 2020) and visualized with the package ‘survminer’ (Kassambara et al., 2020). Most survival analyses must contend with censoring, which occurs when we have some information about individual survival time, but we don’t know the survival time exactly. Our data on time to juvenile starvation are ‘interval-censored’, because counts were done at intervals when water changes happened every two or three days. The exact time of metamorphosis or death is unknown and can only be placed within a specified window of time. Censoring can also occur when an individual withdraws from the study prior to the time of the event. In our analysis of juvenile starvation, the event of interest was death, and no juveniles were censored as all were followed until the time interval of the event. Survival curves for juvenile death probabilities were compared using a non-parametric log-rank test. 3. RESULTS 3.1. Species identity and occurrence of larvae The opaque orange larvae studied here belong to an undescribed species of Amphiodia that occurs at least from southern Oregon to British Columbia. On many occasions since March 2003, we have collected eggs, unhatched and hatched embryos, and larvae from Oregon plankton that we assigned, by color and morphology to A. sp. opaque. These samples were collected 3 1 between January and April, but on one occasion embryos were collected in November (2018). We also collected these larvae in the Salish Sea (Friday Harbor, Washington) in August of 2019 and September of 2020. We have generated COI- barcodes for more than 15 specimens collected from 2013 to 2019 that we assigned to Amphiodia sp. opaque, and their sequences are 98+ % similar (Fig. 2.2). During the times we collected these embryos and larvae we never found similar orange embryos or larvae that had different barcodes and were other species. The sequences of our larvae are a close match (>98% pairwise nucleotide similarity) to that of a juvenile amphiurid specimen (2.5 mm disk diameter) collected from Boundary Bay WA (BOLD specimen record: BBPS564-19). The juvenile is clearly a member of the genus Amphiodia, because it has the appropriate pattern of 3 similarly sized oral papillae per jaw. In BOLD we found another close match (>99%) to a larva with the same morphology from Hyacinthe Bay, Quadra Island, British Columbia (BOLD specimen record: QHAK711-21, Fig. 2.2). A neighbor-joining tree of COI sequences from larvae and adults of other amphiurid species known from the NE Pacific shows A. sp. opaque as the sister species to A. urtica which has obligately feeding larvae (Schiff & Bergen 1996, N. Nakata, unpubl. data; Fig. 2.2). Together these clades are sister to a species complex of A. occidentalis (R. Emlet, unpubl. data) 3 2 Amphichondrius granulatus 99 Amphipholis pugetana feeding ophiopluteus 100 Amphiodia sp. sensu pelagic 100 Emlet 2006 direct99 development 100 Amphiodia occidentalis 91 100 Amphiodia urtica 69 feeding ophiopluteus 100 Amphiodia sp. opaque facultative planktotroph Figure 2.2. Maximum likelihood tree of COI sequences from adult and larval amphiurid spp. in the northeastern Pacific, constructed using the PhyML plug-in in Geneious. Sequences from GenBank (*) and BOLD are labeled with accession number (and see Table 2). Development modes are indicated on the right, with characteristic larvae depicted. Bootstrap values are shown next to nodes. 3.2. Larval developmental mode Larvae of Amphiodia sp. opaque develop from eggs of moderate size (140 µm) and are facultative planktotrophs (Fig. 2.1): larvae can feed but do not require food for metamorphosis 3 3 into juveniles. While percent metamorphosis did differ between years, some larvae developed into juveniles in the absence of food. We observed larvae from unfed culture with empty stomachs, and those from fed cultures with stomachs full of microalgal food (Fig. 2.1E and F). Furthermore, aspects of larval and juvenile performance differed according to larval food treatment (Table 2.3). 3.3. Effects of larval food on developmental timing Fed larvae reached metamorphosis more quickly than unfed larvae (Fig. 2.3). Planktonic duration differed between years but was about a week shorter for fed larvae than for larvae raised in FSW (2020 median 17 days vs 23.5 days; 2021 median 13 days vs 22 days; see Table 2.3). Table 2.3. Summary of traits for larval and juvenile Amphiodia sp. opaque. Larvae were raised with and without microalgal food in 2020 and 2021 and collected as late-stage plutei in 2019. Planktonic duration is given for all larvae that completed metamorphosis. Planktonic duration and time to juvenile starvation are time data and are listed as minimum, median, and maximum. Percent metamorphosis and juvenile aboral surface area are given as means ± standard error for replicate bowls and for individuals, respectively. Year Treatment Total larvae / Planktonic Juv. n Percent Juv. aboral surface Time to # replicate duration meta- area (mm2) starvation bowls (days) morphosis (days) 2019 Wild 80 / 7 - 46 - 0.032 ± 0.001 0–6–38 2020 Food 110 / 11 15–17–21 88 80 ± 3 0.046 ± 0.001 2–59–104 No-food 110 / 11 20–23.5–30 16 14 ± 6 0.030 ± 0.001 0–48.5–76 2021 Food 120 / 7 13–13–19 54 45 ± 4 0.047 ± 0.001 2–23–65 No-food 120 / 7 15–22–33 64 53 ± 3 0.032 ± 0.001 2–18.5–54 3 4 Figure 2.3. Cumulative sum of juveniles over time by treatment and year: (A) 2020, (B) 2021. Each year had the same initial number of larvae in each treatment (2020 n=110 per treatment, 2021 n=120). Final time points represent date of discovery of the last juvenile in that treatment. 3.4. Effects of larval food on percent metamorphosis The effect of treatment on the percent of larvae that completed metamorphosis differed significantly in 2020 but not in 2021 (across all treatments and years: Kruskal-Wallis test: χ2 = 29.25, df = 3, p < 0.001; Fig. 2.4). Pairwise comparisons show a significant difference between treatments in 2020 (Dunn’s test: adj. p < 0.001, Table S2.1), when food had a strong effect: 80% of fed larvae created a juvenile whereas only 15% of unfed larvae were able to do so (Fig. 2.4). By contrast, in 2021 slightly more juveniles resulted from cultures without larval food (food: 45%, no-food: 53%), and percent metamorphosis was not statistically different between treatments (p = 0.44). 3 5 Figure 2.4. Bar plots of percent metamorphosis (= # juveniles / initial # larvae) averaged across experimental bowls for years 2020 (initial larvae n=220/22 bowls) and 2021 (n=240/14 bowls). Error bars represent standard error (SE). Lowercase letters above bars represent significant differences in pairwise comparisons. 3.5. Effects of larval food on juvenile size Juvenile sizes measured as aboral surface area were significantly different between treatments in both years (Kruskal-Wallis test: χ2 = 26.149, df = 4, p < 0.0001). Pairwise comparisons showed that juveniles from fed larvae were larger than juveniles from unfed larvae in both years. Juveniles from wild larvae were not significantly different in size from juveniles from the no-food treatment (Dunn’s test, Table S2.2; Fig. 2.5). We modeled the association between planktonic duration and juvenile aboral surface area (Fig. 2.6A) using a series of GLMs and found that the best fit model included only treatment as a covariate (AIC = -288.3, Table S2.3). 3 6 Figure 2.5. Boxplot of juvenile aboral surface area (SA) at metamorphosis by food treatment, pooled across years. Lowercase letters above boxplots represent significant differences in pairwise comparisons. 3.6. Effects of larval food on juvenile survival The survival time in juveniles that received food as larvae was longer compared to those that did not receive food, but survival times varied between years (2020 median 59 days (n=88 vs 48.5 days, n=16; 2021 median 23 days, n=54 vs 18.5 days, n=64, Table 2.3). Juveniles that resulted from wild-caught larvae had relatively short survival times under starvation conditions (median 6 days, n=80). Bigger juveniles tended to survive longer, and juveniles from fed larvae had greater surface areas than those from larvae that did not receive food (Fig. 2.6B). Juvenile aboral surface area was positively associated with days to juvenile starvation and the best-fit model included an 3 7 Figure 2.6. Scatterplots of (A) planktonic duration by juvenile size, and (B) juvenile aboral surface area (SA) by time to juvenile starvation. Points are mean values for replicate finger bowls (larval culture containers) and bars are standard error. Treatment is coded by color (food: dark gray, no-food: white, wild: ‘w’) and experimental year by shape (2020: circle, 2021: triangle, wild: ‘w’). interaction between treatment and year (GLM, AIC 1672.1; Table S2.4). Treatment (food, no- food) and experimental year (2020, 2021) were both significant covariates (p = 0.02 and p < 0.001, respectively). The survival curves for each treatment and year were significantly different from each other (log-rank test, p < 0.0001) and were significantly different in each pairwise comparison (p < 0.0001. Fig. 2.7). 3 8 a b c d e Figure 2.7. Kaplan-Meier survival curves for Amphiodia juveniles according to larval food treatment and year. No juveniles were censored as they were followed until the time of the event, death. Lowercase letters below survival curves indicate significantly different pairwise comparisons. 4. DISCUSSION 4.1. Species identity Through DNA barcoding of the COI gene, we found a single species-level match (>98% pairwise nucleotide similarity) to our Amphiodia larvae in a juvenile specimen from Boundary Bay, WA (BBPS564-19, provided by G. Paulay), which was difficult to assign to species from its morphology. The specimen was small (2.5 mm disk diameter) and the adult characteristics necessary for identification may have been absent or reduced, as they often are in small 3 9 specimens (Stöhr, 2005). The specimen had three oral papillae of equal size and spacing, as is characteristic of Amphiodia. Its oral shields are pentagonal in shape, and the arm spines are tapered to a point like in A. urtica, but the radial shields are approximately 2 times as long as they are wide, the dorsal arm plates are oblong and the ventral arms plates are squarish like in A. occidentalis (Lambert & Austin, 2007). Furthermore, molecular data for COI delineates this specimen from adults of A. occidentalis or A. urtica (Fig. 2). Rather than signifying a new species, the lack of a molecular match with morphological identification may be due to the limited genetic sampling of the genus Amphiodia (8 of 34 species, N. Nakata unpubl. data) and of amphiurids of the northeast Pacific (5 of 12 species, N. Nakata unpubl. data). 4.2. Larval morphology and developmental mode Using feeding assays, we confirmed that Amphiodia sp. opaque is a facultative planktotroph. Amphiodia sp. opaque has eggs of moderate size (140 µm), which is consistent with other ophiuroids with abbreviated development (Hendler, 1991). Larvae of this species developed into juveniles in the absence of microalgal food but were still capable of planktonic feeding. Percent metamorphosis varied across treatments and study years, but 15 to 53% of larvae given no microalgal food successfully completed metamorphosis, supporting our diagnosis of facultative planktotrophy. This developmental mode cannot be diagnosed from larval morphology alone because larvae retain the structures necessary for feeding. Larvae must be cultured in the presence and absence of microalgal food to determine developmental mode. We did suspect facultative planktotrophy of this larva because it was opaque orange in color and has a reduced pluteus 4 0 morphology (Hendler, 1975), as determined by comparison with a sympatric congener A. urtica, which has a transparent, planktotrophic larva with 8-arms. The posterolateral arms are reduced in length relative to those of A. urtica, and the posterodorsal arms are highly reduced or absent, resulting in 6 larval arms instead of 8. Larvae of Amphiodia sp. opaque have other features that are associated with evolutionary transitions in developmental pattern, including an egg of intermediate size (ca.140 µm) and intermediate planktonic duration. Facultative planktotrophy has been observed in six other marine invertebrates (Table 2.1). Amphiodia sp. opaque is the second facultative planktotroph to be described from the Ophiuroidea and the first from the family Amphiuridae. Only one ophiuroid was previously known to have this developmental pattern: Macrophiothrix rhabdota (Ophiotrichidae) develops via an eight-arm pluteus. The family Ophiotrichidae diverged from Amphiuridae approximately 200 Mya (O’Hara et al., 2017), meaning Amphiodia sp. opaque represents an independent evolution of the facultative planktotroph phenotype. Food-limited growth is common for larvae of benthic invertebrates (Paulay et al., 1985), and facultative planktotrophy may have evolved as a bet-hedging strategy to increase the number of resulting juveniles in regions or spawning times when planktonic food availability is low. 4.3. Development time Fed larvae developed more quickly than larvae that did not receive food. Larvae in the food treatment completed metamorphosis in fewer days than their starved peers (Fig. 2.2). This is consistent with echinoids with feeding larvae, which are known to take longer to form their 4 1 rudiment in no- and low-food conditions (Miner et al., 2005; Sewell et al., 2004; Strathmann et al., 1992). Compared to congenerics of contrasting development modes, Amphiodia sp. opaque has an intermediate planktonic duration (medians of 13 to 17 days when fed). The planktotroph Amphiodia urtica has a median planktonic duration of 20 days (n=10 larvae collected at different times, N. Nakata, unpublished data) and a congeneric, pelagic, direct-developer may be planktonic for 8 days at 15°C (Emlet, 2006). The availability of food for larvae does not always decrease time to metamorphosis in facultative planktotrophs, For example, there was no difference between fed and unfed treatments in the echinoid Clypeaster rosaceus or the gastropods Conus pennaceus and Phestilla sibogae (Emlet, 1986; Kempf & Hadfield, 1985; Perron, 1981). In the ophiuroid Macrophiothtrix rhabdota planktonic duration was decreased in the presence of food (Allen & Podolsky, 2007). Shorter planktonic interval may limit mortality from predation in the plankton (Rumrill, 1990) and may reduce capacity for dispersal in species with shorter developmental times (Hendler, 1991; Shanks, 2009). We interpret larval developmental times as dependent on the accumulation of energetic reserves necessary for metamorphosis and juvenile life, but other factors may have contributed to planktonic durations observed in this study. Little is known about cues for competency or settlement in ophiuroids (Hendler, 1991; Hodin et al., 2015). Furthermore, ophiuroids are known to be capable of undergoing metamorphosis in the plankton and continuing to ride ocean currents as juveniles (Hendler et al., 1999), indicating that some species may not require benthic cues to begin metamorphosis once sufficient food has been consumed in the plankton. 4 2 4.4. Percent metamorphosis The effect of larval feeding on the proportion of larvae that completed metamorphosis was different across years for Amphiodia sp. opaque. In 2020, food had a strong effect on the proportion of larvae able to complete metamorphosis, as was true for the other facultatively planktotrophic ophiuroid (Allen & Podolsky, 2007). In 2021, percent metamorphosis was not significantly different between treatments and slightly more juveniles resulted from the no-food treatment (Figs. 2.2, 2.3). Amongst facultative planktotrophs, larval feeding did not affect percent metamorphosis (referred to as larval survival) in an echinoid (Emlet, 1986) and two gastropods (Kempf & Todd, 1989; Perron, 1981). It is possible that the observed variation between cohorts resulted from differences in culture conditions between years, or from intrapopulation or interannual variation in parental investment and developmental regimes. As all embryos originated from the plankton, we are unable to determine if differences between cohorts reflect variation in maternal provisioning due to nutrition or genetic variation between populations. 4.5. Juvenile size We observed that fed larvae of Amphiodia sp. opaque produced juveniles that were ca. 50% larger in surface area than those of starved larvae (Table 2.3; Fig. 2.5). Increased juvenile size as a consequence of larval feeding has been observed in other facultative planktotrophs 4 3 (Allen & Podolsky, 2007; Emlet, 1986; Hart, 1996; Kempf & Hadfield, 1985; Miller, 1993). Juveniles of Amphiodia sp. opaque from the fed treatment were slightly smaller than those of a sympatric congener with feeding larvae, A. urtica (juvenile aboral surface area 0.047 mm2, n = 18; N. Nakata unpubl. data). Advanced larvae collected from wild plankton (2019) produced juveniles that were not significantly different in size from those from the experimental no-food treatment. This suggests that these larvae had limited opportunity to feed in their natural environment (and they were not fed in the lab). Even low-food conditions can lead to smaller juveniles in a calyptraeid gastropod (Chiu et al., 2007, 2008). Evidence from a barnacle has shown that even food deprivation during a portion of the larval life can lead to smaller juveniles with the early stages being the most important (Emlet & Sadro, 2006). 4.6. Juvenile survival Juveniles of fed larvae lived longer than those from larvae raised without food, suggesting that they gained greater energetic reserves due to larval feeding. However, juvenile survival times differed among treatments and experimental years (Fig. 2.7). Juveniles from both treatments in 2021 had shorter survival times than juveniles from either treatment in 2020. Interestingly, the juveniles that resulted from wild-caught larvae from 2019 fared the poorest of all. They were no different in size from juveniles of lab-reared larvae that received no food (Fig. 2.5), but they died from starvation more quickly (Fig 2.6B, 2.7). This may have been the result of variation in egg composition or size, but whether that was due to intraspecific genetic variation, 4 4 local environmental deficiencies that lowered maternal condition, or some combination was not determined. 5. CONCLUSION In this study, we tested the influence of larval feeding in an unidentified facultatively planktotrophic larva, Amphiodia sp. opaque. We know this animal only in its larval form, the adults have not been collected. We utilized phylogenetic analysis in an attempt to determine species identity and to compare it with closely related ophiuroids from the NE Pacific. Nevertheless, we were able to conduct a series of experiments that showed clear benefits of larval feeding across multiple life history characters: larvae that received food developed more quickly, experienced higher rates of metamorphosis, and produced larger juveniles that evaded starvation conditions for longer than those of larvae that received no food. BRIDGE In Chapter II, I described a facultative planktotroph from the ophiuroids of the northeast Pacific that can feed but can also complete metamorphosis without food. The larva took the form of a reduced pluteus, a relatively rare larval form amongst the brittle stars that may represent an intermediate larval form between obligately feeding ophioplutei and nonfeeding larvae. In the next chapter I built upon Chapter II’s results by describing the early life stages of brittle stars from Oregon, which included several more reduced plutei. 4 5 CHAPTER III BRITTLE STAR LARVAE OF THE NORTHEAST PACIFIC This work includes coauthor Dr. Richard Emlet as principal investigator and contributor to the final manuscript preparation. It is written in the journal style of Invertebrate Biology. 1. INTRODUCTION With ca. 2100 described species, the brittle stars (Ophiuroidea) are the most diverse class of echinoderms and dominate benthic environments across all ocean depths (O’Hara et al., 2019b; Stöhr et al., 2012). Ophiuroids have a wide variety of ecological roles, and when abundant are significant contributors to benthic communities. Brittle stars exhibit diverse reproductive patterns including planktonic feeding and nonfeeding larvae, brooding, and fissiparity. These variations in development are distributed across Ophiuroidea, and closely related species can have very different ontogenies (Allen & Podolsky, 2007; Hendler, 1991; O’Hara et al., 2019a). Brittle stars have great potential for the study of life history evolution, but such evaluations are limited by the available data on phylogenetic relationships (9% of spp.) and data on developmental patterns (15% of spp., Nakata, unpublished). Of the brittle stars for which development is known (approx. 275 species), about half brood their young; the remainder have planktonic larvae. We used DNA barcoding to identify brittle star larvae collected from plankton. Barcoding of all life stages can lead to higher estimates of regional species diversity (Maslakova et al., 2022). Here, barcoding allowed us to identify or place within a phylogeny planktonic larvae for a group of organisms that has proven difficult to study in a laboratory setting. Unlike echinoids, ophiuroids do not have reliable means for inducing spawning in the laboratory so raising larvae of known species is not readily accomplished. Brittle star larvae are diverse in form but are generally divided into two categories: planktotrophs, with an obligately feeding and slow-growing ophiopluteus that comes from small eggs produced in large quantities; and abbreviated developers, who develop from moderate numbers of yolky eggs and can take a variety of planktonic forms, including reduced plutei, vitellaria, and pelagic direct developers (Hendler, 1991). We used morphological and developmental characters (naming scheme after Mortensen, 1921) to describe ophiuroid larvae identified by DNA barcoding. The characteristic feeding larva of ophiuroids is the ophiopluteus, a bilaterally symmetrical larva that usually has eight arms bearing a ciliated band and supported by calcite rods (Fig 3.1A). The longest and most prominent pair of arms are the posterolateral arms. The anterolateral arms extend anteriorly and support the mouth and adjacent ciliary band. The postoral arms extend ventrally to hold up a ciliary band in below of the mouth. Finally, the posterodorsal arms originate from the posterior portion of the anterolateral arm rods and project dorsally. In many species the postoral and posterodorsal arms are mirror imaged across the frontal plane, making it difficult to see both pairs at once. The larval arms are supported by skeletal rods composed of calcite, and whose shape, proportions, and adornments can be used to distinguish between species (Mortensen, 1921). Though branching into multiple rods to give the distinctive shape of the larval arms, there are only two major calcite spicules arranged in mirror 4 7 image (across the sagittal plane) in the ophiopluteus. A third larval spicule is reported in rare cases, see (Mortensen, 1921). Each spicule has rods that support 4 arms on its side of the larva. The posterolateral, anterolateral, and postoral rods join at two posterior junctions on either side of the stomach, and are connected to the body rods, which extend posteriorly and support the bottom half of the larva. Posterior to the stomach, the body rods each bear dorsal and ventral transverse rods, which extend to the larval midline and contact along the larval midline with their counterparts from the opposite spicules. Some ophioplutei have pairs of recurrent rods that connect the postoral and ventral transverse rods on the ventral side, and the anterolateral and dorsal transverse rods on the dorsal side of the larva. Posterior to the transverse rods, the body terminates in end rods. The transverse rods may bear one more median process(es). We refer to the collection of calcite rods at the posterior of the larva as the posterior girdle (Fig. 3.1A) Ophiuroids with abbreviated development usually develop to juvenile more quickly than species with feeding plutei and usually have small post-larvae produced from moderate numbers of yolky eggs. Abbreviated developers are a developmentally heterogenous group taking the form of reduced plutei, vitellaria, demersal larvae, or modified larvae that undergo metamorphosis in an attached fertilization envelope (Hendler, 1975). We have also observed several pelagic direct developers (Emlet, 2006). Reduced plutei have a modified or reduced morphology, and may have two, four or six arms and opaque coloration (Allen & Podolsky, 2007; Fenaux, 1963; Mladenov, 1979). Reduced plutei are often lecithotrophic, and lack digestive structures, but some may be facultative planktotrophs (Allen & Podolsky, 2007; Nakata & Emlet, 2023). 4 8 The typical nonfeeding larva of ophiuroids is the vitellaria (Fig. 3.1C), though some authors include doliolaria (McEdward & Miner, 2001). We distinguish doliolaria as being barrel- shaped and having a series of transverse ciliary rings, while vitellaria are dorsoventrally compressed and may have complete ciliary rings at the anterior and posterior, but the mediary ciliary bands are disjunct (Fig. 2.1 C). The vitellaria lacks a mouth, stomach, and anus as it does not feed. Vitellaria larvae have been observed across many ophiuroid families (Hendler, 1991), and are believed to be derived from an ophiopluteus, which is supported by the presence of bilaterally paired larval spicules in some vitellaria (Hendler, 1982, 1991; Mortensen, 1921; Selvakumaraswamy & Byrne, 2023). Furthermore, in one species a feeding ophiopluteus transforms into a vitellaria prior to metamorphosis (Mladenov, 1985b). In ophiuroids there are several variations of direct development, in which the embryo develops into a pentaradial juvenile without first developing a larval body such as an ophiopluteus or vitellaria. The most common form of direct development in brittle stars is brooding, in which embryos and young juveniles develop in the disc of the adult until they crawl away as advanced juveniles. Another, rarer type is pelagic direct development, in which embryos and early juvenile stages are planktonic (Emlet, 2006; Hendler, 1973; Patent, 1970). In this study we describe the planktonic forms for eighteen species of ophiuroids found in the northeastern Pacific. Our primary sampling location was Coos Bay, Oregon, but we have also observed several of the same larvae described here in the San Juan Islands, Washington. As many as 66 species of ophiuroids are known to occur from California to Alaska (Astrahantseff & Alton, 1965; Hendler, 1996; Kyte, 1969; Lambert & Austin, 2007), of which 25 have records from Oregon. Of those, we observed 11 species, nine of which occur throughout the range. Many 4 9 Figure 3.1. Developmental stages of the Ophiuroidea. (A) Eight-armed ophiopluteus, oral view, with posterolateral (pl), anterolateral (al), postoral (po), and posterodorsal (pd) arm pairs, which support ciliary bands (stippled regions) and mirror each other across the midline. These arms are supported by skeletal rods (solid black lines). The pl, al, and po arm rods on each side grow from a single calcite spicule that extends posteriorly as a body rod (br) and in some species a recurrent rod (rr), terminating as an end rod (er). The left and right spicules (with their arm rods) are in contact at the posterior end where short transverse rods (tr) meet along the midline. The feeding larva has a mouth (m), esophagus (e), and stomach (s), intestine, and anus. (B) Rudiment-stage pluteus, oral view with right and left posterolateral arms and the right anterolateral arm (ral). The larva has a pentagonal juvenile rudiment (jr) with buds for tube feet (tf). (C) The nonfeeding vitellaria, oral view, has several disjunct ciliary bands and complete bands at the anterior (acb) and posterior (pcb). of the species we identified from the plankton occur nearshore. We observed three species that are not listed to occur in Oregon: Amphiura arcystata, Ophiothrix spiculata, and Ophiocten 5 0 hastatum. The species that are listed to occur in the region but that we didn’t observe in the plankton have subtidal depth distributions from 50 to 3000 m. Two notable species absent from this study are Amphipholis squamata and Amphiodia occidentalis. The former is a brooder, which we find intertidally at Cape Arago, and the latter is a well-known species from the west coast of North America that is part of a species complex (see Section 3.4.3.6). Finally, four additional larvae lack molecular matches to adults. 2. METHODS 2.1. Collection of larvae and adults We collected embryos, larvae, and juveniles of local ophiuroids from plankton tows in years 2018-2021. We collected samples in these years with 130 µm mesh net in the Coos Bay estuary (Oregon) approximately 3 km from the entrance to the Pacific Ocean (Charleston Marina: 43°21.2’N, 124°20’W). We collected plankton nearly daily, especially in the winter months, approximately one hour before high tide and examined our catch within two hours using a stereomicroscope. As many planktonic stages of brittle stars are negatively buoyant and poor swimmers, we most often found our quarry at the bottom of a given plankton sample. All brittle star material derived from the plankton was sorted into phenotypes and select individuals were preserved and analyzed using DNA barcoding. To determine species identities of planktonic life stages we compared our sequences with those available from public databases. However, we found that many of our local brittle star taxa were absent from such databases, and thus we had 5 1 to collect our own adults. Ultimately, we barcoded 102 embryos, larvae, and juveniles, and 53 adult brittle stars. When possible, larvae were kept alive and observed through development into a juvenile. We maintained larvae in glass finger bowls at 15°C in an incubator. Feeding ophioplutei were given a tripartite microalgal diet composed of Rhodomonas lens, Dunaliella tertiolecta, and Isochrysis galbana at a combined concentration of 5,000 cells ml-1. Nonfeeding larvae were kept in filtered sea water (FSW) that was changed every 2 to 3 days. Small flakes of cetyl alcohol were added to all cultures to prevent larvae from perishing in the air-water interface. Embryos, larvae, and juveniles were photographed across development using a Zeiss Universal (Carl Zeiss, Inc., Thornwood, NY) coupled with a digital camera (Flir Grasshopper Express GX-FW-28S5C- C) and software (Astro IIDC) to capture images. For the purposes of describing the various developmental stages, we have divided development into the following stages: the early pluteus is from the first sign of bilateral symmetry, usually accompanied by paired calcite spicules in the posterior of the larva, up through the four- and six-armed stages (i.e., posterodorsal arms are absent). Reduced plutei also fit this description but can be distinguished from planktotrophic plutei by their opaque pigmentation and that they do not ever develop the eighth pair of (posterodorsal) arms. Late plutei are recognized by their eight arms (or six in reduced plutei) and visible coeloms, especially the left coelom, which develops five lobes that are the precursor to the water vascular system. We consider larvae as ‘rudiment-stage’ when the left coelom has wrapped around the larval esophagus, a pentaradial juvenile (or parts of its skeleton) are apparent, and one or more of the larval arms has been resorbed. Finally, juveniles have pentaradial symmetry, are free of any 5 2 remnant of the larval body such as skeletal rods or ciliated bands, and locomote via their tube feet. We also reviewed records of daily samples taken at high tide from January to March during each of years 2014, 2015 and 2016 with a diaphragm pump (Emlet, Shanks, and Sutherland, unpublished data; Shanks et al., 2020). In 2014 this sampling coincided with the warm water blob (Bond et al., 2015) that resulted in northward shifts in species ranges from central and northern California to Oregon and beyond (Sanford et al., 2019). We conducted all data validation and plot visualization in the R environment v4.2.2 using the packages ‘ggplot2’ and ‘ggtree’ (R Core Team, 2022; Wickham, 2016). 2.2. Molecular identification of embryos and larvae To verify species identity of planktonic early life stages, we compared our sequences with those available for ophiuroids of the northeast Pacific from public databases, such as GenBank and BOLD (Benson et al., 2017; Ratnasingham & Hebert, 2007) as well as those derived from adults we collected locally. Adult material originated from several sources, including collection by hand in the intertidal and dredging soft-bottomed subtidal habitats. Adults were relaxed in a 50/50 solution of magnesium sulfate or magnesium chloride and FSW for morphological examination, relying on regional guides (Hendler, 1996; Lambert & Austin, 2007). A few podia or an arm tip were removed with fine forceps for DNA extraction. We barcoded select individuals of each larval morphotype and compared sequences to those for adult specimens. Larvae were usually frozen individually in a small amount of seawater at -20°C, until DNA extraction and PCR. We extracted genomic DNA with the Chelex-based 5 3 InstaGene™ Matrix (Bio-Rad) after rinsing each larva in nuclease-free water. We amplified fragments of cytochrome c oxidase subunit I (COI) and 16S rDNA using a variety of primers and thermocycler conditions (Table 3.1). PCR amplification reactions were performed in a 20 µL total reaction volume that included 11.4 µL nuclease-free water, 4 µL 5X Green Buffer, 0.4 µL dNTP 10 mM, 0.2 µL GoTaq Polymerase (Promega), and 1 µL each of forward and reverse 10 µM primers. PCR conditions were as follows: initial step 95°C for 2 min, followed by 34 cycles of denaturation at 95°C for 40 sec, annealing at 45 or 48°C for 40 sec, and extension at 72°C for 1 min, followed by a final extension at 72°C for 2 min. PCR products were cleaned up using the Wizard SV Gel and PCR Clean up System (Promega) prior to Sanger sequencing (Sequetech, Mountain View, CA). Barcode sequences were compared to GenBank (www.ncbi.nlm.nih.gov/genbank/), BOLD (Ratnasingham & Hebert, 2007), and our unpublished dataset of ophiuroid sequences using the BLAST function in Geneious Prime (https://www.geneious.com). We aligned sequences using the Geneious MAFFT plug-in Table 3.1. PCR primers and reaction conditions for molecular identification of ophiuroid larvae. Locus Primer Name Primer Sequence Reference COI jgLCO1490 TITCIACIAAYCAYAARGAYATTGG (Geller et al., jgHCO2198 TAIACYTCIGGRTGICCRAARAAYCA 2013) EchinoF1 TTTCAACTAATCATAAGGACATTGG (Ward et al., EchinoR1 CTTCAGGGTGTCCAAAAAATCA 2008) COIcef ACTGCCCACGCCCTAGTAATGATATTTTTTATGGTNATGCC (Hoareau & COIcer TCGTGTGTCTACGTCCATTCCTACTGTRAACATRTG Boissin, 2010) 16S 16SARL CGCCTGTTTATCAAAAACAT (Palumbi, 1996) 16SBRH CCGGTCTGAACTCAGATCACGT 16Sar GCCTGTTTACCAAAAACAWCG (Kirby & 16Sbr GATCCAACATCTAGGTCGC Lindley, 2005) 5 4 (Katoh & Standley, 2013) created family- or superfamily-level maximum likelihood trees using the PhyML Geneious plug-in or online execution (Guindon et al., 2010), with 100 bootstraps and based on the HKY85 model of sequence evolution. Family names are after O’Hara et al. (2017). For species delimitation, we used the online execution for Assemble Species by Automatic Partitioning (ASAP) to determine the barcoding gap for COI of ophiuroid species (Puillandre et al., 2021). 3. RESULTS and DISCUSSION 3.1. Taxonomic Diversity We collected, raised larvae, and generated barcodes for 18 species from seven families of ophiuroids (Table 3.2). The best represented families were Amphiuridae with eight species, and Ophiuridae with four species. We determined species identity of larvae by creating phylogenetic trees of our larval sequences amongst barcodes for adult specimens or available from public databases. We considered sequence pairs to be molecular matches if they had less than an 8.5% difference in pairwise identity at the COI locus, a threshold we determined using ASAP analysis. See each section for the tree and specimen information therein. 3.2. Developmental Diversity We observed embryos of only a few species, indicating that their natal populations may be close to shore: Amphiodia periercta?, Amphiodia sp. opaque, Amphiodia sp. orange belly, 5 5 Table 3.2. Ophiuroid species of southern Oregon with planktonic development. Species marked with an asterisk (*) are not regularly found in the waters near Coos Bay, OR. Larval forms are planktotrophic (=feeding) ophiopluteus (P), reduced pluteus (RP), pelagic direct (D), and indirect development via a vitellaria (V). Taxonomy Larva Spawning season Planktonic duration (days) Ophiotrichidae Ophiothrix spiculata* P December ? Ophiopholidae Ophiopholis kennerlyi P Sept-Nov; Mar-Apr 83-216 Ophiopholis bakeri P Oct-Jan ? Amphiuridae Amphiodia urtica P Oct-May 20 Amphipholis pugetana P Oct-Mar ? Amphiodia sp. opaque RP Aug-May, especially 13-33 Jan-Mar Amphiodia sp. orange belly RP Jan-Mar 8 (n=1) Amphiodia sp. tan* RP Sept ? Amphiodia periercta? D Oct-Mar 8 Amphioplus sp. vitellaria V Nov, Jan ~7 Amphiura arcystata D Nov-Feb 6 (n=1) Ophiacanthidae Ophiacantha diplasia P Jan-Apr 82 (n=1) Ophiopteridae Ophiopteris papillosa P, V Nov-Apr 65 (n=1) Ophiuridae Ophiocten hastatum* P Jan ? Ophiura leptoctenia* P May ? Ophiura sarsii* P Mar-Jun1 ? Ophiura luetkenii RP Nov, Feb-Apr 5 Gorgonocephalidae Gorgonocephalus eucnemis D Jan-Mar 5 Amphiura arcystata, Gorgonocephalus eucnemis, and Ophiura luetkenii. The fertilized eggs and developing embryos of ophiuroids often have a thick hyaline layer coating the egg, enclosing the cleavage and embryonic stages, and surrounding the blastula and gastrula (Emlet, 2006; Olsen, 5 6 1942; Yamashita, 1984). Embryos of brittle stars could be reliably identified by their thick hyaline layers, which were even visible between blastomeres in the two- to eight-celled stages, especially for A. sp. opaque and A. periercta? (Sections 3.4.3.1 and 3.4.3.3). We observed feeding larvae and different forms of nonfeeding developmental stages in the plankton of southern Oregon. We follow Hendler’s (1975) classification of ophiuroid development into three functional categories: planktotrophy with a feeding ophiopluteus; abbreviated development via a reduced pluteus, vitellaria, or pelagic direct developer; and brooding. We observed ten species with a planktotrophic ophiopluteus. We observed abbreviated development in the form of four reduced plutei, one vitellaria, and three species with pelagic direct development (Table 3.2). The ophiopluteus was the most common larval form, present in fourteen species (Table 3.2, Fig. 3.2). Ten of these ophioplutei are planktotrophic, whereas the remaining four were reduced plutei (see below). The planktotrophic plutei we observed were spread across six families. Ophiotrichidae (1 species), Ophiopholidae (2 species), Ophiopteridae (1 species) and Ophiacanthidae (1 species) contained only feeding plutei. Reduced plutei occurred in Amphiuridae (2 species), and Ophiuridae (1 species). The opacity and color of the larval tissues was correlated with development mode. Planktotrophic plutei generally had transparent larval tissues, while abbreviated forms were translucent or opaque in varying shades of tan to pink. We found the shape and proportions of the larval skeletal rods to be the definitive way to distinguish between plutei of different species (Mortensen, 1921), which could be viewed using cross- polarized light (Fig. 3.2). 5 7 Abbreviated modes of development were present in four of seven families in this study. Reduced plutei were observed in two families, Amphiuridae and Ophiuridae: Amphiodia sp. opaque, Amphiodia sp. orange belly, and Amphiodia sp. tan, and Ophiura luetkenii. Reduced plutei were recognized by their relatively short larval arms, particularly the posterolateral arms, well-developed left coelom, opaque coloration, and short development times. We obtained sufficient material to test for facultative planktotrophy in A. sp. opaque (Nakata & Emlet, 2023), and suspect the remaining reduced plutei to have this developmental mode as well. Another notable form of abbreviated development is the vitellaria, of which we observed in two species: Amphioplus sp. vitellaria and as the secondary larval form (following the planktotrophic ophiopluteus) in Ophiopteris papillosa. The vitellaria may have evolved multiple times as a modification of the pluteus (Hendler, 1991; Mortensen, 1921). We describe the first vitellaria from the Amphiuridae and the second example of an ophiopluteus developing into a vitellaria prior to metamorphosis (Cisternas & Byrne, 2005). 3.3. Patterns in Spawning Phenology We observed embryos, larvae, and juveniles of brittle stars over the course of the years 2014–2016 and 2018–2021. Plankton observations were made daily in the months of January, February, and March of years 2014 (79 days), 2015 (73 days), and 2016 (74 days). In years 2018–2021, sampling occurred almost daily in winter months and approximately weekly from 5 8 Figure 3.2. Ophioplutei of the northeast Pacific, viewed in dark field (A,L) and cross-polarized light (B–K,M,N). (A) Ophiothrix spiculata, scale bar 1 mm, (B) Ophiopholis kennerlyi, scale bar 100 µm, scale same for X-X, (C) another individual. (D) Amphipholis pugetana, (E) Amphiodia urtica, (F) A. sp. tan, (G) A. sp. orange belly, and (H) A. sp. opaque. (I) Ophiacantha diplasia and (J) Ophiopteris papillosa. (K) Ophiura sarsii, (L) Ophiocten hastatum, (M) Ophiura leptoctenia, and (N) O. luetkenii. 5 9 April to December. We examined plankton twice weekly during summer of 2018, but we limited our sampling to a few times a month from May to September during subsequent years due to the absence of ophiuroid larvae and high diatom abundance during those months. Because samples were collected by hand, we did not sample on days in which the high tide occurred between 8 pm and 7 am. We observed strong seasonal patterns in presence of ophiuroid developmental stages. All but three species (Amphiodia sp. tan, Ophiura leptoctenia, Ophiura sarsii, each observed just once; see Fig. 3.4) were observed most frequently in the plankton during the fall and winter months (November to March). During ENSO years we observed rare larvae, perhaps because of altered current patterns. Spawning may have been suppressed in local populations by heat waves (Shanks et al., 2020). We observed several patterns in our spawning records according to developmental mode. Nonfeeding larvae (Amphiura arcystata, Amphioplus sp. vitellaria, Gorgonocephalus eucnemis, Ophiura luetkenii) were observed in the winter months Jan-Mar. One exception is the pelagic direct developer, Amphiodia periercta? (sensu Emlet 2006), whose larvae were observed from October to March. Obligately and facultatively planktotrophic ophioplutei (A. sp. opaque, A. urtica, A. pugetana, and Ophiopholis spp.) occurred in the plankton from Oct to May in Coos Bay, Oregon. Larvae of a few species were observed very rarely, and only in the El Niño years of 2015 and 2019: Ophiothrix spiculata, Ophiura leptoctenia, Ophiura sarsii, and Ophiocten hastatum. We interpret these rare occurrences as larvae advected away from their benthic adult populations during warm-water events. 6 0 6 1 Figure 3.3. Spawning phenology for 14 planktonic ophiuroid spp. of the southern Oregon coast. Days of the month are marked on the x-axis, with each gray box representing five days’ time. The months of June, July, and September are omitted because no ophiuroid larvae were observed in plankton samples from those months. Occurrence data is represented by dots: filled for species with abbreviated development and open for species with planktotrophic development. Abundance of larvae (n) on each collection day is indicated by the size of the dot, with the smallest dot representing collections of larvae fewer than ten, and the largest representing collections of ten or more. Note that Ophiopholis kennerlyi and O. bakeri are combined in a single plot because the ophioplutei of these congeners are morphologically indistinguishable. Larvae with abbreviated development are marked by filled circles and those with planktotrophic development with empty circles. Figure 3.4. Spawning phenology for 4 additional ophiuroid spp. that were each observed on a single occasion. Amphiodia sp. tan has a reduced pluteus and Ophiocten hastatum, Ophiura leptoctenia, and Ophiura sarsii have planktotrophic ophioplutei. 3.4. Species accounts 3.4.1. Ophiotrichidae LJUNGMAN 1867 3.4.1.1. Ophiothrix spiculata LE CONTE 1851 Species Identity. DNA barcoding of a single larva and several juveniles revealed close matches to the species Ophiothrix spiculata. As sequences for O. spiculata were absent from databases at the time, R. Emlet collected adults off Catalina Island, CA, to which our larvae and juveniles were less than 4% different at the COI locus (Fig. 3.5, Table 3.3). Two distinct sequences attributed to O. spiculata from the Caribbean Sea (Bribiesca-Contreras et al., 2013) 6 2 Figure 3.5. Maximum likelihood tree of 16S and COI of Ophiothrix spp. from North America. Bootstrap support values >70 for major nodes are shown to the left of the node. Larvae identified in this study are in bold. Specimen information is listed in Table 3.3. Branch lengths are shown in scale. Atlantic that did not form a clade with the Pacific O. spiculata (Fig. 3.5) and were over 18% different from our specimens at the COI locus. Based on these data and our species-level threshold of 8.5% determined by ASAP analysis, we treat Pacific and Atlantic O. spiculata as different species. Furthermore, the species was described from Pacific, which suggests that animals in the Pacific, while morphologically similar to those in the Atlantic, should be raised to species. Distribution and Local Sites. Ophiothrix spiculata has not been listed as part of the benthic fauna of the southern Oregon coast and is known only from California to Chile. Its Specimen information for Ophiothrix spp. included in Figure 3.5. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. 6 3 Table 3.3. Specimen information for Ophiothrix spp. included in Fig. 3.5. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. Species Specimen Locus BOLDB or Collection Reference Code GenBankG Locality Ophiothrix angulata KC626250 COI KC626250G Cozumel Island, (Bribiesca-Contreras Mexico et al., 2013) Ophiothrix lineata EF053424 COI EF053424G Florida (Richards et al., 2007) Ophiothrix lineata KU895447 COI KU895447G USA (Hugall et al., 2016) Ophiothrix oerstedii KU895445 COI KU895445G USA (Hugall et al., 2016) Ophiothrix spiculata 12-11-1Op 16S, COI OLAB089-23B Charleston, OR This study Ophiothrix spiculata KC626260 COI KC626260G Cozumel Island, (Bribiesca-Contreras Mexico et al., 2013) Ophiothrix spiculata KC626261 COI KC626261G Cozumel Island, (Bribiesca-Contreras Mexico et al., 2013) Ophiothrix spiculata Oj26 16S, COI OLAB090-23B Charleston, OR This study Ophiothrix spiculata Op12-23 16S, COI OLAB091-23B Charleston, OR This study Ophiothrix spiculata OP5-699 16S OLAB092-23B Charleston, OR This study Ophiothrix spiculata OP5-740 16S OLAB093-23B Charleston, OR This study Ophiothrix spiculata Os1 16S, COI OOPH012-18B Catalina, CA This study Ophiothrix spiculata Os2 16S, COI OOPH013-18B Catalina, CA This study Ophiothrix suensoni KC626237 COI KC626237G Cozumel Island, (Bribiesca-Contreras Mexico et al., 2013) Ophiothrix suensoni KU895446 COI KU895446G Cozumel Island, (Bribiesca-Contreras Mexico et al., 2013) previous northern limit was Moss Beach, San Mateo County, California (Austin & Hadfield, 1980; Lambert & Austin, 2007), which was expanded northward to Patrick’s Point State Park, Humboldt County, CA based on iNaturalist records (Sanford et al., 2019). A small number of juveniles were observed in Victoria, BC but surveys conducted in the region since 2001 have not documented any more O. spiculata (Lambert & Austin, 2007; Sanford et al., 2019). Embryonic Description. We did not observe the embryos of this species. Larval Description. Ophiothrix spiculata develops via an eight-armed ophiopluteus, and it is one of the largest of all the plutei we observed (Fig. 3.2A). The epidermis is transparent, and 6 4 the larva develops very long, straight, and wideset posterolateral arms that may be over five times the length of the other arms (Fig. 3.2A). The postoral and posterdorsal arms are approximately the same length, and slightly shorter than the anterolateral arms. The body rods are short and in line with the posterolateral arms (Fig. 3.6J). The left transverse rod may bear a median process, as reported for ophioplutei of this genus (Mortensen, 1921), but cannot be clearly seen in our photos (Fig. 3.6J). Similar Species. We observed O. spiculata only as eight-armed ophiopluteus and juvenile, both of which are similar to that of Ophiopholis spp. (see Section 3.4.1.1 below). In both species, the posterolateral arms are straight, long, and wideset; the remaining arms are similar in length to one another but markedly shorter than the posterolateral arms. Plutei of O. spiculata have posterolateral arms up to five times the length of their other arms, whilst the posterolateral arms of Ophiopholis spp. are up to three times the length of the other arms. This comparison is most useful in advanced larvae, as the ultimate length of the arms is accumulated across time as the larva eats and grows. Development Mode. Ophiothrix spiculata has an obligately feeding, 8-armed ophiopluteus, as expected from a small egg of diameter 110 µm (Pearse, 1994). Feeding larvae are the norm in Ophiothrix (Guille, 1964; Hendler, 2005, 1995; Hendler & Littman, 1986; Kitazawa et al., 2015; Mladenov, 1983, 1985a; Morgan & Jangoux, 2005; Mortensen, 1938; Selvakumaraswamy & Byrne, 2000, 2006), although nonfeeding planktonic larvae (Mladenov, 1979; Richards et al., 2007) and one brooder (Schoppe, 1996) are also known. Another genus in the same family, Macrophiothrix, has feeding ophioplutei and several reduced plutei (Allen & Podolsky, 2007). 6 5 Juvenile Description. The juvenile is large and motile at metamorphosis, with three arm segments, the proximal two of which bear hooked arm spines (Fig. 3.6K’) that can be used to secure the individual in rock crevices or algal holdfasts. The hooked arm spines can be used to distinguish the juveniles of O. spiculata and Ophiopholis spp., as O. spiculata has a single hook, and Ophiopholis spp. have a double hook. The juvenile arms are often folded under the disk while still attached to the larval arms, and this condition may persist following metamorphosis. The stereom of the juvenile disk is shown in Fig. 3.6K”. Reproductive Timing. This larva is not usually observed in the plankton of Charleston, OR, but five larvae or recently metamorphosed juveniles were collected from plankton samples during the marine heatwave known as The Blob from December 2014 and January 2015 (Fig. 3.3). Spawning has been observed from late winter to July in central California (Lambert & Austin, 2007; Rumrill, 1982). Planktonic Duration. We were not able to observe the planktonic duration for O. spiculata, but we suspect it to be a month or longer given the planktotrophic development and large size of the ophiopluteus (see Fig. 3.2A). Figure 3.6. Larvae and juveniles of (A-G) Ophiopholis kennerlyi, (H,I) O. bakeri, and (J,K) Ophiothrix spiculata. Secondary and tertiary views of the same individual are marked with an apostrophe (’) and quotation mark (”), respectively. (A) Aboral view of the early pluteus of O. kennerlyi, (B) oral view of six-armed pluteus, and (C) aboral view of early eight-armed pluteus, with anterolateral (al), postoral (po), posterodorsal (pd), and posterolateral (pl) arms. B and C are same scale as A, scale bar is 100 µm. (D) Oral view of a more advanced eight-armed pluteus and (D’) closeup of the posterior girdle in cross-polarized light with end rods (er), body rods (br) and transverse rods (arrowhead). Scale bars are 1 mm in D and 100 µm in D’. In both D and D’ the posterodorsal arms are largely behind the postoral arms. (E) Another eight-armed pluteus, showing the reddish coloration of the skeleton, especially the posterolateral, body, and end rods, as well as the accumulation of red pigmentation in the distal portion of the posterolateral arms. (F) A rudiment-stage larva with a juvenile rudiment (jr) carried by the posterolateral arms, same scale as D. (G) Juvenile collected from the plankton, and (G’) a closeup of the hooked spines with two teeth on the terminal segment of the arm (open arrowhead). Scale bar in G is 100 µm. (H) Eight- armed pluteus of O. bakeri, oral view at same scale as D and (H’) closeup of the posterior girdle, same scale as D’. (I) Aboral view of another individual, showing the red-orange coloration of the posterolateral, body, and end rods. (J) Closeup of the posterior girdle of O. spiculata, same individual as Fig. 3.2.2A and same scale as G’. (K) Juvenile collected from the plankton, aboral view in transmitted light, and (K’) closeups of the hooked spines on the terminal segments of the arms, and (K”) of the disc stereom in partially cross-polarized light. 6 6 6 7 3.4.2. Ophiopholidae O’HARA, STÖHR, HUGALL, THUY & MARTYNOV 2018 Ophiopholidae was recently raised to family status (O’Hara et al., 2018) and contains a single genus, Ophiopholis, which is comprised of eight species. Five of these occur in the northeastern Pacific: aculeata, bakeri, japonica, kennerlyi, and longispina (Lambert & Austin, 2007). We observed two species, Ophiopholis kennerlyi and O. bakeri, whose larvae were not distinguishable based on morphology (see Fig. 3.6D, H) but barcoding showed to be distinct (Fig 7, Table 3.4). Therefore, we will give a description of both species, although we suspect that the early pluteus material is of O. kennerlyi due to its intertidal to shallow subtidal distribution compared to that of O. bakeri, which occurs subtidally at depths of up to 1200 m (Lambert & Austin, 2007). 3.4.2.1. Ophiopholis kennerlyi LYMAN 1860 and O. bakeri MCCLENDON 1909 Species Identity. Although there are characters listed in the literature that distinguish O. kennerlyi from O. aculeata, (Lambert & Austin, 2007) state that adult specimens of O. aculeata occur in the Bering Sea and into British Columbia but that positively identified specimens of O. aculeata have not been collected in waters of Oregon and Washington. We have not collected O. aculeata in our samples, nor do any of our molecular sequences match those of O. aculeata from the Atlantic (Fig. 3.7). The deep-water species O. bakeri and O. longispina are recognized as separate species and are distinguished by presence or absence of spines on their radial shields, and length of arm spines (Lambert and Austin 2007). They are also acknowledged to be like one another as 6 8 Figure 3.7. Maximum likelihood tree of COI sequences for Ophiopholis spp. Barcodes of larvae identified in this study are in bold. Table 3.4. Specimen information for Ophiopholis spp. included in Fig. 3.7. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. Species Specimen Locus BOLDB or Collection Locality Reference Code GenBankG Ophiopholis aculeata GU670181 COI GU670181G Manitoba, Canada (Corstorphine, 2011) Ophiopholis aculeata HM542280 COI HM542280G New Brunswick, (Corstorphine, Canada 2011) Ophiopholis aculeata KJ620605 COI KJ620605G Iceland (Khodami et al., 2014) 6 9 Ophiopholis aculeata KU895311 COI KU895311G USA (Hugall et al., 2016) Ophiopholis aculeata KX459016 COI KX459016G North Sea (Laakmann et al., 2017) Ophiopholis aculeata MG421109 COI MG421109G Newfoundland, Dewaard, Canada unpublished Ophiopholis bakeri 83P COI OLAB073-23B Charleston, OR This study Ophiopholis bakeri BOIMB-2132 COI Cape Arago, OR G. Paulay, pers. comm. Ophiopholis bakeri HM473935 COI HM473935G British Columbia, Canada Ophiopholis bakeri K01-OP COI OLAB074-23B Charleston, OR This study Ophiopholis bakeri KU895312 COI KU895312G USA Ophiopholis bakeri Obak1 COI OOPH047-23B Florence, OR This study Ophiopholis bakeri Obak2 COI OOPH048-23B Florence, OR This study Ophiopholis bakeri Oph1 COI OLAB075-23B Charleston, OR This study Ophiopholis bakeri R1 COI OLAB076-23B Charleston, OR This study Ophiopholis japonica HM473934 COI HM473934G British Columbia, (Corstorphine, Canada 2011) Ophiopholis kennerlyi Batman 10- COI OLAB077-23B Charleston, OR This study 11-18 Ophiopholis kennerlyi BOIMB-0995 COI G, Paulay, pers. comm. Ophiopholis kennerlyi HM542299 COI HM542299G British Columbia, (Corstorphine, Canada 2011) Ophiopholis kennerlyi HM542302 COI HM542302G British Columbia, (Corstorphine, Canada 2011) Ophiopholis kennerlyi KU495781 COI KU495781G British Columbia, (Corstorphine, Canada 2011) Ophiopholis kennerlyi Oac1 COI OLAB097-23B Cape Arago, OR This study Ophiopholis kennerlyi Oaii COI OOPH007-18B Cape Arago, OR This study Ophiopholis kennerlyi Oj323 COI OLAB078-23B Charleston, OR This study Ophiopholis kennerlyi Ok1 COI OLAB079-23B Charleston, OR This study Ophiopholis kennerlyi Ok2 COI OLAB080-23B Charleston, OR This study Ophiopholis kennerlyi Ophiopholis COI OLAB081-23B Charleston, OR This study 10-11-18 Ophiopholis kennerlyi Ophor COI OLAB082-23B Charleston, OR This study Ophiopholis longispina OE/Oplo1 COI OOPH017-18B Cape Arago Shelf, OR This study Ophiopholis longispina Og/Oplo2 COI OOPH019-18B Cape Arago Shelf, OR This study 7 0 Clark (1911, p123) states that “the line of separation between the two forms is very narrow”. Molecular data we collected and obtained from public databases does not show bakeri and longispina to form reciprocally monophyletic groups, but the data for longispina is limited (Fig. 3.7). Our larvae group most closely with an adult collected and identified by us as O. bakeri. As such, we will refer to the larvae by that name in this manuscript. Distribution and Local Sites. Ophiopholis kennerlyi and O. bakeri occur across similar ranges, from British Columbia to southern California (Lambert & Austin, 2007). They are separated by depth: O. kennerlyi occurs intertidally to 366 m and O. bakeri occurs from 37 to 1,204 m depth (Lambert & Austin, 2007). We encounter adult O. kennerlyi intertidally and from dredges offshore, where we have also collected O. bakeri (Table 3.4). Embryonic Description. We did not observe the embryos of these species, but the embryology of O.kennerlyi (misidentified as O. aculeata) is documented (Primus, 2005; Strathmann, 1987; Strathmann et al., 2020). Larval Description. Ophiopholis spp. develop via an eight-armed ophiopluteus. The larva of O. bakeri was previously unknown, but O. kennerlyi (identified as O. aculeata) has been studied extensively in the laboratory, including the feeding mechanism of the ophiopluteus (Strathmann, 1971, 1978b), and larval cloning (Balser, 1998). The early pluteus of Ophiopholis is transparent, has four arms, which extend anteriorly and terminate at approximately the same height (Fig. 3.6A). The posterolateral arms grow in length and become increasingly wideset as the larva develops. These first two pairs are followed quickly by the postoral arms, which support a ciliary band in front of the mouth (Fig. 3.6B). The posterodorsal rods bud from the anterolateral rods (Fig. 3.6) and eventually grow to be of equal 7 1 anterior height with the postoral arms (Fig. 3.6D). The posterolateral arms of the eight-armed pluteus are greater in height than the rest of the arms, which are similar in length to one another and may be more wideset in O. bakeri, although this was not consistent across all specimens (Fig. 3.6D, H). As the larva develops, the posterolateral, body, and end rods may take on an orange-red hue (Fig. 3.5E, I), and pigmentation that is yellow to red in color may accumulate in the distal portions of the posterolateral arms (Fig. 3.6D, E, I). This appears to be more common and more extreme in kennerlyi, but the number of specimens of bakeri were limited. Furthermore, larvae cultured in the laboratory with microalgal food offered ad libitum develop coloration on the skeletal rods and in the posterolateral tissues to far greater effect than any wild- caught larvae we have observed (N. Nakata, personal observation). Similar species. Early plutei of Ophiopholis spp. larvae can be difficult to distinguish from those of Amphiodia urtica and Amphipholis pugetana. While the plutei of all three species have transparent epidermises, A. urtica and A. pugetana also have red pigmentation at the distal tips of their posterolateral arms. The skeletal rods of Ophiopholis are generally straight, whereas those of other taxa described here are usually curved (Fig. 3.2). Posterolateral arm rods of Ophiopholis larvae may bear very small thorns on their inner side, that are well-spaced (Fig. 3.6, compare with amphiurids in Fig. 3.10). The late ophiopluteus of Ophiopholis spp. is most like that of Ophiothrix spiculata, which occurs rarely in southern Oregon. From the small number of O. spiculata larvae we observed, we determined a few characters in the advanced ophioplutei that distinguish the species: first, posterolateral arms of Ophiopholis spp. never exceed three times the length of the other arms, while posterolateral arms of O. spiculata can be five times as long; second, the 7 2 posterolateral arms of the ophiopluteus (before the juvenile is formed) become increasingly wideset in both species, but in Ophiopholis spp. their angle does not exceed 120°, while in O. spiculata the angle of the posterolateral arms can exceed 120°. Finally, many ophioplutei of Ophiothrix have a medial process on the transverse rod (Mortensen, 1921), this process is absent in early plutei of Ophiopholis, and if present there are two medial processes. Development Mode. Both O. kennerlyi and O. bakeri have a planktotrophic ophiopluteus. This is well known from the literature for O. kennerlyi as it is often called O. aculeata, but we consider to be O. kennerlyi in the Pacific (Austin & Hadfield, 1980; Balser, 1998; Strathmann, 1987). Juvenile Description. Plutei of Ophiopholis undergo a metamorphosis, in which the anterolateral, postoral and posterodorsal arms are all resorbed in the creation of the juvenile rudiment. The juvenile has 2-3 arm segments; the arms develop folded in on the juvenile oral surface. Juveniles have hook-shaped arm spines with two hooks, distinguishing Ophiopholis spp. from O. spiculata, which has arm spines with single hooks (Fig. 3.6G’). Reproductive Timing/Spawning Phenology. Larvae of Ophiopholis spp. occurred in the plankton from October to April, although the timing of their appearance varied from year to year (Fig. 3.3). Ophiopholis kennerlyi is known to spawn throughout the year in the laboratory (Strathmann, 1987). Planktonic Duration. Ophiopholis kennerlyi has a long-lived ophiopluteus, with a planktonic duration of 83-216 days (Austin & Hadfield, 1980; Strathmann, 1978a). 7 3 3.4.3. Amphiuridae LJUNGMAN 1867 We observed eight species of amphiurids in the plankton and were able to match five types of larvae to adult specimens via DNA barcoding: Amphiodia urtica, A. periercta?, and A. sp. opaque, Amphipholis pugetana and Amphiura arcystata. The remaining three species are ‘orphan larvae’ without molecular matches to adult specimens. Two of these ‘orphans’ group with other species of Amphiodia (sp. orange belly and sp. tan), and the last with Amphioplus (sp. vitellaria). See Fig. 3.8, Table 3.5. We observed several developmental patterns amongst the amphiurids of the NE Pacific. The only brittle star known to brood its young in the NE Pacific is the cosmopolitan species Amphipholis squamata (Hugall et al., 2023; Strathmann, 1987), which was not described here because it lacks a planktonic form. Two of the eight plankton ic stages of amphiurids had planktotrophic development via an ophiopluteus, Amphipholis pugetana and Amphiodia urtica. Remarkably, we observed six amphiurids with abbreviated development, including three species that develop via a reduced pluteus (Amphiodia spp. opaque, orange belly, and tan), one via a nonfeeding vitellaria (Amphioplus sp. vitellaria), and the two others that develop directly into juveniles in the plankton (Amphiodia periercta?, Amphiura arcystata). Facultative planktotrophy has been confirmed for A. sp. opaque (Nakata & Emlet, 2023), and the other two reduced plutei may also be facultative planktotrophs because they form complete guts, have ciliated bands and larval arms. Abbreviated forms of development are not unknown from the family Amphiuridae, Amphioplus abditus (Hendler, 1973) and Amphiura chiajei (Hendler, 1975), with several more species inferred to have abbreviated development based on egg size (Hendler, 1995; see Nakata Chapter IV). 7 4 7 5 Figure 3.8. Maximum likelihood tree of 16S and COI sequences for Amphiuridae species from the northeast Pacific. See Table 3.5 for specimen and locus data. Branch lengths denote patristic distances according to the legend in the bottom left. Support values for major nodes are marked when >70. Table 3.5. Specimen information for Amphiuridae spp. included in Fig. 3.8. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. Species Specimen Code Locus BOLDB or Collection Reference GenBankG Locality Amphichondrius Au1 16S, COI OOPH005-18B Catalina Island, This study granulatus CA Amphiodia cf. HM542062 COI HM5420622 British Columbia, (Corstorphine, occidentalis Canada 2011) Amphiodia occidentalis A.occ FHL 6-15- 16S, COI OOPH034-23B Friday Harbor, This study 19 WA Amphiodia occidentalis AoMc1a 16S, COI OOPH020-22B Charleston, OR This study Amphiodia occidentalis AoMc1b 16S, COI OOPH021-22B Charleston, OR This study Amphiodia occidentalis KU495744 COI KU495744G British Columbia, (Corstorphine, Canada 2011) Amphiodia sp. Dome MMB17 16S, COI OOPH032-22B Charleston, OR (Nakata & House Emlet, 2023) Amphiodia sp. Olga B MMB13 16S, COI OOPH039-23B Olga, WA This study Amphiodia sp. Olga C MMB14 16S, COI OOPH040-23B Olga, WA This study Amphiodia sp. opaque 41P COI OLAB099-23B Charleston, OR This study Amphiodia sp. opaque Opaque 11-7-18 COI OOPH029-22B Charleston, OR (Nakata & Emlet, 2023) Amphiodia sp. opaque Opaque FHL 16S OLAB048-23B Friday Harbor, This study WA Amphiodia sp. opaque Opq1 FHL 16S, COI OLAB047-22B Friday Harbor, (Nakata & WA Emlet, 2023) Amphiodia sp. opaque Opq2 16S, COI OLAB004-22B Charleston, OR (Nakata & Emlet, 2023) Amphiodia sp. opaque QHAK711 21 COI QHAK711-21B British Columbia, Hakai Inst., Canada unpublished Amphiodia sp. orange Ob1 16S, COI OLAB050-23B Charleston, OR This study belly Amphiodia periercta? 829 COI OLAB032-22B Charleston, OR This study Amphiodia periercta? 8P COI OLAB033-22B Charleston, OR This study Amphiodia periercta? Amphio egg1 COI OLAB045-22B Charleston, OR (Nakata & Emlet, 2023) Amphiodia periercta? R7 16S OLAB031-22B Charleston, OR This study 7 6 Amphiodia sp. Portside MMB16 16S, COI OOPH031-22B Charleston, OR (Nakata & Emlet, 2023) Amphiodia sp. tan O919 COI OLAB051-23B Charleston, OR This study Amphiodia urtica 988 COI OLAB028-22B Charleston, OR This study Amphiodia urtica A.urt adt 4-23-19 COI, 16S OOPH056-23B Cape Arago This study Shelf, OR Amphiodia urtica A.urt1 FHL COI OLAB053-23B Friday Harbor, This study WA Amphiodia urtica DISA835 COI DISA835-19B Los Angeles, CA DISCO MBC LACM, unpublished Amphiodia urtica HM542068 COI HM542068G British Columbia, (Corstorphine, Canada 2011) Amphiodia urtica urtica 11-5-18 COI OOPH027-22B Charleston, OR (Nakata & Emlet, 2023) Amphioplus KU895009 COI KU895009G Antarctica (Hugall et al., peregrinator 2016) Amphioplus sp. OB 16S OOPH046-23B Newport Valley, This study OR Amphioplus sp. OH 16S, COI OOPH049-23B Newport Valley, This study OR Amphioplus sp. juvA 1-4-19 COI OLAB055-23B Charleston, OR This study vitellaria Amphioplus sp. R38 16S, COI OLAB056-23B Charleston, OR This study vitellaria Amphioplus sp. R39 16S, COI OLAB057-23B Charleston, OR This study vitellaria Amphioplus sp. vit1 16S, COI OLAB054-23B Charleston, OR This study vitellaria Amphioplus sp. Vitellaria 11-14- 16S, COI OLAB058-23B Charleston, OR This study vitellaria 18 Amphioplus E6874 COI NA Northeast Pacific T. O’Hara, strongyloplax pers. comm. Amphipholis pugetana 811 COI OLAB016-22B Charleston, OR This study Amphipholis pugetana A.pug1 FHL 16S, COI OLAB059-23B Friday Harbor, This study WA Amphipholis pugetana Ampu1 16S, COI OOPH003-18B Cape Arago, OR This study Amphipholis pugetana Ampu2 16S, COI OOPH004-18B Cape Arago, OR This study Amphipholis pugetana pugetana 10-25- COI OOPH028-22B Charleston, OR This study 18 Amphipholis pugetana R3 16S, COI OLAB020-22B Charleston, OR This study Amphipholis squamata A.squ1 FHL COI OOPH035-23B Friday Harbor, This study WA 7 7 Amphiura arcystata OpArm 16S, COI OOPH050-23B Charleston, OR This study Amphiura diomedeae Amdi COI OOPH015-18B Newport, OR This study Amphiura arcystata Cross 16S, COI OLAB106-23B Charleston, OR This study Amphiura arcystata H19-Op 16S, COI OLAB060-23B Charleston, OR This study Amphiura arcystata H20-op 16S, COI OLAB061-23B Charleston, OR This study Amphiura arcystata op-10 COI OLAB103-23B Charleston, OR This study Amphiura arcystata Op111 COI OLAB062-23B Charleston, OR This study Amphiura arcystata Op112 COI OLAB063-23B Charleston, OR This study Amphiura arcystata R35 16S OLAB064-23B Charleston, OR This study Amphiura arcystata R42 16S, COI OLAB065-23B Charleston, OR This study 3.4.3.1. Amphiodia urtica (LYMAN 1860) Species Identity. We collected larvae from Charleston, OR and Friday Harbor, Washington whose COI sequences were 2% different adults from southern CA, British Columbia, Canada, and the Cape Arago Shelf, OR (Fig. 3.8, Table 3.5), which falls within the 8.5% threshold determined by ASAP analysis. In our tree of amphiurids from the northeast Pacific, Amphiodia urtica is sister to the clade made up of A. sp. orange belly and A. sp. tan, two species of orphan larvae. Distribution and Local Sites. Benthic populations of A. urtica are known from Alaska to California, from intertidal to 370 m depth (Austin & Hadfield, 1980; Hendler, 1996; Kyte, 1969; Lambert & Austin, 2007). We have collected a few juvenile individuals by dredge at 75– 100 m off Cape Arago, Oregon. Embryonic Description. We did not observe the embryos of these species. Larval Description. Amphiodia urtica develops via an eight-armed planktotrophic ophiopluteus. Planktotrophic development was predicted by small egg size (<100 µm) (Schiff & 7 8 Bergen, 1996). The early pluteus is small, with four larval arms and red pigmentation at the tips of the posterolateral pair (Fig. 3.9A). The postoral arms emerge next, supporting a portion of the ciliated band ventral to the mouth (Fig. 3.9B). Lastly the posterodorsal arms grow out from the bases of the anterolateral arms to make an eight-armed ophiopluteus (Fig. 3.9C). As the larva grows the posterolateral arms become curved and bear thorns (Fig. 3.9C’); anterolateral and postoral arms may also bear thorns, but these tend to be shorter than those on the posterolateral arms. Red pigmentation is often visible at the distal ends of the posterolateral arms, but never spans the length of the arm. The posterolateral and body rods may be orange red in color. The postoral and posterodorsal arms are relatively short, and their distal tips rarely exceed the height of the larval mouth. The body rods curve anteriorly to meet the anterolateral and posterolateral rods and descend posteriorly toward each other at an approximately 45° angle. The body rods are connected by thorny transverse rods and the end rods are pointed but not touching (Fig. 3.9C”). The juvenile rudiment develops to the left of the digestive system. The lobes of the left coelom are visible (Fig. 3.9C). The rudiment-stage larva has three arms - a pair of posterolateral arms and one remaining (right) anterolateral arm - with the pentagonal juvenile rudiment bearing tube feet centered among the arms (Fig. 3.9D). Similar Species. As an early pluteus, A. urtica is most easily confused with Amphipholis pugetana. Both larvae have red-orange skeletons covered in transparent epidermis, with an accumulation of red pigmentation at the distal ends of the posterolateral arms. The epidermis of A. urtica adheres closely to the larval skeleton and follows the curvature of the arms. In A. pugetana, the epidermis is separated from the skeleton and has an inflated appearance, creating a club shape at the distal ends of the anterolateral and posterolateral arms. Finally, the pluteus of A. 7 9 urtica bears many thorns, especially later in development, and thorns are absent or nearly imperceivable in A. pugetana. The skeleton of A. urtica is most like that of A. sp. opaque and A. sp. orange belly, with its curved posterolateral arms and the presence of spines. The pluteus of A. urtica can be distinguished from its congeners by the transparency of its epidermis, which is opaque pink to salmon in A. sp. opaque and orange in the body and white on the arms in A. sp. orange belly. Development Mode. Amphiodia urtica develops as an obligately feeding ophiopluteus (N. Nakata, unpublished). Prior reports for the life history of this species are limited to egg size, approximately 0.12 mm diameter, indicating development as a feeding ophiopluteus (Hendler, 1996). Juvenile Description. Juvenile is star shaped, comprised of a pentagonal disc and triangle-shaped arm plates (Fig 9F). Reproductive Timing. In our data, larvae were present in the Oregon plankton from November to April, with rare observations in October and May. We also have collected these larvae in August and September in the San Juan Islands, WA. In Southern California, where this species is very abundant on the shelf, A. urtica has been observed spawning in November, and in October in British Columbia (Austin & Hadfield, 1980). Planktonic Duration. Planktonic duration for A. urtica is approximately 20 days (Nakata & Emlet, 2023), which is relatively short for an obligate planktotroph. 8 0 Figure 3.9. Planktotrophic plutei of Amphiuridae: Amphiodia urtica (A–F) and Amphipholis pugetana (G–K). Secondary and tertiary views of the same individual are marked with an apostrophe (’), or a quotation mark (”), respectively. Scale bar in A is 100 µm and the same for A-E, G-I, K. Scale bars in F and J are 100 µm. (A) Oral view of the early pluteus of A. urtica with anterolateral (al) and posterolateral (pl) arms, the latter with red pigment accumulated at the distal tips. (B) Early eight-armed (oral view), and (C) advanced eight-arm pluteus (oral view) with anterolateral, posterolateral, postoral (po), and posterodorsal (pd) arm pairs. (C’) Closeups of thorns on posterolateral arms, and (C”) posterior girdle, respectively, in cross-polarized light. The posterior girdle is composed 8 1 of the transverse rods (filled arrowhead), body rods (br) and end rods (er). (D) Oral view of a rudiment-stage larva with right anterolateral arm (ral) and juvenile rudiment (jr) bearing tube foot buds (tf). (E) The juvenile of the same individual, one day later. (F) A closeup of the stereom of another individual in cross-polarized light. (G) Early pluteus of Amphipholis pugetana with red pigmentation at the distal tip of the posterolateral arms. (H, I) Six- and eight-armed plutei, respectively, of A. pugetana. The portion of the ciliated band on the distal end of the anterolateral arms often bends medially (open arrowhead). (J) Rudiment-stage larva with right anterolateral arm (ral), juvenile rudiment (jr) with tube feet (tf), scale 100 µm. (K) Juvenile in partially cross-polarized light highlights the stereom of the disk plates and spine tips. 3.4.3.2. Amphipholis pugetana (LYMAN 1860) Species Identity. There are two species of Amphipholis that occur in the northeast Pacific (26 Amphipholis spp. total): A. pugetana and the cosmopolitan brooder A. squamata. Our specimens of A. pugetana and A. squamata do not group together, and the genus Amphipholis is probably polyphyletic (Fig. 3.8, Table 3.5)(O’Hara et al., 2018). The placement of Amphipholis relative to Amphiodia and Amphiura is uncertain and the family Amphiuridae probably requires revision (O’Hara et al., 2018). Distribution and Local Sites. Amphipholis pugetana occurs from the Gulf of Alaska to southern California, 0-1204 m (Lambert & Austin, 2007). We have collected adults by dredge at approximately 50 m depth off Cape Arago, Oregon. Embryonic Description. We did not observe the embryos of these species. Larval Description. Amphipholis pugetana develops via a planktotrophic eight-armed pluteus, which is recognized by its inflated epidermis that is well separated from the larval arm rods. The epidermis is transparent, often with an accumulation of red pigmentation at the distal ends of the posterolateral arms. This feature is present even in the early pluteus (Fig. 3.9G). 8 2 The posterolateral arms are curved, and any thorns are very small, and triangle shaped, if they are present at all. The anterolateral rods are straight but may become slightly curved proximally and distally in the advanced pluteus (Fig. 3.2D); the anterolateral rods taper to a sharp point, and never bear thorns, unlike plutei from the genus Amphiodia (e.g., Amphiodia urtica, A. sp. opaque, A. sp. orange belly, and A. sp. tan). The ciliated band on the tips of anterolateral arms may extend medially, giving this larva a ‘bunny-eared’ appearance (Fig. 3.9I). The body rods are relatively short, curved, and nearly horizontal. The transverse rods are unadorned. The end rods are short and run parallel to one another, not touching at their distal ends. The postoral and posterodorsal arms remain short, with the postoral arms terminating just above the mouth at their longest, and the posterodorsal arms never terminating above the larval mouth. Similar Species. As an early pluteus, A. pugetana is most likely to be confused with A. urtica or Ophiopholis spp. Amphipholis and A. urtica often have an accumulation of red pigmentation at the distal ends of the posterolateral arms (Fig. 3.9G, I, J). Amphipholis pugetana has thicker arms than A. urtica due to its inflated epidermis, which becomes more apparent as the larva develops. Thorns are present on the skeletal rods of A. urtica but arm rods of A. pugetana lacks thorns. Plutei of Ophiopholis may have red pigmentation on the posterolateral arms and have straight posterolateral and body rods (Fig. 3.6); while these are curved in A. pugetana (Figs. 2D, 9I). Ophiura leptoctenia also has an inflated epidermis, but it is widest at the base of the posterolateral arms and tapers to a point at their distal ends. Furthermore, the pluteus of O. leptoctenia has recurrent rods, which are absent in A. pugetana. 8 3 Development Mode. Amphipholis pugetana broadcast spawns its gametes (Lambert & Austin, 2007) and develops as a planktotrophic ophiopluteus (Shanks, 2001). As the juvenile rudiment develops. The postoral, posterodorsal and left anterolateral are resorbed, leaving a three-armed larva (Fig. 3.9J). Juvenile Description. The juvenile has a roughly pentagonal disc with five arm tips that end bluntly (Fig. 3.9K). The shape of the arm tips and the distinct shape of the red pigmentation often visible through the disc are characteristic for this species. There are several amphiurid juveniles with light pink to cream-colored bodies, often with reddened gut. Only the juveniles of A. pugetana juveniles have extensions of the red gut into the interradii. Juveniles were found in the plankton. Reproductive Timing. The ophioplutei and juveniles of A. pugetana occur in the Charleston plankton from October to March. We also have collected them in August and September in the San Juan Islands. WA. Planktonic Duration. Unknown. 3.4.3.3. Amphiodia sp. opaque Species Identity. The species identity of this larva remains unresolved, but COI and 16S sequences group with species in the genus Amphiodia. Distribution and Local Sites. The location of adult populations in Oregon are not known. The embryos and larvae of this species have been collected off the Oregon coast (see also Nakata and Emlet 2023), at least 10 juvenile and adult stages that have been barcoded have been collected in the Salish Sea (private data BOLD) 8 4 Development Mode. A. sp. opaque is a facultative planktotroph, as determined by feeding experiments. The larva can complete metamorphosis into a juvenile even under starvation conditions, but has quicker development, higher metamorphic success, and larger juveniles when given food (Nakata & Emlet, 2023). Embryonic Description. Embryos of this species are orange to salmon in color and have a thick hyaline layer (Fig. 3.10A). This hyaline layer can be seen between blastomeres at the 2-8 cell stages, like A. periercta? (Emlet, 2006). Embryos usually hatch as blastulae. Gastrulae are shaped like a chicken’s egg, wider toward the posterior and more pointed in the anterior., and uniformly ciliated. The first spicules develop as a pair of spicules in the posterior of the gastrula (Fig. 3.10A’). Larval Description. Amphiodia sp. opaque develops via a six-armed reduced pluteus with opaque epidermis. The early pluteus is dorsoventrally compressed but has the beginnings of the posterolateral and anterolateral arms, supported by paired triradiate spicules (Fig. 3.10B’). The postoral ciliated band is present, stretching across the ventral side of the larva from the tips of the posterolateral arms. The ophiopluteus can be recognized by an epidermis that is light pink to salmon in color, and semi- to fully opaque so that internal structures are not easily viewed. The posterolateral arms are often straight but may be curved or become curved during rudiment formation. The posterolateral and anterolateral rods often bear thorns (Fig. 3.10D’). The anterolateral arms can be straight or curved and bear short thorns (Fig 10C’, D’). The body rods are mostly straight, curving to connect with the posterolateral and end rods, which are of moderate length, connected by thorny transverse rods. The end rods do not touch at their distal ends. 8 5 Juvenile Description. The juvenile is pentagonal juvenile (Fig. 3.10E), very similar to A. sp. orange belly (Fig. 3.10L). Similar Species. Due to their morphology as reduced plutei, their opaqueness and coloration in shades of orange, salmon, pink, and tan the early plutei of A. sp. opaque, A. sp. orange belly, A. sp. tan, and Ophiura luetkenii can be confused with one another. In A. sp. opaque and A. sp. tan the color of the epidermis is uniform across the larval body and arms, light pink to salmon in the former (Fig. 3.10C), tan in the latter (Fig 10O). In A. sp. orange belly and O. luetkenii the larval body is orange, and the posterolateral and anterolateral arms are white (Fig. 3.10H). Reproductive Timing. Embryos and larvae of A. sp. opaque can be found in the Charleston plankton most reliably and in greatest density in the winter, particularly the month of February. Larvae have also been observed in fall (October, November). In Friday Harbor, WA, larvae were collected in late summer (July, August), but this was the only season we were able sample at that location. Larvae of various developmental stages (early pluteus, pluteus, and rudiment-stage larva) were observed on February 24, 2023, just outside of Newport, OR, suggesting that adult populations may be nearby. Planktonic duration. Planktonic duration in this species has been shown to vary according to food availability during the larval stage. Median planktonic durations (PD) for fed larvae were 17 and 13 days in two study years, with a maximum PDs of 21 and 19 days, respectively (Nakata & Emlet, 2023). 8 6 3.4.3.4. Amphiodia sp. orange belly Species Identity. We know Amphiodia sp. orange belly only from its larva, which we collected only four times (Fig. 3.3). Adult populations of A. sp. orange belly have not been identified near Charleston, Oregon. DNA barcoding and larval morphology places the larva within the genus Amphiodia, sister to another orphan larval species, A. sp. tan (Fig. 3.8, Table 3.5). There are at least two described Amphiodia species known to occur in the NE Pacific that are lacking developmental information: digitata and psara (Lambert & Austin, 2007), but we lack sequences from adults to connect these species with our larval morphotypes. Distribution and Local Sites. Adult populations in Oregon are not known. Embryonic Description. We did not observe the embryos of this species. Larval Description. Orange belly develops via a six-armed pluteus. The early pluteus is similar to that of A. sp. opaque, with a well-developed left coelom (Fig. 3.10F) The reduced pluteus is small (Fig. 3.10G, H) and has shortened posterolateral arms (compare with A. urtica) and postoral arms are lacking. It also has a deep orange stomach and left coelom and salmon to white colored larval arms (Fig 10G–K). The pluteus has an open mouth and hollow stomach and intestine. The posterolateral arms are somewhat straight, and bear thorns, as do the anterolateral arms (Fig. 3.10K’). The body rods are curved and similar in shape to those of Amphipholis pugetana. The end rods are of moderate length and united by thorny transverse rods. Similar Species. The reduced pluteus of A. sp. opaque is most similar in size and shape to its congener, A. sp. opaque, but differs in color by having an orange body with white posterolateral arms. It also has posterolateral arms that are more strongly curved than those 8 7 usually found on opaque. The curvature and spination call to mind that of Amphiodia urtica. Amphiodia sp. orange belly has similar coloration to Ophiura luetkenii. Development Mode. We suspect that this larva is a facultative planktotroph, like A. sp. opaque, but have lacked the material to conduct feeding experiments. We have observed larvae metamorphose after raised in FSW without algal cells, but larvae appear to have a complete larval digestive system and may be capable of eating and benefitting from larval feeding. Juvenile Description. Pentagonal juvenile, very similar to that of other Amphiodia spp. (Fig. 3.10L). Reproductive Timing. We have observed the larvae of A. sp. orange belly four times in the Oregon plankton. Collections were made in January and February, and just once in April (Fig. 3.3). Planktonic Duration. The planktonic duration of A. sp. orange belly is about 8 days (n=1). 3.4.3.5. Amphiodia sp. tan Species Identity. We know this species from a single larva that groups with Amphiodia orange belly but is genetically and morphologically distinct (Fig. 3.8, Table 3.5). We have no adult match. Distribution and Local Sites. Local adult populations remain unknown. Embryonic Description. The single specimen was collected in September 2020, as a tan blastula and recognized as an ophiuroid embryo from its thick hyaline coat. When first collected 8 8 it, we assumed incorrectly that it was A. sp. opaque; but the color was not orange or salmon typical of A. sp. opaque. Larval Description. Amphiodia sp. tan develops via a six-armed ophiopluteus with a semi-opaque epidermis tan in color. The early pluteus is dorsoventrally compressed with short posterolateral and anterolateral arms (Fig. 3.10M). The final ophiopluteus has curved posterolateral and anterolateral arms, that bear small thorns (Fig. 3.10O’). The larva has a mouth and stomach, but it is not clear if the larva feeds. The body rods are curved, and the end rods are short and parallel to one another. (Fig. 3.2F) The rudiment stage larva is distinct from the other amphiurid reduced plutei. The rudiment-stage larva retains part of the left anterolateral rod as well as the right anterolateral rod (Fig. 3.10P). Similar Species. The reduced pluteus of A. sp. tan is most like those of A. sp. opaque and A. sp. orange belly. The tan coloration of the epidermis is what distinguishes it, as does the manner of arm resorption. Development Mode. We suspect that A. sp. tan is a facultative planktotroph due to its reduced pluteus morphology and the presence of a mouth and stomach. We did not attempt to give the single individual food, but it successfully made a juvenile without any food. Juvenile Description. The juvenile is small and pentagonal like other Amphiodia spp. Reproductive Timing. We collected one early pluteus on September 19, 2022 (Fig. 3.4). Planktonic Duration. The larva we observed went from early pluteus to juvenile in approximately 11 days. 8 9 9 0 Figure 3.10. Reduced plutei of three species of Amphiodia: (A–E) Amphiodia sp. opaque, (F-L) Amphiodia sp. orange belly, and (M–R) Amphiodia sp. tan. Secondary views of the same individual are marked with an apostrophe (’). Scale bar in A is 100 µm and applies to all images on the plate. (A) Gastrula of A. sp. opaque with thick hyaline layer (hy). (A’) The newly forming pair of spicules are visible at the posterior in partially cross-polarized light. (B) Aboral view of the early pluteus, the gut is medial and not yet complete; (B’) The paired spicules have branched, forming rods that will support larval arms. (C) Oral view of the six-armed pluteus. (C’) In cross-polarized light, the anterolateral (al), postoral (po), and posterolateral (pl) arm rods are visible. The anterolateral and posterolateral arms may be straight or curved. (D) In the rudiment-stage pluteus, the juvenile rudiment (jr) forms around the larval mouth and resorbs all but the right anterolateral arm (ral) and the paired posterolateral arms (aboral view). (D’) The longer arm rods are adorned with thorns (open arrowheads) as the pluteus grows and develops. The body rods (br) and end rods (er) are straight, and the transverse rods (filled arrowhead) are thorny. (E) The juvenile (aboral view) is pentagonal, often with a red-pink coloration at the center of the disk. (F,G) The early pluteus of A. sp. orange belly (F, aboral view, G oral view) has a large left coelom (lc), mouth (m) and stomach (s). (H, I) The left coelom has developed lobes by the time the pluteus develops six arms (H oral view, I oral view of another individual). (J,K) The rudiment-stage pluteus (J oral view, K aboral view another individual) has both posterolateral arms and the right anterolateral arm, which are covered in thorns (K’). (L,L’) The juvenile in aboral view, of the same individual as K and photographed six days later, is very similar to juveniles of other Amphiodia spp. The tube feet (tf) at this stage are active. F-H, J are the same individual, photographed daily over four days; I, K, L are the same individual, photographed on two consecutive days (I, K) and then again six days later (L). (M) An aboral view of an early pluteus of A. sp. tan with anterior and posterior left coeloms (alc, plc). (M’) Same larva under cross-polarized light reveals the larval rods. (N) One day later and in oral view, the pluteus developed the postoral arms (out of focal plane) and the left coelomic lobes. (O,O’) The anterolateral and posterolateral arms continue to grow in length (aboral view) and bear very small thorns. (P) The rudiment-stage pluteus at first retains both right and left anterolateral arms (oral view, P’ aboral view). (Q) Eventually the rudiment-stage pluteus has only the right anterolateral arm (oral view) and tube feet are visible on the oral side of the juvenile rudiment. (R, R’) The pentagonal juvenile, photographed five days after Q, in oral (R) and aboral views (R’). 3.4.3.6. Amphiodia periercta? H.L. CLARK 1911 Species Identity. This larva was initially described as Amphiodia occidentalis (Emlet, 2006), but molecular evidence suggests otherwise. Within NE Pacific Amphiodia, there is a species complex containing at least three species: occidentalis, periercta?, and a population from the Queen Charlotte Islands, BC (Fig. 3.8, Table 3.5). The original discovery of this pelagic direct developer (1991) was the result of a spontaneous spawning in a bucket with many adults collected near Olga on Orcas Island, WA. Most of the adults were identified as A. occidentalis by R. Emlet using the key in Kozloff (Kozloff, 1987); and material collected later from Olga later in the 1990’s was confirmed as A. 9 1 occidentalis by G. Hendler (Los Angeles County Museum of Natural History). The same phenotype was found in the plankton in Charleston Oregon from 1994 onward and assumed to be A. occidentalis (Emlet, 2006). In 2011, R. Emlet barcoded specimens of this phenotype from Charleston, OR only to find that they were not a species-level match with adult specimens of A. occidentalis from our region or the A. occidentalis collected subsequently from the original site at Olga, WA. (Fig. 3.8, Table 3.5). In 2019, BioBlitz researchers (G. Paulay, personal communication) and Nakata collected sub-adults in eelgrass beds in the low intertidal at several sites in Coos Bay estuary and these specimens were a molecular match to this planktonic form (Fig. 3.8, Table 3.5). Based on comparison of this adult material with species descriptions and with material from the USNM (E13853), we have provisionally identified the adult and its larval phenotype as A. periercta? (G. Hendler, personal communication). Distribution and Local Sites. Amphiodia periercta occurs from the intertidal to 92 m depth from Unalaska (Aleutian) Island, Alaska to central California, (Lambert & Austin, 2007). We have collected small, infaunal adults in eelgrass beds at -0.5 m tidal height at sites in the marine dominated part of the Coos Bay estuary. Embryonic Description. The eggs are light pink to salmon colored and 190um in diameter, suggestive of their abbreviated development. See Emlet (2006) for a full description. Larval Description. Embryos and unhatched juveniles of A. periercta? and are easily recognized by the thick hyaline layer surrounding their cells as eggs and embryos (Fig. 3.11A, B). The embryo is also encased in a large, robust, and somewhat sticky fertilization envelope. Later in development, but still prior to hatching, the developing embryo is light pink, flattened along the (juvenile) oral-aboral axis and has characteristic peripheral ring of spicules (Fig. 9 2 3.11D, D'). After hatching, the scallop-shaped juvenile has a ring of skeletal elements supporting its periphery, and a stomodeal invagination (Fig. 3.11E). Similar Species. The unusual developmental pattern of A. periercta? means that it is not mistaken for any other ophiuroids whose development has been described. There is one other directly developing amphiurid in the region, Amphiura arcystata. Amphiodia periercta? is distinct from A. arcystata in the presence of a peripheral skeletal ring, and the absence of the interradial bulges that occur in A. arcystata. Development Mode. Amphiodia periercta? is a pelagic direct developer. It may represent the only of its kind, and was described in detail (Emlet, 2006). Juvenile Description. Development into a juvenile with moving podia and jaws is achieved in 7-8 days (Emlet, 2006) and juveniles at this stage are encountered in plankton tows during the winter months. Ophiuroids are known to occur as plankton even after they metamorphose (Hendler et al., 1999). Motile juveniles of A. periercta? are initially pentagonal but their terminal arms plates develop relatively rapidly into pointed rays (Fig. 3.11F) Reproductive Timing. All stages, from eggs, to embryos and hatched juveniles of A. periercta may occur in the plankton of Charleston, Oregon starting in mid-October, but especially from January to March after storms accompanied by large waves. Our daily plankton records from 2014-2016 suggests that A. periercta? spawns frequently throughout the winter (Fig. 3.3). This species is reproductive in the San Juan Islands of Washington in summer months, June to August (Emlet 2006 and 2019 unpublished observations). 9 3 Planktonic Duration. Planktonic duration is approximately eight days (Emlet, 2006). Cohorts of similarly aged stages were often discernible in our samples, though we could not determine if new cohorts represented a subsequent spawning event by the same adults. Other Remarks. As noted by Emlet (2006), the discovery of a pelagic directly developing larva disagreed with prior – albeit sparsely evidenced – accounts for developmental mode in A. occidentalis. Previous accounts for A. occidentalis list a yellow green egg 90-106 µm in diameter, which presumably developed into an obligately planktotrophic pluteus (Rumrill, 1982; Strathmann, 1987). At present we do not know the development of A. occidentalis, but it does have eggs similar in size and color to those of A. periercta? (R. Emlet, personal communication). We have collected adults often and never found them brooding. Despite continued attempts to induce spawning in gravid A. occidentalis and the extensive planktonic collection and sequencing of ophiuroid larvae presented here, we have never encountered larvae of A. occidentalis. We wonder if A. occidentalis develops in benthic capsules similar to Amphioplus abditus (Hendler, 1977). 9 4 9 5 Figure 3.11. Three nonfeeding planktonic forms of Amphiuridae: (A–F) Amphiodia periercta?, (G–J) Amphioplus sp. vitellaria, and (K–N) Amphiura arcystata. Secondary and tertiary views of the same individual are marked with an apostrophe (’) or quotations marks (”), respectively. (A) The fertilized egg of Amphiodia periercta? has a thick layer of hyaline (hy). The fertilization envelope (fe) is robust and sticky. (B) The two-cell stage. (C) The gastrula has a visible archenteron (ar). (D) Oral view of an unhatched A. periercta?. and (D’) in cross-polarized light. The embryo has a stomadeal invagination (st), left coelom (lc) and peripheral rods (pr) of skeleton. (E) After hatching the early juvenile has elaborate peripheral rods as the radials (r) develop. (F) The juvenile moves using its tube feet (tf) and resorbs the peripheral rods. Lateral views of (G) early and (H) late gastrula of Amphioplus sp. vitellaria, taken of the same individual one day apart. Note the pebbled exterior appearance due to many oil droplets interlaced with green pigmentation that are characteristic of this larva and the first spicules visible in partial cross-polarized light (solid arrowheads). (I) Aboral view of the vitellaria in transmitted light with the ciliary bands (cb) marked by asterisks (*). (I’) The same larva in oral view, with visible tube foot buds. (I”) The position of the juvenile radii (r) and interradii (ir) can be determined by identifying the arm spines under partial cross-polarized light. (J) Aboral view of the vitellaria juvenile. (K) The early juvenile of A. arcystata has a stomadeal invagination, and (K’) five calcified projections that become interradii of the juvenile disk, two of which are connected by ephemeral skeletal elements (closed arrowhead). (L) Another individual, viewed approximately one day later, has assumed pentaradial symmetry and the cross piece skeletal elements are visible from the aboral side under partial cross-polarized light. This stage is marked by rounded interradii that exceed the radii in size initially but that quickly recede in comparison to the growing arm rays. (M) Oral and (M’) aboral views of an early juvenile with buds of tube foot buds, pointed radii, and rounded interradii, one of which is larger and redder in color (open arrowhead). (N) The juvenile has a rounded disc with triangle-shaped terminal arm plates. The disc stereom under cross-polarized light is shown in the inset. 3.4.3.7. Amphioplus sp. vitellaria Species Identity. The species identity of this larva is uncertain, but it groups with other Amphioplus specimens from the northeast Pacific (Fig 8, Table 3.5). The sequences from our larvae are approximately 5% different from adults identified as Amphioplus strongyloplax (E6874, T. O’Hara, personal communication). Larval sequences are 4% different from two adult specimens collected from 450 m depth on the Oregon Shelf (OH, OB), that could not be identified to species because they consisted of a disk only and a young adult that lacked the characters necessary identification (G. Hendler, personal communication). All the members of our Amphioplus clade could be given the identification of A. strongyloplax based on the threshold of 8.5% from our ASAP analysis, but we gave this larva the provisional name of 9 6 Amphioplus sp. vitellaria due to our uncertainty on this identification. The genus is polyphyletic and requires revision (O’Hara et al., 2017) and we have few adult specimens of species from this region for comparison. Distribution and Local Sites. Two species of Amphioplus are known from this region, A. strongyloplax and A. macraspis. Amphioplus macraspis is widespread in the north Pacific, in Asia and from the Queen Charlotte Islands to Washington in the eastern Pacific, 1–876 meters depth. Amphioplus strongyloplax has a greater range in the eastern Pacific, ranging from the Gulf of Alaska to the Mexican border, 40–623 meters (Lambert & Austin, 2007). Adults of both species have been reported from benthic surveys off of Coos Bay, OR (Henkel & Gilbane, 2020). Embryonic Description. We did not observe the embryos of these species. Larval Description. The embryos and larvae of this species can be recognized at any stage by the presence of lipid droplets and dark green speckled pigmentation, visible through the tan epidermis. The vitellaria has disjunct ciliated bands that wrap laterally around the interradii, as well as anterior and posterior ciliated bands. Similar Species. The only other species to have a vitellaria stage in this region was Ophiopteris papillosa, whose vitellaria is preceded by an ophiopluteus. Furthermore, the vitellaria of O. papillosa has remnants of the larval arms and skeletal rods at the anterior that A. sp. vitellaria lacks, Finally, the vitellaria of O. papillosa has a white body with a deep red interior and yellow ciliated bands (Figure 3.14), whereas A. sp. vitellaria is light pink to tan, with ciliated bands slightly darker in color. Development Mode. The larva of A. sp. vitellaria is the first vitellaria larva to be described for the family Amphiuridae (Hendler, 1991). Nonfeeding, planktonic vitellaria have 9 7 evolved many time in the brittle stars, but appear to be restricted to the Ophintegrida O’HARA, HUGALL, THUY, STÖHR & MARTYNOV, 2017, in the genera Ophionereis, Ophioplocus, Ophiolepis, and Ophiacantha (Brogger et al., 2013; Hendler, 1979, 1982, 1995; Komatsu & Shosaku, 1993; Mortensen, 1938; Selvakumaraswamy & Byrne, 2000). At least two species, Ophiopeza spinosa and Ophioplocus esmarki are known to brood vitellaria larvae (Byrne et al., 2008; Sweet et al., 2019). Juvenile Description. The juvenile is tan in color and has a round disc with pointed terminal arm plates (Fig 11J). Reproductive Timing. Amphioplus sp. vitellaria occurs rarely in December and January in the Charleston plankton (Fig. 3.3). We usually find them as single individuals or pairs, not in high abundance. Planktonic Duration. Development of A. sp. vitellaria proceeds from a bullet-shaped gastrula to a nonfeeding vitellaria in 4-5 days. Metamorphosis into a motile juvenile takes another 2-3 days. 3.4.3.8. Amphiura arcystata H.L. CLARK 1911 Species Identity. We have barcoded eight of these larvae since 2012, and sequences generated for COI match to a single young adult identified as Amphiura arcystata (Fig. 3.8, Table 3.5). This may represent a range expansion for this species, which is unexpected due to its abbreviated development and likely limited dispersal. 9 8 Distribution and Local Sites. Amphiura arcystata is known from Mexico to Monterey Bay, California (Hendler, 1996). The presence of the short-lived larvae suggests a possible range expansion, but as we have not yet discovered adult populations locally. Embryonic Description. We have not observed the embryos of these species. Larval Description. We usually find this species as a pentaradial larva with a line of skeletal elements crossing the aboral surface and only visible under polarized light (Fig. 3.11K’, L). Similar skeletal elements have been observed in an undescribed direct developer from Australia (Emlet, unpublished data), and may represent ectopic vestiges of ophiopluteus skeleton. The developing juvenile of A. arcystata has five distal bulges that are the interradii: the juvenile arms soon develop between them (see arm stereom under polarized light in Fig. 3.11N). One of these bulges tends to be pink to orange red in color, in contrast to the mottled tan of the rest of the larva. The buds of the tube feet are often visible at this developmental stage, though they are not mobile or capable of holding on to substrate. Similar Species. The directly developing juvenile of A. arcystata could be mistaken for that of Amphiodia periercta?, which also develops directly in the plankton (see discussion above). However, the larva of A. periercta? has a peripheral ring of spicules while the larva of A. arcystata has a line of skeletal elements that cross the larval body. Amphiura arcystata could also be mistaken for Amphioplus sp. vitellaria, especially at developmental stages where the symmetry is unclear. Both share a tan coloration, but A. sp. vitellaria has a more pebbled exterior appearance. 9 9 Development Mode. We know this animal mainly from its pelagic, directly developing larva, which appears rarely in the Charleston plankton from November to January. The genus Amphiura is known to contain brooding species (n=17), pelagic lecithotrophs (3), and at least one planktotroph (A. filiformis) [citations]. Juvenile Description. The juvenile of this species is similar to other amphiurids, but thicker at the center of the disc (Fig. 3.11N). Reproductive Timing. The larvae and juveniles of A. arcystata occur rarely in the Charleston plankton from November to January. Planktonic Duration. 6 days (n=1). 3.4.4. Ophiacanthina O’HARA, HUGALL, THUY, STÖHR & MARTYNOV 2017 Species Identity. We found two species, Ophiacantha diplasia and Ophiopteris papillosa, that were the sole representatives of the superfamily Ophiacanthina (517 spp.; O’Hara et al. 2017). Despite not being very closely related (Fig. 3.12, Table 3.6), the ophioplutei of these species greatly resemble one another, but may be delineated by features of the skeleton and style of metamorphosis. Development Mode. Ophiacantha diplasia and Ophiopteris papillosa both develop as eight-armed ophioplutei and give rise to an advanced and motile juvenile with multi-segmented arms. However, O. papillosa transforms from a feeding ophiopluteus into a vitellaria before metamorphosing into a juvenile. These two species are the first of their kind to be described for their families. Ophiacantha diplasia is the first planktotroph from the family Ophiacanthidae 10 0 Figure 3.12. Maximum likelihood tree for a concatenated dataset of 16S and COI sequences for superfamily Ophiacanthina from the Pacific Ocean. Barcodes from larvae identified in this study are in bold. Bootstrap values of 70 and higher are shown beside nodes. Table 3.6. Specimen information for species of superfamily Ophiacanthina from the Pacific included in Fig. 3.12. New sequence data from this study is in bolded text. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. 10 1 Species Specimen Locus BOLDB or Collection Reference Code GenBankG Locality Clarkcoma JN593756 COI JN593756G Australia Naughton et al., canaliculata unpublished Clarkcoma pulchra JN593681 COI JN593681G Australia Naughton et al., unpublished Ophiacantha aff. rosea GU806183 COI GU806183G Australia iBOL1, unpublished Ophiacantha antarctica GU806354 COI GU806354G Antarctica iBOL1, unpublished Ophiacantha bidentata HM400539 COI HM400539G Nunavut, (Corstorphine, 2011) Canada Ophiacantha GU806225 COI GU806225G Antarctica iBOL1, unpublished brachygnatha Ophiacantha diplasia 1002 COI OLAB069-23B Charleston, OR This study Ophiacantha diplasia 935 16S, COI OLAB107-23B Charleston, OR This study Ophiacantha diplasia 942 16S, COI OLAB108-23B Charleston, OR This study Ophiacantha diplasia 964 COI OLAB068-23B Charleston, OR This study Ophiacantha diplasia grn 1-5-19 16S OLAB070-23B Charleston, OR This study Ophiacantha diplasia MMB9 16S, COI OOPH045-23B Stonewall Bank, This study OR Ophiacantha diplasia O1 2-5-15 16S, COI OLAB071-23B Charleston, OR This study Ophiacantha opulenta GU806364 COI GU806364G Antarctica iBOL1, unpublished Ophiacantha rosea KU895383 COI KU895383G Australia (Hugall et al., 2016) Ophiacantha GU806187 COI GU806187G Australia iBOL1, unpublished spectabilis Ophiacantha GU806192 COI GU806192G Australia iBOL1, unpublished vepractica Ophiacantha vivipara GU806366 COI GU806366G Antarctica iBOL1, unpublished Ophiocamax drygalskii KU895118 COI KU895118G Antarctica (Hugall et al., 2016) Ophiophthalmus MMB10 16S, COI OOPH041-23B Newport, OR This study cataleimmoidus Ophiophthalmus HM542946 COI HM542946G BC, Canada (Corstorphine, 2011) cataleimmoidus Ophiophthalmus OF COI OOPH018-18B Newport, OR This study normani Ophiopristis luctosa KU895397 COI KU895397G Australia (Hugall et al., 2016) Ophiopristis procera KU895396 COI KU895396G Papua New (Hugall et al., 2016) Guinea Ophiopteris antipodum JN593395 COI JN593395G Australia Naughton et al., unpublished Ophiopteris antipodum JN593396 COI JN593396G Australia Naughton et al., unpublished 10 2 Ophiopteris antipodum JN593397 COI JN593397G Australia Naughton et al., unpublished Ophiopteris antipodum KU895344 COI KU895344G New Zealand (Hugall et al., 2016) Ophiopteris papillosa 1538 COI OLAB083-23B Charleston, OR This study Ophiopteris papillosa H12-op COI OLAB084-23B Charleston, OR This study Ophiopteris papillosa H16-Op 16S OLAB085-23B Charleston, OR This study Ophiopteris papillosa JN593399 COI JN593399G Australia Naughton et al., unpublished Ophiopteris papillosa JN593400 COI JN593400G Australia Naughton et al., unpublished Ophiopteris papillosa JN593401 COI JN593401G Australia Naughton et al., unpublished Ophiopteris papillosa KU895345 COI KU895345G USA (Hugall et al., 2016) Ophiopteris papillosa R48 16S OOPH054-23B Castle Rock, CA This study2 Ophiopteris papillosa yop juve 16S OLAB086-23B Charleston, OR This study Ophiopteris papillosa yop2 16S OLAB087-23B Charleston, OR This study Ophiopteris papillosa yyopDec18 16S OLAB088-23B Charleston, OR This study Ophiotreta larissae KU895402 COI KU895402G Papua New (Hugall et al., 2016) Guinea 1International Barcode of Life, 2Specimen borrowed from the Natural History Museum of Los Angeles County (NHMLAC). LJUNGMAN 1867, and Ophiopteris papillosa is the first vitellaria from Ophiopteridae O’HARA, STÖHR, HUGALL, THUY & MARTYNOV 2018. 3.4.4.1. Ophiacantha diplasia H. L. CLARK 1911 Species Identity. We encountered several larvae of various developmental stages in 2015 and 2019 (Fig. 3.3). Barcoding revealed these larvae to be a match to a specimen we collected on the Stonewall Bank off the coast of Newport, Oregon that we identified as Ophiacantha diplasia (Figure 12, Table 3.6). 10 3 Distribution and Local Sites. Ophiacantha diplasia occurs from Haida Gwaii to Southern California at depths of 71 to 1178 meters on silty sand or hard rock (Astrahantseff & Alton, 1965; Lambert & Austin, 2007). We have encountered the adult animal only once, on Stonewall Bank, 307 m (MMB9, Table 3.6). Embryonic Description. We did not observe embryos of this species. Larval Description. Ophiacantha diplasia develops via an eight-armed feeding ophiopluteus. The epidermis is transparent, with yellow-green pigmentation at the distal ends of the larval arms. The posterolateral arms are curved near the posterior girdle, seen in the early eight-arm stage (Fig. 3.13B), and straight as they lengthen in the advanced ophiopluteus (Fig. 3.13C, D). The anterolateral arms are straight and project anteriorly beyond other arms at all stages of development. The postoral and posterodorsal arms are long, with their distal tips of an equal height with the posterolateral arms. The posterior is supported by the body rods and recurrent rods. The right dorsal recurrent rod may bear a median process, but we have also observed individuals that lack a median process. The skeletal rods lack thorns. The eight-armed pluteus has vibratile lobes, prominent enlargements of the ciliary band at the bases of the posterolateral arms. Vibratile lobes are known also from the ophioplutei from the genus Ophiocoma L. AGASSIZ 1836 (Hendler, 1991). Like the epaulettes of echinoplutei, the ophiopluteus vibratile lobes may act in locomotion (Mortensen, 1921; Strathmann, 1975) The juvenile rudiment is star shaped, and all or most larval arms are retained until metamorphosis (Fig. 3.13E). Similar Species. In the northeast Pacific, the pluteus of O. diplasia is most likely to be confused with that of Ophiopteris papillosa. The larval epidermis of these two species is 10 4 transparent and accented with touches of yellow green at the distal tips of the arms. Their posterolateral arms are not markedly longer than other arms. The advanced larvae of both species have vibratile lobes, at first giving the impression of an echinopluteus. However, careful examination of the placement of the pluteus arms shows them to be ophioplutei. If one has access to a microscope with cross-polarized light, the posterior girdles of the two species can be used to differentiate them at all stages: O. diplasia has recurrent rods, even as a young pluteus, while O. papillosa lacks these. The manner of metamorphosis also distinguishes between the species: O. diplasia forms a juvenile rudiment as a pluteus, while O. papillosa transforms into a vitellaria before metamorphosis. Development Mode. Ophiacantha diplasia has a feeding ophiopluteus larva. The bright red coloration of the stomach in wild-caught specimens is due their algal diet (Fig. 3.13B-D). Juvenile Description. At metamorphosis the juvenile has arms with up to three segments and is highly motile. Juvenile arm tips are a yellow-green color, a feature that is absent from arms of juvenile O. papillosa. Reproductive Timing. Larvae were observed infrequently in the Charleston plankton from January to April. Planktonic Duration. We estimated planktonic duration as 82 days for a single larva collected from the plankton as a young ophiopluteus and raised in the laboratory on a mixed microalgal diet. 10 5 Figure 3.13. Ophioplutei and juveniles of Ophiacantha diplasia, all in aboral view. Secondary and tertiary views of the same individual are marked with an apostrophe (’) and quotation mark (”), respectively. (A, B) Aboral views of six- and eight-armed ophioplutei, respectively (postoral arms cannot be seen). The arms tips are yellow green in 10 6 color. (A’, B’) In cross-polarized light the skeletal rods that support the posterolateral (pl), anterolateral (al), posterodorsal (pd), and postoral (po) arms are visible. (A”, B”) Closeups of the posterior girdles with recurrent rod (rr), transverse rod (filled arrowhead), median process (open arrowhead), body rod (br), and end rod (er) visible. (C) An eight-armed pluteus, with modest vibratile lobes (vl). (C”) Detail of yellow-green pigmentation at the distal end of the arms. (D) Eight-armed pluteus with well-formed vibratile lobes (vl), and (po) arms in hind ground. (D’) Closeup of the posterior girdle in partially cross-polarized light. (E) A rudiment-stage larva, note larval arms are present and still long. (E’) Closeup of the developing juvenile arms in partially cross-polarized light. (F) A juvenile with 3 arm segments. (G) Detail of the disk and arm stereom of a juvenile in partially cross-polarized light. Scale in A is 100 µm and applies for B, D, and E. No scale for C, F, G. 3.4.4.2. Ophiopteris papillosa (LYMAN 1875) Species Identity. We were able to connect larvae at different developmental stages to the species Ophiopteris papillosa through DNA barcoding and comparison with sequences for adult specimens from GenBank or material borrowed from NHMLAC (Fig. 3.12, Table 3.6). The family Ophiopteridae contains just two species, Ophiopteris papillosa throughout the west coast of North America, and O. antipodum to the south (Naughton et al., 2014). Distribution and Local Sites. Coos Bay, Oregon, is well within the range of O. papillosa, which occurs from Barkley Sound, British Columbia, and southern Oregon to Isla Cedros, Baja California (Austin & Hadfield, 1980; Lambert & Austin, 2007). Previously, a planktotrophic larva was predicted by the small egg size, diameter 100 µm, of the species (Pearse, 1994). Embryonic Description. We did not observe the embryos of these species. Larval Description. The larval forms of O. papillosa have a transparent epidermis, sometimes with yellow to green pigmentation on at the distal ends of the larval arms and on the ciliary bands. As an early pluteus O. papillosa resembles Ophiacantha diplasia, but can be distinguished by its single body rods, which are double (body rod and recurrent rod) in O. 10 7 diplasia. In later pluteal stages the posterolateral arms are not markedly longer than the others, and postoral and posterodorsal arms shorter in height than the anterolateral arms. The larval arms may, at their distal end, bend outward except those of the anterolateral arms may point medially (Fig. 3.14). After approximately a month, the ophiopluteus develops into a vitellaria (Fig. 3.14D). The developing juvenile body at the center is white and may have a dark red gut with five lobes projecting into the interradial spaces. The ciliated band of the pluteus reorganizes into complete or partial transverse ciliated bands that are yellow in color and distributed in four locations from anterior or posterior (Fig. 3.14D, D’). The anterior most ciliated band wraps around the distal ends of the remnants of larval arms. The juvenile arms are multisegmented and develop folded toward the center of the oral side. Similar Species. As an ophiopluteus, O. papillosa resembles Ophiacantha diplasia (see above). There is only one other vitellaria that occurs in this region that O. papillosa could be confused with, that of Amphioplus sp. vitellaria (Section 3.4.3.6), which differs from O. papillosa in several ways. First, the vitellaria of O. papillosa has a mostly transparent epidermis with yellow-green coloration at the ciliated bands. Vestiges of the pluteus skeleton are visible as two rods bearing loops of ciliated band projecting from the anterior end of the vitellaria (Fig 14D), a feature absent in Amphioplus sp. vitellaria. Development Mode. Ophiopteris papillosa has a feeding ophiopluteus that transforms into a nonfeeding vitellaria before metamorphosis into a juvenile. This type of development is known previously from Ophiocomella pumila (Cisternas & Byrne, 2005). 10 8 Figure 3.14. Ophioplutei, vitellaria, and juvenile of Ophiopteris papillosa. Secondary and tertiary views of the same individual are marked with an apostrophe (’) and quotation mark (”), respectively. (A) An oral view in DIC optics and (A’) partially polarized light which highlights the larval skeleton of six-armed pluteus at time of collection and (B) eight days later. (B’) The posterior girdle in cross-polarized light with postoral (po) and posterolateral (pl) arm rods, thorny transverse rods (filled arrowhead), body rods (br), and end rods (er). (C) The advanced eight-arm pluteus shown in aboral view has anterolateral (al), posterodorsal, postoral, and posterolateral arms, and vibratile lobes (vl). Note the way the portion of ciliated band at the distal end of the anterolateral arm (open arrowhead) bends medially. (C’) An oral view of the same individual shows the mouth (m), stomach (s), and lobed left coelom (lc). (C”) The vibratile lobes are shown in a posterior view. (D) The eight-armed ophiopluteus develops into a vitellaria, shown in aboral and (D’) oral view. (E) An early juvenile, still bearing remnants of the yellow ciliated bands (*). The scale bar in A is 100 µm and applies to B-E. 10 9 Ophiopteris antipodum is inferred to have planktotrophic development from egg size (Naughton et al., 2014). It is possible that O. antipodum also has an ophiopluteus followed by a vitellaria stage like O. papillosa. Juvenile Description. The juvenile of this species develops from the vitellaria larva. The developing arms are folded between the ciliated bands of the vitellaria on the oral side. The juvenile may be motile while still bearing remnants of the ciliated bands. Reproductive Timing. We found a small number of 8-armed ophioplutei in the winter of 2015, from November to February, and rarely in April (Fig. 3.3). Planktonic Duration. We estimated planktonic duration as 65 days for a single larva collected from the plankton as a young ophiopluteus and raised in the laboratory on a mixed microalgal diet. 3.4.5. Ophiuridae MÜLLER & TROSCHEL 1840 We observed the ophioplutei of four species from the family Ophiuridae, all of which were identified using DNA barcoding (Fig. 3.15, Table 3.7). The family has 65 described species (O’Hara et al., 2017), and is dominated by the genus Ophiura, three of which are known from the northeastern Pacific and documented here: O. leptoctenia, O. luetkenii and O. sarsii (Lambert & Austin, 2007). The final species from the family we observed was Ophiocten hastatum, a deep-water species that occurs at depths of 1000 m and greater. 11 0 3.4.5.1. Ophiocten hastatum LYMAN 1878 Species Identity. We observed a single larva from this species, collected on January 19, 2015. The large ophiopluteus (Fig. 3.2L) was a species-level match at the COI locus with two adult sequences from GenBank (Fig. 3.15, Table 3.7). Distribution and Local Sites. Ophiocten hastatum has a cosmopolitan distribution at bathyal to abyssal depths, occurring at over 2000 m depth in the NE Atlantic, 1408–2877 m in tropical eastern Pacific, 916–2877 m in British Columbia and Washington (Gage et al., 2004; Kyte, 1969; Lütken & Mortensen, 1899). Embryonic Description. We did not observe the embryos of these species. Larval Description. The larva we collected was a large, eight-armed ophiopluteus and obligately planktotrophic. The larval arms were long and bilaterally asymmetric in length, especially the posterolateral arms. This asymmetry could be due to earlier damage. Reddish brown coloration on the posterolateral arms approximately halfway up the length and at the distal tenth (of one arm), distinguish this larva from all other ophioplutei we collected (Fig. 3.2L). The posterior girdle is supported by end rods, relatively short body rods, and recurrent rods of the same length, the oral and aboral pairs of which lie largely over one another so it’s difficult to see both pairs (Fig. 3.16J). Similar Species. The advanced eight-armed ophiopluteus of O. hastatum is not easily confused with any of the other plutei we observed, but the presence of recurrent rods may be used to distinguish it from other large ophioplutei in this study. 11 1 Figure 3.15. Maximum likelihood tree for a concatenated dataset of 16S and COI sequences for Ophiuridae spp. Barcodes for larvae identified in this study are in bold. Table 3.7. Specimen information for Ophiuridae spp. included in Fig. 3.15. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. Species Specimen Locus BOLDB or Collection Reference / Field ID Code GenBankG Locality 11 2 Ophiocten gracilis KU895451 COI KU895451G Ireland (Hugall et al., 2016)12/20/23 3:47:00 PM Ophiocten hastatum HM542940 COI HM542940G British (Corstorphine, 2011) Columbia, Canada Ophiocten hastatum HQ946167 COI HQ946167G Australia iBOL1, unpublished Ophiocten hastatum Otx1 16S, COI OLAB072-23B Charleston, OR This study Ophiocten sericeum HM405894 COI HM405894G Prince of Wales (Corstorphine, 2011) Strait, Canada Ophionotus hexactis KU895454 COI KU895454G Antarctica (Hugall et al., 2016) Ophionotus victoriae KY048258 16S KY048258G Antarctica (Galaska et al., 2017) Ophiura albida KX459039 COI KX459039G North Sea (Laakmann et al., 2017) Ophiura ambigua KU894970 COI KU894970G Antarctica (Hugall et al., 2016) Ophiura flagellata KU894987 COI KU894987G Australia (Hugall et al., 2016) Ophiura flexibilis KU894984 KU894984G Antarctica (Hugall et al., 2016) Ophiura irrorata KU894969 KU894969G Australia (Hugall et al., 2016) Ophiura leptoctenia Opl 16S, COI OOPH051-23B Cape Arago This study Shelf, OR Ophiura leptoctenia OpE 16S, COI OLAB094-23B Charleston, OR This study Ophiura luetkenii BOIMB- COI Cape Arago G. Paulay, personal 2131 Shelf, OR communication Ophiura luetkenii Creamsicle2 16S OLAB095-23B Charleston, OR This study Ophiura luetkenii HM542957 COI HM542957G British (Corstorphine, 2011) Columbia, Canada Ophiura luetkenii KU495745 COI KU495745G British (Corstorphine, 2011) Columbia, Canada Ophiura luetkenii MMB3 COI OOPH042-23B Cape Arago This study Shelf, OR Ophiura luetkenii Ol1 16S OOPH008-18B Cape Arago, Ol1 OR Ophiura luetkenii Olc1 16S OOPH010-18B Catalina Is., CA Olc1 Ophiura luetkenii Olc2 16S OOPH011-18B Catalina Is., CA Olc2 Ophiura ophiura KX459054 COI KX459054G North Sea (Laakmann et al., 2017) Ophiura rouchi KR861611 COI KR861611G Antarctica (Sands et al., 2015) Ophiura sarsii HM473940 COI HM473940G British (Corstorphine, 2011) Columbia, Canada 11 3 Ophiura sarsii KU495754 COI KU495754G Baffin Bay, (Corstorphine, 2011) Canada Ophiura sarsii MMB5 16S, COI OOPH043-23B Cape Arago This study Shelf, OR Ophiura sarsii MMB6 16S, COI OOPH044-23B Cape Arago This study Shelf, OR Ophiura sarsii OD 16S, COI OOPH016-18B This study Ophiura sarsii Ophiura 5- COI OLAB096-23B Charleston, OR This study 20-19 Ophiuroglypha lymani HM425597 COI HM425597G Antarctica iBOL1, unpublished 1International Barcode of Life Development Mode. The larva we found confirms that Ophiocten hastatum is planktotrophic, which was suggested by small size of oocytes seen in histological sections of the gonads of adults from the Porcupine Abyssal Plain in the north Atlantic (Gage et al., 2004). The timing of observation of this larva in the Pacific differs somewhat from spawning phenology in the Atlantic, where spawning is inferred for late winter and planktotrophic development until settlement in summer (Gage et al., 2004). Two congeners, O. gracilis and O. sericeum, also have feeding ophioplutei (Gage & Tyler, 1981; Thorson, 1934; Tyler & Gage, 1982, 1980). Juvenile Description. We did not observe the juvenile of this species. Reproductive Timing. We observed this larva just once, on January 18, 2015, a warm water year. Planktonic Duration. Unknown. 11 4 Figure 3.16. Ophioplutei of four species of Ophiuridae: (A) Ophiura leptoctenia, (B–F) O. luetkenii, (G–I) O. sarsii, and (J) Ophiocten hastatum. Secondary views of the same individual are marked with an apostrophe (’). (A) Ophiura leptoctenia, (B–F) O. luetkenii, (G–I) O. sarsii, and (J) Ophiocten hastatum. Scale bar in A is 100 µm; same scale for A–I. (A) Oral view of eight-armed ophiopluteus of O O. leptoctenia in transmitted light has posterodorsal arms (pd), and small right and left coeloms (rc, lc). (A’) The posterior girdle in cross-polarized light, with transverse rods (filled arrowhead), recurrent rods (rr), body rods (br) and end rods (er). (B) Oral views of O. luetkenii at the gastrula, (C) early pluteus, and (D) six-armed reduced pluteus stages. The reduced pluteus has 11 5 anterolateral (al), postoral (po), and posterolateral (pl) arms, a stomach (s), and left coelom (lc). (D’) The posterior girdle in partial cross-polarized light shows the transverse rods (filled arrowhead), body rods (br), and end rods (er). (E) The rudiment-stage larva has a star-shaped juvenile rudiment with five tubefoot buds (tf) on each radius, with two pairs leading proximally to the mouth and a single terminal podium. (F) The juvenile of O. luetkenii in cross- polarized light, aboral view. Note the honeycomb pattern of the primary plates and disc plate. (G) Pluteus of O. sarsii, aboral view, and (H) aboral view of a rudiment-stage pluteus, and (I) it’s juvenile a day later. (J) The posterior girdle of Ophiocten hastatum is of the same individual as Fig. 3.2L. 3.4.5.2. Ophiura leptoctenia H. L. CLARK 1911 Species Identity. We observed a single larva on May 8, 2019, differed by 0.9% at the COI locus to an adult specimen collected from the continental shelf off Oregon (Fig. 3.15; Table 3.7). Distribution and Local Sites. Ophiura leptoctenia was characterized by Clark (1911) as one of the most common ophiuroid species in the north Pacific, and they occur in large numbers in off California (Astrahantseff & Alton, 1965). This species has a bathyal depth range, 122- 3239 m (Hendler, 1996). We collected one adult from the Cape Arago shelf at 120 m depth. Embryonic Description. We did not observe the embryos of these species. Larval Description. The advanced ophiopluteus of O. leptoctenia has eight arms, and a transparent epidermis that appears inflated due to the distance from the skeletal rods. The epidermis tapers at the distal tip of the posterolateral arms. The skeletal rods lack thorns and curve slightly toward the midline (Fig. 3.16A). The body rods are short and there are ventral and dorsal recurrent rods (Fig. 3.16A’). Similar Species. The ophiopluteus of O. leptoctenia is very similar to that of O. sarsii. Both have eight arms, an inflated transparent epidermis, and a similarly structured posterior girdle with short body rods and recurrent rods. The pluteus of Amphipholis pugetana also has an 11 6 inflated epidermis, but it is of a uniform width along the length of the arm and terminates with red pigmentation at the distal ends of the posterolateral arms. The epidermis around the posterolateral arms of O. leptoctenia is widest at approximately halfway the length, and then tapers to a dull point at the distal end. Development Mode. To our knowledge, this is the first report of the larva of O. leptoctenia. Hendler (1996) reported an egg diameter of ca. 0.2 mm and suggested the species had either abbreviated development or an ophiopluteus. The transparent epidermis, small stomach, and lack of a developing rudiment suggest to us that this species is planktotrophic. Juvenile Description. We did not observe the juvenile of O. leptoctenia. Reproductive Timing. We observed a single larva on May 8, 2019, an El Niño year. Planktonic Duration. Unknown. 3.4.5.3. Ophiura luetkenii (LYMAN 1860) Species Identity. DNA barcoding revealed that this white and orange colored larva is that of Ophiura luetkenii, which is closely related to O. sarsii (Fig. 3.15, Table 3.7). Distribution and Local Sites. Ophiura luetkenii occurs in large aggregations across its range, including off the Oregon coast (Hemery et al., 2018; Henkel et al., 2014) on the continental shelf off Cape Arago. We have collected in large numbers of adults along with those of its congener O. sarsii. Embryonic Description. O. luetkenii spawned in the laboratory on one occasion. After repeat flipping adults under bright lights at room temperature, three females spawned 11 7 approximately eight hours later in the early evening; we fertilized the eggs with sperm dissected from a male. The earliest stage we have observed from the plankton is the gastrula, which is ovoid and dorsoventrally flattened. The gastrula is orange with deeper color at the anterior lightening to the posterior. Larval Description. Ophiura luetkenii has a reduced pluteus. The early pluteus has posterolateral arms, a mouth, and visible left and right coeloms (Fig. 3.16C, C’). At the six- armed stage the pluteus the posterolateral arms are translucent white, and the body region is orange. The places where the posterolateral and anterolateral arms meet are elevated towards the anterior of the larva compared to other ophioplutei. At the posterior end the epidermis is distinctively swollen around the body and end rods (Fig. 3.16D, E). The larva is large and develops into a juvenile in about seven days. Similar Species. The orange body and white arms of the larvae of O. luetkenii give it a superficial resemblance to that of Amphiodia sp. orange belly, and even certain individuals of Amphiodia sp. opaque. However, the swollen posterior end of the larva of O. luetkenii, and the resulting juvenile should all clue the reader in to its identity. Development Mode. Ophiura luetkenii has a reduced pluteus that may also be facultatively planktotrophic (Nakata and Emlet, unpublished data). Juvenile Description. The juvenile is large and has elongate terminal arm segments, approximately one third the length of the diameter of the juvenile disc (Fig. 3.16F). Podia are papillate. 11 8 Reproductive Timing. We observed the embryos and larvae of O. luetkenii in February and March. Planktonic Duration. Approximately 9 days, based on larvae raised in the laboratory at 15°C (N. Nakata, personal observation). 3.4.5.4. Ophiura sarsii LÜTKEN 1855 Species Identity. We identified the larva of O. sarsii from DNA barcoding and by comparing our sequences to GenBank and to those of adults of the Oregon coast (Fig. 3.16G, Table 3.7). Distribution and Local Sites. Ophiura sarsii occurs throughout the Arctic Ocean, the northern Atlantic, and in Pacific, from the Bering Sea to California, Japan and Korea, from 0– 1460 meters (Lambert & Austin, 2007). Adults occur in large aggregations on the continental shelf off the Cape Arago and Columbia River (Astrahantseff & Alton, 1965) on muddy bottoms. Given the abundance of adults we are surprised that we have collected only two larvae of O. sarsii. Embryonic Description. We did not observe the embryos of these species. Larval Description. We collected two larvae on May 20, 2019, including a six-armed pluteus and a rudiment-stage larva of O. sarsii (Fig. 3.16F–H). The ophiopluteus (Fig. 3.16F) has a transparent epidermis and eight arms. The individual we observed had not fully developed the posterodorsal arms, but the skeletal rods are visible on the dorsal side. The body rods are short and bear long end rods that are wide apart at their origin and nearly touching at their distal ends. Images of ophioplutei of O. sarsii prior to rudiment 11 9 formation suggest that this arrangement occurs to accommodate the juvenile rudiment and that the long end rods are closer to one another at the proximal sides in the ophiopluteus (Thorson, 1934). There may also be variation in the transverse rods. In the rudiment-stage larva the right anterolateral and postoral arms had been resorbed and there was a developing juvenile rudiment (Fig. 3.16H). The larva bore a striking resemblance to Mortensen’s (1921) Ophiopluteus fusus. The remnants of the larval body are transparent in color, with a small number of red to orange pigment cells occurring along the skeleton of the posterolateral arms. Similar Species. The pluteus of O. sarsii is very similar to that of O. leptoctenia (see section above, Fig. 3.16A, G). Development Mode. Ophiura sarsii develops via a small egg of 100–110 µm diameter that gives rise to a planktotrophic ophiopluteus (Dautov & Selina, 2009; Kungurtzeva & Dautov, 2001; Strathmann, 1987). Juvenile Description. The juvenile of O. sarsii is very similar to that of O. luetkenii in skeletal structure (Fig. 3.16I, F), but was tan in color in the single specimen we observed, compared to the orange coloration of O. luetkenii. Reproductive Timing. The two larvae we observed, including the rudiment-stage ophiopluteus on May 20, 2019, is in accordance with published accounts of spawning period occurring from March to June (Strathmann, 1987). Planktonic Duration. At least 26 days (R. Emlet, unpublished data). 12 0 3.4.6. Gorgonocephalidae LJUNGMAN 1867 3.4.6.1. Gorgonocephalus eucnemis (MÜLLER & TROSCHEL 1842) Species Identity. The genus Gorgonocephalus is comprised of ten species with an anti- tropical distribution. Five species occur in the northern hemisphere: eucnemis and diomedeae in the north Pacific; arcticus in the Arctic; and lamarcki, caputmedusae in the north Atlantic (D’yakonov, 1967). Gorgonocephalus eucnemis represents several synonymies: G. caryi, japonica, and stimpstonii (Clark, 1911). Our own analysis shows several species within the genus to be closely related at the COI locus (within 6%; Fig. 3.17, Table 3.8). The sequences from our larvae were identical to that of an adult we collected off Cape Arago and another from California. The clade formed by our specimens was divergent from another clade of G. eucnemis from the west coast of Canada and Asia (clades 1, 2 in Fig. 3.17). Distribution and Local Sites. Gorgonocephalus eucnemis is known to occur in the North Atlantic and in the eastern Pacific from the Bering Sea south to Laguna Beach, CA at depths of 8-1850 m (Austin & Hadfield, 1980; Lambert & Austin, 2007). In the northeast Pacific G. eucnemis occurs from Alaska to California, from the shallow subtidal to 1240 meters (Lambert & Austin, 2007). No other basket stars occur in our region; though a congener, G. diomedeae, occurs in Panama. Embryonic Description. Eggs are approximately 0.22 mm diameter and give rise to nonfeeding embryos and juveniles (Emlet, unpublished data). The development has been described previously under the synonymized name G. caryi (Patent, 1970). The embryos of G. eucnemis are pear-shaped (Fig. 3.18A) and have a thick hyaline layer with a textured appearance 12 1 like that of translucent spheres (Fig. 3.18B’). The peach to salmon colored embryo is unciliated and negatively buoyant. Embryos develop into pentaradial juveniles within a week and have five arm buds by three days post spawn, and complete pentaradial symmetry by day four. Juvenile Description. Juveniles have five lobate arms bearing symmetrical claws at their distal ends. The pink to beige epidermis is still semi-transparent, and one can see the juvenile skeleton (Fig. 3.18E). The arms of early juveniles of Gorgonocephalus that have not yet begun to branch are thought to associate with the soft coral Gersemia rubiformis (Austin & Hadfield, 1980). Similar Species. The early juvenile of Amphiodia periercta? is also pink in color, unciliated, and is dorsal-ventrally flattened. However, the early juvenile of G. eucnemis can be distinguished by its textured hyaline layer, which is smooth in Amphiodia, and the pear shape of the embryo. The juvenile can be distinguished from those of other ophiuroids in the region by the absence of stereom in the central disc. Development Mode. Freely spawned, fertilized eggs give rise to nonfeeding embryos and juveniles. Development is unknown for other members of Gorgonocephalus, but G. caputmedusae free-spawns eggs, but it is unknown if they develop into an ophiopluteus larva (Mortensen, 1927). Reproductive Timing. We observed embryos and juveniles of G. eucnemis in the Charleston plankton a few times each year from late January to late March (Fig. 3.3). Planktonic Duration. Approximately 5 days (Nakata and Emlet, unpublished data), similar to that found by (Patent, 1970). 12 2 Figure 3.17. Maximum likelihood tree of COI for 27 spp. of Gorgonocephalus from the Pacific and the outgroup Ophiura sarsii. Barcodes of larvae identified in this study are in bold. Bootstrap values of 70 and higher are shown beside nodes. Table 3.8. Specimen information for Gorgonocephalus spp. included in Fig. 3.17. Additional information for specimens collected in this study can be accessed using the BOLD Project IDs below. Species Specimen Locus BOLDB or Collection Reference Code GenBankG Locality Gorgonocephalus arcticus HM542197 COI HM542197G New Brunswick, (Corstorphine, 2011) Canada Gorgonocephalus arcticus HM543017 COI HM543017G Nunavut, (Corstorphine, 2011) Canada Gorgonocephalus arcticus KR919684 COI KR919684G Korea (Kim & Shin, 2015) Gorgonocephalus arcticus KU495867 COI KU495867G Baffin Bay, (Corstorphine, 2011) Canada 12 3 Gorgonocephalus MG935270 COI MG935270G Sweden Lundin et al., caputmedusae unpublished Gorgonocephalus MG935339 COI MG935339G Sweden Lundin et al., caputmedusae unpublished Gorgonocephalus chilensis AB758812 COI AB758812G Antarctica (Okanishi & Fujita, 2013) Gorgonocephalus chilensis KM491789 COI KM491789G Antarctica (Summers et al., 2014) Gorgonocephalus chilensis KU895116 COI KU895116G New Zealand (Hugall et al., 2016) Gorgonocephalus eucnemis DISA697- COI DISA697-19B San Diego, CA DISCO MBC 19 LACM, unpublished Gorgonocephalus eucnemis G201 COI OLAB104-23B Charleston, OR This study Gorgonocephalus eucnemis gor COI OLAB105-23B Charleston, OR This study Gorgonocephalus eucnemis Gorgono- COI OOPH055-23B Cape Arago This study cephalus Shelf, OR Gorgonocephalus eucnemis GU670194 COI GU670194G Queen Charlotte (Corstorphine, 2011) Is., Canada Gorgonocephalus eucnemis HM473910 COI HM473910G Queen Charlotte (Corstorphine, 2011) Is., Canada Gorgonocephalus eucnemis HM542198 COI HM542198G British (Corstorphine, 2011) Columbia, Canada Gorgonocephalus eucnemis KR919685 COI KR919685G South Korea (Kim & Shin, 2015) Gorgonocephalus eucnemis Op9 COI OLAB066-23B Charleston, OR This study Gorgonocephalus eucnemis R34 COI OLAB067-23B Charleston, OR This study Gorgonocephalus AB758810 COI AB758810G New Zealand (Okanishi & Fujita, pustulatum 2013) Gorgonocephalus KU895114 COI KU895114G New Zealand (Hugall et al., 2016) pustulatum Gorgonocephalus sundanus HM400460 COI HM400460G Australia iBOL1, unpublished Gorgonocephalus sundanus KU895115 COI KU895115G Australia (Hugall et al., 2016) Gorgonocephalus tuberosus AB758811 COI AB758811G Antarctica (Okanishi & Fujita, 2013) Gorgonocephalus. arcticus KU495901 COI KU495901G Baffin Bay, (Corstorphine, 2011) Canada Gorgonocephalus. eucnemis AB758809 COI AB758809G Japan (Okanishi & Fujita, 2013) Gorgonocephalus. eucnemis HM473911 COI HM473911G Queen Charlotte (Corstorphine, 2011) Is., Canada 1International Barcode of Life (iBOL) 12 4 Figure 3.18. Planktonic stages of Gorgonocephalus eucnemis. Secondary views of the same individual are marked with an apostrophe (’). (A) Embryo, lateral view; (B) early juvenile, oral view; (C) one and (D) two days later. (B’) Closeup view of hyaline layer with bubbled appearance. (E) Another individual with mobile arms tipped with hooks (filled arrowhead), also shown in cross-polarized light. Scale bar is 100 µm, same scale for A-E. 4. CONCLUSIONS We observed planktonic developmental stages of 18 species from seven families of brittle stars from the southern Oregon coast and identified them using DNA barcoding. The planktonic stages we observed included embryos, ophioplutei, reduced plutei, vitellaria larvae, pelagic direct developers, and juveniles; for most of these developmental stages we were able to identify morphological features that can be used to identify them to species. We include a key, species- level descriptions, and photo plates to explain how these species can be recognized. 12 5 5. KEYS TO THE OPHIUROID PLANKTONIC FORMS OF THE NORTHEAST PACIFIC OCEAN The keys below can be used to identify the planktonic ophiuroid forms of the northeast Pacific. The keys are divided into ophioplutei (5.1), nonfeeding forms (5.2), and juveniles (5.3). 5.1. Key to the ophioplutei of the NE Pacific 1a. Ophiopluteus with four to eight arms; larval epidermis transparent, such that skeletal rods are visible through the epidermis under transmitted light; pigmentation is restricted to the arms, and may occur at the distal ends or along the length of the posterolateral arms … planktotrophic ophioplutei, 2 1b. Ophiopluteus with four to six arms; epidermis orange to pink in color and opaque enough to obscure the skeletal rods under transmitted light … reduced plutei, 12 2a. Posterolateral (pl) arms are curved … 3 2b. Posterolateral arms are more or less straight … 8 2a 2b 3a. Recurrent rods present … 4 3b. Recurrent rods absent … 6 rr 3a br 3b 4a. Ventral and dorsal pairs of recurrent rods are slightly offset so that both pairs are visible. Recurrent rods articulate with transverse rods at approximately half their length. Ophiopluteus has a transparent epidermis, sometimes with yellow to green pigmentation at 12 6 the distal tips of the larval arms. The pluteus has a transparent epidermis with yellow-green pigmentation at the distal tips of the arms … young Ophiacantha diplasia 4b. Ventral and dorsal pairs of recurrent rods are of the same length and position so that one pair obscures the other. Recurrent rods connect with the transverse rods near their connection point with the body and end rods. The pluteus has a transparent epidermis that has an inflated appearance, that is, there is an obvious gap between the epidermis and the skeletal rods, especially the posterolateral and anterolateral arms; the stomach may be colored by partially consumed microalgae … Ophiura leptoctenia or O. sarsii* *We caution the reader that we observed each of these species from only one or two larvae. Therefore, we cannot make comparisons between species across ontogeny. I.e., young O. hastatum may be very similar in appearance to the specimens we observed of O. leptoctenia and O. sarsii. 6a. Ophiopluteus has a transparent epidermis with yellow-green pigmentation at the tips of the arms … young Ophiopteris papillosa 6b. Ophiopluteus with a mostly transparent epidermis but may have red, orange, or yellow pigmentation, especially on the posterolateral arms … 7 7a. Arm epidermis is close to the larval skeleton; skeletal rods maybe a red-orange color; anterolateral arms curved and bearing thorns; postoral arms at least twice the length of the body rods; thorns present all sets of arms, but are longest on the proximal side of the posterolateral arms; red pigmentation restricted to the distal tips of the posterolateral arms or absent … Amphiodia urtica 7b. Arm epidermis has an inflated appearance; skeletal rods, especially the posterolateral arms and body rods, may be red-orange in color; anterolateral arms straight to curved in advanced plutei; anterolateral arms without thorns; postoral arms approximately the same length as the body rods; red pigmentation begins at the distal tips of posterolateral arms and usually accumulates across the length of the posterolateral arms as the larva develops … Amphipholis pugetana al pl po 7a br 7b 8a. Ophiopluteus is flat, with a silhouette like an inverted triangle … 9 8b. Ophiopluteus is boxy, with arms of similar heights separated by vibratile lobes. The posterolateral arms are relatively short and are held at a narrow angle, creating a rectangular silhouette like that of some echinoplutei … 11 9a. Recurrent rods absent … 10 12 7 9b. Recurrent rods present. The posterolateral arms of the advanced ophiopluteus are much longer than the other arms and may have bands of reddish-brown pigmentation … Ophiocten hastatum* 10a. Posterolateral arms of the late pluteus are up to five times the length of other arms and are so wide that they are almost horizontal. If present, there is a single median process … Ophiothrix spiculata 10b. Posterolateral arms do not exceed three times the length of the other arms, and do not exceed a width of 90°. If present, there are two median processes … Ophiopholis spp. 11a. Recurrent rods present. Pluteus keeps most of the larval arms when forming a juvenile rudiment … late Ophiacantha diplasia 11b. Recurrent rods absent. The skeletal rods of the anterolateral arms are straight, but the ciliated band bends medially at the distal ends, often touching. The pluteus develops into a vitellaria prior to metamorphosis … late Ophiopteris papillosa 12a. Body rods long, of length equal to or greater than the short straight posterolateral arms; epidermis is orange on the body and white on the posterolateral arms; the junction where the posterolateral arms and body rods meet is at about the height of the mouth … Ophiura luetkenii 12b. Body rods approximately one third the length of the posterolateral arms. Coloration of the epidermis either uniformly tan to salmon or orange with white arms; posterolateral arms with curved spines, especially on their inner side … 13 13a. Epidermis uniformly tan to salmon … 14 13b. Epidermis on the body orange, white on the posterolateral arms … Amphiodia sp. orange belly 14a. Epidermis tan, pink, or salmon in color. Body rods and posterolateral rods are both straight (but may be curved in some larvae). Posterolateral arms are wideset, making an angle that is 90° or wider and form a continuous line with the body rods. Posterolateral and anterolateral arm rods often with thorns … Amphiodia sp. opaque 14b. Epidermis tan in color. Posterolateral arms are held at a narrow angle and bearing only minute thorns … Amphiodia sp. tan 5.2. Key to ophiuroid nonfeeding larvae The nonfeeding larvae include pelagic direct developers and vitellaria larvae. The nonfeeding larvae are usually pink, salmon, or orange in color. 1a. Embryo with pentaradial symmetry … 2 1b. Symmetry is semi-bilateral or unclear … 4 12 8 2a. Embryo or juvenile with five arms … 3 2b. Embryo or juvenile approximately the shape of a flattened disc … 5 3a. Five arms covered with pink-orange epidermis … Gorgonocephalus eucnemis 3b. Five arms with multiple arm segments … Ophiopholis spp. or Ophiothrix spiculata 3c. Five arms with single segments … Amphiuridae, Ophiuridae 4a. Embryo is flattened and round … 5 4b. Embryo is ovoid and wider at one end than the other, with disjunct ciliary bands on the body and shelf-like ones on either end; center of body may have pentaradial symmetry and tube feet … vitellaria, 6 5a. Embryo with five rounded lobes, one of which may be orange to pink in color compared to tan in the others; embryo with a line of skeletal elements spanning an arc across the embryo; mouth centered … Amphiuridae sp. cross piece 5b. Embryo light pink in color, very flat with an uneven periphery; with a ring of skeletal elements around periphery of embryo; stomodeal pit not centered (until pentaradial symmetry is well established) … Amphiodia periercta? 6a. Vitellaria light pink in color with darkened ciliated bands. Ciliated bands five in number … Amphioplus sp. vitellaria 6b. Vitellaria that is colorless except for the ciliated bands which are yellow green … Ophiopteris papillosa (the vitellaria in this species is preceded by a feeding ophiopluteus, 6a) 5.3. Key to the juveniles 1a. Juvenile with arms not obviously segmented, covered in pink epidermis, and lacking tube feet but with a pair of distal claws … Gorgonocephalus eucnemis 1b. Juvenile with segmented arms … 2 2a. Juvenile arms with 2 or more segments … 3 2b. Juvenile arms with 1 segment (terminal arm plate) … 6 3a. Arm segments without hooked spines … 4 3b. A pair of hooked spines on the lateral sides of penultimate arm segment … 5 4a. Yellow pigmentation present at the distal arm segments … Ophiacantha diplasia 4b. Arms white throughout … Ophiopteris papillosa 5a. Penultimate arm segments bearing spines with one hook … Ophiothrix spiculata 5b. Penultimate arm segments bearing spines with two hooks … Ophiopholis spp. 12 9 6a. Terminal arm plates are triangular to cone shaped … Amphiuridae 6b. Terminal arm plates long and rectangular, comprised of three parallel skeletal rods bisected by regular rods that form a honeycomb pattern that echoes that of the disc … 7, Ophiuridae 7a. Larval body is orange … Ophiura luetkenii 7b. Larval body is transparent to tan … Ophiura sarsii BRIDGE In Chapter III, I described various planktonic stages of brittle stars in the northeast Pacific. These results beg for analysis of the evolution of development. In Chapter IV, I used comparative phylogenetic analyses to estimate the ancestral state of the Amphiuridae, a diverse family of brittle stars with planktotrophic ophioplutei, reduced plutei, vitellaria larvae, and pelagic direct developers. We derived character state data from the species accounts in Chapter III, published accounts in the literature, and unpublished data from Panama and Australia (R. Emlet, unpublished). To build a phylogenetic hypothesis we used sequence data derived from our DNA barcoding of brittle star larvae, local collections of adult specimens, and from specimens borrowed from museums to build a 39-spp. phylogenetic hypothesis of Amphiuridae. 13 0 CHAPTER IV EVOLUTION OF LARVAL DEVELOPMENT IN THE BRITTLE STAR FAMILY AMPHIURIDAE This work will include Dr. Richard Emlet as coauthor and contributor to the final manuscript preparation. It is written in the journal style of Invertebrate Biology. 1. INTRODUCTION Developmental patterns are diverse in marine invertebrates and include production of larvae that feed in the plankton, planktonic larvae that do not feed and nonplankton development that may be brooded or develop in benthic capsules. These patterns represent differences in parental investment that result in tradeoffs in allocation to individual offspring and fecundity, the planktonic period and offspring mortality and they have consequences for dispersal. Remarkably, disparate strategies are known to occur amongst closely related species in many taxa: e.g., in sacoglossans (Krug et al., 2015), calyptraeid gastropods (Collin, 2004), barnacles (Ewers‐ Saucedo & Pappalardo, 2019), echinoids (Hart et al., 2011; Hoegh-Guldberg & Emlet, 1997), and ophiuroids (Hart & Podolsky, 2005). A large body of literature has sought to understand the ecological and evolutionary mechanisms that underlie transitions between these strategies (Collin & Moran, 2018; Strathmann et al., 2020). Transitions in developmental pattern from feeding to nonfeeding larvae are associated with increases in egg size, nurse eggs or extracellular materials 131 (Sewell & Young, 1997; Strathmann, 1978b). Comparative phylogenetic analytical methods based on molecules have been increasingly used to examine the evolutionary origin, loss, and potential regain of feeding planktonic larvae (planktotrophy). However, sequence data for marine invertebrate taxa are severely underrepresented in public databases, limiting our ability construct phylogenies to test evolutionary hypotheses even in widespread and abundant groups of macrofauna. In many groups of marine invertebrates, another limitation is lack of information on type of development. Phylogenetic analyses are increasingly used to understand taxonomic organization and evolutionary history of marine invertebrates (e.g. Giribet & Edgecombe, 2019; O’Hara et al., 2017). Comparative phylogenetic analytical methods have been used increasingly to examine the evolutionary origin, loss, and potential regain of feeding planktonic larvae (planktotrophy) (Collin, 2004; Ewers‐Saucedo & Pappalardo, 2019; Hart et al., 1997; Hart & Podolsky, 2005; Krug et al., 2015; Wray, 1996). However, sequence data for many marine invertebrate taxa are severely underrepresented in public databases, limiting our ability to test evolutionary hypotheses even in widespread groups of macrofauna. Brittle stars (Echinodermata: Ophiuroidea, ca. 2100 spp.) are the most diverse echinoderm class and are prominent members of benthic marine environments worldwide (Stöhr et al., 2012). Their widespread distribution across ocean depths and habitats make ophiuroids a good group in which to test evolutionary hypotheses (Bribiesca-Contreras et al., 2017; O’Hara et al., 2019b). They have the potential for analysis of evolutionary patterns in reproduction to the diversity of developmental strategies exhibited by the class. Three broad developmental categories are recognized: brooding, feeding planktonic larvae, and abbreviated development. The latter group includes several larval forms including reduced plutei, vitellaria, and pelagic 132 direct developers (Allen & Podolsky, 2007; Hendler, 1975; O’Hara et al., 2019a; Selvakumaraswamy & Byrne, 2000). Ophiuroids have been found to have numerous instances of reduced plutei (Nakata, Ch. III) which may represent transitions from feeding to nonfeeding; understanding these occurrences in a phylogenetic context may help identify circumstances related to transitions in developmental strategy. While brooding occurs throughout echinoderm groups, it is particularly common in ophiuroids, so they are a good group in which to examine the loss (or gain) of planktonic development. Few analyses of evolutionary patterns have been conducted for ophiuroids due to difficulties in obtaining gametes or identifying larvae to species (but see Hart & Podolsky, 2005; Lessios & Hendler, 2022; O’Hara et al., 2019a; Selvakumaraswamy & Byrne, 2004). DNA barcoding has proven to be an effective method for identifying wild-caught embryos and larvae and can greatly increase estimates of regional species and developmental diversity (Collin et al., 2019, 2020b; Hiebert & Maslakova, 2015; Maslakova et al., 2022) as well as help to test hypotheses about the evolution of development. Here we present comparative phylogenetic analyses of developmental traits in the family Amphiuridae, a large family of brittle stars (ca. 457 spp., O’Hara et al. 2017) that burrows in soft sediments. This family has many developmental patterns, including brooding, planktotrophy via an ophiopluteus, facultative planktotrophy via a reduced pluteus, nonfeeding development via a vitellaria, and pelagic direct development (Emlet, 2006; Hendler, 1975, 1978, 1995; Hendler & Bundrick, 2001; Hendler & Littman, 1986; Mortensen, 1924; Nakata & Emlet, 2023; Stancyk, 1973). We compiled published data on larval development and added previously unpublished data on development from Oregon, Panama, and Australia. We acquired adult materia from museum collections and field collections and sequenced four loci to augment molecular sequence 133 data available in published databases. The family Amphiuridae needs taxonomic revision; a phylogenetic analysis based on exon-capture data found important genera in the family such as Amphiura, Amphioplus, and Amphipholis are polyphyletic in their current composition (O’Hara et al., 2018). Using a 4-marker (18S, 28S, 16S, and COI) and 39-species phylogeny, we asked (i) what was the ancestral developmental strategy for the family, (ii) what is the direction of evolutionary transitions between developmental and larval states (and are they reversible), and (iii) did shifts in developmental mode occur with more frequency in certain clades of the tree? 2. METHODS 2.1. Egg size and developmental patterns To infer developmental character states, we analyzed egg size for 37 spp. of Amphiuridae. We collected egg diameter data from the published literature, species accounts in Chapter III, and for seven additional species, data from collaborators (R. Emlet, unpublished data). The datasets for egg size and phylogenetic analysis differed in species composition but 18 species appear in both datasets (Table 4.1). Eggs were collected from free spawn of adults shortly after collection or were collected from plankton. We photographed eggs in transmitted light, and we measured egg diameter from the photos in Adobe Photoshop. For eggs spawned in the laboratory, a minimum of ten egg diameters were measured and the mean calculated; eggs from the plankton were usually few in number, only one to two individuals. Egg size data was used to infer developmental character states for seven species that were part of the phylogenetic analysis of developmental patterns. 134 1 Table 4.1. Egg size, developmental mode, and larval form for species included in the ancestral state reconstructions. Egg size is given as egg diameter. 2 Development modes are as follows: brooder (Br), planktotrophic ophiopluteus (Pl), and abbreviated (Ab). In species marked with an asterisk (*) development 3 was inferred from egg size. Entries in bold represent new developmental data presented in this study. Larva is ophiopluteus (Pl), reduced pluteus (RP), pelagic 4 direct developer (PD), vitellaria (V), and (-) for brooders. Gene sequences used for phylogenetic analysis are marked by an X. Species Egg Dev. Larva Source BOLDB or Loci (µm) GenBankG 18S 28S 16S COI Ophiopholis aculeata 105 Pl Pl (Strathmann, 1987) OOPH007-18 B X X X (LINNEAUS 1767) Ophiothrix spiculata 110 Pl Pl (Hendler, 1996) OOPH012-18B X X X LE CONTE 1851 Amphiodia periercta? 190 Ab PD (Emlet, 2006) OOPH032-22B X X X X Amphiodia pulchella dark - Pl Pl Emlet, unpublished PAOPA001-23B X Amphiodia pulchella light 85 Pl Pl Emlet, unpublished PAOPA002-23B X Amphiodia sp. opaque 140 Ab RP (Nakata & Emlet, 2023) OLAB004-22B X X X X Amphiodia sp. orange - Ab RP Nakata, Ch III OLAB050-23B X belly Amphiodia sp. tan - Ab RP Nakata, Ch III OLAB051-23B X Amphiodia tabogae 85 Pl* - Emlet, unpublished PAOPA003-23B X X (NIELSEN 1932) Amphiodia urtica 100 Pl Pl Nakata, Ch III, (Schiff & OOPH056-23B X X X (LYMAN 1860) Bergen, 1996) Amphioplus sp. vitellaria - Ab V Nakata, Ch III OLAB058-23B X X X X Amphipholis januarii 150 Ab RP Emlet, unpublished PAOPA004-23B X X LJUNGMAN 1866 Amphipholis kochii 90 Pl Pl (Yamashita, 1985) T. O’Hara, X LÜTKEN 1872 unpublished data 135 Amphipholis misera - Br - (Mortensen, 1933) KU895014G X KOEHLER 1899 Amphipholis pugetana - Pl Pl Nakata, Ch III OOPH004-18B X X X X (LYMAN 1860) Amphipholis squamata 130 Br - (Strathmann, 1987) OOPH035-23B X X X (DELLE CHIAJE 1829) Amphipholis torelli - Br - (Mortensen, 1924) T. O’Hara, X LJUNGMAN 1872 unpublished data Amphiura arcystata - Ab PD Nakata, Ch III OLAB060-23B X X X X Amphiura belgicae - Br - (Mortensen, 1936) OOPH057-23B X X X X KOEHLER 1905 Amphiura borealis - Br - (Stöhr, 2005) T. O’Hara, X (G.O. SARS 1872) unpublished data Amphiura capensis - Br - (Hendler, 1975; T. O’Hara, X LJUNGMAN 1867 MacKinnon et al., 2017) unpublished data Amphiura chiajei - Ab RP (Fenaux, 1963) T. O’Hara, X FORBES 1843 unpublished data Amphiura constricta - Br - (O’Loughlin, 1991) T. O’Hara, X X LYMAN 1879 unpublished data Amphiura deficiens - Br - (Mortensen, 1936) OOPH058-23B X X X X KOEHLER 1922 Amphiura elandiformis - Ab PD Emlet, unpublished OOPH059-23B X X X CLARK 1966 Amphiura eugeniae - Br - (Mortensen, 1936) T. O’Hara, X X LJUNGMAN 1867 unpublished data Amphiura grandisquama - Br - (Mortensen, 1933) T. O’Hara, X LYMAN 1869 unpublished data 136 Amphiura magellanica - Br - (Hendler, 1975) OOPH060-23B X X X LJUNGMAN 1867 Amphiura ptena - Ab PD Emlet, unpublished OLAB109-23B X X CLARK 1938 Amphiura rosea 80 Pl* - (Mortensen, 1924) KU895019G X FARQUHAR 1894 Amphiura spinipes 100 Pl* - (Mortensen, 1924) KU895041G X MORTENSEN 1924 Amphiura stictacantha - Ab PD Emlet, unpublished OLAB110-23B X X H.L. CLARK 1938 Amphiura stimpsonii - Br - (Hendler, 1975) OOPH061-23B X X LÜTKEN 1859 Amphiuridae sp. Au284 - Ab PD Emlet, unpublished OLAB111-23B X X Microphiopholis 90 Pl* - Emlet, unpublished PAOPA005-23B X X geminata LE CONTE 1851 Microphiopholis 87 Pl Pl Emlet unpublished, PAOPA006-23B X X X gracillima (Hendler, 1995) (STIMPSON 1854) Ophiocnida scabriuscula 210 Ab* - (Hendler, 1995) OOPH064-23B X X (LÜTKEN 1859) Ophiodaphne formata - Pl Pl (Tominaga et al., 2004) OOPH062-23B X X KOEHLER 1905 Ophiophragmus 220 Ab* - (Stancyk, 1973) KU895058G X filograneus (LYMAN 1875) Ophiophragmus pulcher 200 Ab* - Emlet, unpublished PAOPA007-23B X H.L. CLARK 1918 Ophiostigma isocanthum 187 Ab RP Emlet, unpublished PAOPA008-23B X X (SAY 1825) 5 137 2.2. Sequence dataset and phylogenetic analyses We built a 39-species database of sequences for amphiurids plus two outgroup taxa by extracting DNA from our own larval and adult collections and specimens from museums. We supplemented this dataset with sequences from public databases. We collected specimens from Oregon, Panama, and Australia from wild plankton, intertidal and subtidal zones. Specimens for an additional 9 species were borrowed from museum collections (Table 4.1). Additional collection information and sequences for our specimens can be found in BOLD (Ratnasingham & Hebert, 2007). We extracted DNA from whole larvae and tube feet and parts of arms of adult specimens. Adults collected by R. Emlet and N. Nakata were relaxed in a 50:50 mix of MgCl2 and filtered sea water (FSW) for morphological identification and photography on a Zeiss dissecting microscope. When possible, we sampled podia for DNA extraction while the specimen was still living; otherwise, podia were collected after preservation in 70-95% ethanol. We extracted larval and adult DNA suing the Chelex-based InstaGene™ Matrix (Bio- Rad). We performed polymerase chain reaction (PCR) to amplify COI, 16S, 28S, and 18S using previously published echinoderm-specific primers (Table 4.2) and thermocycler conditions (Folmer et al., 1994; Geller et al., 2013; Kerr et al., 2005; Kirby & Lindley, 2005; Littlewood, 1994; Okanishi et al., 2011; Okanishi & Fujita, 2013). In particular, we had greater success when using echinoderm- or ophiuroid-specific forward primers (Bribiesca-Contreras et al., 2013; Corstorphine, 2011; Heimeier et al., 2010) that we paired with HCO2198 or jgHCO2198. We used a PCR reaction of 20 µl total volume: 11.4 µl nuclease-free water, 4 µl 5X Green Buffer, 0.4 µl dNTP 10 mM, 0.2 µl GoTaq Polymerase (Promega), 1 µl each of forward and reverse primers, and 2 µl of template DNA. We purified the crude PCR products using the Wizard SV 138 Table 4.2. PCR primers used to amplify four loci for phylogenetic analysis. Locus Primer Sequence (5’-3’) Reference COI LCO1490 GGTCAACAAATCATAAAGATATTGG (Folmer et al., 1994) HCO2198 TAAACTTCAGGGTGACCAAAAAATCA jgLCO1490 TITCIACIAAYCAYAARGAYATTGG (Geller et al., 2013) jgHCO2198 TAIACYTCIGGRTGICCRAARAAYCA COIceF ACTGCCCACGCCCTAGTAATGATATTTTTTA (Hoareau & Boissin, 2010) TGGTNATGCC COIceR TCGTGTGTCTACGTCCATTCCTACTGTRAAC ATRTG EchinoF1 TTTCAACTAATCATAAGGACATTGG (Ward et al., 2008) EchinoR1 CTTCAGGGTGTCCAAAAAATCA Echino COI-F TTTCYACYAAACACAAGGAYATTGG (Heimeier et al., 2010) OphiF ATAATGATAGGAGGATTTGGAAA (Bribiesca-Contreras et al., 2013) LCOechlaF1 TTTTTTCTACTAAACACAAGGATATTGG (Corstorphine, 2011) 16S 16SARL CGCCTGTTTATCAAAAACAT (Palumbi, 1996) 16SBRH CCGGTCTGAACTCAGATCACGT 16Sar GCCTGTTTACCAAAAACAWCG (Kerr et al., 2005; Kirby & Lindley, 2005) 16Sbr GATCCAACATCTAGGTCGC 28S LSU5 taggtcgACCCGCTGAAYTTAAGCA (Littlewood, 1994) LSU3 tagaagctTCCTGAGGGAAACTTCGG LSU001 GCTAAGGAGTGTGTAACAACTCACC (Okanishi et al., 2011) LSU002 GCTTTGTTTTAATTAGACAGTCGGA 18S SSU001 GCTTGTCTTAAAGACTAAGCCATGC (Okanishi et al., 2011) SSU002 CCGTGTTGAGTCAAATTAAGCCGC Gel and PCR Clean up System (Promega) and sequenced in both directions using the same primers used in PCR (Sequetech, Mountain View, CA). We conducted all sequence validation and analysis in Geneious Prime v. 2022.1.1 (https://www.geneious.com). We trimmed and aligned individual reads to create consensus sequences and assigned bases with combined PHRED scores <20 as ‘N’. We received additional sequence data for 8 species from colleagues (T. O’Hara, personal communication) and 139 downloaded four from GenBank (Table 4.1). We aligned sequences one gene at a time using the MAFFT plug-in (Katoh & Standley, 2013) with default parameters, and compared individual gene trees for topological similarity (Figs. S4.1–4). Altogether, we had a 1075 bp alignment of 18S for 12 spp., a 1214 bp alignment of 28S for 12 spp., a 513 bp alignment of 16S for 20 spp., and a 1432 bp alignment of COI for 38 spp. We created a Maximum Likelihood phylogeny using a concatenated dataset of the four loci with PhyML (Guindon et al., 2010). Model of sequence evolution was substitution model HKY85. Bootstrap analysis (100 replicates) was used to calculate clade support. For comparison, we also implemented Bayesian Inference using the HKY85 substitution model, gamma rate variation, with 1,100,000 chain length and 100,000 burn-in length in MrBayes v2.2.4 (Huelsenbeck & Ronquist, 2001). We used sequences for Ophiothrix spiculata and Ophiopholis kennerlyi as outgroup taxa. 2.3. Phylogenetic comparative analyses We performed phylogenetic comparative analyses on a 39-taxon tree inferred by maximum likelihood with fixed topology and branch lengths. We used the ace function in ‘ape’ (Paradis & Schliep, 2019) to estimate discrete ancestral states using maximum likelihood and visualized state probabilities at nodes using standard plotting functions in ‘phytools’ (Revell, 2023). We tested for asymmetry in transition rates between character states using a likelihood (ML) approach for evolution of discrete traits. To do this, we fit a series of models using the fitMk function in phytools, which allows the user to specify transition matrices for hypothesis testing (Lewis, 2001; Revell, 2023). We compared model fit using analysis of variance 140 (ANOVA). We assessed ordered (“irreversible”) and unordered versions of two suites of models: three-state models of development mode (brooding, abbreviated, and planktotrophy) and five- state models of larval form (brooding, pelagic direct, vitellaria, reduced pluteus, and ophiopluteus). We used ordered models to prohibit transitions between brooding and planktotrophy states; we used the unordered models to test if allowing such trait reversals would produce a better fit for our data. We inferred development mode from egg size for nine species. As abbreviated developers have eggs of similar sizes that give rise to a variety of larval forms, we could not infer larval form from egg size alone. For the five-state larval form models we removed eight tips from the tree for which we inferred development (Amphiura spinipes, Amphiua rosea, Amphipholis torelli, Microphiopholig geminata, Amphiodia tabogae, Ophiocnida scabriuscula, Ophiophragmus filograneus, Ophiophragmus pulcher). 3. RESULTS 3.1. Egg size and development patterns Egg size was a strong predictor for developmental pattern: taxa with planktotrophic development have small eggs, 50–110 µm in diameter; taxa with abbreviated development (including reduced plutei and pelagic direct developers) have moderately sized eggs, from 110- 190 µm diameter; and taxa that brood had egg sizes from 120 to 700 µm diameter (Fig. 4.1). 141 Figure 4.1. Histogram of egg diameters by development mode for 37 spp. of Amphiuridae. We inferred planktonic development type for seven species based on egg size to include in our phylogenetic comparative analyses, including four planktotrophs (Amphiura spinipes, Amphiura rosea, Microphiopholis geminata, and Amphiodia tabogae), and three abbreviated developers (Ophiocnida scabriuscula, Ophiophragmus filograneus, and Ophiophragmus pulcher; Table 4.1, Development marked with *). We inferred development pattern for one additional species, Amphipholis torelli based on an early account by Mortensen (1924). 3.2. Sequence dataset and phylogenetic analyses We obtained sequences for four loci (COI, 16S, 28S, 18S) for 39 species of amphiurids, including 74 loci obtained from our specimens, and 13 additional sequences from published and unpublished data (Table 4.1). We constructed phylogenetic hypotheses using maximum likelihood (Fig. 4.2, Table 4.1) and Bayesian methods (Fig. 4.3). We had data for all four loci for seven species (Table 4.1). Individual gene trees contained subsets of the species and displayed similar topologies to the four-gene tree (Figs. S4.1–4). 142 Figure 4.2. Phylogenetic hypothesis for 41 species, including 39 species from the family Amphiuridae and two outgroup taxa, Ophiopholis aculeata and Ophiopthrix spiculata. The tree was made using Maximum Likelihood analysis of a concatenated dataset (COI, 16S rDNA, 18S rDNA, and 28S rDNA). Bootstrap support values >70 % are shown underneath the clades. 143 Figure 4.3. Phylogenetic hypothesis for 41 species, including 39 species from the family Amphiuridae and two outgroup taxa, Ophiopholis aculeata and Ophiopthrix spiculata. The tree was constructed by Bayesian methods using MrBayes. Posterior probabilities are shown underneath their respective clades. Our phylogenetic hypothesis included species from nine of the 26 amphiurid genera, including important, diverse, and polyphyletic genera Amphiura, Amphipholis, and Amphiodia. 144 We inferred Amphiura as basal and paraphyletic with respect to the rest of the clade. Amphipholis and Amphiodia were polyphyletic. We inferred that the color morphs (“dark” and “light”) of Amphiodia pulchella represented two species; they were 19.1% different at the COI locus. Three of our specimens were genetically divergent from other published sequences of the same identification: Microphiopholis gracillima, M. geminata, and Amphipholis januarii. As our developmental data was associated with these adults, we used our own sequence data for those species in the tree. 3.3. Comparative phylogenetic analyses The developmental mode character states (planktotrophy, abbreviated, brooding) were all present in the two major clades within the family. The species in the three-state models included 12 brooders, 16 abbreviated developers (three of which were inferred from egg size), and 11 planktotrophs (four of which were inferred from egg size). Most brooding species were in Amphiura; but there were also three brooders in Amphipholis. Abbreviated developers occurred throughout the tree. Feeding development, or planktotrophy, occurred in Amphipholis, Amphiodia, Microphiopholis, and Ophiodaphne genera, and we inferred it for two species in Amphiura from egg size (Mortensen, 1924). In our five-state analysis of larval form, the 33-spp. tree included 11 brooders (nine of which were Amphiura spp., and two Amphipholis spp.), six pelagic direct developers, one vitellaria, five reduced plutei, and eight planktotrophic plutei. The pelagic direct developers occurred in Amphiura taxa and Amphiodia periercta?. Reduced plutei were present in a clade of Amphiodia spp. from the northeast Pacific, and one in Amphiura chiajei. 145 The ancestral state of Amphiuridae was indeterminate in both three-state and five-state ancestral estimations (Figs. 3.3, 3.4). In the three-state developmental mode model, the best-fit model had equal transition rates between all states. In the five-state larval form model, the best- fit model also had equal transition rate between all states. For both the three-state and five-state models, the best-fit model had equal transition rates between all states (Tables 3.3, 3.4). Fitted values for the three-state model were 2.94 for state changes and -5.87. Similarly, in both cases the worst-fit models were the irreversible models that prohibited secondary gains of planktonic and feeding development. 4. DISCUSSION 4.1. Inference of development pattern from egg size Our egg size data suggests slightly different threshold sizes between developmental patterns compared to analyses of other groups of ophiuroids. Amphiurid species with feeding planktonic development produce smaller eggs than those of other taxa (<110 µm). Species of Macrophiothrix H.L. CLARK, 1938 produce planktotrophic eggs up to 200 µm in diameter, and surveys across Ophiuroidea found planktotrophic eggs up to 170 µm in diameter (Allen & Podolsky, 2007; Hendler, 1991). Two species for which we inferred planktotrophy based on egg size, Amphiura spinipes and A. rosea, have reliable reports for egg size (Mortensen, 1924), but we cannot be sure if those animals are molecular matches for those represented in our tree. Amphiuridae was similar to Macrophiothrix in that egg size does not predict larval form of 146 14 7 Figure 4.4. Estimated ancestral character states for a three-state model of development modes mapped on a phylogeny of 39 spp. from Amphiuridae and two outgroup taxa, Ophioholis aculeata and Ophiothrix spiculata. Nodes with inferred development mode based on egg size are marked with a hollow center. Table 4.3. Summary of the six three-state transition models tested using ML. Model types were equal rates (ER), all rates different (ARD), symmetric (SYM), and irreversible (IRR). Development modes are given as brooding (Br), abbreviated (Ab), and planktotrophy (Pl). The best fit model is in bold text. Type No. Transition matrix Character Description lnL AIC DAIC rates ER 1 Br Ab Pl unordered All transitions -42.76 87.52 0.00 Br - a a occurred at Ab a - a the same rate Pl a a - ARD 6 Br Ab Pl unordered All rates -39.47 90.94 3.42 Br - a b different Ab c - d Pl e f - SYM 3 Br Ab Pl unordered Symmetric -42.52 91.04 3.52 Br - a b rates Ab a - c Pl b c - IRR1 4 Br Ab Pl ordered Abbreviated is -42.10 92.19 4.67 Br - a 0 intermediate Ab b - c Pl 0 d - IRR2 4 Br Ab Pl ordered No secondary -46.47 100.93 13.41 Br - 0 0 gain of Ab a - b planktonic Pl c d - development IRR3 3 Br Ab Pl ordered No secondary -53.67 113.34 25.82 Br - 0 0 gains of Ab a - 0 feeding Pl b c - development 14 8 Figure 4.5. Estimated ancestral character states for a five-state model of larval form mapped on a phylogenetic hypothesis for 31 spp. of Amphiuridae. Table 4.4. Summary of the six five-state transition models tested using ML. Model types were equal rates (ER), all rates different (ARD), symmetric (SYM), and irreversible (IRR). Larval forms are given as brooding (Br), pelagic direct developer (D), vitellaria (V), reduced pluteus (RP), and ophiopluteus (Pl). The best fit model is in bold text. 14 9 Type No. Transition matrix Description lnL AIC DAIC rates ER 1 Br D V RP Pl All -46.68 95.73 0.00 Br - a a a a transitions D a - a a a occurred at V a a - a a the same RP a a a - a rate Pl a a a a - ARD 20 Br D V RP Pl All rates -37.54 115.08 19.35 Br - a b c d different D e - f g h V i j - k l RP m n o - p Pl q r s t - SYM 10 Br D V RP Pl Symmetric -41.71 103.41 7.68 Br - a b c d rates D a - e f g V b e - h i RP c f h - j Pl d g i j - IRR1 18 Br D V RP Pl Abbreviated -45.46 126.92 31.19 Br - a b c 0 is D d - 0 0 0 intermediate V h i - j k RP l m n - o Pl 0 p q r - IRR2 16 Br D V RP Pl No -39.49 110.99 15.26 Br - 0 0 0 0 secondary D a - 0 0 0 gain of V e f - 0 0 planktonic RP 0 j k - l development Pl m n o p - IRR3 13 Br D V RP Pl No -44.91 117.83 22.10 Br - 0 0 0 0 secondary D a - 0 0 0 gains of V d e - 0 0 feeding RP g 0 i - 0 development Pl j 0 0 m - abbreviated developers (Allen & Podolsky, 2007). Egg sizes for brooders are similar to other analyses (Hendler, 1991). 15 0 4.2. Phylogenetic analyses While this work presents the largest comparative phylogenetic analysis for the Amphiuridae to date, it encompasses only a small fraction of the family diversity. Our phylogenetic hypothesis represents less than 9% of the described species of Amphiuridae (O’Hara et al., 2017), as we included only species with both molecular and developmental data. Future work should focus on collecting molecular and developmental data for all species in the family. For example, although there was an 18-spp. overlap between our phylogenetic and egg size datasets, there were an additional 19 species for which developmental pattern can be inferred from egg size, but currently lack molecular data. DNA barcoding of larvae from plankton is an effective method for simultaneously capturing developmental pattern and species diversity (Collin et al., 2020a, 2021; Maslakova et al., 2022; Nakata, Ch III). Additional molecular data is also necessary to resolve taxonomic relationships. The family Amphiuridae requires revision, and contains many polyphyletic taxa (O’Hara et al., 2018). Our results show several genera – including Amphiura, Amphipholis, and Amphiodia – are polyphyletic, in agreement with a transcriptome-based analysis of 69 species (O’Hara et al., 2017, 2018). A consequence of this analysis was the discovery of divergent sequences of specimens given the same identification based on morphology. We found two lineages of Amphiodia pulchella from Panama that may be distinguishable based on color. Specimens collected in Panama for Microphiopholis gracillima, M. geminata, and Amphipholis januarii were over 5% different from sequences with the same species identification in GenBank. 15 1 4.3. Ancestral state estimations We were not able to infer the developmental pattern for the ancestor of Amphiuridae in either analysis of ancestral state estimations. This was influenced by the basal placement of Amphiura, a genus with over half the species brooded development. We inferred Amphiura to be paraphyletic to the rest of the clade. Two species, A. borealis and A. chiajei were inferred to be basal. The remaining 15 Amphiura spp. formed a clade that was sister to a clade formed by the other genera. The addition of more taxa may cause tree topology to change and move Amphiura from its basal position. In a 1484-exon dataset for 576 ophiuroid species, the two main Amphiura clades had a derived position, but two additional Amphiura species were inferred to be in the most basal clade of the family with high support (O’Hara et al, 2017, Fig. S3). This phylogenetic hypothesis included only 14 species with known development, and the three developmental patterns were distributed across the tree, suggesting multiple gains and losses of feeding development similar to our data (see below). 4.4. Repeated gains and losses of planktonic and feeding development The family Amphiuridae is part of the suborder Gnathophiurina MATSUMOTO, 1915, which also includes families Ophiotrichidae, Ophiopholidae, Ophiactidae, and Ophiothamnidae (O’Hara et al., 2017, 2018). These taxa have mostly feeding development, with nonfeeding development inferred to be derived in the Ophiotrichidae (Allen & Podolsky, 2007; Hendler, 2005, 1995; Kitazawa et al., 2015; Mortensen, 1938; Nakata, Ch III; Pearse, 1994; Selvakumaraswamy & Byrne, 2000). This suggests a planktotrophic ancestor for Amphiuridae, as does the presence of feeding larvae in the family. Similarities of feeding larval forms across the classes of echinoderms, and essential body plan similarities among all feeding ophioplutuei 15 2 make it unlikely that feeding larvae evolved multiple times within echinoderm or with in the ophiuroids or that the ancestor of Amphiuridae was a brooder (Strathmann, 1978b; Strathmann et al., 2020). We inferred frequent transitions in development mode in this subsample of Amphiuridae. If the ancestor was a planktotroph, in the 39-spp. amphiurid tree (Fig. 4.3), the distribution of developmental patterns we observed would require an initial loss of feeding development followed by, four secondary gains (Amphiura spinipes, Amphiura rosea, the ancestor to the Amphipholis and Amphiodia clade, and Amphiodia urtica), and five other secondary losses of feeding. If a feeding ancestor is present in all branches before it disappears completely then there are over 15 losses of feeding. Evolutionary transitions between development modes are widespread across marine invertebrates, but they tend to be unidirectional. Transitions from nonfeeding to feeding planktonic larvae are rare (Collin, 2004; Emlet, 1995; Krug et al., 2015; Wray, 1996), probably because complex feeding structures may be difficult to re-evolve once lost (Strathmann, 1985; Strathmann et al., 2020). When feeding evolves de novo the morphology is likely to be different, e.g., feeding larvae of different phyla don’t look or function the same. However, in our estimations of ancestral character states there are many inferred gains and reversals in developmental mode, which were supported by the best-fit models (Table 3.3, 3.4). The models in which transitions from brooding to planktonic development were prohibited were consistently the worst fits for our data. Our data suggest several secondary gains of both nonfeeding and feeding larvae in lineages with brooding. Gains of nonfeeding larvae from brooding lineages have been inferred in barnacles (Ewers‐Saucedo & Pappalardo, 2019), but secondary gains of feeding larvae are 15 3 exceedingly rare (Collin, 2004). We suspect that the gains of feeding larvae from brooding lineages inferred in this analysis may be an artifact of our limited taxonomic representation and large amount of missing data. The transition from brooding to pelagic nonfeeding larvae appears more likely. The abbreviated larvae in Amphiura are pelagic direct developers (except A. chiajei), rather than reduced plutei or vitellaria. It is possible that these species accomplished the transition from brooding to planktonic development by release of eggs that develop into juveniles while in the plankton rather than re-evolving a ciliated larval form. In contrast, species with abbreviated development that come from inferred planktotrophic ancestors (Amphipholis, Ophiophragmus, and Amphiodia) are a mix of reduced plutei (n=4 spp.), pelagic direct developers (n=2), and vitellaria (n=1). This suggests the reduced pluteus, possibly facultatively feeding, as the intermediate larval form in the evolution of nonfeeding larvae. Future work should investigate the occurrence of reduced plutei in other ophiuroid taxa. Prior to this analysis, only one reduced pluteus was known from Amphiuridae, Amphiura chiajei (Fenaux, 1963); we found four new reduced plutei from the Oregon plankton using DNA barcoding, demonstrating that this larval form is more prevalent in ophiuroids than previously known. This work shows evidence for repeated gains and losses of feeding development in a family of brittle stars or if feeding larvae only evolved one time, then there are over 10 losses of feeding among the 39 taxa examined here. However, inferring developmental transitions from such an incomplete dataset can be misleading. We include only 9% of species from the family. Future work should focus on the collection of molecular data across Amphiuridae to resolve taxonomic relationships within the family. There is also much to be done to characterize development patterns in the family. We show how DNA barcoding is a powerful tool for 15 4 identifying larvae from plankton, but this type of analysis can be expensive, time consuming, and limited to particular regions where there is regular access to wild plankton. 15 5 CHAPTER V CONCLUSION This work highlights the potential of brittle stars as a system to study transitions in developmental patterns in marine invertebrates. Despite their diversity of developmental patterns and evidence of repeated evolution of nonfeeding development, most groups of ophiuroids have not been analyzed for evolution of development. One reason for this is the lack of observations on development pattern and larval form across ophiuroid taxa, which is known for less than 15% of species (N. Nakata, unpublished data). A primary cause is that unlike for echinoids and asteroids, ophiuroids have no reliable means for obtaining mature gametes in the laboratory. In the research presented here, we relied instead on observations of larvae collected from plankton identified with DNA barcoding. One type of larva, which at present could not be identified, was shown to be a facultative planktotroph, and used to test the effects of larval feeding on developmental timing, juvenile size, and survival. Amphiodia sp. opaque has a reduced pluteus that can make a juvenile without food but benefits from larval feeding. On average, individuals that fed as larvae had shorter development times, higher percent metamorphosis, larger juveniles, and greater capacity to avoid juvenile starvation by relying on energetic reserves accumulated during the larval period (Nakata & Emlet, 2023). Facultatively planktotrophy has only been observed eight times across marine invertebrates, including the brittle star Macrophiothrix rhabdota, which showed similar responses to food (Allen & Pernet, 2007; Allen & Podolsky, 2007). In addition to Amphiodia sp. opaque, we found seventeen other ophiuroid species in the coastal plankton. In Chapter III we summarized over a decade’s worth of observations to describe the taxonomic, developmental, and morphological diversity of brittle star larvae. Using DNA barcoding of larvae and adults, we identified the developmental patterns of 14 described species. Four of these are currently only known in their larval form: Amphiodia sp. opaque, A. sp. orange belly, A. sp. tan, and Amphioplus sp. vitellaria. These larvae may not represent new species as many brittle stars species are missing from molecular databases for comparison, highlighting a limitation of barcoding for species identification. Still, we were able to triple the number of larval descriptions from the most recent summary of ophiuroid development in the northeast Pacific (Strathmann, 1987). By barcoding multiple specimens of each species, we were able to connect multiple life stages – embryo, larva, rudiment-stage, and juvenile – without needing to spawn animals in the lab and culture larvae over weeks or months. After identifying individual larvae by barcoding, we used our observations and photos to describe morphological features that can be used to identify larvae found in plankton. In planktotrophic ophioplutei, we identified features of the larval skeleton that were characteristic of families and species. Abbreviated developers varied in color and shape. One of the primary findings of Chapter III was that abbreviated development is common amongst the brittle stars of the northeast Pacific, present in nine of the eighteen species we observed. We observed four reduced plutei, two vitellaria, and three pelagic direct developers. The vitellaria were the first to be described for their respective families: Amphioplus sp. vitellaria, Amphiuridae, and Ophiopteris papillosa, Ophiopteridae. In the case of O. papillosa, we were also able to use barcoding to connect divergent larval forms from the same species: Ophiopteris papillosa has a feeding ophiopluteus that develops into a vitellaria prior to metamorphosis, only the second ophiuroid recorded to do so to date (Cisternas & Byrne, 2005). 15 7 DNA barcoding is a powerful tool for identifying larval stages of benthic invertebrates, and it can also be an important source of phylogenetic data for understudied taxa. Sampling across life stages increases estimates of biodiversity (Maslakova et al., 2022), and many brittle star species are absent from molecular databases. Molecular data increasingly inform and determine our understanding of taxonomic relationships. Molecular phylogenies have led to recent reorganization of major clades within the Ophiuroidea (O’Hara et al., 2017, 2018). In addition to the ophiuroids of southern Oregon we described in Chapter III, R. Emlet used similar methods to collect molecular and developmental data for ophiuroids of Panama and Australia. By utilizing this dataset with data from the published literature, we built a 39-species phylogenetic hypothesis for the family Amphiuridae in Chapter IV and used it to test hypotheses of the evolution of development. We inferred frequent transitions between developmental types and larval forms, even from nonfeeding (abbreviated) to feeding development. In Amphiodia abbreviated development mostly took the form of reduced plutei, but a pelagic direct developer is also known (Emlet, 2006). In Amphiura abbreviated development was restricted to pelagic direct developers, but one reduced pluteus is known. 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Comparison Z P P adjusted Food 2020 – food 2021 2.97 0.003 0.015 * Food 2020 – no food 2020 5.35 < 0.001 < 0.001 * Food 2021 – no food 2020 1.75 0.080 0.160 Food 2020 – no food 2021 2.12 0.034 0.102 Food 2021 – no food 2021 -0.76 0.444 0.444 No food 2020 – no food 2021 -2.60 0.009 0.038 * 17 6 Table S2.2. Pairwise comparisons from Dunn’s test for juvenile aboral surface area according to treatment and year: 2020 (20), 2021 (21). Comparison Z P.unadj P.adj Food 20 – food 21 -0.558 0.577 1.000 Food 20 – no food 20 3.641 0.000 0.002 * Food 21 – no food 20 3.814 0.000 0.001 * Food 21 – no food 21 2.765 0.006 0.028 * Food 21 – no food 21 3.006 0.003 0.019 * No food 20 – no food 21 -1.071 0.284 1.000 Food 20 – wild 19 2.766 0.006 0.034 * Food 21 – wild 19 3.007 0.003 0.021 * No food 20 – wild 19 -0.520 0.603 1.000 No food 21 – wild 19 0.443 0.657 0.657 17 7 Table S2.3. Akaike’s (AIC) and Bayesian Information Criteria (BIC) values for generalized linear models of juvenile aboral surface area (juv.size) in response to planktonic duration (p.d.). Models with the lowest AIC or BIC values are bolded. Model df AIC DAIC BIC DBIC Juv. size ~ p.d. 3 -244.4 43.9 587.2 37.2 Juv. size ~ p.d. + treatment 4 -288.3 - 550.0 - Juv. size ~ p.d. + year 4 -286.3 2 570.7 20.7 Juv. size ~ p.d. + treatment + year 5 -268.4 19.9 553.3 3.3 17 8 Table S2.4. Akaike’s (AIC) and Bayesian Information Criteria (BIC) value for generalized linear models of juvenile starvation time (days) based on juvenile aboral surface area. AIC weights are w. Models with the lowest AIC or BIC values are bolded. Model df AIC DAIC BIC DBIC Starve time ~ juv. size 3 2120.9 459.2 2125.4 453.2 Starve time ~ juv. size + treatment 5 1978.7 317.0 1986.2 314.0 Starve time ~ juv. size + year 5 1703.7 42.1 1711.2 39.1 Starve time ~ juv. size + treatment + year 6 1686.1 24.5 1695.1 23.0 Starve time ~ juv. size + treatment * year 7 1661.7 - 1672.1 - 17 9 Figure S4.1. Maximum likelihood tree for 18S of Amphiuridae. Figure S4.2. Maximum likelihood tree for 28S. 18 1 Figure S4.3. Maximum likelihood tree for 16S. 18 2 Figure S4.4. Maximum likelihood tree for 38 amphiurid spp., COI locus. 18 3