Understanding the Morphology and Distribution of Nematocysts in Sea Anemones and Their Relatives

Home , Acanthastrea, Actinia fragacea, Actinoscyphiidae, Actinostola, Actinostolidae, Actinothoe

Understanding the morphology and distribution of nematocysts in sea anemones and their relatives

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Abigail Julia Reft

Graduate Program in Evolution, Ecology and Organismal Biology

The Ohio State University

2012

Dissertation Committee:

Dr. Marymegan Daly, Advisor

Dr. John V. Freudenstein

Dr. William Ausich

Abigail Julia Reft

2012

Abstract

Cnidaria includes organisms diverse in body form, life-cycle, and ecology and includes

corals, sea anemones, Hydra, and jellyfish. Despite this diversity, cnidarians are easily

recognized by the presence of small intracellular stinging capsules called nematocysts.

This structure, which consists of a tubule attached at one end that typically bears spines, are a synapomorphy for the group as all members of the phylum produce them. These structures are used in many aspects of everyday biology including defense against predators, attachment to substrate, capture of prey, and aggression against other cnidarians. Although the basic construct of the nematocyst is simple, high amounts of morphological variation in tubule and spine features are found throughout the phylum.

This variation has been difficult to interperate for several reasons including the need for advanced microscopy techniques to visualize the morphology, disagreements among authors as to how to best circumscribe and organize the variation that is observed, and the lack of many broad, phylogenetically based analyzes to put this diversity in an evolutionary context. Because the interpretation of nematocyst diversity is so problematic, the utility of nematocyst as phylogenetic characters for cnidarians is unclear.

To determine if nematocysts can be used phylogenetically requires a better understanding of the morphological variation itself and its distribution within cnidarians.

To address these issues, I performed a morphological survey of nematocysts using advanced microscopy techniques (differential inference contrast, scanning and ii

transmission electron microscopy). This work included assessing the distribution and

morphology of a single nematocyst character, the apical structure, to determine if it

contained any phylogenetic signal, fully documenting the diversity of a group of

nematocyst morphologies that have been particularly confusing (the rhabdoids or

nematocysts with a wider diameter basal tubule), placing this diversity on phylogenetic

trees to look for patterns of evolution, and utilizing a multivariate approach to assess the

morphological groups implied by qualitative characters.

The apical structure study found that an individual morphological character does provide strong phylogenetic information. Three morphological states are present and they each represent a monophyletic group (Medusozoa, Anthozoa, and the anthozoan order Actiniaria). For the Actiniaria, this provides a morphological synapomorphy that this group has previously lacked.

In fully describing the variation of rhabdoid nematocyst forms, I recognize nine distinct morphologies. Some of these morphologies (particularly some of the p-rhabdoid forms) agree closely with previously hypothesized morphological groupings. However, other morphologies are recognized as distinct for the first time. New names are given to these nematocyst morphologies to help clarify the exact distribution of variation.

Placing this variation in a phylogenetic context reveals that the presence/absence of certain morphologies does define monophyletic groups. Clarifying the confusion over which cnidarians have each morphology results in a clearer pattern morphotypes shared amongst certain taxa (i.e. the p-rhabdoid B forms in the Metridiodea).

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Finally, using quantitative characters in a multivariate analysis reveals support for many groups of nematocyst morphology. Using qualitative features to group the variation is effective in finding groups of morphologies that share shape features as quantified by a multivariate analysis.

Dedication

Dedicated to my parents, Joyce A. and Chester S., for all their love and support, and Prof.

Michael LaBarbera for introducing me to the wonderful world of invertebrates.

Acknowledgments

A project such as this one is impossible to complete without the help and support of many people both professionally and personally. First and foremost, I would like to thank my advisor, Meg Daly for taking me under her wing while I was working on my masters and supporting me through the completion of my Ph.D. I know of no other scientist could nurture my love of all things invertebrate and Asian like she has; I could not have asked for a better academic mother. I also want to thank the other members of my committee,

John Freudenstein and Bill Ausich. In and out of class, I have learned much about both systematics and being a scientist from both and have always enjoyed learning about the plant and paleontological perspectives on the field.

Also, I thank those that assisted me in fieldwork and hosted me while I was in foreign countries. Specifically, I thank Dr. Shin Kubota of the Seto Marine lab of Kyoto

University for his assistance (and for the onsen) during my time in Japan, Dr. Jun-Im

Song and her lab at Ewha Womens University for their assistance during my time in

South Korea (and for the lovely trip to Jeju), and Dr. Bernard Picton of the National

Museums of Northern Ireland for his assistance (and for the warm hospitality of his family) in Northern Ireland. Others to thank in collecting material include Anthony

Montgomery, Estefanía Rodríguez, Neil Blackstone, and Paulyn Cartright; thank you for providing some of the raw material needed to complete my project.

The Daly lab has been my home for the last six years and I thank all members both past and present for their help and friendship. Specifically, thanks to Annie

Lindgren for always providing perspective, Luciana Gusmaõ for the many long discussions about anything and everything, Estefanía Rodríguez for endless help and support, Paul Larson for fun atmosphere in lab, Nick Skomrock for the fun and gossip, and Jason Macrander for the events and parties.

Finally, I would like to thank my family for being so supportive even if they do not completely understand what I am doing here. My parents in particular have always encouraged me to grow and explore the world around me and never discouraged me from trying new (and sometimes hard things). Thanks so much for all the love and support.

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Vita

October 1980...... Born: Chicago, IL, USA

2002...... B.A. Biology, University of Chicago

2005...... M.A. Ecology and Evolutionary Biology,

University of Kansas

2006 to present ...... Graduate Teaching Associate, Department

of Evolution, Ecology and Organismal

Biology, The Ohio State University

Publications

Voight JA, Lee RW, Reft AJ, Bates AE. 2012. Scientific gear as a vector for alien species at deep-sea hydrothermal vents. Conserv Biol DOI: 10.1111/j.1523-1739.2012.01864.x

Reft AJ, Daly M. 2012. Morphology, distribution, and evolution of apical structure of nematocysts in Hexacorallians. J Morph. 273: 121-136.

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Daly M, Gusmao LC, Reft AJ, Rodriguez E. 2010. Phylogenetic Signal in Mitochondrial and Nuclear Markers in Sea Anemones (Cnidaria, Actiniaria). Integr Comp Biol 50: 371-

388.

Reft, AJ, Westfall, JA, Fautin DG. 2009. Formation of the apical flaps in nematocysts of sea anemones (Cnidaria: Actiniaria). Biol Bull 217: 25-34.

Reft AJ, Voight J. 2009. Sensory structures on the siphons of wood-boring bivalves

(Pholadidae: Xylophagainae: Xylophaga). Nautilus 123:43-48.

Sierwald P, Reft AJ. 2004. The Millipede Collections of the World. Fieldiana, Zoology

New Series 103, Publication number 1532, 1-100.

Fields of Study

Major Field: Evolution, Ecology, and Organismal Biology

Table of Contents

Abstract...... ii

Dedication...... v

Acknowledgments...... vi

Vita...... viii

List of Tables ...... xv

List of Figures...... xvii

Chapter 1: Introduction...... 144

Overview of the chapters...... 3

References...... 6

Chapter 2: Morphology, distribution, and evolution of apical structure of nematocysts in

Hexacorallia...... 8

Introduction...... 8

Materials and Methods...... 11

Materials...... 11

Methods...... 12

Results...... 13

SEM: Hydrozoa...... 13

SEM: Actiniaria...... 14

SEM: other anthozoans...... 15

TEM: Medusozoa...... 16

TEM: Anthozoa...... 17

Discussion...... 20

The three forms of nematocyst apical structure...... 20

Functional differences in apical structure...... 23

Distribution of apical structure and its significance...... 26

Implications for evolution of cnidae and their structures...... 29

References...... 33

Chapter 3: Gross morphology and ultrastucture of anthozoan nematocysts...... 52

Introduction...... 52

Historical perspective on the classification of mastigophores...... 53

Materials and Methods...... 56

Materials...... 56

Methods...... 57

Results...... 59

Nematocyst morphologies observed...... 60

Morphology 1: Acanthoneme...... 60

Morphology 2: Colloponeme...... 63

Morphology 3: Hadroneme...... 65

Morphology 4: p-rhabdoid A...... 67

Morphology 5: p-rhabdoid B1a...... 69

Morphology 6: p-rhabdoid B2d ...... 71

Morphology 7: p-rhabdoid B2s (variant a and b)...... 73

Morphology 8: Diakaneme ...... 76

Morphology 9: Aphylloneme...... 78

Basal tubule of hydrozoan heteroneme nematocysts...... 79

Discussion...... 80

Historical perspective on observed nematocyst morphologies...... 80

p-mastigophores, amastigophores, and the subdivision of these types...... 92

Basitrichs and b-mastigophores...... 94

Light microscopy of undischarged nematocysts...... 96

Relationships among morphologies...... 98

Nature of the shaft...... 103

References...... 107

Chapter 4: Distribution of nematocyst morphology and implications for phylogeny and evolution...... 144

Introduction...... 144

Materials and Methods...... 147

Materials...... 147

Methods...... 148

Sample preparation...... 148

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Phylogeny...... 150

Results...... 152

Variation within nematocyst morphotype...... 152

Distribution of types...... 157

Phylogenetic tree...... 157

Mapping nematocyst characters...... 158

Discussion...... 160

Trees...... 160

Anthozoan cnidomes...... 162

Actiniarian cnidomes...... 165

Taxon sampling...... 169

Evolution of morphologies...... 170

References...... 175

Chapter 5: Multivariate analysis of nematocyst variation visible in light and electron microscopy...... 209

Introduction...... 209

Materials and methods...... 212

Materials...... 212

Methods...... 213

Sample preparation...... 213

Morphological measurements...... 214

Multivariate analysis...... 216

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Results...... 217

Normality...... 217

PCA and NMS of SEM dataset of all morphologies...... 218

PCA and NMS of SEM dataset of annulated basal tubule

morphologies...... 220

PCA and NMS of light microscopy dataset...... 220

Discriminate function analysis of all three datasets...... 221

Discussion...... 222

Normality of nematocyst data...... 222

Support for morphological groups...... 224

Acanthonemes, Aphyllonemes, and Colloponemes...... 224

Diakanemes...... 226

Hadronemes...... 227

p-rhabdoids A...... 227

p-rhabdoids B1a...... 228

p-rhabdoids B2d and B2s...... 229

Further directions in multivariate analysis of nematocyst variation...... 229

References...... 231

Bibliography...... 254

Appendix A: Raw data for multivariate analysis...... 269

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List of Tables

Table 2.1 Summary of taxa and collection locality used in apical structure study...... 41

Table 2.2 Summary of nematocyst SEM apical structure data...... 43

Table 2.3 Summary of nematocyst TEM apical structure data...... 44

Table 3.1 Summary of taxa and collection locality and body part sampled for SEM and

TEM for nematocyst monograph...... 113

Table 3.2 Morphologies described in monograph, their distribution in cnidarian body

tissues, and their categorization under previous classifications...... 116

Table 3.3 Light microscope details that differentiate the nine morphologies observed in monograph study...... 117

Table 4.1 Summary of taxa and collection locality and body part sampled for SEM for distribution study...... 181

Table 4.2 Summary of taxa and genes sampled to create matrix for phylogenetic

analysis...... 184

Table 4.3 Distribution of nematocyst morphologies by actiniarian family...... 189 xv

Table 4.4 Summary of nematocyst distribution by species and tissue type...... 190

Table 5.1 Summary of taxa and collection locality used in the multivariate study...... 235

Table 5.2 List of variables used in all three multivariate datasets...... 236

Table 5.3 List of all tissues and taxa from which the multivariate datasets were derived with numbers of individual nematocysts studied...... 237

Table 5.4 Results from principle components analysis...... 239

Table 5.5 Results from discriminant function analysis of the all morphology dataset using cross validation...... 240

Table 5.6 Results from discriminant function analysis of the annulated basal tubule dataset using cross validation...... 241

Table 5.7 Results from discriminant function analysis of the light microscopy data using cross validation...... 242

Table A.1 Raw data for multivariate analysis of the all morphology dataset...... 269

Table A.2 Raw data of extra variables for annulated dataset...... 274

Table A.3 Raw data for multivariate analysis of the light microscopy dataset...... 276

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List of Figures

Figure 2.1. Basic nematocyst morphologies identified with differential interference contrast light microscopy...... 46

Figure. 2.2 Scanning electron micrographs of apical structure in cnidae from across the phylum Cnidaria...... 48

Figure. 2.3 Transmission electron micrographs of apical structure in cnidae across

Cnidaria...... 50

Figure. 2.4 Phylogenetic interpretation of apical structures...... 51

Figure 3.1. Flow chart of the Weill (1934) classification system for nematocysts...... 118

Figure 3.2. Nematocyst types. Light micrographs of nematocysts as traditionally classified using the modified Weill system...... 120

Figure 3.3. Transmission electron microscope images of some of the morphologies observed in monograph study...... 122

Figure 3.4. Acanthoneme...... 124

Figure 3.5. Colloponeme...... 126

Figure 3.6. Hadronemes...... 128

Figure 3.7. p-rhabdoid A...... 130

Figure 3.8. p-rhabdoid B1a...... 132

Figure 3.9. p-rhabdoid B2d...... 134 xvii

Figure 3.10. p-rhabdoid B2s (variants a and b)...... 137

Figure 3.11. Diakaneme...... 139

Figure 3.12. Aphylloneme...... 141

Figure 3.13. Morphology and ultrastructure of hydrozoan (medusozoan) rhopaloid nematocysts (those with basal tubule that changes in diameter)...... 143

Figure 4.1. Nematocyst morphotypes documented in distribution study...... 196

Figure 4.2. Variation among the b-rhabdoid types...... 198

Figure 4.3. Variation among the p-, amastigophore types...... 200

Figure 4.4. Holotrichous isorhiza (holotrich) variation...... 202

Figure 4.5. Highest likelihood tree...... 203

Figure 4.6. Cnidome distribution across Hexacorallia...... 204

Figure 4.7. Phylogenetic perspective on the cnidome of mesenterial filaments of actiniarians...... 205

Figure 4.8. Phylogenetic perspective on the cnidome of tentacles of actiniarians...... 206

Figure 4.9. Phylogenetic perspective on the cnidome of the body column of actiniarians...... 207

Figure 4.10 Phylogenetic perspective on the cnidome of acontia and acrorhagi of actiniarians...... 208

Figure. 5.1. Morphometic measurements taken for light and scanning electron microscope datasets...... 244

Figure 5.2. Plot of principle component 1 vs. 2 for the dataset that included all morphologies...... 245

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Figure 5.3. Three-dimensional plot of principle components analysis for the dataset that included all morphologies...... 246

Figure 5.4. NMS plot using Euclidian distance and three dimensions of the dataset the included all morphologies...... 247

Figure 5.5. Plot of principle component 1 vs. 2 for the annulated tubule dataset...... 248

Figure 5.6. Three-dimensional plot of principle components analysis for the annulated basal tubule dataset...... 249

Figure 5.7. NMS plot using Euclidian distance and three dimensions of the annulated basal tubule dataset...... 250

Figure 5.8. Plot of principle component 1 vs. 2 for light microscope data set...... 251

Figure 5.9. Three-dimensional plot of principle components analysis for the light microscope dataset...... 252

Figure 5.10. NMS plot using Euclidean distance and three dimensions of the light microscope data set...... 253

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Chapter 1: Introduction

The phylum Cnidaria is diverse in life-cycle, body-form and ecology. However,

the group is united by a strong synapomorphy: all cnidarians produce cnidae. All

members of the phylum produce these complex secretory products that are packaged with

a tubule inverted into the capsule. Under appropriate stimulus, the tubule everts,

discharging the contents of the capsule. Nematocysts, one of three kinds of capsules

collectively called cnidae, are produced by all members of the phylum. In contrast, spirocysts and ptychocysts are more restricted in their distribution (Mariscal 1974, 1984,

Fautin and Mariscal 1991). Nematocysts serve a diversity of functions, including defense

against predators, capture of prey, intracnidarian aggression, and larval attachment (e.g.

Mariscal 1974, 1984, Bigger 1988). Spirocysts are only known from members of the

anthozoan subclass Hexacorallia (Schmidt 1974, Mariscal 1974, 1984), and are

hypothesized to have an adhesive function (Schmidt 1974, Mariscal 1974, Mariscal et al.

1977a). Ptychocysts are even more restricted in their distribution, being known only

from members of the hexacorallian order Ceriantharia (tube anemones); the everted

tubules of ptchocysts trap sediment and thus build the tube in which the animal lives

(Mariscal et al. 1977b; Mariscal 1984).

Because they are so important to the biology of cnidarians and only this group is

known to produce them, nematocysts can be considered a key innovation of the phylum.

Key innovations in morphology or physiology spur organismal diversification by opening

up new ecological niches, enabling new interactions between organisms and their

environment, and allowing new modes of feeding, defense, or reproduction.

Understanding the evolution of these key innovations is thus critical to understanding the

diversification of a group, as the key innovations are integral to both the pattern of

diversification and the process that drives it. Therefore, understanding the morphology

and evolution of nematocysts will provide insight into the evolution of the phylum itself.

Nematocysts show greater morphological diversity than spirocysts or ptychocysts

(e.g. Weill 1930, 1934, Carlgren 1940, 1945, Mariscal 1974, 1984, Fautin and Mariscal,

1991, Rifkin 1991). Unsurprisingly given the wide variety of uses, nematocysts vary greatly in the morphology of the tubule, in the shape, size and arrangement of spines, and in ultrastructural features of these elements (Weill 1930, 1934, Carlgren 1940, 1945,

Mariscal 1974, Fautin and Mariscal 1991). In fact, the variation is so extensive that authors have disagreed on how to classify and understand it (Weill 1934, Carlgren 1940,

Cutress 1955, Schmidt 1969, Östman 2000, see England 1991 for a discussion of this issue). This confusion has confounded interpretation of any patterns of evolution and made evaluating the phylogenetic use of nematocyst characters difficult.

This study seeks to address this confusion by using modern microscopy methods

(i.e. scanning and transmission electron microscopy and differential interference contrast,

DIC, light microscopy) to clearly document the morphological diversity, analyze it in a phylogenetic context, quantify it using multivariate techniques, and evaluate the historical interpretations of diversity.

Overview of the chapters

To gain a better understanding of the diversity and evolution of nematocysts in sea anemones and other hexacorallians, I documented the feasibility of using an individual nematocyst feature in a phylogenetic context, undertook a monograph of nematocyst diversity across Actiniaria, analyzed the distribution of nematocyst morphology with the use of a phylogenetic tree, and utilized multivariate statistics to assess whether quantitative data supports the morphological groups determined by qualitative assessment. I follow with a summary of the goals of each chapter and a brief summary of the major findings of each.

Chapter 2 serves as a case study for the use of nematocyst features as a phylogenetic character by documenting the exact morphology and distribution of one aspect of nematocyst morphology: the apical structure. I used scanning and transmission electron microscopy to document the exact nature of the apex of undischarged capsules to determine the distribution of the two known apical structures, apical flaps and operculum.

A specific objective was to determine if apical flaps are a synapomorphy of Anthozoa or of a subset of the class. I found that there are three apical structures, not two, with the third, the apical cap, previously known from spirocysts. Apical flaps are in fact restricted to Actiniaria and serve as a morphological synapomorphy for the group. Also, the apical cap is ancestral for cnidae, spirocysts and ptychocysts arose from a nematocyst that is most parsimoniously assumed to have an apical cap.

Chapter 3 is a monograph of “rhabdoid” nematocyst morphologies across

Actiniaria with some data on other anthozoans. I used scanning and transmission electron microscopy as well as light microscopy to document morphological variation in all nematocysts hypothesized in one of the two major classification systems (Weill 1934 and Schmidt 1969) to have a shaft (which are the groups that have historically caused the most confusion). A specific objective was to evaluate how well or poorly the two systems categorized this variation into types and determine ways these systems could be improved. I found nine distinct morphologies that vary in characters visible by both light and scanning electron microscopy. Some of these categories agreed with previously identified categories (such as Schmidt’s (1969) p-rhabdoid A and p-rhabdoid B1a), while others did not. To better classify the variation clearly without implying an evolutionary relationship or similarity amongst types, several new categories are described and named.

Chapter 4 takes the morphologies outlined in chapter 3 and looks at the distribution of each type in Actiniaria in a phylogenetic context. I used three genes (16S,

18S, 28S) to construct a phylogenetic tree of Actiniaria using maximum likelihood.

Among the taxa included in the tree were 26 actiniarians for which I obtained detailed morphological data for nematocysts of distinct body regions (acontia, acorhaghi, body column, mesentrial filaments, and tentacles). A specific objective was to evaluate the evolution of cnidomes (i.e. the complement of cnidae) as well as the evolution of distinct morphological types. Additionally, data was collected from 17 other anthozoans, allowing for some analysis of the evolution of cnidomes throughout Hexcorallia. I found that nematocyst morphologies have distributions that are highly related to phylogeny,

4 with the presence of certain types in particular tissues defining monophyletic groups.

Therefore, better categorization of nematocyst morphology results in a clearer pattern of evolution amongst the morphologies.

In chapter 5, multivariate analyses were utilized to assess the robustness of the morphological groups formed in chapter 3. I took detailed measurements of several features that vary amongst the morphologies (e.g. basal tubule length and width, spine length and width) creating a dataset of scanning electron microscope observed variables and light microscopy observed variables. These datasets (and another which included a subset of the morphologies that had additional SEM variables measured) were analyzed using principle components analysis (PCA) and non-metric multidimensional scaling

(NMS) to look for patterns amongst the nematocysts. A specific objective was to assess how well the morphologies defined in chapter 3 cluster in a multivariate analysis when using qualitative data. A linear discriminant function analysis provided an opportunity to evaluate the ability of chosen variables to classify my a priori groups (the morphology classification). This study allows for a somewhat independent evaluation of the morphological groupings based on qualitative data and provides data on which morphologies resemble each other morphologically. I found support for several of the nine morphologies in both the clustering of each morphology in PCA and NMS as well as in the success of the discriminant function analysis to correctly classify the morphologies.

Some groups however (such as the hadroneme and p-rhabdoid B2d) are not so easily distinguished and therefore may not currently be well circumscribed.

References

Bigger CH. 1988. The role of nematocysts in anthozoan aggression. In: Hessinger DA, Lenhoff HM, editors. The Biology of Nematocysts. New York: Academic Press. p 295- 308.

Carlgren O. 1940. A contribution to the knowledge of the structure and distribution of the cnidae in the Anthozoa. K Fysiogr Sällsk Handl 51:1-62.

Carlgren O. 1945. Further contributions to the knowledge of the cnidom in the Anthozoa especially in the Actiniaria. K Fysiogr Sällsk Handl 56:1-24.

Cutress CE. 1955. An interpretation of the structure and distribution of cnidae in Anthozoa. Syst Zool 4:120-137.

Fautin DG, Mariscal RN. 1991. Cnidaria: Anthozoa. In: Harrison FW, Westfall JA, editors. Microscopic Anatomy of Invertebrates, Vol. 2. Placozoa, Porifera, Cnidaria, and Ctenophora. New York. p 267-358.

Mariscal RN. 1974. Nematocysts. In: Muscatine L, Lenhoff HM, editors. Coelenterate Biology: Reviews and New Perspectives. New York: Academic Press. p 129–178.

Mariscal RN. 1984. Cnidaria: Cnidae. In: Bereiter-Hahn J, Matoltsy AG, Richards KS, editors. Mariscal RN, Conklin EJ, Bigger CH. 1977b. The ptychocyst, a major new category of cnidae used in tube construction by a cerianthid anemone. Biol Bull 152:392- 405.

Mariscal RN, McLean RB, Hand C. 1977a. The form and function of cnidarian spirocysts. Cell Tiss Res 178:427-433.

Östman C. 2000. A guideline to nematocyst nomenclature and classification, and some notes on the systemic value of nematocysts. Sci Mar 64(Sup 1): 31-46.

Rifkin JF. 1991. A study of the spirocytes from the Ceriantharia and Actiniaria (Cnidaria: Anthozoa). Cell Tiss Res 266:365-373.

Rifkin J, Endean R. 1983. The structure and function of the nematocysts of Chironex fleckeri Southcott, 1956. Cell Tiss Res 233:563–577.

Schmidt H. 1969. Die Nesselkapseln der Aktinien und jhre differentialdiagnostische Bedeutung. Helgol Meeresunters 19:284-317.

Schmidt H. 1974. On evolution in the Anthozoa. Proc. 2nd Intl Coral Reef Symp 1:533– 560.

Weill R. 1930. Essai d’une classification des nématocystes des cnidaires. Bull Biol France Belg 64:141-153.

Weill R. 1934. Contribution à l’étude des cnidaires et de leurs nématocystes. Trav Sta Zool Wimereux 10/11:1–701.

Chapter 2: Morphology, distribution, and evolution of apical structure of nematocysts in Hexacorallia

Introduction

With extensive variation in several morphological features of nematocysts, determining if that variation contains phylogenetic signal has been the focus of several morphological studies (Weill 1934, Carlgren 1940, 1945, Cutress 1955, Westfall and

Hand 1962, Westfall 1965, Schmidt 1969, 1972, 1974, den Hartog 1980, Östman 1983,

1987). The structure of the apex of the capsule is one of the few aspects of nematocyst morphology that seems to provide phylogenetic information.

A nematocyst consists of a capsule with a tubule attached at one end. The attached tubule is contained within an undischarged capsule. Under appropriate stimulation, the apex of the capsule opens and the tubule everts from the capsule, typically revealing spines and releasing toxin (Mariscal 1974). The apex has two known forms: an operculum or flaps. An operculum is a plug-like structure varying in form from circular to trilobed and in size from about 0.8 to 1.5 µm in diameter (Östman 1982, Rifkin and

Endean 1983, Yanagihara et al. 2002, Reft et al. 2009). Regardless of its shape or size, an operculum is hinged, swinging to one side to allow the tubule to evert. In contrast, apical flaps consist of three triangles that fold outward (Westfall and Hand 1962, Westfall 1965,

Godknecht and Tardent 1988, Salleo et al. 1991). An individual flap is about 1-1.26 µm from its base to its tip (Reft et al. 2009).

The apical structure of the nematocyst capsule has been inferred to reflect the phylogenetic relationships of four classes of the phylum, Hydrozoa, Scyphozoa,

Cubozoa, and Anthozoa (Westfall and Hand 1962, Westfall 1965, Mariscal 1984). From electron microscopy, Westfall (1965) concluded that hydrozoan and scyphozoan nematocysts have opercula whereas anthozoan nematocysts have three apical flaps. From this evidence, she inferred that the Hydrozoa and Scyphozoa are more closely related to each other than either is to Anthozoa. Cubozoa was not defined as a class until 1975: at the time of Westfall’s study, Cubozoa was considered an order of Scyphozoa (Werner

1975). Subsequently, cubozoan nematocysts have been found to bear opercula (e.g.

Rifkin and Endean 1983; Yanagihara et al. 2002). Westfall’s (1965) hypothesis was corroborated by genetic evidence, which supports the monophyly of a clade that includes

Scyphozoa, Cubozoa, and Hydrozoa (Bridge et al. 1995, Collins 2002). Nothing has been reported about the apical structure of nematocysts from animals belonging to

Staurozoa (Marques and Collins 2004), a recently-erected class that consists of animals that were formerly ranked as an order of Scyphozoa.

Nonetheless, the inference about the phylogenetic import of apical structures is flawed. Apical flaps are not characteristic of the nematocysts of all anthozoans. Renilla, an anthozoan of the subclass Octocorallia, has an “operculum-like” structure, not apical flaps (Ivester 1977). Schmidt (1969) claimed that nematocysts of the orders Scleractinia and Corallimorpharia (class Anthozoa, subclass Hexacorallia) have neither apical flaps nor opercula; den Hartog (1980) confirmed this claim for Corallimorpharia. However,

neither Schmidt (1969) nor den Hartog (1980) described or depicted the morphology of

the apex of the nematocyst from these groups.

den Hartog (1980) stated that apical flaps are confined to the hexacorallian order

Actiniaria (sea anemones), the one group in which apical flaps had been documented.

However, even within Actiniaria, apical morphology is poorly studied. Schmidt (1969)

noted that in actiniarians, one type of nematocyst (his p-rhabdoid A) lack apical flaps

(which he called opercular valves). Furthermore, although Westfall (1965) found that

five types of nematocysts from Metridium senile fimbriatum have apical flaps, she

studied nematocysts from only tentacles and acontia, and did not assay nematocysts

belonging to the category Schmidt (1969) identified as lacking apical flaps. Godkneckt

and Tardent (1988) documented apical flaps in Anemonia sulcata Pennant, 1777, but examined only tentacles, and only one type of nematocyst, the microbasic mastigophore.

No one has reported apical structure of nematocysts from throughout the body, and the

nematocysts of some nematocyst-dense structures, such as acrorhagi or catch tentacles

have never been assayed for apical structures.

We survey the apical structure of nematocysts across a broad sample of Anthozoa,

focusing most heavily on members of subclass Hexacorallia. As a secondary objective,

we perform a more thorough survey of apical structure of nematocysts throughout the

body of actiniarians. Both scanning and transmission electron microscopy (SEM and

TEM) provide ways to image the apex of a nematocysts that allow full description of the

morphology. Comparative studies of nematocysts and of ptychocysts and spirocysts

10 provide the necessary information to evaluate the phylogenetic utility of apical structure of nematocysts.

Materials and methods

Materials

We examined two representative octocorallians, at least one representative of each order of Hexacorallia, and one representative stauromedusan (Table 2.1). Hydrozoans provide comparative data on the structure of the operculum. A complete survey of all tissues and types of nematocysts was performed for the actiniarians Metridium senile

(Linnaeus 1761) and Bunodosoma cavernata.

Specimens were collected in the field from various localities (see Table 2.1) or obtained from commercial sources such as Aquarium Adventure (Columbus, OH), Gulf

Specimen Marine Lab (Panacea, FL), and Reef Hot Spot (Inglewood, CA). A few samples, such as the stoloniferan octocoral Clavularia sp. A (sensu Parrin et al., 2010), were obtained from cultures maintained by Neil Blackstone at Northern Illinois

University. The antipatharian samples were collected, fixed, and provided by Anthony

Montgomery (see Opresko 2009 for collecting details). All specimens were identified to genus by the collector (or faculty of Kyoto University Seto Marine Lab) using morphology. Specimens from Aquarium Adventure, Reef Hot Spot, and the identification by the Gulf Specimen were verified by the authors. Two specimens are not fully identified; one belongs to the genus Actinostola and is therefore listed as Actinostola sp., the other is only known as an acontiarian and is therefore known as Acontiaria sp.

Methods

For each species, the cnidom was initially determined with differential interference contrast microscopy at Ohio State University using a Leica DM LB2 microscope (see

Fig. 2.1 for examples of each nematocyst type identification with light microscopy) and confirmed with scanning electron microscopy. We follow the classification system for nematocysts developed by Weill (1930, 1934), as modified by Carlgren (1940) and

Mariscal (1974). The initial determination of apical structure was based on SEM.

Because apical flaps consist of three parts, at least one flap will be in view from any angle when imaging the apical structure of a nematocyst using SEM. Although the lack of an obvious structure at the apex eliminates apical flaps as a possibility, it does not eliminate the operculum as a potential apical structure. In discharged nematocysts, the operculum can only be seen on the side of the capsule on which the hinge of the operculum is attached. TEM is necessary to definitively determine the apical structure of a nematocyst lacking apical flaps.

Nematocysts listed in Table 2.2 from various species and body regions were prepared for SEM. After the dissection of the body regions, each sample was placed in a 1M sodium citrate solution for 15 min to induce expulsion of nematocysts from nematocytes

(Yanagita 1959, Reft et al. 2009). Following three washes with distilled water, most samples were placed in 70% ethanol. Samples from Bunodosoma cavernata, Hydra sp., and Metridium senile were placed in a 1% OsO4 solution overnight before being placed in 70% ethanol. All specimens were dehydrated in ethanol, then critical-point dried with

CO2. Samples from B. cavernata, Hydra sp., and M. senile and were sputter-coated with

gold palladium in a Hummer sputter coater and examined using a LEO 1550 field

emission scanning electron microscope at the University of Kansas, Lawrence. All other

samples were sputter-coated with gold palladium in a Cressington sputter coater and

examined using a FEI NOVA nanoSEM at the Ohio State University, Columbus.

A subset of the samples were also prepared for TEM (see Table 2.3). Samples from

Haliclystus borealis were initially preserved in 70% ethanol. These specimens were

rehydrated through a graded ethanol series back to water before being placed in

phosphate buffer. All specimens were fixed with gluteraldehyde: small bits of tissue

were fixed in 2.5% glutaraldehyde in phosphate buffer at pH 7.4 and postfixed in 1%

OsO4 in phosphate buffer. All specimens were dehydrated in a graded ethanol series and embedded in Epon. Thin sections (70 nm) cut on a Leica EM UC6 Ultramicrotome were stained with 2% aqueous uranyl acetate for 15 min followed by 5 min in Reynold’s lead citrate. Micrographs were taken on a Technai G2 Spirit TEM at the Ohio State

University, Columbus. Size and shape differences in nematocyst capsules usually (but not always) enabled the identification of different nematocyst types when more than one was present.

Results

SEM: Hydrozoa

The apical structure of hydrozoan nematocysts was similar in all samples examined.

The apex of nematocysts of Hydra sp. (Fig. 2.2A, B) is representative of all hydrozoans

examined (i.e., Cordylophora caspia; Ectopleura larynx; and Neoturris breviconis).

Each hydrozoan nematocyst has a singular apical structure, i.e. the operculum, visible

with SEM (Fig. 2.2A, B). The hinged operculum varies in shape from ovoid (Fig. 2.2A)

to triangular (Fig. 2.2B), with shape corresponding to nematocyst morphological type.

At the capsule apex, the capsule walls not attached to the operculum form a thickened

ridge with which the operculum lies flush before discharge (Fig. 2.2A). After the tubule

has everted, the operculum lies in the same plane as the tubule, usually against the tubule

itself, and does not flex away from the tubule (Figs. 1.2A, B).

SEM: Actiniaria

We found two apical morphologies in the examined nematocysts from Actiniaria. The majority of actiniarian nematocysts examined had a three part apical structure known as apical flaps, as seen in Bartholomea annulata (Fig. 2.2C). Each of the three flaps is triangular. After discharge, the flaps flex away from the tubule rather than lie against it, and at least one flap is always visible (Figs. 1.2C, D). Often a central ridge is apparent on at least one of the flaps (Fig. 2.2C). Although most actiniarian nematocysts studied here had the tripartite flaps regardless of nematocyst type or body location (Table 2.2), the survey of nematocysts of all body regions of Bunodosoma cavernata revealed that microbasic p-mastigophores of the mesenterial filaments do not have the tripartite apical structure (Fig. 2.2E). This nematocyst has no obvious structures at the apex. The microbasic p-mastigophores of the mesenterial filaments of Actinostola sp. also lack any conspicuous apical structure. Rather, the capsule ends without any extension of the

capsule wall into a visible apical structure (Fig. 2.2E). In contrast, the mesenterial

filaments of Metridium senile contain microbasic p-mastigophores that do have obvious

apical flaps (Fig. 2.2F). Nematocysts in other body regions of B. cavernata and

basitrichs of this species and S. coccinea in the same body region all had the three-flap

apical structure (Table 2.2).

SEM: Other anthozoans

Nematocysts (microbasic b-mastigophores) in all body regions of the cerianthid

Ceriantheopsis americanus show no sign of any visible apical structure (Fig. 2.2G): the

capsule apex is smoothly rounded and bears no trace of a thickened rim, a flap, or hinge.

Nematocysts from the tentacles of another cerianthid, Cerianthus filiformis, are identical

in apical structure to those of C. americanus. Ptychocysts of Ceriantheopsis americanus

also reveal no evidence of flaps or an operculum (Fig. 2.2O). Similarly, both types of

nematocysts (microbasic p-mastigophores and basitrichs) of the antipatharians Antipathes

grandis and Antipathes griggi also have no obvious structures at the apex of the capsule

(Fig. 2.2H). Furthermore, the holotrichous isorhizas (holotrichs) of the octocoral Renilla

mulleri lack evident apical structures (Fig. 2.2I).

Microbasic p-mastigophores of the corals Platygyra astreiformis and Gonipora sp. are identical in having a smooth end to the capsule with no obvious projections (Fig. 2.2J).

The apex of these nematocysts closely resembles the apex of the microbasic p- mastigophores in the mesenterial filaments of Bunodosoma cavernata and Actinostola sp. but does not resemble the apex of nematocysts having flaps or an operculum. However,

in the holotrichs of P. astreiformis, the apical wall may extend into a thin collar around

the tubule (Fig. 2.2K). Other than this collar, no other apical structures are evident in the

holotrichs of either coral studied. The nematocysts of corallimorpharians Discosoma sp.

and Rhodactis sp. show no evidence of any extensions of the capsule wall (Fig. 2.2L).

Some discharged holotrichs of Discosoma sp. bear cracks in the apical capsule wall near

the everted tubule (Fig. 2.2L). These cracks appear in an extension of the capsule wall

that is similar to the collar seen in some holotrichs of P. astreiformis (Fig. 2.2K).

Holotrichs of the zoanthids Protopalythoa mutuki and Zoanthus pulchellus have no obvious apical structure (Fig. 2.2M). Basitrichs of these species sometimes have roughly triangular structures at the apex of the capsule (Fig. 2.2N). These structures are irregular in shape and number, in some cases appearing to have only one element, in other cases having at least two. These structures, when present, always lie against the tubule and never flex outward from the tubule as do apical flaps. Furthermore, deep cracks in basitrich capsule apices (as in Fig. 2.2N) are occasionally present.

TEM: Medusozoa

Transmission electron micrographs of stenoteles from the tentacles of Ectopleura larynx and Sertularella sp. depict nematocysts that have identical apical structures (Fig.

2.3A). The capsule wall extends out and then curves back down towards the capsule, forming a concave space at the apex of the nematocyst. This space is filled with a material that appears more electron-dense than the capsule wall under TEM. This material forms the operculum, and the capsule wall extension forms the rim seen in the

scanning electron micrographs. The tubule wall attaches to the infolded section of the

apical capsule wall (Fig. 2.3A).

The tentacles of the staurozoan Haliclystus borealis contain two types of nematocysts: euryteles (Fig. 2.3B) and holotrichs (Fig. 2.3C). Micrographs of the euryteles (Fig. 2.3B) depict an extension of the apical capsule walls similar to that seen in hydrozoan nematocysts, creating a central apical space filled with electron-dense material, i.e. operculum. The material that makes the operculum is more electron-dense than the capsule wall but is not uniform in its electron-density: striations of more and less electron-dense material are evident (Fig. 2.3B). The tubule attaches to the apical extensions of capsule wall directly beneath the operculum.

We obtained no clean sections through undischarged nematocysts that could be clearly identified as holotrichs in H. borealis. However, a micrograph depicting a partially discharged holotrich (Fig. 2.3C) provides some evidence of apical structure. The capsule wall is continuous at the apical end, ending with a thickened part similar to the rim seen in hydrozoan nematocysts with opercula.

TEM: Anthozoa

Basitrichs from the tentacles of both Anthopleura elegantissima and Condylactis gigantea have similar apices when viewed with TEM (Fig. 2.3D). The apex is occluded by structures (flaps) containing electron -dense and -lucent layers. Striated material fills the central space where the tips of the flaps meet: this material extends down into the

tubule. The tubule layer can be traced on the capsule wall below the apical flaps (Fig.

2.3D).

All non-actiniarian anthozoan nematocysts studied here with TEM had a similar apical

structure, which is neither an operculum nor apical flaps (Table 2.3, Fig. 2.3F-J).

Basitrichs from the tentacles of the zoanthid Protopalythoa mutuki (Fig. 2.3F), b- mastigophores from all tissues of Ceriantheopsis americanus (Fig. 2.3G), microbasic p- mastigophores from the tentacles of Goniporia sp. (Fig. 2.3H), and holotrichs (Fig. 2.3I) and microbasic p-mastigophores (Fig. 2.3J) from the mesenterial filaments of Discosoma sp. all lack an apical structure distinct from the general capsule morphology. In these nematocysts, the thick capsule wall ends abruptly on either side of the apex (Fig. 2.3F-J).

At this junction, a thinner tubular layer attaches to the capsule wall, extending to the

center of the nematocyst apex before curving down into the central space (Fig. 2.3F-J).

The tubule contains electron-dense spines (Fig. 2.3F-H, J). The apex is filled with an

electron-dense material that forms a cap for the nematocyst. The tubule layer underlies

this cap. The cap extends down the sides of the nematocyst to attach to the outer sides of

the capsule walls forming a structure hereafter referred to as an apical cap. This layer of

electron dense material does not appear to line the outside of the whole capsule (Fig.

2.3F-J): either it is only present at the apex or it forms a very thin layer on the outer wall

of the nematocyst that is difficult to visualize. Often, the spines contained inside the

inverted tubule abut the cap (Fig. 2.3F, G). In some micrographs (such as Fig. 2.3G), the

central-most portion of the cap does not appear. Presumably, this is either an artifact of

fixation or the section was not cut all in the same plane.

Micrographs of microbasic p-mastigophores from the mesenterial filaments of the sea anemone Actinostola sp. (Fig. 2.3E) have an apical cap similar to that of non-actiniarian anthozoan nematocysts (Fig. 2.3F-J) rather than apical flaps found in other sea anemones

(Fig. 2.3D). However, unlike the nematocysts of other hexacorallians, the microbasic p- mastigophores of S. coccinea have a central thickening of the apex of the cap, forming a mound of material (Fig. 2.3E). The material that makes the cap also appears to fill the inverted tubule and surrounds the anterior end of the spines.

In the holotrichs of the octocoral Clavularia sp. A, the tubule does not attach to the center of the capsule apex (Fig. 2.3K). Micrographs of these nematocysts reveal no evidence of structures or material that occlude the apex as in actiniarian nematocysts bearing apical flaps. Furthermore, the capsule wall does not fold to create a convex space that is filled by a plug of material. A mat of material extends over the capsule walls, forming a structure that is consistent with the apical cap seen in non-actiniarian hexacorallians.

All spirocysts examined have the same apical structure (Fig. 2.3L). The inner capsule wall is serrated. The capsule wall does not extend to the apex of the capsule: the wall stops abruptly on both sides and the tubule layer attaches at these points. A cap of electron dense material covers the apex with the tubule layer underlying it. The structure resembles that of the apical cap of nematocysts of non-actiniarian anthozoans.

Although the large size of ptychocyst capsules makes obtaining a clean section difficult, we saw no evidence of capsule wall extension into apical flaps in C. americanus ptychocysts (Fig. 2.3M). Furthermore, no plug of material similar to an operculum is

19 evident. Instead, the ptychocyst has a thick mat of material at the apex similar (though thicker) to the apical cap seen in spirocysts and the nematocysts of non-actiniarian anthozoans. This material is of similar electron-density to the material that forms the cap in other cnidae.

Discussion

The three forms of nematocyst apical structure

We document three distinct apical structures: operculum, apical flaps, and apical cap.

An operculum and apical flaps can usually be clearly identified with SEM whereas the apical cap requires TEM for definitive identification.

Operculated nematocysts have been documented in detail before, and our findings agree with previous descriptions of the structure from both SEM (e.g. Rifkin and Endean

1983, Östman 1982) and TEM (e.g. Chapman and Tilney 1959a, b, Mattern et al. 1965,

Westfall 1970, Burnett 1971a, b, Holstein 1981, Hessinger and Ford 1988). We document the structure for the first time in nematocysts of Cordylophora caspia,

Ectopleura larynx, Neoturris breviconis, and Sertularella sp. We have documented apical structure in a staurozoan, Haliclystus borealis. Euryteles of this species are clearly operculated. The apical space created by the extension of the capsule wall (Fig. 2.3B) is never as large or broad as in hydrozoans (Fig. 2.3A), but is structurally very similar.

Although the images are imperfect, current evidence suggests that holotrichs of H. borealis are operculated. The capsule wall extends across the apex without protrusion, unlike an apical flap or an apical cap. Furthermore, the capsule wall ends apically in a

20 thickened cup-shape, similar to the rim seen in hydrozoan nematocysts and in euryteles of

H. borealis.

Apical flaps have also been previously documented both with SEM (e.g. Mariscal

1974, Godknecht and Tardent 1988, Salleo et al. 1991, Reft et al. 2009) and TEM

(Westfall and Hand 1962, Westfall 1965, Watson and Mariscal 1985, Blake et al. 1988,

Reft et al. 2009). The nematocysts described and depicted here agree with previous reports in terms of the morphology and structure of the flaps. Here we document this structure for the first time in nematocysts of Anthopleura elegantissima, Bartholomea annulata, Bunodosoma cavernata, Condylactis gigantea., Acontiaria sp., and Halcurias levis.

The apical cap has previously been identified in spirocysts (Westfall 1965, Mariscal et al. 1976, Mariscal and McLean 1976, Rifkin 1991). It has not been clearly documented in nematocysts prior to this study. Here, using TEM, we definitively document this structure in Ceriantheopsis americanus, Discosoma sp., Goniporia sp., Protopalythoa mutuki, and the octocoral Clavularia sp. A. SEM reveals no evidence of either apical flaps or an operculum and therefore strongly implies the presence of the apical cap in the nematocysts of several more species: Antipathes grandis and A. griggsi, Cerianthus filiformis, Platygyra astreiformis, Renilla mulleri, Rhodactis sp., and Zoanthus pulchellus. In addition, TEM reveals that the microbasic p-mastigophores of the mesenterial filaments in Actinostola sp. also have an apical cap, although it differs slightly in forming a central mound at the apex. Despite the difference in shape, the relationship and relative positions of the tubule and capsule layers in these microbasic p-

mastigophores are identical to the relationship and relative positions of those layers in

other nematocysts with apical caps. The cap consists of a mat of electron-dense material

that curves across the apex of the capsule. This mat is distinct from and overlaps the

capsule walls (Fig. 2.3E-L). The serrated inner wall of spirocysts (Westfall 1965,

Mariscal and McLean 1976, Rifkin 1991) makes it clear that the tubule layer underlies

the cap on the inside of the spirocyst (Fig. 2.3L). Likewise, the tubule layer underlies the

apical cap in nematocysts. In TEM pictures, the apical cap is identical in structure in

both spirocysts and non-actiniarian nematocysts.

The apical structure of ptychocysts remains unclear. Neither this study (Figs. 1.2O,

1.3M) nor earlier work by Mariscal et al. (1977) found evidence of either apical flaps or

an operculum on ptychocysts. Mariscal et al. (1977) describe the apex of ptychocysts as

having apical folds that are “sealed along several suture planes” (p. 397). These apical

folds (Mariscal et al. 1977: Fig. 2.11) resemble the triangular structures often seen in

basitrichs of zoanthids (Fig. 2.2N), in that they are irregularly shaped and lie against the

tubule; apical flaps as previously defined have a very regular shape and always project

away from the tubule. Furthermore, apical flaps open cleanly, without any evidence of

material sealing the flaps together, whereas the apical folds of Mariscal et al. (1977) more

closely resemble a rupture of a seam (Fig. 2.11) with the sealant leaving some residue where the folds had previously been sealed. Furthermore, the TEM images (Fig. 2.2M)

presented here reveal a thick mat of material at the apex similar in placement and

electron-density to the apical cap. This mat is thicker than the apical cap of spirocysts or

nematocysts, however, the large size of these ptychocysts (often over 60 µm) might

influence the size of the mat. Therefore, the most likely apical structure is an apical cap

similar to that found in spirocysts and non-actiniarian nematocysts. However, more TEM

study is necessary to fully characterize the apical structure of ptychocysts.

Functional differences in apical structure

Although all three apical structures perform the same function, structural differences

among them necessitate differences in how they achieve that function. The apical

structure in any form functions to hold the contents inside the capsule until the

appropriate time. When stimulated, the apical structure opens the apex to allow for the

eversion for the tubule. Previous work in operculated hydrozoan nematocysts has shown

that mechanical energy is stored in the capsule walls; this energy in addition to the high

osmotic pressure of the capsule is thought to power discharge (Dujardin 1845, Jones

1947, Carré 1980, Weber 1989, Holstein and Tardent 1984, Tardent 1988, Holstein et al.

1994, Szczepanek et al. 2002, Özbek et al. 2009). The discharge of hydrozoan nematocysts has been documented with high-speed cameras (Holstein and Tardent 1984,

Nüchter et al. 2006). The entire process takes less than 3 ms, and the operculum opens too quickly to be distinguished from the eversion of the basal part of the tubule (Holstein and Tardent 1984, Tardent 1988, Nüchter et al. 2006). The discharge of the tubule occurs immediately after the opening of the apical structure.

However, the tubule eversion does not seem to always follow immediately after the opening of the apical flaps, and may have a significant delay, 60 s or longer (Godknecht and Tardent 1988, Salleo et al. 1991). Whether nematocysts with apical flaps or caps are

23 held under the same order of osmotic pressure (15 MPa) as operculated nematocysts

(Weber 1989, Holstein et al. 1994, Nüchter et al. 2006) has not been demonstrated, though presumably discharge is affected similarly mechanically and chemically in all nematocysts including those with apical flaps (Tardent 1988).

Unlike the other two apical structures, the apical cap does not appear to have a hinge

(as with the operculum) or base (as with apical flaps) that would allow it to move cleanly out of the capsule opening and thus out of the path of an everting tubule. Therefore, not all nematocysts have a lid structure as has been assumed (Özbek et al. 2009). SEM reveals that the apex of discharged nematocysts with caps do not have any regular form.

In some nematocysts, roughly triangular shapes may form (Fig. 2.2N), but in others the apex forms a smooth (Fig. 2.2K) or broken collar (Fig. 2.2L) around the tubule, break in the capsule apex (Fig. 2.2N), or have nothing obvious at all (Fig. 2.2E, G-I, J, M).

Presumably, the cap breaks open in some way to allow the tubule to evert, therefore these differences in form reveal that although the cap is shared by these nematocysts, the way in which the cap breaks may differ among nematocysts. The cap may function as suggested by Mariscal et al. (1977) for ptychocysts, having weak points that can be ruptured with the force of the discharging tubule. However, we saw no evidence of suture lines or areas of thinner material of the cap in TEM as would be expected of structurally weaker areas. If this force is important in opening the apex of the nematocyst, then we would predict that discharge in a nematocyst with an apical cap will be more similar to that of the operculum, in which the tubule eversion is indistinguishable

from the opening of operculum, than to apical flaps, which may have a delay in tubule

eversion.

Conversely, the cap, which consists of less structured electron-dense material than the

operculum or an apical flap, could dissolve during the discharge process. In some cases

(Figs. 1.2E, J), the apex of the discharged nematocyst appears to lack any sort of apical

structure, as would be expected if the cap had dissolved. However, other nematocysts do

have some sort of apical structure, presumably the remnants of the cap (Fig. 2.2K, N),

although the form that structure takes is inconsistent. More than one discharge

mechanism could be possible within Anthozoa, with different lineages utilizing different

strategies.

Furthermore, on the basis of unpublished data, Özbek et al. (2009) argue that the operculum changes in molecular arrangement during discharge into a more loose arrangement of the structural proteins. Therefore, they propose that ionic changes that trigger discharge will cause molecular changes in the operculum creating designated weak points that will break to allow the operculum to open while preventing ruptures in the capsule walls (Özbek et al. 2009). The cap may function in a similar way, with the discharge process initiating changes in some or all of the proteins that form the structure.

However, if this process is similar in capped nematocysts, occasional ruptures in the capsules of these discharged nematocysts reveal that the process is not as clean as in operculated nematocysts. Whether a similar process occurs with nematocysts with apical flaps is unknown as we lack a full characterization of the discharge process in non- operculated nematocysts.

Distribution of apical structures and its significance

All nematocysts of all medusozoans studied to date have an operculum (e.g.

Scyphozoa: Burnett 1971a, b, Cubozoa: Rifkin and Endean 1983, Yanagihara et al. 2002,

Hydrozoa: Chapman and Tilney 1959a, b, Mattern et al. 1965, Westfall 1970, Holstein

1981, Östman 1982, Hessinger and Ford 1988). No nematocyst from any anthozoan has been found to have an operculum. This suggests that operculated nematocysts are a synapomorphy for Medusozoa, an idea that has previously been suggested (Westfall and

Hand 1962, Westfall 1965, Mariscal 1984). This study clarifies one critical point: the apical structure of nematocysts of members of Staurozoa, a class erected by Marques and

Collins (2004) for members of the scyphozoan order Stauromedusa. The finding of an operculum in staurozoan nematocysts is critical because Staurozoa has been interpreted to be the sister group to other Medusozoa (Collins and Daly 2005, Collins et al. 2006, van

Iten et al. 2006). Furthermore, the operculum appears to be the only apical structure present in this clade: all types of nematocysts in Medusozoa have an operculum.

However, the polarity and transformation of apical structures cannot be fully understood without a fuller appreciation of the apical structure of nematocysts in anthozoans or of ptychocysts or spirocysts.

Anthozoan nematocysts have generally been inferred to bear apical flaps (Westfall

1965, Westfall 1966a, Mariscal 1974, 1984, Werner 1984, Kass and Scappaticci 2002).

Although some authors have disagreed with this generalization (e.g., Ivester 1977;

Schmidt 1969, 1981, den Hartog 1980), the actual morphology of the apical structure has

26 not been well described for these taxa, and no images have been published. Our data clarifies the distribution of apical flapped-nematocysts within Anthozoa. We find that apical flaps are restricted to nematocysts of Actiniaria. Other hexacorallians have nematocysts with apical caps; no hexacorallian we examined has nematocysts with an operculum. Tripart apical flaps are characteristic of members of all major lineages of

Actiniaria studied to date, and represent a morphological synapomorphy for Actiniaria.

Although widely recovered as monophyletic in DNA-based studies, Actiniaria does not have an anatomical synapomorphy and its members are more diverse in polyp anatomy and life history than those of any other anthozoan group (reviewed in Daly et al. 2007).

From an anatomical standpoint, the affiliation of enigmatic, morphologically distinct suborders Protantheae and Endocoelantheae with Nynantheae, the group to which the majority of Actiniaria belong has been particularly problematic, with some authors suggesting that these suborders are more closely allied with other hexacorallian lineages than with nynanthean actiniarians (Carlgren 1914, 1949, Stephenson 1920, Schmidt

1974). The apical flaps seen on the nematocysts of the endocoelanthean Halcurias levis

(Fig. 2.2D) ally this enigmatic taxon with Nynantheae, and the lack of an apical cap rules out association with any other hexacorallian order. The nematocysts of protanthean actiniarians have not yet been studied.

Although apical flaps appear to only be found in nematocysts of Actiniaria, not all actiniarian nematocysts have apical flaps. In some taxa, microbasic p-mastigophores

(Fig. 2.1F, G) do not have apical flaps (Figs. 2E, 3E); in others this same kind of nematocyst (Fig. 2.1D, E) clearly has flaps (Fig. 2.2F). All actiniarian microbasic p-

27 mastigophores documented here are from a single region, the mesenterial filaments; this and the actinopharynx are the only regions from which Schmidt (1969) reported p- rhabdoids A, which he described as lacking flaps. The microbasic p-mastigophore nematocysts that lack apical flaps (from B. cavernata and S. coccinea in this study) are consistent in morphology with Schmidt’s (1969) p-rhabdoid category A. Therefore, this study provides some support for Schmidt’s argument that there are subcategories of the microbasic p-mastigophore (which he called p-rhabdoides). However, Schmidt (1969) also characterized the family to which Metridium senile belongs as having these p- rhabdoids A, and the microbasic p-mastigophores of M. senile clearly bear apical flaps

(Fig. 2.2F). A more extensive survey of this particular nematocyst type within Actiniaria is necessary to determine whether the cap (versus flap) is generally characteristic of this kind of nematocyst, and to accurately document its taxonomic occurrence.

The apical structure of octocorals has not been fully explored. In the species examined here, apical structure is inconsistent with either apical flaps (no large structures occluding the apex) or an operculum (no extension of the capsule walls to make a convex space that can be filled with and operculum). Although Ivester (1977) claimed that nematocysts of the octocoral Renilla reniformis do not have apical flaps and suggested that the apical region is more similar to the operculum documented in Obelia longissima

(Pallas 1766) by Westfall (1966b), his images (Ivester 1977: Figs. 10, 12) provide no evidence of a single apical plug such as seen for an operculum nor of a space in which the operculum would fit. Schmidt (1981) further described the development and ultrastructure of octocoral nematocysts, and clearly documented an apical cap (though he

28 did not use the term) in nematocysts of Alcyonium digitatum Linnaeus, 1758 (Schmidt,

1981: Figs. 21 and 23). Furthermore, Schmidt (1981) observed that unlike in medusozoans, the apical material is not gathered in an area formed by the capsule wall

(which forms the operculum), but rather, that apical material extends over the capsule walls at the apex to form this cap.

Implications for evolution of cnidae and their structures

Nematocysts are found in all members of Cnidaria, and are most diverse in the

Hydrozoa (Mariscal 1974). Cnidae, the more general category to which nematocysts belong, are most diverse in Anthozoa, specifically in Hexacorallia. Determining the ancestral state for attributes of nematocysts requires knowledge about those features common to all cnidae.

The apical cap is an ultrastructural feature that is shared between spirocysts and most nematocysts of Hexacorallia (with the exception of most nematocysts of Actiniaria), implying that this structure preceded the divergence of spirocysts and nematocysts.

Current data suggests that ptychocysts also bear an apical cap. Thus, the apical cap can be inferred to predate the divergence of ptychocysts as well as spirocysts (Fig. 1.4A).

Furthermore, the current best estimate of phylogeny within Hexacorallia has Ceriantharia as the sister group to the rest of the Hexacorallia (Won et al. 2001, Daly et al. 2003,

Brugler and France 2007), which means that the apical cap is present at the base of the hexacorallian tree (Fig. 1.4B), being later modified into flaps in some nematocysts of

Actiniaria. Evidence that supports an apical cap structure for octocorals places the

character at the base of the Anthozoan tree, making the cap the plesiomorphic

morphology for the apex of a cnida in Anthozoa (Fig 1.4A).

Although this represents a major step in understanding the evolution of cnidae,

interpreting the ancestral state of nematocysts for the phylum Cnidaria is not yet possible.

Anthozoan nematocysts have one state, the apical cap (with one lineage within the group

evolving another state, apical flaps), whereas medusozoan nematocysts uniformly have

the third state, the operculum. With a clear phylogenetic bifurcation between Anthozoa

and Medusozoa (reviewed in Daly et al. 2007), and no known variation in apical

morphology among the classes of Medusozoa, neither state can clearly be identified as

ancestral (Fig. 2.3B). Either the apical cap or the operculum is the ancestral state for

nematocysts; apical flaps are clearly a derived state, having evolved from apical caps.

Restricting the operculum to Medusozoa has implications beyond interpreting intracnidarian diversification. Several authors (Siddall et al. 1995, Zrzavý et al. 1998, Zrzavý,

2001, Jiménez-Guri et al. 2007) have suggested that the enigmatic metazoan phylum

Myxozoa are highly derived cnidarians. Historically, this conclusion has been based on the observation that myxozoan polar capsules are developmentally and morphologically extremely similar to nematocysts (e.g. Stolc 1899, Weill 1938, Lom and Dyková 1997).

DNA based studies have failed to resolve the issue, as some datasets and methods place this taxon within Cnidaria (Zrzavý et al. 1998, Zrzavý 2001, Jiménez-Guri et al. 2007), whereas others place the group with bilateria (Smothers et al. 1994, Zrzavý and Hypša

2003). Holland et al. (2011) showed that myxozoans have a homologue to the gene that produces minicollagen, one of the primary components of the nematocyst. Furthermore,

30 this minicollagen gene is more like those found in medusozoans than anthozoans, as would be predicted by the nature of the nematocyst apex.

Closer inspection of the polar capsules provides some information that pertains to this question. Micrographs of polar capsules from several different genera of Myxozoa depict apical structure identical to the operculum, i.e. the capsule walls extend apically to create a space filled by a plug (e.g. Dresser et al. 1983: Fig. 20, Lom 1990: Figs. 12, 14, Siddall et al. 1995: Fig. 2D, Cannon and Wagner 2003, Holland et al. 2011: Fig. 2.1A). Because the operculum is characteristic of Medusozoa, this morphological feature provides support for the placement of Myxozoa not only within Cnidaria, but specifically within

Medusozoa. However, if the operculum is the ancestral state for nematocysts, then the similarity between the apex of myxozoan polar capsules and the apex of medusozoan nematocysts is less significant, and would only provide evidence against grouping

Myxozoa within Anthozoa.

If the apical cap were the plesiomorphic apical structure, then the operculum would have evolved from this starting point. The major change required to transform these morphologies is further growth of the apical ends of the capsule wall. This growth would result in an extension of the capsule wall and the creation of a central space that would be filled with the operculum. To transform an operculum to a cap, the capsule wall would need to develop less apically, such that the wall does not fold to create the central space into which the operculum fits. Furthermore, the capsule walls would need to end development earlier, so that the walls do not extend across the apex, but end more abruptly at the sides. Finally, the compact operculum would need to become less

31 organized, losing the repeating striations characteristic of opercular material. The apical cap shows no evidence of any layers or organization as found in the operculum; rather, the cap appears to just be a collection of electron-dense material.

The transition from an apical cap to flaps requires growth from the capsule walls to create the flaps. Furthermore, the central space where all the flaps meet is filled with a material that seems to line a pore leading from the apex into the tubule (Westfall and

Hand 1962, Westfall 1965, Blake et al. 1988, Reft et al. 2009). This central opening makes the flap structure distinctly different from an operculum or cap, neither of which have any obvious openings. However, the shape of the cap in the microbasic p- mastigophore of Actinostola sp. reveals a form that is similar in shape to apical flaps

(with the material of the cap forming a central point) if not in structure. This variant form of the cap might be pertinent to understanding the evolution of the apical flaps in

Actiniaria. However, full characterization of the development of these two structures will be necessary before any conclusions can be made. Characterization of all the molecular steps in nematocyst development has focused on only medusozaoan nematocysts (e.g.

Holstein et al. 1994, Koch et al. 1998, Szczepanek et al. 2002, Özbek et al. 2009). The full development of anthozoan nematocysts, both with caps or with flaps, is not currently understood but necessary to understand the evolution of these structures.

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Table 2.1. Summary of taxa and collection locality used in apical structure study. Higher taxon Species Locality Ceriantharia Ceriantheopsis Gulf Specimen Marine Lab, americanus (Agassiz in Panacea, FL Verrill, 1864) Cerianthus filiformis Kyoto University Seto Marine Lab, Carlgren, 1924 Shirahama, Japan Actiniaria Aiptaisa sp. Aquarium Adventure, Columbus, OH Anthopleura San Juan Island, WA elegantissima (Brandt, 1835) Bartholomea annulata University of Virgin Islands; St. (Le Sueur, 1817) Thomas, US Virgin Islands Bunodosoma cavernata Galveston, TX (Bosc, 1802) Calliactis tricolor (Le Galveston, TX Sueur, 1817) Condylactis gigantea Aquarium Adventure, Columbus, OH Diadumene lineata Jakyakdo, S. Korea (Verrill, 1869) Acontiaria sp. Jakyakdo, S. Korea Halcurias levis Uchida, Kyoto University Seto Marine Lab, 2004 Shirahama Japan Metridium senile Lynn, MA (Linnaeus, 1761) Actinostola sp. San Juan Island, WA Zoanthidea Protopalythoa mutuki Neil Blackstone culture (Haddon and Shackleton, 1891) Zoanthus pulchellus University of Virgin Islands; St. (Duchassaing and Thomas, US Virgin Islands Michelotti, 1864) Antipatharia Antipathes grandis Auau Channel, HI Verrill, 1928 Antipathes griggi Auau Channel, HI Opresko, 2009 Corallimorpharia Discosoma sp. Aquarium Adventure, Columbus, OH Rhodactis sp. Reef Hot Spot, Inglewood, CA Scleractinia Goniporia sp. Reef Hot Spot, Inglewood, CA Continued 41

Table 2.1 continued

Platygyra astreiformis (Milne Edwards and Gulf Specimen Marine Lab, Haime, 1849) Panacea, FL Octocorallia Clavulariid sp. A Neil Blackstone culture Renilla mulleri Kölliker, Gulf Specimen Marine Lab, 1872 Panacea, FL Medusozoa Cordylophora caspia (Hydrozoa) (Pallas, 1771) Exeter, NH Ectopleura larynx (Ellis Darling Marine Center, Walpole, and Solander, 1786) ME Hydra sp. Paulyn Cartwright culture Neoturris breviconis (Murbach and Shearer, 1902) San Juan Island, WA Darling Marine Center, Walpole, Sertularella sp. ME Medusozoa Haliclystus borealis Usujiri Field Station, Hokkaido, (Staurozoa) Uchida 1933 Japan

Table 2.2. Summary of nematocyst SEM apical structure data. Higher taxon Species Body part: Nematocyst/cnidae type Apical Structure Ceriantharia Ceriantheopsis americanus C, MF, T: b-mastigophore* No flaps/oper. C: ptychocyst* No flaps/oper. Actiniaria Aiptaisa sp. MF, T: amastigophore, MF: basitrich Flaps Bartholomea annulata T: basitrich, Flaps A, T: amastigophore Flaps Bunodosoma cavernata MF: p-mastigophore No flaps/oper. MF, T: basitrich, Ac: Acrorhagi Flaps Calliactis tricolor A: basitrich Flaps Condylactis gigantea T: basitrich* Flaps Diadumene lineata T: amastigophore Flaps

43 Acontiaria sp. A: amastigophore, basitrich Flaps

Halcurias levis T: basitrich Flaps Metridium senile A, C, T, MF: amastigophore, Flaps b-mastigophore, MF: p-mastigophore Flaps Actinostola sp. MF, T: basitrich Flaps MF: p-mastigophore No flaps/oper. A: acontia, Column: body column, MF: mesenterial filament, T: tentacle. Asterisk (*) indicates samples for which TEM data is also available, oper.: operculum

Table 2.3. Summary of nematocyst TEM apical structure data. Higher taxon Species Body Structure: Apical Nematocyst/cnidae type Structure Ceriantharia Ceriantheopsis americanus C, MF, T: b-mastigophore* Cap T: spirocyst, C: ptychocyst* Cap Actiniaria Anthopleura elegantissima T: basitrich/spirocyst Flaps/Cap Condylactis gigantea T: basitrich*/spirocyst Flaps/Cap Actinostola sp. MF: p-mastigophore* Cap Zoanthidea Protopalythoa mutuki T, MF: basitrich*, MF: spirocyst Cap Corallimorpharia Discosoma sp. MF: holotrich*, p-mastigophore* Cap Scleractinia Goniporia sp. T: p-mastigophore*/spirocyst Cap/cap Octocorallia Clavularia sp. A T: holotrich Cap? Medusozoa (Hydrozoa) Ectopleura larynx T: stenotele* Operculum 44 Sertularella sp. T: stenotele Operculum Medusozoa (Staurozoa) Haliclystus borealis T: eurytele, holotrich Operculum Body structure from which nematocysts came is indicated by letters in third column, C: column, MF: mesenterial filaments, T: tentacles. Asterisk (*) indicates that SEM data on that nematocyst were also collected.

Figure 2.1. Basic nematocyst morphologies identified with differential interference contrast light microscopy. See Table 2.1 for collection information. Scale Bar = 20 µm. A. b-mastigophore from tentacle of Ceriantheopsis americanus. B. Basitrichous isorhiza (B) and microbasic amastigophore (A) from acontia of Flosmaris mutsuensis. C. Microbasic p-mastigophore from tentacle of Gonipora sp. D. Microbasic p-mastigophore from mesenterial filament of Metridium senile. E. Discharged microbasic p-mastigophore from mesenterial filament of Metridium senile. F. Microbasic p-mastigophore from mesenterial filament of Bunodosoma cavernata. G. Microbasic p-mastigophore from mesenterial filament of Stomphia coccinea. H. Holotrichous isorhiza from tentacle of Protopalythoa mutuki. I. Desmoneme from tentacle of Neoturris breviconis. J. Euryteles from tentacle of Neoturris breviconis. K. Stenotele from tentacle of Ectopleura larynx. L. Holotrichous isorhizas (H) and eurytele (E) from tentacle of Haliclystus borealis. M. Spirocysts (S) and basitrichous isorhizas (B) from tentacle of Condylactis gigantea N. Ptychocysts from column of Ceriantheopsis americanus.

Figure 2.1

Figure 2.2. Scanning electron micrographs of apical structure in cnidae from across the phylum Cnidaria. A. Ovoid operculum of a basitrichous isorhiza from tentacle of Hydra sp. Note the rim into which the operculum fits (arrow). B. Triangular operculum of a stenotele from a tentacle of Hydra sp. C. Apical flaps of an unidentified nematocyst from a tentacle of Bartholomea annulata. Note the ridge visible on one flap (arrow). D. Apical flaps of a basitrichous isorhiza from a tentacle of Halcurias levis. E. Apex of a microbasic p-mastigophore of a mesenterial filament of Budodosoma cavernata. F. Apical flaps of a microbasic p-mastigophore of a mesenterial filament of Metridium senile. G. Apex of a microbasic b-mastigophore from a tentacle of Cerianthopsis americanus. H. Apex of a basitrichous isorhiza from a tentacle of Antipathes grandis. I. Apex of a holotrichous isorhiza from a tentacle of Renilla mulleri. J. Apex of a microbasic p-mastigophore from a tentacle of Platygyra astreiformis. Note the thin collar of capsular material that surrounds the tubule (arrow). K. Apex of a holotrichous isorhiza from a tentacle of Platygyra astreiformis. L. Apex of a holotrichous isorhiza from a mesenterial filament of Discosoma sp. Note the cracks in the apical end of the capsule (arrow). M. Apex of a holotrichous isorhiza from a tentacle of Protopalythoa mutuki. N. Apex of a basitrichous isorhiza from a tentacle of Protopalythoa mutuki. Note the roughly triangular structure (T) and the crack in the capsule wall (arrow). O. Apex of a ptychocyst from the column of Cerianthopsis americanus. (O operculum, F apical flap) Scale Bars = 2 µm (A, C-H, J-N), 5 µm (B, O), 1 µm (I).

Figure 2.2

Figure 2.3. Transmission electron micrographs of apical structure in cnidae across Cnidaria. A. Operculum of a stenotele from a tentacle of Ectopleura larynx. B. Operculum of eurytele from a tentacle of Haliclystus borealis. C. Partially discharged holotrichous isorhiza from a tentacle of Haliclystus borealis. D. Apical flaps of a basitrichous isorhiza from a tentacle of Condylactis gigantea. Striated material (arrow) lines the central space where the flaps meet. E. Apical cap of a microbasic p- mastigophore from a mesenterial filament of Actinostola sp.. Note the central mound of cap material (arrows). F. Apical cap of a basitrichous isorhiza from a tentacle of Protopalythoa mutuki. G. Apical cap of a b-mastigophore from a tentacle of Cerianthopsis americanus. H. Apical cap of a basitrichous isorhiza from a tentacle of Gonipora sp. I. Apical cap of a holotrichous isorhiza from a mesenterial filament of Discosoma sp. J. Apical cap of a microbasic p-mastigophore from a mesenterial filament of Discosoma sp. K. Apical structure of a holotrichous isorhiza from a tentacle of the octocoral Clavularia sp. A. L.Apical cap of a spirocyst from a tentacle of Condylactis gigantea. M. Mat (M) of apical material of a ptychocyst from the column of Cerianthopsis americanus. (O operculum, F apical flap, W capsule wall, T tubule wall, S spines, C cap, asterisks edges of the apical cap). Scale Bars = 500 nm.

Figure 2.3

Figure 2.4. Phylogenetic interpretation of apical structures. A. Distribution of apical structure in cnidae within Anthozoa. Nematocysts have three forms of apical structure whereas both spirocysts and ptychocysts have one one kind, the apical cap, which they share with nematocysts. B. Distribution of apical structure in nematocysts within Cnidaria. Apical flaps and operculum are synapomorphies for subsets of the phylum (Actiniaria and Medusozoa respectively), whereas the apical cap is ancestral in Anthozoa.

Chapter 3: Gross morphology and ultrastucture of anthozoan nematocysts

Introduction

Given the wide variety of uses in cnidarian biology and the phylum-wide distribution, the diversity of nematocysts is unsurprising. To make sense of the extensive morphological diversity of nematocysts, Weill (1930, 1934) created a classification system that, with some modification (Carlgren 1940, Mariscal 1974) remains the primary classification system used today. Confusion has arisen over the identity of various kinds of nematocyst because other classification systems have also been developed (e.g.

Stephenson 1929, den Hartog 1980, Schmidt 1969), and the categories of each system contain different assemblages of morphologies (England 1991). The authors of these systems disagree about which aspects of nematocyst morphology are most important for classification and differ in the interpretation of several aspects of nematocyst morphology. These differences also result in differing numbers of morphologies recognized, as some classifications subdivide smaller morphological variations than others (England 1991).

Among types of nematocysts, mastigophores, i.e. those nematocysts with a thickening at the base of the tubule (usually called a shaft), have been important in taxonomy and phylogenetic assessment for Hexacorallia (Carlgren 1940, 1945, Cutress

1955, Schmidt 1969, 1974). For example, Daly et al. (2003) found that within

Hexacorallia, large groups are defined by the presence of two particular mastigophores.

However, the categorization of mastigophores is the most controversial aspect of Weill’s

(1930, 1934) classification system and the element most revised by subsequent authors

(see England 1991 for a summary). Because mastigophores are important to hexacorallian systematics, it is imperative to clarify the distinction between and among mastigophores and to determine if any classification system consistently captures the underlying morphological variation.

Historical perspective on the classification of mastigophores

The classification of nematocysts created by Weill (1930, 1934) was among the first and most comprehensive classification system and a slightly modified version of it is the most widely used today (see Fig. 3.1 for a summary). After dividing nematocysts into those that have an opening at the tip from those that do not, he separated the former nematocysts into two categories: haplonemes, which have no swellings or thickenings along the tubule (e.g. Fig. 3.2A), and heteronemes, which have a thickening of the basal tubule, i.e. a shaft (e.g. Fig. 3.2D-I). Weill (1930, 1934) further subdivided the heteronemes: rhabdoides that have an isodiametric shaft, whereas rhopaloids that have bulges or extra thickenings along the shaft. Anthozoans have only the former, whereas medusozoans have great diversity in the latter (Mariscal 1974). Rhabdoides are further subdivided by Weill (1930, 1934): mastigophores have a tubule that continues from the shaft (e.g. Fig. 3.2E) and amastigophores lack a tubule (e.g. Fig. 3.2H-J). Carlgren (1940)

53 noted that mastigophores could be subdivided further yet: undischarged p-mastigophores

(and amastigophores) have a v-shaped notch at the end of the shaft (Fig. 3.2 F, H) and b- mastigophores have a blunt end to the shaft (no v-shaped notch) (e.g. Fig. 3.2 D, E). The notch is formed from the shaft folding in on itself in the undischarged capsule (Gosse

1860, Bedot 1896, Moebius 1866, Cutress 1955, Tardent 1988) as the shaft is tightly packed into the capsule.

These distinctions notwithstanding, detailed microscopic investigations have shown that amastigophores do have a tubule: it is short and usually breaks off during discharge (Cutress 1955, Schmidt 1969, Östman 2000) (e.g. Fig. 3.2I). Thus, there is no difference between amastigophores and p-mastigophores as currently defined: all heteronemes have a tubule distal to the thickened shaft. Nonetheless, the relationship between amastigophores and mastigophores is controversial. Cutress (1955) and Schmidt

(1969) proposed that amastigophores are a particular kind of p-mastigophore. Östman

(2000) concurred that the p-mastigophores and amastigophores were similar, and suggested calling amastigophores p-amastigophores to signify that these have a short distal tubule that often breaks off the shaft, in contrast to the long distal tubule that remains attached to the shaft of a p-mastigophore. Subsuming amastigophores within p- mastigophores implies that the two types are morphologically similar and differ only in the length of the distal tubule. England (1991) proposed maintaining the two categories to clearly identify the different morphologies.

Rather than combine mastigophores into fewer categories, Schmidt (1969) subdivided p-mastigophores into several new categories. Unfortunately, Schmidt’s

54 system requires examination of discharged nematocysts (England 1991) and so does not correspond to the distinctions made by Carlgren (1940) and subsequent workers based on the morphology of undischarged capsules. Furthermore, because most cnidarian biologists identify nematocysts from the undischarged (and un-dischargeable) capsules of preserved specimens, Schmidt’s (1969) subcategories of p-mastigophore are often not used. Therefore, whether this finer subdivision captures the underlying variation in nematocyst morphology has yet to be fully explored.

A related issue pertains to the other kind of mastigophore, called a b- mastigophore by Carlgren (1940) (Fig. 3.2D, E). This nematocyst resembles another type, the basitrichous isorhiza (basitrich) (Figs. 3.2B, C). A basitrich lacks a shaft but has a tubule with large proximal spines (Fig. 3.2C). Although he distinguished between them,

Carlgren (1940) acknowledged that undischarged basitrichs and b-mastigophores resemble each other: when folded inside the capsule, the dense spines on the proximal tubule of basitrichs look like a thickened shaft (see Fig. 3.3 A-C). Although Cutress

(1955) noted that most nematocysts classified as basitrichs have a widened, rigid proximal tubule and should therefore be called b-mastigophores, he did not formally reject basitrich as a category because he recognized that some nematocysts classified as basitrichs do not have a rigid, widened proximal tubule. Schmidt (1969) completely rejected basitrich as a category, arguing that a larger diameter of the tubule (i.e. shaft) is a direct result of having larger spines on the proximal tubule (Schmidt 1969, 1974).

Therefore, any nematocyst with large spines at the base must have a corresponding

55 widening of the tubule. England (1991) stated that the two categories have a distinct morphology when discharged and should be maintained.

The shaft and its defining features are fundamental to the distinguishing of basitrichs from b-mastigophores. Weill (1930: 145) defined the shaft as an expanded segment of the proximal tubule. In p-mastigophores or amastigophores, the proximal tubule is much wider than the distal tubule; however, in b-mastigophores, the change can be more gradual, even as small as 0.1 µm (Cutress 1955). If, as Cutress (1955) and

Östman (1983, 1987, 2000) suggest, a distinct shaft is not present, then there is no clear distinction between a basitrich and a b-mastigophore.

To clarify the differences and similarities of the nematocysts variously classified as basitrichs, b-mastigophores, p-mastigophores, and amastigophores, I have conducted a careful survey of these morphologies from exemplars across Hexacorallia. In the process, features of the shaft have been closely examined to determine the exact nature of this character. These investigations enable me to evaluate how effectively previous classification systems for nematocysts capture the diversity of form among

“mastigophore” nematocysts.

Materials and Methods

Materials

I examined nematocysts from species spanning Anthozoa: 27 actiniarians, 2 antipatharians, 3 cerianthids, 2 corallimorpharians, 2 scleractinians, 2 zoanthids, and two octocorallians (Table 1). Additionally, nematocysts from four hydrozoan taxa were

56 studied for comparison of ultrastructure and morphology. These specimens were collected from various localities (Table 1) or obtained from commercial sources such as

Aquarium Adventure (Columbus, OH), Gulf Specimen Marine Lab (Panacea, FL),

Marine Biological Lab (Woods Hole, MA) and Reef Hot Spot (Inglewood, CA). Two samples, the stoloniferan octocoral Clavulariid sp. A (sensu Parrin et al. 2010) and

Protopalythoa mutuki, were obtained from cultures maintained by Neil Blackstone at

Northern Illinois University and the Hydra sp. was obtained from a culture maintained by

Paulyn Cartwright at University of Kansas. The antipatharian samples were collected, fixed, and provided by Anthony Montgomery (see Opresko, 2009 for collecting details).

Species identifications were determined by the collector or verified in the case of commercial sources for all but three specimens. One sea anemone was not fully identified beyond belonging to the Acontiaria; it is listed as Acontiaria sp. in tables and figures and another sea anemone is identified only to genus and listed as Actinostola sp. Finally, one species of cerianthid remains unidentified, but is known to belong to the family

Cerianthidae (listed as Cerianthidae sp. in tables).

Methods

After collection, most samples were dissected to isolate the key body regions: mesenterial filaments, body column, tentacles, and acrorhagi or acontia if present (see Table 1 for summary of collections). After dissection, samples for SEM and light microscopy were placed in a 1M sodium citrate solution for 10-15 minutes (larger tissues required more time) to induce expulsion of nematocysts from nematocytes. Samples were washed three

57 times with distilled water and most were then placed in 70% ethanol. Samples from

Bunodosoma cavernata and Metridium senile were placed in a 1% OsO4 solution overnight before being placed in 70% ethanol. SEM samples were dehydrated in ethanol, then critical-point dried with CO2. Samples from B. cavernata and M. senile were sputter-coated with gold-palladium in a Hummer sputter coater and examined using a

LEO 1550 field emission scanning electron microscope at the University of Kansas,

Lawrence. All other samples were sputter-coated with gold-palladium or palladium in a

Cressington sputter coater and examined using a FEI NOVA nanoSEM at the Ohio State

University, Columbus. Squash preparations of undischarged capsules were made using samples fixed for SEM as above, but before critical-point drying, and maintained in 70% ethanol. A very small piece of tissue was floated in a droplet of water and then compressed between a coverslip and a microscope slide; squash preparations were examined under DIC at 1000X.

Some samples were also prepared for TEM (Table 1). After dissection, these specimens were fixed in 2.5% glutaraldehyde in phosphate buffer at pH 7.4 and postfixed in 1% OsO4 in phosphate buffer. All specimens were dehydrated in a graded ethanol series and embedded in Epon. Thin sections cut on a Leica EM UC6 Ultramicrotome were stained with 2% aqueous uranyl acetate for 15 min followed by 5 min in Reynold’s led citrate. Micrographs were taken on a Technai G2 Spirit TEM at the Ohio State

University, Columbus.

Nematocyst morphologies were binned into like categories based on both light and electron (SEM and TEM) microscopy data. Morphologies were named using

58 established nomenclature when previous, unambigious names exist. For morphologies with confusing or misleading nomenclature, new names were assigned. In assigning new names, the greek word neme, from the root nema, meaning thread, was used to avoid using trich, tele, mastigophore or rhabdoid. These names might unintentionally imply a relationship between the newly named morphologies and previously named ones. All new names were based on features of the basal tubule (or thread). See results for details on specific names for morphologies.

Results

The nematocysts I studied (basitrichs, b-mastigophores, p-mastigophores, and amastigophores) using various microscopy techniques could be separated into basic morphologies with some variation within these categories. A summary of the features of each morphological type as seen by light and scanning electron microscopy are summarized below. For five of these morphologies, transmission electron microscopy was also available; for these TEM investigations, I use holotrichous isorhizas

(holotrichs), which are not the focus of this study, for comparison. In holotrichs, the tubule is uniform throughout and therefore, unlike the other morphologies detailed here, there is no obvious and distinctive tubule descending from the apex (Fig. 3.3A, B). All the other morphologies discussed here (Fig. 3.3C-P) have a basal tubule that descends from the apex and that has features that distinguish it from the distal tubule.

Nematocyst morphologies observed

Morphology 1: Acanthoneme

Distribution: The acanthoneme occurs in the hexacorallian orders Actiniaria,

Antipatharia, and Zoanthidea, and can be found in all body regions of a polyp.

Capsule shape and size: In its undischarged state, the acanthoneme has an elongated, symmetrical capsule (Figs. 2B, 4A, B). Usually the capsule is broadest at its midsection, tapering evenly at each end (Fig. 3.4A, B). In some cases, the capsule is more ovoid than elongate (Fig. 3.4C). These range in size from a length of 11.8-35.9 µm and a width of

1.6-3.8 µm.

Undischarged tubule: A thickened portion of the tubule descends down the middle in the

undischarged state (Fig. 3.4A, B). From the apex, the tubule descends almost the entire

length of the capsule and has a blunt end to its visible portion (i.e. no v shaped notch at

the distal end). Although always clearly visible, the central tubule is sometimes not very

robust or thick (Fig. 3.4B) and is not always smooth under the light microscope (Fig.

3.4A). Furthermore, the central tubule sometimes narrows along its length from the apex

and may not remain in the center of the capsule (Fig. 3.2B). Therefore, in its

undischarged state, this morphology varies with respect to shape or size for the thickened,

descending tubule.

Ultrastructural features of undischarged capsule: An ultrastructural view of the basal portion of the acanthoneme reveals a tubule that contains long spines (Fig. 3.3B). These long spines are collapsed tightly against each other and begin immediately beneath the apex (Fig. 3.3C-F). The tubule encases these spines and follows the contour of these

60 spines closely. In some cases, no extra projections of the tubule surrounding these spines are evident (Fig. 3.3C), or very few (Fig. 3.3D). In some sections, a few longer projections of the tubule are evident, however these are widely spaced apart (Fig. 3.3E).

Furthermore, regions between these periodic projections have a smooth tubule which follows the cylinder formed by the packaged spines with no evident bumps or folds (Fig.

3.3F).

A distal tubule is always present and is evident to either side of the descending tubule

(Fig. 3.3C-F) and at the bottom of the capsule (Fig. 3.3E). Under light microscopy, the distal tubule is sometimes visible inside the capsule (Fig. 3.2B), but often is not (Fig.

3.4A, B).

Discharged tubule: After discharge, the tubule can be divided into a basal portion and distal portion that differ primarily in spine pattern (see below). The length of the basal portion of the tubule is typically very close to the length of the capsule, often being only slightly shorter or longer than the length of the capsule and can therefore be quite long

(about 40 µm in Fig. 3.4D), or quite short (about 8 µm in Fig. 3.4E).

A distal tubule continues beyond this basal region (Figs. 3.2C, 3.4D, E, I). In the transition from the basal to distal portions, the tubule narrows slightly so that the distal tubule is smaller in diameter than the basal portion (Fig. 3.4I). No other changes in the tubule are commensurate with this transition, indicating continuity between the basal and distal portions.

Spines and spine attachment: In the discharged state, the acanthoneme has long spines on the basal tubule which typically flare outward and are visible under both light microscopy

(Fig. 3.2C) and SEM (Fig 3.4D, E). The basal spines vary in length and width but are

always longer than the diameter of the basal tubule and are usually more than 1 µm in

length (Fig. 3.4E-G). Basal spines are wider at the base and narrow evenly to a point

(though the degree to which they narrow varies, compare Fig. 3.4F-G). Often the spines

curve up (in the direction of the distal tubule), seemingly as a result of this narrowing

(Fig. 3.4G). The attachment of these spines to the tubule is straight, with no folds in the

tubule accompanying this attachment (see spine scars in Fig. 3.4H). In fact, the whole

tubule of the basal region is smooth in fully discharged nematocysts, with no apparent

folds or annulations in the surface (Fig. 3.4H-I).

Spine pattern changes at the transition from basal to distal tubule. The spines

become shorter and smaller and abruptly stop (Fig. 3.4I). Shortly thereafter, a new type of

spine appears and continues along the entire length of the distal tubule (Fig. 3.4J, K).

These distal spines differ in size from the basal spines, being less than 1 µm in length,

narrow and somewhat hook-like in shape (Fig 3.4K), though they are never very thick in

cross section (Fig. 3.4J, K). The points of these spines are oriented toward the capsule

rather than toward the end of the tubule (Fig. 3.4J, K).

Apical structure: Actinarian nematocysts of this morphology have apical flaps (Figs.

3.3C, D, 3.4L), whereas nonactinarian ones have an apical cap (Figs. 3.3E, F, 3.4M).

Name: This morphotype has long spines on the basal tubule that flare outward giving a bottle brush appearance. Therefore, it is given the new name acanthoneme based on the root word akantha meaning thorn or spine.

Morphology 2: Colloponeme

Distribution: The colloponeme occurs only in actiniarians, primarily in the acontia of

some species; occasionally it occurs in the tentacles, although only in those species that

also have them in the acontia (e.g. Metridium senile).

Capsule shape and size: This morphology consists of a capsule (Fig. 3.5A, B) similar in

basic form to that seen in the first morphology described (Fig. 3.4A, B): elongate,

symmetrical, and broadest at its midsection (Fig. 3.5A, B). These range in size from a

length of 20.1-37 µm and a width of 1.3-2.78 µm.

Undischarged tubule: A thickened portion of the tubule descends down the middle in the

undischarged state, is robust and consistent in shape, and fills almost the entire capsule

(Figs. 3.5A, B, 3.3G). Sometimes the tubule shows evidence of being folded within the

capsule (Fig. 3.5C). The tip of descending tubule either appears to end bluntly (Fig. 3.5A)

or with a point (Fig. 3.5B), but never with a v-shaped notch. The distal tubule is

sometimes apparent in the capsule (Fig. 3.5A).

Ultrastructural features of undischarged capsule: An ultrastructural view of the colloponeme reveals a tubule that contains spines (Fig. 3.3G-H). However, the spines of the basal tubule do not begin immediately beneath the apex (Fig. 3.3H). Rather, folds of the tubule are always evident directly underneath the apical flaps, which are always present (Fig. 3.3H). About 1-3 µm beneath the apex, short spines arise: the end of the spines that is attached to the tubule is not tightly packed against other spines and each spine flares outwards at the edges of the capsule (Fig. 3.3H). Therefore, the tubule is not smooth against the cylinder of spines but instead has a frilly, highly folded appearance

(Fig. 3.3H). These folds of the tubule never form long extentions (such as in Fig. 3.3F), but remain tight against the spines (Fig. 3.3H). The distal tubule is evident in the space between the basal tubule, which is centered in the capsule, and the capsule wall (Fig.

3.3G, H).

Discharged tubule: In the discharged state, the tubule can be divided into two sections on the basis of spine (see below) and tubule surface differences (Fig. 3.5D-F). The basal portion of the tubule ranges from about the same length (Fig. 3.5D-G) to much longer

(Fig. 3.5H) than the capsule. Furthermore, the tubule of this region is not smooth, but rather has annulations that encircle the diameter of the tubule (Fig. 3.5I). These annulations are visible even if the spines are flush against the tubule (Fig. 3.5J). This basal region sometimes discharges with kinks rather than straight (Fig. 3.5H).

A distal tubule continues from this basal portion. At the transition, the tubule does not change in size but remains isodiametric (Fig. 3.5K). However, the regular annulations or folds stop (Fig. 3.5K, L); unlike the basal tubule, the distal tubule is smooth (Fig.

3.5L). There is no evidence for narrowing of the distal tubule: even further along its length, the distal tubule maintains the same diameter as the basal tubule (Fig. 3.5L, M).

Spines and spine attachment: In the discharged state, the colloponeme has spines on the basal tubule (Fig. 3.5D-F, H), and these sometimes remain tightly associated with the tubule (Fig. 3.5E, J). The spines of the basal tubule are close in size to the diameter of the tubule, typically being under 1 µm. Each is slightly wider at the base than the tip, however the spine only narrows at the very tip (Fig. 3.5I) and although the very edges may curve up slightly, the whole spine does not curve inward but rather typically lays flat

against other spines (Fig. 3.5J, K). The base of each spine of the basal tubule is attached

in the folds created by the regular annulations in the surface.

At the transition from basal to distal tubule, the spines also change in shape and

size from a longer, broader, flatter spine to a shorter, conical, thick spine (Fig. 3.5L, M).

Spines of intermediate length and thickness are evident between the two final shapes and

sizes (Fig. 3.5L). Further along the distal tubule, these spines have a thick, knobby profile

but are short (Fig. 3.5L, M). These short, thick spines continue until the end of the distal

tubule.

Apical Structure: Nematocysts of this morphology always have apical flaps (Fig. 3.5N).

Name: This morphology has spines that lie in close association against the basal tubule after discharge creating a screw-like pattern. Therefore, it is given the new name colloponeme based on the root kollopos meaning peg or screw.

Morphology 3: Hadroneme

Distribution: The hadroneme is seen only in cerianthids but is found in all three tissues

studied (tentacles, mesenterial filaments, column).

Capsule shape and size: The shape of the undischarged capsule can vary, sometimes

being more elongate (Fig. 3.6A), and other times being more ovate (Fig. 3.6B). In both

cases, the capsule is symmetrical and typically broadest at its midsection. In the taxa

studied here, the capsules range in size from 14.5-30.3 µm in length and 1.55-3.6 µm in

width.

Undischarged tubule: A thick descending tubule with an obvious spiral pattern that runs

down its length is always evident (Fig. 3.2D, 6A, B). The thickened portion of the tubule

usually is more than half as long as the capsule (Figs. 3.2D, 3.6A, B) and narrows distally

(Fig. 3.2D). No v-shaped notch is ever evident in the transition from proximal to distal

tubule, just a narrowing of the diameter of the tubule (Figs. 3.2D, 3.6A, B). A distal

tubule is sometimes evident in the capsule (Fig. 3.6B).

Ultrastructural features of undischarged capsule: An ultrastructural view of the hadroneme reveals long spines encased by the descending tubule for most of the length of the capsule (Fig. 3.3O). These spines begin almost immediately beneath the apical cap

(Fig. 3.3P). The ends of the spines that are attached to the tubule do not lie flat against each other, but flare out towards the edges of the capsule (Fig. 3.3P, Q). The tubule may have long extensions (Fig. 3.3P). In between spines that flare out, extra folds can be seen in the tubule (Fig. 3.3Q). Even if these longer extensions are not evident (as in Fig. 3.3Q), smaller folds in the tubule are evident in-between insertions of the spines. The distal tubule is sometimes evident at the bottom of the capsule (Fig. 3.3O).

Discharged tubule: In discharge, the hadroneme has a basal tubule much larger in

diameter than the distal tubule (Fig. 3.6C, D). The basal tubule is about the length of the

capsule. Folds are evident in the tubule surface at the points of attachment of the spines

(see below), but the tubule is fairly smooth elsewhere.

Spines and spine attachment: The basal tubule bears large spines (Fig. 3.6C, D). These

spines have a broad base, narrowing to form the blade of the spine, giving the whole

spine a T shape. The head of the T of each spine is embedded in a fold in the tubule. The

blade of the spine narrows gradually, forming a tip that typically points back towards the

capsule (Fig. 3.6C). However, in some capsules, the tips of these long spines curl upward

(Fig. 3.6D). The sides of these spines also curl inward, giving them a central groove (Fig.

3.6E). These spines shorten in length at the transition between basal and distal tubule and

the entire transitional region bears these spines (Fig. 3.6F). The distal tubule has very

small spines that lie close to the surface of the tubule, but seem flat in profile (Fig. 3.6G).

Apical Structure: The hadroneme always has an apical cap rather than flaps (Fig. 3.3P).

Name: This morphology has a stout, thick basal tubule with a blunt end usually visible under the light microscope. Therefore, it is given the new name hadroneme from the root hadros meaning well-developed, bulky, stout, large.

Morphology 4: p-rhabdoid A

Distribution: This p-rhabdoid A occurs primarily in the mesenterial filaments of all orders

of Hexcorallia except Ceriantharia. It is also found in the tentacles of corals, zoanthids

and a few actinarians.

Capsule shape and size: In actiniarians, the p-rhabdoid A consists of capsule that is wider

at the non-apical end (Fig. 3.7A, B). Capsule shape varies: in some taxa, the capsule is

more elongate (Fig. 3.7A), whereas in others it is more ovate (Fig. 3.7B). In non-

actiniarians, the capsule wall is generally thicker and more elongate (see Fig. 3.2D).

Regardless of shape, the capsule wall is thin and the capsule itself sometimes contains

some substance (Fig. 3.7A). In the taxa studied here, the capsules range in size from 15.6-

26.0 µm in length and 1.88-4.1 µm in width.

Undischarged tubule: The thickened basal portion of the tubule is corkscrew-like in its

undischarged state (Fig. 3.7A, B). The basal tubule ends in a v-shaped notch (Fig. 3.7A)

from which a thin distal tubule emanates (Fig. 3.5B); this distal tubule is not always be

evident with light microscopy of undischarged capsules (Fig. 3.7A). In non-actinarians,

the notch is often more pronounced and the distal tubule thicker and more substantial

(Fig. 3.2D). In both actinarian and non-actiniarian nematocysts of this type, the basal

tubule descends to about the midpoint of the capsule and its proximal end attaches to the

capsule just beneath the capsule apex.

Ultrastructural features of undischarged capsule: An ultrastructural view of this morphology reveals a descending tubule encasing large spines (Fig. 3.3I). These spines typically begin directly beneath the apical cap (Fig. 3.3J, K). The end of a spine that is attached to the tubule does not lie flat against other such spines and its point of attachment correspond to a bump or fold in the basal tubule (Fig. 3.3J, K). The tubule has long, regularly-spaced extensions that are closer together than those of the basal tubule of morphology 1 (compare Fig. 3.3I-K to Fig. 3.3E, F). In between these longer tubule extensions, the tubule has small, regular folds corresponding to the attachment of spines

(Fig. 3.3J, K, see below). The distal tubule is not evident to the sides of the descending tubule (Fig. 3.3J, K) but is evident at the end of the capsule (Fig. 3.3I).

Discharged tubule: In the discharged state, the p-rhabdoid A has a basal tubule that is

much thicker than the distal tubule (Fig. 3.7C); this basal tubule also bears spines. The

basal tubule is equal to or slightly longer in length than the capsule. At the distal end, the

basal tubule narrows dramatically, the large spines taper in size until they cease,

68 concomitant with the change of diameter of the tubule (Fig. 3.7F). This change in diameter and spination marks the transition from basal to distal tubule (Fig. 3.7G).

Spines and spine attachment: Long spines are attached to the thick basal tubule (Fig.

3.7C). The flaring spines are perpendicular to the surface of the tubule (Fig. 3.7C). An individual spine has a broad base, but narrows immediately to form a thin long spine in the shape of a T (Fig. 3.7D), which slowly tapers to a point (Fig. 3.7D, E). Each spine attaches to the tubule by inserting its broad base (the head of the T) into a fold in the tubule (Fig. 3.7F). These folds do not encircle the basal tubule (Fig. 3.7F) and smooth regions can be seen on the surface of the tubule in between rows of spines (Fig. 3.7C-F).

Spines taper at either end of the basal tubule and therefore are shorter at the beginning (Fig. 7D) and at the end of the basal tubule (Fig. 3.5G). Scleractinian nematocysts of this morphology have spines on the distal tubule that are small, thin, and hook-like with the point oriented towards the capsule (Fig. 3.7H). Nematocysts of this type in actiniarians do not appear to have spines on the distal tubule.

Apical Structure: Apical flaps are never present in the p-rhabdoid A, even when in actiniarians (Figs. 3.3K, 3.7E).

Name: This morphology matches the description given by Schmidt (1969). Therefore his name, p-rhabdoid A is maintained.

Morphology 5: p-rhabdoid B1a

Distribution: This p-rhabdoid B1a is found only in actiniarians, and is found primarily in the mesenterial filaments and occasionally in the body column.

Capsule shape and size: The undischarged capsule has a distinctive teardrop shape in which the non-apical end is broadest (Fig 3.8A, B). The capsule is symmetrical and usually small, typically between 10-20 µm. The size range of capsules studied here is 8.0-

20.9 µm in length and from 1.7-2.6 µm in width.

Undischarged tubule: The basal tubule extends more than half-way down the capsule and often spans the entire length of the capsule (Fig. 3.8A, B). The apical end of the basal tubule forms a point (Fig. 3.8A); the tubule widens as it descends through the undischarged capsule and bears a v-shaped notch at its end (Fig. 3.8A, B). In some nematocysts, a distal tubule is evident (Fig. 3.8A), while in others it is not (Fig. 3.8B).

Discharged tubule: In discharge, the basal tubule has two portions, a much thicker basal portion with large spines (see below) and a thinner distal portion. In discharge, the basal tubule is about the same length as the capsule (Fig. 3.8C, D) and has a wide diameter

(Fig. 3.8C) before narrowing at the distal tubule (Fig. 3.8D). In nematocysts where the spines have been disrupted, annulations are evident on the tubule surface between whorls of spines (Fig. 3.8F). These annulations are regularly spaced and encircle the entire basal tubule. Distally, the tubule narrows (Fig. 3.8G) as it transitions from the basal to distal tubule (Fig. 3.8H). The distal tubule is much narrower than the basal tubule, has no annulations, and is unarmed (Fig. 3.8H). A distal tubule may not always be evident (Fig.

3.8I) but this fact may be the result of incomplete discharge rather than evidence that the tubule consistently breaks off during discharge.

Spines and spine attachment: Long spines that project towards the capsule are densely packed on the basal tubule typically obscuring the view of the tubule surface (Fig. 3.8E).

The spines appear to have edges that curl upward forming an indentation down the center

of the spine (Fig. 3.8E). These long spines form an acute angle with the tubule surface

with the points oriented toward the capsule (Fig. 3.8D, E). Spines have a broader base

and narrow uniformly to the tip therefore not really forming a T shape (Fig. 3.8F, H).

They attach to the surface through the folds created by the annulations. The spines

shorten in the region of transition from basal to distal tubule until they stop abruptly (Fig.

3.8G, H). No spines are evident on the distal tubule.

Apical Structure: The p-rhabdoid B1a always has apical flaps (Fig. 3.8E, I).

Name: This morphology matches the description given by Schmidt (1969). Therefore his name, p-rhabdoid B1a, is maintained

Morphology 6: p-rhabdoid B2d

Distribution: This morphology is found only in actiniarians and is found in tentacles,

body column, and mesenterial filaments.

Capsule shape and size: Before discharge, this capsule is distinguished by its fairly

consistent size (usually just under or about 20 µm) and a asymmetrical appearance

created by one side of the capsule curving out more than the other (Fig. 3.9A, B). As a

result, the non-apical end of the nematocyst is the narrowest of the capsule in both

undischarged (Fig. 3.9A, B) and discharged nematocysts (Fig. 3.9C, D). Size ranges from

9.5-24.1 µm in length, and 1.5-3.6 µm in width.

Undischarged tubule: Just beneath the apex in undischarged nematocysts, the thickened

descending tubule is narrow but it widens towards the middle of the capsule before

71 ending in a v-shaped notch in the bottom third of the capsule (Fig. 3.9A, B). No distal tubule is clearly evident in undischarged nematocysts (Fig. 3.9A, B).

Discharged tubule: In discharge, the basal tubule is at least as long as the capsule (Fig.

3.9C), though usually longer (Fig. 3.9D), and it consists of two regions distinguished by diameter of the tubule. The surface of the basal tubule has folds or annulations that encircle the entire tubule (Fig. 3.9E) and are evident at the base due to the sparsity of large spines in this region (Fig. 3.9E, F). The annulations of the more distal portion of the basal tubule (Fig. 3.9H, I) appear similar in size and spacing to those of the more proximal portion. In some cases, as the basal tubule narrows distally, the terminal tubule present breaks off, rendering the terminus of the tubule cone-shaped (Fig. 3.9H). This cone never bears spines. In other cases, a terminal tubule far narrower than the basal tubule remains attached (Fig. 3.9I).

Spines and spine attachment: The spines of the apical-adjacent region are narrow and typically oriented so that they point towards the capsule (Fig. 3.9F) but are easily ripped off the surface (Fig. 3.9E). This short region of the basal tubule has sparse spines, and transitions abruptly to a region with longer, though still narrow, spines (Fig. 3.9E, F).

These longer spines each taper to a point, tend to curl up at the free edges to form an indentation or groove down its center of the blade (Fig. 3.9G), and attach to the tubule with a slightly wider base in a fold created by the annulations (Fig. 3.9F, H, I). The terminal tubule is unarmed (Fig 3.9I).

Apical Structure: This nematocyst morphology always has apical flaps (Fig. 3.9E).

Name: This morphology and the following both are very similar to the description given by Schmidt (1969) for p-rhabdoid B2a. However, I find that this category can be split into two based on light and scanning electron microscopy into this category and the next one. Therefore the original name is modified to p-rhabdoid B2d. p-rhabdoid B2c has already been used by den Hartog (1995) to describe an isophellid nematocyst type.

Morphology 7: p-rhabdoid B2s (variant a and b)

Distribution: This nematocyst morphology occurs in all structures studied here of some

actinarians.

Capsule shape and size: In the undischarged state, the capsule is symmetrical, elongate,

and widest at its midpoint (Figs. 3.2H, 3.10A, B). Size ranges from 16.5-62.7 µm in

length, and 1.4-3.9 µm in width.

Undischarged tubule: The tubule has two clearly distinct regions, both of which can be seen just beneath the apex where a narrow inner and wider outer region can be distinguished (Fig. 3.10A). The descending tubule is at least two-thirds the length of the capsule (Fig. 3.10B) though it often fills almost the entire length of the capsule (Fig.

3.10A), and ends in a v-shaped notch. A small length of distal tubule may be visible from the notch (Fig. 3.2H), or may not be (Fig. 3.10A, B).

Although only oblique sections of this morphology were obtained (Fig. 3.3L-N), some distinctive features are evident. The descending tubule consists of two parts (Fig.

3.3L). The region of the tubule directly beneath the apex is filled with highly folded tubule with no evidence of spines (Fig. 3.3M). Spines are present along the whole basal

73 tubule (Fig. 3.10C-E). Presumably these spines would be visible in a section of the center of the nematocyst. However, the extra folds of the tubule captured in oblique section imply that the portion of the descending tubule directly beneath the apex must not smoothly outline those spines but must have numerous extra folds. Distally, long spines are evident in the descending tubule (Fig. 3.3N). The tubule closely follows the contours of the spines in this region with larger folds in the tubule spaced regularly apart and smaller folds filling the regions of tubule in between the larger folds (Fig. 3.3N). No long extensions of the tubule are evident. No distal tubule is evident although a more electron- dense material often is present to one side of the descending tubule near the bottom of the capsule (Fig. 3.3L, N).

Discharged tubule: In discharge, the tubule has two distinct regions defined by differences in tubule diameter. The basal tubule is slightly (Fig. 3.10C) to greatly longer

(Fig. 3.10D, E) than the capsule and can be subdivided into two distinct regions (Figs.

3.2I, 3.10C-E). The relative length of these regions varies among nematocysts of these morphologies; in most cases the basal-most region is shorter than the distal region and is identified as variant “a” (Fig. 3.10C, D), however, in some cases this pattern may be reversed and is considered variant “b” (Fig. 3.10E). These two regions are marked by differences in spine size and pattern (see below) and by differences in the tubule surface.

In the apical-adjacent region, the basal tubule has annulations or folds that encircle the entire tubule and is often not smooth and even in its diameter, but rather has bulges (Fig.

3.10G). Some nematocysts have regular, circular indentations in between whorls of spines in this region (Fig. 3.10H). The tubule of the more distal region of the basal tubule

74 also has annulations that encircle the tubule, but they are closer together than those of the apical-adjacent region (Fig. 3.10J). Also, the tubule here is smoother and seems more rigid (i.e. no bulges are evident) (Fig. 3.10J).

In some nematocysts the terminal tubule remains attached (Fig. 3.10M, N), while in others it breaks off just leaving the conical end to the basal tubule (Fig. 3.10O). This small amount of distal tubule may be visible left behind in the capsule (Fig. 3.2I). Among nematocysts that retain the terminal tubule, the width and robustness of that terminal tubule varies, with some having a thinner, more malleable tubule with a smooth surface

(Fig. 3.10M, P), and others having a thicker, more rigid structure with some evidence of folds in the surface (though not annulations that encircle the tubule) (Fig. 3.10N, Q).

Spines and spine attachment: Spines of the apical-adjacent region are thin, and each has a slightly wider base that attaches to the tubule at the fold of an annulation (Fig. 3.10F-H).

Typically these spines are narrow and long (Fig. 3.10G), although not nearly as long as the spines of the more distal region, however, some spines are shorter (Fig. 3.10H).

Spines in this region are not densely packed (Fig. 3.10G, H). The transition to the more distal portion of the basal tubule is demarcated by an abrupt change from short spines to much longer spines (Fig. 3.10I). These longer spines are so densely packed that the tubule surface is often not visible (Fig. 3.10I). The spines on this part of the basal tubule have a wider attachment point, narrow to form a neck, and then widen again to form the blade of the spine; they are not T-shaped (Fig. 3.10J). These long, thin spines curl upward at the edges to form a groove down the center of the spine (Fig. 3.10K, L). These spines

become shorter as the basal tubule narrows into a spine-free conical end (Fig. 3.10M-O).

The terminal tubule is always without visible spines (Fig. 3.10P, Q).

Apical Structure: Nematocysts of this morphology always have apical flaps (Fig. 3.10F).

Name: This morphology is very similar to the description given by Schmidt (1969) for p- rhabdoid B2a and B2b (the distinction being the relative length of the two portions of the distal tubule). However, I find that this category can be split into two based on features detailed here. While one morph is designated the p-rhabdoid B2d (detailed above), this morphology is called the p-rhabdoid B2s, the “s” a reference to the fact that unlike the

2Bd form this one always has a symmetrical capsule. Individual nematocysts of this type are designated as variant a or b to maintain the original a/b distinction.

Morphology 8: Diakaneme

Distribution: Among the specimens sampled here, the diakaneme is only found in the

mesenterial filaments of Urticinia felina.

Capsule shape and size: These nematocysts are large in size, typically about 40 µm in

length before discharge. The capsule is elongate with the non-apical end being broader

than the apical end (Fig. 3.11A, B).

Undischarged tubule: A thickened descending tubule is narrow near the apical end (Fig.

3.11A), widening as it descends to the midway point of the capsule (Fig. 3.11A, B).

Sometimes this tubule has a distinct v-shaped notch (Fig. 3.11B), sometimes this notch is

less apparent (Fig. 3.11A).

Discharged tubule: The tubule is divisible into two regions, based on spine pattern (see

below) and tubule diameter. The basal tubule is smooth with no folds in the surface (Fig.

3.11D-F) though spine scars may be visible (Fig. 3.11G). The transition between basal to

distal portions of the tubule is marked by a slight narrowing of the tubule (Fig. 3.11G).

The distal tubule has a smaller diameter than the basal tubule and is unarmed (Fig.

3.11H).

Spines and spine attachment: In discharge, this form has long spines at the base of the

tubule, but none on the distal tubule (Fig. 3.11C). The region of the basal tubule nearest

to the capsule has narrow, smaller spines (Fig. 3.11D) whereas those further along the

basal tubule are wider, and larger (Fig. 3.11E, F). These wide spines flare outward to give

the nematocyst tubule a full look in discharge, although the spines are not usually

perpendicular to the tubule surface (Fig. 3.11C). Each of these spines has a wider base

and narrows uniformly to the tip, therefore distinctly not T-shaped (Fig. 3.11D-F). The

spine attaches to a tubule without any folds or annulations, and leaves a small spine scar

on a tubule stripped of spines (Fig. 3.11G). As the tubule diameter narrows to transition

from basal to distal tubule, the long spines become shorter and abruptly stop.

Apical Structure: This nematocyst morphology has apical flaps.

Name: This morphology has two types of spines on the basal tubule, shorter, thinner spines initially which transition to larger, thicker, more robust spines. Therefore, this morph is given the new name diakaneme from the root words di meaning two and akantho meaning spine.

Morphology 9: Aphylloneme

Distribution: This nematocyst morphology occurs only in the mesenterial filaments of some members of the family Actiniidae.

Capsule shape and size: The undischarged capsule ranges in length from 32.8-36.6 µm and in width from 2.5-3.6 µm. The capsule is elongate in shape with the non-apical end being distinctly wider than the apical end (Fig. 3.12A).

Undischarged tubule: A thickened descending tubule descends a little more than half-way down the capsule but never fills the whole length of the capsule. The basal tubule ends bluntly with no v-shaped notch apparent at the end of the basal tubule in the undischarged capsule (Fig. 3.12A).

Discharged tubule: In discharge, the tubule of the aphylloneme is divisible into two regions based on the presence of spines (Fig. 3.12B). The tubule surface is smooth with no folds or annulations that encircle the tubule (Figs. 3.12C-E). The transition from basal to distal tubule is not indicated by any obvious change in tubule surface or diameter (Fig.

3.12D). The tubule appears to be isodiametric for both basal and distal tubule with any changes in size being minimal (Fig. 3.12B, D).

Spines and spine attachment: The basal portion of the tubule bears spines (Fig. 3.12C), whereas the distal portion does not (Fig. 3.12E). This morphology seems particularly prone to the stripping of spines and consistently appears spineless along the entire length

(Fig. 3.12B). The spines are similar in shape and size for the whole length of the basal tubule and do not taper at the transition to the distal tubule (Fig. 3.12D). The spines are much longer than they are wide, giving them a skinny appearance, and taper gradually

only towards the end of the blade (Fig. 3.12E). The base of the spine is squared off rather

than t-shaped (Fig. 3.12F).

Apical structure: The aphylloneme has apical flaps (Fig. 3.12C).

Name: One of the most notable features of this morphology is its tendency to lose all its spines on the basal tubule. Therefore, this morphology is given the new name aphylloneme based on the root words a- and phyllos, together meaning without leaves.

Basal tubule of hydrozoan heteroneme nematocysts

The basal tubule of the two heteroneme types called rhopaloids commonly found in medusozoans reveal several commonalities. Both stenotele (Fig. 3.13A) and eurytele

(Fig. 3.13B) have a basal tubule of much larger diameter than the distal tubule. The basal tubule is not isodiametric, having bulges (e.g. 3.13B), and its surface is smooth, lacking regular folds or annulations, but in some instances having longitudinal furrows (3.13A,

B). Adjacent the capsule apex, the basal tubule is spineless; spines are situated on the bulge of the basal tubule (Fig. 3.13A, B). In these medusozoan heteronemes, a spine is wide at the base, tapering uniformly, to form a broad, short, and flat spine (Fig. 3.13A,

B). In addition to these spines of the basal tubule, a stenoteles has three thick conical spines, or stylets (Tardent 1988) basal to the more lamellar spines (Fig. 3.13A). The basal tubule can be clearly distinguished from the distal tubule in TEM (Fig. 3.13 C, D). The basal tubule is centered in the capsule, and in stenoteles, the region just beneath the apex contains the stylets, the bases of which are visible towards the posterior end of the capsule (Figure. 3.13 C, D). The longitudinal furrows of the basal tubule are visible under

TEM as folds just beneath the apex (Fig. 3.13D). The tubule expands in diameter to form

the bulge that bears the lamellar spines. Small, regular folds in the surface are evident at

the posterior end of the basal tubule (Fig. 3.13C, D). Near the end of the capsule, the

basal tubule dramatically changes in diameter as the tubule transitions from basal to distal tubule (Fig. 3.13 D). More distal tubule is evident on either side of the basal tubule (Fig.

3.13C, D).

Discussion

The nematocyst morphologies studied here (basitrichs, b-mastigophores, p-

mastigophores, and amastigophores) vary considerably. How authors categorize this

variation is fundamental to understanding how the various classification systems differ

and to determining which is most effective at summarizing this variation. Table 2

summarizes how the morphologies seen here relate to other classification systems.

Historical perspective on observed nematocyst morphologies

Acanthoneme

Under the Weill system, this morphology has generally been categorized as a basitrichous

isorhiza (Weill 1930, 1934, Mariscal 1974). By the original definition, a bastrich does not

have a shaft, and thus has no change in diameter between the basal and distal tubule

(Weill 1930, 1934, Carlgren 1940, Mariscal 1974). However, although there is no change

in the tubule surface, there is a slight narrowing of the diameter of the basal tubule as it

transitions to distal tubule (Fig. 3.4I), therefore the tubule is not isodiametric as required

80 by Weill’s (1930, 1934) definition. Furthermore, as other authors have noted (Cutress

1955, Westfall 1965, Schmidt 1969), the distal tubule is armed (= hoplotelic sensu Weill

(1934)). By definition, however, a basitrich only has spines on the base, or is anoplotelic

(Weill 1934). Therefore there are two problems with defining this morphology of nematocyst as a basitrich: the tubule changes in diameter and bears spines distally.

Although some authors have maintained usage of the name basitrich despite recognizing some problems with the definition (e.g., Cutress 1955, England 1991), Weill’s system includes a category that better describes this morphology: heterotrichous anisorhiza. By definition, a heterotrichous anisorhiza has a slightly dilated basal tubule that narrows to a distal tubule, and has different spines on the basal and distal tubules (Weill 1934).

In contrast to Weill (1930, 1934) and his followers (Carlgren 1940, Mariscal

1974), Schmidt (1969) contended that long spines define the tubule, not a change in diameter (see discussion on the nature of shaft below for more on this point). Therefore, he created the category b-rhabdoid for nematocysts of this type (Schmidt 1969); this descriptor also encompasses other morphologies (such as colloponemes, hadronemes, and aphyllonemes). Similarly, den Hartog (1977) also has one term for all the morphologies with a thickened basal tubule and no v-shaped notch, but he uses Stephenson’s (1928) term, spirula. However, combining this morphology with others (such as colloponemes and hadronemes) obscures the clear morphological distinctions among these morphologies visible under TEM and SEM (see below for more discussion on light microscopy vs. undischarged capsule data).

Considering the broad distribution of this morphology (it is found in Actiniaria,

Antipatharia, and Zoanthidea), the variation within the morphology is not surprising. The most variable aspect is the length of the basal tubule, although length of spines on the base can also vary. One distinction in actiniarian nematocysts of this type is apical flaps rather than a cap as the apical structure. However, this character is believed to be a synapomorphy for Actiniaria (Reft and Daly 2012) and is always present in nematocysts of this morphology in Actiniaria. Despite the variation, the construction remains the same even at the ultrastructural level, therefore these differences should be considered variation within the type rather than separate morphologies.

Colloponeme

As for the acanthoneme, Weil’s (1930, 1934) system would classify this type as a basitrich. Carlgren (1940) modified the Weill system because he perceived a type that superficially resembles basitrich in having a thickened descending tubule with no v- shaped notch, but which actually is a mastigophore with a shaft (defined by him as a basal tubule much thicker than the distal tubule). He termed this a b-mastigophore

(Carlgren 1940) and did consider the genus Metridium to have this type, however

Aiptasia, Cerus, and other genera that share colloponeme were not deemed to have b- mastigophores but rather basitrichs (Carlgren 1949).

Under Schmidt’s (1969) system, this morphology would be considered a b- rhabdoid like the acanthoneme. However, Schmidt (1969) notes that there is a b-rhabdoid type in the acontia of Aiptasiidae (and some mesenterial filaments of a few other

82 anemones) with a distinct morphology, a shorter, more dense armature, and an unusually short “filament.” The filament (i.e. distal tubule) is scarcely thinner than the shaft

(Schmidt 1969). He also notes that Metridium senile has a b-rhabdoid in the acontia in which the shaft is much longer than the capsule, though he describes the shaft as having two parts (Schmidt 1969). Stephenson (1928) also noted that the acontia of M. senile has a distinct morphology. Based on my investigations, I argue that these two variants of

Schmidt’s b-rhabdoides represent variation within what is described here as the colloponeme. The armature is short (compared to that of the acanthoneme), may appear dense because the spines are often sealed against each other in discharge, and no change in size occurs between basal and distal tubules (Fig. 3.5). I disagree with Schmidt’s

(1969) assessment that the basal shaft has two regions in M. senile, although it is true that the basal tubule is extremely long in this type. In M. senile, the nematocysts of this morphology resemble those found in the acontia of other sea anemones (such as Aiptasia sp. and Bartholomea annulata), differing only in the length of the basal tubule.

One detail of this morphology not previously described is the regular annulations or folds that encircle the basal tubule. These folds can be seen in TEM where the spines insert or attach to the tubule, and were documented by Westfall and Hand (1962) and

Westfall (1965). These folds resemble those characteristic of other nematocyst morphologies, particularly of the p-rhabdoid B1a and B2d forms in which the basal tubule has a similar folded pattern, although the spine pattern differs. The p-rhabdoid B2s also has similar folds on the basal tubule, though the pattern of folds changes with the spine pattern, which does not occur on the colloponeme. This may indicate an

evolutionary relationship between these morphologies or may just be a convergent

feature (discussed more below). These other morphologies differ from the colloponeme

in that they do have a v-shaped notch in undischarged form (this morphology does not),

and a distal tubule that is noticeably smaller in diameter than the basal tubule. Assessing

which features share an evolutionary history and which are convergent requires putting

these nematocysts in a phylogenetic context.

Hadroneme

This morphology was one that formed the basis of Carlgren’s (1940) argument for creating the category b-mastigophore, as he recognized that this morphology is distinct from what Weill (1930, 1934) described as a basitrich (here, acanthoneme). Schmidt

(1969, 1972), however, categorizes this nematocyst as a b-rhabdoid, grouping it with the acanthoneme and colloponeme. Clearly this morphology is distinct from these morphs as it has spines on the basal tubule that are more reminiscent of spines in a similar position in p-rhabdoid A (with a T-shaped head and obvious folds in the tubule surface at the point of attachment). Furthermore, the ultrastructure of the tubule as revealed by TEM is more similar to morphology p-rhabdoid A than the colloponeme or acanthoneme. Unlike the former morphology, basal spines of the hadroneme are not perpendicular to the tubule surface and the distal tubule always bears small spines. Furthermore, the lack of a v- shaped notch at the end of the basal tubule in an undischarged nematocyst of this morph distinguishes it from p-rhabdoid A. However, this notch is formed by the basal tubule folding back within itself (Gosse 1860, Bedot 1896, Moebius 1866, Cutress 1955,

Tardent 1988). Perhaps the presence of a notch is simply a function of the length of the

thickened basal tubule, with longer basal tubules forming the v-shaped notch whereas

shorter ones do not. Perhaps because he saw this difference as minor, Cutress (1955) did

not consider cerianthid nematocysts to be b-mastigophores, rather he considered them

macrobasic p-mastigophores.

p-rhabdoid A

Using the Weill system, this morphology would be identified as a microbasic p- mastigophore. Schmidt (1969) considered this a p-rhabdoid, noting several distinct characteristics confirmed by my investigations. As he noted, this type is the only nematocyst in Actinaria to lack apical flaps. Also, the capsule wall is often thin and seems more fragile than in other nematocyst types, although ultrastucturally it does not obviously differ from other types (Fig. 3.3). The spines are perpendicular to the shaft unless obstructed in contrast to the p-rhabdoid B1a, B2d, and B2s, in which the spines of the basal tubule are at an acute angle and point towards the capsule. Finally, the shaft typically does not extend to the end of the undischarged capsule but usually stops half- way. To these characters described by Schmidt (1969) and supported by den Hartog

(1995) I can add one more: the head of the t-shaped spines (noted by Schmidt 1969) of the basal tubule are embedded into tubule via folds in the tubule surface. Unlike in other morphologies, these folds do not form annulations that encircle the basal tubule. All of these distinctive features result in a distinctive morphology, relatively easy to identify with light, scanning electron, or and transmission electron microscopy.

Schmidt (1974) separates the actiniarian nematocysts of this morphology (called

p-rhabdoid As) from those in non-actiniarian groups such as corallimorphs (called p-

rhabdoid D) because of differences in spine length on the basal tubule and because non-

actiniarian nematocysts of this morphology have spines on the distal tubule (Fig. 3.7H).

Although I also observed these differences, I think that creating different categories

obscures the large overall similarity between these nematocysts in terms of shape, size,

surface of basal tubule, morphology, distribution of spines and ultrastructural features

such as the small folds in the tubule surface spaced between larger folds (see Fig. 3.3) visible using TEM. Therefore, I think the two morphologies should be similarly classified and these features that differ should be treated as variation within the same type. If the same type, with minor variations, is found across Hexacorallia, then the absence of apical flaps in actiniarian nematocysts of this type, held to be a synapomorphy of Actiniaria

(Reft and Daly 2012), is logical. The nematocyst would be a plesiomorphic morphology in Actiniaria that simply retained the ancestral apical cap while developing some variation in other features.

p-rhabdoid B1a

This morphology has a thickened basal tubule (historically defined as a shaft), a v-shaped notch, and a distal tubule evident from the notch, features that define a p-mastigophore in the Weill (1934) system, but Schmidt (1969) identifies it as part of a larger category he calls p-rhabdoid B (to which the p-rhabdoides B2d and B2s obviously belong). The p- rhabdoid B (or penicilli B for den Hartog (1977)) is defined as having a thickened basal

tubule that usually reaches the end of the undischarged capsule (longer in the macrobasic

case), apical flaps, spines that point toward the capsule after discharge, and a conical,

unarmed midpiece at the transition between basal and distal tubule (though a distal tubule

may not always be attached). In some nematocysts of this group, the basal tubule has two

distinguishable regions (discussed in detail below) (Schmidt 1969). Schmidt (1969)

named this morphology and defined it as having a uniform basal tubule. In fact, the B1

type is distinct from the other p-rhabdoides B in that it only has one region of the basal

tubule, it lacks the more proximal region (Schmidt (1969) defined the proximal region as

the Faltstück, which means pleated part), only having the Hauptstück (meaning main

part), which is about as long as the capsule (Schmidt 1969). The Hauptstück is defined as

having long, densely packed spines that taper in length just before the conical midpiece,

and is less elastic than the Faltstück. The “a” subdivision of the B1 category refers to the

fact that this morphology always has an evaginated terminal tubule (although den Hartog

(1995) disagrees that there is a B1b category and just calls this morphology penicilli B1).

The morphology seen here agrees with Schmidt’s (1969) identification. The basal tubule only has one type of spine which is longer than the diameter of the tubule and tapers towards the end just before the midpiece. This morphology also has a distinctive capsule shape making the identification of this type straightforward (see below for more discussion on this point). Additionally, the tubule has annulations that encircle the basal tubule and into which the spines insert. These annulations seem to be characteristic of the

nematocysts categorized by Schmidt (1969) as p-rhabdoid Bs, as they are also shared by

the B2d and B2s variants (which are also of this category, according to Schmidt 1969)

but absent in p-rhabdoid A. The folds in p-rhabdoid A where the spines insert may be a

precursor to the complete annulations seen in p-rhabdoid B (since the distribution of

types implies that A preceded B due to the more restricted distribution of B), but

character mapping on a robust phylogenetic tree would be needed to support this

interpretation. As previously mentioned, colloponemes also have these annulations which

may indicate a relationship between them and p-rhabdoid Bs.

p-rhabdoid B2d

Using the Weill (1934) system, this morphology can be classified as two different types.

If a terminal tubule is visible from the v-shaped notch then it has been considered a p- mastigophore (Weill 1934, Carlgren 1940, England 1991). However, more often, a tubule is not visible or is short, leading to its classification as an amastigophore (Weill 1934,

Carlgren 1940, Carlgren 1949, England 1991). Like the previous morph, Schmidt (1969) considered this morphology a p-rhabdoid B. However, this type has two distinct regions of the tubule (Faltstück and Hauptstück) demarcated by different spines. The more basal

Faltstück has shorter, narrower spines that are sparsely packed onto the tubule. In contrast, the more distal Hauptstück has very long, wider spines that tend to curl upward to form a bottle-brush like appearance. Because the basal tubule has the two regions, this morphology would be classified as p-rhabdoid B2, and since the Faltstück is shorter than the Hauptstück, it would be further classified as the b variant of B2 (Schmidt 1969). A terminal tubule may or may not remain attached, but Schmidt (1969) claims it is always short.

Although the spines do distinguish regions of the basal tubule, the surface of the tubule

remains the same. Unlike p-rhabdoid B2s, the annulations that encircle the basal tubule

maintain the same spacing and pattern for the entire length. Therefore, this division

between Faltstück and Hauptstück is based on spine pattern alone.

p-rhabdoid B2s

As with the previous morphology, the p-rhabdoid B2s has been identified as both p- mastigophore and (more commonly) amastigophore, depending on whether a terminal tubule could be discerned in the undischarged state (Weill 1934, Carlgren 1940, Carlgren

1949, England 1991). The basal tubule in this morphology is often long, sometimes much longer than the capsule; these attributes in addition to capsule shape, distinguish this morphology from 6. Similarly to morphology 6, some of these nematocysts would be classified as p-rhabdoid B2a. The nematocyst of Fig. 3.10C would be considered a B2a as a result of the basalmost Faltstück being shorter than the more distal Hauptstück.

However, the nematocyst of Fig. 3.10E would be considered a B2b due to a Faltstück that is longer than the Hauptstück. I choose to group these morphologies together and distinguish them from the p-rhabdoid B2d due to the reasons given above and due to the pattern of annulation on the surface of the basal tubule that distinguishes the two regions.

Unlike in p-rhabdoid B2d, the change in spine shape and pattern as the tubule transitions from Faltstück to Hauptstück has a corresponding change in annulation pattern. In the

Faltstück, the annulations are more widely spaced apart and form a tubule that seems flexible with buldges and bends, whereas in the Hauptstück, the annulations are closer

89 together and the tubule seems more smooth and rigid. Schmidt (1969) commented on the differences between the regions calling the pleats on the Faltstück coarser and less closely spaced. However this distinction is not evident in morphology of all things identified as p-rhabdoid B2a as it is not seen in morphology 6.

Diakaneme

This morphology does not fit any existing classification and is probably an example of an innovation in this lineage. Carlgren (1949) did not particularly mention this type in his account of cnidom in Urticina, saying that this species bears only basitrichs and microbasic p-mastigophores, the latter of which could refer to this morphology and/or morphology 4, which this species also have. However, others have noticed this distinct type (Sanamyan and Sanamyan 2006, den Hartog and Ates 2011). Typically it is defined as a p-mastigophore, and den Hartog and Ates (2011) classify it as a large penicilli B1, the largest known to occur. This treats this morphology as a scaled up version of the p- rhabdoid B1a, however, there are morphological differences that make this equation doubtful. Fine detail of the U. felia nematocyst reveals differences from the p-rhabdoid

B1a. The tubule surface does not have the annulations that characterize p-rhabdoid Bs, nor does it have the conical midpiece. Although the basal tubule does narrow at the transition to the distal tubule, the change in diameter is not as dramatic as in p- rhabdoides. Although Schmidt (1969) claims that p-rhabdoid B1 was observed in Actinia equina, a variety from the Mediterranean (Schmidt 1971), and Anthopleura rubripunctata, other members of the family to which Urticina belongs, I did not observe

90 this type in any other member of Actiniidae including Actinia equina. This nematocyst type is also found in the mesenterial filaments of species in the genus Cribinopsis which provides support for the hypothesis that the two genera are closely related (Sanamyan and

Sanamyan 2006).

Because nematocysts are so important to the biology of cnidarians and are ubiquitous, it is likely that some lineages have evolved novel nematocysts for particular purposes or even through stochastic processes as populations become divided. Therefore, there are likely to always be a few nematocyst types restricted to a few lineages that are unique and not comparable to other types. The diakaneme, which is only known to occur in the genera Urticina and Cribinopsis, seems to be an example of this phenomenon.

Obviously, this nematocyst evolved from some pre-existing form, but without a well- supported tree with mapped nematocyst features, it is difficult to determine the transformation series.

Aphylloneme

Like the diakaneme, this morphology seems to be restricted to a particular group (the family Actiinidae) and body structure (mesenterial filaments). The capsule shape of this morph does resemble that of the diakaneme with an elongate form widest at the non- apical end. In fact, the general similarity of the undischarged form makes it difficult to determine if the morphologies observed by Schmidt (1969) in taxa such as Bunodactis rubripunctata should be classified as a diakaneme or aphylloneme. den Hartog (1987,

Fig. 3.5E) documents a nematocyst in the mesenterial filaments of Bunodosoma

91 biscayensis that he describes at the same morphology as that in Urticina which he considers a p-rhabdoid B1a (although I have identified it as a distinct morphology called the diakaneme). Although the light microscope data provides little detail, it is clear that the shape and size of this morphology is more consistent with the aphylloneme than p- rhabdoid B1a, particularly when considering the discharged nematocyst (den Hartog

1987, Fig. 3.5E compare to Fig. 3.2F). However, this morphology might also have been a diakaneme as the light microscope data is not detailed enough to eliminate this possibility

(discussed below). Therefore, the distribution of the aphylloneme and what it has been previously identified as is unclear.

p-mastigophores, amastigophores, and the subdivision of these types

Historically, the morphological types called p-mastigophore and amastigophore have been defined by the presence or absence of a terminal thread (Weill 1930, 1934,

Carlgren 1940, Mariscal 1974, also see Fig. 3.1). This character has caused confusion

(Cutress 1955, Östman 2000) because a terminal filament can be very difficult to see under the light microscope (e.g. it is visible in Fig. 3.2G though not in Fig. 3.11A even though both of these do have a terminal thread). However, previous authors have demonstrated that there is always (barring one exception, den Hartog’s (1995) penicilli

2Bc ) a terminal thread (Cutress 1955, Schmidt 1969, Östman 2000) although often this thread is short and breaks off. Schmidt (1969) noted that this character was too highly variable to be useful in classifying nematocysts and only used it for lower level subdivisions (see previous discussion on p-rhabdoid B1b). This opinion is not universal:

England (1991) suggested that Schmidt (1969) did not subdivide enough using this criteria.

Because this distinction is not particularly helpful in classifying nematocyst variation, some authors have suggested either renaming all amastigophores p- mastigophores (Cutress 1955) or creating a category called p-amastigophores (Östman

2000). The latter solution indicates a similarity between p-mastigophores and amastigophores while still maintaining distinct morphological categories (Östman 2000).

However, not all p-mastigophores resemble those nematocysts traditionally classified as amastigophores and the unilateral solutions of Cutress (1955) and Östman (2000) pool unlike morphologies (e.g. p-rhabdoides A and B2s).

Schmidt’s (1969) categories of the p-rhabdoides are often justified (at least from a morphological level, the homology of these characters still need to be assessed phylogenetically). Specifically, p-rhabdoid A (and D) is clearly distinct and should not be confused with the p-rhabdoid Bs. Within the latter category, morphological justification exists for the division between p-rhabdoid B1a, and p-rhabdoid B2a/b. However, the division between B2a and B2b seems less clear as it is only based on relative length differences and not any qualitative differences. Furthermore, with the B2a category, there seem to be two distinct forms (morphology B2d vs. B2s). Within this category more subdivision may be justified. However, a multivariate assessment of these quantitative features may help determine how much subdivision is justified as these different morphologies could represent points in a continuum rather than discrete groupings.

Basitrichs and b-mastigophores

In Stephenson’s (1928) two-part classification system of nematocysts, the morphology later identified as basitrichous isorhizas was identified and given the name spirulae, clearly separate from what would later (Weill 1934) be called the mastigophores

(called penicilli by Stephenson). The spirula was recognized to be common across the anthozoan subclass Hexacorallia (Weill 1934, Carlgren 1940, 1945). However, Carlgren

(1940) argued this category could be confused with a similar looking form which he determined actually has a shaft (or larger diameter basal tubule) and called this the b- mastigophore, although he acknowledged that the dilation of the shaft could be small and in undischarged form, basitrichs and b-mastigophores can closely resemble each other.

Cutress (1955) differentiated basitrichs and b-mastigophores based on the on the apparent rigidity and folding of the basal tubule. According to Cutress (1955), basitrichs have a completely coiled tube, and the descending tubule is not rigid and thick, but of equal diameter with the distal tubule (see Cutress 1955, Fig. 3.3). Conversely, in b- mastigophores, the basal tubule forms a rigid rod that tapers to a thinner distal tubule (see

Cutress 1955, Fig. 3.4), though the shaft can be only minutely larger than the distal tubule

(as small as 0.1 µm). Östman (2000) supported Cutress’ (1955) argument, even going so far to argue that there may be no difference at all between the basal and distal tubule and that a “distinct shaft is not present in all discharged b-mastigophores” in hydrozans (p.

42), an odd statement given that she defines the shaft as an enlarged basal portion of tubule and that highlights questions about the very nature of the shaft (discussed separately below). Östman (1979, 1982, 1983, 1987, 1988) has focused on medusozan

94 nematocysts, and, as she notes (Östman 2000), there are differences in shape and construction that render homology of nematocyst types between Medusozoa and

Anthozoa unclear. Cutress (1955) and Östman (2000) consider b-mastigophores more widespread than basitrichs; Östman (2000) considers it is one of two categories present throughout Cnidaria. Also important to note is that Östman (2000) does not comment on this morphology. In contrast, Schmidt (1969), in part because he defines the shaft as being based more on the large basal spines than on any dilation of the shaft (see discussion below on the nature of the shaft), argues that the two types should be considered one, the b-rhabdoid. Under his defintion, acanthonemes, aphyllonemes, colloponemes, and hadronemes) would all belong to this category.

As discussed previously, Weill’s (1934) definition of this type is problematic and does not properly describe the morphology. Schmidt (1969) also fails to adequately segregate the morphology into like groups; his general category b-rhabdoid would group morphologies 1-3 despite their clear distinctions. Cutress (1955) correctly distinguished cerianthid nematocysts from these morphologies, but the rod-like basal tubule is not a way to consistently distinguish between three distinct morphologies, acanthonemes, colloponemes and aphyllonemes, particularly given that the basal tubule may not always be rigid and may show evidence of folding as in Fig. 3.5C, although he and Östman

(2000) were correct that colloponemes do not have basal tubules with larger diameters than the distal tubules. An exact description of the fine details of these morphologies with broad sampling in hexacorallia has been lacking. In Hexacorallia, the acanthoneme is the

95 most widespread of the forms that fall into these confusing categories (basitrich, b- mastigophore, b-rhabdoid).

Unlike the other nematocysts studied here, features of the undischarged capsule

(such as shape) of acanthonemes and colloponemes do not consistently distinguish the two types (see below for more discussion on this point) making easy determination from preserved specimens difficult (aphyllonemes, however, can be distinguished) . The features that distinguish the two groups are not discernible under the light microscope and they have convergent undischarged form. Therefore the identification of these will always be difficult unless phylogenetic distribution allows for an inference on which form is more likely in a particular group. Also, unfortunately the “b-mastigophores” found in the tentacles of the genera Stomphia and Actinostola (Carlgren 1949) were not obtained for study here, therefore it is impossible to say where they fit in regards to these categories (or if they represent a distinct category).

Light microscopy of undischarged nematocysts

One of the main problems with creating an integrated and practical classification system is simply that to fully categorize the morphological diversity, discharged nematocysts must be used. Spine arrangement and form and tubule surface are rarely obvious in the undischarged form. Also, nematocysts that are not fully mature may be misclassified, an issue that den Hartog (1995) cites as part of the problem in Schmidt’s

(1969) misinterpretation of nematocysts of the genus Telmatactis. Proper discharge (i.e. discharge through the space created by apical structure and not created by a broken

96 capsule) is thought to only be possible with fully mature nematocysts which nullifies this problem for discharged nematocysts. However, most cnidarian biologists are working with preserved material.

The importance of features visible only in discharged capsules is most acute in cases where distinct morphologies converge on a similar undischarged form. This issue has long led to the confusion between basitrichs and b-mastigophores as the thickness of the “shaft” in b-mastigophores has been noted to be quite small in some situations making it difficult to distinguish “shaft” from an undischarged tubule stretched to accommodate large spines (Carlgren 1940, Cutress 1955, Schmidt 1969). Acanthonemes and colloponemes here illustrate this problem: no obvious features to differentiate the undischarged capsules under light microscopy. However, for the former, the thick descending tubule is the result of large spines; the latter has relatively short spines and the thickness of the descending tubule is the result of the annulations of the basal tubule.

In cases with similar-looking (though actually distinct) morphology, no classification system will resolve the problem for undischarged nematocysts.

In other cases, a study such as this one which looks at variation visible at all levels (light, scanning electron, and transmission electron microscopy) can help identify what features are correlated and provide insight for future classification if only undischarged nematocysts are available. Most of the major morphological divisions seen here can be identified using capsule shape and length of descending tubule (see Table 3 for summary of these features). Using these features the following morphologies can be distinguished: p-rhabdoid A has an elongate-shaped, symmetrical capsule with the central

97 tubule only descending half-way down the capsule, p-rhabdoid B1a has an ovate or egg- like capsule shape with the central tubule almost filling the length of the capsule, p- rhabdoid B2d has a capsule broader near the apex than at the other end and often looks slightly asymmetrical with a central tubule that reaches the bottom third of the capsule, p- rhabdoid B2s which has an elongate capsule that is symmetrical being widest at the midpoint and a central tubule that reaches the bottom third of the capsule (and typically fills the length of the capsule), the diakaneme which has a capsule broadest at the non- apical end and a central tubule that descends about half-way down the capsule.

Associating these capsule shapes with discharged nematocyst morphologies as done here will assist in creating a system by which nematocysts are identified more consistently even if only using undischarged material. How successful capsule shape is at classifying these morphologies needs to be assessed using a quantitative morphometric analysis with known/identified nematocysts.

Relationships among morphologies

Although documenting morphological differences is important to understanding the evolution of nematocysts, separating these morphologies into discrete categories does not achieve that goal. To track the evolution of these structures, we need to understand the transitions between character states that lead to different morphs as newer morphologies evolved from old ones. Although a full phylogenetic analysis of nematocyst morphology is beyond the scope of the present work, initial statements of homology can be made based on observations of this study. These can serve as hypotheses of homology

(Patterson 1988, de Pinna 1991) to be tested in a phylogenetic framework. Characters are

evaluated for homologies follow.

Basal tubule (and spine attachment): The basal tubule surface has three states among these morphologies. Some (acanthonemes and diakanemes) are smooth with only small, slit-like openings in the tubule surface into which the base of the spine attaches to the tubule (Figs. 4, 11). One type (aphyllonemes) is completely smooth with no markings at all (Fig. 3.12). Others (hadronemes, p-rhabdoides A) have large folds in the tubule surface into which the head of a spine inserts, but these folds never encircle the entire tubule (Figs. 6, 7). Finally, some morphologies (colloponemes, the p-rhabdoides B) have annulations that encircle the entire tubule and through which the spines attach (Figs. 5, 8-

10). Furthermore, most morphologies here have one region of the basal tubule (all but p- rhabdoides B2d and B2s). However, these morphologies have two distinct regions, the

Faltstück and Hauptstück (defined by spine pattern). In the p-rhabdoid , these two regions are also discernable by a change in the annulation pattern.

Distal tubule: Of the morphologies presented here, only a few have spines on the distal

tubule. Acanthonemes, colloponemes, hadronemes, and the non-actiniarian examples of

p-rhabdoid A all have spines on the distal tubule whereas all other morphologies and the

actiniarian examples of p-rhabdoid A do not. These spines are always small and never

protrude very far above the tubule surface. These spines in acanthonemes, hadronemes

and p-rhabdoides A (that have them) are more similar in shape than any are to those of

colloponemes. Also, a dramatic change in diameter during the transition from basal to

distal tubule is evident in hadronemes and all p-rhabdoid forms, whereas a subtler

narrowing is evident in acanthonemes and diakanemes. Colloponemes and aphyllonemes

do not have any obvious change in tubule diameter.

Basal tubule spine shape: The spines of the basal tubule vary in size in ways that can confound easy classification. However, at least two major morphs of spines can easily be recognized: the T-shaped spines and the non-T-shaped spines. Hadronemes and p- rhabdoides A have the distinctive T-shaped spines identified by Schmidt (1972), and observed but not noted by Skaer and Picken (1965), as occurring in haplonemes such as holotrichs and a few heteronemes. These have a broad base that quickly narrows to form the head of the T.

In the non-T-shaped spines (in all other morphs), the base of the spine is usually a bit broader than the base of the blade, but narrowing is either more gradual

(acanthoneme, Fig. 3.4) or less pronounced (colloponemes, p-rhabdoid B forms, Figs.

3.5H; 3.8F; 3.10G, J; 3.11D, E) such that no obvious T-shape is created at the base. The spines within this category vary in the exact shape, and are continuum of variation that is difficult to segregate.

Taking these characters together, some morphologies can be hypothesized to be closely related to each other evolutionarily. For example, p-rhabdoid B2d and B2s share basal tubule and spine characters, although the basal tubule annulations do not change in pattern as in the B2s form and these two types most closely resemble each other. The p- rhabdoid B1a shares some tubule and spine features with morphs B2d and B2s, though this type only has one region of tubule, which Schmidt (1969) homologizes with the

Haupstück of other types, which is logical given the type of spines in this region.

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Although superficially the p-rhabdoid B1a shares an overall similarity with the p- rhabdoid A, the differences in spine shape, tubule surface, and apical structure all indicate that it is more like other p-rhabdoid B forms than the A form.

The colloponeme is difficult to place in the context of other morphologies. Spine shape and tubule surface would seem to indicate a close relationship with the p-rhabdoid

B2d and B2s (as the basal tubule and spines all appear very similar to the Faltstück of these morphs). However, the presence of a distal tubule that is no different in size from the basal tubule and bears spines is not consistent with these other morphs. In fact, these features and those of the undischarged capsule make this morphology more similar to acanthonemes than to the p-rhabdoid B2d or B2s. Therefore, which placement is preferred will depend on which characters are given more weight.

The hadroneme and p-rhabdoid A have some interesting similarities. Both have the T-shaped spines that are attached through folds in the tubule, and the distal tubule

(which is of much smaller diameter than the basal tubule) of both may be spined (always for hadronemes and for non-actiniarians for p-rhabdoides A). Furthermore, this nematocyst always has an apical cap, the plesiomorphic state for Anthozoan apical structure, rather than flaps which would be expected in Actiniaria (Reft and Daly 2012).

Given that the presence of a v-shaped notch has been demonstrated to result from basal tubule folding in on itself (Cutress 1955), the lack of this feature in hadronemes could indicate that the basal tubule is shorter or just packaged differently than in p-rhabdoid A while remaining structurally the same. Although a full phylogenetic analysis is beyond the scope of the current work, the distribution of these two morphologies across

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Hexacorallia suggests that the features shared between these morphologies may be plesiomorphic for the group. Hadronemes are found only in cerianthids (Carlgren 1940,

Schmidt 1974), which are currently believed to be the sister group to the rest of the hexacorals (Won et al. 2001, Daly et al. 2003, Brugler and France 2007). The transition from hadronemes to p-rhabdoides A in non-actiniarians (antipatharians, zoanthids, corallimorpharians, and scleractinians) would require a change in the basal tubule to create the v-shaped notch (discussed previously) and a change in the orientation of the spines to the tubule surface (from acute to perpendicular). Within actinarians, the only major change in this form is the loss of spines on the distal tubule, which seems to be common in the group as only acanthonemes have distal spines of the morphologies studied. Therefore, morphology p-rhabdoid A could be interpreted as a derived form of the hadroneme.

Finally, as discussed previously, the diakaneme is unusual. Features of the tubule and spine attachment discussed here make it similar to acanthonemes, whereas the shape of the undischarged capsule implies a similarity to aphyllonemes. However, having two differently shaped spines on the basal tubule, the spine shape generally, and the presence of a v-shaped notch in undischarged form distinguish it from both acanthonemes and aphyllonemes. Other than lacking T-shaped spines, which makes it dissimilar from hadronemes and p-rhabdoides A, this morphology has little in common with the p- rhabdoid B morphologies. Therefore, the placement of this morphology amongst the others is unclear.

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Nature of the shaft

Another of the main points of confusion in classifying nematocysts is the definition of a shaft. Weill (1930, 1934) identified the shaft as a swelling or thickening of the tubule at the base; this can be cylindrical and essentially isometric or have bulges/dilations (Weill

1934, Mariscal 1974). Cutress (1955) agreed, defining the shaft as the portion of the tubule with a greater diameter, though he acknowledged that degree to which the two portions differ can be quite small (0.1um). If the shaft is defined as a widening of the basal tubule, then morphologies 3-8 have shafts whereas morphologies 1-2 do not. The colloponeme is usually identified as a b-mastigophore (Carlgren 1940, Westfall and Hand

1962, 1965, Mariscal 1974) or b-rhabdoid (Schmidt 1969), both of which are defined as having a shaft. However, the basal tubule is not wider than the distal tubule in this morphology. Cutress (1955) may be correct that there is a minute change in the diameter as basal tubule transitions to distal tubule, however, if any small change is considered enough distinguish these two regions then morphology 1 would be included in this definition as well. Applying this standard results in no differentiation among nematocyst types, as the shaft would be a character shared amongst all types; such a definition of the shaft is not helpful in categorizing or understanding nematocyst morphological diversity.

Schmidt (1969) claimed that a basal shaft, whether weakly or strongly developed, does not correspond as much to a conspicuous thick portion but more to a conspicuous armature (Schmidt 1969). Therefore, in a heteroneme, long spines at the base define the shaft, with a transition to small spines or the loss of spines demarcating the beginning of the distal tubule; whether the change in diameter is gradual or abrupt is irrelevant

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(Schmidt 1969, p. 288). In haploneme nematocysts, in contrast, either the size of the spines does not change over the whole length of the tubule (as in holotrichous isorhizas) or the spines gradually taper in size as the tubule tapers in diameter with no abrupt change in size at any point. Under Schmidt’s (1969) definition, the defining characteristic of the shaft is not any intrinsic quality of the tubule but rather of the spines borne on the basal tubule. Under this defintion, morphology 1 now would be considered a rhabdoid with a shaft. However like Cutress’s (1955) 0.1um cutoff for the dilation of a shaft, this definition is unsatisfactory, as its application means that all morphologies studied here have shafts.

Another way to define the shaft would be to describe all nematocysts with annulations that encircle the basal tubule as shaft-bearing. This defines the shaft using an intrinsic feature of the tubule and means that morphologies 2, 5-7 would be characterized as having a shaft. However, this definition would exclude the p-rhabdoid A, which has always been considered to have a shaft (Weill 1934, Carlgren 1940, Schmidt 1969,

Mariscal 1974) and certainly does have a distinct, robust basal tubule. A more inclusive definition would be one that defines the shaft as the portion of the tubule on which spines are attached via folds in the tubule surface. Such a definition would include all morphologies here except acanthonemes, diakanemes and aphyllonemes. Such a definition incorporates both a spine character, i.e the attachment of the spine, and an intrinsic feature of the tubule surface itself, the fold. This classification would then include all nematocysts traditionally classified as shaft-bearing. Furthermore, the ultrastructural data from the transmission electron microscope distinguishes the

104 descending tubule in this way as well. Extra folds in this central tubule that appear to correspond to spine attachment differentiate morphologies that under the scanning electron microscope have folds for spine attachment. The only morphology studied here which lacked these spine-associated folds under the TEM was the acanthoneme (and the holotrichs which are provided for comparison).

Another aspect of this discussion is the question of homology between nematocysts with an isodiametric basal thickened tubule or rhabdoides (the only kind found in Anthozoa) and those with bulges as part of the shaft or rhopaloids (which is characteristic of nematocysts common to medusozoans such as stenoteles and euryteles).

Using the term shaft for the basal tubule of both types of nematocysts implies a homology between the two, a view explicitly supported by the Weill (1934) classification system in which all nematocysts with wide diameter basal tubules are grouped as heteronemes (see Fig. 3.1). However, evidence supporting that conclusion has never been formally evaluated, and comparison of basal portions of these two types of nematocysts reveals some differences. Longitudinal furrows in the discharged stentotele (and eurytele) tubule have been previously observed (stenotele: Tardent and Holstein 1982, Östman et al. 1991, eutytele: Yanagihara et al. 2002) and the extra folds corresponding to these furrows are often observed with TEM (e.g. stentotele: Chapman and Tilney 1959,

Tardent and Holstein 1982, eurytele: Yanagihara et al. 2002). These folds are not observed in hexacorallian heteronemes (or rhabdoides), which have instead small folds around the base of the spines. Longer extensions are common on hexacorallian nematocysts, but these are off to the side of the tubule, not longitudinal in orientation (see

105

Fig. 3.3, Westfall and Hand 1962, Westfall 1965, Watson and Mariscal 1985, Blake et al

1988, Reft et al. 2009). Although similar small folds in the tubule are present in the rhopaloids, these occur more posteriorly than in the rhabdoides, often being evident only just before the transition to the distal tubule (see Fig. 3.13, Chapman and Tilney 1959,

Campbell 1977, Tardent and Holstein 1982, Clausen 1991). Furthermore, spines seem to associate with the bulging portions of the tubule (see Fig. 3.13, Chapman and Tilney

1959, Tardent and Holstein 1982, Östman et al. 1991, Yanagihara et al. 2002) rather than other portions of the basal tubule. In discharged form, no folds or annulations are evident in the surface of the tubule where the spines attach, rather only a small spine scar similar to that seen in acanthonemes (which is not traditionally considered to bear a shaft) is evident if the spine is stripped off (see Fig. 3.13, Chapman and Tilney 1959, Yanagihara et al. 2002, Östman et al. 1991). All the morphologies studied here that are traditionally considered to be shaft-bearing (hadronemes, all p-rhabdoid forms and usually colloponemes) either have large folds or annulations in the tubule where the spines attach. Taken together, the morphological and ultrastructural features of the basal tubule of the rhopaloides seem to distinguish this group from the basal tubule of the rhabdoides, therefore using the same terminology for both structures implies a homology where there may be none. Medusozoans have rhabdoides too, although the shape of these can be quite different from anthozoan examples (Skaer 1973, Östman et al. 1979, Östman 2000). The relationship between medusozoan and anthozoan rhabdoides needs to be assessed to determine what morphological similarities exist between the two.

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Ultimately, regardless of what definition is used to for the shaft, the question of homology remains. Ideally, terms used to describe nematocyst features across various morphological types should indicate homology, i.e. that the feature is not only morphologically similar but also shares an evolutionary history. If “shaft” is used for features that do not share recent and exclusive evolutionary history, then the term is not useful in classifying morphological diversity in a way that helps us understand nematocyst evolution. The only way to assess this homology is to test the homology statement made by these definitions on a robust phylogenetic tree. Coding the shaft character in all the variations listed above will be necessary to determine which causes the least amount of homoplasy on the tree.

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Table 3.1. Summary of taxa and collection locality and body part sampled for SEM and TEM for nematocyst monograph. A: acontia, Ac: acrorhagi, C: body column, MF: mesenterial filament, T: tentacle. S indicated that the species was studied by Schmidt (1969) whereas (S) indicates that Schmidt studied the same genus.

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Table 3.1 Higher Taxon Species Location collected SEM TEM Actiniaria Actinia equina S SW of Mahee Island, Strangford Lough, N. C, MF, T Ireland Adamsia palliata Ringhaddy Sound, Strangford Lough, N. A, C, MF, T Ireland Aiptasia sp. (S) Aquarium Adventure, Columbus, OH, A, C, MF, T USA Anemonia sulcata S Ballymacormick Pt, Groomsport, N. C, MF, T Ireland Anthopleura San Juan Island, WA, USA Ac, C, MF, T T elegantissima (S) 113 Bartholomea annulata University of Virgin Islands, St. Thomas, A, C, MF, T

US Virgin Islands Boloceroides mcmurrici S Kyoto University Seto Marine Lab, T Shirahama, Japan Bunodosoma cavernata Galveston, TX Ac, C, MF, T Calliactis polypus (S) Jeju Island, S. Korea A, C, MF, T Calliactis tricolor (S) Galveston, TX, USA A, C, MF, T Cereus pedunculatus S SW of Mahee Island, Strangford Lough, N. A, C, MF, T Ireland Condylactis gigantea Aquarium Adventure, Columbus, OH, A, C, MF, T T USA Continued

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Table 3.1 Continued Diadumene leucolena S Marine Biological Lab, Woods Hole, MA, USA A, C, MF, T T Diadumene sp. (S) Jakyakdo Island, S. Korea A, C, MF, T Entacmaea quadricolor Jeju Island, S. Korea C, MF, T Epiactis proliferia San Juan Island, WA, USA C, MF, T Acontiaria sp. Jakyakdo Island, S. Korea A, C, MF, T Halcurias levis Kyoto University Seto Marine Lab, Shirahama, T Japan Haliplanella lineata Jakyakdo Island, S. Korea A, C, MF, T Metridium senile S Marine Biological Lab, Woods Hole, MA, USA A, C, MF, T A, T Nemanthus nidtus Jeju Island, S. Korea C, MF, T Paracondylactis hertwigi Jakyakdo Island, S. Korea C, MF, T

114 Sagartia elegans S SW of Mahee Island, Strangford Lough, N. A, C, MF, T Ireland

Sagartigeton lacerates S Ringhaddy Sound, Strangford Lough, N. Ireland A, C, MF, T Sagartigeton undatus S Ringhaddy Sound, Strangford Lough, N. Ireland A, C, MF, T Actinostola sp. San Juan Island, WA, USA C, MF, T T, MF Urticina felina S SW of Mahee Island, Strangford Lough, N. C, MF, T Ireland Antipatharia Antipathes grandis Auau Channel, HI, USA T Antipathes griggi Auau Channel, HI, USA T Ceriantharia Ceriantheopsis Gulf Specimen Marine Lab, Panacea FL, USA C, MF, T C, MF, americanus T Cerianthus filiformis Kyoto University Seto Marine Lab, Shirahama, T Japan Cerianthidae sp. Reef Hot Spot, Inglewood, CA, USA T T Continued

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Table 3.1 Continued Corallimorpharia Discosoma sp. Aquarium Adventure, Columbus, OH, USA MF MF Rhodactis sp. Reef Hot Spot, Inglewood, CA, USA MF Scleractinia Platygyra asteriformis Gulf Specimen Marine Lab, Panacea, FL, USA T Goniporia sp. Reef Hot Spot, Inglewood, CA, USA T T Zoanthidea Protopalythoa mutuki Neil Blackstone culture C, MF, T C, MF, T Zoanthus pulchellus University of Virgin Islands, St. Thomas, US C, MF Virgin Islands Octocorallia Clavulariid sp. A Neil Blackstone culture T Renilla mulleri Gulf Specimen Marine Lab, Panacea FL, USA T Medusozoa: Cordylophora caspia Exeter, NH T Hydrozoa Ectopleura larynx Darling Marine Center, Walpole, ME T T

115 Hydra sp. Paulyn Cartwright culture T

Sertularella sp. Darling Marine Center, Walpole, ME T

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Table 3.2. Morphologies described in monograph, their distribution in cnidarian body tissues, and their categorization under previous classifications.

Morphology Weill (1930, 1934; Schmidt den Hartog Body Carlgren 1940; (1969, 1974) (1980, 1987, Structure Mariscal 1970) 1995, 2011) acanthoneme basitrich b-rhabdoid Spirula A, Ac, C, MF, T colloponeme b-mastigophore/ b-rhabdoid Spirula Ac, T basitrich hadroneme b-mastigophore b-rhabdoid Spirula C, MF, T p-rhabdoid A p-mastigophore p-rhabdoid Penicilli A C, MF, T A/D p-rhabdoid B1a p-mastigophore p-rhabdoid Penicilli B1 MF, T B1a p-rhabdoid B2d p-mastigophore/ p-rhabdoid Penicilli B2a C, MF, T amastigophore B2a p-rhabdoid B2s amastigophore p-rhabdoid Penicilli B2a/b Ac, C, (var. a and b) B2a/b MF, T diakaneme p-mastigophore p-rhabdoid Penicilli B1 MF B1a? aphylloneme basitrich p-rhabdoid Penicilli B1? MF B1a? The ? reflects the uncertainty of which morphology (diakaneme or aphylloneme) Schmidt (1969) and den Hartog (1987) documented. A: acontia, Ac: acrorhagi, C: body column, MF: mesenterial filament, T: tentacle.

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Table 3.3. Light microscope details that differentiate the nine morphologies observed in monograph study. Nematocyst Capsule shape Length of basal V-shaped morphology tubule in capsule notch in basal tubule acanthoneme Variable: usually Variable, usually more no elongate than 1/2 the length colloponeme Elongate and narrow in Variable, usually more no width than 1/2 the length hadroneme Variable: usually Variable: usually 1/2 no elongate, but can be more to 2/3 the length, ovoid rarely 1/3 the length p-rhabdoid A Elongate, though can be 1/2 the length yes broad p-rhabdoid B1a Usually egg/ovate Almost full length yes p-rhabdoid B2d Asymmetrical, widest 2/3 to full length yes point near apical end p-rhabdoid B2s Symmetrical, widest 2/3 to full length yes point at midpoint diakaneme Elongate with wide non- 1/2 the length Usually apical end (sometimes not observed) aphylloneme Elongate, widens from 2/3 the length no midpoint of capsule to the non-apical end

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Figure 3.1. Flow chart of the Weill (1934) classification system for nematocysts.

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Figure 3.2. Nematocyst types. Light micrographs of nematocysts as traditionally classified using the modified Weill system. Scale bars = 20 µm. A. Holotrichous isorhiza of zoanthid tentacle Protopalythoa mutuki. The tubule has no swellings or thickened parts (no shaft). B. Undischarged basitrichous isorhiza from tentacle of Protopalythoa mutuki. There appears to be a thicker part of the tubule in the undischarged state which corresponds to the region with large spines at base of tubule. C. Discharged basitrichous isorhiza from tentacle of Protopalythoa mutuki with large spines at base on display. D. Microbasic b-mastigophore from tentacle of Cerianthidae sp. Arrow indicates the end of the shaft which has no v-shaped notch. E. Microbasic p-mastigophore from tentacle of scleractinian Goniopora sp. Arrow indicates the v-shaped notch which marks the end of the shaft. The tubule continues from that notch. F. Discharged microbasic p- mastigophore from mesenterial filaments of actiniarian Metridium senile. G. Microbasic amastigophore from mesentearial filaments of actiniarian Sagartia elegans. Arrow indicates short tubule that continues from the notch. I. Discharged microbasic amastigophore from mesenterial filaments of S. elegans. Arrow indicates remnant of tubule left behind in capsule. H. Macrobasic amastigophore from tentacle of actiniarian Diadumene sp. Arrow indicates a loop in the bent shaft.

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Figure 3.2

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Figure 3.3. Transmission electron microscope images of some of the morphologies observed in monograph study. A., B. Holotrichs from tentacles of octocoral Clavulariid sp. A. The tubule has no noticeable change in spine pattern or diameter so that tubule packaged at the bottom of the capsule as in A. is no different from tubule that descends down the middle of the capsule (as in B.). Scale bar = 500 nm. C. Acanthoneme from tentacle of actiniarian Condylactis sp. Large spines situated on the base of tubule are indicated with arrow, distal tubule indicated with arrowheads. Base of tubule is smooth around the spines. Scale bar = 500 nm. D. Apical region of acanthoneme from the tentacle of actiniarian Actinostola sp. Proximal tubule descends down the middle of the capsule (arrow) with distal tubule to either side (arrowheads). Few projections of the basal part of the tubule are visible. Apical flaps (A). Scale bar = 500 nm. E. Overview of acanthoneme from a tentacle of the zoanthid Protopalythoa mutuki. Both the basal region of tubule with large spines (arrow) and the distal tubule (arrowheads) are visible. Only a few projections of the basal tubule are evident. Scale bar =2 µm. F. Apical region of acanthoneme from a tentacle of the zoanthid Protopalythoa mutuki. Tubule is smooth between distantly spaced projections (arrow) of the basal tubule. Scale bar = 500 nm. G. Overview of colloponeme from acontia of Metridium senile. A long shaft (arrow) fills the center of the capsule. Scale bar = 10 µm. H. Apical region of colloponeme from acontia of Metridium senile. Note the spines of the shaft region (arrow) and the numerous folds of the shaft region (arrowheads), the distal part of the tubule (T) can be seen to either side of the shaft. Scale bar = 2 µm. I. Overview of p-rhabdoid A from a mesenterial filament of Discosoma sp. Both the shaft region with long spines (arrow) and the distal tubule (arrowhead) are visible. Scale bar = 10 µm. J. Apical region of p-rhabdoid A from a mesenterial filament of corallimorphorian Discosoma sp. Small folds in the surface of the basal tubule (arrowhead) fill the regions between long projections of the same tubule (arrow). Scale bar = 500 nm. K. Apical region of p-rhabdoid A from a mesenterial filament of Actinostola sp. Large spines (arrow) fill the shaft. Scale bar = 500 nm. L. Oblique section of p-rhabdoid B2s from acontia of Metridium senile. Note the two regions of basal shaft (arrows). Scale bar = 2 µm. M. Apical region of nematocyst in L, note the numerous folds in the tubule (arrow). Scale bar = 500 nm. N. More distal region of basal tubule of nematocyst in L, note spines encased by tubule. Small folds (arrow) in the tubule are evident between regularly spaced larger folds. Scale bar = 500 nm. O. Hadroneme from a tentacle of cerianthid Ceriantheopsis americanus. The long spines of the shaft are indicated with an arrow. Arrowheads indicate the distal tubule. Scale bar = 2 µm. P. Detail of apical region of hadroneme from a mesenterial filament of Ceriantheopsis americanus. Like in p-rhabdoid A, this nematocyst has periodic long extension of the tubule (arrow), and also some smaller extra folds of the tubule (arrowhead). Scale bar = 500 nm. Q. Detail of more distal region of the basal tubule of hadroneme from tentacle of Cerianthidae sp. Smaller folds of the tubule are evident between longer folds of the tubule where spines insert. Scale bar = 500 nm.

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Figure 3.3 122

Figure 3.4. Acanthoneme. Typically classified as basitrichous isorhizas or b-rhabdoids. A. Light microscope image from a tentacle of actinarian Actinostola sp. Scale bar = 20 µm. B. Light microscope image from a tentacle of actinarian Paracondylactis hertwigi. Scale bar = 20 µm. C. Light microscope image from tentacle of Entacmaea quadricolor. Scale bar = 20 µm. D. SEM of nematocyst from tentacle of zoanthid Zoanthus pulchellus. Note the two distinct regions of the tubule, the basal part with large spines and the distal part with short spines. Scale bar = 20 µm. E. SEM of nematocyst from tentacle of antipatharian Antipathes grandis. Scale bar = 5 µm. F. SEM of nematocyst from mesenterial filament of zoanthid Protopalythoa mutuki. Scale bar = 5 µm. G. SEM of nematocyst from acontia of actinarian Calliactis polypus. Scale bar = 2 µm. H. SEM of proximal part of tubule (with large spines) of a nematocyst from a tentacle of Nemathus niditus. Note that the tubule is smooth except for the spine scars indicated by arrows. These mark places where the spines were once attached. Scale bar = 4 µm. I. SEM of the transitional area of nematocyst from a tentacle of actinarian Actinostola sp. A slight change in diameter of the tubule occurs here becoming narrower as it transitions to the distal tubule, a change in size and orientation of the spines also occurs at this transition. Scale bar = 4 µm. J. SEM of the distal tubule of the nematocyst in I that shows the small, hook-like spines on the distal tubule. Scale bar = 4 µm. K. SEM of distal tubule of nematocyst from tentacle of zoanthid Zoanthus pulchellus. Scale bar = 3 µm. L. SEM of basal potion of the tubule of nematocyst from tentacle of Nemanthus niditus. Note the smooth surface of the tubule, arrows indicate apical flaps which open to allow tubule to evert. Scale bar = 4 µm. M. SEM of the apex of the capsule and base of the basal tubule of a nematocyst from tentacle of zoanthid Zoanthus pulchellus. Note the lack of apical flaps. Scale bar = 4 µm.

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Figure 3.4

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Figure 3.5. Colloponeme. Typically classified as b-mastigophores, b-rhabdoids, or sometimes basitrichous isorhizas. All images here are from the acontia although some species also have this type in the tentacles. A. Light microscope image of nematocyst from actinarian Bartholomea annulata. Scale bar = 20 µm. B. Light microscope image of actinarian Cereus pedunculatus nematocysts. Scale bar = 20 µm. C. Light microscope image of an undischarged nematocyst from Aiptasia sp. where the descending tubule is somewhat folded and not rigid and straight. Scale bar = 20 µm. D. Light microscope image of discharged nematocysts from Aiptasia sp. with detail of basal tubule. Scale bar = 20 µm. E. Light microscope image of actinarian Sagartiogeton laceratus showing discharged nematocysts. Because the spines can remain tightly sealed together after discharge, spiral patterns often appear on discharged nematocysts of this morphology. Scale bar = 20 µm. F. SEM overview of a nematocyst of Aiptasia sp. showing the two regions of the tubule, the shaft and distal tubule. Scale bar = 10 µm. G. SEM overview of nematocyst from Sagartia elegans. As in C, the spines are tightly sealed against the shaft region giving the tubule the appearance of having a wider basal section. Scale bar = 50 µm. H. SEM of nematocysts from Metridium senile. The shaft region of these nematocysts can be quite long. Scale bar = 10 µm. I. SEM detail of mid-section of shaft region of a nematocyst of Aiptasia showing spines that are typically under 1 µm long. Scale bar = 3 µm. J. SEM detail of nematocyst from Sagartia elegans with the spines sealed against the shaft. The spines are short here, typically less than 1 µm, and the ridging/annulation of the shaft is apparent between whorls of spines. Scale bar = 2 µm. K. SEM detail of transitional section tubule in nematocyst in D. The ridging/annulation found in the shaft region does not continue into the distal tubule and the spines change form and size. No obvious narrowing of the diameter from shaft to tubule occurs. Scale bar = 5 µm. L. SEM detail of distal tubule of nematocyst in D. No annulations/ridging is apparent here and the spines are short, but robust, knobby or conical in shape. Scale bar = 5µm. M. SEM of the distal tubule of nematocyst from Bartholomea annulata where the knobby shape of the spines of the distal tubule are clear. Scale bar = 3 µm. N. SEM detail of apical region of discharged nematocyst of Aiptasia sp. showing the apical flaps and beginning of the shaft region. Scale bar = 4 µm.

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Figure 3.5

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Figure 3.6. Hadronemes. Typically classified as b-mastigophores or b-rhabdoids. A. Light microscope image of nematocysts from tentacle of Cerianthus filiformis. Note the spiral pattern on the thickened basal tubule and the two distinct sized and shaped nematocysts. Scale bar = 20 µm. B. Light microscope image of nematocyst from tentacle of Cerianthidae sp. Note the elongate shape of the capsule, short thickened basal tubule, and distal tubule indicated with arrow. Scale bar = 20 µm. C. SEM overview of nematocyst from tentacle and Cerianthidae sp. Note the wider basal tubule with large spines and the narrower distal tubule. Scale bar = 20 µm. D. SEM overview of nematocyst from tentacle of Ceriantheopsis americanus. Note the longer basal spines that curl upward. Scale bar = 5 µm. E. SEM detail of basal tubule of nematocyst from Cerianthidae sp. Note the T shape of the spines with a wide base and the fold in the tubule that corresponds to the attachment of the spines. No annulations encircle the entire tubule. Scale bar = 5 µm. F. SEM of transition from basal to distal tubule. Note the reduction in the size of the spines commensurate with the narrowing of the tubule. Scale bar = 4 µm. G. SEM of distal tubule of nematocyst from Ceriantheopsis americanus. Very small spines are evident. Scale bar = 2 µm.

127

Figure 3.6

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Figure 3.7. p-rhabdoid A. Typically classified as p-mastigophore. A. Light microscope image of nematocyst from mesenterial filament of Actinia equina. Scale bar = 20 µm. B. Light microscope image of nematocyst from mesenterial filament of Bunodosoma cavernata. Arrow indicates distal tubule that is present Scale bar = 20 µm. C. SEM of nematocyst from tentacle of coral Platygyra asteriformis. Note the large diameter basal tubule with large spines. A smaller distal tubule is indicated with and arrow. Scale bar = 10 µm. D. SEM of base of nematocyst from mesenterial filaments of Urticina felia. Spines project away from the tubule at right angles and tubule surface in- between spines is smooth. Scale bar = 5 µm. E. SEM of apical end of nematocyst capsule from mesenterial filament of Bunodosoma cavernata. No apical flaps are present. Note the T shape spine with the head of the T attached to the tubule. Scale bar = 1 µm. F. SEM of basal tubule from tentacle of Actinostola sp. Spines have been stripped off in some places revealing the folds of the tubule into which the head of the T of the spine attaches (arrows). The tubule is smooth in-between rows of spines. Scale bar = 5 µm. G. SEM of transition from basal to distal tubule from the tentacle of Actinostola sp. Spines become smaller through the transitional region. The distal tubule is smaller in diameter than the basal tubule and appears to be without spines. Scale bar = 5 µm. H. SEM of distal tubule of nematocyst from tentacle of Platygyra asteriformis. Small hook-like spines are evident on the distal tubule (arrow). Scale bar = 10 µm.

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Figure 3.7

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Figure 3.8. p-rhabdoid B1a. This type has been classified as a p-mastigophore. All images are of nematocysts from mesenterial filaments. A. Light microscope image of nematocyst from Cereus pedunculatus. Note the teardrop shape if the capsule and the thickened descending tubule which almost reaches the distal end of the capsule. Scale bar = 20 µm. B. Light microscope of nematocyst from Sagartiogeton undatus. A v-shaped notch is apparent at the far end of the thickened descending tubule. Scale bar = 20 µm. C. Light microscope image of discharged nematocyst from mesenterial filament of Sagartiogeton undatus. Scale bar = 20 µm. D. SEM of two nematocysts from Diadumene leucolena showing the basal portion of the tubule and the capsule shape. No smaller diameter distal tubule is evident in these nematocysts. Scale bar = 5 µm. E. SEM of spines of the basal tubule of a nematocyst from Nemanthus niditus. Scale bar = 4 µm. F. SEM detail of the basal tubule of nematocyst from Calliactis polypus. Arrow indicates the annulations that encircle this portion tubule. Scale bar = 3 µm. G. SEM detail of the transition of basal to distal tubule in a nematocyst of Acontiaria sp. Scale bar = 4 µm. H. SEM detail of transition of basal to distal tubule in a nematocyst of Calliactis polypus. This particular nematocyst has a smaller diameter distal tubule still attached. Note the lack of annulations and smooth, spineless surface of the distal tubule Scale bar = 2 µm.

131 Figure 3.8

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Figure 3.9. p-rhabdoid B2d. This type is usually classified as an amastigophore (occasionally as a p-mastigophore) or a p-rhabdoid B2a by Schmidt (1969). A. Light microscope image from tentacle of Sagartiogeton lacerates. Note the slight asymmetry of the capsule shape where the left side curves more than the right side. Scale bar = 20 µm. B. Light microscope image of nematocyst from a tentacle of Cereus pedunculatus. The thickened basal tubule extends for almost the whole length of the capsule and ends in an obvious v-shaped notch. Scale bar = 20 µm. C. Light microscope image of discharged nematocyst from mesenterial filament of Sagartia elegans. Note the two regions of the basal tubule, the Faltstück (F) and Hauptstück (H). The Faltstück is shorter than the Hauptstück. Scale bar = 20 µm. D. SEM image of nematocyst from a tentacle of Haliplanella lineata. Both the basal tubule and smaller diameter distal tubule (D) are evident. The basal tubule has two regions, the Faltstück (F) and the Haupstück (H). Scale bar = 10 µm. E. SEM detail of basal tubule of nematocyst from mesenterial filaments of Sagartia elegans. The Faltstück (F) is adjacent to the apex of the capsule and has shorter more sparsely distributed spines than the Haupstück (H). Regular annulations (unlabeled arrow) are evident on the Faltstück. An apical Flap (A) is evident. Scale bar = 5 µm. F. SEM detail of the Faltstück of nematocyst from D. Scale bar = 2 µm. G. SEM detail of the longer, densely arranged spines of the Haupstück on a nematocyst from D. Scale bar = 2 µm. H. SEM detail of the Haupstück of the nematocyst in E, which ends in a conical, unspined section (arrow). No distal tubule is attached. Annulations (A) are evident on the Haupstück. Scale bar = 5 µm. I. SEM detail of transition from basal to distal tubule in the nematocyst from D. Annulations (arrow) are apparent on the basal tubule (Haupstück), but not the distal tubule (which is also unspined). Scale bar = 2 µm.

133 Figure 3.9

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Figure 3.10. p-rhabdoid B2s (variants a and b). This nematocyst type is usually classified as an amastigophore, or as a p-rhabdoid B2a or B2b (depending on relative lengths of Faltstück and Hauptstück). A. Light microscope image of nematocyst from mesenterial filaments of Sagartia elegans. A v-shaped notch is evident at the end of this region of the tubule. The basal tubule is folded in on itself, which is evident in the region indicated by the arrow. Scale bar = 20 µm. B. Light microscope image of nematocyst from acontia of Sagartiogeton undatus. The descending basal tubule is about two-thirds the length of the capsule. Scale bar = 20 µm. C. SEM image of nematocyst from a tentacle of Bartholomea annulata. No distal tubule is attached to a basal tubule segregated into two regions (Faltstück: F, Hauptstück: H). Faltstück is shorter than Hauptstück making this a p-rhabdoid B2s variant a. Scale bar = 20 µm. D. SEM overview of nematocyst from mesenterial filaments of Sagartiogeton undatus. Spines are shorter and more sparsely distributed on the Faltstück (F), than on the Hauptstück (H). Faltstück is shorter than Hauptstück making this a p-rhabodoid B2s variant a. Scale bar = 15 µm. E. SEM overview of a nematocyst from a mesenterial filament of Diadumene sp. Faltstück (F) is longer than Hauptstück (H) in this nematocyst making this a p-rhabdoid B2b. Also, the distal tubule is attached (D). Scale bar = 50 µm. F. SEM of basal-most region of the Faltstück in nematocyst from an aconitum of Metridium senile. Apical flaps (A). Scale bar = 5 µm. G. SEM of the Faltstück of a nematocyst from acontium of Cereus pedunculatus. Regular annulations that encircle the tubule (arrow). Spines of this region thin, flat, and have a narrow blade. Scale bar = 4 µm. H. SEM of Faltstück of nematocyst in E. Spines are short, thin, flat with a narrow blade. Circular indentations (arrows) in the tubule are evident in-between rows of spines and interrupt the annulations. Scale bar = 5 µm. I. SEM of transition from Faltstück to Hauptstück in nematocyst in E, marked by the change in size of the spines. Scale bar = 5 µm. J. SEM of transition from Faltstück (F) to Hauptstück (H) in a nematocyst from a mesenterial filament from Aiptasia sp. The two regions are distinguished by spine length, and by the width of annulations in the tubule surface. Some of the spines of the Hauptstück have been stripped off during discharge. These spines have a slightly broader base, and a blade that narrows before widening. Scale bar = 5 µm. K. SEM of spines of the Hauptstück on the nematocyst from E. These spines always form an acute angle with the tubule and are never perpendicular to it. Scale bar = 5 µm. L. SEM detail of Hauptstück of a nematocyst from acotium of Aiptasia sp. These spines are long and thin and typically curl up at the ends. Scale bar = 5 µm. M. SEM detail of the transition from basal to distal tubule in a nematocyst from an aconitum of Aiptasia sp. Arrow indicates the unspined transitional region. Scale bar = 5 µm. N. SEM detail of transition from basal to distal tubule in nematocyst from E. The regular annulations of the Hauptstück do not continue on the distal tubule. Scale bar = 5 µm. O. SEM detail of transitional region of a nematocyst that has no distal tubule still attached. When the distal tubule breaks off, an unspined conical region with no annulations (unlike the Hauptstück) remains at the tip of the tubule (arrow). From a aconitum of Cereus pedunculatus. Scale bar = 4 µm. P. SEM detail of distal tubule of nematocyst in M. The distal tubule is smooth with no spines or folds. Scale bar = 5 µm. Q. SEM detail of distal tubule of nematocyst in E. No spines are 135 present, though folds in the surface are visible, interrupted by creases (arrows) that spiral around the tubule. Scale bar = 5 µm.

136 Figure 3.10

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Figure 3.11. Diakaneme. All images are of nematocysts from mesenterial filaments of Urticina felia. A. Light microscope image of nematocyst with the characteristic shape of this morphology. The non-apical end is widest and the thickened basal tubule extends a little more than half the length of the capsule. Scale bar = 20 µm. B. Light microscope image nematocyst in which the v-shaped notch (arrow) is apparent at the end of the descending basal tubule. Scale bar = 20 µm. C. SEM image showing the overall morphology of this type. Spines are sparse in the apical adjacent region of the basal tubule, and are more densely distributed further along the tubule. A distal tubule is always present. Scale bar = 20 µm. D. SEM detail of apical adjacent region of the basal tubule. The base of the spine (arrow) is wider and the blade narrows uniformly along the length (therefore no T shaped base is apparent). Overall these spines are long and very narrow. The tubule is smooth with no annulations. Scale bar = 5 µm. E. SEM detail of the transitional region of the basal tubule where spines change size. Although the large spines have a similar shape, they become wider, thicker and generally larger distant to the apex of the capsule. These spines often flare outward away from the tubule but are not typically perpendicular to it. Scale bar = 5 µm. F. SEM image of the basal tubule distant from the apex of the capsule where large spines are found. Spines are wider at the base but are not T-shaped. Scale bar = 4 µm. G. SEM image of transition from basal to distal tubule. The tubule narrows and the spines become somewhat smaller in size before abruptly stopping. The tubule is smooth in between whorls of spines, spine scars (arrow) are evident. Scale bar = 5 µm. H. SEM image of distal tubule. The surface is smooth and bears no spines. Scale bar = 5 µm.

138 Figure 3.11

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Figure 3.12. Aphylloneme. All images are of nematocysts from the mesenterial filaments of Anemonia sulcata. A. Light microscope image of nematocyst with characteristic elongated capsule shape with a wider non-apical end. No v-shaped notch is present at the end of the basal tubule. Scale bar = 20 µm. B. SEM overview of the aphylloneme morphology. The basal tubule has long spines that are typically stripped off so it does not have a full, bushy appearance. Scale bar = 30 µm. C. SEM detail of apical end of nematocyst and the spines on the basal tubule. Two apical flaps (arrows) are evident. Scale bar = 5 µm. D. SEM detail of transition from basal to distal tubule. The diameter of the tubule does not change noticeably but the spines are no longer present. Scale bar = 4 µm. E. SEM detail of distal tubule. The aphylloneme bears no spines on the distal tubule. Scale bar = 2 µm. F. SEM detail of long spines on the middle region of the basal tubule. Scale bar = 5 µm.

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Figure 3.12

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Figure 3.13. Morphology and ultrastructure of hydrozoan (medusozoan) rhopaloid nematocysts (those with basal tubule that changes in diameter). A. SEM of a stenotele nematocyst from a tentacle of Hydra sp. The tubule has a basal region that changes in diameter and a distal tubule (arrow) with a consistent, smaller diameter. The tubule surface is smooth with no annulations or folds where the spines attach. Scale Bar = 3 µm. B. SEM of a eurytele nematocyst from a tentacle of Cordylophora caspia. This nematocyst morphology has a characteristic bulge (arrow) in the basal tubule. The tubule surface is smooth with no annulations or folds where the spines attach. Scale Bar = 2 µm. C. TEM of stenotele nematocyst from a tentacle of Sertularella sp. The basal tubule is smooth around the spines packaged more apically, but has folds toward the non-apical end (arrow). Scale Bar = 500 nm. D. TEM of stenotele from tentacle Ectopleura larynx. The basal tubule contains spines shorter than those seen in Anthozoans, packaged against each other. The tubule is smooth apically with regular folds towards the non-apical end (arrow). Scale Bar = 1 µm.

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Figure 3.13

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Chapter 4: Distribution of nematocyst morphology and implications for phylogeny and evolution

Introduction

As one of the earliest branches on the metazoan tree, cnidarians represent an ancient lineage (e.g. Edgecombe et al. 2011). Members of this group share a simplified body form with two embryonic cell layers (Hyman 1940) and limited morphology to use for phylogenetic study and taxonomic classification. However, all members of the phylum Cnidaria produce nematocysts, and typically produce forms of these structures that are highly variable in capsule shape and size, in tubule morphology, and in the size, shape, and distribution of spines (e.g. Weill 1930, 1934, Schmidt 1969, 1974, Mariscal

1974). Additionally, cnidarians typically have distinct nematocyst morphologies in different regions of the body; together these comprise the cnidome, which may also vary between species (Weill 1934).

Classification schemes (e.g. Stephenson 1928, Weil 1934, Schmidt 1969, den

Hartog 1977) have been created to organize and interpret this variation. In all classification systems, nematocysts are classified as a particular type and the presence or absence of the type is assessed for each body region that bears nematocysts (Carlgren

1949). Disagreements on the relative importance of different characteristics of nematocysts, differences in how to interpret some features, and a lack of resolution of critical features with light microscopy has led to a proliferation of classification systems,

144 resulting in confusion (see Chapter 3). Two competing systems are most widely used: that of Weill (1930, 1934), modified by Carlgren (1940) and others (see Mariscal 1974), and that of Schmidt (1969, 1972, 1974). The two systems classify the same morphology in different ways, sometimes joining morphologies that the other system considers separate (as with the b-rhabdoides of Schmidt’s (1969) system), or subdividing more general categories (as with p-rhabdoides of Schmidt (1969) (see England 1991 and

Chapter 3 for further discussion). The distillation of the complex morphology into a single presence/absence character (the definition of which varies from author to author) creates problems when synthesizing past work on the distribution of nematocyst diversity in the phylum.

Nematocyst morphology has the potential to serve as an important source for taxonomic and phylogenetic characters, and in fact, is often treated this way (Seifert

1928, Weill 1934, Carlgren 1900, 1940, 1945, Schmidt 1969, 1974, Östman 1982, 1983,

1987, Picciani et al 2011). Nematocysts have been particularly important in the taxonomy and systematics of hexacorallians (Cnidaria: Anthozoa: Hexacorallia), and especially in the hexacorallian order Actiniaria (Fautin 1989). For example, nematocysts are an important component in the definition of actiniarian families (Carlgren 1949), and have been used as evidence for scenarios of evolution of various hexacorallian groups

(Hand 1956, Schmidt 1969, 1974, England 1987, den Hartog 1977, Picciani et al 2011).

However, whether nematocyst characters are phylogenetically informative for

Actiniaria is not yet clear. The confusion over nematocyst morphology makes large-scale conclusions about nematocysts as synapomorphies difficult. For example, Shostak and

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Kolluri (1995) attempted to use literature surveys to determine the evolution of cnidomes within Cnidaria, but in the process, mixed data from Carlgren, who uses Weill’s (1934) system and Schmidt, who uses a system that is not entirely compatible with that of Weill

(1934), with no acknowledgement of the disagreement. This conflation of systems severely weakened their conclusions about which nematocyst morphologies are shared.

Furthermore, the advent of modern phylogenetic analyses of molecular and morphological data has revealed that the taxonomic classification of sea anemones

(Carlgren 1949) does not reflect the evolutionary history of the group very closely; many of the higher-level taxa are non-monophyletic (Daly et. al 2008, 2010; Rodriguez et al.

2008, 2012). To assess the phylogenetic value of cnidomes and of nematocyst morphology requires both a clear understanding of the morphology and its distribution in the taxa and a well-supported hypothesis of relationships among the taxa. To achieve the former goal, I determined nematocyst morphology using light and scanning electron microscopy from a diversity of actiniarians, enabling me to fully characterize morphological diversity. I built a robust phylogeny for the taxa under study using two mitochondrial and one nuclear markers, and then mapped the nematocyst characters onto the phylogenetic tree. This enabled me to determine if changes in nematocyst morphology and cnidome track phylogenetic change.

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Materials and Methods

Materials

I examined nematocysts from species belonging to four lineages of Anthozoa: 26 actiniarians, 2 antipatharians, 3 cerianthids, 3 corallimorphorpharians, 5 scleractinians, 2 zoanthideans, and 2 octocorals (Table 4.1). Specimens were collected from the wild

(Table 4.1) or obtained from commercial sources such as Aquarium Adventure

(Columbus, OH), Gulf Specimen Marine Lab (Panacea, FL), Marine Biological Lab

(Woods Hole, MA) and Reef Hot Spot (Inglewood, CA). The stoloniferan octocoral

Clavularid sp. A (sensu Parrin et al. 2010) and the zoanthidean Protopalythoa mutuki were obtained from cultures maintained by Neil Blackstone at Northern Illinois

University. The antipatharian samples were collected, fixed, and provided by Anthony

Montgomery (see Opresko, 2009 for collecting details). Species identifications were determined by the collector or verified in the case of commercial sources.

A few samples were problematic in terms of their identification. One specimen originally identified as Stomphia coccinea (Müller 1776) has nematocysts in the tentacles (p-rhabdoid A), which are inconsistent with the cnidome attributed to the species circumscription (Carlgren 1940); the occurrence of this nematocyst indicates that the specimen may belong to Actinostola which has this nematocyst (Carlgren 1945,

Fautin 1984) and is inferred to be the sister to Stomphia (Rodríguez et al. 2012).

Furthermore, Ross and Sutton (1967), in studying specimens collected in the same

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locality, acknowledged that there is an unnamed Actinostola that is easily confused with

S. coccinea. Because Stomphia and Actinostola are well-supported as sister taxa

(Rodríguez et al. 2012), the interpretation of nematocyst characters will not be affected

by the small change in position in the tree. This specimen is labeled as Actinostolid sp.

in Table 4.1 and is treated as an Actinostola in the phylogenetic tree. Another specimen

was originally identified as Flosmaris mutsensis, but DNA and nematocyst characters,

such as the p-rhabdoides B2d in the tentacles (Song 1992), do not support this taxonomic

assignment; it is referred to as Acontiarian sp. Finally, the cerianthid remains

unidentified beyond family (listed as Cerianthidae sp.).

Methods

Sample preparation

After collection, all actiniarian samples were dissected to isolate key body regions: mesenterial filaments, body column, tentacles, and acrorhagi or acontia if present (see

Table 4.2 for body regions surveyed). Non-actiniarian specimens were sampled primarily for tentacles and/or mesenterial filaments (see Table 4.1). Some of these body regions, particularly the column, can have specialized sub-regions that may contain nematocysts of distinct morphologies (e.g. Rodríguez and López-González 2005, Zelnio et al. 2009), Furthermore nematocysts have been demonstrated to vary along the length of column and some nematocyst morphologies are rare in certain regions (Fautin 1989).

Thus, I attempted to collect from the upper half of the column in all samples and avoided specialized regions (other than acrorhagi, which were collected in addition to the body

148 column when present). However, to obtain samples from a wider taxonomic scope, each body region may have been sampled only once; therefore, rare nematocysts or those found in specialized parts of these body regions may have been missed.

After dissection, samples for SEM and light microscopy were placed in a 1M sodium citrate solution for 10-15 minutes (larger tissues required more time) to induce expulsion of nematocysts from nematocytes. Samples were washed three times with distilled water and most were then placed in 70% ethanol. Samples from Bunodosoma cavernata and Metridium senile were placed in a 1% OsO4 solution overnight before being placed in 70% ethanol. SEM samples were dehydrated in ethanol and then critical- point dried with CO2. Samples from B. cavernata and M. senile were sputter-coated with gold-palladium in a Hummer sputter coater and examined using a LEO 1550 field emission scanning electron microscope at the University of Kansas, Lawrence. Three scleractinian samples (Echinophyllia aspera, Caulastrea tumida, and Acanthastrea sp.) were processed using a Freeze Dryer (Hitachi ES-2030) rather than a critical point drier after being dehydrated through an ethanol series. These samples were sputter coated with gold-palladium in a Hitachi E-1030 sputter coater, and examined with a Hitachi S-4300

SEM at Kyoto University’s Seto Marine Laboratory, in Shirahama Japan. All other samples were sputter-coated with gold-palladium or palladium in a Cressington sputter coater and examined using a FEI NOVA nanoSEM at The Ohio State University,

Columbus. Squash preparations of undischarged capsules were made using samples fixed for SEM as above, using samples taken from ethanol before critical-point drying. For these light microscopy samples, a very small piece of tissue was floated in a droplet of

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water and then compressed between a coverslip and a microscope slide; squash

preparations were examined under DIC at 1000X.

The nematocyst morphotypes distinguished here are based on the thorough

morphological analysis performed in Chapter 3 with the addition of two more types, the

macrobasic amastigophore, and the holotrichous isorhiza, or holotrich. The macrobasic

mastigophore is defined as having a thickened basal tubule that is so much longer than

the capsule that twists and folds are required for it to fit within the capsule. The holotrich

is identifiable using light microscopy by the absence of any differentiation of the tubule

(i.e. no thickened descending tubule as the tubule appears to be the same throughout the

capsule). Figure 4.1 summarizes the features visualized through light and scanning

electron microscopy that identify the ten morphotypes used here. As in Chapter 3, the

names of each morphotype reflect a combination of classification systems, primarily

those of Weill (1934, modified by Carlgren 1940, Mariscal 1974) and Schmidt (1969).

Phylogeny

To place the nematocyst morphologies in a phylogenetic context, I built a tree of actiniarian taxa on which to map nematocyst morphologies and cnidome characters. The

DNA matrix of Rodríguez et al. (2012) was obtained and all taxa were included (see

Table 4.2), however only three of the five genes were used: 16S, 18S, 28S. Eight additional taxa were included : Aiptasia sp., Bunodosoma cavernata, Condylactis gigantea, Diadumene sp. Korea, Entamaea quadricolor, Epiactis prolifera,

Paracondylactis hertwigi, and Urticina felina. Some of these new sequences were

150 obtained from Genbank and others were collected specifically for this study (see Table

4.2).

For new sequences, genomic DNA was obtained from column tissue following the “Rat tail” protocol supplied with the Qiagen DNeasy kit. Standard techniques and published primers (Daly et al. 2008) were used to amplify template DNA. Both mitochondrial (16S) and nuclear (18S, 28S) partial sequences were obtained and sequenced using an ABI 3730xl by staff at the sequencing facility of Beckman-Coulter

Genomics, Danvers, MA. Forward and reverse sequences were assembled in Sequencher ver. 4.8 (Gene Codes Corp., Ann Arbor, MI) and blasted against the nucleotide database of Genbank to ensure that target locus and organism were sequenced rather than symbiont or other contaminant.

New sequences were aligned to those of the Rodríguez et al. (2012) matrix using

Muscle (Edgar 2004). The concatenated dataset was analyzed in RAxML 7.2.8

(Stamatakis 2006) with default settings (GTR + Γ) and rapid hill climbing using RAxML blackbox (Stamatakis et al. 2008). Each run produced 100 bootstrap analyses and clade support was assessed by combining the bootstrap analyses of ten independent runs in

Consense, part of the Phylip v. 3.69 package, to obtain a total of 1000 bootstrap replicates. The zoanthidean Savalia savaglia served as the outgroup. Two long-branch taxa, Boloceroides mcmurrichi and Acontiaria sp., were sequentially removed and the analysis re-run to determine their respective positions in the tree.

Nematocyst morphotypes of all taxa were manually mapped onto ML tree for the cnidome overall and for each body region. For the few focal taxa for which I lacked

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DNA suitable material (Actinia equina, Anemonia sulcata, and Actinostolid sp.), I included sequences from a representative of the same genus (respectively Actinia fragacea, Anemonia virdis, and two species of Actinostola). Actinia fragacea is closely related to Actinia equina and shares the complement of nematocyst types (Carter and

Thorpe 1981), Anemonia sulcata and A. virdis are part of the same species complex

(Bulnheim and Sauer 1984, Wiedenmann et al 2000), and the Actinostola species has already been addressed (see materials). One specimen was not identified beyond belonging to the group Acontiaria, but is labeled as Acontiaria sp. in tables and trees.

Shared characters were placed on branches only if they could be unambiguously optimized to that branch. Some characters were ambiguous and were therefore not specifically mapped. Autapomorphies were not noted on branches but are evident as a change in color changes or the addition of a dot to signify the gain or loss of a nematocyst morphology.

A broader perspective on the evolution of nematocyst morphology and cnidome is gained through mapping nematocyst morphologies onto a published tree of hexacorallian relationships (Brugler and France 2007).

Results

Variation within nematocyst morphotype

For each body region studied, nematocysts were categorized into one of nine morphotypes based on features visible under light microscopy, such as capsule shape and features of the basal tubule that correspond to features visible under SEM, such as surface

152 tubule features, spine shape, etc (see Chapter 3). In the process of determining the cnidome of each species, I observed significant variation in capsule and tubule shape under the light microscope; this variation had little or no consequence in terms of features observed under SEM. I summarize here some of the variation visible with light microscopy of undischarged capsules which does not affect their placement in the morphotype category based on SEM data.

Acanthoneme: Capsule shape and size varies considerably in this morphotype. Often elongate as in Fig. 4.1A, it can also take a more rounded form (Fig. 4.2A, B). The basal tubule may span the length of the capsule (Fig. 2B), or only part of the length (Fig. 4.2A,

C, D). The descending tubule is usually straight, but can be dramatically curved (Fig.

1C). Usually, the widest point of the undischarged capsule is at mid-length, but in some examples, the apical end is wider (Fig 4.2D).

Aphylloneme: This morphotype generally varies little in form. It is always larger than 20

µm and is distinctively wider at the non-apical end (Figs. 4.2E-G). The descending tubule often appears to be patterned (Figs. 4.2E, F,), but occasionally does not (Fig.

4.2G).

Colloponeme: This morphotype varies in size, but has a consistent elongate shape with the midpoint in length being the widest part of the undischarged capsule (Fig. 4.2H-J). In some instances, the descending tubule appears rigid and straight (Fig. 4.2J), but in others it appears flexible and slightly wavy (Fig. 4.2I).

Hadroneme: This morphotype varies considerably in size and shape, from ovoid (Fig.

4.2K) to elongate (Fig. 4.2L). Typically, the basal tubule is robust, has a corkscrew

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pattern (Fig. 4.2K), and narrows distinctly at the non-apical end (Fig. 4.2M). However,

in smaller or more elongate capsules, the basal tubule is less robust and any pattern or

narrowing is less obvious (Fig. 4.2L).

p-rhabdoid A: This morphotype can have a more elongate (Fig. 4.3A-C) or a more ovoid

form with a wide midpoint (Fig. 4.3D, E). Even in the elongate form, it never is as

slender as 1 and 3. Usually, some material, often granular in appearance, fills the

capsular space (Fig. 4.3A-E). The descending tubule passes the widest point of the

capsule before ending in a v-shaped notch.

p-rhabdoid B1a: The capsule of this morphotype generally has an egg or teardrop shape

in which the apical end is narrow and the capsule flares dramatically towards the

midpoint (Figs. 4.1J, 4.3F). However, the capsule can be ovoid, almost round (Fig.

4.3G), or more elongate (Fig. 4.3H). In Nemanthus nitidus, this morphotype is

considerably elongated, with the widest portion near the midpoint (Fig. 4.3I). In the more

elongate form (Fig. 4.3I), the descending shaft does not span the capsule as it does in

other forms of this morph (Fig. 4.3F-H). This elongate variant is common but extreme in

form; other p-rhabdoid B1a nematocysts in N. nitidus more closely resemble the normal

morphology (Fig. 4.3J).

p-rhabdoid B2d: The capsule of this morphotype has an asymmetric shape in which one side of the capsule curves more than the other (Fig. 4.3K-N). The degree to which the two sides differ can be extreme (Fig. 4.3K-M) or subtle (Fig. 4.3N). Often, as a result of this curve, the descending tubule angles to one side, appearing to be closer to one side of the capsule (Fig 4.3K, L).

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p-rhabdoid B2s (variant a): The capsule of this morphotype is symmetrical when compared to the p-rhabdoid B2d as both sides of the capsule appear to curve to the same degree. The descending tubule is typically centered in and often as long as the capsule

(Fig. 4.3M, N), although the tubule may descend to just past the midpoint of the capsule

(Fig. 4.3O). In one species, two variants of p-rhabdoid B2s were observed in the same tissue: one, in which the descending tubule is straight (Fig. 4.1N) and one in which the tubule is slightly longer and thus looped to fit within the capsule (Fig. 4.3P). The length of this loop varies but is never as extensive as it is in the macrobasic amastigophore (Fig.

4.1R). Because this variant differs from others of the same morph in the same tissue only by the slightly longer tubule, the two variants could not be distinguished when discharged. However, extensive SEM study revealed no structural differences among any of the nematocysts of this general form, suggesting that the variant with a slightly longer tubule does not differ structurally from those having a shorter tubule.

Diakaneme: This morphotype varies only in the distinctiveness of the v-shaped notch at

the end of the basal tubule. In size and shape, this form is consistent.

macrobasic amastigophore: This morphotype varied with respect to the number and form

of the twists and loops of the descending tubule. No SEM data was obtained of this

morphology, which was only observed in one taxon (in tentacles only of Diadumene sp.

Korea).

Holotrich: This morphotype was observed to have three basic forms. In one, found in

acrorhagi of actiniarians, the capsules are long and thin, with the tubule usually

discernible (Fig. 4.4A) and often packaged in loops inside the capsule (Fig. 4.4B). SEM

155 reveals the presence of very small spines flush to the surface of the tubule and knobby in shape (Fig. 4.4C). Also, the tubule is smooth between whorls of spines. In the octocoral

Renilla mulleri, the nematocysts are small, have small diameter tubules (Fig. 4.4D), and are similar to the actiniarian examples in having small spines that are flush to the surface

(4.4D, E).

Another form in the zoanthideans has a capsule less elongate than wide and the tightly packed tubule fills the undischarged capsule (Fig. 4.4F). In the undischarged form, the tubule bears spines that project away from the surface (Fig 4.4G) and have a characteristic form. These spines are thick and three dimensional rather than flat, and have a visible central groove (Fig. 4.4H, I). This spine form is seen in both large examples of zoanthidean holotrichs (Fig. 4.4I) and small ones (Fig. 4.4J), however the tubule surface is smooth in the small ones but has extra folds in the large ones. The base of the spine is broader giving the spine a T-shape (Fig. 4.4I).

The final holotrich form is seen in both corallimorpharians and scleractinians.

The capsule of the undischarged holotrich may be ovoid (Fig. 4.4K) or elongate (Fig.

4.4L), but both are tightly packed with a wide diameter tubule. The spines on the undischarged tubule are stiff and usually project at a ninety-degree angle from the tubule surface (Fig. 4.4M, N). The spines are flat rather than three-dimensional and form a T- shape with the wide base of attachment (Fig. 4.4O, P). Although the spines and tubule may be smaller in scleractinians, the form is the same (Fig. 4.4P). Whether large or small, the tubule surface is always smooth with no evidence of extra folds (Fig. 4.4O, P).

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Distribution of types

Although there are some commonalities across lineages, at least some nematocyst morphotypes are taxonomically restricted (Table 4.3), and others are functionally or otherwise restricted in terms of their distribution in actiniarian tissues (Table 4.4). The acanthoneme is the only morphotype found in all families studied here (Table 4.3). In contrast, the aphylloneme and diakaneme are only found in Actiniidae, and the colloponeme, p-rhabdoid B2d and B2s are characteristic of different acontiate families

(Table 4.3). Acanthonemes and p-rhabdoid B2s are reported from all tissues (Table 4.4), although they are not always in the same tissues in all taxa and may not occur in some lineages (Table 4.3). Other morphs are restricted by body region: for example, the p- rhabdoid B2d is not seen in the acontia, the colloponemes are typically found only in acontia (and occasionally in tentacles), p-rhabdoid As and p-rhabdoid B1as are usually restricted to mesenterial filaments (with a few exceptions), and the aphylloneme and diakaneme are found only in mesenterial filaments.

Phylogenetic tree

As has been seen in other DNA-based phylogenetic analyses (e.g., Daly et al. 2008,

Rodriguez and Daly 2010, Rodriguez et al. 2012), I find strong support for the monophyly of Edwardsiidae, Endomyaria, and the superfamily Metridioidea (Fig. 4.5).

Actinostolina is paraphyletic with respect to Endomyaria and Metridioidea. Within

Metridioidea, support is high for a monophyletic grouping of Basitrina, Graspina, and

Phellina (as defined in Rodriguez et al. 2012). A clade within Metrioidea with consistent

157 albeit low support includes members of the families Diadumenidae, Haliplanellidae,

Metridiidae, and Sagartiidae (labeled as Clade A in Fig. 4.5). Across the tree, bootstrap support is generally higher for shallower groups and weakest at the deeper nodes.

Two analyses were run to see the effect of taxa with highly divergent sequences

(Boloceroides mcmurrichi and Acontiaria sp.) on the topology. These taxa are sisters in the tree based on the full matrix. The removal of Boloceroides mcmurrichi affected only the placement of Acontiaria sp., which is also a long branch. In the absence of B. mcmurrichi, Acontiaria sp. is sister to Sagartia troglodytes. In the analysis run without

Acontiaria sp., Boloceroides mcmurrichi is the sister to a group including members of the genera Metridium, Diadumene, and Haliplanella.

Mapping nematocyst characters

Based on the current hypothesis of hexacorallian relationships (Brugler and France 2007),

I infer that acanthonemes and p-rhabdoides A arose after the split between Ceriantharia and all other hexacorals, and are thus homologous across Actiniaria, Antipatharia,

Corallimorpharia, Scleractinia, and Zoanthidea (Fig. 4.6). Although the exact relationship of homology among holotrichs cannot be decisively determined given the tree here, morphotypes of holotrichs are not inferred to be homologous among lineages: holotrich 1 is most parsimoniously mapped as arising twice, once in octocorals and once in Actiniaria, and there is no evidence to suggest that holotrichs 2 and 3 share an evolutionary history.

158

Acanthonemes are inferred to be the common element of the cnidome of tentacles

in Actiniaria (Fig. 4.7), being common to the tentacles of all taxa studied here. Clade A

shows the greatest diversity of morphotypes in the tentacles, with two basic complements

of morphotypes: 1) acanthonemes and p-rhabdoides B2d; and 2) Acanthonemes, p-

rhabdoides B2d and B2s (variant a). The few other nematocyst morphotypes that occur

in tentacles are autapomorphies for particular taxa or lineages.

I observe five cnidomes of mesenterial filaments of actiniarians (Table 4.4, Fig.

4.8): 1) acanthoneme, p-rhabdoid A; 2) acanthoneme and p-rhabdoid B1a; 3) acanthoneme, p-rhabdoid B1a, and B2s (variant a); 4) acanthoneme, p-rhabdoid B1a, b2d, and B2s (variant a for all but Diadumene species that have variant b); and 5) acanthoneme, p-rhabdoid B2d and B2s (variant a). The aphylloneme is found in the filaments of a few endomyarian taxa but cannot be unambiguously mapped: it is equally parsimonious given the taxa studied here to infer that it arose once (with two losses) or arose three times (Fig. 4.8). The clade Metridioidea is defined by the loss of p-rhabdoid

A and the gain of p-rhabdoid B1a in the filaments (Fig. 4.8).

Among actiniarians that do not belong to Clade A, acanthonemes constitute the primary cnidom for the body column (Fig. 4.9), although some taxa also have p- rhabdoides A. Clade A is defined by the gain of the p-rhabdoid B2b in the column (Fig.

4.9); the family Aiptasiidae loses this morph and gains p-rhabdoid B2s (variant a). Two other taxa gain p-rhabdoid B2s (variant a) as autapomorphies.

Acrorhagi and acontia are nematocyst-dense structures found only in subsets of

Endomyaria and Metridoidea, respectively; both structures are unique to Actiniaria and

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they never co-occur in an animal. Acrorhagi contain holotrichs alone or holotrichs and

acanthonemes (Fig. 4.10). Whether the acanthonemes are ancestral to this structure or

have been gained and lost several times is difficult to determine with the limited amount

of sampling of the acrorhagi. The acontia of members of the Basitrina have only

acanthonemes, whereas the acontial cnidome of members of Clade A comprises

colloponemes and p-rhabdoides B2s (variant a). Within Clade A, the two species of

Diadumene are distinguished by having variant b of p-rhabdoid B2s.

Discussion

Trees

Although the tree in this study used a subset of the data in Rodríguez et al. (2012) and is generally very similar in overall topology, the two trees differ in the placement of taxa within Metridioidea. The deep-sea and chemosynthetic taxa (Jasonactis erythraios,

Alvinactis chessi, Cyananthea hourdezi, and Kadosactis antarctica) are sister to all other

Metridioidea (Fig. 4.5); in contrast to the tree of Rodríguez et al. (2012), in which these taxa are sister to the Basitrina + Graspina + Phellina group within Metridioidea.

Furthermore, although the members of Clade A here and the Clade 1 of Rodríguez et al.

(2012) are mostly the same, some taxa, such as the Aiptasiidae and a few Sagartiidae fall out of the group in the latter tree. However, these differences occur in areas with low bootstrap support for both trees, indicating that a stable, well-supported tree topology has

yet to be found. In both analyses, the Basitrina + Graspina + Phellina grouping has high

bootstrap support; this support is corroborated by morphological data that further

160 supports the group as distinct (see below). In Rodríguez et al. (2012), Nemanthus nitidus lies within a monophyletic Basitrina, whereas my analyses (Fig. 4.5) place it between

Graspina and Phellina, all of which are nested within Basitrina (making it paraphyletic).

Again, these are locations in the tree with low bootstrap support on both trees.

The placement of Acontiaria sp. is unresolved. This specimen was initially identified in the field as Flosmaris mutsensis but both DNA and nematocyst data clearly distinguish it from the other Isophellid (Telmatactis sp.) in the tree. The nematocyst data support its placement in Clade A. The DNA data are inconclusive: this taxon has very divergent sequences and groups with either the other long-branch species or with the sagartiids Sagartia and Cereus (Fig. 4.5), a position that makes sense in light of the nematocyst characteristics: like Sagartia elegans and Cereus pedunculatus, this species has acanthonemes and p-rhabdoid B2d in its tentacles. However, other nematocyst characters differ, including the cnidome of the mesenterial filaments and body column.

Obtaining more molecular information (this taxa is missing 28S data) may resolve the position of this taxon more effectively.

Boloceroides mcmurrichi has a relatively simple morphology, and lacks acontia, and has historically been considered an early branch in the actiniaria tree (Carlgren 1924,

1949, Schmidt 1974). However, DNA-based phylogenetic analyses have consistently placed this taxon with acontiate anemones in the clade now called Metridioidea (Daly et al. 2008, Rodríguez and Daly 2010, Rodríguez et al. 2012). The nematocyst data agrees with this interpretation; it has has both p-rhabdoides B2d and B2s (variant a), morphotypes restricted to Metridioidea. Schmidt (1969) noted that B. mcmurrichi lacks

161 p-rhabdoides A which is significant in that p-rhabdoides A appear to be present at the base of the actiniarian tree but is absent in the Metridioidea. Because this species has a highly divergent sequence, its precise placement is unclear, but its placement as sister to

Metridiidae + Diadumene + Haliplanella (when Acontiaria sp. is removed from the matrix) is supported by the cnidome of the body column. This placement is contradicted by the cnidome of the tentacles: B. mcmurrichi has an additional nematocyst morphotype

(p-rhabdoid B2s).

Anthozoan cnidomes

Although my sampling of body tissues of non-actiniarians was not exhaustive, the data collected here agrees with the literature in terms of the overall cnidome for the other hexacorallian orders with the exception of some rare types that I did not observe

(Carlgren 1940, Schmidt 1974, Ryland and Lancaster 2004). From these data a few important conclusions can be made. First, the ancestral cnidome for Actiniaria likely comprised acanthonemes and p-rhabdoides A, and possibly included holotrichs 1.

Although I did not observe acanthonemes in corallimorpharians, previous work by

Carlgren (1940, 1945) suggests that some corallimorpharians do bear this morphology.

Therefore, all hexacorallians other than cerianthids have these two morphologies, though some features can vary (Schmidt 1974, Picciani et al. 2011, see Chapter 3).

The cnidome of Ceriantharia is distinctive, and not just in lacking acanthonemenes and p-rhabdoides A. This lineage is the only cnidarian group in which ptychocysts occur; this kind of cnida is inferred to be distinct from nematocysts, and

162 differs from nematocysts in capsule and tubule morphology (Mariscal et al. 1977). Two of the three families of cerianthids have only one type of nematocyst, the hadroneme; members of the family Arachnactidae have this and an additional form (Carlgren 1940), defined by den Hartog (1977) as a pencillus (or mastigophore) type. Although no comment can be made on this additional form, the hadroneme is shared amongst the three families and is likely to be ancestral for the group. As previously discussed (Chapter 3), the hadroneme has intriguing similarities to the p-rhabdoid A, particularly in spines and spine attachment, and when phylogeny is considered (Fig. 4.6), the p-rhabdoid A can be interpreted as the sister to the hadroneme (in fact, Cutress (1955) called this morph a p- mastigophore). Daly et al. (2003) claimed that b-mastigophores were part of the ancestral cnidome of hexacorals, but this conclusion was based on homologizing hadronemes and acathonemes as b-mastigophores under the Weill (1930, 1934) classification scheme.

Based on this more detailed analysis, it seems more likely that the hadroneme/p-rhabdoid

A form is ancestral for hexacorals.

The phylogenetic analyses also indicate that the various forms of holotrich have arisen independently within Hexacorallia. As Cutress (1955) observed, the category

“holotrich” is heterogenous; I find at least three distinct morphologies among hexacorallians. The holotrichs of the zoanthideans are not similar to those of scleractinians and corallimopharians, differing minimally in spine structure. Although

Cutress (1955) suggested that there is a large holotrich morphology in common amongst these three lineages that might represent a “prototype of all other nematocysts” (p. 135), the morphological and phylogenetic evidence does not support homologizing these

163 forms. Although sampling in Actiniaria was not exhaustive, the holotrichs studied here do not resemble those of zoanthideans, corals, or corallimorphs, but rather resemble the holotrichs of octocorallians. Furthermore, large holotrichs sensu Cutress (1955) are not known to occur in Actiniaria (Daly et al. 2003). Therefore, the “holotrich” form has two additional forms in Hexacorallia that must have arisen independently.

Determining what is ancestral for Hexacorallia is difficult given the confusion about which nematocyst types occur in its sister group, Octocorallia. Minimally, octocorals have holotrichs; in the literature, this morphology has been called an atrichous isorhiza (Weill 1934, Carlgren 1940, Hyman 1940), as the spines are too small to visualize with the light microscope (Westfall 1965). At least this form is present ancestrally as hypothesized by Carlgren (1940) and supported by Daly et al. (2003).

Others have claimed that this single morphotype is a b-mastigophore/rhabdoid (Schmidt

1974, Shostak and Kolluri 1995) or a basitrich (Cutress 1955). Although most studies

(e.g., Weill 1934, Carlgren 1940, Hyman 1940, Cutress 1955, Schmidt 1974, Sammarco and Coll 1992, Shostak and Kolluri 1995) report that octocorallians bear a single morphotype (regardless of the name used), at least some species have multiple morphotypes (e.g., Fautin and Mariscal 1991,Yoffe et al. 2012). Therefore the cnidome of Octocorallia is unresolved and cannot currently provide much perspective (beyond the presence of the holotrich 1) on the ancestral nematocyst complement for Hexacorallia.

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Actiniarian cnidomes

Cnidome diversity is far higher in Actiniaria than in other hexacorallian taxa (Carlgren

1940, Schmidt 1974). The group can be divided into three major groups based on cnidome: 1) Endomyaria + Actinostolidae; 2) Hormathia + Nemathidae; and 3)

Aiptasiidae + Diadumenidae + Haliplanellidae + Metridiidae +Sagartiidae (Clade A in

Fig. 4.5). The first group has acanthonemes and p-rhabdoides A; the second also has acanthonemes, but has p-rhabdoides B1a rather than the p-rhabdoides A. The third group has the most diverse cnidome, having acanthonemes, colloponemes, and p-rhabdoides

B1a, B2d, and B2s (with both a and b variants). Because the variety of nematocysts is greatest in this third group, its members also have the highest diversity in the cnidome of the body tissues.

Despite an overall lack of diversity in nematocyst morphotypes in Endomyaria, its members evince some diversity in the nematocysts of the mesenterial filaments. In this group, and more specifically, in the family Actiniidae, two other morphologies are sometimes present, the aphylloneme and the diakaneme. Schmidt (1969) claimed that

Actina equina mediterranea (Schmidt 1971), now Actina schmidti (Monteiro et al. 1997), and Bunodactis rubripunctata have p-rhabdoides B1a in the mesenterial filaments, which is unlikely given that p-rhabdoid B forms in general are not found in Actiniidae. What he described as an ovoid or elliptical p-rhabdoid B is far more likely an aphylloneme or possibly a diakaneme, both of which have a capsule that is narrowest at its apical end, similar to (though generally more elongate than) the common p-rhabdoid B1a (Schmidt,

1969). I find no evidence of any form of p-rhabdoid B in members of Actiniidae, despite

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the claims of Schmidt (1969) that they occur, if only rarely. Most other variation in this

group is restricted to occurrence of p-rhabdoides A in the column; this feature cannot be

unambiguously mapped onto the tree so its value in defining subgroups within

Endomyaria is unclear.

Cnidomes in Actinostolidae (sensu Rodríguez et al. 2012; excludes Antholoba

achates and Paranthus niveus) studied here and in reports from the literature,

Hormosoma scotti (Carlgren, 1949, 1949, Dunn 1983) and Anthosactis janmayeni

(Carlgren 1945, 1949), vary little from those in Endomyaria, differing mostly in having

large “b-mastigophores” in the tentacles (capsules not studied here), and in occasionally

(e.g. Actinostola) having p-rhabdoides A in the tentacles (Carlgren 1945, Fautin 1984).

Schmidt (1969) allied this group with Sagartiidae and Metridiidae based on nematocysts,

however, this interpretation was based on the mistaken idea that Actinostolidae has p-

rhabdoides B1a and B2s (variant a), an idea that this study rejects. Schmidt (1969) also

likened the “large b-mastigophores” of actinostolid tentacles to the special b-rhabdoides

in the acontia of the Metridiidae. Given the large phylogenetic distance between the two

lineages and that these nematocyst morphs occur in different body structures, these

nematocyst forms are likely not homologous.

Members of the Hormathiidae + Nemanthidae group have a fairly invariant

cnidome, consisting mostly of acanthonemes in all body structures, with the addition of

p-rhabdoides B1a in the mesenterial filaments. Nemanthus nitidus also has the latter type in the tentacles. The lack of p- or amastigopores (=p-rhabdoid B forms) in the acontia has

166 been previously observed (Daly et al. 2008, Gusmão and Daly 2010) and Schmidt (1969) noted the lack of any other p-rhabdoid (A or B) forms in members of the group.

Clade A has all the nematocysts present in the Hormathiidae + Nemathidae clade, and four more: colloponeme and p-rhabdoides B2d and B2s (variants a and b). Having acontia with colloponemes and p-rhabdoid B2s (variant a or b) differentiates this group, and all members have p-rhabdoid B2d in one of the studied body structures. However, the distinction between this clade and Hormathiidae + Nemanthidae is most likely a loss of these morphotypes in Hormathiidae + Nemanthidae rather than a gain in Clade A, when the morphological sampling here is taken into account (see taxon sampling below for a discussion on this point).

Within Clade A, the complement of nematocysts in various body regions provides support for the close relationship of certain taxa. For example, Bartholomea annulata and Aiptasia sp. share several nematocyst characters, including loss of p-rhabdoides B1a in the mesenterial filaments, gain of p-rhabdoides B2s in the tentacles, and coupled gain of p-rhabdoides B2s and loss of p-rhabdoides B2d in the body column; these similarities bolster the hypothesis that they are sister taxa. Similarly, the two Diadumene species share a morphological change in p-rhabdoid B2s from variant a (where the Faltstück is shorter than the Hauptstück) to variant b (where the Faltstück is longer than the

Hauptstück). Schmidt (1969) claimed that the b variant occurs only in Diadumenidae, although according to Hand (1956) this morph is not present Diadumene cincta or D. lighti. Haliplanella lineata, which has been suggested to be a Diadumene (e.g., Manuel

1981, den Hartog 1987, Rodríguez et al. 2012), also does not have this form, but given

167 that not all members of the genus have variant b form, the utility of this feature for defining or diagnosing Diadumene is unclear.

A necessary point of clarification for the members of Clade A is their lack of p- rhabdoides A. Schmidt (1969) claimed that members of families Isophellidae,

Sagartiidae and Metridiidae have p-rhabdoides. However, den Hartog (1995) documented via light microscopy that the isophellid Telmatactis has only one morphotype of nematocyst that could be possibly be a p-rhabdoid A (from the actinopharynx); my assessment on the capsule shape is that this is more likely to be a large p-rhabdoid B1a than a p-rhabdoid A. Species of Sagartiidae and Metridiidae

(including species studied here: Sagartia elegans, Sagartiogeton undatus, Metridium senile) were interpreted to have p-rhabdoides A by den Hartog and Ates (2011). Although there is a morphotype of nematocysts in the mesenterial filaments in these animals that resembles the p-rhabdoid A in capsule shape and length of the basal tubule, (see Fig. 4.3

Q vs. Fig. 4.3A-E), SEM demonstrates that these are clearly a variant of p-rhabdoid B2s

(see Chapter 3 Fig. 2.10D). Therefore, no evidence supports the interpretation that any member of Metridioidea has p-rhabdoides A. Schmidt (1969) suggested that

Sagartiogeton laceratus is actually an aiptasiid based on the perception that both lack p- rhabdoid A, but DNA and cnidome characters link it tightly with S. undatus and with other sagartiids, rather than to the aiptasiids. Finally, Schmidt (1974) proposed that

Isophellidae, Sagartiidae, and Metridiidae all belong to a group identified as “late

Mesomyaria” in association with Actinostolidae and Halcampidae, based on the presence of both p-rhabdoides A and B (and small a B specifically in the mesenterial filaments) in

168 these taxa. Since neither fact is true (p-rhabdoides A do not occur in Sagartiidae or

Metridiidae, and p-rhabdoides B do not occur in Actinostolidae), this hypothesis cannot be supported (Schmidt 1974).

Taxon sampling

Although taxonomic coverage within Clade A is reasonable, sampling of other acontiarians is sparse, limiting the conclusions that can be made about the phylogenetic signal of nematocysts. Particularly problematic is the lack of data from deep sea and chemosynthetic sea anemones such as Jasonactis erythraios, Kadosactis antarctica, or

Alvinactis chessi. In the DNA tree, these deep-sea taxa are sister to the rest of

Metridioidea and thus important in determining the polarity of nematocyst characters.

The difference between Hormathiidae and Nemanthidae and the rest of the Metridioidea with respect to p-rhabdoides B2d and B2s cannot be polarized; it is unclear whether these capsules were gained in Clade A or lost in Hormathiidae + Nemanthidae. Kadosactis antarctica, Cyananthea hourdezi, and Jasonactis erythraios clearly have p-rhabdoid B2s

(presumably variant a, since variant b is likely restricted to Diadumene) based on high quality images from which nematocyst morphology can be evaluated (Rodríguez and

López-González 2005, Zelnio et al. 2009). All probably also have p-rhabdoides B2d; this is almost certain for K. antarctica (see Fig. 6d in Rodríguez and López-González (2005) and Zelnio et al. (2009). Furthermore, Phellia and Andvakia, which consitute part of the sister group to the Basitrina are both reported to have basitrichs and p-mastigophores

(likely to be p-rhabdoid B2s) in the acontia (Daly et al. 2008). Therefore, the proliferation

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of p-rhabdoid B forms most likely occurred at the base of Metridioidea and the

differences in cnidome are most likely the result of a loss of morphotypes in

Hormathiidae and Nemathidae rather than a gain in Clade A.

Although p-rhabdoid B morphotypes (B1a, B2d, B2s) can often be distinguished

in images of undischarged capsules under light microscopy, this is not the case with the

distinction between the acanthoneme and the colloponeme. Furthermore, the

colloponeme is most often restricted to the acontia in most taxa and many of these deep-

sea taxa do not bear acontia (Rodríguez and López-González 2005, Zelnio et al. 2009,

Rodríguez et al. 2012). Therefore the polarity of this character is unclear, it may have arisen at the base of the Metridioidea or Clade A. Taxa such as Amphianthus sp., J.

erythraios, and K. antarctica will be important in addressing this question.

Evolution of morphologies

Weill (1934) and Carlgren (1940) both suggested that nematocyst evolution followed a

path from simple (e.g., atrichs and holotrichs), to complex (e.g., the various forms of p-

rhabdoides B, called amastigophores in their terminology). However, only in the

transition of nematocysts from Octocorallia to Hexacorallia is this likely because more

complex morphologies (the hadroneme) are present early on in Hexacorallia. Even at

that transition, a lack of data on the cnidome of octocorals hinders this hypothesis (see

above). The three different holotrich forms are highly distinct in spine morphology

raising the question as to whether these forms are homologous. Holotrichs 2 and 3 could

be highly derived forms of the ancestral holotrich (which may or may not be holotrich 1).

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Conversely, these could be independently derived morphotypes. If holotrichs 2 and 3 are

not related to each other or to holotrich 1, then they likely evolved from either the

acanthoneme or p-rhabdoid A. As Schmidt (1974) noted, morphologies with thickened

basal tubules and two types/sizes of spines are present in the earliest-branching lineage of

Hexacorallia (= the hadroneme of Ceriantheria); he considered T-shaped spines and rhabdoides with long shafts (i.e. basal tubule of wide diameter) to be plesiomorphic.

With respect to the possible connections between morphotypes, the spine shape of the large holotrichs of Corallimopharia and Scleractinia (Skaer and Picken 1965, Schmidt

1974) more closely resemble the spines of the p-rhabdoid A than they do the spines of the other two holotrich forms seen here. In both the large holotrichs and p-rhabdoides A, the spines are T-shaped, with the head of the spine bearing narrow projections to each side

(Fig. 4.4O, P). This similarity was also noted by Cutress (1955), who suggested that some “holotrichs” of Scleractinia or Corallimorpharia might actually be macrobasic p- mastigophores (his terminology for what is here called a p-rhabdoid A). Although I see no evidence of a smaller diameter distal tubule in any coral or corallimorph holotrich as would be the case if they were p-rhabdoides, his hypothesis does recognize that there are some similarities between this holotrich form and p-rhabdoides A. Furthermore, the latter form is common in corals and corallimorphs and could provide the source for this derived form.

Based on the tree and the mapping of characters, the various forms of p-rhabdoid

B are likely to have been derived from a form similar to either an acanthoneme or a p- rhabdoid A. As their similar names suggest, Schmidt (1969, 1974) inferred that the

171

dramatic thickening of the basal tubule compared to the distal tubule was homologous for

p-rhabdoides A and B and thus evidence of their close relationship. My investigations

provide some support for this connection: the p-rhabdoid A has large folds in the tubule

surface where the spines attach (see Chapter 3); if these folds extended to encircle the

basal tubule they could create the annulations seen on the surface of all p-rhabdoides B.

However, the shape of the base of the spine of p-rhabdoid B morphotypes is more similar

to the shape of spines on an acanthoneme than to the T-shaped spines of a p-rhabdoid A.

Furthermore, both acanthonemes and p-rhabdoid B forms have the derived apical

structure (apical flaps) rather than the ancestral form (the apical cap) that is found in p-

rhabdoides A. The spine and apical structure features provide some support for the

hypothesis that the p-rhabdoid B forms are derived from acanthonemes rather than p-

rhabdoides A. Following Schmidt (1969, 1974) and treating the dilation of the basal

tubule as a distinct feature (the shaft) that is homologous in those nematocysts that bear it may lead to more value placed on this character, and therefore bolster the interpretation that B forms of p-rhabdoides arose from A forms. However, the exact nature and definition of this “shaft” feature has been problematic and inconsistent (see Chapter 3).

Determining which of the p-rhabdoid B forms evolved first is problematic with

out a clear placement on the tree of the origin of each form. The p-rhabdoid Bs all arise

within Metridioidea, but determining where exactly in the tree they originate is

confounded by taxon sampling issues (see above). The p-rhabdoid B1a evolved early in

Metridioidea and is shared by all but the aiptasiids, which are inferred to have lost this

morphotype. The p-rhabdoid B1a has more in common with p-rhabdoides A than do the

172 other forms: both have only one region of the basal tubule and one type of spine (though not the same spine), and the distal tubule always remains attached in both (see Chapter

3). However, without knowing the full distribution of p-rhabdoid B2d and B2s, the B1a form cannot be assumed to be the first of the p-rhabdoid B forms in Metridioidea.

Greater taxon sampling for morphology should help clarify the node of origin for p- rhadoids B2d and B2s. The two variants of p-rhabdoides B2s are less ambiguous in their relationship: variant b is clearly a derived form of p-rhabdoid B2s variant a. This hypothesis is supported by the phylogenetic distribution where the latter form replaces the former within the Diadumenidae.

As to the evolution of other morphs such as the colloponeme, acanthoneme, and hadroneme, little can be hypothesized given the current data. The full distribution of the colloponeme is not currently understood in Metridioidea (particularly among the deep-sea and chemosynthetic taxa), which confounds any attempt to understand its origins. As for the acanthoneme and hadroneme, both morphologies arise early in Hexacorallia, and are pervasive once they arise. There are no obvious morphological features shared by the two forms other than both having a thick basal tubule in the undischarged capsule, although the structure of this basal tubule is obviously different between the two forms

(Fig. 4.1A, F). Acanthonemes may indeed be related to atrichs or holotrich 1 as hypothesized by Carlgren (1940), with the cerianthid lineage losing this form (or losing the holotrich from which acanthonemes evolved).

Finally, the tree has implications for understanding the evolution of the two unique nematocysts of Endomyaria. The similarities in capsule shape, basal tubule, and

173

spines of the aphylloneme and diakaneme provide morphological evidence that the two

forms are related. The fact that they occur in the same body structure (mesenterial

filaments) in taxa that are all part of the same monophyletic group (Endomyaria) provides

more support that these two forms are evolutionary related. The most likely tree from

this data provides information on the polarity of any evolutionary relationship, suggesting

that the aphylloneme is derived from the diakaneme, and represents an instance of simplification, as it has one type of spine rather than two and an isodiametric rather than anisometric tubule.

174

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Table 4.1. Summary of taxa and collection locality and body part sampled for SEM for distribution study. A: acontia, Ac: acrorhagi, C: body column, MF: mesenterial filament, T: tentacle.

180

Table 4.1 Higher Taxon Species Location collected Tissues sampled Actiniaria Actinia equina SW of Mahee Island, C, MF, T Strangford Lough, N. Ireland Acontiaria sp. Jakyakdo Island, S. Korea A, C, MF, T Actinostolid sp. San Juan Island, WA, USA C, MF, T Adamsia palliata Ringhaddy Sound, Strangford A, C, MF, Lough, N. Ireland T Aiptasia sp. Aquarium Adventure, A, C, MF, Columbus, OH, USA T Anemonia Ballymacormick Pt, C, MF, T sulcata Groomsport, N. Ireland Anthopleura San Juan Island, WA, USA Ac, C, elegantissima MF, T Bartholomea University of Virgin Islands, A, C, MF, annulata St. Thomas, US Virgin Islands T Boloceroides Kyoto University Seto Marine C, T mcmurrichi Lab, Shirahama, Japan Bunodosoma Galveston, TX, USA Ac, C, cavernata MF, T Calliactis Jeju Island, S. Korea A, C, MF, polypus T Calliactis Galveston, TX, USA A tricolor Cereus SW of Mahee Island, A, C, MF, pedunculatus Strangford Lough, N. Ireland T Condylactis Aquarium Adventure, A, C, MF, gigantea Columbus, OH, USA T Diadumene Marine Biological Lab, Woods A, C, MF, leucolena Hole, MA, USA T Diadumene sp. Jakyakdo Island, S. Korea A, C, MF, T Entamaea Jeju Island, S. Korea C, MF, T quadricolor Epiactus San Juan Island, WA, USA C, MF, T prolifera Haliplanella Jakyakdo Island, S. Korea A, C, MF, lineata T Metridium senile Marine Biological Lab, Woods A, C, MF, Hole, MA, USA T Continued

181

Table 4.1 continued Nemanthus Jeju Island, S. Korea C, MF, T nidtus Paracondylactis Jakyakdo Island, S. Korea C, MF, T hertwigi Sagartia elegans SW of Mahee Island, A, C, MF, Strangford Lough, N. Ireland T Sagartigeton Ringhaddy Sound, Strangford A, C, MF, lacerates Lough, N. Ireland T Sagartiogeton Ringhaddy Sound, Strangford A, C, MF, undatus Lough, N. Ireland T Urticina felina SW of Mahee Island, C, MF, T Strangford Lough, N. Ireland Antipatharia Antipathes griggi Auau Channel, HI, USA T, MF Antipathes Auau Channel, HI, USA T, MF grandis Ceriantharia Ceriantheopsis Gulf Specimen Marine Lab, C, MF, T americanus Panacea FL, USA Cerianthus Kyoto University Seto Marine T filiformis Lab, Shirahama, Japan Cerianthidae sp. Reef Hot Spot, Inglewood, T CA, USA Corallimorpharia Discosoma sp. Aquarium Adventures, MF Columbus, OH, USA Rhodactis sp. Reef Hot Spot, Inglewood, MF CA, USA Ricordia sp. Aquarium Adventures, MF Columbus, OH, USA Scleractinia Platygyra Gulf Specimen Marine Lab, T asteriformis Panacea, FL, USA Goniporia sp Reef Hot Spot, Inglewood, T CA, USA Echinophyllia Kyoto University Seto Marine MF aspera Lab, Shirahama, Japan Caulastrea Kyoto University Seto Marine MF tumida Lab, Shirahama, Japan Acanthastrea sp. Kyoto University Seto Marine MF Lab, Shirahama, Japan Zoanthidea Protopalythoa Neil Blackstone culture C, MF, T mutuki Zoanthus University of Virgin Islands, C, MF pulchellus St. Thomas, US Virgin Islands Octocorallia Renilla mulleri Gulf Specimen Marine Lab, T Panacea FL, USA 182

Table 4.2. Summary of taxa and genes sampled to create matrix for phylogenetic analysis. Genbank numbers are given for all published genetic data. NEW designates a new sequence.

183

Table 4.2 Higher Taxon Family Species 16S 18S 28S Athenaria Edwardsiidae Edwardsia elegans EU190770 EU190857 EU190815 Edwardsia japonica GU473288 GU473304 GU473321 Edwardsia timida GU473299 GU473315 JF832999 Edwardsianthus gilbertensis EU190772 EU190859 EU190817 Nematostella vectensis AY169370 AF254382 EU190838 Haloclavidae Haloclava producta EU190779 AF254370 EU190823 Boloceroides Boloceroidaria Boloceroididae mcmurrichi EU190852 EU190810 Endomyaria Actiniidae Actinia fragacea EU190756 EU190845 EU190802 184 Anemonia viridis EU190760 EU190849 EU190806 Anthopleura elegantissima EU190755 EU190844 EU190801 Bunodactis verrucosa EU190766 EU190854 EU190812 Bunodosoma cavernata NEW NEW Bunodosoma grandis EU190765 EU190853 EU190811 Condylactis gigantea NEW NEW NEW Entacmaea quadricolor NEW Epiactis lisbethae EU190771 EU190858 EU190816 Epiactis prolifera NEW NEW NEW Isosicyonis striata EU190781 EU190864 FJ489463 Macrodactyla doreenensis EU190785 EU190867 EU190828 Paracondylactis hertwigi NEW NEW NEW Urticina coriacea EU190797 EU190877 EU190840 Continued

184

Table 4.2 continued Urticina felia U91751 NEW NEW Actinodendridae Actinostephanus haeckeli EU190762 EU190808 Aliciidae Triactis producta EU190876 EU190839 Phymanthidae Phymanthus loligo EU190791 EU190871 Preactiidae Dactylanthus antarcticus AY345877 AF052896 AY345873 Mesomyaria Actinostolidae Actinostola crassicornis EU190753 EU190843 EU272904 Actinostola chilensis GU473285 GU473302 Antholoba achates GU473284 GU473301 GU473318 Anthosactis janmayeni GU473292 GU473308 GU473324 Hormosoma scotti EU190778 EU190863 EU190822 Paranthus niveus GU473295 GU473311 GU473327

185 Stomphia didemon EU190795 EU190875 EU190837

Stomphia selaginella GU473298 GU473314 GU473331 Metridioidea Actinoscyphiidae Actinoscyphia plebeia EU190754 FJ489437 EU190800 Aiptasiidae Aiptasia mutabilis FJ489418 FJ489438 FJ489469 Aiptasia pulchella EU190757 EU190846 EU190803 Aiptasia sp. NEW NEW NEW Bartholomea annulata EU190763 EU190851 EU190809 Neoaiptasia morbilla EU190788 EU190869 EU190831 Amphianthus Amphianthus sp. FJ489432 FJ489450 FJ489467 Andvakiidae Andvakia boninensis EU190759 EU190848 EU190805 Andvakia discipulorum GU473287 GU473316 GU473320 Telmatactis sp. JF832979 JF833001 Antipodactinidae Antipodactis awii GU473286 GU473303 GU473319 Continued

185

Table 4.2 continued

Bathyphellidae Bathyphellia australis FJ489422 EF589063 EF589086 Diadumenidae Diadumene cincta EU190769 EU190856 EU190814 Diadumene leucolena JF832977 JF832986 JF832995 Diadumene sp. JF832976 JF832980 JF832990 Diadumene sp. Korea NEW NEW Halcampidae Halcampa duodecimcirrata EU190776 AF254375 Cactosoma sp. nov. GU473297 GU473313 GU473329 Halcampoides purpureus EU190780 AF254380 EU190824 Haliplanellidae Haliplanella lineata (USA) EU190774 EU190860 EU190819 Haliplanella lineata (Japan) JF832973 JF832987 JF832998 Hormathiidae Actinauge richardi EU190761 EU190850 EU190807 186 Adamsia palliata FJ489419 FJ489436 FJ489452 Allantactis parasitica FJ489420 FJ489439 FJ489454 Calliactis japonica FJ489423 FJ489441 FJ489456 Calliactis parasitica EU190752 EU190842 EU190799 Calliactis polypus (HI) FJ489427 FJ489445 FJ489459 Calliactis tricolor FJ489425 FJ489443 FJ489458 Chondrophellia orangina FJ489426 FJ489444 Hormathia armata EU190775 EU190861 FJ489460 Hormathia lacunifera FJ489428 FJ489446 FJ489461 Hormathia pectinata FJ489430 FJ489448 FJ489465 Paracalliactis japonica FJ489429 FJ489447 FJ489464 Paraphelliactis sp. FJ489431 FJ489449 FJ489466 Continued

186

Table 4.2 continued

Isanthidae Isanthus capensis GU473291 GU473307 GU473323 Paraisanthus fabiani GU473283 GU473300 GU473317 Kadosactinidae Alvinactis chessi GU473296 GU473312 GU473328 Cyananthea hourdezi GU473293 GU473309 GU473325 Kadosactis antarctica EU190782 EU190865 EU190825 Jasonactis erythraios GU473289 GU473305 GU473330 Metridiidae Metridium senile EU190786 AF052889 EU190829 Metridium senile (WA) JF832972 JF832982 JF833000 Metridium s. fibratum (Japan) JF832974 JF832988 JF832996

187 Metridium s. lobatum JF832971 JF832981 JF832991 (Argentina)

Nemathidae Nemanthus nitidus EU190787 EU190868 EU190830 Ostiactinidae Ostiactis pearseae EU190798 EU190878 EU190841 Phelliidae Phellia gausapata EU190790 EU190870 EU190833 Phellia exlex JF832978 JF832984 JF832993 Sagartiidae Anthothoe chilensis FJ489416 FJ489434 FJ489453 Actinothoe sphyrodeta FJ489421 FJ489440 FJ489455 Cereus pedunculatus EU190767 EU190855 EU190813 Cereus herpetodes JF832969 JF832983 JF832992 Sagartia elegans JF832970 JF832989 JF832994 Sagartia troglodytes EU190792 EU190872 EU190834 Sagartia ornate JF8329975 JF832985 JF832997 Sagartiogeton lacerates EU190794 EU190874 EU190836 Continued

187

Table 4.2 continued Sagartiogeton undatus FJ489417 FJ489435 FJ489462 Verrillactis paguri FJ489433 FJ489440 FJ489468 Unknown Acontiaria sp. NEW NEW Zoanthidea Savalia savaglia DQ825686 HM044299 HM044298 188

188

Table 4.3. Distribution of nematocyst morphologies by actiniarian family. Family Aca Aph Coll pA pB1a pB2d pB2s (a) pB2s (b) Dia Mac Holo Actiniidae X X X X X Actinostolidae X X Aiptasiidae X X X X Boloceroididae X X X Diadumenidae X X X X X X X (X) Haliplanellidae X X X X X X (X) Hormathiidae X X Metridiidae X X X X X (X) Nemathidae X X Sagartiidae X X X X X (X) X indicates that morphology observed in representatives of that family, (X) indicates those families with catch tentacles and therefore with holotrichs that were not observed in this study. Aca: acanthoneme, Aph: aphylloneme, Coll: colloponeme, pB1a: p- 189 rhabdoid B1a, pB2d: p-rhabdoid, pB2s (a): p-rhabdoid B2s variant a, pB2s (b): p-rhabdoid B2s variant b, Dia: diakaneme, Mac: macrobasic mastigophore, Holo: holotrich

189

Table 4.4. Summary of nematocyst distribution by species and tissue type. Unless noted, all p-rhabdoid B2s are of the a variety. Higher Taxon Species Acrorhagi Acontia Column Mesenterial Tentacle filament Boloceroidaria: Boloceroides NA NA acanthoneme, Not sampled acanthoneme, Boloceroididae mcmurrichi p-rhabdoid p-rhabdoid B2d, B2d, B2s B2s Endomyaria: Actinia equina holotrich NA acanthoneme acanthoneme, acanthoneme Actiniidae p-rhabdoid A Anemonia Not sampled NA acanthoneme, acanthoneme, acanthoneme sulcata p-rhabdoid A aphylloneme, p-rhaboid A Anthopleura acanthonem NA acanthoneme acanthoneme, acanthoneme

190 elegantissima e, holotrich p-rhabdoid A Bunodosoma acanthonem NA acanthoneme acanthoneme, acanthoneme

cavernata e, holotrich aphylloneme, p-rhabdoid A Condylactis NA NA acanthoneme, acanthoneme, acanthoneme gigantea p-rhabdoid A p-rhabdoid A Entacmaea NA NA acanthonemes, acanthoneme, acanthonemes quadricolor p-rhabdoid A p-rhabdoid A Epiactis NA NA acanthoneme acanthoneme, acanthoneme prolifera p-rhabdoid A Paracondylactis NA NA acanthoneme acanthoneme, acanthoneme hertwigi p-rhabdoid-A, aphylloneme Continued

190 Table 4.4 continued Urticina felia NA NA acanthoneme, acanthoneme, acanthoneme p-rhabdoid A p-rhabdoid A, diakaneme Mesomyaria: Actinostolid sp. NA NA acanthoneme acanthoneme, acanthonemes Actinostolidae p-rhaboid A p-rhabdoid A Metridioidea: Aiptasia sp. NA colloponeme acanthoneme, acanthoneme, acanthoneme, p- Aiptasiidae p-rhabdoid B2s p-rhabdoid p-rhabdoid B2d, rhabdoid B2d, B2s B2s B2s Bartholomea NA colloponeme acanthoneme, acanthoneme, p- acanthoneme, annulata p-rhabdoid B2s p-rhabdoid rhabdoid B2d, colloponeme, B2s B2s p-rhabdoid B2d, 191 B2s

Metridioidea: Diadumene NA colloponeme acanthoneme, acanthoneme, acanthonemes, p- Diadumenidae leucolena p-rhabdoid B2s p-rhabdoid B1a, p-rhabdoid rhabdoid B2d (b) B2d B2d, B2s (b)

Diadumene sp. NA colloponeme acanthoneme, acanthoneme, acanthoneme, p- Korea p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, rhabdoid B2d, (b) B2d B2d, B2s (b) macrobasic amastigophore, Metridioidea: Haliplanella NA colloponeme acanthoneme, acanthoneme, acanthoneme, Haliplanellidae lineata p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, p-rhabdoid B2d B2d B2d, B2s Metridioidea: Adamsia NA acanthoneme acanthoneme acanthoneme, acanthoneme Hormathiidae palliata p-rhabdoid B1a Continued

191

Table 4.4 continued Calliactis NA acanthoneme acanthoneme acanthoneme, acanthoneme polypus B1a Calliactis NA acanthoneme, Not sampled Not sampled Not sampled tricolor Metridioidea: Metridium NA acanthoneme, acanthoneme, acanthoneme, acanthoneme, Metridiidae senile colloponeme p-rhabdoid p-rhabdoid B1a, colloponeme, p-rhabdoid B2s B2d B2s p-rhabdoid B2d Metridioidea: Nemanthus NA NA acanthoneme acanthoneme, acanthoneme, Nemathidae nitidus p-rhabdoid B1a p-rhabdoid B1a Metridioidea: Cereus NA colloponeme acanthoneme, acanthoneme, acanthoneme, 192 Sagartiidae pedunculatus p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, p-rhabdoid B2d B2d B2s Sagartia NA colloponeme acanthoneme, acanthoneme, acanthoneme, p- elegans p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, rhabdoid B2d B2d, B2s B2d, B2s Sagartigeton NA colloponeme acanthoneme, acanthoneme, acanthoneme, lacerates p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, colloponeme, p- B2d B2s rhabdoid B2d, B2s Sagartigeton NA colloponeme acanthoneme, acanthoneme, acanthoneme, undatus p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, colloponeme, B2d B2s p-rhabdoid B2d, B2s Continued

192

Table 4.4 continued Metridioidea: Acontaria sp. NA colloponeme acanthoneme, acanthoneme, acanthoneme, Unknown p-rhabdoid B2s p-rhabdoid p-rhabdoid B1a, p-rhabdoid B2d B2s B2d, B2s Antipatharia Antipathes NA NA Not sampled Acanthoneme, acanthoneme grandis p-rhabdoid A Ceriantharia Ceriantheopsis NA NA hadroneme hadroneme hadroneme americanus C filiformis NA NA hadroneme hadroneme hadroneme Corallimorpharia Discosoma sp. NA NA Not samples holotrich, 3 NA p-rhabdoid A 193 Rhodactis sp. NA NA Not sampled holotrich, 3 p-rhabdoid A Ricordea sp. NA NA Not sampled holotrich, 3 p-rhabdoid A Octocorallia Renilla mulleri NA NA Not sampled Not sampled holotrich Scleractinia Acanthastrea NA NA holotrich 3 Not sampled sp. p-rhabdoid A Caulastrea NA NA Not sampled acanthoneme Not sampled tumida holotrich 3, p-rhabdoid A Echinophyllia NA NA Not sampled holotrich 3, Not sampled aspera p-rhabdoid A Goniporia sp. NA NA Not sampled Not sampled holotrich, p-rhabdoid A

193 Table 4.4 continued Platygyra NA NA Not sampled Not sampled holotrich, astreiformis p-rhabdoid A Zoanthidea Protopalythoa NA NA Not sampled acanthoneme, acanthoneme, mutuki holotrich 2, holotrich p-rhabdoid A Zoanthus NA NA Not sampled acanthoneme, acanthoneme, pulchellus holotrich 2, holotrich p-rhabdoid A

194

Figure 4.1. Nematocyst morphotypes documented in distribution study. Light and scanning electron microscope images showing the characteristic features of 12 identified morphotypes. A. Light microscope image of an acanthoneme from a tentacle of Epiactis prolifera. Scale Bar = 20 µm. B. Scanning Electron microscope image of an acathoneme from a tentacle of Actinostolid sp. Scale Bar = 10 µm. C. Light microscope image of an aphylloneme from a mesenterial filament of Anemonia sulcata. Scale Bar = 20 µm. D. Light microscope image of a colloponeme from aconitum of Bartholomea annulata. Scale Bar = 20 µm. E. Scanning electron microscope image of a colloponeme from acontium of Aiptasia sp. Scale Bar = 5 µm. F. Light microscope image of a hadroneme from a mesenterial filament of Cerianthopsis americanus. Scale Bar = 20 µm. G. Scanning electron microscope image of a hadroneme from mesenterial filament of Cerianthosis americanus. Scale Bar = 10 µm. H. Light microscope image of p-rhabdoid A from mesenterial filament of Entacmaea quadricolor. Scale Bar = 20 µm. I. SEM image of a p-rhabdoid A from a mesenterial filament of Bunodosoma cavernata. Scale Bar = 2 µm. J. Light microscope image p-rhabdoid B1a from mesenterial filament of Acontiaria sp. Scale Bar = 20 µm. K. Scanning electron microscope image of p-rhabdoid B1a from mesenterial filement of Diadumene lecuolena. Scale Bar = 5 µm. L. Light microscope image of p-rhabdoid B2d from column of Sagartia elegans. Scale Bar = 20 µm. M. Scanning electron microscope image of p-rhabdoid B2d from mesenterial filament of Sagartia elegans. Scale Bar = 10 µm. N. Light microscope image of p- rhabdoid B2s (variant a) from aconitum of Sagartia elegans. Scale Bar = 20 µm. O. Scanning electron image of p-rhabdoid B2s (variant a) from mesenterial filament of Bartholomea annulata. Scale Bar = 25 µm. P. Scanning electron image of p-rhabdoid B2s (variant b) from mesenterial filament of Diadumene sp. Korea. Scale Bar = 50 µm. Q. Light microscope image of diakaneme from mesenterial filament of Urticina felina. Scale Bar = 20 µm. R. Light microscope image of macrobasic amastigophore from tentacle of Diadumene sp. Korea. Scale Bar = 20 µm. S. Light microscope image of holotrichous isorhiza (holotrich) from mesenterial filament Protopalythoa mutuki. Scale Bar = 20 µm.

195

Figure 4.1 196

Figure 4.2. Variation among the b-rhabdoid types: A-D Acanthonemes, E-G aphyllonemes, H-J colloponemes, K-M hadronemes. Scale Bar = 20 µm. A. Light microscope image of acanthoneme from tentacle of Entacmaea quadricolor. B. Light microscope image of acanthoneme from mesenterial filament of Aiptasia sp. C. Light microscope image of acanthoneme from mesenterial filament of Urticina felina. D. Light microscope image of acanthoneme from tentacle of Antipathes grandis. E. Light microscope image of aphylloneme from meseterial filament of Bunodosoma cavernata. F. Light microscope image of aphylloneme from mesenterial filament of Paracondylactis hertwigi. G. Light microscope image of aphylloneme from mesenterial filament of Paracondylactis hertwigi. H. Light microscope image of colloponeme from acontia of Aiptasia sp. I. Light microscope image of colloponeme from acontia of Aiptasia sp. µm. J. Light microscope image of colloponeme from acontia of Sagartiogeton laceratus. Scale Bar = 20 µm. K. Light microscope image of hadroneme from tentacle of Cerianthopsis americanus. L. Light microscope image of colloponeme from tentacle of Cerianthidae sp. M. Light microscope image of colloponeme from tentacle of Cerianthidae sp.

197

Figure 4.2

198

Figure 4.3. Variation among the p-, amastigophore types: A-E p-rhabdoid A, F-J p- rhabdoid B1a, K-N p-rhabdoid B2d, O-R p-rhabdoid 2Bs (variant a). Scale Bar = 20 µm. A. Light microscope image of p-rhabdoid A from mesenterial filament of Actinostola sp. B. Light microscope image of p-rhabdoid A from mesenterial filament of Urticina felina. C. Light microscope image of p-rhabdoid A from mesenterial filament of Bunodosoma cavernata. D. Light microscope image of p-rhabdoid A from mesenterial filament of Epiactis prolifera. E. Light microscope image of p-rhabdoid A from mesenterial filament of Antipathes grandis. F. Light microscope image of p-rhabdoid B1a from mesenterial filament of Sagartiogeton undatus. G. Light microscope image of p- rhabdoid B1a from mesenterial filament of Diadumene leucolena. H. Light microscope image of p-rhabdoid B1a from meseneterial filament of Aiptasia sp. I. Light microscope image of p-rhabdoid B1a from mesenterial filament of Nemanthus nidtus. J. Light microscope image of p-rhabdoid B1a from mesenterial filament of Nemanthus niditus. K. Light microscope image of p-rhabdoid B2d from body column of Haliplanella lineata. L. Light microscope image of p-rhabdoid B2d from tentacle of Sagartia elegans. M. Light microscope image of p-rhabdoid B2d from body column of Diadumene sp. Korea. N. Light microscope image of p-rhabdoid B2d from body column of Sagartia elegans. O. Light microscope image of p-rhabdoid B2s from tentacle of Bartholomea annulata. P. Light microscope image of p-rhabdoid B2s from acontia Haliplanella lineata. Q. Light microscope image of p-rhabdoid B2s from mesenterial filament of Sagartiogeton undatus. R. Light microscope image of p-rhabdoid B2s from mesenterial filament of Sagartia elegans.

199

Figure 4.3. 200

Figure 4.4. Holotrichous isorhiza (holotrich) variation A. Light microscope image of holotrich 1 from acrorhagi of Anthopleura elegantissima. Scale Bar = 20 µm. B. Light microscope image of holotrich 1 from acrorhagi of Actinia equina. Scale Bar = 20 µm. C. Scanning electron microscope image of holotrich 1 from acrorhagi of Bunodosoma cavernata. Scale Bar = 2 µm. D. Scanning electron microscope image of holotrich 1 from tentacle of Renilla mulleri. Scale Bar = 5 µm. E. Scanning electron microscope image of holotrich 1 from tentacle of Renilla mulleri. Scale Bar = 3 µm. F. Light microscope image of holotrich 2 from tentacle of Protopalythoa mutuki. Scale Bar = 20 µm. G. Scanning electron microscope image of holotrich 2 from tentacle of Zoanthus pulchellus. Scale Bar = 10 µm. H. Scanning electron microscope image of holotrich 2 from tentacle of Protopalythoa mutuki. Scale Bar = 5 µm. I. Scanning electron microscope image of holotrich 2 from tentacle of Protopalythoa mutuki. Scale Bar = 3 µm. J. Scanning electron microscope image of holotrich 2 from tentacle of Zoanthus pulchellus. Scale Bar = 3 µm. K. Light microscope image of holotrich 3 from tentacle of Gonipora sp. Scale Bar = 20 µm. L. Light microscope image of holotrich 3 from mesenterial filament of Ricordia sp. Scale Bar = 20 µm. M. Scanning electron microscope image of holotrich 3 from tentacle of Platygyra asterformis. Scale Bar = 5 µm. N. Scanning electron microscope image of holotrich 3 from mesenterial filament of Rhodactis sp. Scale Bar = 20 µm. O. Scanning electron microscope image of holotrich 3 from tent of Platygyra asterformis. Scale Bar = 5 µm. P. Scanning electron microscope image of holotrich 3 from mesenterial filament Rhodactis sp. Scale Bar = 5 µm.

201

Figure 4.4

202

Figure 4.5. Highest likelihood tree. Branch lengths are proportional; see scale below. ML bootstrap values from 1000 replicates shown for nodes having values >50%. Dots represent nodes bootstrap values >95%.

203

Figure 4.6. Cnidome distribution across Hexacorallia. Cnidome features are mapped onto tree of Brugler and France (2007); only unambiguous optimizations indicated.

204

Figure 4.7. Phylogenetic perspective on the cnidome of mesenterial filaments of actiniarians. Tree is from Fig. 4.5 with Edwardsiidae collapsed and branch lengths rendered equal. Only unambiguous optimizations indicated.

205 Figure 4.8. Phylogenetic perspective on the cnidome of tentacles of actiniarians. Tree is from Fig. 4.5 with Edwardsiidae collapsed and branch lengths rendered equal . Only unambiguous optimizations indicated.

206 Figure 4.9. Phylogenetic perspective on the cnidome of the body column of actiniarians. Tree is from Fig. 4.5 with Edwardsiidae collapsed and branch lengths rendered equal. Only unambiguous optimizations indicated.

207 Figure 4.10 Phylogenetic perspective on the cnidome of acontia and acrorhagi of actiniarians. Tree is from Fig. 4.5 with Edwardsiidae collapsed and branch lengths rendered equal. Only unambiguous optimizations indicated. Note that these structures do not occur in the same taxa, being restricted to some representatives of Endomyaria (acrorhagi) and Metridioidea (acontia).

208

Chapter 5: Multivariate analysis of nematocyst variation visible in light and electron

microscopy

Introduction

Because all cnidarians bear nematocysts, these capsules are one of the few morphological features that can be assessed and analyzed across the phylum. Like DNA, these characters provide an opportunity to code characters from organisms highly diverse in other features. However, as with DNA where the alignment is key to determining homology of the base pairs, the assessment of nematocyst morphological type is a fundamental step where homology is assessed.

Individual features of nematocysts are not coded as phylogenetic characters; rather, all aspects of morphology are evaluated qualitatively and simultaneously so that morphological variation is binned into a few small categories (Weill 1934; Carlgren

1940, 1949; Hand 1956; Schmidt 1969, 1974; den Hartog 1980, Shostak and Kolluri

1995). Individual nematocysts are classified into these categories using an overall similarity criterion, to determine the “type” of nematocyst; this is a composite character sensu Wilkinson (1995). This approach creates issues in the use of nematocysts as phylogenetic characters. Because characters themselves are not being evaluated and coded, any small change in the morphology may cause confusion as to its placement.

The exact circumscription of the nematocyst morphology categories can be difficult to define for all cases leaving some morphologies ambiguous and difficult to interpret, and 209 therefore unusable for any phylogenetic analysis. Furthermore, the relationship between

morphologies is unclear. Transformation series for individual characters can easily be

hypothesized, but a series by which one distinct morphology transforms several features

to transition to another morphology is far more difficult.

Because nematocyst characters are treated as composite characters and almost

always coded as presence/absence of morphological types (Weill 1934; Carlgren 1940,

1949; Hand 1956; Schmidt 1969, 1974; den Hartog 1980, Shostak and Kolluri 1995) the

assignment to type is the key step where homology is hypothesized. Therefore,

confusion or disagreement as to how to group the morphologies (as described in Chapter

3) will directly affect the homology statements made about these morphologies. These

groupings have historically been based on qualitative assessments of the morphologies

(Weill 1934, Carlgren 1940, Cutress 1955, Schmidt 1969, Mariscal 1974, Östman 2000),

but this need not be the case. The nematocyst consists of several morphological features including the shape of the capsule, features of the tubule surface and diameter, and shape, size and distribution of spines. All of these features can be measured and a biometric approach utilized to assess morphological groupings instead of a qualitative one.

The use of measurements of nematocyst features has a long history in cnidarian

biology (e.g. Carlgren 1900, Calder 1972, 1977, Calder and Peters 1975, Avian et al.

1991, Chintiroglou 1996, Avian et al. 1997, Ardelean and Fautin 2004, see Fautin 1988

for a summary of the use of these measurements specifically in Actiniaria). However,

previous studies have focused solely on capsule length and width (or the ratio of the two)

of undischarged nematocysts studied with light microscopy (Calder 1972, 1977, Calder

and Peters 1975, Chintiroglou 1996, Williams 1996, 1998, 2000, Acuña et al. 2003, 210 Ardelean and Fautin 2004, Ryland and Lancaster 2004, Acuña et al. 2004). Furthermore, authors have been interested in using nematocyst biometric data primarily in differentiation species or populations of cnidarians (Calder 1972, 1977, Williams 1996,

1998, Acuña et al. 2003, 2004, Ryland and Lancaster 2004), or in determining if a relationship exists between the size of the organism and the cnidae it bears (Chintiroglou

1996, Acuña et al. 2007). Whether the classification of the nematocyst morphologies that is based on qualitative data is supported by quantitative (continuous, measured) data has not been determined. Presumably, finding such support would strengthen confidence that these morphological groupings represent natural and real objects that can be identified objectively rather than unnatural groupings forced by our need to organize the observed variation.

A multivariate biometric approach can be used to explore relationships among morphological features (e.g., Neff & Marcus 1980) and assess how morphologies group in morphospace. Rather than limit the assessment of nematocyst shape to only length and width of undischarged capsules, electron microscopy enables precise measurements of details not visible under the light microscope and thus allows for a measurement of far more nematocyst features (such as tubule width and spine size). With the addition of new features, a more comprehensive biometric analysis can be performed to better assess the grouping of morphologies. Furthermore, homology statements are initially made by assessing shared similarity (Patterson 1988, de Pinna 1991) and a multivariate approach will provide data by which similarity of nematocysts can be assessed.

Reliance on this typological framework for nematocyst diversity obscures the diversity of form in these remarkable structures, complicates their utility as taxonomic 211 characters, and makes it impossible to interpret their diversity in a phylogenetic context.

To rectify this situation I document the microscopic diversity using a biometric approach to better characterize variation in nematocyst shape and assess how qualitatively well- defined groups (i.e. the nine morphologies discussed in Chapter 3) match up to quantitatively defined groups. A multivariate approach identifies clusters of nematocysts with similar morphologies providing information on what features distinguish different morphological groups of nematocysts. This will inform the evaluation of current nematocyst classification systems and nematocyst features to be homologized for phylogenetic analyses.

Materials and methods

Materials

I examined nematocysts from twenty-two actiniarians, one antipatharian, two cerianthids, and one scleractinian (Table 5.1). These specimens were collected from various localities

(Table 5.1) or obtained from commercial sources such as Aquarium Adventure

(Columbus, OH), Gulf Specimen Marine Lab (Panacea, FL), Marine Biological Lab

(Woods Hole, MA) and Reef Hot Spot (Inglewood, CA). The antipatharian samples were collected, fixed, and provided by Anthony Montgomery (see Opresko, 2009 for collecting details). Species identifications were determined by the collector or verified in the case of commercial sources wherever possible. In one case, the actiniarian could not be identified beyond belonging to the group Acontiaria (Acontiaria sp. in tables). In another, a cerianthid could only be identified to family (Cerianthidae sp. in tables).

212 Methods

Sample preparation

After collection all actiniarian samples were dissected to isolate key body regions: mesenterial filaments, body column, tentacles, and acontia if present (see Table 5.2 for body regions collected).

CO2. Samples from B. cavernata and were sputter-coated with gold-palladium in a

Hummer sputter coater and examined using a LEO 1550 field emission scanning electron microscope at the University of Kansas, Lawrence. All other samples were sputter- coated with gold-palladium or palladium in a Cressington sputter coater and examined using a FEI NOVA nanoSEM at the Ohio State University, Columbus. Squash preparations of undischarged capsules were made using samples fixed for SEM as above, but before critical-point drying and maintained in 70% ethanol. A very small piece of tissue was floated in a droplet of water and then compressed between a coverslip and a microscope slide; squash preparations were examined under DIC at 1000X.

213 Morphological measurements

Nematocyst morphotypes were identified based on the thorough morphological analysis performed in Chapter 3. After being categorized as one of the nine morphologies, each nematocyst was measured for a set number of variables in both SEM and light microscopy (or LM) preparations (Table 5.2). All nematocysts investigated by

SEM were measured for 15 different variables that focused on different aspects of the morphology of the basal tubule and spines, whereas nematocysts investigated by LM were measured for eight variables focused on the shape of the capsule and basal tubule.

Figure 5.1 provides a pictorial representation of some of the variables measured.

All measurements were performed using high quality images of each individual nematocyst, usually multiple images of each one to enable accurate measurement of all features. These images were transformed from tiff to tps files using the program tpsUtil version 1.50 (Rohlf 2012), and measurements were made in the tpsDig version 2.16

(Rohlf 2010). An image of a micrometer was used to set the scale for the light microscope images; the scale bar of each individual SEM image was used to calibrate the tpsDig program for measurement.

In SEM images, the capsule length and width were measured to give some sense of overall nematocyst size; other measurements included the width of the basal tubule at three different points along its length, lengths and widths of three different spines each progressively more distal, and the widths of the base and tip of spines of the spine in the middle region of the basal tubule. Width of the distal tubule was measured if present, however, for most of the p-rhabdoid B morphologies, the distal tubule was absent or

214 could not be measured. Wherever possible, the lack of a distal tubule measurement was

treated as missing data (see below).

For nematocysts with annulations along the basal tubule (i.e. colloponemes, all p-

rhabdoid B morphologies), five extra variables were measured, most of which are

inapplicable to other morphs of nematocysts (Table 5.2). These included width of the

annulations at both ends of the basal tubule (proximal and distal), the length of the

faltstück, and the width of the base and tip of spine 1, the spine measured that is closest to

the capsule (see Fig. 5.1). The faltstück is the proximal portion of the basal tubule when

two parts are present. As discussed in Chapter 3, colloponemes can be interpreted as

having a basal tubule that consists only of faltstück whereas the p-rhabdoid B1a form can

be interpreted as having only haupstück (the distal portion). Therefore, for the former,

the length of faltstück was equal to the length of the basal tubule and the latter had a

value of zero for this variable. The two portions of the basal tubule have different sized

spines (Schmidt 1969, see Chapter 3), therefore, measuring the width of the base and tip

of spine 1 will provide data on faltstück spine shape (if the nematocyst morph has this

feature) whereas the same variables for spine 2 will provide data on the haupstück spine shape.

Eight measurements were made on the light microscopy nematocysts. In an attempt to capture more of the shape of the capsule, the width of both the apical and non-

apical ends were measured in addition to the width of the midpoint of the capsule. The length of the basal tubule was also assessed and width of this feature was measured for three different regions (see Fig. 5.1).

215

Multivariate analysis

All variables were first tested for univariate normality using the Shapiro-Wilk (Shapiro and Wilk 1965) and the Jarque-Bera (Jarque and Bera 1987) tests, then tested for multivariate normality using Mardia’s (1970) multivariate skewness and kurtosis and the

Doornik and Hansen (1994) omnibus, all performed in PAST version 2.15 (Hammer et al.

2001). The focus of this study was not on size differences among morphs, but rather shape differences, therefore ratios were utilized to help study shape independently of size

(James and McCulloch 1990). To minimize the effect of overall size on the analysis, all variables in SEM and LM datasets were divided by capsule length and thereby scaling for size. Finally, the scaled data was log transformed (the faltstück variable was transformed as log (1 + x) as there were zero values in this variable) in order to help correct for the positive skew and bring in outliers (Bland and Altman 1996, Manikandan 2010), both of which were common in the distributions of these variables.

The transformed data were analyzed in three different ways: principle components analysis (PCA), nonmetric multidimensional scaling (NMS) using Euclidean distances, and linear discriminant function analysis. The first two analyses were performed in

PAST, and the first three axes were mapped in three dimensions to visualize the spatial relationships among the individual nematocysts. PCA is a standard data reduction technique however it can be sensitive to outliers and the lack of normality (James and

McCulloch 1990). Therefore, NMS, which is based on rank order only and does not require any underlying distribution, was also used to look for relationships among 216 nematocyst morphs (James and McCulloch 1990). The linear discriminant function analysis was performed in Minitab 15 version 1.30 (www.minitab.com) using the nematocyst morphologies as the a priori groups. Because this analysis does not handle missing data, the width of the distal tubule was not included in the two SEM analyses.

Cross validation was used to estimate misclassification probabilities. Due to the lack of normality in the data (see below), no tests of statistical significance were performed.

Results

The raw data that comprise all three data sets are found in Appendix A. A summary of nematocyst morphotypes and the taxa from which they were collected can be found in

Table 5.3. The SEM data was collected from 124 individual nematocysts from four body regions (acontia, body column, mestenterial filaments, and tentacles) in 26 different taxa.

Light microscope data was collected from 309 individual nematocysts from the same body regions and taxa as the SEM data (see Table 5.3). However, insufficient material was available to perform light microscopy on the coral Platygyra astreiformis for the p- rhabdoid A, and for the sea anemone Bartholomea annulata mesenterial filaments

(although only for p-rhabdoid B2s; there was enough material of the latter to do LM for the p-rhabdoid B2d).

Normality

For all data sets, the normal distribution was rejected for all raw variables using various normality tests, including the Shapiro-Wilk (Shapiro and Wilk 1965) and the Jarque-Bera

(Jarque and Bera 1987) tests both performed in PAST. Scaling the variables by capsule 217 length and performing a log transformation affected the distribution of some but not all variables. A normal distribution could not be rejected for the following variables once transformed: distal tubule width, spine lengths 1, 2 and 3, spine widths 1 and 3, spine 2 base and tip in the complete SEM dataset, capsule w basal tubule w2, distal tub w, spine

1 w, spine 2 w, spine 2 tip, spine 1 base spine 1 tip, and proximal annulation size in the annulated basal tubule dataset, and capsule width 3 in the light microscopy data set. In part due to the lack of univariate normality in all variables, multivariate normality is rejected using tests based on Mardia’s (1970) multivariate skewness and kurtosis and the

Doornik and Hansen (1994) omnibus computed in PAST. The lack of normality requires that the principle components and discriminant function analyses be treated as exploratory only; no statistical significance in groupings can be gleaned from this data.

PCA and NMS of SEM dataset of all morphologies

The principle components analysis of the SEM dataset containing all morphologies resulted in plots seen in Figs. 5.2 and 5.3. With three axes, a total of 83.39% of the variation is explained (Table 5.4). Only the diakanemes form a tight group separate from all other morphs (Figs. 5.2, 5.3), other morphs have a wider dispersion in the morphospace. However, most morphologies do group in particular portions of the plot.

Acanthonemes have a strong cluster below the x-axis (PCA 1) and to the left of the y-axis

(PCA 2) (Fig. 5.2). Some nematocysts of this morphology do map away from this main cluster, and there is no pattern to these outliers (they do not all come from the same taxon). The colloponemes cluster above the acanthonemes and form a tight group with the exception of one outlier (the Acontiaria sp. nematocyst from the acontia) that doesn’t 218 fall particularly close to any other groupings. The aphyllonemes cluster close to the acanthonemes, though they do separate somewhat along the z-axis (PCA 3, Fig. 5.3).

Although the diakanemes fall on the same side of the plot as the acanthonemes and aphyllonemes, they occupy a distinct portion of the morphospace. Hadronemes fall into two distinct groups, one close to the acanthonemes (which consists of all the

Ceriantheopsis americanus nematocysts) and another near various p-rhabdoids (which consists of all the Cerianthidae sp. nematocysts).

Among the p-rhabdoid forms, the majority of p-rhabdoid B2s samples fall in a cluster on the left side of the plot, above the x-axis. Three nematocysts of this morph are scattered on the right side of the plot but do not themselves form a tight grouping. These outliers are from different taxa, Boloceroides mcmurrici, Haliplanella lineata, and

Sagargetion lacerates, but are all from the tentacles. The p-rhabdoids A and B1a both are spread out on the right side of the graph along the x-axis, but separate somewhat along the y and z-axis (Fig. 5.2, 5.3). The three p-rhabdoid A nematocysts that cluster to the far right side of the graph are all from coral Platygyra astreiformis. Finally, the p- rhabdoid B2d are spread above the x-axis and do mix with outliers of other morphologies.

The NMS plot (Fig. 5.4) is very similar in how the individual nematocysts group to the PCA. The groupings are a bit neater and tighter in the NMS analysis than in PCA which may be an indication that outliers are having an effect on the principle components analysis. In particular, the p-rhabdoid A nematocysts form a tighter group in NMS, though the coral nematocysts are still outliers. Also, the p-rhabdoid B2s form a cleaner grouping, though again, the three outliers remain. 219 PCA and NMS of SEM dataset of annulated basal tubule morphologies

The principle components analysis of the annulated dataset containing additional variables resulted in the plots seen in Figures 5.5 and 5.6, and the results from NMS are in Figure 5.7. With three axes, a total of 87.9% of the variation is explained, an increase from the previous dataset (Table 5.4). The groupings from this more extensive dataset did not differ from those that resulted from the dataset with fewer variables but more morphologies. Adding these variables did not appear to improve the tightness of the clustering within the morphotypes.

PCA and NMS of light microscopy dataset

The principle components analysis of the light microscopy dataset resulted in the plots seen in Figures 5.7 and 5.8 and NMS results in Figure 5.9. With three axes, a total of

92.56% of the variation is explained (Table 5.4). In these plots, some morphotypes form distinct groups such as aphyllonemes and diakanemes. Colloponemes, p-rhabdoids A, and p-rhabdoids B1a have a broad spread in the x-axis, but still cluster in distinct portions of the plot. The p-rhabdoids B2s and B2d cluster on opposite ends of the x-axis with the

B2s morphs forming a large cluster on the one side of the morphospace whereas the B2d morphs cluster on the other side. However, both forms have a few individual nematocysts that cross to the other side and mingle with the other form. There is no particular taxonomic pattern in the p-rhabdoid B2d or B2s forms that fall outside the main cluster. Acanthonemes form a strong cluster on the left side of the plot, though outliers spread to the right. Some acanthonemes fall amongst the colloponemes to the far left, but these are all from one taxon and tissue (Halcurias levis tentacle). Most of the rest 220 of the ancanthonemes differ from colloponemes in the z-axis. Hadronemes are scattered throughout the plot and show no obvious clustering either by taxon or tissue structure indicating a high amount of variation in the morphology of this morph.

Discriminant function analysis of all three datasets

The results of the discriminant function analysis for the SEM all morphologies dataset are summarized in Table 5.5. Overall, the procedure had a correct classification rate of 75%.

Some groups are quite robust and have a high proportion of correct classification even with the stringent cross validation procedure. Acanthonemes, aphyllonemes, colloponemes, and p-rhabdoids B1a all have a proportion of correct classification of 80% or higher while diakanemes are always correctly classified. Three of the misclassifications of acanthonemes are of the same taxon and tissue (Bartholomea annulata tentacle). Hadronemes have a low proportion of correct classification at 57%; two of the three misclassifications are correctly predicted and only fail to be grouped using cross validation. No taxonomic pattern is evident in the three failed classifications.

However, in the p-rhabdoids A, three of the four misclassifications are from the coral

Platygyra astreiformis. Once again, these three nematocysts were correctly predicted but failed cross validation. The p-rhabdoid B2d has several misclassifications scattered in several different groups with no obvious taxonomic pattern. The one p-rhabdoid B2d that classified as an acanthoneme is a Haliplanella lineata tentacle nematocyst.

With the removal of other morphologies and the addition of a few variables, the proportion of correct classifications increased for all groups (annulated basal tubule dataset: Table 5.6). Overall, 85.7% of nematocysts are correctly classified. With the 221 removal of acanthonemes, colloponemes are always classified correctly, and other groups have gained a few correct classifications. Although classification improves for the p- rhabdoid B2d, it remains much lower than the proportion for other morphs.

The results of the discriminant function analysis for the light microscopy data set are summarized in Table 5.7. Overall, 85.4% are correctly classified. With this analysis both aphyllonemes and diakanemes are always correctly assigned, whereas acanthonemes, colloponemes, and p-rhabdoid B2s all have correct classification rates of over 82%. Similarly to the SEM dataset, p-rhabdoid B1a has a high successful classification rate. In contrast to the SEM results, the proportion of correctly classified p- rhabdoids A is high (91%), although light microscopy data for P. astreiformis was not included. Misclassifications between p-rhabdoids B2d and the p-rhabdoids B1a and B2s caused the rate of correct classifications to be lower for the B2d (though the rate is higher than in the SEM dataset). Finally, the hadroneme has a higher correct classification rate than in the SEM dataset, but is still lower than other categories. Misclassifications for the hadronemes are scattered across five different groups (more than any other morph) with no clear pattern.

Discussion

Normality of nematocyst data

Previous studies have demonstrated conflicting results when evaluating the normality of nematocyst length and widths and therefore the suitability of parametric statistical methods. Williams (1996, 1998, 2002) maintains that these variables have normal frequency distributions in the taxa he studied with no significant within-sample 222 variability. However, Acuña et al. (2003, 2004, 2007) consistently finds higher within sample than between sample variation and therefore used non-parametric methods.

Because the focus of this study is on the morphotypes themselves, nematocysts were pooled from across many taxa and body structures, and evaluation of within-sample variation is not informative given the low number of data points for any particular taxon.

However all variables measured have a strong positive skew in the raw state that was not always eliminated by the log transformation.

This skew in the data could be caused by several factors but two in particular seem most likely. First, these datasets pool all nine morphologies even though there are some distinct differences amongst those morphs and the samples are pulled from a wide phylogenetic range. Therefore, we might expect there to be different patterns of variation among the different morphologies as a result of different biological forces driving variation whereas for a variable to have a normal distribution, we would expect only one underlying force. Pooling the data in such a way may mix variables that are responding differently and therefore are not distributed normally. Second, particularly for the SEM dataset, some of these variables have quite small values. For some structures such as the spine there may be a biological/developmental limit as to how small the feature can be.

Most of these variables lack a left tail, instead stopping abruptly once a certain size is reached. Such a biological constraint will necessarily prevent the variables from having a normal distribution. Regardless of the cause, the lack of normality makes statistical conclusions unjustified but should not prevent the exploration of data by these methods.

223 Support for morphological groups

The classification of nematocyst morphological types has always been based on qualitative assessments of the variation (e.g. Stephenson 1928, Weill 1934, Carlgren

1940, Mariscal 1974, Schmidt 1969, 1974, Östman 2000, Chapter 3). A multivariate approach provides an opportunity to evaluate these groupings independently of human classification bias (although choice of characters and measuring mistakes prevent any study from being completely free of bias). Furthermore, the use of two kinds of data, scanning electron microscopy and light microscopy, in which different variables

(although not necessarily independent ones) are measured, provide two opportunities to evaluate these groupings and look for similarities between groups using quantitative data.

Although no statistical significance can be associated with this study, most morphologies studied here do show clustering in morphospace and can be distinguished from each other with a linear discriminant function. All morphologies (except the tightly clustered diakaneme and the diffusely distributed hadroneme) have outliers that may scatter far from its main cluster and makes the exact circumscription of each morphology difficult to determine. However, all but the two mentioned exceptions do have specific areas of morphospace where a main cluster forms and this cluster can form the basis of our organization of nematocyst morphologies. Specific details of each morphology are detailed below.

Acanthonemes, Aphyllonemes, and Colloponemes: These morphologies (with the addition of the hadroneme) make up a category that has been called “b-rhabdoids”

(Schmidt 1969), an amalgamation of two categories that others consider distinct, the 224 basitrichous isorhiza and the b-mastigophore (Carlgren 1940, Mariscal 1974, England

1991, see Chapter 3). Therefore, it is not surprising that these morphs all map close to each other in multivariate space. However, although none of the groups are exclusive of outliers, they do generally cluster in different spaces adjacent to each other (see Fig. 5.2-

5.9). The discriminant function analysis was highly successful in classifying each of these morphologies using both light and scanning electron microscopy data indicating that there are consistent morphological differences between the groups.

The acanthoneme is common across the entire order Actiniaria (see Chapter 4); therefore the high amount of variation seen in this morphotype is not surprising.

However, acanthonemes do cluster in morphospace with outliers generally showing no taxonomic pattern (except for the previously mentioned group of Halcurias levis outliers).

Colloponemes can be difficult to differentiate from acanthonemes using single length and width measurements with light microscopy (see Chapter 3) although most are clearly separated from the latter in SEM. Even with LM, when more measurements are taken, most, though not all, colloponemes cluster separately from acanthonemes. The discriminant function analysis was successful in classifying colloponemes with light microscopy data (with slightly higher success than with SEM data). Therefore, this morphology may be successfully identified from light microscopy alone if given the right data. As SEM study is often not feasible for most actiniarian biologists, the ability to use light microscopy to get reliable nematocyst classifications will be important in better understanding the distribution of the morphology (see Chapter 4 for a discussion of the problem). 225 Aphyllonemes are an interesting case in that they do not clearly separate from

acanthonemes in the SEM dataset yet are clearly distinct using the light microscope data.

The lack of differentiation with SEM may be in part due to the choice of characters. The lack of spines on the distal tubule, for example, is a character that would clearly differentiate it from acanthonemes. Therefore, a more thorough SEM analysis may yet find characters that result in this morphology clustering separately from acanthonemes in a multivariate analysis.

Diakanemes: In all analyses using either SEM or LM data, this group remains distinct

and even with cross validation members of this group are always correctly classified by

the linear discriminant function. The small sample size (one individual from one

species) raises the question as to whether this morphology would remain so distinct with

more samples. Diakanemes have an interesting morphology where they have a capsule

shape and distribution similar to aphyllonemes (elongate with a wider non-apical end

found only in mesentrial filaments of some actiniids), however they often have a visible

v-shaped notch at the end of the basal tubule and a wider diameter basal tubule which are

characters shared by p-rhabdoids (see Chapter 3). In both SEM and LM datasets,

diakanemes consistently cluster in morphospace closer to the “b-rhabdoid” types

(discussed above) than the p-rhabdoid types. However, despite sharing a similar

undischarged capsule shape, they do not appear to be morphologically similar to

aphyllonemes.

226 Hadronemes: Although this morphology is one of the easiest to differentiate qualitatively, no clustering for all samples was evident in either the SEM or LM data. In the SEM dataset, the nematocysts from the two taxa grouped in two distinct clusters: one near the

“b-rhabdoids” and another closer to the p-rhabdoids. However, making conclusions based on this data is difficult given that these two groupings also correspond to the two different sampled taxa. More extensive sampling may help determine if there truly are two distinct clusters or if these two taxa simply have nematocysts that represent ends of a continuum.

The hadroneme is variable in form with a stouter, wider form and an elongate thin form (See Fig. 4.2 in Chapter 4). Although it is possible that the two groupings in the

SEM morphospace are associated with these different forms, the light microscopy data supports the idea of a continuum. Both morphs were measured for SEM but no obvious clusters formed. Instead, this nematocyst is widely dispersed in morphospace. None seem to cluster particularly with any other morphology as the light microscope discriminant analysis exemplifies; five hadronemes are misclassified into five different morphologies. Possibly the use of other characters would result in clearer clustering (e.g. the absence of a v-shaped notch at the end of the basal tubule). Regardless, of the morphologies studied here, the hadroneme clearly most needs more data to clarify its placement in morphospace.

p-rhabdoids A: Although this group was spread over a large amount of morphospace, it

generally is separate from other morphologies, though it has some interesting outliers. In

the SEM dataset, the three coral examples are extreme outliers (unfortunately no light 227 microscopy data of this type was obtained) and are misclassified in the discriminant

function analysis. Schmidt (1974) distinguished the coral p-rhabdoid from those of

Actiniaria calling it a p-rhabdoid D, a distinction based on the length of the basal tubule

spines and the presence of spines on the distal tubule. Although in gross morphology the

p-rhabdoids A and D share several similarities, the multivariate analysis does show the

coral nematocysts do group separately off to one end of the axis. Therefore, calling them

by another name could be justified. However, because the distribution of the morph is

relatively dispersed, I prefer to interpret the coral nematocysts as an end to a continuum.

A Bunodosoma cavernata nematocyst maps to an area of morphospace adjacent to the

coral examples; therefore, although they are not near the main cluster of p-rhabdoids A,

they do not cluster with anything else and are near another representative of the type. In

contrast, the antipatharian p-rhabdoid A falls close to several actiniarian samples:

Schmidt (1974) considered it to be the same form as that found in Actinaria.

p-rhabdoids B1a: Like the p-rhabdoid A, this morphology is widely dispersed and has some distant outliers. Despite not appearing to be morphologically distinct and lacking a main cluster of nematocysts in morphospace, the linear discriminant function is relatively highly successful in correctly classifying this morphology. This procedure maximizes differences between groups rather than the reduction of the data (Manly 2005), which may cause the discrepancy. Regardless, the B1a forms occupy their own morphospace and do not cluster with any other morphology so although they do not form a tight cluster, seem to be a consistent group.

228 p-rhabdoids B2d and B2s: These two morphologies are distinguished by differences in capsule shape and some features specific to p-rhabdoid B forms (specifically, B2d usually has a short faltstück and no change in annulation size along the basal tubule).

The extra data set had characters that pertained to these features and could potentially help separate the groups. However, this added data did not substantially improve groupings or classification (the increase in correct classifications for p-rhabdoid B2d in the discriminant function analysis is more likely to be the result of other morphologies not being included in the analysis rather than an effect of the extra variables). In all plots, the B2d and B2s have large clusters at opposite ends with outliers crossing into each other’s morphospace. The two forms together could be interpreted as a continuum with the outliers representing transitional forms between the two. Although the p-rhabdoid

B2ds do occupy their own morphospace, their distribution is broad with no obvious clustering. The discriminant function analysis had the most difficulty outside the hadronemes in correctly classifying this morphotype. Taken together, the biometric analysis suggests that I may be misinterpreting this group and what defines it morphologically. In contrast, the p-rhabdoids B2s did form a distinct cluster (though they also had some members dispersed away from this cluster). This morphology seems more likely to capture some natural grouping.

Further directions in multivariate analysis of nematocyst variation

This study focused on using measured variables to access shape information about the nematocysts. However, other techniques may be superior at fully capturing shape information, such as the use of landmark-based geometric morphometrics (see Adams et 229 al. 2004 for review). The use of landmarks in techniques such as the thin plate spline does require an assumption of homology at the chosen landmarks (Brookstein 1986

1991). Determining which points of the capsule are homologous to points on another, differently shaped capsule is difficult outside of the apical structure (which is presumed to be homologous for all nematocysts even if it does take different forms as discussed in

Chapter 2). Therefore, outline methods where the outline of a shape is quantified with no regard for homologous points might be more appropriate (Adams et al. 2004).

Particularly with the light microscopy data where capsule shape is of interest in discriminating amongst nematocyst morphologies, techniques such as eigenshape analysis or elliptical Fourier analysis might be more informative in a PCA than the variables used here and therefore represent a possible future direction in nematocyst biometric research.

Furthermore, even with the addition of these new variables in a biometric context, the full variation of nematocysts is not captured here. Some variables, such as the presence or absence of a v-shaped notch at the end of the basal tubule in undischarged nematocysts studied by light microscopy, could be coded for as a presence-absence character and may affect these groupings (particularly of the hadroneme which does not have such a notch and which groups near things that do in this study). Furthermore, some of the variables from the annulated dataset could be coded as presence-absence (such as the presence of a Faltstück, or annulations on the basal tubule) and incorporated into the larger dataset. This inclusion may better separate some morphologies such as the hadroneme and p-rhabdoid A from the p-rhabdoid B forms. Finally, some features of the nematocysts clearly vary among morphotypes but are difficult to measure consistently. 230 For example, distance between individual spines and between whorls of spines on the basal tubule varies among nematocyst types (Schmidt 1969, 1974, Chapter 3), however the dense packing of spines in some morphs prevented reliable measurement of these features. If these characters could be reliably measured then they might prove to be useful in distinguishing morphs using a multivariate approach. This study was intended as an initial foray into using biometric approaches to better evaluate and understand the groupings of morphologies. Even with limited data, the results indicate some success in finding quantitative support for qualitatively based nematocyst type distinctions. More extensive datasets and approaches may prove to be even more successful.

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234 Table 5.1. Summary of taxa and collection locality used in the multivariate study Higher Taxon Species Location collected Actiniaria Acontiaria sp. Jakyakdo Island, S. Korea Actinostolid sp. San Juan Island, WA, USA Adamsia palliata Ringhaddy Sound, Strangford Lough, N. Ireland Aiptasia sp. Aquarium Adventure, Columbus, OH, USA Anemonia sulcata Ballymacormick Pt, Groomsport, N. Ireland Bartholomea annulata University of Virgin Islands, St. Thomas, US Virgin Islands Boloceroides Kyoto University Seto Marine Lab, mcmurrici Shirahama, Japan Bunodosoma Galveston, TX, USA cavernata Calliactis polypus Jeju Island, S. Korea Cereus pedunculatus SW of Mahee Island, Strangford Lough, N. Ireland Condylactis gigantea Aquarium Adventure, Columbus, OH, USA Diadumene leucolena Marine Biological Lab, Woods Hole, MA, USA Diadumene sp. Jakyakdo Island, S. Korea Entamecea quadcolor Jeju Island, S. Korea Halcurias levis Kyoto University Seto Marine Lab, Shirahama, Japan Haliplanella lineata Jakyakdo Island, S. Korea Nemanthus nidtus Jeju Island, S. Korea Paracondylactis Jakyakdo Island, S. Korea hertwigi Sagartia elegans SW of Mahee Island, Strangford Lough, N. Ireland Sagartigeton lacerates Ringhaddy Sound, Strangford Lough, N. Ireland Sagartigeton undatus Ringhaddy Sound, Strangford Lough, N. Ireland Urticina felia SW of Mahee Island, Strangford Lough, N. Ireland Antipatharia Antipathes griggi Auau Channel, HI, USA Ceriantharia Ceriantheopsis Gulf Specimen Marine Lab, Panacea FL, americanus USA Cerianthidae sp. Reef Hot Spot, Inglewood, CA, USA Scleractinia Platygyra asteriformis Gulf Specimen Marine Lab, Panacea, FL, USA 235

Table 5.2. List of variables used in all three multivariate datasets. All morph SEM Annulated SEM (additional Light microscope to all morph variables) Capsule length Length of Faltstück Capsule length Capsule width at midpoint Length of base of spine 1 Width at apex of capsule Basal tubule length Width of tip of spine 1 Width of midpoint of capsule Basal tubule width 1 Width of proximal Width of non-apical end annulations on basal tubule of capsule Basal tubule width 2 Width of distal annulations Length of basal tubule on basal tubule Basal tubule width 3 Basal tubule width 1 Distal tubule width Basal tubule width 2 Spine length 1 Basal tubule width 3 Spine width 1 Spine length 2 Spine width 2 Spine length 3 Spine width 3 Length of base of spine 2 Width of tip of spine 2 See Fig. 5.1 and text for details on each variable. The annulated SEM dataset includes all the variables in the all morph dataset in addition to the ones listed here.

236 Table 5.3. List of all tissues and taxa from which the multivariate datasets were derived with numbers of individual nematocysts studied. Nematocyst Taxon Body Structure Microscopy type Technique: Count acanthoneme Acontiaria sp. Tentacle SEM: 1, LM: 5 Actinostola sp. Tentacle SEM: 7, LM: 8 Adamsia p. Acontia SEM: 1, LM: 5 Anemonia s. Tentacle SEM: 2, LM: 7 Bartholomea a. Tentacle SEM: 4, LM: 5 Calliactis p. Tentacle SEM: 1, LM: 10 Calliactis p. Acontia SEM: 4, LM: 6 Condylactis g. Tentacle SEM: 8, LM: 6 Entacmaea q. Tentacle SEM: 6, LM: 6 Halcurias l. Tentacle SEM: 2, LM: 6 Nemanthus n. Mesenterial fil SEM: 3, LM: 7 Urticina f. Tentacle SEM: 4, LM: 5 aphylloneme Anemonia s. Mesenterial fil SEM: 4, LM: 8 Paracondylactis h. Mesenterial fil SEM: 1, LM: 5 colloponeme* Acontiaria sp. Acontia SEM: 1, LM: 4 Aiptasia sp. Acontia SEM: 3, LM: 5 Cereus p. Acontia SEM: 1, LM: 9 Sagartia e. Acontia SEM: 2, LM: 4 Sagartiogeton l. Tentacle SEM: 1, LM: 3 diakaneme Urticina f. Mesenterial fil SEM: 7, LM: 10 hadroneme Ceriantheopsis a. Column SEM: 1, LM: 5 Ceriantheopsis a. Mesenterial fil SEM: 2, LM: 7 Cerianthidae sp. Tentacle SEM: 4, LM: 12 p-rhabdoid A Actinostola sp. Tentacle SEM: 3, LM: 7 Anemonia s. Mesenterial fil SEM: 3, LM: 13 Antipathes g. Mesenterial fil SEM: 1, LM: 5 Bunodosoma c. Mesenterial fil SEM: 3, LM: 7 Platygyra a. Tentacle SEM: 3, LM: 0 p-rhabdoid B1a* Calliactis p. Mesenterial fil SEM: 3, LM: 7 Diadumene l. Mesenterial fil SEM: 4, LM: 8 Nemanthus n. Mesenterial fil SEM: 1, LM: 9 Sagartia e. Mesenterial fil SEM: 4, LM: 5 p-rhabdoid B2d* Aiptasia sp. Mesenterial fil SEM: 3, LM: 4 Bartholomea a. Mesenterial fil SEM: 2, LM: 6 Diadumene l. Mesenterial fil SEM: 1, LM: 4 Diadumene sp. Tentacle SEM: 2, LM: 5 Continued

237

Table 5.3 continued Haliplanella l. Tentacle SEM: 1, LM: 6 Sagartia e. Column SEM: 1, LM: 3 Sagartia e. Mesenterial fil SEM: 1, LM: 5 Sagartiogeton l. Column SEM: 1, LM: 4 p-rhabdoid B2s* Acontiaria sp. Acontia SEM: 1, LM: 5 Aiptasia sp. Acontia SEM: 2, LM: 6 Bartholomea a. Mesenterial fil SEM: 1, LM: 0 Boloceroides m. Tentacle SEM: 2, LM: 6 Cereus p. Acontia SEM: 2, LM: 5 Cereus p. Mesenterial fil SEM: 1, LM: 6 Diadumene sp. Mesenterial fil SEM: 1, LM: 5 Haliplanella l. Tentacle SEM: 1, LM: 3 Sagartia e. Acontia SEM: 2, LM: 6 Sagartiogeton l. Acontia SEM: 1, LM: 5 Sagartiogeton l. Tentacle SEM: 1, LM: 3 Sagartiogeton u. Acontia SEM: 1, LM: 5 Sagartiogeton u. Mesenterial fil SEM: 1, LM: 5 Total 26 taxa 4 Body structures SEM: 124, LM:309 SEM =scanning electron microscopy, LM= light microscopy. * indicate nematocyst morphs that were included in the annulated basal tubule dataset.

238

Table 5.4. Results from principle components analysis. Dataset PC Eigenvalue % variance SEM: all morphologies 1 0.358 69.99 2 0.0411 8.037 3 0.0274 5.366 Total: 83.39 SEM: annulated basal tubule 1 0.631 77.06 2 0.0559 6.825 3 0.0329 4.014 Total: 87.9 Light microscopy 1 0.1454 74.00 2 0.02841 14.46 3 0.008053 4.099 Total: 92.56

239 Table 5.5. Results from discriminant function analysis of the all morphology dataset using cross validation. True Group Assigned Acan Aphy Coll Diak Hadr p-A p- p- p- Group B1a B2d B2s Acan 36 1 1 1 Aphy 5 4 Coll 1 7 2 Diak 1 7 Hadr 4 4 1 p-A 1 7 1 p-B1a 2 2 10 2 p-B2d 1 7 4 p-B2s 1 11 Total N 43 5 8 7 7 13 12 12 17 N correct 36 4 7 7 4 7 10 7 11 Prop 0.84 0.80 0.86 1 0.57 0.54 0.83 0.58 0.65 Acan=Acanthoneme, Aphy=Aphylloneme, Coll=Colloponeme, Diak=diakaneme, Hadr=hadroneme, p-A= p-rhabdoid A, p-B1a= p-rhabdoid B1a, , p-B2d= p-rhabdoid B2d, p-B2s= p-rhabdoid B2s

240

Table 5.6. Results from discriminant function analysis of the annulated basal tubule dataset using cross validation. True Group Assigned Coll p- p- p- Group B1a B2d B2s Coll 8 1 1 p-B1a 11 1 p-B2d 1 9 2 p-B2s 1 14 Total N 8 12 12 17 N correct 8 11 9 14 Prop 1 0.92 0.75 0.82

Acan=Acanthoneme, Aphy=Aphylloneme, Coll=Colloponeme, Diak=diakaneme, Hadr=hadroneme, p-A= p-rhabdoid A, p-B1a= p-rhabdoid B1a, , p-B2d= p-rhabdoid B2d, p-B2s= p-rhabdoid B2s

241 Table 5.7. Results from discriminant function analysis of the light microscopy data using cross validation True Group Assigned Acan Aphy Coll Diak Hadr p-A p- p- p- Group B1a B2d B2s Acan 63 2 1 1 Aphy 13 1 Coll 9 22 1 Diak 1 10 1 1 Hadr 3 19 p-A 1 31 2 2 p-B1a 1 25 3 p-B2d 1 1 2 29 7 p-B2s 4 50 Total N 76 13 25 10 24 34 29 37 60 N correct 63 13 22 10 19 31 25 29 50 Prop 0.83 1 0.88 1 0.79 0.91 0.86 0.78 0.83 Acan = Acanthoneme, Aphy = Aphylloneme, Coll = Colloponeme, Diak = diakaneme, Hadr = hadroneme, p-A = p-rhabdoid A, p-B1a = p-rhabdoid B1a, , p-B2d = p-rhabdoid B2d, p-B2s = p-rhabdoid B2s

242

Figure. 5.1. Morphometic measurements taken for light and scanning electron microsope datasets. A. Acanthoneme from Actinostola sp. tentacle, scale bar =20 µm. Measurements on the capsule included length (L), apical w (A), midpoint width (M), and Non-apical width (N). B. p-rhabdoid B2s from Aipstasia sp. acontia, scale bar =20 µm. Measurements on the basal tubule included length (L), width just under the apex (A), width at the midpoint (M) and width of the distal tip (D). C. p-rhabdoid B2s from Cereus pedunculatus acontia, scale bar = 20 µm. Measurements on the basal tubule included length (L), width near apex (A), midpoint width (M), and width at the end of the basal tubule (E). Whenever possible, width of the distal tubule was included as well. D. Acanthoneme from Urticina felia mesenterial filament, scale bar = 4 µm. Measurements on spines included length (L), width at the midpoint (M), width at the base of the spine (B), and width at the tip (T). M and L were measured for a spine near the start of the basal tubule, at the midpoint, and near the end of the basal tubule. Base and tip widths were taken from the spine at the midpoint of the basal tubule.

243

Figure 5.1

244

Figure 5.2. Plot of principle component 1 vs. 2 for the dataset that included all morphologies. See key for labels of individual morphotypes.

245

Figure 5.3. Three-dimensional plot of principle components analysis for the dataset that included all morphologies.

246 Figure 5.4. NMS plot using Euclidian distance and three dimensions of the dataset the included all morphologies.

247 Figure 5.5. Plot of principle component 1 vs. 2 for the annulated tubule dataset.

248 Figure 5.6. Three-dimensional plot of principle components analysis for the annulated basal tubule dataset.

249 Figure 5.7. NMS plot using Euclidian distance and three dimensions of the annulated basal tubule dataset.

250 Figure 5.8. Plot of principle component 1 vs. 2 for light microscope data set.

251

Figure 5.9. Three-dimensional plot of principle components analysis for the light microscope dataset.

252 Figure 5.10. NMS plot using Euclidean distance and three dimensions of the light microscope data set.

253

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