CONTRIBUTIONS Phylogenetic Relationships of the Earliest Anisostrophically Coiled Gastropods . T)T] ,r PT]'T) 1 I n tr -a ||f I Ppj • a*' tAjfy. j vi J UK ri Cl ^4' . \ J m J 1 K ! Ifi f j wT lllr/^7 V M VO \\ W SERIES PUBLICATIONS OF THE SMITHSONIAN INSTITUTION Emphasis upon publication as a means of “diffusing knowledge” was expressed by the first Secretary of the Smithsonian. In his formal plan for the Institution, Joseph Henry outlined a program that included the following statement: "It is proposed to publish a series of reports, giving an account of the new discoveries in science, and of the changes made from year to year in all branches of knowledge." This theme of basic research has been adhered to through the years by thousands of titles issued in series publications under the Smithsonian imprint, commencing with Smithsonian Contributions to Knowledge in 1848 and continuing with the following active series: Smithsonian Contributions to Anthropology Smithsonian Contributions to Botany Smithsonian Contributions to the Earth Sciences Smithsonian Contributions to the Marine Sciences Smithsonian Contributions to Paleobiology Smithsonian Contributions to Zoology Smithsonian Folklife Studies Smithsonian Studies in Air and Space Smithsonian Studies in History and Technology In these series, the Institution publishes small papers and full-scale monographs that report the research and collections of its various museums and bureaux or of professional colleagues in the world of science and scholarship. The publications are distributed by mailing lists to libraries, universities, and similar institutions throughout the world. Papers or monographs submitted for series publication are received by the Smithsonian Institution Press, subject to its own review for format and style, only through departments of the various Smithsonian museums or bureaux, where the manuscripts are given substantive review. Press requirements for manuscript and art preparation are outlined on the inside back cover. Lawrence M. Small Secretary Smithsonian Institution SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY • NUMBER 88 Phylogenetic Relationships of the Earliest Anisostrophically Coiled Gastropods Peter J. Wagner Smithsonian Institution Press Washington, D.C. 2002 ABSTRACT Wagner, Peter J. Phylogenetic Relationships of the Earliest Anisostrophically Coiled Gastro¬ pods. Smithsonian Contributions to Paleobiology, number 88, 152 pages, 37 figures, 3 tables, 2002.—In order to explore the phylogenetic relationships among early gastropods, cladistic analyses were conducted of nearly 300 “archaeogastropod” species known from the latest Cambrian through the Silurian. The study includes an extended outgroup analysis of Cambrian molluscs. The resulting estimates of gastropod phytogeny differ not only from traditional ideas about early gastropod relationships, but also from most alternative notions. Outgroup analyses suggest that gastropods had ancestors among the Tergomya (= Monoplacophora of many work¬ ers) of the Middle or Late Cambrian. Putative gastropods from older strata (e.g., the Pelagiel- lida and early Onychochilidae) apparently are not closely related to gastropods. The hypothesized ancestor of gastropods possessed dextral-coiling, septation, a deep sinus, and a peripheral band. An anal slit is commonly described as a synapomorphy of gastropods that many clades subsequently lost; however, this study suggests that the slit is a rare, highly derived, and polyphyletic character among early Paleozoic species, and that the ancestors of most “advanced” clades (e.g., the Apogastropoda) never had slits. This study suggests that two major subclades evolved by the earliest Ordovician. The diag¬ noses and definitions of these two subclades best correspond to the traditional diagnoses and definitions of the Euomphalina and Murchisoniina. The Pleurotomarioidea is not a paraphyletic ancestral taxon as typically suggested, but instead it is a polyphyletic assemblage derived multi¬ ple times from “euomphalinae” and “murchisoniinae” species. The Bellerophontina is at least diphyletic, as the taxon includes both the ancestors of “archaeogastropods” and a clade of planispiral species that is secondarily derived from “archaeogastropods.” Macluritoids sensu stricto represent a restricted subclade of the “euomphalinae”; other supposed macluritoids evolved among different euomphalinae subclades or are not gastropods. Early Paleozoic species previously classified as caenogastropods (i.e., the Loxonematoidea and Subulitoidea) represent separate murchisoniinae subclades, with some putative members of the Subulitoidea derived within the Loxonematoidea. Early Paleozoic species assigned to the Trochoidea also represent several subclades, with most of those clades having evolved from the “euomphalinae.” An extensive taxonomic revision is presented, which removes all early Paleozoic taxa from the Pleurotomariina and broadly expands the definitions of the Euomphalina and Murchisoniina. Official publication date is handstamped in a limited number of initial copies and is recorded in the Institution’s annual report, Annals of the Smithsonian Institution. SERIES COVER DESIGN: The trilobite Phacops rana Green. Library of Congress Cataloging-in-Publication Data Wagner, Peter J. Phylogenetic relationships of the earliest anisostrophically coiled gastropods / Peter J. Wagner, p. cm.— (Smithsonian contributions to paleobiology ; no. 88) Includes bibliographic references. I. Gastropoda, Fossil. I. Title. II. Series. QE808.W34 1999 560 s-dc21 [564\scl36\.3] 98-52464 © The paper used in this publication meets the minimum requirements of the American National Standard for Permanence of Paper for Printed Library Materials Z39.48—1984. Contents Page Introduction. 1 Acknowledgments. 1 “Archaeogastropods”—A Temporary Definition . 2 Review of Previous Phylogenetic Hypotheses. 2 Material. 4 Specimens. 4 Biogeography of Analyzed Species. 4 Cladistic Characters. 5 Homology Versus Architecture. 5 The Paucity of Shell Characters Revisited . 5 Sinuses, Slits, Selenizones, and Peripheral Bands . 9 Shell Mineralogy and Protoconchs. 11 Phylogenetic Analysis. 11 Character Deweighting 1—Balancing Continuous Characters. II Character Deweighting 2—Asymmetry and Changing Homologies . 12 Character Analyses . 13 Cambrian Molluscs and the Choice of an Outgroup. 15 Results. 16 Cladogram Statistics and Descriptions . 16 Outgroup Analyses . 17 Relationships among Early Ordovician Species. 21 I. “Euomphalinaes”. 21 1.1. “Ophiletoids”. 21 1.2. “Macluntoids’'. 23 1.3. “Ceratopeoids” . . .. 26 1.3.1. “Raphistomatids” . 26 1.3.1.1. “Lesueurillines”. 26 1.3.1.2. “Holopeines” . 29 1.3.2. “Helicotomids”. 29 1.3.2.1. “Ophiletinines”. 31 1.3.2.2. “Euomphalopterines”. 31 1.3.2.2.1. “Anomphalides”. 31 1.3.2.2.2. “Poleumitides” . 31 1.3.2.2.3. “Pseudophorides” . 33 II. “Murchisoniinaes”. 35 II. 1. “Plethospiroids” . 39 11.2. “Straparollinoids”. 39 11.3. “Hormotomoids”. 39 11.3.1. “Subulitids”. 39 11.3.2. “Cyrtostrophids” . 43 11.3.2.1. “Goniostrophines” . 46 11.3.2.2. “Omospirines” . 46 11.4. “Eotomarioids” . 50 11.4.1. “Lophospirids”. 50 11.4.2. “Clathrospirids”. 52 11.4.2.1. “Liospirines’ 7 . 52 11.4.2.2. “Brachytomariines” . 55 iii IV SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 11.4.2.2.1. “?Palaeoschismides” . 55 11.4.2.2.2. “Phanerotrematides” . 57 11.4.2.2.3. “Luciellides” . 58 11.4.2.2.4. “Planozonides”. 58 Problematica . 61 Platyceratoidea . 61 Neritoidea . 63 Oriostomatoidea . 63 Discussion. 65 Paleozoic Pleurotomarioids: An Oxymoron?. 65 “Euomphalinaes”: Two Gills or One?.65 Implications for Relationships among Extant Gastropods . 66 Systematic Paleontology. 68 Class Gastropoda Cuvier, 1797 . 69 Order “Archaeogastropoda” Thiele, 1925 . 69 Family jSlNUOPElDAE Wenz, 1938 . 69 Genus ^Sinuopea Ulrich, 1911 . 69 Genus f Schizopea Butts, 1926 . 70 Genus Euconia Ulrich in Ulrich and Scofield, 1897 . 70 Genus Gasconadia Ulrich in Weller and St. Clair, 1928 . 70 ?Genus Calaurops Whitfield, 1886 . 70 Suborder Euomphalina de Koninck, 1881 . 70 Superfamily fOPHiLETOiDEA trans. nov. Knight, 1956 . 70 Genus |Ophileta Vanuxem, 1842 . 71 Genus f Lecanospira Butts, 1926 . 71 Genus Ecculiomphalus Portlock, 1843 . 71 Genus t Asgardaspira, new genus. 71 Genus Lytospira Koken, 1896 . 71 Superfamily Macluritoidea Fischer, 1885 . 71 Genus ^Macluritella Kirk, 1927 . 72 Genus ^Teiichispira Yochelson and Jones, 1968 . 72 Genus f Maclurites Le Sueur, 1818. 72 Genus Maclurina Ulrich and Scofield, 1897 . 72 Genus Palliseria Wilson, 1924 . 72 ?Genus Rousseauspira Rohr and Potter, 1987 . 73 Superfamily fEuOMPHALOiDEA de Koninck, 1844 . 73 Family IRaphistomatidae Koken, 1896 . 73 Genus ^Ceratopea Ulrich, 1911 . 73 Genus Bridgeites Flower, 1968a. 73 Genus Orospira Butts, 1926 . 73 Genus Raphistoma Hall, 1847 . 73 Genus fScalites Emmons, 1842 . 74 Family Holopeidae Wenz, 1938 . 74 Genus Raphistomina Ulrich and Scofield, 1897 . 74 Genus f Pachystrophia Pemer, 1903 . 74 Genus Sinutropis Pemer, 1903 . 75 Genus Umbospira Pemer, 1903 . 75 Genus Holopea Hall, 1847 . 75 Family Lesueurillidae, new family. 75 Genus ^Eccyliopterus Remele 1888 . 75 Genus \Lesueurilla Koken, 1898 . 76 Genus Mestoronema, new genus. 76 Genus Pararaphistoma Vostokova, 1955 . 76 Family tHELlCOTOMIDAE Wenz, 1938 . 76 NUMBER 88 V Genus |Lophonema Ulrich in Purdue and Miser, 1916 . 76 Genus Linsleyella Rohr, 1980 . 76 Genus f Helicotoma Salter, 1859 . 77 Genus Palaeomphalus Koken, 1925 . 77 Genus Ophiletina Ulrich and Scofield, 1897 . 77 Family fEuOMPHALlDAE de Koninck, 1881 . 77 Genus \Boucotspira Rohr, 1980 . 77 Genus ^Euomphalopterus Roemer, 1876 . 77 Genus Spinicharybdis Rohr and Packard, 1982 . 78 Genus f Poleumita Clarke and Ruedemann, 1903 . 78 Genus Nodonema Linsley, 1968 . 78 Genus Centrifugus Bronn, 1834 . 78 Genus Euomphalus Sowerby, 1814. 78 Genus Straparollus de Montfort, 1810 . 79 Genus Micromphalus Knight, 1945 . 79 Family Anomphalidae Wenz, 1938 . 79 Genus ^Trochomphalus Koken, 1925 . 79 Genus Pycnomphalus Lindstrom, 1884 . 79 Family Pseudophoridae Miller, 1889 . 79 Genus Pseudophorus Meek, 1873 . 79 Genus Pseudotectus Pemer, 1903 . 80 Genus \Discordichilus Cossmann, 1918. 80 Genus Hystricoceras Jahn, 1894 . 80 Genus Streptotrochus Pemer, 1903 . 80 Genus Elasmonema Fischer, 1885 . 80 Suborder Murchisoniina Cox and Knight, 1960 . 80 Superfamily fMuRCHiSONiOiDEA Koken, 1896 . 81 Family fHORMOTOMlDAE Wenz, 1938 . 81 Genus ’fHormotoma Salter, 1859 . 81 Genus f Coelocaulus Oehlert, 1888. 81 Genus Catazone Pemer, 1903 . 81 Genus Mesocoelia Pemer, 1907 . 81 Genus Plethospira Ulrich in Ulrich and Scofield, 1897 . 81 Family Murchisoniidae Koken, 1896 . 82 Genus 'fMurchisonia d’ Archaic, 1841. 82 Genus Morania Homy, 1953 . 82 Genus Michelia Roemer, 1854 . 82 Superfamily Loxonematoidea Koken, 1889 . 82 Family Loxonematidae Koken, 1889 . 82 Genus ^Loxonema Phillips, 1841 . 82 Genus ^Omospira Ulrich and Scofield, 1897 . 83 Genus Diplozone Pemer, 1907 . 83 Genus Rhabdostropha Donald, 1905 . 83 Genus Spiroecus Longstaff, 1924 . 83 Genus Macrochilus Lindstrom, 1884 . 83 Genus Stylonema Pemer, 1907 . 84 Superfamily EOTOMARIOIDEA Ulrich and Scofield, 1897 . 84 Family Eotomariidae Wenz, 1938 . 84 Genus "\Clathrospira Ulrich and Scofield, 1897 . 84 ?Genus Spirotomaria Koken, 1925 . 84 Genus f Eotomaria Ulrich and Scofield, 1897 . 84 Genus Paraliospira Rohr, 1980 . 84 Genus Liospira Ulrich and Scofield, 1897 . 85 Family IGOSSELETINIDAE Wenz, 1938 . 85 VI SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Subfamily fEuRYZONlNAE, new subfamily . 85 Genus | Deaechospira, new genus . 85 Genus f Cataschisma Branson, 1909 . 86 ?Genus Palaeoschisma Donald, 1902 . 86 Genus \Pleurorima Pemer, 1907 . 86 Genus Euryzone Koken, 1896 . 86 Genus Latitaenia Koken, 1925 . 86 Subfamily GOSSELETININAE Wenz, 1938 . 86 Genus \Stenoloron Oehlert, 1888 . 86 Genus Platyloron Oehlert, 1888 . 87 Genus Umbotropsis Pemer, 1907 . 87 Family IPhanerotrematidae Knight, 1956 . 87 Genus ^Brachytomaria Koken, 1925 . 87 Genus Phanerotrema Fischer, 1885 . 87 Genus Ulrichospira Donald, 1905 . 88 Family Luciellidae Knight, 1956 . 88 Genus Conotoma Pemer, 1907 . 88 Genus Prosolarium Pemer, 1907 . 88 Genus Oehlertia Pemer, 1907 . 88 Superfamily Lophospiroidae Wenz, 1938 . 89 Family LOPHOSPIRIDAE Wenz, 1938 . 89 Genus Ectomaria Koken, 1896 . 89 Genus Donaldiella Cossmann, 1903 . 89 Genus Lophospira Whitfield, 1886 . 89 Genus Proturritella Koken, 1889 . 89 Genus Eunema Salter, 1859 . 89 Genus Gyronema Ulrich in Ulrich and Scofield, 1897 . 89 Genus Ruedemannia Foerste, 1914. 89 Genus Loxoplocus Fischer, 1885 . 90 Genus Longstaffia Cossman, 1908 . 90 Genus Arjamannia Peel, 1975 . 90 Genus Trochonemella Okulitch, 1935 . 90 Family Trochonematidae Zittel, 1895 . 90 Genus Trochonema Salter, 1859 . 90 Genus Globonema Wenz, 1938 . 90 Superfamily Straparollinoidea, new superfamily. 90 Family STRAPAROLLrNlDAE, new family. 90 Genus ^Straparollina Billings, 1865 . 90 Genus Daidia Salter, 1859 . 90 Genus Haplospira Koken, 1897 . 91 Superfamily SUBULITOIDEA Lindstrom, 1884 . 91 Family SUBULITIDAE Lindstrom, 1884 . 91 Genus ^Eroicaspira, new genus . 91 Genus 'fSubulites Emmons, 1842 . 91 Genus Fusispira Hall, 1872 . 91 Genus Cyrtospira Ulrich in Ulrich and Scofield, 1897 . 91 Conclusions. 91 Appendix 1: Characters and Character States . 93 Appendix 2: Data Matrix . 100 Appendix 3: Stratigraphic Data . 134 Literature Cited.. Phylogenetic Relationships of the Earliest Anisostrophically Coiled Gastropods Peter J. Wagner 1 Introduction The renewed interest in gastropod phylogenetics (see Bieler, 1992, for a review) has generally neglected fossil taxa. This is unfortunate because solely neontological studies exclude many interesting gastropod clades and likely underestimate the com¬ plexity of gastropod evolution. The earliest members of di¬ verse, long-lived clades also might possess informative combi¬ nations of plesiomorphies and apomorphies (see Gauthier et al., 1988; Donoghue et al., 1989). Many major taxa (extinct and extant) apparently diverged very early in gastropod history (Knight et al., 1960) without obvious intermediates (Erwin, 1990a); therefore, a phylogenetic analysis of the earliest gastro¬ pods could contribute much to contemporary ideas about gas¬ tropod relationships. The last phylogenetic study to concentrate on Early Paleo¬ zoic gastropods was by Knight (1952). Later workers presented alternative ideas about relationships among particular taxa (e.g., Yochelson, 1967, 1984; Runnegar, 1981; Linsley and Kier, 1984), but none have conducted large-scale phylogenetic analyses. In this paper, I discuss the results of phylogenetic analyses that encompass 295 species of early anisostrophically 'Author’s Note: The scientific content of this paper originally appeared as a chapter in the author’s 1995 dissertation, “The Generation and Maintenance of Morphologic and Phylogenetic Diversity among Early Gastropods” (Uni¬ versity of Chicago). This paper was slightly modified from that chapter and was accepted for publication in 1996. Thus, this work is older than studies published by the author since 1997, and readers should consider conclusions in those papers to supercede conclusions in this work. Also, this paper refers to no studies published after 1997 (except in cases where this paper originally referred to works in press or in preparation, or to published abstracts now rep¬ resenting journal articles). As a result, several studies using similar methods and/or data sets are not mentioned. The author regrets any confusion that might arise because of the misleading publication date. Peter J. Wagner, Department of Geology, Field Museum of Natural History, 1400 South Lake Shore Drive, Chicago, Illinois 60605-2496. E-mail address: pwagner@fmnh.org. coiled gastropods. These results are contrasted with the many previous phylogenetic estimates that gastropod systematists have presented. Although the primary goal of the study is to es¬ timate relationships among early gastropods, the study ad¬ dresses (by necessity) some larger phylogenetic issues. These topics include the relationship of early gastropods to other early Paleozoic molluscs, the relationships of the Paragas- tropoda (Linsley and Kier, 1984) to each other and to gastro¬ pods and other molluscs, and the relationships of the problem¬ atic bellerophonts to gastropods, other molluscs, and (to a much lesser extent) each other. This study differs from its predecessors in two ways. First, it includes only species that appeared from the Cambrian through the Silurian, whereas studies such as those cited above esti¬ mated gastropod phylogeny using species that appeared long after major taxa (i.e., orders and suborders) diverged (Yochel¬ son, 1984). Second, this study is essentially a species-level analysis, whereas previous studies typically used one or two exemplar species to represent each higher taxon. Erwin (1990b) rendered both strategies suspect, as a cladistic analysis of higher taxa that used late Paleozoic exemplar species sug¬ gested very different relationships than did an analysis of the same higher taxa that used early Paleozoic exemplars. The analysis presented herein avoids both problems by making no assumptions about the definitions or diagnoses of higher taxa. Acknowledgments A Smithsonian Predoctoral Fellowship during the summer of 1991, sponsored by D.H. Erwin and E.L. Yochelson, funded much of the research presented herein. Additional research funding was provided by an NSF doctoral dissertation im¬ provement grant, the Geological Society of America, Sigma Xi, the Hinds Fund from the Division of the Biological Sci¬ ences at the University of Chicago, National Aeronautics and Space Administration (U.S.A.) grant NAGW-1693 to J.J. Sep- koski, Jr., and NSF grant #EAR-84-177011 to D. Jablonski. I 1 2 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY thank the following people for allowing me access to the col¬ lections housed at their institutions (and also for outstanding hospitality): J. Cooper and N.J. Morris at The Natural History Museum, London; R. Owens at the Welsh Museum of National History in Wales; J. Bergstrom and V. Jaanuson at the Swedish Natural History Museum in Stockholm; R.J. Homy at the Nar- odni Museum, Prague; and the staff at the Okresni Museum, Rokycany, Czech Republic. The scanning equipment and gray¬ scale printers that made many of the figures possible were made available by D. Rowley and D. MacAyeal. I thank the following for extended to near-endless discussions about gas¬ tropod biology and evolution, and/or phylogenetic theory and methods in general, or for suggesting useful revisions to this manuscript: W. Allmon, J. Alroy, R. Bieler, D.H. Erwin, M. Foote, D. Jablonski, R. Homy, M. LaBarbera, D. Lindberg, R. Linsley, D. Miller, N.J. Morris, P.J. Morris, J.S. Peel, J.A. Schneider, J.J. Sepkoski, Jr., S. Suter, E.L. Yochelson, and the Field Museum systematics discussion group. By no means do I wish to imply that any of the above agree with all (or any) of my results, methods, and/or conclusions. “Archaeogastropods”—A Temporary Definition Workers have classified most early Paleozoic gastropod spe¬ cies within the order Archaeogastropoda; however, the defini¬ tions and diagnoses of the taxon has changed drastically in re¬ cent years, and there is little consensus on the taxon’s meaning or utility. Hickman’s (1988) monophyletic definition of extant taxa (Pleurotomarioidea + Fissurelloidea + Haliotoidea + Scis- surelloidea + Trochoidea; = the Vetigastropoda of Salvini-Pla- wen and Haszprunar, 1987) is not obviously applicable to early Paleozoic species. Neither is Haszprunar’s (1988) paraphyletic definition, which is based on the nervous system. Gastropod systematics is weighed down with excessive names and an un¬ stable taxonomy (Bieler, 1992), so retaining a definition of the “Archaeogastropoda” that is monophyletic through the Silurian (and hence used in quotes) is useful, if only for purposes of this discussion. For this paper, I define “archaeogastropods” as dex- trally coiled molluscs with an anal emargination (i.e., a sinus) and a peripheral band (see diagnosis below), plus all of their descendants through the Silurian. This definition differs from that of other paleontologists (e.g., Knight et al., 1960) and also from the paleontological definition of the Vetigastropoda given by Tracey et al. (1993) in that “archaeogastropods” include not only putative pleurotomarioids and trochoids, but also the earli¬ est species assigned to the Apogastropoda 2 and other orders. “Archaeogastropods” might be synonymous with a crown- group definition of gastropods (i.e., the last common ancestor 2 The Apogastropoda include fossil taxa, such as the Loxonematoidea, that Wenz (1938) and Knight et al. (1960) classified in the Caenogastropoda as well as modem caenogastropods, allogastropods, opisthobranchs, and pulmonates (Tracey et al., 1993). Therefore, I use the Apogastropoda as a replacement for the traditional paleontological definition of the Caenogastropoda. of all extant gastropods and all of its descendants), but this analysis cannot demonstrate that possibility. Review of Previous Phylogenetic Hypotheses A standard neontological depiction of gastropod phylogeny suggests that the planispiral Bellerophontina 3 were the earliest gastropods and that bellerophontinae were ancestral to anisos- trophic pleurotomarioids (e.g., Barnes, 1987; see Figure lA). Pleurotomarioids later produced other archaeogastropods, apo- gastropods, and the Patellogastropoda (e.g., Fretter and Gra¬ ham, 1962). The traditional paleontological depiction is simi¬ lar, essentially differing only by suggesting that a second extinct group of anisostrophic species (i.e., the Macluritoidea and Euomphaloidea) evolved from bellerophontinae (e.g., see Knight, 1952; Knight et al., 1960; Figure lB). Proposed alternatives exist to nearly every relationship shown in Figure lA,B. Some alternatives are only slightly dif¬ ferent. For example, Koken (1898, 1925; also N.J. Morris and Cleevely, 1981; P.J. Morris, 1991) thought that macluritoids and pleurotomarioids shared a helically coiled ancestor (Figure lc). Other proposals are radically different. A key controversy concerns the affinities of the Bellerophontina. Yochelson (1967, 1984) suggested that bellerophontinae evolved from (rather than giving rise to) pleurotomarioids (Figure ID). Ul¬ rich and Scofield (1897) considered bellerophontinae derived, but they thought that bellerophontinae and pleurotomarioids evolved independently from limpet gastropods (Figure lE). Haszprunar (1988) also considered limpet gastropods to be the ultimate gastropod ancestors of bellerophontinae, without com¬ menting on the relationship between bellerophontinae and heli- 3 In this discussion and elsewhere, the Bellerophontina refers to a diagnosed taxon, which presumably should represent a monophyletic or paraphyletic group. “Bellerophont” denotes a grade of bilaterally symmetrical, planispiral molluscs that might be polyphyletic. I use the former when discussing potential clades and the latter when discussing a morphologic type. FIGURE 1 (opposite).—Summary of previous phylogenetic hypotheses for the Gastropoda. “T” denotes the hypothesized onset of torsion, which is the chief synapomorphy of gastropods. A, Traditional neontological hypothesis. B, Tra¬ ditional paleontological hypothesis (e.g., Knight, 1952), with bellerophontinae as gastropods that give rise to macluritoids and pleurotomarioids separately, with pleurotomarioids giving rise to murchisonioids. (Murchisonioids later gave rise to apogastropods.) C, Macluritoids and pleurotomarioids sharing a pleurotomarioid-like ancestor (e.g., Koken, 1898, 1925; N.J. Morris and Cleevely, 1981; P.J. Morris, 1991). D, Bellerophontinae as derived pleuroto¬ marioids (Yochelson, 1967, 1984). E, Bellerophontinae and pleurotomarioids derived separately from early limpet gastropods (Ulrich and Scofield, 1897). F, Bellerophontinae as monoplacophorans with no close relationship to gastro¬ pods (e.g., Wenz, 1938; Runnegar and Pojeta, 1974; N.J. Morris, 1990). G, Bel¬ lerophontinae as monoplacophorans and gastropods and macluritoids (includ¬ ing onychochilids) evolving torsion independently from a pelagiellid ancestor (Runnegar, 1981). H, Bellerophontinae as a collection of monoplacophorans and primitive gastropods (e.g., Homy, 1965; Peel, 1991a). i, Macluritoids as paragastropods and not closely related to gastropods (Linsley and Kier, 1984). NUMBER 88 3 Patellogastropods AT S' A Monoplacophorans Bellerophonts —^ Pleurotomarioids —^ Other Archaeogastropods --<■ Caenogastropods + “Higher” Gastropods Macluritoids Euomphaloids B Monoplacophorans X^ Bellerophonts Murchisonioids —^ Loxonematoids /C Ni Pleurotomarioids —-* Trochoids N Subulitoids Other Caenogastropods Patellogastropods Euomphaloids —^ Macluritoids C Monoplacophorans X^ Bellerophonts —^ Raphistomatoids Loxonematoids Murchisonioids Pleurotomarioids ' Subulitoids Trochoids \i ^ jl m ._■.! j. ' Bellerophonts Caenogastropods D Monoplacophorans X^ Pleurotomarioids Macluritoids ■ Euomphaloids Trochoids Patellogastropods -^Patellogastropods v —Euomphaloids—* Macluritoids A / E Monoplacophorans —^ Limpet Urgastropod—^Raphistomatoids V \ -^Eotomarioids ^Bellerophonts Pleurotomarioids —*■ Trochoids Murchisonioids — * Loxonematoids —* Subulitoids -» F Monoplacophorans Bellerophonts Pleurotomarioids Murchisonioids ■ y Loxonematoids Caenogastropods f Patellogastropods 'other Archaeogastropods Macluritoids —^ Euomphaloids Subulitoids Bellerophonts G Monoplacophorans Onychochilids Pellagiellids Macluritoids Pleurotomarioids Other Gastropods H Monoplacophoran T V Gastropod V , V ^_ Bellerophonts ~~T Bellerophonts “f Pleurotomarioids—^ Other Gastropods , s* 1 Monoplacophorans Macluritoids Other Paragastropods Bellerophontoids -t Pleurotomarioids \ Other Gastropods 4 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY cally coiled gastropods. Conversely, segmented muscle scars in Devonian Cyrtonella Hall, 1879, led Wenz (1938) to conclude that the Bellerophontina were untorted molluscs. Runnegar and Pojeta (1974, 1985; also Runnegar, 1981, 1996; Dzik, 1981; N.J. Morris, 1990) advocated similar views and considered bel- lerophontinae and gastropods to be distant relatives (Figure 1f,G). Finally, some workers considered traditional definitions of the Bellerophontina to include both untorted and torted mol¬ luscs (e.g., Horny, 1965, 1991, 1992b; Harper and Rollins, 1982; Peel, 1991a,c, 1993; Wahlman, 1992). The last author suggested that bellerophont gastropods evolved from anisos- trophically coiled gastropods, whereas the first three authors considered some bellerophont gastropods ancestral to all other gastropods (Figure lH). The second model is not inconsistent with most traditional models (Figure lA,B), because traditional models often do not specify the gross morphology of the imme¬ diate ancestor of gastropods. The affinities of the Macluritoidea represent another source of contention. Traditionally, workers considered macluritoids to be one of the earliest offshoots within the gastropod clade (e.g., Knight, 1952; N.J. Morris and Cleevely, 1981). Consider¬ ations about molluscan functional biology led Linsley and Kier (1984) to suggest that macluritoids belonged to the Paragas- tropoda, a group of untorted molluscs (Figure ll). The Hyper- strophina (= Mimospirina of Dzik, 1982, plus the Omphalocir- ridae), a group of molluscs with highly ultra-dextral shells 4 , are pivotal in this controversy. Workers traditionally assumed that macluritoids evolved from the Onychochilidae (e.g., Knight, 1952; Linsley and Kier, 1984; Runnegar, 1981, 1996), an early family of the Hyperstrophina. Runnegar (1981) also considered onychochilids to include the ancestors of true macluritoids. In this scheme, torsion evolved independently in true gastropods and onychochilids, with untorted pelagiellids being the com¬ mon ancestor of both torted onychochilids and true gastropods. Functional analyses by P.J. Morris (1991), however, suggested that taxa such as Maclurites Le Sueur were torted, whereas hy- perstrophinae were untorted. Morris and others (e.g., Peel, 1991 a,b) considered hyperstrophinae and gastropods (includ¬ ing Maclurites ) as only distant relatives. Material Specimens I based the phylogenetic analysis on specimens housed in the type, biologic, and stratigraphic collections at the National Mu¬ seum of Natural History (Washington, D.C.), the Field Mu¬ seum of Natural History (Chicago), the Natural History Mu- “Ultradextral” shells often are described as “hyperstrophic.” Hyperstrophy and orthostrophy, however, refer to the orientation of the internal anatomy, whereas “dextral” and “ultra-dextral” refer to whether the shell coils “down” or “up” the coiling axis. Opercula indicate that ultra-dextral species, such as Ma¬ clurites, had orthostrophic organizations (Yochelson, 1990), so I use the latter set of terms instead. seum, London, the National Museum of Wales (Cardiff), the Natural History Museum of Sweden (Stockholm), the Narodni Museum (Prague), and the Okresni Museum (Rokycany, Czech Republic). These specimens included material from North America (including Alaska), China, Malaysia, the British Isles, Scandinavia, Estonia, and western and central Europe. I also examined material collected in Kentucky (U.S.A.) by the United States Geological Survey (Wagner, 1990). Published photographs were used when available. The analyses presented herein include 295 species. I ex¬ cluded most members of the Bellerophontinae, although sev¬ eral were used as outgroups (see below). I also excluded most species assigned to the Subulitidae, except for the earliest and also some later ones suspected of belonging to other clades (Er¬ win, 1992). Finally, I also omitted species assigned to the Platyceratidae. In this case, I attempted to include some of the earliest species, but 1 could not identify their relationships sat¬ isfactorily (see below). In addition to the deliberately excluded taxa, I also excluded at least 50 “archaeogastropod” species that likely represent valid taxa. Nearly all of those species are known from only one or a few localities, and I was not able to examine enough specimens to code them adequately; however, these species also appear to have been short-lived and apomor- phic. As I could include close relatives (which often were po¬ tential ancestors of the short-lived species), the exclusion of the short-lived species should not interfere with the stated goal of this paper, i.e., identifying the relationships among major gas¬ tropod taxa in the early Paleozoic. Notably, this analysis ex¬ cluded no species known from five or more localities. I examined multiple specimens of most species while coding the character states, which allowed me to evaluate ranges of in- traspecific variation and also to observe ontogenetic variation within species. I granted no special status to type material be¬ yond determining species assignments. This is important be¬ cause much of the material that I examined has not been de¬ scribed, especially from the Early Ordovician. Nearly all of the specimens examined fit within the diagnoses of previously de¬ scribed species. A question mark precedes a species name throughout the paper if I was unable to examine the type speci¬ men (or if the type was so poor as to be uninformative). Biogeography of Analyzed Species The biogeographic affinities of the examined species are somewhat complex, largely because of geographic evolution from the latest Cambrian through the Silurian. “Archaeogastro- pods” from the latest Cambrian through the early Arenig (Early Ordovician) appear to have been restricted to the Laurentian fauna (i.e., eastern North America and Scotland). Even after the Early Ordovician, the tropical Laurentian fauna appears to have maintained the highest “archaeogastropod” diversity of the early Paleozoic realms. By the late Early Ordovician (i.e., middle Arenig), “archaeogastropods” also existed in the Baltic, Toquima-Table Head, Celtic, and Gondwanan faunas (see Neuman and Bruton, 1989; Cocks and Fortey, 1990, for general NUMBER 88 5 descriptions of those faunas). The Toquima-Table Head fauna represents a tropical to equatorial assemblage, whereas the Bal¬ tic fauna represents a temperate fauna, and the Celtic and Gondwanan faunas represent a temperate to near-polar assem¬ blage. The Toquima-Table Head and Celtic faunas (i.e., gastro¬ pod species from western North America, the northern east coast of North America, Wales, and parts of Norway) are some¬ what problematic. Some workers recognize both faunas as dis¬ tinct provinces that were unique to the late Early to Middle Or¬ dovician (i.e., Neuman and Bruton, 1989; Neuman and Harper, 1992). Others consider both faunas to be mixtures of Lauren- tian and Baltic faunas (McKerrow and Cocks, 1986; Cocks and McKerrow, 1993). Gastropods from the Celtic fauna are not well known, but the few described species (e.g., Neuman, 1964) are also known from the Toquima-Table Head faunas (pers. obs.). Although these gastropods do have affinities with both Laurentian and Baltic species, they appear to represent a separate fauna. Ordovician gastropods from the temperate-to- polar Gondwanan realm appear to have been rare, and I exam¬ ined species only from the Middle Ordovician of western and central Europe. Early to Middle Ordovician gastropods also have been reported from the Gondwanan faunas of South America (e.g., Beresi and Rigby, 1993), but I was not able to examine any specimens. Paleogeographic reconstructions sug¬ gest that those South American faunas should have been equa¬ torial, and general published descriptions suggest that the gas¬ tropods belonged to the tropical Toquima-Table Head fauna. “Archaeogastropod” biogeography simplified in the Late Or¬ dovician, which witnessed an increasing homogeneity among realms, not only for gastropods, but also for bryozoans (Anstey, 1986), trilobites (Cocks and Fortey, 1990), and brachiopods (Cocks and Ruang, 1988; Cocks and Fortey, 1990). Notably, all of the faunas are thought to have been closer to the equator dur¬ ing the Late Ordovician than they had been in the Early and Middle Ordovician (Scotese, 1989). The Early Silurian (i.e., Llandovery-Wenlock) shows still greater homogeneity, as “ar- chaeogastropods” appear to have represented a single equato¬ rial fauna. Some differentiation is noticeable by the Late Sil¬ urian (Ludlow-Pridoli), with the temperate Gondwanan fauna distinct from the tropical Laurentian and Baltic faunas. Cladistic Characters Homology Versus Architecture It is important to discuss codings and a priori hypotheses of homology when conducting cladistic or phenetic analyses. Un¬ fortunately, most of the terms used to describe gastropod shells (or portions of those shells) refer to architectural features that might or might not be present depending on the interactions of different character suites (e.g., coiling parameters and aperture shape). For example, terms such as “columella” or “umbilical carina” are not used to label homologous regions of the shell on species with very different gross morphologies. To avoid con¬ fusion, I avoid these terms and instead use slightly less com¬ mon terms. For example, the inner margin forms a columella on a “typical” gastropod shell (e.g., Lophospira perangulata (Hall, 1847)) and thus usually is labeled either the columella or the columellar lip (Cox, 1960; e.g., Figure 2A). On species with sufficiently low shell curvature, such as Clathrospira el- liptica (Hisinger, 1829) (Figure 2b) or Spiroraphe bohemica Barrande in Pemer, 1907 (Figure 2c), the “columellar lip” fails to form a columella unless it is extremely thick. A columella might still be formed by the parietal inductura (i.e., a funicle) on species with the same basic shell geometry (e.g., Siluripho- rus gotlandicus (Lindstrom, 1884)) (Figure 2d). Finally, the homologous region on nearly planispiral taxa, such as Barne- sella llecanospiroides Bridge and Cloud, 1947 (Figure 2e) or Palliseria robusta Wilson, 1924 (Figure 2f), forms the base of the shell rather than a columella (see, e.g.. Figure 2f). Given the inconsistent relationship between this feature and the col¬ umella (and the fact that the columella is an architectural fea¬ ture rather than a true homology), I refer to that portion of the aperture as the inner margin. A columella at the base of the inner margin typically encir¬ cles the umbilicus and hence is labeled a basal carina. On nearly planispiral forms, however, the carina is at the periphery of the shell base. Accordingly, I refer to the feature as a basal carina. In addition, the “upper” and “lower” ramps of normally coiled species (e.g., Figure 2a-d) are the “right” and “left” ramps of nearly planispiral species (Figure 2e,F). As the latter terminol¬ ogy also refers to the post-torsional orientation of internal or¬ gans, I use right ramp throughout this paper. Batten (1989) la¬ beled the left ramp the alveozone, a term that I use herein. The Paucity of Shell Characters Revisited Appendix 1 gives the characters and character states used in this study. Appendix 2 gives the data matrix. I discuss addi¬ tional data relevant to both appendices below. I used 143 char¬ acters encompassing 352 character-states for this study (Ap¬ pendix 1; note that I count continuous characters as only one state). There are four reasons why I used so many characters. First, the species analyzed herein encompass sufficient mor¬ phological disparity to be classified in multiple orders. One usually can describe any specimen with fewer than 50 charac¬ ters, but one needs many more characters to describe the entire spectrum of gastropod shell morphologies. The second reason for the high number of characters is that this is a species-level analysis. Supraspecific phylogenetic analyses of gastropods average less than one shell character per operational taxonomic unit (OTU), whereas species-level anal¬ yses average more than one shell character per OTU (Table 1). This likely is because many shell characters vary within clades as well as among clades, so if a study uses only a few exemplar species to represent higher taxa, then many shell characters be¬ come uninformative. Thus, many shell characters that are use¬ ful in phylogenetic analyses of closely related species are not useful in phylogenetic analyses of distantly related ones. 6 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY FIGURE 2. —Some basic terms and characters used in the analyses. AC = Carina at base of Alveozone; AL = Alveozone (= Post-Torsional Left Ramp; Batten, 1989); B = Base of Inner Margin; BC = Basal Carina (typically an umbilical carina); IM = Inner Margin; PB = Peripheral Band (almost always located at the sinus apex); RR = Right Ramp; SA = Sinus Apex (for specimens without a peripheral band). A, Lophospiraperangulata (Hall); IM thick, nearly straight, trends nearly parallel to coiling, reflects around coiling axis and forms columella; BC absent; AC present, sharp; left and right ramps symmetrical in shape (concave) and length; PB trilineate (i.e., bearing both peripheral lira and a medial lirum), bisects left and right ramps and oriented approximately 30° adapically from perpendicular to IM. B, Clathrospira elliptica (Hisinger) IM little thicker than rest of shell, curved and nearly parallel to coiling axis; BC and AC absent; AL and RR convex, but AL shorter than RR; PB bilineate (i.e., peripheral lira present), bisecting AL and RR and oriented nearly perpendicular to IM. C, Spiroraphe bohemica Barrande in Pemer: IM curved, oriented nearly 30° off parallel to coiling axis; BC and AC absent; AL and RR convex, but RR more convex and longer than AL; PB bilineate, falling partially on RR and oriented approxmiately 10° abapically from perpendicular to IM. D, Siluriphorus gotlandicus (Lindstrom): IM strongly curved, thicked above base, approximately 15° off perpendicular to coiling axis; BC present, thin and weak; AC present, “squared” and strong; AL and RR convex, with RR very short relative to AL; PB absent, with SA near suture. E, Barnesella llecanospiroides Bridge and Cloud: IM runs nearly perpendicular to coiling axis (forming flat base of shell); BC thick but weak; AC absent; AL and RR flat and long, symmetric in shape and length; PB monolineate, sharp, weak. F, Palliseria robusta Wilson: IM thick, curved and oriented at very high angle (-120°) relative to coiling axis (forming curved base of the aperture); BC dull thickening; AC absent; AL and RR equal in length, but with convex AL and flat or concave RR. The third reason for the large number of characters is that this analysis used many traits that are inapplicable to extant species. Characters describing sinuses, peripheral bands, slits, and differential shell asymmetry are not relevant to studies of most extant taxa, but they are very important to the study of early Paleozoic species (Table 2). For example, only a few types of peripheral bands exist on a handful of extant species; however, most early Paleozoic species possessed peripheral NUMBER 88 7 Table 1.—Operational taxonomic units (OTUs) versus shell characters for some previous phylogenetic analyses of gastropods. Included are the number and taxonomic level of taxa and the number of shell characters that they utilized. Only characters that could be utilized in this study were counted, so some characters, such as shell min¬ eralogy, were excluded. “No. of OTUs” is the number of taxa that were analyzed, including outgroup taxa. Study Taxon Taxic level of OTUs No. of OTUs No. of shell characters Ponder and Lindberg (1996) Gastropoda “superfamily” 22 3 Haszprunar (1988) Gastropoda “superfamily” 15 3 Houbrick(1988) Cerithioidea superfamily 15 15 Ponder and Lindberg (1997) Gastropoda “family” 25 5 Davis et al. (1985) Rissoidea subfamily 8 1 Hickman and McLean (1990) Trochoidea subfamily/tribe 29 26 Hickman (1996) Trochoidea subfamily/tribe 20 20 Hickman (1996) Turbinidae subfamily/tribe 9 13 Davis and Pons da Silva (1984) Hydrobiidae genus 8 4 Ponder (1984) Iravadiidae genus 14 9 Bieler (1988) Architectonicidae genus 12 12 Reid (1989) Littorinidae genus 36 2 Jung(1992) Planorbidae genus 9 27 Kool (1993b) Ocenebrinae genus 5 3 Kool (1993a) Rapaninae genus 24 4 Houbrick (1984)/ Cerithidea subgenus 4 9 Erwin (1988) Glyptospira species 8 21 Michaux (1989) Ancillinae species 32 32 Wagner (1995a) Lophospiridae species 42 79 This study “Archaeogastropods” species 295 352 Table 2. —Importance of “archaeogastropod” shell characters in this analysis versus their importance in other analyses. Columns give the number of character states used in each study for the particular trait. Asterisks denote examples discussed in the text. PB denotes characters describing peripheral bands, and InAn denotes characters describing apertural inclination. Coiling denotes coiling and/or growth parameters. Study Characters Slit Sinus PB Asymmetry InAn Coiling Ornament Ponder and Lindberg (1996) 0 0 0 0* 0 0 0 Haszprunar (1988) 1* 0 0 0 0 2 0 Houbrick (1988) 0 0 0 0 0 1 1 Ponder and Lindberg (1997) 0 0 0 1 0 0 0 Davis et al., 1985 0 0 0 0 0 1 0 Hickman and McLean (1990) 1 0 0 0* 1 2 4 Hickman (1996) 1 0 0* 0 1 1 1 Davis and Pons da Silva (1984) 0 0 0 0 0 1 1 Ponder (1984) 0 0 0 0 3 0 4 Bieler (1988) 0 0 0 0 0 1 8 Reid (1989) 0 0 0 0 0 1 1 Kool (1993a) 0 0 0 0 0 1 0 Kool (1993b) 0 0 0 0 0 0 2 Houbrick (1984) 0 0 0 0 0 0 2 Erwin (1988) 0 0 3 0 0 1 11 Jung (1992) 0 0 0 0 0 5 4 Michaux (1989) 0 0 0 0 0 6 1 Wagner (1995a) 1 3 5 0 4 5 4 This study 9 21 58 15 5 11 16 bands, and several different types existed. The transition from symmetric morphologies to asymmetric ones introduces addi¬ tional character states, as both the left and right sides of several features must be coded independently. Some additional impli¬ cations of this pattern are discussed in detail below. Finally, this study used finer divisions of shell characters than employed by previous workers. Gastropods can produce very similar shell shapes using different combinations of growth parameters, aperture shapes and orientations, and shell thicknesses. Character complexes often have been treated as single characters, but I divided these into several characters and multiple states. Hickman and McLean’s (1990; see also Hick¬ man, 1996) cladistic analysis of the Trochoidea used tangential and radial apertures (i.e., inclined versus noninclined) as two states of one character. Early Paleozoic gastropods produced inclined apertures in many ways (Table 2; Appendix 1, charac- 8 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY ters 109-119). Figure 3 illustrates some examples. The aper¬ tures of some species, such as Siluriphorus gotlandicus, form a plane that is inclined relative to the coiling axis (Figure 3a). For other species, different parts of the aperture are inclined to different degrees. On species such as the Middle Ordovician Clathrospira conica Ulrich and Scofield, 1897, the left 5 side of the aperture is much more inclined than the right side (Figure 3b). The opposite condition exists on Pleurorima migrans (Barrande in Pemer, 1907), where the right side provides most of the inclination (Figure 3c). The base of the aperture also can deviate from radial by pro¬ jecting either anteriorly or posteriorly. An example of the former is the Middle Ordovician species Helicotoma tennes- seensis Ulrich and Scofield, 1897 (Figure 3d). The Early Or¬ dovician species Pararaphistoma qualteriata (Schlotheim, 1820) is an example of the latter (Figure 3e). Note that the pos¬ terior projection is not the same as a basal excavation (where the base has a sigmoidal or U-shape), which is coded as a sepa¬ rate character. Posterior projection of the base is especially common in species with very low spire heights (especially planispiral species) and might serve to alter the center of grav- 5 “Left” and “right” here and elsewhere refer to the post-torsional left and right. For species with selenizones, the right corresponds to the area between the selenizone and the coiling axis (i.e., usually above the “upper whorl”). ity within a nearly planispiral shell in a manner that is analo¬ gous to the standard inclination of the aperture on a normally coiled shell (e.g., Linsley, 1977). Anterior projection of the base occurs in both low-spired and high-spired species and also might serve to enhance apertural inclination. Aperture shape is another trait that previous workers have used as a single character (e.g., Houbrick, 1984; Hickman and McLean, 1990). Different parts of “archaeogastropod” aper¬ tures (or, more appropriately, different portions of the outer and inner whorl faces) can have different shapes (e.g., convex, flat, concave) on the left and right side of the aperture owing to the decoupling of left-right homologies with increasing asymme¬ try. Accordingly, I coded the shape of the left and right sides of the aperture separately rather than trying to code a whole shape for the aperture. An additional important point to this breakdown of character complexes concerns the functional biology of gastropod shells. Features such as spire height, apertural inclination, aperture shape, and columellar type strongly affect how easily snails can balance and move their shells (e.g., Linsley, 1977, 1978; Mc¬ Nair et al., 1981; Signor, 1982). Thus, convergent evolution of similar functional complexes among different clades is a major concern. The characters comprising functional complexes are used herein, however, rather than the complexes themselves. Because different combinations of characters will yield very ix-x x y y i rzzzzzzzzzi '*-■ L .. . .7^3 Inclination of whole aperture Inclination of right side Inclination of left side Projection of the base Plane radial to the coiling axis FIGURE 3. —Different ways in which early Paleozoic gastropods produced tangential apertures. A, Inclination of the whole aperture. B, Inclination provided primarily by the left side of the aperture only. Note that the lower part of the aperture has a stronger inclination than the upper part. C, Inclination provided primarily by the right side of the aperture. D, Anterior projection of the aperture base. E, Posterior projection of the aperture base. NUMBER 88 9 similar character complexes (e.g., gross aperture shapes, spire heights, overall apertural inclinations, and columellas), two distantly related but grossly similar morphologies that func¬ tional biologists might consider to be the “same” probably will have very different character codings. Similarly, two closely re¬ lated but grossly different morphologies might have very simi¬ lar character codings. This does not mean that the coding scheme used herein will identify all functional convergences. Functional convergence between closely related clades or in¬ volving simplification of the shell (and necessarily reducing applicable characters) will confound any coding scheme. The breakdown of the characters (coupled with the use of strati¬ graphic data as tests of parsimony estimates), however, should increase the accuracy of the phylogenetic analyses. Sinuses, Slits, Selenizones, And Peripheral Bands Among modem gastropods, the sinus (a broad, acute emar- gination culminating at the presumed site of the exhalent cur¬ rent; Figure 4a) exists only on some vetigastropods (i.e., pleu- rotomarioids and some scissurelloids) and on some basal caenogastropods (e.g., turritellids), whereas the slit (a thin lin¬ ear cleft, also culminating at the presumed exhalent current; Figure 4b) occurs on other vetigastropods and on species in some other clades, such as the Architectonicidae. Neontolo- gists generally have ignored the sinus and have considered the slit to be a synapomorphy of the earliest gastropods (e.g., Hasz- prunar, 1988). Paleontologists noted that the sinus evolved prior to the slit, but most considered the two features to be ho¬ mologous (e.g., Knight, 1952). Horny (1962), however, sug¬ gested that the two features represented independent homo- logues, at least in the case of bellerophonts. This study supports Horny’s idea. Although much attention has been paid to the slit, the sinus is far more informative phylogenetically, as si¬ nuses provide multiple characters that are distinguishable among nearly all “archaeogastropod” species (Table 2; see also Appendix 1, characters 1-11). Slits provide far fewer traits (Ta¬ ble 2; see also Appendix 1, characters 34-36), but as slits are much rarer than sinuses, they tend to be highly informative where they exist (see below). An associated issue herein is the relationship between the slit and the peripheral band (i.e., the “slit-band” of 19th and early 20th century literature). Knight (1934) relabeled the feature the “selenizone” and defined it as a structure generated by a slit. This assumed that the band was simply a distortion produced by a linear cleft in the shell. Several previous workers (e.g., Lindstrom, 1884; Ulrich and Scofield, 1897; Donald, 1902, 1906) had noted that slit-bands predate slits in the fossil record. Although slits were not common before the Devonian and were very rare during the Ordovician, the ubiquitous peripheral band appeared by the Late Cambrian. Knight (1941, 1952) later rec¬ ognized this and considered the band to be homologous on both slit-bearing and slitless species. Instead of abandoning the pre¬ vious morphogenetic hypothesis, Knight (1952) abridged it by inferring an unseen notch in the aperture that generated the pe¬ ripheral band for slitless species. Knight (and subsequent work¬ ers) used the term “pseudoselenizone” to describe such periph¬ eral bands. Despite frequent allusions to a notch, this feature has never been observed and its existence has been inferred solely on the assumption that a peripheral band is an artifact of a cleft in the aperture. Knight and others apparently did not consider the possibility that peripheral bands had no morphogenetic relation to slits or notches. One line of evidence suggesting that this is the case is that the pseudoselenizone of species with no such slit (e.g., “ Longstaffia ” “laquetta ” (Lindstrom, 1884) (Figure 4a) typi¬ cally differs little from selenizones of closely related species with slits (e.g., “ Seelya ” lloydi (Sowerby in Murchison, 1839) (Figure 4b). A related point is that when the slit is a variable feature on individual specimens, the selenizone is unaffected. The growth lines of some species (e.g., Clathrospira subconica (Hall, 1847) (Figure 4C) suggest that the specimens had incon¬ sistent slits, i.e., the presence or absence of the slit varied. The growth lines within the peripheral band (i.e., lunulae) corre¬ spond with the growth lines outside the band, which indicates that the animal did not have a slit (or a “notch”) when that part of the shell was secreted. There are fewer lunulae than growth lines, however, and the initial growth did not include shell dep¬ osition within the selenizone. This suggests that deposition within the peripheral band was halted for a period, resulting in the production (and subsequent lengthening of) a slit. The short slit later is filled, leaving the shell temporarily slitless. Despite the inconsistent nature of the slit, the peripheral band of C. subconica remains the same, which indicates that the slit is not responsible for the structure. Species such as Pararaphistoma qualteriata provide an ex¬ ample similar to that of Clathrospira Ulrich and Scofield (Fig¬ ure 4f). Pararaphistoma Vostokova species and their relatives lack slits on their juvenile whorls, and the presence of a slit sometimes is erratic on the adult whorls. The nature of the sele¬ nizone, however, does not vary over ontogeny or change with the production of a slit. The apex of a sharp (e.g., V-shaped) sinus, such as seen on Clathrospira, Pararaphistoma, and most other early Paleozoic taxa, might act as Knight’s notch. Several species with very sharp sinuses (e.g., the Silurian Sinuspira tenera Barrande in Pemer, 1907), however, lose the peripheral band over ontogeny without any corresponding ontogenetic changes in sinus mor¬ phology. If the band were purely an artifact of sinus morphol¬ ogy, this would not be possible. If a slit generated the peripheral band, then we would expect the position of the two features to coincide on the shell. This usually is true, as the peripheral band usually borders the slit (e.g., Figure 4b-d), especially on species with bilineate bands (i.e., two carinae). Species such as Pararaphistoma qualteri¬ ata, however, have slits that are wider than the peripheral band and are positioned somewhat differently on the aperture (Fig¬ ure 4f). Another extreme is shown by Oehlertia scutulata 10 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY FIGURE 4.—Sinuses, slits, and periph¬ eral bands. A, Gastropod with a sinus (the curved emargination), a periph¬ eral band (the paired threads at the apex of the sinus), but no slit. B, Gas¬ tropod with a sinus, peripheral band, and periodic production of a slit in the middle of the peripheral band. C, D, Gastropods with sinus, peripheral bands, and slits. E, Gastropod with a narrow, shallow sinus, and a broad peripheral band lunulae angling straight into a very narrow slit that is bordered by two sharp threads. F, Gas¬ tropod with a sharp, single carina for a peripheral band, and a slit that begins above the base of the peripheral band and terminates above the top of the peripheral band. In this case, separate views of the right (upper) and left (lower) ramps are shown, as well as a profile with arrows denoting the posi¬ tions of the ramps. Note that the slit is not always produced, as some growth lines give way immediately to sigmoi¬ dal-shaped lunulae. A and C-F taken from Lindstrom (1884); B taken from Ulrich and Scofield (1897). (Lindstrom, 1884), which has a very thin slit (shown by the in¬ ner pair of sharp lira in Figure 4 e) that is much narrower than the peripheral band (shown by the outer pair of sharp lira in Figure 4 e). Note that the outer pair appear to be homologous with the peripheral band of other gastropods, whereas the inner pair represent a feature unique to Oehlertia and related taxa. In summary, (1) peripheral bands evolved long before slits did, (2) peripheral bands on early Paleozoic gastropods cannot have been created by slits or notches, and (3) there is no evi¬ dence that “notches” of any sort ever existed. The definition of a “pseudoselenizone” is rendered logically complex and cer¬ tainly must be discontinued. Knight’s definition of the seleni- zone also is logically complex (as the peripheral bands do not coincide with slits in either ontogeny or phylogeny). Knight (1952) and previous workers did, however, recognize the ho¬ mology of selenizones and pseudoselenizones. Although a cor- NUMBER 88 11 rected definition of “selenizone” could be used, the term has become too strongly identified as a by-product of a slit rather than an independent anatomical feature. Accordingly, the use of selenizone also should be abandoned, at least in reference to the peripheral band of most Paleozoic gastropods. The term “fasciole” suffers from the same problems. In the absence of a more appropriate term, I will use the cumbersome “peripheral band” to label the band at the apex of the sinus (and presum¬ ably denoting the location of the anus). In addition to clarifying terminology, the importance of this lengthy discussion is that it indicates the need for a slightly broader coding scheme than is implied by the earlier literature. The morphogenetic scheme of Knight (1952) implies that the peripheral band is an architectural artifact similar to a col¬ umella, which therefore would require coding only the pres¬ ence/absence of a notch/slit (e.g., Hickman, 1996). Instead, the presence or absence of a slit represents one character and the presence/absence of a peripheral band represents another char¬ acter. As noted above, the presence/absence of a sinus is a com¬ pletely independent third character. Shell Mineralogy and Protoconchs Many workers consider shell mineralogy and protoconch morphology to be phylogenetically informative (e.g., Batten, 1972, 1984; Bandel, 1988, 1991; Ponder, 1990a, 1990b); how¬ ever, I did not use these characters in this analysis. Shell miner¬ alogy probably varied widely among “archaeogastropods,” but it rarely is possible to identify the exact types of shell mineral¬ ogy. Taphonomic characteristics reveal which species had at least partially calcitic shells, but recrystalization usually ob¬ scures the relative amount of calcite or its exact nature. Most of the specimens that I examined were silicified. Silicification can reveal the number of mineral layers and their relative thick¬ nesses, but it leaves no other evidence about those layers (see Carson, 1991). When possible, I do discuss some basic aspects of mineralogy (i.e., aragonitic versus calcitic shells), especially if they support or contradict the results presented herein. I omitted protoconchs for a different reason. A distinct proto¬ conch morphology typifies modem species, but not early Pale¬ ozoic ones. I examined many extremely well-preserved speci¬ mens representing a number of different taxa, but the boundaries between protoconch and teleoconch usually were vague at best (some exceptions are discussed below). I also ex¬ amined microfossils from beds rich in gastropods. Sinus and peripheral band morphologies usually were not observable on microfossils, but the basic profiles of the shells matched those of macrofossils known from the same beds. Dzik (1978) fig¬ ured protoconchs of two early Paleozoic “archaeogastropod” species, which also possessed adult profiles but lacked clearly defined sinuses and peripheral bands. Many species show distinct ontogenetic changes. These dif¬ fer from the abrupt transitions between protoconchs and teleo- conchs because the changes are gradual. For example, species classified as Macluritella Kirk, Teiichispira Yochelson and Jones, and Malayaspira Kobayashi have juvenile shells that are similar in overall morphology to Prohelicotoma Flower (Figure 5a). Similarly, species of Pararaphistoma Vostokova and Cli- macoraphistoma Vostokova have juvenile shells similar to that of Lesueurilla Koken (Figure 5b). Therefore, I did not code ju¬ venile shell types as separate characters. Instead, I coded the types of ontogenetic changes (e.g., counter-clockwise rotation of the aperture and differential expansion of the left side in ma- cluritids, or clockwise rotation of the aperture and increased translation in raphistomatids) as present or absent. I then coded the traits that changed during ontogeny as polymorphic, with all the states produced by an ontogenetic trajectory coded as present. Phylogenetic Analysis Character Deweighting 1—Balancing Continuous Characters There are two standard justifications for character weight¬ ing: accounting for differential homoplasy among characters (e.g., Farris, 1969; Goloboff, 1993) and mitigating the effects of ordered characters (Thiele and Ladiges, 1988; Chappill, 1989; Hauser and Presch, 1991; Skelton and McHenry, 1992). I did not weight or reweight characters because of homoplasy. I did deweight ordered characters, however, to accommodate continuous characters. In all cases these represented continu¬ ous morphologic features, such as shell growth parameters or apertural inclination. An ordered trait with 10 characters will result in a nine-step difference between species coded as “1” and those coded as “10”; however, the maximum difference A B FIGURE 5.— Ontogenetic changes in shell morphology. A, Morphology of adult Prohelicotoma and juvenile Macluritella or Teiichispira. B, Morphology of adult Teiichispira, with negative translation, the left side of the aperture expanded, and the entire aperture rotated counter-clockwise. C, Morphology typical of adult Lesueurilla and juvenile Climacoraphistoma or Pararaphis¬ toma. D, Morphology of adult Climacoraphistoma, with higher translation and the aperture rotated clockwise. 12 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY between a presence/absence character is only one step (i.e., “0” to “1”). The concern herein is that ordered characters with many states will have an excessively strong effect on a cladis- tic analysis. The “over-weighting” effects of ordered charac¬ ters noted by Hauser and Presch (1991) is of particular con¬ cern herein, as malacologists have documented the potential plasticity of shell characters within species (e.g., Kemp and Bertness, 1984; Palmer, 1985; Boulding and Hay, 1993). Un¬ weighted continuous characters then might separate close rela¬ tives by several steps due to evolutionarily minor differences. Therefore, I deweighted continuous characters as follows: continuous character weight =--- no. of character states -1 (see Thiele and Ladiges, 1988). One presenee/absenee synapo- morphy always contributes at least as much (and usually more) to the parsimony analysis as does a continuous synapo- morphy. Deweighted continuous characters are denoted with a “C” in Appendix 1. Farris’ (1990) contentions against the deweighting of con¬ tinuous characters assume that discrete characters are recog¬ nizable. This assumption is unsound for the continuous char¬ acters used herein. Techniques used to categorize quantitative characters (e.g., Thorpe, 1984; Archie, 1985; Goldman, 1988) all assume that intraspecific variation is roughly equal for all species, but intraspecific variation can vary among different morphotypes (Schindel, 1990) and at different times in a clade’s history (Hughes, 1991). The latter problem appears to have been the case among early gastropods (Wagner, 1995b, 1996). These factors also discouraged the use of simple maxi¬ mum likelihood methods for quantitative characters (e.g., Felsenstein, 1981, 1988); therefore, I used segment coding (Chappill, 1989), which simply divides continuous characters into equal distributions (e.g., “narrow” <0.1, “medium” > 0.1 and < 0.2, etc.). This defines no discrete states, so the charac¬ ters are meaningful only if ordered. Segment coding links ad¬ jacent characters so that “narrowness” is a synapomorphy of a “narrow” and “very narrow” species relative to a “wide” out¬ group. This is an inexact method of describing character states, but it makes the fullest use of the available data and thus should be used in analyses such as this one. Several authors have criticized segment coding (e.g., Pimen¬ tel and Riggins, 1987; Cranston and Humphries, 1988) on the grounds that because one cannot define homologies in a repeat- able manner for continuous characters, such features lack phy¬ logenetic information. These arguments assume that meaning¬ ful evolution always proceeds like a cladistic character state optimization, with states clicking on and off without intermedi¬ ate forms (see Janvier, 1984; Gayon, 1990). This is a conten¬ tious view of morphologic evolution that is better examined within the contexts of phylogenetic estimates than assumed when conducting phylogenetic analyses. Character Deweighting 2—Asymmetry and Changing Homologies I employed character deweighting for another situation that previous workers have not discussed. Evolutionary events can couple or decouple homologies (e.g., Schaeffer and Lauder, 1986; Wake and Roth, 1989; Atchley and Hall, 1991), result¬ ing in multiple characters becoming one or one character be¬ coming several. In such cases, some species possess multiple characters for a morphologic structure whereas other species have only one. Among gastropods, the loss of bilateral sym¬ metry represents an example of the decoupling of homologies. This has had appreciable effects on the internal anatomy. Among gastropods, left and right homologues often serve dif¬ ferent functions (or the right organ is absent); these organs are paired in other molluscs. Indeed, trends toward asymmetrical conditions have been so pervasive that parsimony optimiza¬ tion finds bilaterally symmetrical conditions to be derived within gastropods (e.g., Ponder and Lindberg, 1996). This pat¬ tern also is reflected in shell morphology. For example, the left and right sides of the sinus and aperture are symmetrical on most early appearing species, but asymmetrical morphologies appeared by the Early Ordovician. This leads to a coding para¬ dox. The left and right can change independently on asymmet¬ rical species and, thus, can represent separate character states. If one codes the left and right sides as separate traits, however, then symmetric species have many of the same characters coded twice. Ideally, one would code features that vary in symmetry in the following manner. First, one would distinguish “symmet¬ ric” versus “asymmetric” as a presence/absence character. There are two mutually exclusive types of asymmetries: greater development of the right side and greater development of the left. Without a priori evidence that one type cannot evolve from the other, one should code these as unordered character states. Among species with the same type of asymmetry, any dif¬ ference on the left or right side should be coded as one step (or ln th of a step for continuous characters with n+1 states). When describing the difference between asymmetric and sym¬ metric species, the coding must avoid assuming how asymme¬ try evolved. An aperture with a more pronounced left side can be produce by enlarging the left side or contracting the right side. Thus, one should code such asymmetric species so that they are equally close to symmetric species with identical left or right sides. For example, consider a species on which the right side of the sinus retreated at 30° whereas the left side re¬ treated at 50°. That species should be considered equally sim¬ ilar to species with symmetric sinuses retreating at either 30° or 50°. The 30°:50° species should be coded as one step away from either the 30°:30° or 50°:50° species (i.e., the absence versus the presence of symmetry), with the 20° difference on the left or right side considered to be produced by the onset of asymmetry. NUMBER 88 13 If a symmetrical species differs from both the left and right sides of an asymmetrical species, then one should code species so that there is a difference of one step (symmetry versus asymmetry) plus the minimum number of steps needed to make either the left or right side of the symmetrical species identical to the left or right side of the asymmetrical one. Con¬ tinuing the example started above, the 30°:50° species would differ from a 40°:40° species by one step (symmetry to asym¬ metry) plus l/n th steps (either 30° to 40° on the left side or 50° to 40° on the right side, with l/n th being one over the number of character states +1; see the discussion of continuous charac¬ ters, above). The 30°:50° species would differ from a 20°:20° species by one step (symmetry to asymmetrically deep left side) plus two more steps (i.e., 20° to 40° on the left side). The major difference between species for the other side is attrib¬ uted to the change in symmetry. This scheme is the most parsi¬ monious possible because it assumes the minimum differences between species. Step matrices (Swofford and Olsen, 1990; Maddison, 1993) permit the character coding scheme described above. Unfortu¬ nately, using step matrices slows down computer analyses so much that I could not analyze even small data sets; therefore, I could not use step matrices in this analysis. Instead, I coded the symmetry as “present,” “absent (left side greater)” or “absent (right side greater).” I then weighted the left and right sides of potentially asymmetric characters as separate characters. The deweighting means that 30°:30° species and 50°:50° species differ by 2n steps (30° to 50°) instead of 4n (30° to 50° on the right side and 30° to 50° on the left side). The 30°:50° species, however, differs from either species by 1 step (symmetry to asymmetry) plus 2n (the difference on the right or left side). There are two disadvantages to this scheme. First, the number of differences between symmetrical and asymmetrical species is slightly greater than it should be (i.e., l+2n steps instead of 1), as differences in both the left and right sides are tallied. More importantly, this scheme means that the cladistic analysis can imply chimeras. Figure 6a gives an example where parsi¬ mony will predict an ancestor with symmetrical sinus but dif¬ ferent left and right sides. In these cases, I had to reoptimize the character states so that no chimeras existed (e.g., Figure 6b). I then kept the shortest trees without chimeras. Until cladistic programs can implement step matrices efficiently, the imper¬ fect approach used herein represents the best solution for this type of problem. Character Analyses I analyzed the data using PAUP 3.1 (Swofford, 1993). I used heuristic searches, with multiple replications and random se¬ quence addition of species employed to account for islands of similar trees (see D.R. Maddison, 1991). This does not guaran¬ tee finding the shortest trees, especially for a matrix of this size; therefore, I reanalyzed smaller portions of the data, with the initial results providing estimates of subclade membership and appropriate outgroups. Ultimately, these analyses were re¬ duced to four clades, which are referred to below as the “helic- otomatids,” “euomphalinae” (minus “helicotomatids”), “eoto- marioids,’' and "murchisoniinae” (minus “eotomarioids”). A disadvantage of this strategy is that the basic clade assign¬ ments might represent local minima rather than global solu¬ tions (D.R. Maddison, 1991). This also permitted the analysis of a far greater number of trees within those local minima. The phylogenetic estimate presented is not derived from the most-parsimonious cladograms, but instead from the most- parsimonious trees that stratigraphic data could not reject. Many systematists have asserted that stratigraphic data cannot > > > > > > sym: sym: asym: sym: sym: asym: L: shallow L: deep L: mod L: shallow L: deep L: mod R: shallow R: deep R: deep R: shallow R: deep R: deep Figure 6. —Effects of asymmetrical characters. Hypothetical cladogram for species on which the sinus becomes asymmetrical. A, The most parsimonious optimization of characters states, given that symmetry/asymmetry is coded as a presence/absence feature, and the left and right sides are coded independently. Note that the node pre¬ dicts an impossible collection of character states, as the hypothesized ancestor is symmetrical yet features differ¬ ent left and right sides. B, The shortest acceptable interpretation, in which the hypothetical ancestral morphology is assumed to be identical to the middle morphology. Coding with step matrices (Swofford and Olsen, 1990) would permit parsimony analyses to produce these results; however, these were not computationally feasible in this study. 14 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY offer tests of parsimony estimates, on the assumption that the effects of incomplete sampling distorts stratigraphic data more than it does parsimony estimates (e.g., Smith, 1994). Simulations studies indicate the opposite, however, at least when sampling levels are comparable to those observed among most shelly invertebrates (Huelsenbeck, 1991a; Wag¬ ner, 2000). Several methods that integrate stratigraphic data into phylogenetic analyses have been proposed to date (e.g., Fisher, 1991; Huelsenbeck, 1994), but like parsimony itself, these methods are fundamentally ad hoc because they use stratigraphy as an arbitrary parsimony or reweighting crite¬ rion. A non-ad hoc method uses confidence intervals on strati¬ graphic ranges to test parsimony assessments of phylogeny (Wagner, 1995a). I saved all trees as short as or shorter than the shortest tree known to be fully consistent with strati¬ graphic data and then used a computer program that searched for trees that implied no statistically significant stratigraphic gaps. I evaluated the significance of stratigraphic gaps using confidence intervals on stratigraphic ranges (Strauss and Sa¬ dler, 1989; Marshall, 1990). As a first step, I used Appearance Event Ordination (AEO; Alroy, 1994a) to array the fossilifer- ous horizons that preserve “archaeogastropods,” bellero- phontinae, tergomyans, bivalves, and rostroconchs with ara- gonitic shells 6 . I then used the AEO ordination as a substitute for stratigraphic ranges and calculated confidence intervals using the formula given by Strauss and Sadler (1989) and Marshall (1990) (Appendix 3). The calculation of confidence intervals assumes that hori¬ zons are distributed randomly throughout stratigraphic ranges (Strauss and Sadler, 1989; Marshall, 1990; but see Marshall, 1994). Differences in ecology, taphonomy, and/or biogeogra¬ phy will violate this assumption (Wagner, 1995a). Closely re¬ lated species might be assumed to have similar ecologies and taphonomies owing to phylogenetic autocorrelation; however, different clades and morphotypes likely will have different ecologic distributions and taphonomies, and hence have dif¬ ferent fossil records. The absence of species from horizons in which no close relatives or morphologically similar snails oc¬ cur are not meaningful (e.g., Bottjer and Jablonski, 1988) and to include such horizons inflates confidence intervals unrealis¬ tically. To control for such differences in expected preserva¬ tion patterns, I described the stratigraphic ranges of species based on only horizons including members of their more in¬ clusive clades (see Appendix 3). To control for differences in biogeography, I determined stratigraphic ranges and confi- 6 Species with calcitic shells occur in horizons in which aragonitic shells are absent or rendered unrecognizable. Including these horizons biases the statisti¬ cal analyses of stratigraphic ranges for calcitic species by giving such species more finds (and thus shorter confidence intervals) than is possible for arago¬ nitic species with identical stratigraphic ranges. Using only horizons that in¬ cluded noncalcitic “archaeogastropods” offers a taphonomic control and, thus, each species is evaluated based on fossiliferous horizons that could have pre¬ served any gastropod shell. dence intervals within the individual provinces discussed above (see “Biogeography of Analyzed Species,” above). As a result, some very widespread species have stratigraphic ranges in multiple provinces. When sister species existed in different provinces, the stratigraphic ranges could not be contrasted di¬ rectly. In such cases, stratigraphic correlations based on Har- land et al. (1990) were used to determine whether any strati¬ graphic gap was significant. This was done loosely, so if parsimony considered a Caradocian species from Baltica to be the sister species of a Llanvirn species from Laurentia, then the relationship was rejected only if the confidence interval’s lower bound for the Baltic species was restricted to the Lland- eilo or Caradoc. If that lower bound for the Baltic species ex¬ tended into the Late Llanvirn, then the estimated relationship was accepted even if the Laurentian species was known from the Early Llanvirn. This accommodates the imprecision of cross-provincial stratigraphic correlations. Appendix 3 gives the stratigraphic ranges of analyzed spe¬ cies plus the 95 percent confidence intervals on those strati¬ graphic ranges. Appendix 3 also includes species that could not be included in the cladistic analyses due to inadequate numbers of well-preserved specimens, but that might bridge strati¬ graphic gaps. I used ACCTRAN character optimization, which favors par¬ allelisms to reversals (Swofford and Maddison, 1987). The choice of character optimization does not affect the initial re¬ sults of parsimony analyses. Ancestor-descendant hypotheses eliminate many significant stratigraphic gaps, however, and ACCTRAN makes it less likely that putative ancestors will have apomorphies. Any putative autapomorphies of ancestral species must be considered reversals in descendants, which lengthens a cladogram. Therefore, ACCTRAN ultimately can imply shorter lengths for identical cladistic topologies (Wagner, 1995a). Culling trees that implied statistically significant strati¬ graphic gaps (i.e., gaps greater than the 95% confidence inter¬ val extensions on the stratigraphic ranges of relevant species) resulted in the analysis finding very few equally parsimonious alternatives. Typically only a few trees out of several thou¬ sand would contain no significant stratigraphic inconsisten¬ cies. Some subclades did produce multiple trees of equal length and no significant inconsistencies. In these cases, I used the following criteria. First, I selected the tree that re¬ quired the fewest unknown ancestors. This represents a sec¬ ondary parsimony criterion that other workers have advocated (Alroy, 1995; see also Fisher, 1994; Smith, 1994). Second, I chose trees that had the smallest stratigraphic parsimony debt (Fisher, 1994; Suter, 1994). Any stratigraphic gaps at this point were not statistically significant; however, given the choice of two otherwise equal assessments of a phylogeny, it is logical to select the one that comes closest to predicting the observed pattern in the fossil record (Fisher, 1991; Smith, 1994). Finally, I chose some trees simply because a particular set of synapomorphies led me to prefer that topology over its NUMBER 88 15 rivals. This last criterion is obviously ad hoc; therefore, when discussing these trees below, I describe the rival topologies and detail why I accepted one over the other. Other phylogenetic methods do yield results that have some notable differences from those presented herein. This is partic¬ ularly true of strict parsimony, as characters or (especially) suites of characters identified by some methods as parallelisms between sister clades are identified as synapomorphies of more inclusive clades by parsimony. The general results de¬ scribed herein (i.e., the membership of basic clades and the re¬ lationships among those basic clades) are replicated by other phylogenetic methods, including strict parsimony. Thus, a ma¬ jor change in phylogenetic methods would be required to ob¬ tain radically different results using this character data. Sam¬ pling is another concern, as the ability of parsimony to reconstruct phylogeny decreases with decreasing sample size (Lanyon, 1985; Lecointre et al., 1993). Sampling also de¬ creases the accuracy of methods incorporating stratigraphic data, albeit to a lesser extent (Wagner, 2000). The sampling density of early gastropods, however, appears to be quite good. Based on the metrics of Foote and Raup (1996), be¬ tween 50% and 60% of the broadly distributed species appar¬ ently are included in this analysis (estimates vary according to binning criteria and subclade). These levels are even higher during the crucial early phases of gastropod evolution (i.e., the latest Cambrian and Early Ordovician of Laurentia, where es¬ timates improve to 70%). This means that we should have sampled many direct and indirect ancestors (see Foote, 1996), especially from the critical intervals during which the major groups were diverging. Such sampling greatly increases the efficacy of phylogenetic methods (Huelsenbeck, 1991a) and greatly reduces the concern that new finds will radically alter the estimates presented herein. I place least confidence in the estimated relationships among groups appearing in the earliest Silurian, as this represents the interval of poorest sampling. Unfortunately, this poor sampling seems to coincide with rapid diversification during the rebound from the end-Ordovician mass extinction. As a result, relation¬ ships among clades appearing in the Early Silurian form many polytomies, and there are several species and very small clades with very uncertain affinities. The discovery of heretofore un¬ known Llandovery species undoubtedly will change some of the relationships proposed herein. Many of the cases where this is especially true are emphasized in the text, with the accepted and alternative estimates presented. It should be stressed that these difficulties concern relatively fine-scale relationships, however, and do not effect the estimated relationships among the major groups that were established in the Ordovician. Thus, Silurian sampling is unlikely to affect the basic results of this analysis. As noted above, obtaining radically different results probably will require radically different reinterpretations of shell characters. Cambrian Molluscs and the Choice of an Outgroup Smith (1994) suggested that, of the available methods for rooting a cladogram, the outgroup method of polarizing charac¬ ters makes the fewest a priori assumptions. Choosing an out¬ group however, makes a major assumption about a group’s phylogeny, namely, that the close relatives of the group of in¬ terest (i.e., the ingroup) are known (Adrain and Chatterton, 1990). This often makes the choice problematic (e.g., Ballard et al., 1992). The different phylogenetic models summarized above and in Figure 1 suggest very different outgroups for the “archaeogastropods”; essentially, nearly every possible rela¬ tionship among bellerophontinae, macluritinae, and “archaeo- gastropods” has been proposed at one point. Ballard et al. (1992) addressed a similar problem for arthropods by using a wider phylogenetic analysis to establish an appropriate out¬ group. I adopted a similar strategy by including over 20 differ¬ ent Cambrian molluscs as potential outgroups. These included species assigned to the Bellerophontina, Onychochilida, Pe- lagiellida, Helcionelloida, and Tergomya. I also included two Late Cambrian members of the Hypseloconidae, which might represent the ancestors of cephalopods (Yochelson et al., 1973; Webers and Yochelson, 1989; but see Teichert, 1988). These species are important because cephalopods are likely gastro¬ pods’ closest relatives among the major extant molluscan classes (Naef, 1911; Wingstrand, 1985). I included an Early Ordovician cyrtonelloid, as the only known Late Cambrian rep¬ resentative of the group is represented by only a few poorly preserved specimens (McGhee, 1989). Finally, I also included some early Ordovician bellerophonts, owing to the relatively poor preservation of known Cambrian bellerophonts. I coded all outgroup species as if they were gastropods. For example, I treated the circumbasal carinae of onychochilids as a gastropod peripheral band. Thus, the analysis would not sepa¬ rate onychochilids and pelagiellids from “archaeogastropods” based on a priori interpretations of homology or assumptions about higher taxonomic associations. Density of species sampling strongly affects the precision of phylogenetic analyses (Lecointre et al., 1993). The outgroup analysis represents a very incomplete sample of the known Cambrian molluscs, which in turn represents only a portion of the species that actually existed. One might worry about the potential affect of species that I did not include. To address this, I reran the analyses multiple times after jackknifing the out¬ group (i.e., deleting one of the species; see Lanyon, 1985). I also employed rudimentary rarefaction (see Raup, 1975; Lecointre et al., 1993), running 50 analyses with random sub¬ sets of 50% and 75% of the outgroup species each. The rarefied analyses test whether the cladistic relationships among the in¬ group species are dependent on a few outgroup species. If the relationships within the ingroup vary widely depending upon the outgroup species included (especially at the 75% level), then estimated relationships among ingroup species likely de¬ pend upon the inclusion of particular outgroup species. This 16 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY suggests that adding more species could easily change esti¬ mates of relationships among “archaeogastropods” and that more outgroup species are needed before the results should be trusted. Conversely, if the relationships are stable (especially at the 50% level), then the ingroup results obviously are not strongly dependent on particular outgroup species. In this case, adding new Cambrian molluscs is not likely to affect phyloge¬ netic inferences among “archaeogastropod” species. Results Cladogram Statistics and Descriptions I subjected the character-state matrix to two tests for phylo¬ genetic signal, thegj skewness test (Huelsenbeck, 1991b; Hil- lis and Huelsenbeck, 1992) and the Permutation Compatibility (PC) test (Alroy, 1994b). PAUP estimated the g t statistic based on 10,000 random trees. For the entire matrix, gj = -0.234 (p < 0.0001, Sokal and Rohlf, 1981:174). The significant result indicates that the character-state matrix is significantly more structured than a random collection of characters. Phylogenetic autocorrelation (and thus signal) is a primary candidate for that structure, although nonphylogenetic signals also will produce similar structure. Simulation studies suggest that the most par¬ simonious trees derived from structured matrices are likely to be good approximations of the true phylogeny (albeit, not nec¬ essarily exactly correct; Huelsenbeck, 1991a). The PC test examines whether a matrix contains greater char¬ acter compatibility and hierarchical signal than randomly scrambled matrices. Unlike other permutation tests (e.g., the Permutation Tail Test of Faith, 1991; Faith and Cranston, 1991), the PC test does not assume a particular model of tree¬ building. Also, although phylogenetic autocorrelation (see Raup and Gould, 1974; Felsenstein, 1985) will produce charac¬ ter correlations, other nonrandom but nonphylogenetic signals (e.g., allometry or functional complexes) also will produce character correlations (Kallersjd et al., 1992). The PC test, how¬ ever, should be less susceptible to nonphylogenetic signals (Al¬ roy, 1994b). Finally, the PC test is computationally feasible and could be applied to the whole matrix. Ten thousand permuta¬ tions found that none of the random permutations of the origi¬ nal data matrix possessed either the character compatibility or the hierarchical signal of the original data matrix. This indicates that the character state matrix possesses the type of character correlation that is expected if a phylogenetic signal is present. Each figure includes the consistency index (C.I.) and reten¬ tion index (R.I.) for the species shown, plus the immediate out¬ group (given at the base of the figure). The C.I. is simply the reciprocal of the average number of steps per derived state (Kluge and Farris, 1969); the R.I. is the difference between hy¬ pothesized tree length and theoretical minimum tree length di¬ vided by the difference between the theoretical maximum tree length and the theoretical minimum tree length (Farris, 1989; Archie, 1989). Note that the uninformative characters were ex¬ cluded when calculating C.I. and R.I. These homoplasy indices (especially C.I.) are meaningful only when compared to other analyses (Archie, 1989; Sanderson and Donoghue, 1989), and I include them for that reason only. Observed C.I.’s did typically fall within the range expected given the number of OTUs (see Sanderson and Donoghue, 1989). Workers discussing individual genera and species often have made suggestions about the phylogenetic relationships of those taxa. Whenever possible, I discuss whether this analysis cor¬ roborates those ideas. Also, I treat the taxonomic schemes pre¬ sented in major works (e.g., Knight et al., 1960, in the Treatise on Invertebrate Paleontology) as general phylogenetic schemes. In an evolving system, such as early Paleozoic gas¬ tropods, higher taxonomic definitions should be either mono- phyletic or cohesively paraphyletic (sensu Estabrook, 1986). Some of the higher taxa defined by Knight et al. (1960) were explicitly polyphyletic (see Tracey et al., 1993). I will treat all other taxonomic schemes, however, as general phylogenetic es¬ timates (i.e., proposed monophyly or cohesive paraphyly) when contrasting my results with the ideas of previous authors. Figures 7 through 35 present the accepted cladograms. The nodes are numbered uniformly throughout these figures: for example, node 1 is the same in Figures 7 and 8 and node 11 is the same in Figures 9 and 11. Nodes from the outgroup analysis are lettered rather than numbered. Species with open boxes are identical to the nodes to which they are attached; these species are considered to be ancestors in this study. Black boxes denote species with sufficient autapomorphies to be considered not an¬ cestral to any other taxa included in this analysis. Gray boxes denote equivocal cases (i.e., the taxa lack autapomorphies but there are missing data). The cladogram includes line sketches of each species. Note that these are drawn to a roughly uniform size, not to actual or relative scale. The figure captions give the morphologic changes for each node, as hypothesized by ACCTRAN parsimony (Swofford and Maddison, 1987). I have edited these so that the captions do not simply list the character-state changes, but instead de¬ scribe the types of morphologic change. Because one type of change can affect two otherwise independent characters (e.g., see the above discussions about the right and left sides of asymmetric morphologies), this presents a more realistic metric of the amount of morphologic change than does a simple count of synapomorphies. Several points about my use of taxonomy in the following sections need to be stressed here. When I use the Linnaean tax¬ onomy formally (e.g., the Murchisoniina, Murchisonioidea, Murchisoniidae, etc.), I am referring to taxa previously defined by other workers. The one exception is the boxed generic names shown on the cladograms in Figures 7-35, which outline the taxonomic revision discussed below. When I use suprage- neric names in quotes, I am labeling noteworthy clades defined by this analysis. Clade names are entirely informal and are de¬ signed to describe the phylogeny, obviating the need to follow conventional taxonomic rules. To avoid confusion, all sub¬ codes have different names than their more inclusive clades. NUMBER 88 17 For example, the “murchisoniinae” clade does not include a “murchisonioid” or “murchisoniid” subclade simply because the names look too similar in print. Also, not every species is assigned to a subclade at every level. This means that the para- phyletic collection of “raphistomatids” that do not belong to the “lesueurillines” or “holopeines” (see below) are not as¬ signed to any “-me” subclade. I usually name clades after their least derived members in¬ stead of following the conventions of priority. 1 use previously named higher taxa whenever possible, but in many cases, no previously named higher taxa are appropriate. Naming clades after their plesiomorphic members is the only way that mor¬ phologic grade affects clade names. As a result, the “murchiso¬ niinae” included many species with morphologies that are very different from that of traditional diagnoses of murchisonioids. Finally, note that “subclade” describes a monophyletic group within the clade that is being discussed immediately. Thus, the “lesueurillines” are a “raphistomatid” subclade if I am discuss¬ ing relationships among the “raphistomatids,” but it is a clade if I am discussing relationships among the “lesueurillines.” Ta¬ ble 3 summarizes the “archaeogastropod” clades and subclades discussed herein. As noted above, this classification is for use in this discussion only and must not be interpreted as a formal taxonomic revision. I also include a numerical ranking of the clades and sub¬ clades in Table 3 and in the discussions below. This follows the scheme proposed by Hennig (1969) as a replacement for the Linnaean hierarchy. Again, this is not meant as a formal reclas¬ sification but simply as an aid for the reader. A formal taxonomic revision for the early gastropods based on this analysis and adhering to traditional Linnaean taxonomy is presented at the end of the paper. As noted above, the revised generic taxonomy also is presented in Figures 7-35. Outgroup Analyses Figure 7 shows the results of the outgroup analysis, with “ar- chaeogastropods” condensed into the Schizopea typica Ulrich and Bridge in Ulrich et al., 1930, plus Dirhachopea normalis Ulrich and Bridge in Ulrich et al., 1930, clade. Jackknifing the outgroups and rarefying them to 75% of the initial total had no effect on the relationships within the ingroup (Figure 8). Rar¬ efying the outgroup species to 50% of the initial total produced the same ingroup relationships in 46 of 50 analyses and in 205 of the 216 total trees produced by those analyses. The runs that did not result in S. typica being the stem-“archaeogastropod” did not include tropidodiscines, such as Strepsodiscus major Knight, 1948. In all other runs, those species are considered the immediate outgroups of the “archaeogastropods.” Thus, the re¬ sults shown in Figure 8 depend only on our knowing about bel- lerophonts, such as S. major, and do not depend on the inclu¬ sion of any one species of Cambrian mollusc. This reduces the concern that adding more Cambrian molluscs will affect as¬ sessments of gastropod relationships. Table 3.—“Archaeogastropod” clades and subclades discussed herein. Note that I have added “-itcs” to described subclades within “-ides.” “Archaeogastropods” (Figures 7-35, nodes 1-215) I. “Euomphalinae” (Figures 8-19, nodes 2, 11-108) 1.1. “Ophiletoids” (Figure 9, nodes 12-24) 1.1.1. “Ecculiomphalids” (Figure 9, nodes 15-18) 1.1.2. “Lytospirids” (Figure 9, nodes 19-24) 1.2. “Macluritoids” (Figure 10, nodes 25-36) 1.3. “Ceratopeatoids” (Figure 11-19, nodes 37-108) 1.3.1. “Raphistomatids” (Figures 11-14, nodes 42-69) 1.3.1.1. “Lesueurillines” (Figure 12, nodes 44-54) 1.3.1.2. “Scalitines” (Figures 13, 14, nodes 55-69) 1.3.1.3. “Holopeids” (Figure 14, nodes 62-69) 1.3.2. “Helicotomids” (Figures 11, 15-19, nodes 41,70-108) 1.3.2.1. “Ophiletinines” (Figure 15, nodes 74-80) 1.3.2.2. “Euomphalopterines” (Figures 16-19, nodes 81-108) 1.3.2.2. L “Anomphalides” (Figure 16, nodes 83-87) 1.3.2.2.2. “Poleumitides” (Figures 15, 17, 18, nodes 81, 88-96) 1.3.2.2.3. “Pseudophorides” (Figure 19, nodes 97-108) II. “Murchisoniinae” (Figures 8, 20-35, nodes 7, 109-215) 11.1. “Plethospiroids” (Figure 20, nodes 110, 111) 11.2. “Straparollinoids” (Figure 21, nodes 112-118) 11.3. “Hormotomoids” (Figures 22-26, nodes 119-154) 11.3.1. “Subulitids” (Figure 22, nodes 120-123) 11.3.2. “Cyrtostrophids” (Figure 23-26, nodes 124-154) 11.3.2.1. “Goniostrophines” (Figure 24, nodes 131-138) 11.3.2.2. “Omospirines” (Figures 25, 26, nodes 139-154) 11.3.2.2.1. “Loxonematides” (Figure 25, nodes 141-145) 11.3.2.2.2. “Rhabdostrophides” (Figure 26, nodes 146-154) 11.4. “Eotomarioids” (Figures 27-35, nodes 155-215) 11.4.1. “Lophospirids” (Figure 27, nodes 158-165) 11.4.2. “Clathrospirids” (Figures 27-35, nodes 157, 168-215) 11.4.2.1. “Liospirines” (Figures 28, 29, nodes 170-181) 11.4.2.2. “Brachytomariines” (Figures 28, 30-35, nodes 167, 168, 182-215) 11.4.2.2.1. “?Palaeoschismides” (Figures 30, 31, nodes 183, 188) 11.4.2.2.2. “Phanerotrematides” (Figures 30, 32, nodes 185, 186, 190, 191) 11.4.2.2.3. “Luciellides” (Figures 30, 33, nodes 184, 197-201) 11.4.2.2.4. “Planozonides” (Figures 34, 35, nodes 202-215) 11.4.2.2.4.1. “Coelozonites” (Figure 34, nodes 202-208) 11.4.2.2.4.2. “Gosseletinites” (Figure 35, nodes 209-215) Figure 7 places onychochilids and pelagiellids as distant out¬ groups of tergomyans (nodes E-J), and considers “archaeogas¬ tropods” to be a tergomyan subclade (i.e., node L and above). This matches the predictions of Peel (1991a). In addition, these results support the hypothesis that gastropods originated in the late Middle to Late Cambrian (e.g., Linsley and Kier, 1984; Peel, 1991a; Tracey et al., 1993) rather than in the Early Cam¬ brian (e.g., Knight, 1952; Runnegar and Pojeta, 1974, 1985; Pojeta, 1980). The exact relationships among tergomyans are vague as there are no obvious synapomorphies linking hypselo- conoids, cyrtonelloids, or the clade of bellerophontinae + “ar¬ chaeogastropods”. The sole synapomorphy linking Cyrtolites Conrad, 1838, to early gastropods at node N is the presence of a peripheral band, and most workers would not interpret the Ca¬ rina of Cyrtolites as homologous with the peripheral band of gastropods. If the features are convergent, then it would be 18 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Cl: 0.464 RI: 0.677 Mr Taxon I I Pelagielloid WM\ Helcionelloid F7771 Tergomyan Onychochilid Et3t£1 Hypseloconoid E23 Bellerophontoid Lj Euomphaloid BH Macluritoid Equivocal Helcionella subrugosa ^ Oelandia rugosa 2- Latouchella merino o 3 o> 5* Coreospira rugosa/ Sinuella minuata Euomphalopsis involuta Macluritellal walcotti v r? Costipelagiella L. (jq zazvorkai ^ Pelagiella S- subangulata “ Maclurites" thomsoni Scaevogyra swezeyi g Kobayashiella circe zr 5. Matherellina walcotti on Matherella saratogensis Kiringella pyramidalis P CTO ^ o Q 3 Hypseloconus elongatus Knightoconus ^ antarcticus \ Cyrtolites sp. ( “ Chippewaella / 3 patellitheca 2 —=—P U / g- Strepsodiscus major \ g' Strepsodiscus a paucivoluta TO •= Chalarostrepsis praecursor Eobucania mexicana o 3 »—► C/5 _ Peelerophon oehlerti r*T Schizopea typica o Dirhachopea normalis " (go to Node 4) NUMBER 88 19 Figure 7 (opposite).—Outgroup analysis. Nodes with letters denote the out¬ groups, whereas nodes with numbers denote “archaeogastropods.” Planispiral species are shown from a side view, anisostrophic species are shown from an apertural view. In this and subsequent figures the following conditions apply: (1) line drawings are not to scale; (2) numbers (or letters in small capitals) denote nodes, whereas numbers in parentheses denote characters (see Appen¬ dix 1); (3) abbreviations for synapomorphy linking include the following: ACh = anal channel beneath peripheral band; (i = angle between inner margin and peripheral band (90° = peripheral band perpendicular to inner margin; 0° = par¬ allel); BC = basal carina; BL = paired peripheral lira of peripheral band; CA = coiling axis; E = shell expansion; GL = growth lines; IM — inner margin (= col- umellar lip of “normally” coiled species); K = shell curvature; LR = alveozone; LRC = alveozone carina; ML = single peripheral lira of peripheral band; PB = peripheral band; PI = parietal inductura; RR = right ramp; RRC = right ramp carina; T = shell torque; || = parallel to the coiling axis; _L = perpendicular to the coiling axis; and (4) branch patterns denote previous higher taxonomic assign¬ ments (see text). Node A (“Helcionelloids”), tiny univalve shell with no sinus or PB, very strong growth lines, symmetric aperture with moderately broad, short ramps, posterior projection of the aperture, and channeled, curved IM. Node B (Latouchella merino clade), straight IM (95). Node C, very long ramps (55, 56); moderately broad aperture (58, 59); IM channel lost (102). Node D, ACh at top of aperture (48); IM\base angle = 60° (94); IM 1 to CA (98); base projected posteriorly -10° (116, 117); extremely low E (121); open coiling (123); anisostrophic shell (?) (125). Node E (“Paragastropods”), asymmetric aperture (contracted right side) with very long, narrowly projected ramps (55-59); IM at -45° to CA (98); PI || to aperture (104); moderate K (123); ani¬ sostrophic shell (125); loss of septation (128). Node F (Pelagielloids), sigmoi¬ dal aperture (13); ACh rotated towards top of aperture (48); IM at high angle relative to CA (98); inclined aperture (109); moderate K. (123); anisostrophic shell (125). Node G (Onychochiloids), sharp ML (28); (i = -10° (48); flat RR (51, 52); asymmetric aperture with longer right side (54-56); IM -15° past _L to CA (98); aperture inclined backwards (115); anterior projection of aperture (116, 118); high K (123); moderate ultradextral coiling (126). Node H, long LR (56); asymmetric aperture (right side broader) (57-59); IMXbase angle = 105° (94); curved IM (95). Node I, extremely strong GL (15); concave RR (52); very low E (121); high ultradextral T (126). Node J, (3 = -30° (48); very narrow broad right side of aperture (58). Node K. ( Sinuella minuata + Coreospira rug- osa), moderately long ramps (55, 56); very broad aperture (58, 59); RR swell¬ ing present (60); moderate E (121); small size (141). Node L (Tergomyans), very narrow aperture (58, 59); curved IM (95); base projected posteriorly -50° (117). Node M (Hypseloconoids), low K (123); large size (141). Node N, PB present (19). Node O (?Gastropods), presence of a sinus (1); dull, lump-like ML (28); extremely long ramps (55, 56); RR swelling present (60). Node P, swollen base of RR and LR (60, 73); low E (121); low K (123); curvature decreases over ontogeny (124). Node Q, narrow sinus (6, 7); BL present (21); slit present (34); weak swelling atop RR (60, 61); straight IM (95). Node R, moderate K (123); isometric curvature (124). Node 1 (“Archaeogastropods”), round ML (28); (3 = 100° (48); RR swelling dulls over ontogeny (62); moderate swelling at LR base (73, 74); BC present (89); IM 15° off parallel to CA (98); base projected posteriorly -30° (117); anisostrophic shell (125); low T (126). more parsimonious to consider cyrtonelloids and gastropods to have evolved separately from a limpet-like tergomyan similar to Kiringella Rosov, 1975. Homy (1965, 1991) objected to the suggested link between cyrtonelloids and gastropods in part be¬ cause of the absence of Late Cambrian cyrtonelloids. Since then, Berg-Madsen and Peel (1994) described Telamocornu, a cyrtonelloid from the Late Cambrian. Telamocornu existed in the Avalonian province, whereas early gastropods and their other tergomyan relatives dwelt in the Laurentian province, so there is still a biogeographic gap. Extended species-level analy¬ ses of Cambrian tergomyans obviously are needed to make any firm estimates of exact tergomyan relationships. Although this should be a priority for future research, the resampling results described above suggest that such research will not greatly af¬ fect estimates of relationships within the Gastropoda. The limpet-like bellerophont Chippewaella patellitheca Gunderson, 1993, is the least derived member of the tergomyan subclade that includes “archaeogastropods.” Synapomorphies uniting this clade include a strongly curved sinus and a dull (i.e., broad but weakly expressed) peripheral band. This is a noteworthy result because C. patellitheca has an external mor¬ phology appropriate for Haszprunar’s (1988:407) “urgastro- pod.” Chippewaella patellitheca is only known from one Late Cambrian specimen (Gunderson, 1993), so I consider this re¬ sult tentative. Excluding C. patellitheca produced the same in¬ group topologies, so even if C. patellitheca is important for un¬ derstanding the immediate relationships and origins of gastropods, it does not affect assessments of “archaeogastro- pod” relationships. Perhaps the most intriguing result of the outgroup analysis is that the Bellerophontina appear to be diphyletic, including a cohesive paraphylum of species assigned to the Tropidodisci- nae (i.e., Chalarostrepsis Knight, 1948, Eobucania Yochelson, 1968, and Peelerophon Yochelson, 1982) and a secondarily de¬ rived clade (Figure 8, node 10, which includes Sinuites Koken, 1896, and Owenella Ulrich and Scofield, 1897). Essentially, this results from the hypothesis that lenticular-shaped apertures with deep, hyperbolically curving sinuses, monolineate periph¬ eral bands, and posterior projection of the aperture (e.g., Strep- sodiscus or Schizopea Butts, 1926) are plesiomorphic, whereas apertures with swollen left and right ramps and compressed midsections, U-shaped, shallow sinuses, and bilineate or absent peripheral bands (e.g., Taeniospira Ulrich and Bridge in Ulrich et al., 1930, Sinuopea, or Sinuites) are derived. This means that the definition of “archaeogastropods” used herein includes many species assigned to the Sinuitidae. Four of the outgroup analyses run at 50% rarefaction considered Sinuites and rela¬ tives to be the immediate outgroup of coiled gastropods and Sinuopea sweeti Whitfield, 1882, to be the most primitive coiled gastropod; however, as noted above, these runs did not include species such as Strepsodiscus. Characters shared among Strepsodiscus and species such as Schizopea typica re¬ sult in a phylogeny that considers species with round apertures, U-shaped sinuses, and no peripheral bands to be highly derived rather than primitive. This study does not resolve whether taxa such as Strepsodis¬ cus or Tropidodiscus Meek and Worthen, 1866, were gastro¬ pods. Interpretations of Sinuites as untorted center partially on a hypothesized association between of the origin of torsion and anisostrophic coiling (e.g., Ghiselin, 1966; Runnegar, 1981). Regardless of the merits of that hypothesis, its implications are irrelevant for any bellerophonts that evolved from anisostroph- ically coiled species. The muscle scars of some sinuitids are segmented, which has been cited as evidence that sinuitids 20 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY □ □ □ □ Strepsodiscus major (see Figure 7) Schizopea typica (see Figure 7) Ophileta supraplana (go to Node 11) Prohelicotoma uniangulata (go to Node 25) Jar lops is conicus Euconia etna m Rhombella ® umbilicata Dirhachopea i_I normalis (see Figure 7) - Gasconadia , ^ putilla _ Dirhachopea I_I subrotunda Sinuopea I_I basiplanata ■—- Taeniospirp I—I 1st. clairi Hormotoma _j Isimulatrix (go to Node 109) j—| Taeniospira emminencis □ Sinuope< sweeti Owenella antiquata Cloudia buttsi Sinuites sowerbyi NUMBER 88 21 FIGURE 8 (opposite).—Relationships among the earliest “archaeogastro- pods.” Genus names in boxes denote proposed generic revisions (see “Sys¬ tematic Paleontology,” below). For abbreviations, see legend to Figure 7. Node P, swollen base of RR and LR (60, 73); low E (121); low K. (123); cur¬ vature decreases over ontogeny (124). Node 1 “Archaeogastropods,” round ML (28); (3 = 100° (48); RR swelling dulls over ontogeny (62); moderate swelling at LR base (73, 74); BC present (89); IM 15° off parallel to CA (98); base projected posteriorly -30° (117); anisostrophic shell (125); low T (126). Node 2 (“Euomphalinae”), moderately strong, flange-like ML (28, 29); PB curves adapically (42); ACh present (45); P = 70° (48); very long ramps (55, 56); weak LR swelling (71, 74); IM at -45° to CA (98); curved base (120). Node 3 (Rhombella clade), asymmetric sinus (shallower left side, wider right side) (2-7); P = 110° (48); asymmetric aperture (left side shorter and nar¬ rower) (54-59); lM\base angle = 120° (94); IM 30° off parallel to CA (98); thin PI (103); inclined aperture (109); moderate K (123); moderate T (126). Node 4 (Dirhachopea normalis clade), BL present (21); IM\base angle = 75° (94). Node 5, sinus angle = 50° (3, 4); PB bilineate throughout ontogeny (27, 41); P = 90° (48); RR swelling with isometric shape (62). Node 6, sinus angle = 40° (3, 4); sinus curvature continuous (9, 10); PB width = 20° (20); strong LR swelling (71, 74); IM || to CA (98); radial base (116, 118); septation absent (128). Node 7, PB width = 25° (20); elongated RR (54-56); broad symmetric aperture (58, 59); weak swelling atop RR (60, 61) and base of LR (74); IM thicker than shell (87); lM\base angle = 90° (94); thin PI (103); IM reflected around umbilicus (106); high K (123); very high T increasing over ontogeny (126, 127). Node 8, strong RR swelling (61). Node 9, narrow, U- shaped sinus (6, 7, 9, 10); PB lost (19); weak lunulae (39); high K. (123); small size (141). Node 10 (Sinuitids), sinus angle = 30° (3, 4); isostrophic, planispiral shell (125, 126). were untorted (e.g., Runnegar, 1981) and as evidence that sinu¬ itids were highly derived gastropods (e.g., Haszprunar, 1988; Peel, 1991c; Horny, 1992b, 1995a). This analysis obviously supports the latter scenario. Given the sparse number of sinu- itid species included in this analysis, however, broader analyses of sinuitids and other bellerophonts obviously are necessary. Relationships among Early Ordovician Species Two major subclades of “archaeogastropods” diverged by the Tremadoc (Figure 8). These correspond to nodes 2 and 7 on the cladogram. One clade contains species classified in Plethospira Ulrich in Ulrich and Scofield, 1897, Hormotoma Salter, 1859, and Turritoma Ulrich in Ulrich and Scofield, 1897 (Figures 8 and 20, nodes 7 and 109). Workers have classified the above genera in the Murchisoniina, so I refer to their clade as the “murchisoniinae” hereafter. Synapomorphies include a bilineate peripheral band with two moderately strong, rounded threads, a thickened and reflected inner margin, a relatively shallow, continuously curving sinus, and high translation that increases over ontogeny. The other clade includes species clas¬ sified in genera, such as Ophileta Vanuxem, 1842, Ceratopea Ulrich, 1911, and Lecanospira Butts, 1926 (Figures 8-11, nodes 2, 11, 37). Previous workers provided no consensus about the superfamilial classifications of these genera: Knight et al. (1960) considered Ophileta to be a pleurotomarioid, Cer¬ atopea to be an euomphaloid, and Lecanospira to be a macluri- toid; Yochelson (1973, 1984) considered all three to be pleuro- tomarioids, and N.J. Morris and Cleevely (1981; also Ulrich and Scofield, 1897; Koken, 1925; Wenz, 1938) considered all three to be euomphaloids. The phylogeny suggested in Figures 8-19 best matches the phylogenetic model of N.J. Morris and Cleevely, so it seems most appropriate to designate the second clade the “euomphalinae.” There also is no evidence suggest¬ ing that the Mesozoic Pleurotomaria Defrance, 1824, evolved within this clade rather than from the “murchisoniinae”; in fact, as discussed below, the opposite seems more likely. “Eu¬ omphalinae” synapomorphies include strong, flange-like monolineate peripheral bands that curve abapically, an inner margin that projects away from the coiling axis, curved bases, and circumumbilical thickenings. I. “EUOMPHALINAES” There are several “euomphalinae” subclades, including three that approximate traditional definitions of the Raphistomatidae, Macluritoidea, and Euomphaloidea. The “euomphalinae” also include most of the putative trochoids of the early Paleozoic. As noted above, the “euomphalinae” clade defined herein is very similar to the Euomphaloidea sensu N.J. Morris and Cleevely (1981). Those authors suggested that the Euompha¬ loidea included two basic clades, the Ophiletidae and the Helic- otomidae (with two other families, the Euomphalidae and the Omphalotrochidae, considered to have evolved from helicoto- mids). This analysis suggests that most of the taxa assigned by N.J. Morris and Cleevely (1981) to the Schizopea group of the Ophiletidae (e.g., Schizopea, Ophileta, Ceratopea, Dirhacho¬ pea) represent a paraphylum relative to all later gastropods: Ophileta species (e.g., O. supraplana Ulrich and Bridge in Ul¬ rich et al., 1930, and O. complanata (Miller, 1889)) are plesio- morphic relative to later-appearing “euomphalinae,” Dirhacho¬ pea species are plesiomorphic relative to the “murchisoniinae,” and Schizopea is plesiomorphic relative to all other “archaeo¬ gastropods.” 1.1. “Ophiletoids” Most of the genera assigned by N.J. Morris and Cleevely (1981) to two other ophiletid groups (e.g., the Lecanospira and Lytospira groups) represent a single clade of early “euomphali¬ nae” (Figure 9, nodes 11-24). The earliest member of this clade is the type species of the genus Ophileta, so I refer to the clade as the “ophiletoids.” “Ophiletoid” synapomorphies include a very lenticular aperture, the loss of the swelling at the base of the alveozone (i.e., left ramp), and a very strong peripheral band. More-derived species (e.g., the Lecanospira compacta clade) share a very narrow, sharp monolineate peripheral band that is near the top of the whorl, a concave right ramp and slightly ultra-dextral coiling that give specimens a “bowl” shape, and a flat “base” that is perpendicular to the coiling axis. Septation is plesiomorphic to “archaeogastropods,” but it is es¬ pecially prominent in many “ophiletoids.” Some derived spe¬ cies assigned to the genus Lytospira display a long channel 22 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY □ □ Ophileta supraplana (see Figure 8) Ophileta complanata Lecanospira compacta Lecanospira nereine I I Barnesella llecanospiroides | | Malayaspira rugosa |jgj Maclurina lannulata □ Malayaspira hintzei |—| Rossospira harrisae Ecculiomphalus bucklandi | Barnesella measuresae □ □ Lytospira yochelsoni Lytospira gerrula Lytospira Inorvegica Ophiletina aff. O. sublaxa (.sensu Rohr Lytospira angelini Lytospira subrotunda NUMBER 88 23 FIGURE 9 (opposite).—Relationships among the “ophiletoids.” For abbrevia¬ tions, see legend to Figure 7. Node 11 (“Euomphalinae”), strong BC (91); thin PI (103); very low E (121). Node 12 (“Ophiletoids”), strong ML (29); LR flat¬ ter than RR (51, 52, 72); long ramps (55, 56); loss of LR swelling (73). Node 13 ( Lecanospira compacta clade), PB width = 10° (20); sharp ML (28); weak lunulae (39); PB on top of aperture = 0° (48); extremely narrow aperture (58, 59); weak RR swelling (61); 1M 1 to CA (98); low ultradextral T (126). Node 14 (Barnesella Vecanospiroides clade), sinus angle = 45° (3, 4); juvenile GL stronger (16); V-shaped lunulae (38); ACh lost (45); asymmetric aperture (LR contracted) (54-56); RR swelling dulls over ontogeny (62). Node 15 (“Eccu- liomphalids”), strong lunulae (39); |3 = 10° (48); asymmetric ramp shapes (RR more convex) (52, 53, 72); narrow aperture (58, 59); RR becoming shorter and rounder over ontogeny (63); BC lost (89); curved 1M (95); IM becoming thicker and rounder over ontogeny (97); IM nearly 1 to CA (98). Node 16, left side of sinus sharper than right (2-4); weakly imbricated GL (17); LR wider than RR (57-59); RRC and LRC present (64, 75). Node 17, moderately broad left side of aperture (59); small size (141). Node 18, basal GL strength same as rest of shell (18); RR swelling acute through ontogeny (62); IM _L to CA (98); open coiling (123). Node 19 (“Lytospirids”), sinus curvature continuous (9, 10); flat RR (52); projecting, peg-like BC beneath outer margin (90, 91, 93). Node 20, very narrow aperture (58, 59); RR swelling becoming acute over ontogeny (62); open coiling (123); carrier-shell scars (140). Node 21, PB width = 05° (20); PB X to aperture (42). Node 22, (Ophiletina cf. O. sublaxa clade), sinus angle -30° on right side (3) with little curve on right side of sinus (9). Node 23, left side of sinus sharper than right (2-4); sinus angle -50° on left side (4); left angle of sinus sharper (6—8); extremely narrow aperture (58, 59); extremely low E (121); open coiling (123). Node 24 ( Lytospira angelini clade), no ontogenetic change in GL strength (16); concentric lunulae (38); convex RR (52); symmetric ramp widths (54—56). along the base (see Rohr, 1993). This likely is a site of muscle attachment, which suggests that the base of “ophiletoid” spe¬ cies is homologous with the inner margin of more typical gas¬ tropods. Although Rohr (1993) described this channel as unique to Lytospira, similar channels exist on the columellas of species classified in Ceratopea, Pararaphistoma, and Pachys- trophia. Thus, the columellar channel seems to have been a re¬ curring, polyphyletic character among the “euomphalinae.” A noteworthy feature in “ophiletoid” evolution is the parallel development of open-coiling in the “ecculiomphalids” (Figure 9, nodes 15-18) and “lytospirids” (Figure 9, nodes 19-24). This shell form evolved only infrequently after the Early Or¬ dovician, but it appeared many times among Early Ordovician “euomphalinae.” The open-coiled forms illustrated in Figure 9 almost certainly were sessile filter-feeders, as the animals would not have been able to effectively balance their shells (Yochelson, 1971; Linsley, 1977; N.J. Morris and Cleevely, 1981). This also has been considered the likely mode of life for nearly planispiral but close-coiled (i.e., coiled with whorls in contact) gastropods, such as those that occupy the base of the “ophiletoid” cladogram (Linsley, 1978). By deriving these more specialized sessile morphologies from the close-coiled forms more than once, these results support the idea that sessile filter feeding first appeared early in “ophiletoid” evolution. “Lytospirid” species bear external shell scars suggesting that they were carrier shells, i.e., small shells, shell fragments, or pebbles were cemented onto the side of the shell as the animal grew (Rohr, 1993; see Linsley and Yochelson, 1973). The one exception is the slightly problematic species Ophiletina cf. O. sublaxa (sensu Rohr, 1988, not Ulrich and Scofield, 1897), which possesses a broad frill in the same position as the shell scars on Lytospira species. Linsley and Yochelson (1973) sug¬ gested that attached shells and large shell frills might serve a similar functional role, i.e., serving as a support or a “snow- shoe” on soft substrates (see also Linsley et al., 1978). In addi¬ tion, early Lytospira species possess a moderately strong, peg¬ like (i.e., with a parallel surface) carina in nearly the same loca¬ tion (at the alveozone-base intersection), which is just below the scars. Therefore, it seems plausible that the frill is function¬ ally homologous with shell carrying and phylogenetically ho¬ mologous with the carina of Lytospira. There has been little discussion of the phylogenetic relation¬ ships among these snails. In general, the analysis supports the suggestion that Lecanospira and its relatives are not closely re¬ lated to macluritoids (Linsley and Kier, 1984; Yochelson, 1984; contra Knight et al., 1960). It also supports the idea that Lecanospira and Barnesella Bridge and Cloud, 1947, are closely related (node 14; Bridge and Cloud, 1947; Knight et al., 1960). Rohr (1994) suggested a close relationship between Rossospira Rohr and Malayaspira, which this analysis also supports; however, this analyses’ suggestion that Eccu- liomphalus Portlock evolved from that clade (see node 18) ap¬ pears to be novel. Finally, there are Silurian and Devonian spe¬ cies assigned to Lytospira (e.g., Homy, 1992a), but this study suggests that those species belong to the “raphistomatid” clade (see below). Thus, it appears that the “ophiletoids” were en¬ tirely extinct by the Early Silurian. 1.2. “Macluritoids” This analysis indicates that Prohelicotoma, Macluritella, Tei- ichispira, Monitorella Rohr, 1994, Maclurites, and Palliseria Wilson, 1924, represent a subclade of “euomphalinae” (Figure 10, node 25). “Macluritoid” synapomorphies include a straight, shallow sinus with a weak monolineate peripheral band, exag¬ gerated basal growth lines, a nearly round aperture, and nearly planispiral coiling that opens in the adult whorls. Many of these characters are apparent in the juvenile whorls of species classified as Macluritella or Teiichispira, which produced char¬ acters typical of more-derived macluritids later in ontogeny. More-derived species (e.g., nodes 27-36) share distinctive on¬ togenetic changes involving differential expansion of the base and left side, counter-clockwise rotation of the aperture, and ul- tra-dextral coiling. Still more-derived species (e.g., nodes 31-36) have narrow sinuses and complete filling of juvenile whorls instead of septa. Additional synapomorphies not used in this analysis are as¬ sociated with the calcareous opercula of “macluritoid” species (Yochelson, 1975). Opercula associated with Teiichispira spe¬ cies are fibrous, horn-shaped structures, which include knob¬ like muscle attachments on later species. The opercula of more- derived species (i.e., Maclurites, Monitorella, and possibly 24 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Prohelicotoma [ ] uniangulata (see Figure 8) Macluritella _ Teiichispira I_I loceana |—| Teiichispira '—'odenvillensis _ Monitorella I_I auricula Teiichispira kobavashi ‘Eccyliopterus I ornatus ” Mitrospira □ longwelli Palliseria robusta I—- Teiichispira I—I sylpha ___ Maclurites I_I magna Maclurites M expansa Maclurites bigsbyi Maclurina logani _ Maclurites I_I sedgewicki iT *\ Maclurina manitobensis NUMBER 88 25 FIGURE 10 (opposite).—Relationships among the “macluritoids.” For abbrevia¬ tions, see legend to Figure 7. Node 25 (“Macluritoids”), sinus angle = 30° (3, 4); sinus curve nearly straight (9, 10); PB width = 10° (20); ML becoming weaker over ontogeny (30); weak lunulae (39); P = 40° (48); globular ramps (52, 72); moderately long ramps (55, 56); IM nearly 1 to CA (98); moderately high E (121). Node 26, sinus angle = 20° (3, 4); weak GL (15); PB width = 05° (20); PB on top of aperture = 0° (48); counter-clockwise rotation of aperture over ontogeny (49); RR becoming more concave over ontogeny (53); IM ± to CA (98); low ultradextral T (126). Node 27, extremely narrow aperture (58, 59); differential ontogenetic expansion of LR (86, 122). Node 28, loss of RR and LR swellings (60, 73); convex LR (72); base projected posteriorly -10° (117); straight base (120). Node 29 (Monitorella auricula clade), PB width = 10° (20); round ML (28); p = -10° (48); IM\base angle = 75° (94); weakly curved IM (95). Node 30 (Teiichispira kobayashi clade), PB _L to aperture (42); moderately long RR and LR (55, 56); IM nearly 1 to CA (98); high E (121). Node 31 (Teiichispira sylpha clade), narrow sinus (6, 7); isometric ML strength (30); lunulae same strength as GL (39); ACh lost (45); symmetric ramp shapes (52, 53, 72); asymmetric ramp lengths (LR longer than RR) (54-56); asymmet¬ ric aperture (RR wider than LR) (57-59); thickened IM (87); complete infilling of juvenile whorls (128). Node 32 (Palliseria clade), weak ML (29); flat RR (52); BC lost (89); strongly curved IM with very thick middle (87, 95, 96); IM -30° past -L to CA (98); isometric E (121, 122); moderate ultradextral T (126). Node 33 (Maclurites clade), very narrow sinus (6, 7); fine, sharp GL (15); PB width = 05° (20); PB _L to aperture (42); no ontogenetic rotation of aperture (49); asymmetric ramp shapes (RR more convex) (52, 53, 72); base projected posteriorly -20° (117); high E (121). Node 34 (Maclurites bigsbyi clade), weak ML (29); flat RR (52); moderately long RR (55); narrow right side of aperture (58); no differential expansion of left side (86, 121, 122); straight IM (95); IM 1 to CA (98); base projected posteriorly -10° (117); ornate base (129). Node 35, very weak ML (29); stronger ornament (131). Node 36, ornate LR (129). Palliseria; but see Rohr, 1994) are shield-like rather than horn¬ like, but retain the knob. The cladogram suggests that the shield-like operculum is derived from the horn-like one. Unfor¬ tunately, there are too few exact shell-operculum associations to use opercular characters, but more detailed analyses of ma- cluritoid opercula and associated shells could be used to test these results in the future. Although Knight (1952) and Erwin (1990b) previously sug¬ gested a close relationship between euomphaloids and macluri¬ toids, this study proposes a very different model of “macluri- toid” evolution than those authors suggested. Knight (1952) and others (e.g., Wangberg-Eriksson, 1979; Runnegar, 1981, 1996; Linsley and Kier, 1984) assumed that Maclurites and re¬ lated species evolved from onychochilids and, therefore, di¬ verged from the main gastropod clade during the Early to Mid¬ dle Cambrian. This analysis, however, suggests that onychochilids and pre-Ordovician species assigned to the Ma- cluritidae (e.g., Macluritellal walcotti Yochelson and Stinch- comb, 1987, and Euomphalopsis involuta Ulrich and Bridge in Ulrich et al., 1930) are related only distantly to the “macluri¬ toids” and other gastropods (see Figure 7, nodes E, G-J). The results of this study corroborate the predictions of RJ. Morris (1991) and, in part, McLean (1981), Linsley and Kier (1984), and Yochelson (1984), but the traditional models merit further investigation. This is best done by examining whether a more traditional estimate of phylogeny produces a signifi¬ cantly longer tree than the one accepted herein. The shortest tree linking the two clades assumes that the Middle Ordovi¬ cian Palliseria and Maclurites are the least derived members of the clade and that the earliest species (e.g., Macluritella and early Teiichispira) are the most derived. Sampling of “maclu- ritoid” species (especially of Palliseria and Maclurites ) is dense, so stratigraphic data reject at high levels any tree that considers the morphologic intermediates to be phylogenetic intermediates (see Appendix 3). Accordingly, I used the short¬ est non-rejectable tree that linked the “macluritoids” with members of the Onychochilidae (which placed “macluritoids” outside the Gastropoda) and compared this tree to the accepted tree (including the outgroups and “macluritoid” species that appeared through the Early Arenig (Early Ordovician)). To test whether a less-parsimonious tree is significantly longer than a more-parsimonious tree, Alroy (pers. comm., 1994) proposed bootstrapping character-states matrices to determine whether random sections of the matrix suggested that one esti¬ mate of phylogeny is as short as another. Bootstrapping is ap¬ propriate only when resampling independent units. This as¬ sumption is violated by the additive coding schemes used herein (see O’Grady et al., 1989); for example, it is impossible to know that some species have a particular type of peripheral band without simultaneously knowing that those species have a peripheral band. Thus, if we sampled character 21 (the pres¬ ence or absence of peripheral lira on the peripheral band), then we know that we have sampled character 19 (the presence or absence of the peripheral band itself). Similarly, if one ran¬ domly samples a character describing the left side of some fea¬ ture, one should know that the species has a right side of that same feature. As an alternative, I bootstrapped the branch lengths on the two different cladograms. For two trees with the same taxa and characters, greater branch lengths indicate some combination of greater homoplasy and poorer sampling of evolutionary transitions (i.e., with poorer sampling, more mor¬ phologic change happens between sampled species). This means that bootstrapping branch-lengths tests whether one tree requires significantly more homoplasy and/or worse sam¬ pling than another. None of the 1000 bootstrap analyses of the accepted tree are as long as the alternative tree, nor are any of the 1000 bootstrap analyses of the alternative tree as short as the accepted tree. This apparently is because the putative homologies between the “macluritoids” and onychochilids exist largely in late ap¬ pearing “macluritoids,” not in the early appearing species. Overall, these data suggest that the traditional hypothesis is sig¬ nificantly less parsimonious than the tree presented herein and also strongly implies that “macluritoids” represent a very re¬ stricted clade of Ordovician gastropods that are not closely re¬ lated to other ultra-dextral molluscs of the early Paleozoic. Thus, contrary to Runnegar (1981, 1996), there is neither a morphologic nor a stratigraphic progression linking ony¬ chochilids and “macluritoids.” 26 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 1.3. “Ceratopeoids” The other major “euomphalinae” subclade (Figures 11-14, nodes 37-69) contains a number of species classified in the genus Ceratopea. This genus has not been used previously to label any higher taxon, but the plesiomorphic nature of Cer¬ atopea species makes it the most appropriate label for this clade. The earliest “ceratopeoids” (Figure 11) possess deep, strongly curved sinuses, strong abapically hooked peripheral bands (see Figure 11), a sharp, thick basal carina, a crenulated base, and a Lytospira- like channel within the columella. Syna- pomorphies linking later members of the clade include similar ontogenetic changes in aperture orientation, asymmetrical si¬ nuses, and peripheral bands placed asymmetrically onto the right ramp. “Ceratopeoids” feature two major subclades, the “raphisto¬ matids’' and “helicotomids,” and one smaller subclade. This small subclade includes species that have been classified under a variety of generic names, including Bridgeites Flower, 1968a, and Bridgeina Flower, 1968a (Figure 11, nodes 38, 39). The “bridgeitids” are diagnosed by a sharply hooked peripheral band, strongly exaggerated basal and juvenile growth lines, and an extremely lenticular aperture. More-derived species feature nearly planispiral shells with open-coiling in adult stages. This short-lived subclade was relatively diverse during the early Arenig (Early Ordovician) but apparently became extinct shortly thereafter. 1.3.1. “RAPHISTOMATIDS” One of the major “ceratopeoid” clades includes a number of taxa that workers from the turn of the century (e.g., Koken, 1898, 1925) associated with the Raphistomatidae. Accord¬ ingly, this clade is labeled the “raphistomatids” (Figures 11-14, nodes 42-68). The “raphistomatid” clade is much broader than the most recent definitions of the family (i.e., Wenz, 1938; Knight et al., 1960), but it is similar to the defini¬ tions presented herein. Synapomorphies include a very high si¬ nus angle, pronounced growth lines on juvenile whorls, and a crenulated base. 1.3.1.1. “LESUEURILLINES”. —Species classified in the genus Lesueurilla dominate one of the major “raphistomatid” sub¬ clades (Figure 12, nodes 44-54). That genus name has never been used to label a suprageneric taxon, but it is the most ap¬ propriate name for this particular clade. “Lesueurilline” syna¬ pomorphies include very strong, hooked peripheral bands and a distinctive sigma-shaped lunulae (see Figure 4 f). An interest¬ ing feature of “lesueurilline” evolution is that nearly planispiral coiling is the primitive condition both ontogenetically and phy- logenetically. Juvenile and early forms possess Lesueurilla- like morphologies, whereas adult forms (among derived species) possess more lenticular morphologies (e.g., see Figure 3). The “lesueurillines” also include the earliest clade diagnosed by a slit (the L. prima clade, Figure 12, node 48). The slit of these species is not a distinct feature, but instead it is an extension of the sinus where the left and right halves run parallel to each other near the apex. The morphogenetic development of the slit from the sinus is most obvious on species such as Pararaphis- toma qualteriata (Schlotheim) and P. schmidti (Koken, 1925) on which the slit appears later in ontogeny. This analysis supports Yochelson’s (1982, 1984) proposition that Lesueurilla and Climacoraphistoma are close relatives. All of the characters that Yochelson cited as linking the two taxa are synapomorphies of relevant nodes, and there are additional synapomorphies that Yochelson did not list, such as the nar¬ rower, somewhat asymmetrically shaped sinus and the loss of the columellar channel. Another pertinent implication is that Eccyliopterus Remele, 1888, is closely related to Eccu- liomphalus Portlock, 1843 (Knight et al., 1960). Eccu- liomphalus (or at least the type species of that genus, E. buck- landi Portlock, 1843) apparently is an “ophiletoid” (see discussion above), whereas Eccyliopterus represents a sister group to derived Lesueurilla plus the clade of Climacoraphis¬ toma + Pararaphistoma. A more tentative suggestion by Yoch¬ elson and Copeland (1974) is that Ceratopea and Pararaphis¬ toma are synonymous. This analysis implies that the similarities between Pararaphistoma and Ceratopea are due to reversals during the evolution of Climacoraphistoma and Pararaphistoma. The second “lesueurilline” subclade includes species classi¬ fied as Raphistoma Hall, 1847, and Scalites Emmons, 1842. Synapomorphies of the clade include increased curvature and differential expansion of the lower portion of the shell, which increased both the whorl expansion and translation rates (sensu Raup, 1966) and provided a more oval aperture shape. Some later species (e.g., R. striata Emmons, 1842, R. peracuta Ulrich and Scofield, 1897, and S. katoi Kobayashi, 1934) possess si¬ nuses with a distorted right half and a sigmoidal left half (Fig¬ ure 13, nodes 55-59). “Lesueurillines” were relatively diverse during the late Arenig through the Llandeilo (Middle Ordovician). Although a few species survived into the Caradoc (Middle to Late Or¬ dovician), there are no known Silurian “lesueurillines.” It also FIGURE 1 1 (opposite).—Relationships among the “ceratopeoids.” For abbrevia¬ tions, see legend to Figure 7. Node 11 (“Euomphalinae”), strong BC (91); thin PI (103); very low E (121). Node 37 (“Ceratopeoids”), long ramps (55, 56); weak RR swelling becoming acute over ontogeny (61, 62); flat LR with no swelling (72, 73); thickened IM (87); moderate K (123). Node 38 (Bridgeites clade), (3 = 50° (48); IM nearly 1 to CA (98); base projected posteriorly -30° (117); low K (123); curvature decreases over ontogeny (124); nearly planispi¬ ral coiling (126). Node 39, ACh lost (45); asymmetric ramp shapes (RR rounder (51-53)), ramp lengths (LR contracted (54-56)), and aperture breadth (RR wider than LR (57-59)); strong RR swelling that dulls over ontogeny (61, 62); IM _L to CA (98); IM channel lost (102). Node 40 (Ceratopea llaurentia clade), strong basal GL (18); strong, flange-like ML (28, 29); asymmetric ramp shapes (RR rounder) (51-53); pronounced, channeled BC (90); IM\base angle = 105° (94); base projected posteriorly -30° (117); moderate E (121). Node 41 (“Helicotomids”), fine, sharp GL (15); clockwise rotation of aperture over ontogeny (49); ontogenetic increase in T (127). Node 42 (“Raphistomatids"), juvenile GL stronger (16); counter-clockwise rotation of aperture over ontog¬ eny (49); elongated RR (54-56); RR swelling dulls over ontogeny (62). NUMBER 88 27 Ophileta supraplana (see Figure 8) Ceratopea canadensis Orospira bigranosa □ Bridgeites planodorsalis □ Bridgeites supraconvexa Bridgeites Idisjuncta □ Ceratopea llaurenha □ Ceratopea pygmaea Ceratopea unguis (go to Node 70) □ Pararaphistoma lemoni Helicotoma medfraensis Palaeomphalus giganteus (go to Node 43) 28 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY NUMBER 88 29 FIGURE 12 (opposite).—Relationships among the “lesueurillines.” For abbrevi¬ ations, see legend to Figure 7. Node 43 ( Palaeomphalus giganteus clade), sinus angle = 50° (3, 4); stronger juvenile GL (16); sigma-shaped lunulae (38); loss of RR swelling (60); curved IM (95). Node 44 (“Lesueurillines”), PB width = 10° (20); [3 = 30°-^t0° (48); asymmetric ramp shapes (LR rounder than RR) (51, 52, 72); IM thickness same as rest of shell (87); IM\base angle = 90° (94); IM 60° off parallel to CA (98); nearly planispiral coiling (126). Node 45 (Eccylioplerus regularis clade), sinus angle = 40° (3, 4); very strong ML (29); no ontogenetic rotation of aperture (49); PI thickness same as rest of shell (103); extremely low K (123). Node 46, asymmetric aperture (LR contracted) (54-56). Node 47, PB width = 05° (20); extremely strong ML (29); asymmetric aperture (RR wider than LR) (57-59); thickened IM (87); IM at -45° to CA (98). Node 48 ( Lesueurilla prima clade), left side of sinus more obtuse (2^4); left side of sinus narrower (5-7); slit present, leaving weak lunulae (34, 39); PB partially on RR (43); IM channel lost (102). Node 49 ( Eccylioplerus louder- backi clade), PB slightly raised relative to whorl (33); RR becoming more con¬ cave over ontogeny (53); BC lost (89); curved base (120); high E (121); large size (141). Node 50, wide left side of sinus (7); no ontogenetic rotation of aper¬ ture (49); very high E (121); isometric T (127). Node 51 ( Lesueurilla margin¬ al is clade), IM at -45° to CA (98); isometric T (127). Node 52, fine, sharp GL (15); no ontogenetic change in GL strength (16); ACh lost (45); flat LR (72). Node 53 ( Climacoraphistoma clade), high rotation of aperture (50); asymmet¬ ric ramp shapes (RR more convex) (52, 53, 72); IM at -45° to CA (98). Node 54, basal GL strength same as rest of shell (18); symmetric ramp shapes (51); IM 30° off parallel to CA (98); straight base (120); low T (126). bears noting that the two basic “lesueurilline” subclades gen¬ erally were restricted to different biogeographic realms from their origins in the middle Arenig (Early Ordovician) through the Llandeilo (Middle Ordovician). The Lesueurilla clade (Figure 12) occurs predominantly in the Baltoscandian faunas of Europe, whereas early members of the Scalites clade (Fig¬ ure 13) exist predominantly in the early Laurentian fauna of North America. Some geographic overlap between the two clades does occur in Malaysia and western North America during that time, and members of both clades occur in both faunas from the Caradoc through the Ashgill (Middle to Late Ordovician). 1.3.1.2. “HOLOPEINES”. —The second “raphistomatid” sub- clade comprises species classified in Raphistomina Salter, 1859, Pachystrophia Pemer, 1903, Sinutropis Pemer, 1903, and Holopea Hall, 1847 (Figures 13, 14, nodes 60-69). This clade is diagnosed primitively by a completely U-shaped sinus (nodes 60-69), although some derived species have no sinus. Pachystrophia includes the least derived members of a clade diagnosed by the loss of a peripheral band (Figure 14, nodes 62-69). Later species (i.e., Pachystrophia gotlandica (Lind- strom, 1884) and Lytospira subuloides Barrande in Perner, 1903) (Figure 14, nodes 68, 69) include the only open-coiled, nearly planispiral species known from the Silurian. In addi¬ tion, Silurian species placed by Knight et al. (1960) in the fam¬ ily Sinuopeidae (e.g., Horiostomella Perner, 1903, and Sellinema Pemer, 1903) likely also belong to this clade. The status of the post-Silurian members of that family are not known, but it is much more likely that they are “holopeines” than it is that they are related to the Late Cambrian-earliest Ordovician Sinuopea species. Species assigned to the genus Holopea are a particularly con¬ spicuous development within the “holopeines” (Figure 14, nodes 64, 65). The earliest Holopea species are nearly identical to contemporaneous Pachystrophia species, save for the ab¬ sence of the sinus (with the absence of a peripheral band being a synapomorphy that links Pachystrophia and Holopea). The trochiform shell typically associated with the genus is shared among derived Holopea species (node 65). Holopea is an im¬ portant genus when discussing gastropod phylogeny because its family (the Holopeidae) represents the earliest putative members of the Trochoidea in some classification schemes (e.g., Knight et al., 1960). None of the other genera typically assigned to the Holopeidae, however, seem to have been close relatives of Holopea. Although the genus is reported through the Devonian (Knight et al., 1960), this analysis did not find any post-Ordovician members of the H. insignis clade. This makes it seem unlikely that Holopea represents an early tro¬ choid. Other phylogenetic scenarios concerning the Trochoidea are discussed below. Some general phylogenetic proposals concerning the genus Pachystrophia are not supported here. Wenz (1938) considered Pachystrophia to be a close relative of “ophiletoid” genera such as Ecculiomphalus and Lytospira. Also, some “hol- opeides” from the Silurian (at least one of which extends into the early Devonian; see Homy 1992a) have been classified in the genus Lytospira. A very different suggestion was made by Knight et al. (1960), who considered Pachystrophia to be a jun¬ ior synonym of Lesueurilla. This analysis contradicts all of these ideas, suggesting instead that Pachystrophia is distantly related to the “ophiletoids” and that Lesueurilla and Pachystro¬ phia have closer relatives than one another. 1.3.2. “HELICOTOMIDS” The other major “ceratopeoid” subclade includes many of the traditional euomphaloid genera (sensu Knight et al., 1960), plus most of the early Paleozoic taxa classified in the Tro¬ choidea (also sensu Knight et al., 1960; Tracey et al., 1993). The earliest species of the clade agree well with the definition of the Helicotomidae (sensu Knight et al., 1960, not Wenz, 1938), so I refer to the clade as the “helicotomids.” The clade presented herein (Figures 15-19, nodes 70-108) is very similar to N.J. Morris and Cleevely’s (1981) genus-level definition of the Euomphalidae + Helicotomidae clade. “Helicotomid” syna- pomorphies include a sigmoidal aperture shape with the base of the aperture projected in front of the inner margin, a broadly expanded (i.e., nearly round or square instead of lenticular) ap¬ erture, a shallow, weakly curved sinus, a weak monolineate pe¬ ripheral band, a weak thread-like basal carina, and strong chan¬ nels beneath the right and left carina. This analysis also corroborates N.J. Morris and Cleevely’s (1981) opinion that derived “euomphalinae,” such as Po- leumita Clarke and Ruedemann, evolved from early members of the Helicotomidae. Those authors divided the Euompha- 30 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY NUMBER 88 31 FIGURE 13 (opposite).— Relationships among the “scalitines” and “holoepeids.” For abbreviations, see legend to Figure 7. Node 42 (“Raphisto- matids”), juvenile GL stronger (16); counter-clockwise rotation of aperture over ontogeny (49); elongated RR (54-56); RR swelling dulls over ontogeny (62). Node 43 (Palaeomphalus giganteus clade), sinus angle = 50° (3, 4); stronger juvenile GL (16); sigma-shaped lunulae (38); loss of RR swelling (60); curved IM (95). Node 55 (“Scalitines”), basal GL strength same as rest of shell (18); PB width = 10° (20); very strong ML (29); symmetric ramp shapes (51); asymmetric ramp lengths (right contracted) (54-56); very thick IM (87); IM\base angle = 75° (94); IM -15° off parallel to CA (98); IM channel lost (102). Node 56, wrinkled right side of sinus (?) (11); ACh lost (45). Node 57, squared ML (28); very short RR (55); asymmetric ramp projection (RR higher) (57-59); whole aperture inclined -20° (109, 110); noncrenulated base (120); high E (121). Node 58, U-shaped left side of sinus (10); BL present (21); PB slightly raised relative to whorl (33); slit present (34); convex RR (52); strong, nonprojecting BC (91). Node 59 (Raphistoma tellerensis clade), U-shaped left side of sinus (10). Node 60 (“Holopeids”), sinus angle = 25° (3, 4); U-shaped sinus (9, 10); no ontogenetic change in GL strength (16); basal GL strength same as rest of shell (18); (3 = 70° (48); asymmetric ramp shapes (RR rounder) (51-53); symmetric ramp lengths (54-56); RRC present (64); strong, non¬ projecting BC (91); IM 30° off parallel to CA (98); low E and low K (121, 123); isometric T (127). Node 61, PB _L to aperture (42); moderately long ramps (55, 56); extremely narrow aperture (57, 58); IM thickness same as rest of shell (87); nonchanneled, weak BC (90, 91); lM\base angle = 105° (94); IM 15° off parallel to CA (98); curved base (120). loidea into two early families, the Ophiletidae and the Helicot- omidae, which they considered to be sister clades. This analy¬ sis suggests that N.J. Morris and Cleevely's definition of the Ophiletidae (which is very close to the “euomphalinae” as de¬ fined herein, but without species from the Middle Ordovician and later) is paraphyletic relative to their Helicotomidae. 1.3.2.1. “OPHILETININES”. —Two “helicotomid” subclades evolved by the Middle Ordovician. The first of these includes species assigned to Helicotoma Salter, 1859, Palaeomphalus Koken, 1925, and Ophiletina Ulrich and Scofield, 1897 (Figure 15, nodes 74-80). As I previously labeled the more inclusive clade the “helicotomids,” I designate this clade the “ophiletin- ines.” “Ophiletinine” synapomorphies include a moderately wide, flange-like peripheral band that curves abapically and overlies a weak channel, a wide, obtuse left carina that overlies a shallow channel, and very low translation. No known species matches the hypothetical common ancestor of the “ophiletin- ines,” although it likely was most similar to Oriostoma bro- midensis Rohr and Johns, 1992, of the Early Caradoc. Synapo¬ morphies uniting the O. bromidensis clade (Figure 15, nodes 75, 76) include a very shallow, very narrow sinus and a strong, sharp basal carina. Ophiletina species (node 76) share a peg¬ like left carina and a bilineate peripheral band. These species also appear to have had partially calcitic shells, although this was not used in the cladistic analysis. Synapomorphies uniting the Helicotoma tennesseensis clade (Figure 15, nodes 77-80) include a rounded right carina that weakens over ontogeny, an expanded, rounded base, very faint growth lines, an unusually strong, hooked peripheral band, and extreme swelling of the left carina. A disc-like paucispiral operculum is associated with one Helicotoma species (Yochel- son, 1966a). This operculum is very different from the horn¬ shaped opercula associated with early “helicotomids” and other early “ceratopeoids” (e.g., Ceratopea unguis Yochelson and Bridge, 1957; see Yochelson and Wise, 1972). If the phy- logeny presented herein is reasonably accurate, then a pau¬ cispiral, disc-like operculum is derived, but it unfortunately is not known how common that operculum was among the “heli- cotomatid” clade. “Ophiletinines” were moderately diverse from the Middle Ordovician through the Late Ordovician, but this study found no Silurian members of the clade. 1.3.2.2. “Euomphalopterines”. —The second “helicoto¬ mid” subclade includes a diverse array of Silurian species that typically have been assigned to the Anomphalidae, Elasmone- matidae, Euomphalopteridae, and Pseudophoridae (Figures 16-19, nodes 81-108). The Ordovician precursors of this clade have been classified in the genus Euomphalopterus Roemer, 1876 (e.g., Euomphalopterus cariniferus Koken, 1925). I cate¬ gorize the clade as the “euomphalopterines,” even though this clade is very different from previous definitions of the Eu¬ omphalopteridae (e.g., see Wenz, 1938, or Knight et al., 1960). “Euomphalopterine” synapomorphies include a very strong left carina, a shallow V-shaped sinus with a very narrow sharp pe¬ ripheral band, and very broadly projecting ramps. 1.3.2.2.1. “Anomphalides”.—Three major “euomphalopter¬ ine” subclades arose during the latest Ordovician and Early Silurian from an Euomphalopterus cariniferus-hke ancestor (Figures 16-19, nodes 81-108). One of these subclades (Fig¬ ure 16, nodes 83-87) consists predominately of species as¬ signed to the Anomphalidae (e.g., Pycnomphalus Lindstrom, 1884, and Grantlandispira Peel, 1984a). Characters diagnos¬ ing the “anomphalide” clade include a strong but dull carina on a swollen alveozone, a strong lirum on the inner margin that partially fills the umbilicus, and a U-shaped sinus with an apex near the suture. Among more-derived species (e.g., nodes 86, 87), the umbilicus is entirely filled and the periph¬ eral band is highly reduced or lost. Kase (1989) noted that some of the earliest species assigned to the Omphalotrochidae (e.g., Middle Devonian species assigned to Labrocuspis Kase) have synapomorphies with “anomphalide” species; however, the majority of omphalotrochids, including Devonian species assigned to Pseudomphalotrochus Blodgett, 1992, appear to share synapomorphies with Straparollus de Montfort, 1810, and Euomphalus Sowerby, 1814 (Erwin, in prep.; Wagner, in prep.). 1.3.2.2.2. “Poleumitides”.—The second major “euompha¬ lopterine” subclade contains species assigned to Poleumita, Euomphalopterus, Centrifugus Bronn, 1834, and (possibly) Spinicharybdis Rohr and Packard, 1982 (Figures 17, 18, nodes 88-96). This clade bears little resemblance to the Poleumitidae of Wenz (1938), which links Poleumita with the problematic Oriostoma Munier-Chalmas, 1876 (see also Boucot and Yoch¬ elson, 1966). Because the Poleumitidae is an established name, I designate this clade as the “poleumitides.” “Poleumitides” ev¬ idently shared a common ancestor in the Early Silurian that 32 SMITHSONIAN CONTRIBUTIONS TO Pachystrophia □ devexa (see Figure 13) d □ achystrophia contigua Holopea insignis Holopea pyrene l—. Holopea I—I rotunda Holopea symmetrica Pachystrophia □ spiralis Sinutropis lesthetica Euomphalus tubus Pachystrophia j gotlandica Lytospira □ triquestra Lytospira subuloides NUMBER 88 33 Figure 14 (opposite).—Relationships among the derived “holopeids.” For abbreviations, see legend to Figure 7. Node 62 (“Pachystrophides”), fine, sharp GL (15); PB lost (19); ACh lost (45); p = 60° (48); no ontogenetic rotation of aperture (49); very narrow aperture (58, 59); RRC lost (64); convex LR (72); BC lost (89); thickened PI (103). Node 63 ( Pachystrophia contigua clade), asymmetric aperture (left side broader) (57-59); IM 15° off parallel to CA (98); whole aperture inclined -20° (109, 110); moderate K (123). Node 64 (Holopea insignis clade), sinus lost (1); asymmetric ramp lengths (RR con¬ tracted) (54-56); broad left side of aperture (59); lM\base angle = 60° (94); moderate T (126). Node 65, P = 40° (48); symmetric ramp shapes (51-53); very short RR (55); moderately wide right side of aperture (58); very convex LR (72); thin PI (103); whole aperture inclined -30° (109, 110); base projected posteriorly -10° (117); straight base (120); moderate E (121); high T (126); ontogenetic increase in T (127); septation absent (128). Node 66 ( Sinulropis ?esthetica clade), asymmetric sinus shape (left curve stronger) (8-10); p = 30° (48); asymmetric ramp shapes (LR rounder than RR) (51, 52, 72); long ramps (55, 56); asymmetric aperture (RR wider than LR) (57-59); base projected pos¬ teriorly -20° (117). Node 67, asymmetric ramp lengths (LR longer than RR) (54—56); very convex LR (72). Node 68 (Pachystrophia gotlandica clade), IM channel present (102). Node 69, sinus angle = 20° (3, 4); narrow sinus (6, 7); weak GL (15); p = 20° (48); moderately long ramps (55, 56); moderately wide aperture (58, 59); IM 1 to CA (98); base projected posteriorly -10° (116, 117); open coiling (123); low ultradextral T (126). was much like E. cariniferus but with a frill-like left carina, slightly imbricated growth lines, ontogenetic weakening of the right carina and peripheral band, and strong increases in whorl convexity over ontogeny. A calcitic shell also diagnoses this clade, although again I did not use this as a character state. In the Euomphalopterus subcarinatus clade (Figure 17, nodes 88-92), the frill becomes strongly developed (see also Linsley et al., 1978). This frill is highly crenulated on species such as E. praetextus (Lindstrom, 1884) or replaced with a series of tubes on others such as E. togatus (Lindstrom, 1884). The lat¬ ter feature is best developed in Spinicharybdis, although it is possible that this represents a parallelism. Rohr and Packard (1982) commented on similarities between Spinicharybdis and Euomphalopterus, but they did not explicitly suggest that the two genera were related. The elongate, widely spaced tubes of S. wilsoni Rohr and Packard, 1982, are more similar to those seen on species of Hystricoceras Jahn than they are to the short, tightly-spaced tubes of E. togatus. Yochelson (1966b) cited this as evidence that the species since classified as Spini¬ charybdis are related to Hystricoceras. Spinicharybdis wilsoni shares other synapomorphies with E. togatus, however, includ¬ ing the shape of the ramps, the development of the sinus, and the complete absence of a peripheral band. There are other species of Spinicharybdis that I could not include in this analy¬ sis because the only known specimens are too incomplete. These species do show a series of tubes that are very similar to the tube-bearing frill of E. togatus, save that they are much longer; however, there is no evidence of frill-bounding lira on any of these species, which exist on E. togatus and its rela¬ tives. Only the bases and lower whorls are visible on any of the pertinent specimens, so it is not known if the sinuses and pe¬ ripheral bands of these specimens are like those of Euompha¬ lopterus or like those of Hystricoceras. Therefore, it is con¬ ceivable that Spinicharybdis actually is a “pseudophoride” (see below). Finally, Spinicharybdis also shares features with contempo¬ rary Straparollus (e.g., S. paveyi Foerste, 1924), and the spac¬ ing of the tubes on S. wilsoni is similar to the spacing of carrier shell scars of S. paveyi (see further discussion, below). The long tubes of Spinicharybdis might have served the same func¬ tional purpose as the agglutinated shells on Straparollus spe¬ cies (e.g., Linsley and Yochelson, 1973). Therefore, another possibility is that the spines of Spinicharybdis represent a mor¬ phologic novelty that maintained a functional “homology” with carrier-shell ancestors such as S. paveyi. In this case, the more frill-like tubes of other species represent convergence to¬ ward a Euomphalopterus- like morphology. As a similar func¬ tional interpretation applies to the extended frill of some Eu¬ omphalopterus species (but not to the short tubular frill of species such as E. togatus ; e.g., Linsley et al., 1978), however, it seems as or more likely that the spines and frills represent different adaptations on the same homologies that were func¬ tional parallelism. Euomphalopterus apparently is the sister taxon of a Silurian clade that includes Poleumita and Centrifugus (Figure 18, nodes 93-96). The earliest known species from this clade, P. alata (Lindstrom, 1884), retains a strong lower ramp carina, but later species possess a wide, dull swelling in this region. Other synapomorphies include a flat, very shallow sinus, a thickened and rounded inner margin, well-developed ornament, and near planispiral coiling with very low curvature. The diag¬ nosis of Poleumita differs from that of Euomphalus primarily in that Poleumita possesses ornamentation. Excluding orna¬ ment (which is plesiomorphic above node 94), “poleumitide” species, such as E. walmstedti Lindstrom, 1884, lack any obvi¬ ous autapomorphies relative to Euomphalus. Thus, the "po- leumitides” likely represent the “euomphalinae” in the truest sense. As noted above, Late Silurian species classified as Straparol¬ lus possess carrier shell scars. This feature is retained on many Devonian species (e.g., Linsley and Yochelson, 1973) and might be a synapomorphy between these species and early omphalotrochids (pers. obs.). 1.3.2.2.3. “Pseudophorides”.—The final “euomphalopter- ine” subclade (Figure 19, nodes 97-108) includes species as¬ signed to Pseudophorus Meek, 1873, Discordichilus Coss- mann, 1918, Hystricoceras, Siluriphorus Cossmann, 1918, and Streptotrochus Pemer, 1903. Previous workers classified many of these species in the Pseudophoroidea (e.g., Knight et al., 1960), so I refer to this clade as the “pseudophorides.” The clade also includes the Late Ordovician Euomphalopterus lor- dovicius Longstaff, 1924. “Pseudophoride” synapomorphies include a tangential aperture (i.e., the inclination of the entire aperture rather than just portions of the aperture), a thickened inner margin with little projection relative to the coiling axis that fills the umbilicus, and moderately high translation. The clade initially retains the sharp peripheral band and the sharp 34 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY NUMBER 88 35 FIGURE 15 (opposite).—Relationships among the “helicotomids.” For abbrevi¬ ations, see legend to Figure 7. Node 70 (“Helicotomids”), sinus angle = 50° (3, 4); moderately strong, round ML (28, 29); asymmetric ramp lengths (RR nar¬ rower than left) (54-56); asymmetric aperture (right side wider) (57-59); lM\base angle = 90° (94); curved IM (95); base projected posteriorly -20° (117); curved base (120). Node 71, sigmoidal aperture (13); basal GL strength same as rest of shell (18); PB width = 10° (20); PB 1 to aperture (42); strong ACh (46); P = 60° (48); symmetric ramp shapes (51,52, 72); moderately broad (slightly asymmetric) aperture (58, 59); loss of RR swelling (60); sharp carina on RR and LR (64, 75); IM thickness same as rest of shell (87); IM\base angle ~ 75° (94); IM channel lost (102); base projected anteriorly -10° (116, 118, 119); low E (121); moderate T (126). Node 72 ( Polehemia taneyensis clade), ornament throughout left side of aperture (129). Node 73 ( Boucolspira aff. B. fimbriata clade), sinus angle = 30° (3, 4); weak GL (15); V-shaped lunulae (38); p = 50° (48); no ontogenetic rotation of aperture (49); left and right widths of aperture symmetrical (57-59); base projected anteriorly -20° (119); isometric T (127). Node 74 (“Ophiletinines”), weak ACh (46); low K. (123); low T (126); septation lost (128). Node 75 (Oriostoma bromidensis clade), nar¬ row sinus (6, 7); strong GL (15); projecting BC (91); base projected anteriorly -10° (119); small size (141). Node 76, BL present (21); concentric lunulae (38); very strong lunulae (39); P = 50° (48); highly asymmetric ramps (LR nearly twice as long as RR) (55, 56); slightly asymmetric aperture (left side contracted slightly) (57-59); squared ridge-like LRC (76); thickened IM (87); BC beneath outer margin (93); lM\base angle = 105° (94); inclined aperture (109). Node 77 ( Helicotoma tennesseensis clade), weak RRC (65); LR swell¬ ing present with weak LRC (73, 77); dull, thickened BC (90). Node 78 ( Palae- omphalus Igradatus clade), PB width = 05° (20); BC lost (89); strongly curved IM (95). Node 79 ( Helicotoma planulata clade), asymmetric aperture (LR moderately contracted) (57-59); thickened IM (87); ontogenetic increase in T (127); ornate LR (129). Node 80, ACh lost (45); asymmetric ramp lengths (LR strongly contracted) (54-56); moderately strong RRC (65); strong, squared ridge-like LRC (76, 77). right and left carinae that are common to plesiomorphic “eu- omphalopterines.” The left carina, however, becomes promi¬ nent with a square periphery, whereas the right carina and the peripheral band are strongly reduced or lost in the Siluriphorus gotlandicus clade (Figure 19, node 104). Also, the sinus is re¬ duced to a shallow kink near the suture, and the sigmoidal shape of the aperture becomes extreme. Among very derived members of the Discordichilus clade (Figure 19, node 106), the peg-like left carina is very weak and obtuse, whereas the ramps become strongly rounded. Among other members of the Pseudophorus clade, the peg-like carina becomes a hood-like frill (Figure 19, node 105). A second “pseudophoride” subclade (Figure 19, node 101) includes species assigned to Streptotrochus and Hystricoceras. This clade’s synapomorphies include an inner margin that re¬ flects around the coiling axis and that is thickened at the top and bottom, a thin parietal inductura, and notable increases in both shell expansion and shell torque over ontogeny. The most derived species of this clade possess a projected, strongly chan¬ neled left carina (e.g., S. lundgreni (Lindstrom, 1884)), which forms a series of closely connected tubes on Hystricoceras. Knight et al. (1960) considered Raphistomina to be the earli¬ est member of the Pseudophoroidea, based on the assumption that the peripheral band on Raphistomina species is homolo¬ gous with the strong lower ramp carina or frill on species of Sil¬ uriphorus and Pseudophorus. Previous workers had interpreted this band as a peripheral band because it lies in the middle of a prominent sinus (e.g., Ulrich and Scofield, 1897; Wenz, 1938). I follow the latter interpretation, which leaves Raphistomina species without any important “pseudophoride” synapomor¬ phies; therefore, this analysis contradicts the relationships im¬ plied by the taxonomy of Knight et al. (1960). Knight et al. also considered Trochomphalus Koken to be a pseudophoroid. These results agree better with that idea, but they suggest that Trochomphalus is a member of the “anomphalide” clade. Knight et al. (1960) assigned both Streptotrochus and Dis¬ cordichilus to the Microdomatoidea. This analysis supports a close relationship between these two genera, but it also implies that they have closer relatives than each other among the Pseudophoroidea. The analysis includes only one other puta¬ tive microdomatoid, Daidia Wilson, 1951, but species belong¬ ing to that genus are considered to be “murchisoniinae” and not at all closely related to the “pseudophorides” (see the “strap- arollinoids” below) N.J. Morris and Cleevely (1981) previously linked the Pseudophoroidea to the Euomphaloidea. Morris and Cleevely, however, thought that the Pseudophoroidea diverged from those species very early, i.e., prior to the divergence of taxa such as Ophiletina and Helicotoma. This analysis suggests that “pseudophorides” actually are highly derived “euomphali- nae,” evolved within the “helicotomids.” N.J. Morris and Cleevely (1981) also suggested that another taxon, the Trocho- nematoidea, are even more closely related to the Euompha¬ loidea than are the Pseudophoroidea. My analyses, however, suggest that the Trochonematoidea evolved from “lophos- piroids” (Wagner, 1995a; see also Ulrich and Scofield, 1897; Knight et al., 1960). As discussed below, lophospirids appar¬ ently evolved from the “murchisoniinae,” so this study does not support that part of Morris and Cleevely's phylogenetic scheme. II. “Murchisoniinaes” The “Murchisoniinae” contain several important taxa, in¬ cluding the earliest putative apogastropods and most of the early Paleozoic taxa assigned to the Pleurotomarioidea (Fig¬ ures 20-35, nodes 109-215). The relationships among the ma¬ jor “murchisoniinae” subclades (Figure 20, node 109) ap¬ proaches a star phylogeny (sensu Felsenstein, 1985), with several taxa derived from the same ancestor. Although system- atists usually interpret such polytomies as unresolved relation¬ ships, the analysis suggests that the polytomy represents the real relationship among these taxa. The polytomy includes Hormotoma Isimulatrix (Billings, 1865), which is plesiomor¬ phic relative to all “murchisoniinae” subclades. This suggests that the long-lived and geographically wide ranging H. Isimul- atrix is ancestral to these clades through separate cladogenetic events (see Hoelzer and Melnick, 1994; Wagner and Erwin, 1995). The earliest members of the major subclades all co-oc¬ cur with H. Isimulatrix, and it often was difficult for me to de- 36 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Family Euomphalopteridae Euomphalidae Anomphalidae Holopeidae Planitrochidae Equivocal Euomphalopterus cariniferus (see Figure 15) Euomphalopterus H lordovicius (go to Node 97) Euomphalopterus □ subcarinatus (go to Node 88) Figure 15 Node 74 y CI: 0.697 RI: 0.743 Poleumita Kj alata (go to Node 93) Trochomphalus W Idimidiatus Straporillina IQ cf. S. circe ( (sensu Rohr 1988) Grantlandispira Si christei FIGURE 16.—Relationships among the “anompha- T 85 I jr lides.” For abbreviations, see legend to Figure 7. Node jm&r 81 (“Euomphalopterines”), sinus angle = 20° (3, 4); PB width = 05° (20); P = 40° (48); broad symmetric aperture (58, 59); strong, peg-like frill LRC with channel (76, 78); round \P QjL A thread-like BC becoming weaker over ontogeny (90, 92); IM\base JB&r angle = 105° (94); PI thickness same as rest of shell (103). Node 82 (“Poleumitides”), sinus angle = 10° (3, 4); frilled LRC (78). Node 83 \8 7) (“Anomphalides”), sinus angle = 30° (3,4); rounded ML (28); weak ML (29); concentric lunulae (38); convex RR (52); loss of LRC channel (78); BC lost (89); IM\base angle = 75° (94); IM 30° off parallel to CA (98); strong columellar lira present (100). Node 84, RRC lost (64); convex LR (72). Node 85, PB lost (19); ACh lost (45); (3 = 30° (48); highly asymmetric ramp lengths (LR very long, RR extremely short) (55, 56); very broad aperture (58, 59); callous-like columellar lira (100, 101). Node 86, sinus angle = 10°; strongly curved IM (95). Node 87, LRC lost (75). Pycnomphalus □ acutus o Pycnomphalus [~| obesus Turbocheilus I immaturum NUMBER 88 37 Cl: 0.695 RI: 0.646 Family Euomphalopteridae 0 Euomphalidae Uncertain Figure 16 Node 81 Figure 17.—Relationships among the “poleumiti- des,” part 1. For abbreviations, see legend to Figure 7. Node 82, sinus angle = 10° (3, 4); frilled LRC (78). Node 88 {Euomphalopterus subcarinatus clade), weakly imbricated GL (17); PB fades over ontogeny (30); RR becoming much more con¬ vex over ontogeny (53); asymmetric aperture (right side broader) (57-59); RRC becoming weaker over ontogeny (67); very strong, chan¬ neled frill (LRC) (77, 79, 82); very low E and high K (121, 123). Node 89 (Euomphalopterus togatus clade), dull, lump-like ML (28); right half of aper¬ ture very broad (58); dull, thickened BC beneath IM (90, 93). Node 90, ACh lost (45); very convex RR becoming flatter through ontogeny (52, 53); crenulated LRC frill (81). Node 91 ( Euomphalopterus praetextus clade), U-shaped sinus (8, 9); fine, sharp, nonimbricated GL (15, 17); PB lost (19); concentric lunulae (38); strongly curved IM (95). Node 92 ( Euomphalopterus frenatus clade), right side of aperture extremely broad (58); LRC frill forming tubes (85); high T (126). 13 JS- 13 ■¥ £ a S- a a_ a. a S 13 $ Euomphalopterus □ subcarinatus (see Figure 16) Euomphalopterus alatus Euomphalopterus □ togatus Euomphalopterus praetextus Euomphalopterus undulans Euomphalopterus n frenatus Spinicharybdis wilsoni Poleumita alata (go to Node 93) 38 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Cl: 0.643 RI: 0.637 Figure 16 Node 81 Family Euomphalidae mSM Euomphalopteridae Euomphalopterus 3 subcarinatus (see Figures 16,17 Poleumita alata (see Figures 16,17) Poleumita discors Poleumita granulosa Centrifugus planorbis Poleumita rugosa Poleumita octavia Euomphalus walmstedti Straparollus bohemicus NUMBER 88 39 FIGURE 18 (opposite).—Relationships among the “poleumitides,” part 2. For abbreviations, see legend to Figure 7. Node 82, sinus angle = 10° (3, 4); frilled LRC (78). Node 93 (Poleumita alata clade), sinus nearly absent (3, 4); crenu- lated aperture (12); highly sigmoidal aperture (14); IM\base angle = 90° (94); straight IM (95); thickened middle of IM (96); 1M 30° off parallel to CA (98); very thick PI in concentrated strip, projecting in front of aperture (103, 104); strongly curved base with slight posterior projection (116, 118, 120); moder¬ ately dense ornament on LR and RR (129, 130, 133, 134). Node 94 (Poleumita discors clade), periodically flared ML (32); weak ACh (46); (3=10° (48); long LR (56); weak RRC (65); thick contusion LRC (76); thickened IM (87); BC lost (89); IM\base angle = 75° (94); low T (126); large size (141). Node 95 (Poleumita rugosa clade), extremely strong GL (15); strongly imbricated GL (17); convex RR (52); dull, lump-like LRC (66); convex LR (72); moderate E (121); denser ornament on LR (130). Node 96 (Euomphalus marix clade), non- imbricated GL (17); convex LR (72); LRC lost (75); uniform thickness of IM (96); IM at -45° to CA (98); base projected posteriorly -10° (116, 117); straight base (120); loss of ornament (129, 133). termine whether a specimen should be classified as H. Isimula- trix or as another Hormotoma species (e.g., H. confusa Cullison, 1944, or H. dubia Cullison, 1944). Hormotoma con¬ fusa and H. dubia, however, share synapomorphies with differ¬ ent “murchisoniinae” subclades, so I treated them as separate species based on the “phylogenetic species” concept (de Queiroz and Donoghue, 1988; Nixon and Wheeler, 1990). II. 1. “Plethospiroids” One early “murchisoniinae” subclade includes species classi¬ fied in the Plethospiridae (Figure 20, nodes 110, 111). “Plethos- piroid” synapomorphies include a shallow sinus and a thick¬ ened, siphonate inner margin. This morphologically novel clade appears to include no members younger than the Early Ordovician, even though Knight et al. (1960) assigned several other genera to the Plethospiridae. Seelya ventricosa Ulrich in Ulrich and Scofield, 1897, belongs to the “plethospiroids,” but none of the other Ordovician or Silurian species assigned to Seelya Ulrich in Ulrich and Scofield belong to the clade. An¬ other putative plethospirid, Diplozone Pemer, 1907, appears to be related to Loxonema Phillips (see below). Erwin (1992) sug¬ gested that the Plethospiridae include the sister taxa of the apo- gastropod-like subulitoids. This analysis suggests that the two clades are closely related, but that other “murchisoniinae” likely are more closely related to subulitids. This is discussed further, below. II.2. “Straparollinoids” Hormotoma dubia represents the stem-member of a moder¬ ately diverse “murchisoniinae” subclade that includes species classified as Daidia Salter, Haplospira Koken, 1897, and Straparollina Billings, 1865 (Figure 21, nodes 112-118). Pre¬ vious workers assigned these taxa to the Holopeidae and the Microdomatidae, which in turn were classified in the Tro- choidea (Knight et al., 1960; Tracey et al., 1993). As noted above, the present definitions of those families (and the Paleo¬ zoic trochoids) is highly polyphyletic, and it is not clear which of these species, if any, represent the precursors of true micro- domatids or trochoids. Accordingly, I label the clade the “strap¬ arollinoids,” which is appropriate given that species assigned to Straparollina represent some of the least derived members of the clade. “Straparollinoid” synapomorphies include a dull monolin- eate peripheral band (which is bilineate on the juvenile whorls of early species and becomes monolineate with age) and a nar¬ row sinus. Slightly more-derived “straparollinoids,” such as Lophospira grandis Butts, 1926 (nodes 114-118), share a very narrow, sharp peripheral band, a nonreflected inner margin, a well-developed left carina, and higher shell torque. Even more- derived species (e.g., Straparollina pelagica Billings, 1865, and more-derived species, nodes 115 and above) possess very narrow sinuses and reduced right sides of the aperture. The most derived “straparollinoids” (e.g., Daidia and Haplospira species, nodes 116, 117) have a weak monolineate peripheral band near attenuated sutures, no sinus, and an inner margin that is entirely contiguous with the previous whorl. Previous workers (e.g., Knight et al., 1960; Erwin, 1988; Tracey et al., 1993) classified Daidia in the Microdomatidae, but the next oldest microdomatids first appear in the Devonian (Blodgett and Johnson, 1992; Blodgett, 1993). The statistical significance of this gap is difficult to assess without data for those Devonian species. It seems unlikely that Daidia is closely related to those species. The “pseudophoride” clade in¬ cludes Silurian species assigned to the Microdomatoidea (al¬ beit, to the Elasmonematidae). Overall, it appears more likely that true microdomatoids arose in that clade rather than from “straparollinoids.” II.3. “Hormotomoids” Hormotoma confusa Cullison, 1944, is the least derived member of a clade that includes most of the species classified in the Murchisonioidea, Subulitoidea, and Loxonematoidea (Figure 8, node 7; Figures 22-26, nodes 119-154). The earliest species in this clade have been classified in the genus Hormo¬ toma by numerous authors, so I refer to the clade as the “hor¬ motomoids.” Note that the definition of this clade is much broader than that of the Hormotominae sensu Wenz (1938). The chief synapomorphy uniting the clade is an asymmetrical sinus that is much broader and much sharper on the left side than on the right. The “hormotomoids” represent the earliest species in which the left side of the aperture is more pro¬ nounced than the right side, which is a feature associated with reduction of the right organs. II.3.1. “SUBULITIDS” The Subulitoidea represent one of the most poorly under¬ stood but intriguing products of early gastropod evolution. The earliest subulitoids appeared in the Arenig (Early Ordovician), 40 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Cl: 0.582 RI: 0.733 Figure 16 Node 81 Family Euomphalopteridae Elasmonematidae Pseudophoridae ? R 0 > o I s a “t. -t ■8 o S' r Co 5 Co o Co n O S. . a s~ . s: <>> £ 05 R §■ 05 05 R Co £ 05 S- S' o s R Co Euomphalopterus I I lordovicius (see Figure 16) Streptotrochus &«! lamellosus _ Streptotrochus ? I—I visbeyensis _ Streptotrochus I_I incisus Streptotrochus I lundgreni Hystricoceras astraciformis _ Streptotrochus I—I aff. S. incisus _ Siluriphorus I—I gotlandicus Siluriphorus undulans Discordichilus ! da lli __ Discordichilus. I—I mollis ._ Discordichilus I—I kolmodini Pseudotectus comes I—| Pseudophorus '—' stuxbergi I Pseudophorus ' profundus NUMBER 88 41 FIGURE 19 (opposite).—Relationships among the “pseudophorides.” For abbreviations, see legend to Figure 7. Node 97 (“Pseudophorides”), funicle present (107); base projected anteriorly -30° (119); high K. (123). Node 98, asymmetric ramp shapes (RR more convex) (52, 53, 72); strong, sharp LRC (76); BC lost (89); IM -15° off parallel to CA (98); aperture inclined -20° (110); high T (126). Node 99 (Streptotrochus? visbeyensis clade), moder¬ ately strong LRC (77); variable RRC (weak to absent) (63, 64). Node 100, straight lunulae (38); (3 = 30°-40° (48); asymmetric aperture (RR wider than LR) (57-59); RRC lost (64). Node 101 (Streptotrochus incisus clade), 1M thicker at top and bottom and reflected around umbilicus (96, 106); thin, complete PI (103); ontogenetically increasing E and T (122, 127). Node 102, convex RR and LR (52, 72); moderately asymmetric aperture (right side extremely broad, left side broad) (58, 59); very strong LRC (77). Node 103 ( Streptotrochus aff. 5. incisus clade), PB lost (19); little ontogenetic change in RR convexity (53); right side of aperture inclined (113). Node 104, highly sigmoidal aperture (14); kinked lunulae (38); acute suture (69, 70); strong, squared LRC (76, 77); PI projecting in front of aperture (104); funicle present (107); base projected anteriorly -50° (119); moderate K (123). Node 105 ( Pseudophorus clade), broad symmetric aperture (57-59); very strong, hood-like LRC frill (76, 77, 79, 84); both IM and base thick¬ ened (88); BC present (89); IM 30° off parallel to CA (98); moderate T (126). Node 106 ( Discordichilus clade), convex RR and LR (52, 72); very asymmetric ramp lengths (RR extremely short, LR long) (55, 56); weak, squared LRC (77); whole aperture inclined -50° (109, 110, 113). Node 107 (Discordichilus dalli clade), fine, sharp GL (15); thin, complete PI (103); small size (141). Node 108 (Discordichilus kolmodini clade), moderately asymmetric aperture (right side extremely broad, left side broad) (58, 59); thick, contuse LRC (76). Hormotoma Isimulatrix (see Figure 8) □ Turritoma aff. T. acrea Hormotoma confusa Turritoma Plethospira Cotter Fm. sp. cannonensis (go to Node 112)(go to Node 119)(go to Node 155) Plethospira Seelya cassina ventricosa Figure 20.—Relationships among basal “murchisoniinae.” For abbreviations, see legend to Figure 7. Node 109, weak GL (15); extremely low E and very high T (121, 126). Node 110 (“Plethospiroids”), sinus angle = 30° (3, 4); narrow sinus (6, 7); PB width = 20° (20); P = 70° (48); slightly asymmetric aperture (58, 59); IM\base angle = 60° (94); slightly twisted siphon (99); nonreflected IM filling umbilicus (106, 108); low E (121). Node 111, loss of RR and LR swellings (60, 73); aperture inclined -20° (109, 110); large size (141). 42 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY NUMBER 88 43 Figure 21 (opposite).—Relationships among the “straparollinoids.” For abbre¬ viations, see legend to Figure 7. Node 112 (“Straparollinoids”), increasing prominence of PB over ontogeny with ML developing (27, 40,47); asymmetric ramp shapes (RR flatter) (51-53); symmetric, moderately long ramps (54-56). Node 113 ( Hormotoma dubia clade), narrow sinus (6, 7); PB width = 15° (20); asymmetric aperture (left side broader) (57, 58). Node 114, sinus angle = 30° (3, 4); (3 = 60° (48); asymmetric ramp shapes (LR rounder than RR) (51, 52, 72); very asymmetric aperture breadth (expanded right side, contracted left side) (57-59); loss of RR and LR swellings (60, 73); nonreflected IM (106); high K (123); high T (126). Node 115 ( Straparoltina pelagica clade), sinus angle = 10° (3, 4); PB width = 10° (20); BL lost (21); highly asymmetric ramp lengths (RR extremely narrow, LR moderately long) (54-56); BC present (89); inclined aperture (109). Node 116 ( Haplospira ?nereis clade), sinus lost (1); PB width = 05° (20); nonprominent PB (33); IM\base angle = 75° (94); aper¬ ture contiguous with previous whorl (108); very low E (121); very high K (123). Node 117 ( Daidia clade), concave RR (52); very asymmetric aperture breadth (left side much broader) (58, 59); RRC present (64); attenuated suture (69, 70); very high T (126). Node 118 ( Straparoltina erigione clade), PB lost (19); ACh lost (45); convex RR (52); moderate T (126). with seemingly very “modern” synapomorphies (e.g., a si¬ phon, no sinus or peripheral band, and an elongated aperture). These simplified morphologies leave subulitids with few obvi¬ ous homologies with other gastropods. The distinctive twisted inner margin of the earliest subulitoids, however, which con¬ tributes to a weak siphon, also exists on Early Ordovician spe¬ cies typically classified as Hormotoma (Figure 22, nodes 120-123). The earliest known subulitoid (identified as Hormo¬ toma sp. in Bridge and Cloud, 1947) co-existed with one of these species ( H. zelleri Flower, 1968a), differing “only” in the absences of the peripheral band and the right side of the aper¬ ture. These early, sinus-bearing “subulitids” possess the asym¬ metric sinus and wide, weak, and swollen bilineate peripheral band diagnostic of the H. confusa clade. As noted above, the highly asymmetric aperture and sinus implies that the organs on the right side of the anus were reduced relative to those on the left side. Apogastropod-like species, such as Subulites Em¬ mons, 1842, show this to an extreme, as the right side of the aperture is absent. For features involving the sinus, I did not code asymmetry as homologous among subulitids and other “hormotomoids” because the subulitids lack a sinus. The aper¬ ture of early subulitids, however, does curve backward (i.e., ophisthocyrt) to the suture, which could represent a remnant of the left side of the sinus. Sinus-bearing, siphonate “subulitid” species (nodes 122, 123) were not simply ephemeral intermediates, as some species (e.g., Hormotoma augustina (Billings, 1865), and H. bel- licincta (Hall, 1847) existed into the late Caradoc (Late Ordov¬ ician). This subclade was never diverse, but its species were the dominant gastropod components of many Ordovician fossil as¬ semblages. “Subulitids” represent one of the few cases where the results of this analysis do not agree with any previous interpretation of gastropod phylogeny. Wenz (1938) linked subulitids to the ra- phistomatids, apparently deeming the asymmetric apertures and lack of a slit to be synapomorphies between the two taxa, and the apertural asymmetry of some raphistomatids to be a precursor to an apogastropod-like morphology. This analysis obviously suggests that “scalitines” and “subulitids” are only distant relatives. Others (e.g., Ulrich and Scofield, 1897; Knight et al., 1960) proposed that subulitids evolved from the Loxonematoidea, which many consider to be the earliest apo- gastropods (e.g., Haszprunar, 1988). This analysis suggests that loxonematoids and “subulitids” shared a Hormotoma- like an¬ cestor (see discussion below), but that “subulitids” evolved long before the Loxonematoidea. Notably, this pattern also is consistent with the fossil record. As noted above, Erwin (1992) considered the plethospirids to be the sister taxon of subulitids. The main synapomorphy be¬ tween plethospirids and subulitids is the presence of a siphon. Although I scored the siphon as homologous among “plethos- piroids” and “subulitids,” more synapomorphies exist between “subulitids” and other “hormotomoids” than between “subulit¬ ids” and “plethospiroids.” Therefore, this analysis suggests that “plethospiroid” and “subulitid” siphons are convergent. Finally, this analysis suggests that the traditional definition of the Subulitoidea is polyphyletic, because some Silurian spe¬ cies (i.e., Auriptygma Perner, 1903, and Macrochilus Lind- strom, 1884) appear to have evolved from “hormotomoid” sub¬ codes other than “subulitids.” These species are discussed further below. II.3.2. “CYRTOSTROPHIDS” The sister taxon of “subulitids” (Figures 23-26, nodes 124-154) is diagnosed by an extremely asymmetric sinus, with the left side being much deeper and much more strongly curved than the right side. Most of these species also have been classi¬ fied in Hormotoma. I use that name to designate the more in¬ clusive clade, so I refer to the subclade as “cyrtostrophids.” Cyrtostropha Donald, 1912, is a Silurian genus that has not previously been used to label a higher taxon; Knight et al. (1960) even considered the genus to be a junior synonym of Murchisonia d’Archaic and de Verneuil in d’Archaic. Cyr¬ tostropha retains many of the features that are plesiomorphic to node 131, so it is the most appropriate label available. Addi¬ tional “cyrtostrophids” synapomorphies include a wide periph¬ eral band with thin sharp lira, a round, relatively symmetric ap¬ erture, and a strongly reflected, L-shaped inner margin. Note that the inner margin is distinctly nonsiphonate. Two major “cyrtostrophid” subclades evolved by the Middle Ordovician. These are discussed below. Smaller subclades ap¬ peared in the Late Ordovician and Early Silurian. One of these (Figure 23, node 126) includes Cyrtostropha coralli (Sowerby in Murchison, 1839) and is diagnosed by a sharp lirum in the middle of the right ramp. The second (Figure 23, nodes 127-130) includes species classified as Hormotoma and Cata- zone Perner. This clade is diagnosed by strongly asymmetrical ramp shapes and lengths (with the right ramp being flatter and much longer), somewhat reduced shell torque, and the periph- 44 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY o 2 Sv. ■—» O ^ a: a a, c c« S g o £ S - 3 >, (D "L GO a a o .S o 5 | $> o a >3 a □ 2 ,| 2 C3 3 1 2 -C5 2 CO 2 • >»*» & a * Kl 2 t*3 C cS w a 2 05 05 I O 00 oo r- in CO U & NUMBER 88 45 a 5 a o o =C Ov cu "O C O a z | 2 o W) □ □ □ □ □ □ 2 §■ Cl £ O in oo O CM «n in o d 46 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY eral band being very low on the whorl. A more-derived sub- clade ( Catazone\ nodes 129, 130) share a reduced peripheral band, a slit, and an inner margin that is contiguous with the pre¬ vious whorl rather than reflected. 11.3.2.1. “Goniostrophines”. —One of the major “cyr- tostrophid” subclades includes species classified in Murchiso¬ nia, Hormotoma, Goniostropha Oehlert, and Sinuspira Pemer (Figure 24, nodes 131-138). The last genus is highly derived and not typical of the clade, so I label this clade the “goniostro¬ phines.” Synapomorphies of the clade include an asymmetric whorl shape, with a rounded left side and a flat or concave right side that culminates in an acute, attenuated suture. The clade is composed primarily of Silurian species. One Silurian subclade, the H. attenuata clade (node 138), is diagnosed by the loss of the peripheral lira that are primitive to “murchisoniinae.” Spe¬ cies in another Silurian clade (the H. subplicata clade, nodes 132-137) possess very strong, sharp peripheral band lira, flat ramps, a more symmetrical shallower sinus, an acute suture, and an acute basal portion of the inner margin. The last feature becomes somewhat siphonate in the Donaldiella declivis clade (node 136), and a slit is present just under the suture on those species. The definition of Murchisonia given by both Knight et al. (1960) and Wenz (1938) includes Goniostropha. The type spe¬ cies of Murchisonia, M. bilineata (Dechen in De la Beche, 1832) is known from the Middle Devonian, so it could not be included here. Murchisonia bilineata possesses Goniostropha synapomorphies, such as a wide peripheral band with strong lira and a reduced sinus, so extended analyses probably will support the shared opinion of Wenz and Knight et al. The latter authors also included Cyrtostropha within Murchisonia, but these results suggest that Cyrtostropha evolved independently from Hormotoma. “Goniostrophines” (= Hormotoma salteri clade) include the only early Paleozoic species known to have had distinct proto¬ conch morphologies. Unfortunately, the high-spired “cyrtostro- phids” rarely preserve the apex. At least two species in the Hormotoma salteri clade ( Sinuspira tenera Barrande in Pemer, 1907, and Donaldiella declivis (Barrande in Pemer, 1907)) possess large, planispiral protoconchs; however, an interesting implication of this analysis is that this distinctive protoconch morphology arose among some “cyrtostrophids” by the Middle Ordovician. 11.3.2.2. “OMOSPIRINES”. —The earliest members of the other major “cyrtostrophid” subclade (Figures 25, 26, nodes 139-154) include species classified in Omospira Ulrich in Ul¬ rich and Scofield, so I refer to the clade as the “omospirines.” “Omospirine” synapomorphies include increased shell expan¬ sion, reduced reflection and thickness of the inner margin, and a more symmetrical sinus. Omospira itself (node 139) is linked by a weaker peripheral band that is closer to the suture, a U-shaped sinus, and increased left to right asymmetry of the aperture. Most previous authors considered Omospira as an aberrant raphistomatid (e.g., Knight et al., 1960), but this analysis cor¬ roborates Wenz’s (1938) classification of Omospira as a murchisonioid. It also should be noted that the “omospirines” bear little resemblance to the definition of the Omospirinae given by Knight et al. (1960), which included putative Silurian Omospira and post-Silurian genera. This analysis suggests that none of the post-Caradoc (Middle-Late Ordovician) members of this clade retained Omospira- like morphologies. Instead, the other "omospirine” species closely match traditional defini¬ tions of the Loxonematoidea. Most workers (e.g., Ulrich and Scofield, 1897; Koken, 1898; Wenz, 1938; Knight et al., 1960; Erwin, 1990b) considered loxonematoids to be derived murchisonioids. This analysis corroborates that view, with the caveat that Omospira- like “murchisoniinae” species are inter¬ mediate between Loxonema and traditional murchisonioid spe¬ cies, such as Hormotoma gracilis (Hall, 1847). Two “omospirine” subclades evolved during the Ordovician. One of these includes Loxonema, so I designate it the “loxone- matides.” This clade is discussed below. The sister clade (Fig¬ ure 26, nodes 146-154) of the “loxonematides” is diagnosed by a very weak peripheral band and weak ornament. Previous workers placed these species in the Loxonematidae (e.g., Donald, 1905; Wenz, 1938; Knight et al., 1960), and this analy¬ sis suggests that they form a sister clade to the one including Loxonema. The genus Rhabdostropha Donald, 1905, appears to be typical of the group, so I refer to this clade as the “rhab- dostrophides” for lack of a more appropriate name. Synapomorphies of the “loxonematides” (Figure 25, nodes 142-145) include the loss of the peripheral band, a thick induc- tura, and a slightly flaring aperture. In the Loxonema berault- ensis clade (Figure 25, node 145), the sinus becomes deeper and culminates close to the suture. This culminates in Diploz- one crispa (Lindstrom, 1884), which has a siphonate base and a nearly slit-like sinus just under the suture. The “rhabdostrophides” are diagnosed by an inner margin that is contiguous with previous whorls, anterior production of FIGURE 24 (opposite).—Relationships among the "goniostrophines.” For abbreviations, see legend to Figure 7. Node 125 ( Hormotoma gracilis clade), asymmetric sinus shape (left curve stronger) (8-10); (3 = 70° (48); moderately asymmetric ramp lengths (right side long, left side short) (55, 56). Node 131 (“Goniostrophines”), dull, wide ML present between BL (27); attenuated suture (69, 70). Node 132 ( Hormotoma subplicata clade), sharp BL (22); slightly asymmetric aperture (right side broad, left side moderately broad) (58, 59). Node 133, fine, sharp GL (15); ML lost (27). Node 134 ( Goniostropha cava clade), thin, incomplete PI (103). Node 135, left sinus angle = 40° (4); P = 90° (48); long, asymmetric ramps (RR moderately long, LR long) (55, 56); flat LR (72). Node 136 ( Murchisonia paradoxa clade), PB width = 10° (20); slit present (34); nonattenuated acute suture (69, 70); strong, sharp LRC present (75-77). Node 137, very asymmetric ramp lengths (RR short, LR very long) (56, 57); very asymmetric aperture (right side extremely broad, left side nar¬ row) (58, 59); IM\base angle = 60° (94). Node 138 ( Hormotoma attenuata clade), fine, sharp GL (15); PB width = 10° (20); BL lost (21); IM\base angle = 60° (94); moderate size (141). 48 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Hormotoma Omospira Omospira Rhabdostropha Loxonema Loxonema Loxonema Loxonema Diplozone trentonensis alexandra laticincta primitiva murrayana crossmanni sinuosa beraultensis crispa (see Figure 23) (go to Node 146) □ □■■□□□■a FIGURE 25.—Relationships among the “omospirines” and “loxonematides.” For abbreviations, see legend to Fig¬ ure 7. Node 139 (“Omospirines”), left sinus angle -40° (4); nonreflected IM (106); very high T increasing over ontogeny (126, 127). Node 140 ( Omospira clade), (3 = 50° (48); very asymmetric ramp lengths (RR very short, LR long) (55, 56); large size (141). Node 141 ( Rhabdostropha primitiva clade), left sinus begins at base of LR (7); nonthickened IM (87). Node 142 (“Loxonematides”), PB lost (19); very asymmetric ramp lengths (RR very short, LR very long) (55, 56); broad (asymmetric) aperture (58, 59); acute suture (69); nonthickened IM (87); arched IM (95); PI thickness same as rest of shell (103); weakly flaring aperture (105); very low E (121). Node 143, left side of sinus beginning near base of LR (7); fine, sharp GL (15); IM 15° off parallel to CA (98). Node 144 ( Loxonema sinuosa clade), sinus angle = 30° (3, 4); IM fills umbilicus (108). Node 145 ( Loxonema beraultensis clade), sinus angle = 40° (3,4); symmetric sinus width (5-7); attenuated suture (69, 70); strongly curved IM (95); IM contigu¬ ous with previous whorl (108); inclined aperture (109). FIGURE 26 (opposite).—Relationships among the “rhabdostrophides.” For abbreviations, see legend to Figure 7. Node 146 (“Rhabdostrophides”), nearly symmetric sinus angles = 20° (3, 4); very weak BL (23); aperture contiguous with previous whorl (108); isometric T (127); sparse, fine ornament present (129-133). Node 147 ( Girvania excavata clade), symmetric U-shaped sinus (5-7, 9, 10); PB width = 05° (20); dense ornament (130, 134). Node 148, very shallow sinus angles = 20° (3, 4); fine, sharp GLs (15); loss of RR ornament (133). Node 149 (Holopella regularis clade), sinus angle = 10° (3,4); U-shaped sinus (9, 10). Node 150 (Kjerulfonema clade), sinus nearly absent (3, 4); thick¬ ened IM (87); extremely low E (121); strong ornamentation (129-137). Node 151 ( Macrochilus buliminus clade), sinus nearly absent (3, 4); P = 30° (48); highly asymmetric ramp lengths (RR highly contracted) (55, 56); thickened middle of IM (96); PI absent (103). Node 152, extremely asymmetric ramp lengths (RR extremely contracted) (55, 56); IM || to CA (98). Node 153, sinus lost (1); IM\base angle = 45° (94); straight IM (95); IM reflected around umbili¬ cus (106); open umbilicus (108). Node 154 ( Stylonema clade), symmetric sinus width (5-7); P = 70° (48); slightly asymmetric, very broad aperture (very broad right side, broad left side) (58, 59); RR swelling present (60); swollen base of LR (73); thin, incomplete PI (103); IM contiguous with previous whorl (108); curved base (120); ornament throughout left side of aperture (129). 50 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY the aperture (e.g., Figure 3 d), and well-developed ornamenta¬ tion. Early species in this clade might retain a very weak bilin- eate peripheral band, but it is not clear whether the feature is a peripheral band or general ornament. Among more-derived species, such as the Holopella regularis clade (Figure 26, nodes 149-154), the sinus is extremely reduced and broadly U- shaped. Stylonema Pemer, 1907 (Figure 26, node 154) is some¬ what more divergent, possessing a curved base, a much rounder and slightly more asymmetric aperture, and marked swelling near the suture. Homy (1955) discussed the phyloge¬ netic position of Stylonema, but only in general terms relative to Loxonema, so this exact estimate cannot be compared to that assessment. The most noteworthy subdivision within the H. regularis clade is the Macrochilus buliminus subclade (Figure 26, nodes 151- 153). This last subclade possesses a gross mor¬ phology that converges strongly on that of derived “subulitids” (see Figure 22, node 122). Synapomorphies include the near loss of the sinus (which is completely lost in the most derived species), a inner margin that is differentially thickened in the middle, the loss of the parietal inductura, and an extremely asymmetric aperture with the right side strongly reduced. This analysis supports the commonly held view that the Loxonematoidea evolved from the Murchisonioidea (e.g., Wenz, 1938; Knight et al., 1960). I noted above that these re¬ sults contradicted the proposal by Knight et al. (1960) that the Subulitoidea evolved from the Loxonematoidea. This study does suggest, however, that a clade of species classified in the Subulitoidea (i.e., the Macrochilus buliminus clade) did evolve from the “loxonematides.” This corroborates Erwin’s (1992) claim that traditional definitions of the Subulitoidea are polyphyletic. In addition to the general suggestions about loxonematoid re¬ lationships, authors also have discussed more specific relation¬ ships of these species. As noted above, the idea of Knight et al. (1960) that Diplozone evolved among the Plethospiridae is re¬ jected by this study. Their suggestion that both Holopella M’Coy, 1851, and Rhabdostropha are closely related to Lox¬ onema is upheld, although this analysis indicates that the former genera belong to a different subclade than does Lox¬ onema. The suggestion by Peel and Yochelson (1976) that the Silurian genus Kjerulfonema Peel and Yochelson is related to Girvania is supported herein. II.4. “Eotomarioids” Most of the taxa classified in the Pleurotomarioidea by Knight et al. (1960) form a major clade that includes species classified in the Eotomariidae, Gosseletinidae, Lophospiridae, Luciellidae, and Phanerotrematidae (Figures 27-35, nodes 155-215). This clade is similar to Ulrich and Scofield’s defini¬ tion of the Eotomarioidea, although their definition does in¬ clude many “raphistomatids” (see above). “Eotomarioid” syna¬ pomorphies include a peripheral band with stronger, sharper lira and weak channels beneath those lira, a straight inner mar¬ gin, flatter ramps, and the loss of swellings on top of the ramps. The earliest species belonging to this clade fit the general de¬ scription of Turritoma, which corroborates Ulrich and Scofield’s (1897:1013) suggestion that eotomarioids evolved from Turritoma. II.4.1. “LOPHOSPIRIDS” “Eotomarioids” include two major subclades. One of these contains the Lophospiridae plus species assigned to the genera Solenospira Ulrich and Scofield, 1897, and Ectomaria Koken, 1896 (Figure 27, nodes 156-165). “Lophospirid” synapomor¬ phies include flat to slightly concave right and left ramps, a strong lower ramp carina, a deep V-shaped sinus, and sharp threads on a bilineate peripheral band. An early appearing “lo¬ phospirid” subclade that includes several species classified as Solenospira and Ectomaria (node 160 and above) represents the sister taxon of traditional lophospirids. The primary syna- pomorphy of this subclade is an extremely wide peripheral band with very strong peripheral lira. Most workers considered Solenospira to be redundant with Ectomaria (e.g., Wenz, 1938; Knight et al., 1960), a view that this analysis supports. Those authors also considered Ectomaria to be a close relative of Hormotoma and other traditional murchisoniids, which this analysis contradicts. Node 160 and its descendants represents the most diverse “archaeogastropod” clade of the Ordovician, including species classified as Arjamannia Peel, 1975a, Donaldiella Cossmann, 1903, Eunema Salter, 1859, Globonema Wenz, 1938, Gy- ronema Ulrich in Ulrich and Scofield, 1897, Longstaffia Coss¬ mann, 1908, Lophospira Whitfield, 1886, Loxoplocus Fischer, 1885, Pagodospira Grabau, 1922, Proturritella Koken, 1889, Ptychozone Pemer, 1907, Ruedemannia Foerste, 1914, Schizol- opha Ulrich in Ulrich and Scofield, 1897, Trochonema Salter, 1859, and Trochonemella Okulitch, 1935. The primary synapo¬ morphies of lophospirids are a trilineate peripheral band with a Figure 27 (opposite).—Relationships among the “eotomarioids.” For abbrevi¬ ations, see legend to Figure 7. Node 155 (“Eotomarioids”), sharp, moderately strong BL (22, 23); LR swelling lost (73). Node 156, flat ramps (52, 72). Node 157 (“Clathrospirids”), left side of sinus sharper and narrower than right (2-7); very asymmetric ramp lengths (RR very long, LR short) (55, 56); left and right widths of aperture symmetrical (57-59). Node 158 (“Lophospirids”), weak GL (15); P = 70° (48); asymmetric ramp lengths (RR shorter) (54-57); RRC and LRC present (64, 75); both 1M and base thickened (88); isometric T (127). Node 159 (Pagodospira cicelia clade), PB width = 15° (20); ML present (27); ACh present (45); asymmetric ramp shapes (RR more convex) (52, 53, 72). Node 160 ( Ectomaria adelina clade), PB width = 30° (20); extremely strong BL (23). Node 161, sharp, thread-like RRC and LRC (66, 76). Node 162 (Murchisonia callahanensis clade), sinus angle = 30° (3, 4); concave ramps (52, 72); RR becoming more concave over ontogeny (53); very low E (121); ontogenetically increasing expansion (122); high K (123); isometric T (127). Node 163 ( Ectomaria pagoda clade), moderately strong LRC (77). Node 164 (Ectomaria nieszkowskii clade), symmetric ramp lengths (54-56). Node 165 (Ectomaria prisca clade), RRC lost (64). 52 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY pronounced channel. The phylogenetics of this clade are dis¬ cussed elsewhere (Wagner, 1990, 1995a, 1999). II.4.2. “CLATHROSPIRIDS” The second major “eotomarioid” subclade includes the re¬ mainder of the taxa previously classified in the Pleurotomario- idea (Figures 28-35, nodes 166-215). I designate the clade as “clathrospirids” because most of the earliest species have been classified in the genus Clathrospira. “Clathrospirid” synapo- morphies include an extended right ramp, placement of the pe¬ ripheral band partially on the right ramp, and an asymmetrical sinus with a wider, shallower right half. Like the “raphistomatids,” two early “clathrospirid” sub¬ codes existed in different biogeographic realms, with one pri¬ marily North American (Laurentian) and the second primarily Baltic; however, there is substantial biogeographic overlap be¬ tween the two clades by the end of the Ordovician. II.4.2.1. “LIOSPIRINES”. —The Laurentian “clathrospirid” subclade includes species classified as Eotomaria, Liospira Ulrich and Scofield, and Paraliospira Rohr, with the first ge¬ nus representing a paraphylum relative to the latter two (Figure 29, nodes 172-181). I label this clade as the “liospirines,” al¬ though the clade differs greatly from the definition of the Li- ospirinae sensu Knight et al. (1960). “Liospirine” synapomor- phies include the projection of the inner margin at a high angle relative to the coiling axis (e.g., Figure 3c), the presence of a channel under the peripheral band, and a dull carina on top of the right ramp, which fills the suture. Two “liospirine” sub- clades were moderately diverse during the Caradoc and Ash- gill (Middle-Late Ordovician). One includes true Liospira (Figure 29, nodes 174-177). Liospira synapomorphies include a thickened inner margin, very weak threads on the peripheral band, a shallower, more symmetric sinus, isometric translation (in contrast to the increasing translation shown over the ontog¬ eny by more plesiomorphic members of the clade), and the fill¬ ing of the umbilicus by the inner margin and an extended in- ductura. The last features suggest that in life Liospira wore their shells like caps on top of the body rather than carrying them by the aperture. A slit does not diagnose the clade, but it is present on the most derived species. The second “liospirine” clade includes species assigned by Rohr (1980) to Paraliospira (Figure 29, nodes 178-181). Paraliospira synapomorphies include very strong square¬ shaped peripheral band lira, a very shallow right side of the si¬ nus, and a nearly planispiral, widely umbilicate form produced by very low curvature and translation. In addition, the base is projected much further in front of the inner margin. More-de¬ rived species (i.e., the P. mundula clade) have much stronger channels beneath the peripheral band, a channeled carina on top of the right ramp, a dull, channeled left carina, and a more symmetrical sinus. It is noteworthy that this clade includes Li¬ ospira larvata (Salter, 1859) and Eotomaria supracingulata Ulrich and Scofield, 1897, as this substantiates Ulrich and Scofield’s (1897) suggestion that those species are closely re¬ lated to P. mundula (Ulrich and Scofield, 1897), P. angulata (Ulrich and Scofield, 1897), and their allies (which Ulrich and Scofield tentatively placed in Liospira). Ambiguity about character-state polarity affects the exact clade structure within the L. larvata clade. Paraliospira mun¬ dula and P. rugata (Ulrich and Scofield, 1897) share a very thick inner margin that is folded back into the umbilicus to form a funicle. This feature obscures the inner margin and makes it impossible to determine if an inner margin lirum is present or absent. To be more exact, if the feature was present, then it would be obliterated by the umbilicus. If the feature is assumed to be absent, the best reconstruction is to link P. angu¬ lata to the Eotomaria rupestris clade. This reconstruction is supported by the fact that some specimens of P. mundula lack¬ ing funicles (owing either to intraspecific variation or differ¬ ences in taphonomy) lack the lirum, suggesting that the feature is actually absent. This analysis supports the idea that Liospira and allies are closely related to Eotomaria, which was the opinion of Ulrich and Scofield (1897) and Wenz (1938). The analysis strongly differs from the views of Knight et al. (1960) in two ways. First, those authors considered the Liospirinae to be a subdivision of the Raphistomatidae. There are strong similarities between some “raphistomatids” (e.g., Pararaphistoma) and Liospira. In fact, the widely distributed and somewhat variable species P. qualteriata appears to have been classified as Liospira by some authors (e.g., Longstaff, 1924). Only the most derived Liospira species, such as L. micula (Hall in Hall and Whitney, 1862), dis¬ play “lesueurilline” synapomorphies. Linking these species di¬ rectly (i.e., so that L. micula is considered a plesiomorphic member of Liospira or so that L. micula is removed from the “liospirines” to the “lesueurillines”) also invokes statistically significant stratigraphic gaps (see Appendix 3). The second difference between these results and the taxon¬ omy of Knight et al. (1960) is that those authors included post- Ordovician species in the Liospirinae. Although Liospira com¬ monly is reported from Silurian strata (e.g., Peel, 1977), this analysis found no Silurian “liospirines.” This might be due to the poor preservation of the relevant specimens, as several of the important synapomorphies (e.g., peripheral band type and Figure 28 (opposite).—Relationships among basal “clathrospirids.” For abbreviations, see legend to Figure 7. Node 166, PB width =15° (20); slightly prominent PB (33); left side of aperture inclined -20° (111, 112); base pro¬ jected anteriorly -20° (118, 119); moderate E (121); moderate K (123); high T (126). Node 167 ( Clathrospira Itrochiformis clade), [) = 80° (48); both 1M and base thickened (88). Node 168 ( Clathrospira euconica clade), symmetric sinus depth (2), ACh lost (45); asymmetric aperture (RR wider than LR) (57-59); RRC present (64). Node 169 ( Clathrospira injlata clade), sinus angle -60° on left side (4); asymmetric aperture (RR much wider than LR) (57-59); moder¬ ately convex whorls (52, 72). Node 170 (Mourlonia mjoela + Eotomaria con- vexa), narrow PB with moderately strong PL (20, 23); very convex whorls (52, 72); IM 15° off parallel to CA (98); moderate size (141). 54 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 1 Cbthrospira J conica (see Figure 28) - Clathrospira I subconica 1 Eotomaria -I canalifera | Eotomaria supracingulata Paraliospira angulata j—j Paraliospira m undub Paraliospira rugata _ Paraliospira I I aff.F. angulata (sensu Rohr 1988) h Eotomaria ^ rupestris NUMBER 88 55 FIGURE 29 (opposite).—Relationships among the “liospirines.” For abbrevia¬ tions, see legend to Figure 7. Node 171 (“Liospirines”), thick, squared BL (22); thin, complete PI (103). Node 172 ( Eotomaria canalifera clade), left angle of sinus sharper (6-8); asymmetric aperture (RR wider than LR) (57-59); RRC present (64); 1M at -45° to CA (98); moderate T (126). Node 173 ( Eotomaria dryope clade), very asymmetric aperture (contracted left side); weak, round RRC filling suture (65); dull, thick LRC present (75); 1M -15° off parallel to CA (98); left side of aperture inclined -10° (112). Node 174 ( Eotomaria labrosa clade), PB width = 10° (20); moderately strong, sharp BL (23); asymmetric ramp shapes (LR rounder) (51); middle of IM very thick (87, 96); IM extended and folded back into umbilicus (108); base projected anteriorly -30° (119); moderate E (121); decrease in T over ontogeny (127). Node 175 ( Liospira ), symmetric sinus width (5-7); weak GL (15); PB largely on RR (43); ACh lost (45); nearly symmetric, narrow aperture (58, 59); weak RRC (65); no ontogenetic change in RRC strength (67); IM\base angle = 105° (94); left side of aperture inclined -10° (112). Node 176, symmetric sinus cur¬ vature (8-10); extremely weak BL (23); PI projecting in front of aperture (104); funicle present (107); base projected anteriorly -50° (119). Node 177, slit present (34); left and right widths of aperture symmetrical (57-59); uni¬ formly thick IM (87, 96). Node 178 ( Liospira larvata clade), asymmetric sinus shape (left side much sharper) (8-10); strong GL (15); strong ACh (46); P = 60° (48); asymmetric aperture (right side contracted) (57-59); IM\base angle = 105° (94); top and bottom of IM thicker (96); left side of aperture inclined -30° (112); base projected anteriorly -40° (119); low T (126); iso¬ metric T (127); small size (141). Node 179 ( Paraliospira angulata clade), very asymmetric sinus widths (right side narrowed) (5-7); RR becoming more concave over ontogeny (53); very asymmetric ramp lengths (RR long, LR short) (55, 56); RRC rounded and channeled with no ontogenetic change in RRC strength (66-68); LRC present (75); BC present (89); IM projected 30° relative to CA (98). Node 180 (Paraliospira mundula clade), PB width = 20° (20); strong RRC (65); very thick IM (87); IM extended and folded back into umbilicus (108). Node 181 ( Eotomaria rupestris clade), clockwise rotation of aperture over ontogeny (49); convex ramps (52, 72); left side of aperture inclined -10° (112). orientation, and sinus shape) are visible only on well-preserved Liospira. The filling of the umbilicus by the shell produces a dense, easily preserved core, which is unknown from Silurian strata. The Late Silurian species, L. marklandensis McLeam, 1924, however, possesses a right ramp, peripheral band, and slit similar to that of L. micula, with a funicle (i.e., shell mate¬ rial plugging the umbilicus) being a variable feature. There¬ fore, it is possible that the clade survived the end-Ordovician extinction. Baltic “liospirines” are not well known, although two species {Eotomaria notablis Eichenwald, 1859, and Eotomaria rupes¬ tris Koken, 1925) are included herein. These species do not ap¬ pear to have been closely related, suggesting that they represent separate incursions into Baltica by a primarily Laurentian clade. This finding corroborates Koken’s assessment of gastro¬ pod phylogeny, which separated the “ notablis ” group from the “rupestris ” group (Koken, 1925:190-194). Koken, however, derived Stenoloron from the latter group. Although there are possible synapomorphies linking Stenoloron to some Paraliospira species (especially involving the thin but strong bilineate peripheral band), Early Silurian Stenoloron species appear to share more characters with another “clathrospirid” clade (see discussion below). Thus, it appears that a “Stenoloron ” morphology is grossly convergent between two distantly related “clathrospirid” clades. II.4.2.2. “Brachytomariines”. —The Baltic “clathrospir- ids” include most of the Ordovician and Silurian taxa classified in the Gosseletinidae, Luciellidae, and Phanerotrematidae, as well as most of the non-Laurentian taxa classified in the Eoto- mariidae (Figures 28, 30-35, nodes 169, 170, 182-215). The earliest clear members best fit descriptions of the genus Brachytomaria Koken, so I refer to the clade as the “brachy¬ tomariines.” The clade appears to have been derived from Bal¬ tic species classified as Clathrospira, being united by charac¬ ters including convex whorls (especially in comparison with Clathrospira from the Early Ordovician of Laurentia) and a pe¬ ripheral band with only moderately strong lira that bisects the right and left ramps (rather than sloping onto the right ramp as on Clathrospira from the Early Ordovician of Laurentia). The biogeographic affinities of the clade are nearly opposite those of the “liospirines,” with most species known from Baltica and a few incursions into Laurentia. It appears that all Silurian “eotomarioids” are “brachytomariines.” Relationships among the major “brachytomariine” subclades are among the most difficult to assess. The phylogenetic inter¬ pretation accepted herein suggests that the “brachytomariines” began diversifying during the Late Ordovician (i.e., late Cara- doc and Ashgill), even though many of the subclades do not appear in the fossil record until the Silurian. Unfortunately, sampling is very poor for the latest Ordovician and Early Sil¬ urian, which is an interval of importance here. Thus, a greater number than usual of intermediate species probably are ex¬ cluded from the analyses of these species. Consequently, better sampling of the Llandovery (Early Silurian) might easily result in different estimates about basic relationships among “brachy¬ tomariines.” The probable outcome of such changes will be to decrease the apparent intensity of pre-Silurian diversification and to increase the apparent intensity of Early Silurian evolu¬ tion. II.4.2.2.1. “?Palaeoschismides”.—One “brachytomariine” clade is confined almost exclusively to the Ordovician (Figures 30, 31, nodes 188, 189). This clade includes species classified in a potpourri of eotomariid genera. One of the important plesi- omorphic species in the clade, “ Bembexia ” globosa Longstaff, 1924, certainly does not belong to the Devonian genus, and it might have some affinities with Palaeoschisma Longstaff, 1924. Unfortunately, Palaeoschisma is known from only a sin¬ gle specimen of the species P. girvanensis Longstaff, 1924, and that specimen was too incomplete to include in this analysis. Thus, this suggestion is highly tentative at best. Some of these species have been linked with Bembexia Oehlert, 1888; how¬ ever, that genus sensu stricto first appears in the Early Devo¬ nian and those species appear to have affinities with other “eotomarioids.” All the other genus names have been used to label other clades, so I refer to this clade as the “?palaeo- schismides.” Synapomorphies of this clade include a strongly developed carina on the upper ramp and a very asymmetrical 56 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY NUMBER 88 57 FIGURE 30 (opposite).—Relationships among basal “brachytomariines.” For abbreviations, see legend to Figure 7. Node 182 (“Brachytomariines”), sinus angle -30° on right side (3); PB bisects aperture (43); ACh lost (45); arched IM (95); right side of aperture inclined (113); base projected anteriorly -10° (119). Node 183, right sinus angle -20° (3); (3 = 70° (48); asymmetric aperture (RR wider than LR) (57-59); RRC present (64); IM || to CA (98). Node 184 (Euryzone kiari clade), convex RR and LR (52, 72). Node 185 (“Luciellides”), very narrow PB (6, 7); extremely asymmetric ramp lengths (RR extremely long, LR very short) (55, 56); ornament throughout left side of aperture (129); orna¬ ment on RR (133). Node 186 (“Phanerotrematides”), symmetric sinus width (5-7); strong GL (15); symmetric ramp widths (54-56); LRC present (75). Node 187 (“ Longstaffia ” “laquetta ” clade), narrow sinus (6, 7); PB width = 10° (20); (3 = 60° (48); asymmetric ramp shapes (RR flatter) (51); RR shorter than LR (54-56); strong RRC (65); strong LRC (77); moderate E (121); large size (141). aperture with a reduced left side. Some of the more-derived species (node 187) are highly asymmetrical, with the peripheral band much closer to the suture and the inner margin extended and siphon-like. This line of morphologic evolution is particu¬ larly interesting because it converges on the asymmetric mor¬ phologies seen in “plethospiroids,” some “scalitines,” and vari¬ ous “hormotomoids.” II.4.2.2.2. “Phanerotrematides”.—Another major “brachy- tomariine” subclade (Figures 30, 32, nodes 186, 187, 190-196) closely matches the Phanerotrematidae as defined by Knight et al. (1960) and later authors (e.g.. Peel, 1984b). Synapomor- phies of “phanerotrematides” include a narrow, symmetrically shaped sinus and apertures, very strong growth lines, and a prominent carina on the lower ramp. The stem species, all classified as Brachytomaria, are known from the Late Ordovician, but the primary subclades appear in the Silurian. One “phanerotrematide” subclade includes Sil¬ urian species with prominent right ramp and alveozone carinae (Figure 30, node 187), reminiscent of Ordovician Brachy- Clathrospira Euryzone “ Bembexia ” convexa kiari globosa (Figures 30, 34-35) (go to Node 184) (see Figure 30) Clathrospira Eotomaria thraivensis elevata Figure 31.—Relationships among the “?palaeoschismides.” For abbrevia¬ tions, see legend to Figure 7. Node 182 (“Brachytomariines”), sinus angle -30° on right side (3); PB bisects aperture (43); ACh lost (45); arched IM (95); right side of aperture inclined (113); base projected anteriorly -10° (119). Node 183, right sinus angle -20° (3); (3 = 70° (48); asymmetric aperture (RR wider than LR) (57-59); RRC present (64); IM || to CA (98). Node 188 (“?Palaeoschismides”), asymmetric sinus (left side wider) (5); moderately strong BL (23); PB entirely on RR (43); (3 = 60° (48); asymmetric ramp shapes (RR flatter) (51); asymmetric ramp lengths (RR contracted) (54-57). Node 189 (Eotomaria elevata + Clathrospira thraivensis), right sinus angle -35° (3); nonprominent PB (33); concave RR (52); moderately asymmetric aperture (broad right side, narrow left side) (58, 59); sharp, thread-like RRC (66); acute suture (69, 70); LRC present (75); IM\base angle = 75° (94); PI lost (103); IM fills umbilicus (108); small size (141). 58 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY tomaria species. Additional synapomorphies include flatter and wider right ramps, and inclination of the entire aperture. The clade includes species assigned to Promourlonia Longstaff, 1924, which Knight et al. (1960) considered to be a junior syn¬ onym of Mourlonia de Konick, 1883. This analysis implies that the two taxa are not closely related. Another “phanerotrematide” subclade includes species clas¬ sified in Phanerotrema Fischer, 1885, and Ulrichospira Donald, 1905 (Figure 32, nodes 193-196). The primary charac¬ ter linking this clade is the presence of a slit within the periph¬ eral band. The aperture of these species also is more asymmet¬ ric than on Brachytomaria species, which is a feature that becomes very pronounced on some derived Phanerotrema (e.g., node 195). That, coupled with the extended, nearly sipho- nate inner margin, represents another parallel development of the asymmetric morphology discussed above. II.4.2.2.3. “Luciellides”.—Perhaps the most problematic “brachytomariine” subclade includes all of the Silurian species classified in the Luciellidae (Figures 30, 33, nodes 185, 197-201). “Luciellide” species share extremely narrow, very asymmetrical sinuses and well-developed ornament. The re¬ sults indicate that two “luciellide” subclades evolved by the late Llandovery (Early Silurian). One of these includes species of Oehlertia Pemer, 1907 (Figure 33, nodes 197, 198). These species possess strong swellings on either side of the peripheral band, straight lunulae, and a slit bordered by very strong lira that is within a much broader peripheral band. The final set of features are illustrated in Figure 4 e. The sister group of Oehler¬ tia is the Conotoma claustrata clade (Figure 33, nodes 199-201), whose synapomorphies are a greatly extended right ramp, a peripheral band very low on the whorl, and frill-like peripheral band lira with the upper lirum noticeably stronger than the lower one. More-derived species of the Crenilunula hallei clade (nodes 200, 201) have the peripheral band lira de¬ veloped into a small frill. In addition, species within this clade possess a slit that produced unusual zipper-shaped lunulae. Linking Crenilunula Knight, 1945, to Prosolarium Pemer, 1907, is consistent with the opinion of Linsley et al. (1978) that Crenilunula is related to the Luciellidae rather than to Euomph- alopterus (Knight et al., 1960). This analysis also supports Flower's (1968a) and Linsley et al.’s suggestions that the Early Ordovician Rhombella (Figure 8, node 3) is not closely related to the Luciellidae. Neither the estimates of the relationships between the two “luciellide” subclades nor the hypothesized relationship of “luciellides” to other “brachytomariines” are well-resolved. This analysis implies that the clade diverged from other “brachytomariines” by the late Caradoc (Late Ordovician), even though the earliest known species do not appear until the late Llandovery (Early Silurian). This gap is somewhat dis¬ concerting, but it is not quite statistically significant. “Luciel¬ lides,” however, are a highly derived clade relative to Ordovi¬ cian “brachytomariines,” and this long-branch problem (Felsenstein, 1978) creates two difficulties. First, many “lu¬ ciellide” synapomorphies are homoplastic on even the most parsimonious tree. Second, the highly derived nature of “lu¬ ciellides” means that some of their seemingly singular features might be homologous with traits on other species. For exam¬ ple, Oehlertia species possess strong swellings on either side of the peripheral band, which I coded as unique characters (Appendix 1, character 47). They also lack the strong carina on the right and left ramps that diagnose the derived “brachy¬ tomariines” of the Ordovician (i.e., nodes 190-192); however, the strong right ramp and left ramp carinae of “phanerotrema¬ tide” species, such as “ Longstaffia ” “laquetta ” and Promour¬ lonia aff. P. furcata Longstaff, 1924, are similar to those swellings. If one codes the swellings of Oehlertia as homolo¬ gous with the carinae of other “brachytomariines,” then it be¬ comes most parsimonious to place Oehlertia within the “L.” “laquetta ” clade. Similarly, some members of the rather vari¬ able species Crenilunula hallei (Whiteaves, 1895) appear to bear slightly weaker versions of the carina displayed by P. aff. P. furcata. These are lost amidst strong ornament in other members of the species, however, and on species such as Cre¬ nilunula limata (Lindstrom, 1884). Again, if one assumes that these carina are present, then it becomes more parsimonious to assume that C. hallei is the stem-member of a clade derived from a species similar to P. aff. P furcata. In this case, Cono¬ toma claustrata (Lindstrom, 1884) is considered to be a some¬ what derived member of the same clade (albeit plesiomorphic relative to Oehlertia gradata ), whereas Oehlertia is consid¬ ered to be a separate derivative within the “L.” “ laquetta ” clade. II.4.2.2.4. “Planozonides”.—The final “brachytomariine” subclade (Figures 34, 35, nodes 202-215) includes species classified in the Gosseletinidae by Knight et al. (1960). Be¬ cause the least derived members of this clade (i.e., species clas¬ sified as Cataschisma Branson, 1909, and Globispira Koken, 1925) have been classified as the Planozonides (a tribe within the Gosseltinidae), I label the clade the “planozonides.” I re¬ serve “gosseletinite” for one of the two major “planozonide” FIGURE 32 (opposite).—Relationships among the “phanerotrematides.” For abbreviations, see legend to Figure 7. Node 190 ( Brachytomaria semele clade), symmetric sinus angle -20° (2—4); symmetric sinus width (5); very strong lunulae (39); LRC present (75); both IM and base thickened (88); nonarched 1M (95); right side of aperture not inclined (113); no anterior projection of base (118). Node 191, narrow sinus (6, 7); [3 = 50° (48); flat LR (72); very thick IM (87). Node 192 ( Brachytomaria striata clade), thin, thread-like RRC (66); weak LRC (77); left side of aperture not inclined (111). Node 193, PB width = 15° (20); slit present (34). Node 194 ( Ulrichospira similis clade), moderately strong BL (23); lunulae same strength as GL (39); very asymmetric aperture (narrow left side) (58, 59); thick IM (87); only IM (not base) thickened (88); very low E (121); isometric T (127). Node 195 ( Phanerotrema clade), very nar¬ row sinus (6, 7); crenulated aperture (12); PB width = 10° (20); PB partially on RR (43); asymmetric ramp shapes (RR rounder) (51-53); acute suture (69, 70); LRC lost (75); thickened middle of IM (96); IM 15° off || to CA (98); PI thick¬ ness same as rest of shell (103); left side of aperture inclined -20° (111, 112); moderate E (121); ornament throughout left side of aperture (129). Node 196, oblique suture (69); IM || to CA (98); no anterior production of aperture (118). .3 S § S •S o ^ o ^ i- Ci, 3 ~ s S -3 Q o ^ O S u ^ co o q a ^ S « a 3 is P. r> 05 bJ 5 § JE O U g C/5 ■3 0 ° 1 . - 10 ° 2 . - 20 ° 3. -30° 4. -40° 5. -50° 6. -60° 7. -70° s>c 4. Sinus depth on left side (= angle of sinus retreat) 0 . > 0 ° 1 . - 10 ° 2 . - 20 ° 3. -30° 4. -40° 5. -50° 6. -60° 7. -70° 5. Sinus width symmetry 1. Right side wider (as on Clathrospira species and de¬ scendants) 2. Symmetric 3. Left side wider (as on Hormotoma confusa and rela¬ tives) s ’ c 6. Width of right side of sinus (Note: variation in the ramp length affects sinus width without changing the char¬ acter state.) 1. Just above peripheral band 2. Between peripheral band and top of right ramp 3. At top of right ramp 4. Above top of right ramp s,c 7. Width of left side of sinus 1. Just below peripheral band 2. Between peripheral band and alveozone (= left ramp) 3. At base of alveozone 4. Between alveozone and inner margin 8. Symmetry of sinus shape 1. More acute curve on the left (as on Hormotoma spe¬ cies) 2. Symmetric 3. More obtuse curve on the left (as on Clathrospira or Pleurorima) s 9. Shape of right side of sinus 1. Plateaus before apex (half a “U” as on Raphistoma and Raphistomina) 2. Straight into apex (as on Ectomaria) 3. Continuous curve towards apex (as on Hormotoma) 4. Hyperbolic curve towards apex (as on Climacora- phistoma) s 10. Shape of left side of sinus 1. Plateaus before apex (half a “U” as on as on Raphis¬ tomina) 2. Straight into apex 3. Continuous curve towards apex (as on Clathrospira) 4. Hyperbolic curve towards apex (as on Hormotoma) 11. Wrinkled right sinus (= a strong kink in the sinus, as on Raphistoma) 1. Absent 2. Present 12. Crenulated aperture (usually represented by undulat¬ ing growth lines) 1. Absent 2. Present 13. Aperture with reverse sigma shape (when viewed from the side) 1. Absent 2. Present (as on “helicotomids”) 14. Extent of sigmoidal aperture 1. Absent 2. Sigmoidal 3. Hypersigmoidal 15. Growth-line prominence 1. Not visible 2. Weak 3. Fine sharp 4. Strong 5. Extremely strong 16. Ontogenetic change in growth-line strength 1. No change over ontogeny 93 94 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 2. Stronger on juveniles (as on Lesueurilla) 17. Imbricated growth lines 1. Absent 2. Weak 3. Moderate 4. Strong 18. Growth lines on base 1. Same 2. Stronger than on the rest of shell (as on Malayaspira) 19. Peripheral band 1. Absent 2. Present c 20. Peripheral-band width 1. E = 05° 2 . 1 = 10 ° 3. E = 15° 4. E = 20° 5. E = 25° 6. E = 30° 21. Peripheral lira (i.e., lira bordering the peripheral band; present with “single” lirum (27) only if there is on¬ togenetic change (41) or if the peripheral band is trilineate) 1. Absent 2. Present 22. Peripheral-lira shape 1. Round (as on Hormotoma) 2. Sharp (as on Ectomaria and Lophospira ) 3. Square (i.e., with edges as on Eotomaria and Parali- ospira ) 23. Peripheral-lira strength 1. Extremely weak 2. Weak 3. Moderate 4. Strong 5. Extremely strong 24. Peripheral lira forming a frill 1. Absent 2. Present (as on Conotoma and Crenilunula) 25. Frill strength 1. Weak 2. Strong 26. Relative strength of right and left peripheral-band lira 1. Same 2. Right lirum stronger than left lirum (as on Conotoma and Crenilunula) 27. Medial lirum (= single keel at apex of sinus; present with peripheral lira (21) only if there is ontogenetic change (41) or if the peripheral band is trilineate) 1. Absent 2. Present 28. Medial lirum type 1. Rounded (as on Schizopea ) 2. Sharp (as on Barnesella and Malayaspira) 3. Flange (as on Lesueurilla) 4. Squared ridge (as on Ophiletina) 5. Dull lump (as on Euomphalopterus) 29. Single peripheral-lira strength 1. Very weak 2. Weak 3. Moderate 4. Strong 5. Very strong 6. Extremely strong 30. Ontogenetic change in single peripheral-lira strength 1. No change over ontogeny 2. Becomes weaker 3. Becomes stronger 31. Single peripheral-lira tubes 1. Absent 2. Present 32. Imbricated single peripheral lira 1. Consistent 2. Flares periodically (as on Poleumita discors and rela¬ tives) 33. Peripheral-band prominence 1. None 2. Slight 3. Strong 34. Slit 1. Absent 2. Present 35. Continuity of slit 1. Periodic slit (i.e., absent periodically as on some Clathrospira) 2. Continuous slit (as on Pleurorima and relatives) 36. Width of slit relative to peripheral band 1. Subsuming peripheral band (as on Pararaphistoma) 2. Within peripheral band (as on Pleurorima and rela¬ tives) 3. Thinner than peripheral band (as on Oehlertia) 37. Lira within peripheral band (usually bordering thin slits (36)) 1. Absent 2. Two lira bordering slit (as on Oehlertia) 38. Lunulae shape 1. Concentric 2. Zipper-like (as on Crenilunula) 3. Sigma-shaped (as on Pararaphistoma) 4. V-shaped (as on Eccyliopterus) 5. Kinked (as on onychochilids) 6. Straight (as on Oehlertia) 39. Lunulae strength relative to the growth lines 1. Weaker than growth lines (as on Liospira or Pleuror¬ ima) 2. Same as growth lines 3. Stronger than growth lines (as on Brachytomaria bal- tica and relatives) NUMBER 88 95 40. Ontogenetic change in peripheral-band prominence 1. None 2. Increasing prominence (as on Straparollina) 3. Decreasing prominence (as on Catazone) 41. Ontogenetic change from a bilineate to a monolineate pe¬ ripheral band 1. Bilineate throughout ontogeny 2. Becoming monolineate over ontogeny (as on early Straparollina) 42. Peripheral-band attitude 1. Projecting normally to the aperture 2. Peripheral band curves adapically (as on “lesueur- illids” or Helicotoma ) 43. Symmetry of peripheral band relative to the aperture 1. Asymmetrical on alveozone (as on “gosseletides”) 2. Bisects whorl 3. Asymmetrical on right ramp (as on Lesueurilla and Eccyliopterus ) 4. Entirely on right ramp (as on Eotomaria and Para- liospira ) 44. Ontogenetic change in peripheral-band symmetry 1. No change 2. Moving from the right ramp to bisecting aperture (as on Paraliospira) 45. Channel beneath peripheral band 1. Absent 2. Present (as on Lophospira perangulata) 46. Channel strength 1. Weak 2. Strong 47. Thickenings on either side of peripheral band (Note: these might be homologous with the right and left ramp carina) 1. Absent 2. Present (as on Oehlertia) c 48. Position of anal notch (based on plane passing through centroid and sinus apex; if perpendicular to inner margin, (3 = 0°; if parallel to inner margin and ori¬ ented adapically, (3 = 0°; if parallel to inner margin and oriented abapically, (3 = 180°). 1. (3 = 150° (e.g., Prosolarium) 2. (3 = 140° 3. |3 = 130° 4. (3= 120° 5. (3= 110° 6. (3= 100° 7. |3 = 90 o 8. (3 = 80° 9. (3 = 70° A. (3 = 60° B. [3 = 50° C. (3 = 40° D. (3 = 30° E. (3 = 20° F. (3=10° (e.g., Centrifugus) 49. Ontogenetic change in 3 (due to rotation of the aperture clockwise or counter-clockwise) 1. Absent 2. Decreasing |3 (as on basal “raphistomatoids”) 3. Increasing [3 (as on Paraliospira) 50. Magnitude of ontogenetic rotation 1. Small (<30°) 2. Great (>30°) 51. Symmetry of ramp shapes 1. Right ramp rounder (as on “holopeides”) 2. Symmetric 3. Alveozone rounder (as on “lesueurillids”) s 52. Right-ramp shape (see also 72) 1. Extremely globular (forming an acute angle) 2. Globular 3. Convex 4. Flat 5. Concave 53. Ontogenetic change in right-ramp shape 1. Ramp becoming more concave 2. Ramp becoming slightly more convex (as on Pachys- trophia) 3. Ramp becoming much more convex (as on Parara- phistoma) 54. Symmetry of ramp lengths 1. Right ramp longer (as on “luciellides”) 2. Symmetric 3. Alveozone longer (as on Raphistoma striata and rela¬ tives) S ’ C 55. Right-ramp length (= degrees between peripheral band and the top of ramp) 0 . < 20 ° 1. -30° 2. -40° 3. -50° 4. -60° 5. -70° 6. -80° 7. -90° 8 . - 100 ° s,c 56. Alveozone length (= degrees between peripheral band and the base of ramp) 0 . < 20 ° 1. -30° 2. -40° 3. -50° 4. -60° 5. -70° 6. -80° 7. -90° 8 . - 100 ° 9. -110° 57. Symmetry of ramp projections (see 58, 59) 1. Right higher (as on “helicotomids”) 2. Symmetric 96 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 3. Left higher (as on “holopeides”) s,c 58. Right-ramp projection (= angle between peripheral band and top of ramp) 1 . - 20 ° 2. -30° 3. -40° 4. -50° 5. -60° 6. -70° 7. -80° 8. -90° 9. -100° s ’ c 59. Alveozone projection (= angle between peripheral band and base of ramp) 0 . - 20 ° 1. -30° 2. -40° 3. -50° 4. -60° 5. -70° 6. -80° 7. -90° 60. Thickening of shell at the top of right ramp 1. Absent 2. Present (as on early “archaeogastropods” and bellero- phonts) 61. Degree of thickening 1. Weak 2. Moderate 3. Strong 4. Very strong c 62. Ontogenetic change in right-ramp swelling 1. Always acute (as on Ecculiomphalus) 2. Going from acute to obtuse over ontogeny (as on Ma- layaspira) 3. Always obtuse 63. Ontogenetic change in right ramp 1. None 2. Ramp becoming shorter and rounder (as on Mala- yaspira) 64. Right-ramp carina (RRC in figure captions) 1. Absent 2. Present 65. Right-ramp carina strength (in terms of prominence) 1. Weak 2. Moderate (roughly equal to a weak-to-moderate pe¬ ripheral band) 3. Strong (roughly equal to a strong peripheral band) 66. Right-ramp carina type 1. Dull thick thread 2. Dull thin thread 3. Round thin-to-moderately wide lira 4. Sharp thin-to-moderately wide thread 67. Ontogenetic change in right-ramp carina strength 1. None 2. Becomes weaker (as on Raphistomina species) 3. Becomes sharper (as on Raphistoma species) 68. Channel beneath right-ramp carina 1. Absent 2. Present (as on “poleumitids”) 69. Suture type 1. Right ramp oblique at suture 2. Right ramp acute at suture 70. Degree of acuteness 1. Acute (as on subulitids) 2. Attenuated (as on Hormotoma salteri ) 71. Aperture curving back at suture 1. Absent 2. Present (as on Liospira) 72. Alveozone shape (see also 52) 1. Extremely globular (forming an acute angle) 2. Globular 3. Convex 4. Flat 5. Concave 73. Thickening of shell at the base of alveozone 1. Absent 2. Present 74. Degree of thickening 1. Weak 2. Moderate 3. Strong 4. Very strong 75. Alveozone carina 1. Absent 2. Present 76. Alveozone-carina type 0. Dull thick thread 1. Thick swelling of shell 2. Sharp thin-to-moderately wide thread 3. Frill (as on Euomphalopterus species—go to charac ter 79) 4. Square thread (as on Ophiletina sublaxa ) 77. Alveozone-carina strength 1. Weak 2. Moderate (= peripheral band) 3. Strong 4. Very strong 78. Channeled alveozone carina 1. Absent 2. Present 79. Frilled alveozone carina 1. Peg (as on Euomphalopterus cariniferus) 2. Weak frill (as on Poleumita alatum) 3. Extended frill (as on Euomphalopterus alatus ) 80. Frill bordered by lira 1. Absent 2. Present (as on Euomphalopterus togatus) 81. Crenulated frill 1. Absent NUMBER 88 97 2. Present (as on Euomphalopterus undulans) 82. Channeled frill 1. Absent 2. Present (as on Euomphalopterus alatus) 83. Projection of frill relative to coiling axis 1. Behind aperture 2. Parallel to aperture 3. In front of aperture 84. Hood-like frill 1. Absent 2. Present (as on Pseudophorus praetextus) 85. Alveozone-carina tubes 1. Absent 2. Present (as on Euomphalopterus togatus) 86. Increasing expansion at base of alveozone over ontogeny (inducing a more “bowl”-like apical umbilicus) 1. Absent 2. Present (as on Teiichispira ) 87. Inner-margin (= columellar lip of most species) thickness, relative to rest of shell 1. No different 2. Thicker 3. Much thicker 88. Thickened region(s) of inner margin 1. Only top of inner margin thicker 2. Entire inner margin thicker 3. Inner margin and base thicker 4. Only base of inner margin thicker 89. Basal carina (BC in figure captions) 1. Absent 2. Present 90. Basal-carina type 1. Narrow thickening on the base 2. Dull extension with channel 3. Sharp thread 4. Rounded thread 5. Peg 91. Prominence of basal carina 1. Weak 2. Strong 3. Projecting 92. Ontogenetic change in basal carina 1. None 2. Becomes weaker with age 3. Becomes stronger with age 93. Position of basal carina 1. Beneath inner margin of aperture 2. Middle of whorl 3. Beneath outer margin of aperture c 94. Angle between inner margin and base 0. <45° 1. >45° 2. >60° 3. >75° 4. >90° 5. >105° 6 . > 120 ° 95. Shape of inner margin on the inner margin of aperture 1. Round 2. Curved 3. Straight 4. Arched inwards 96. Shape of inner margin on the outer margin of aperture 1. More obtuse than the inner margin 2. Same 3. More acute than the inner margin 97. Ontogenetic change in inner-margin shape 1. None 2. Becomes rounder with age c 98. Inner-margin attitude relative to coiling axis 1. -0° 2. -15° 3. -30° 4. -45° 5. -60° 6. -75° 7. -90° 8. -105° 9. -120° 99. Siphon 1. Absent 2. Slight twist of inner margin 3. Strong extension of inner margin 100. Inner-margin lira 1. Absent 2. Present 101. Type of inner-margin lira 1. Lira (as on Paraliospira) 2. Callous (as on Pycnomphalus) 102. Inner-margin channel 1. Absent 2. Present 103. Parietal-inductura strength 1. Absent 2. Thin and incomplete 3. Thin 4. Same as shell 5. Thicker than shell 104. Inductura projection 1. Parallels rest of aperture 2. Extended in front of aperture (as on Liospira) 3. A thick, concentrate strip extended in front of aper ture (as on Poleumita ) 4. Fills umbilicus (as on Eotomaria) 105. Flaring aperture 1. Absent 2. Weak (as on Loxonema) 3. Strong (as on Gasconadia) 106. Reflected inner-margin lip 1. Absent 98 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY 2. Present (as on basal “hormotomoids”) 107. Funicle 1. Absent 2. Present (as on Liospira) 108. Inner margin fills umbilicus 1. Umbilicus open or filled by inductura 2. Inner margin fills umbilicus 3. Inner margin contiguous with previous whorl 4. Inner margin folds back into umbilicus 109. Whole aperture inclined (species with left and right halves both inclined (111 and 113), but to different degrees, were scored as unknown (?)) 1. Radial/backwards inclination (see 115) 2. Left and right side inclined to same degree (as on Stra- parollina ) C 110. Angle of inclination of whole aperture 1. InAn = 10° 2. InAn = 20° 3. InAn = 30° 4. InAn = 40° 5. InAn = 50° 6. InAn-60° 111. Inclination of left half of aperture 1. Radial 2. Inclined (right side different or radial; e.g., Clath- rospira) C 112. Inclination of left side of aperture 0. InAn = 5° 1. InAn = 10° 2. InAn = 20° 3. InAn = 30° 4. InAn -40° 113. Inclination of right half of aperture 1. Radial 2. Right side inclined (left side different or radial; e.g., Pleurorima) c 114. Inclination of right side of aperture 1. InAn = 10° 2. InAn = 20° 3. InAn = 30° 4. InAn-40° 115. Aperture inclined “backwards” 1. Absent/forward inclination 2. Present (as on onychochilids) 116. Posterior projection of aperture base 1. Absent (as on all species with anterior projections; see also 120) 2. Present (as on Schizopea) c 117. Magnitude of posterior projection 1. --10° 2 . -- 20 ° 3. —30° 4. —40° 5. --50° 118. Anterior projection of aperture base 1. Absent 2. Present (as on “helicotomids” and “eotomarioids”) C 119. Magnitude of anterior projection 1. -10° 2 . - 20 ° 3. -30° 4. -40° 5. -50° 120. Shape of inner-margin base 1. Straight 2. Excavated (curved) (as on Spiroraphe) 3. Excavated (crenulated) (as on Lesueurilla ) c 121. Shell expansion (in radians) 1. E < 0.05 (extremely low) 2. 0.05 < E < 0.10 (very low) 3. 0.10 0.25 (very high) 7. E > 1.0 (extremely high) 122. Ontogenetic change in expansion 1. Decreasing expansion over ontogeny 2. None 3. Increasing expansion over ontogeny c 123. Shell curvature around coiling axis (in radians) 0. K < 0.40 (open coiling) 1. K < 0.5 (extremely low) 2. 0.5 < K < 0.55 (very low) 3. 0.55 0.95 (extremely high) 124. Ontogenetic change in curvature 1. No change over ontogeny 2. Decreasing curvature over ontogeny 125. Anisostrophy (applies only to coiling, not left-right asymmetry) 1. Isostrophic 2. Anisostrophic c 126. Shell torque 0. High ultradextral 1. Moderate ultradextral 2. Low ultradextral 3. Nearly planispiral 4. Low dextral 5. Moderate dextral 6. High dextral 7. Very high dextral 8. Extremely high dextral 127. Ontogenetic change in shell torque 1. Increasing torque over ontogeny 2. Isometric NUMBER 88 99 3. Decreasing torque over ontogeny 128. Septation 1. Absent 2. Septation present (as on most “euomphalinae”) 3. Complete filling of juvenile whorls (as on Palliseria) 129. Ornament on left side of aperture 1. Absent 2. Present throughout the alveozone and base 3. Present on the alveozone only 130. Density of ornament 1. One thread per 20° 2. One thread per 10° 3. One thread per 5° 4. One thread per 1° 131. Strength of alveozone ornament 1. Fine threads 2. Weak lira 3. Strong lira 132. Consistency of alveozone ornament 1. Uniform 2. Threads stronger higher on the alveozone 3. Threads stronger lower on the lower ramp 4. Highest threads strong, the rest uniform 133. Ornamentation of right ramp 1. Absent 2. Present throughout 3. Ornament present on upper half of right ramp only 134. Density of ornament on right side 1. One thread per 20° 2. One thread per 10° 3. One thread per 5° 4. One thread per 1° 135. Strength of ornament on right side 1. Fine threads 2. Weak lira 3. Strong lira 136. Pattern of ornament on right side 1. Uniform 2. 2:1:2 (every other thread twice as strong as interme¬ diate thread) 137. Type of right-ramp ornament 1. Local thickenings on shell 2. Local changes in aperture shape 138. Transverse ornament 1. Absent 2. Present (as on Cataschisma) 139. Peripheral-band ornament 1. Absent 2. Present (as on Crenilunula) 140. Carrier-shell scars 1. No scars 2. Scars (as on Lytospira ) 141. Size (= shell volume) 0. Very small (micro-mollusc: <10 mm 3 ) 1. Small (> 10 mm 3 < 10 2 mm 3 ) 2. Moderate (> 10 2 mm 3 < 10 3 mm 3 ) 3. Large (> 10 3 mm 3 < 10 4 mm 3 ) 4. Huge (> 10 4 mm 3 < 10 5 mm 3 ) 142. Protoconch coiling 1. Like teleoconch 2. Planispiral (as on Sinuspira) 143. Protoconch size 1. Small (< 10 _l mm) 2. Large (> 10 -1 mm, as on Murchisonia and Loxonema) Appendix 2. Data Matrix Data matrix for the analyzed species. Character numbers correspond to those in Appendix 1, and species numbers correspond to those in Appendix 3. Outgroup species have letters rather than numbers. 100 NUMBER 88 101 Appendix 2.—Continued. Number Species 1 2 3 4 5 6 7 8 9 1 0 1 1 1 2 1 3 1 4 1 5 1 6 1 7 1 8 1 9 2 0 2 1 2 2 2 3 2 4 2 5 AC. Mollusc ? ? ? ? ? ? ? ? ? 9 ? ? ? ? ? 9 ? ? ? ? ? ? 9 9 9 AB. Helcionella subrugosa 1 n n n n n n n n n n n 1 n 4 ? ? ? 1 n n n n n n AA. Latouchella merino 1 n n n n n n n n n n n 1 n 5 1 4 1 1 n n n n n n Z. Pelagiella subangulala 1 n n n n n n n n n n n 2 1 2 1 1 1 1 n n n n n n Y. Costipelagiella zazvorkai 1 n n n n n n n n n n n 2 1 4 1 1 1 1 n n n n n n X. Oelandia rugosa 1 n n n n n n n n n n n 1 n 4 ? ? ? 1 n n n n n n W. Coreospira rugosa 1 n n n n n n n n n n n 1 n 4 1 2 1 1 n n n n n n V. Sinuella minuata 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 1 n n n n n n u. Chippewaella paiellitheca 2 2 4 4 2 2 2 2 4 4 i 1 1 n 4 1 1 1 2 2 i n n n n T. Strepsodiscus major 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 3 i n n n n S. Chalarostrepsis praecursor 2 2 6 6 2 2 2 2 4 4 i 1 1 n 4 1 1 1 2 3 2 2 i i n R. Eobucania mexicana 2 2 6 6 2 2 2 2 4 4 i 1 1 n 3 1 1 1 2 2 2 2 i i n Q Strepsodiscus paucivoluta 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 4 1 n n n n p. Modestospira poulseni 2 2 6 6 2 3 3 2 3 3 i 1 1 n 4 1 1 1 1 n n n n n n 0. Peelerophon oehlerti 2 2 6 6 2 2 2 2 4 4 i 1 1 n 5 1 1 1 2 3 2 2 4 i n N. Kiringella pyramidalis 1 n n n n n n n n n i n 1 n 3 1 2 1 1 n n n n n n M. Cyrtolites sp. 1 n n n n n n n 11 n i n 1 n 4 1 3 1 2 3 1 n n n n L. Macluritellal walcotti 1 n n n n n n n n n n n ? n 4 1 1 2 2 2 1 n n n n K. Euomphalopsis involuta 2 2 i i 2 1 1 2 3 3 i 1 1 n 2 1 1 1 1 n n n n n n J. “ Maclurites " thomsoni 1 n n n n n n n n n n n 1 n 4 1 1 1 2 1 i n n n n I. Kobayashiella circe 1 n n n n n n n n n n n 2 n 5 1 1 1 1 n n n n n n H. Scaevogyra swezeyi 1 n n n n n n n n n n n 2 n 4 1 1 1 2 2 i n n n n G. Matherella saratogensis 1 n n n n n n n n n n n 1 n 5 1 1 1 1 n n n n n n F. Matherellina walcotti 1 n n n n n n n n n n n 1 n 5 1 1 1 2 2 i n n n n E. Hypseloconus elongatus 1 n n n n n n n n n n n 1 n 4 1 1 1 1 n n n n n n D. Knightoconus antarcticus 1 n n n n n n n n n n n 1 n 4 1 1 1 1 n n n n n n C. Sinuites sowerbyi 2 2 3 3 2 2 2 2 i i i 1 1 n 4 1 1 1 1 n n n n n n B. Cloudia buttsi 2 2 3 3 2 2 2 2 i i i 1 1 n 4 1 1 1 1 n n n n n n A. Owenella antiquata 2 2 2 2 2 2 2 2 i i i 1 1 n 4 1 1 1 1 n n n n n n 1. Dirhachopea normal is 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 3 2 i 2 1 n 2. Dirhachopea subrotunda 2 2 5 5 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 3 2 i 2 1 n 3. Schizopea typica 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 3 1 n n n n 4. Sinuopea sweeti 2 2 4 4 2 2 2 2 1 1 i 1 1 n 4 1 1 1 1 n n n n n ii 5. Taeniospira emminencis 2 2 4 4 2 3 3 2 3 3 i 1 1 n 4 1 1 1 2 4 2 i 3 i n 6. Ceratopea canadensis 2 2 6 6 2 3 3 2 4 4 i 1 1 n 4 1 1 1 2 3 1 n n n n 7. Gasconadia putilla 2 2 6 6 2 1 1 2 4 4 i 1 1 n 1 1 1 1 2 1 2 i 2 n n 8. Jarlopsis conicus 2 1 6 4 1 2 2 2 4 4 2 1 1 n 4 1 1 1 2 2 1 n n n n 9. Ophileta supraplana 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 1 1 1 2 3 1 n n n n 10. Rhombella umbilicata 2 1 6 4 1 3 3 2 4 4 1 1 1 n 4 1 1 1 2 4 1 n n n n 11. Prohelicotoma uniangulata 2 2 3 3 2 3 3 2 2 2 1 1 1 n 4 1 1 1 2 2 1 n n n n 12. Sinuopea basiplanata 2 2 4 4 2 3 3 2 3 3 1 1 1 n 4 1 1 1 2 4 2 i 2 I n 13. Taeniospira 1st. clairi 2 2 4 4 2 3 3 2 3 3 1 1 1 n 4 1 1 1 2 5 2 i 2 1 n 14. Bridgeites Idisjuncta 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 ? 1 ? 2 2 1 n n n n 15. Bridgeites planodorsalis 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 1 1 1 2 3 1 n n n n 16. Bridgeites supraconvexa 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 1 1 1 2 3 1 n n n n 17. Euconia etna 2 1 6 4 1 3 3 2 4 4 1 1 1 n 2 1 1 1 2 3 1 n n n n 18. Ceratopea llaurentia 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 1 1 2 2 3 1 n a n n 19. Ceratopea pygmaea 2 2 6 6 2 3 3 2 4 4 1 1 1 n 3 1 1 2 2 3 1 n n n n 20. Orospira bigranosa 2 2 5 5 2 3 3 2 4 4 1 1 1 n 3 1 3 1 2 3 1 n n n n 21. Macluritella stantoni 2 2 2 2 2 3 3 2 2 2 1 1 1 n 2 1 3 1 2 1 1 n n n n 22. Teiichispira odenvillensis 2 2 2 2 2 3 3 2 2 2 1 1 1 n 2 1 1 1 2 1 1 n n n n 23. Teiichispira loceana 2 2 2 2 2 3 3 2 2 2 1 1 1 n 2 1 1 1 2 1 1 n n n n 24. Palliseria robust a 2 2 1 1 2 2 2 2 2 2 1 1 1 n 2 1 1 1 2 2 1 n n n n 25. Mitrospira longwelli 2 2 1 1 2 2 2 2 2 2 1 1 1 n 2 1 1 1 2 2 1 n n n n 26. Teiichispira kobayashi 2 2 2 2 2 3 3 2 2 2 1 1 1 n 2 1 1 1 2 1 1 n n n n 27. Teiichispira sylpha 2 2 1 1 2 2 2 2 2 2 1 1 1 n 2 1 1 1 2 2 1 n n n n 28. Monitorella auricula 2 2 2 2 2 3 3 2 2 2 1 1 1 n 2 1 1 1 2 2 1 n n n n 29. Maclurites magna 2 2 2 2 2 1 1 2 2 2 1 1 1 n 3 ? 1 2 2 1 1 n n n n 30. “Eccyliopterus omatus" 2 3 1 3 2 3 3 3 3 2 1 2 1 n 2 1 1 2 2 2 1 n n n n 31. Maclurites bigsbyi 2 2 2 2 2 1 1 2 2 2 1 1 1 n 4 1 1 2 2 1 1 n n n n 102 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 AC. Mollusc ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? ? 7 ? ? ? 1 7 ? 2 AB. Helcionella subrugosa n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n AA. Latouchella merino n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n Z. Pelagiella subangulata n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n Y. Costipelagiella zazvorkai n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n X. Oelandia rugosa n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n W. Coreospira rugosa n n n n n n n n 1 n n n n n n n n n n ? n 1 7 1 n V. Sinuella minuata n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 7 1 n U. Chippewaella patellitheca n 2 5 1 1 i 1 1 1 n n n 1 2 1 n 1 2 i 1 n 1 7 1 n T. Strepsodiscus major n 2 5 2 1 i 1 1 1 n n n 1 2 1 n 1 2 i 1 n 1 7 1 n s. Chalarostrepsis praecursor n 2 5 1 1 i 1 1 2 2 2 1 1 2 1 1 1 2 i 1 n 1 7 1 n R. Eobucania mexicana n 2 5 1 1 i 1 1 2 2 2 1 1 2 1 1 1 2 i ? n 1 7 1 n Q Strepsodiscus paucivoluta n 2 5 1 1 i 1 1 1 n n n 1 2 1 n 1 2 i 1 n 1 7 1 n p. Modestospira poulseni n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 7 1 n o. Peelerophon oehlerti n i n n n n n 1 2 2 2 1 1 1 i 1 1 2 i 1 n 1 7 1 n N. Kiringella pyramidalis n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n M. Cyrtolites sp. n 2 i 4 1 i 1 n 1 n n n 4 2 i n 1 2 i 2 2 1 7 1 n L. Macluritellal walcotti n 2 i 2 1 i 1 1 1 n n n 5 n i n 1 2 i 1 n 1 7 1 n K. Euomphalopsis involuta n n n n n n n n 1 n n n 4 n n n n n n 1 n 1 7 1 n J. “Maclurites" thomsoni n 2 2 2 1 1 1 i 1 n n 1 5 2 1 n 1 2 i 2 i 1 7 1 n I. Kobayashiella circe n n n n n n n n I n n 1 5 2 n n n n n 1 n 1 7 1 n H. Scaevogyra swezeyi n 2 2 2 1 1 i i 1 n n 1 5 2 i n i 2 i 2 i 1 7 1 n G. Matherella saratogensis n n n n n n n n 1 n n 1 5 2 n n n n n 1 n 1 9 1 n F. Matherellina walcotti n 2 2 2 i 1 1 1 1 n n 1 5 2 i n i 2 i 1 n 1 9 1 n E. Hypseloconus elongatus n n n n n n n n 1 n n 1 n n n n n n n 1 n 1 7 1 n D. Knightoconus antarcticus n n n n n n n n 1 n n 1 n n n n n n n 1 n 1 7 1 n C. Sinuites sowerbyi n n n n n n n n 1 n n n i i n n n n n 1 n 1 7 1 n B. Cloudia buttsi n n n n n n n n I n n n i i n n n n n 1 n 1 7 1 n A. Owenella antiquata n n n n n n n n 1 n n n i i n n n n n 1 n 1 7 1 n 1. Dirhachopea normalis n 2 i 2 i i 1 1 1 n n 1 i 2 1 2 i 2 i 1 n 1 5 1 n 2. Dirhachopea subrotunda n 1 n n n n n 1 1 n n 1 i 2 1 1 i 2 i 1 n 1 6 1 n 3. Schizopea typica n 2 1 2 1 i i 1 1 n n 1 i 2 1 n i 2 i 1 n 1 5 1 n 4. Sinuopea sweeti n n n n n n n n 1 n n n i 1 1 n n n n 1 n 1 6 1 n 5. Taeniospira emminencis n i n n n n n 1 1 n n i i 2 1 1 1 2 i 1 n 1 6 1 n 6. Ceratopea canadensis n 2 2 3 1 1 1 1 1 n n i i 2 1 n 2 2 i 2 1 1 5 1 n 7. Gasconadia putilla n 2 1 1 1 1 1 1 I n n i i 2 1 2 1 2 i 1 n 1 5 1 n 8. Jarlopsis conicus n 2 1 3 1 1 1 1 2 2 2 i i 2 1 n 1 2 i 1 n 1 2 1 n 9. Ophileta supraplana n 2 3 3 1 1 1 1 1 n n i i 2 1 n 2 2 i 2 1 1 5 1 n 10. Rhombella umbilicata n 2 1 2 1 1 1 1 1 n n i i 2 1 n 1 2 i 1 n 1 2 1 n 11. Prohelicotoma uniangulata n 2 3 3 2 1 1 1 1 n ■ i i i 1 1 n 2 2 i 2 1 1 5 1 n 12. Sinuopea basiplanata n 1 n n n n n 1 1 n n i i 2 1 1 1 2 i 1 n 1 7 1 n 13. Taeniospira 1st. clairi 1 1 n n n n n 1 1 n n i i 2 1 1 1 2 i 1 n 1 7 1 n 14. Bridgeites Idisjuncta n 2 4 3 1 1 1 I 1 n n i i 2 1 n 2 2 i 1 n 1 4 1 n 15. Bridgeites planodorsalis n 2 2 3 1 1 1 1 1 n n i i 2 1 n 2 2 i 2 1 1 4 1 n 16. Bridgeites supraconvexa n 2 2 3 1 1 1 1 1 n n i i 2 1 n 2 2 i 1 n 1 2 1 n 17. Euconia etna n 2 1 2 1 1 1 1 1 n n i i 2 1 n 1 2 i 1 n 1 2 1 n 18. Ceratopea llaurentia n 2 3 4 1 1 1 1 1 n n i i 2 1 n 2 2 i 2 1 1 5 1 n 19. Ceratopea pygmaea n 2 3 4 1 1 I 1 1 n n i i 2 1 n 2 2 i 2 1 1 5 3 i 20. Orospira bigranosa n 2 2 3 1 1 1 1 1 n n i 3 2 1 n 2 2 i 1 n 1 8 1 n 21. Macluritella stantoni n 2 1 3 2 1 1 1 1 n n i 1 1 1 n 1 2 i 2 1 1 7 2 n 22. Teiichispira odenvillensis n 2 3 3 2 1 1 1 1 n n i 1 1 1 n 2 2 i 2 1 1 7 2 n 23. Teiichispira loceana n 2 3 3 2 1 1 1 1 n n i 1 1 1 n 2 2 i 2 1 1 7 2 n 24. Palliseria robusta n 2 1 2 I 1 1 1 1 n n i 1 2 1 n 2 2 i 1 n 1 7 2 n 25. Mitrospira longwelli n 2 I 2 1 1 1 1 1 n n i 1 2 1 n 2 2 i 2 1 1 7 2 n 26. Teiichispira kobayashi n 2 1 2 2 1 1 1 1 n n i 1 1 1 n 1 2 i 2 2 1 B 1 n 27. Teiichispira sylpha n 2 1 3 1 1 1 1 1 n n i 1 2 1 n 2 2 i 1 n 1 9 2 n 28. Monitorella auricula n 2 1 3 2 1 1 1 1 n n i 1 1 1 n 2 2 i 2 1 1 9 2 n 29. Maclurites magna n 2 1 3 1 1 1 1 1 n n i 1 2 1 n 1 2 i 1 n 1 9 1 n 30. “Eccyliopterus ornatus" n 2 1 3 n 1 1 1 1 n n i 1 1 1 n 1 2 i 2 1 1 8 2 n 31. Maclurites bigsbyi n 2 1 2 1 1 1 1 1 n n i 1 2 1 n 1 2 i 1 n 1 9 1 n NUMBER 88 103 Appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 6 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 AC. Mollusc ? ? ? 7 ? 7 ? ? 7 7 7 7 I 7 7 ? 7 ? 7 7 1 7 7 7 7 AB. Helcionella subrugosa 2 2 2 2 2 2 2 5 5 1 n n 1 1 n n n n i n 1 2 1 n 1 AA. Latouchella merino 2 1 2 2 2 2 2 5 5 1 n n 1 1 n n n n i n 1 1 1 n 1 Z. Pelagiella subangulata 3 3 2 2 7 7 2 2 2 1 n n 1 2 2 i 1 i i n 1 2 1 n 1 Y. Costipelagiella zazvorkai 2 2 2 2 7 7 2 2 2 1 n n 1 1 n n n n i n 1 2 1 n 1 X. Oelandia rugosa 2 2 2 2 2 2 2 5 5 1 n n 1 1 n n n n i n 1 2 1 n 1 W. Coreospira rugosa 2 2 2 2 3 3 2 7 7 2 4 1 1 1 n n n n i n 1 2 1 n 1 V. Sinuella minuata 2 3 2 2 4 4 2 7 7 2 4 1 1 1 n n n n i n 1 3 2 4 ? u. Chippewaella patellitheca 2 2 2 2 8 8 2 2 2 2 1 3 1 1 n n n n 7 n 1 2 1 1 1 T. Strepsodiscus major 2 2 2 2 8 8 2 2 2 2 2 2 1 1 n n n n 1 n 1 2 2 2 1 s. Chalarostrepsis praecursor 2 2 2 2 8 8 2 2 2 2 1 3 1 1 n n n n 1 n 1 2 2 1 1 R. Eobucania mexicana 2 2 2 2 8 8 2 2 2 2 1 3 1 1 n n n n 1 n 1 2 2 1 1 Q Strepsodiscus paucivoluta 2 2 2 2 8 8 2 1 1 2 2 3 1 1 n n n n 1 n 1 2 2 1 1 P. Modestospira poulseni 2 2 2 2 7 7 2 4 4 2 2 3 1 1 n n n n 1 n 1 2 2 3 1 0. Peelerophon oehlerti 2 2 2 2 8 8 2 2 2 2 1 3 1 1 n n n n 1 n 1 2 2 1 1 N. Kiringella pyramidal is 2 2 2 2 7 7 2 2 2 1 n n 1 1 n n n n ? n 1 2 1 n ? M. Cyrtolites sp. 2 3 2 2 7 7 2 2 2 1 n n 1 1 n n n n ? n 1 3 1 n 2 L. Macluritella ? walcotti 1 2 2 ? 8 8 7 2 2 1 n n 1 1 n n n n 1 n 1 3 1 n 1 K. Euomphalopsis involuta 2 1 2 2 7 7 2 4 4 1 n n 1 1 n n n n 1 n 1 1 1 n 1 J. “Maclurites" thomsoni 3 3 2 1 7 7 2 1 1 1 n n 1 1 n n n n 1 n 1 1 1 n 1 I. Kobayashiella circe 3 4 2 1 7 5 3 3 4 1 n n 1 1 n n n n 1 n 1 1 1 n 1 H. Scaevogyra swezeyi 3 3 2 1 7 3 1 3 1 1 n n 1 1 n n n n 2 i 1 2 1 n 1 G. Matherella saratogensis 3 4 2 1 7 6 1 5 0 1 n n 1 1 n n n n 2 i 1 2 1 n 1 F. Matherellina walcotti 3 4 2 1 7 4 1 6 5 1 n n 1 1 n n n n 2 i 1 2 1 n 1 E. Hypseloconus elongatus 2 2 2 2 7 7 2 3 3 1 n n 1 ? n n n n ? n 1 2 1 n ? D. Knightoconus antarcticus 2 2 2 2 7 7 2 2 2 1 n n 1 ? n n n n 7 n 1 2 1 n 1 C. Sinuites sowerbyi 2 2 2 2 7 7 2 3 3 2 3 3 1 1 n n n n ! n 1 2 2 3 ? B. Cloudia buttsi 2 2 2 2 7 7 2 4 4 2 3 3 1 1 n n n n 1 n 1 2 2 3 ? A. Owenella antiquata 2 2 2 2 7 7 2 6 6 2 3 3 1 1 n n n n 1 n 1 2 2 3 ? 1. Dirhachopea normal is 2 2 2 2 8 8 2 2 2 2 2 2 1 1 n n n n 1 n 1 2 2 2 1 2. Dirhachopea subrotunda 2 2 2 2 7 7 2 3 3 2 2 3 1 1 n n n n 1 n 1 2 2 2 1 3. Schizopea typica 2 2 2 2 8 8 2 2 2 2 2 2 1 1 n n n n 1 n 1 2 2 2 1 4. Sinuopea sweeti 2 2 2 2 7 7 2 5 5 2 3 3 1 1 n n n n 1 n 1 2 2 3 1 5. Taeniospira emminencis 2 2 2 2 7 7 2 4 4 2 3 3 1 1 n n n n 1 n 1 2 2 3 1 6. Ceratopea canadensis 2 2 2 2 6 6 2 2 2 2 1 3 1 1 n n n n 1 n 1 2 1 n 1 7. Gasconadia putilla 2 3 2 1 6 4 1 4 4 1 n n 1 1 n n n n 1 n 1 3 2 1 1 8. Jarlopsis conicus 2 2 2 1 8 4 1 2 1 2 2 2 1 1 n n n n 1 n 1 2 2 2 1 9. Ophileta supraplana 2 2 2 2 6 6 2 2 2 2 2 2 1 1 n n n n 1 n 1 2 2 1 1 10. Rhombella umbilicala 2 2 2 1 8 4 1 2 1 2 2 2 1 1 n n n n 1 n 1 2 2 2 1 11. Prohelicotoma uniangulata 2 1 2 2 4 4 2 2 2 2 2 2 1 1 n n n n 1 n 1 1 2 1 1 12. Sinuopea basiplanata 2 2 2 2 5 5 2 3 3 2 2 3 1 1 n n n n 1 n 1 2 2 3 1 13. Taeniospira 1st. clairi 2 2 2 1 4&5 3 2 5 5 2 1 3 1 1 n n n n 1 n 1 2 2 1 1 14. Bridgeites Idisjuncta 1 2 2 1 6 4 1 3 2 2 3 2 1 1 n n n n 1 n 1 3 1 n 1 15. Bridgeites planodorsalis 2 2 2 2 5 5 2 1 1 2 1 3 1 1 n n n n 1 n 1 3 1 n 1 16. Bridgeites supraconvexa 1 2 2 1 5 4 1 1 1 2 3 2 1 1 n n n n 1 11 1 3 1 n 1 17. Euconia etna 1 2 2 1 8 3 1 3 1 2 2 2 1 1 n n n n 1 n 1 3 1 n 1 18. Ceratopea llaurentia 1 2 2 2 6 6 2 1 1 2 1 3 1 1 n n n n 1 n 1 3 1 n 1 19. Ceratopea pygmaea 1 2 2 2 5 5 2 1 1 2 1 n 1 1 n n n n 1 n 1 3 1 n 1 20. Orospira bigranosa 2 3 2 2 5 5 2 3 3 2 1 3 1 2 3 4 l 1 1 n 1 3 1 n 1 21. Macluritella stantoni 3 1 1 2 4 4 2 2 2 2 1 2 1 1 n n n n 1 n 1 1 2 1 1 22. Teiichispira odenvillensis 3 2 1 2 5 5 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 23. Teiichispira loceana 2 1 1 2 5 5 2 1 1 2 1 2 1 1 n n n n 1 n 1 1 2 i 1 24. Palliseria robusta 3 3 2 3 5 6 1 3 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 25. Mitrospira longwelli 3 3 2 3 5 6 1 2 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 26. Teiichispira kobayashi 1 4 1 2 4 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 27. Teiichispira sylpha 3 4 2 3 5 6 1 2 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 28. Monitorella auricula 3 4 1 2 5 5 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 29. Maclurites magna 3 4 3 3 5 7 1 2 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 30. “Eccyliopterus omatus" 3 3 1 2 4 4 2 1 1 2 2 3 1 1 n n n n 1 n 1 2 1 n 1 31. Maclurites bigsbyi 3 3 3 3 3 7 1 3 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 104 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 7 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 1 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 AC. Mollusc 1 ? ? ? ? ? ? ? ? ? 7 ? ? ? ? ? ? 7 ? ? ? 1 1 ? ? AB. Helcionella subrugosa n n n n n n n n n n 1 1 n 1 n n n n 3 1 2 1 1 1 1 AA. Latouchella merino n n n n n n n n n n 1 I n 1 n n n n 3 3 2 1 1 1 1 Z. Pelagiella subangulata n n n n n n n n n n 1 2 n 1 n n n n 3 3 2 1 4 1 1 Y. Costipelagiella zazvorkai n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 4 1 1 X. Oelandia rugosa n n n n n n n n n n 1 1 n 7 ? ? ? 7 3 1 2 1 1 1 ? W. Coreospira rugosa n n n n n n n n n n 1 1 n 1 n n n n 5 3 2 1 1 1 2 V. Sinuella minuata ? ? ? n n n n n n n 1 ? ? 1 n n n n 3 3 2 1 1 1 1 u. Chippewaella patellitheca n n n n n n n n n n 1 7 ? 1 n n n n 5 2 2 1 1 1 1 T. Strepsodiscus major n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 1 1 1 S. Chalarostrepsis praecursor n n n n n n n n n n 1 1 n 1 n n n n 4 3 2 1 1 1 1 R. Eobucania mexicana n n n n n n n n n n 1 1 n 1 n n n n 4 3 2 1 1 I 1 Q. Strepsodiscus paucivoluta n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 1 1 1 p. Modestospira poulseni n n n n n n n n n n 1 1 n 1 n n n n 4 7 2 1 1 1 1 0. Peelerophon oehlerti n n n n n n n n n n 1 1 n 1 n n n n 4 3 2 1 1 1 1 N. Kir ingel la pyramidal is ? 7 ? n n n n n n n 1 1 n 1 n n n n 5 ? 2 1 1 1 1 M. Cyrtolites sp. 1 1 n n n n n n n n 1 1 n 1 n n n n 3 ? 2 1 1 1 2 L. Macluritella ? walcotti n n n n n n n n n n l 1 n 1 n ii n n 1 3 2 1 7 1 1 K. Euomphalopsis involuta n n n n n n n n II n 1 1 n 1 n n n n 2 1 2 1 7 1 1 J. “ Maclurites ” thomsoni n n n n n n n n n n 1 1 n ? ? ? ? ? 4 3 2 I 8 I 1 I. Kobayashiella circe n n n n n n n n n n 1 ? ? ? ? ? ? ? 5 2 2 1 7 1 1 H. Scaevogyra swezeyi n n n n n n n n n n 1 1 n ? ? ? 7 ? 5 2 2 1 8 1 1 G. Matherella saratogensis n n n n n n n n n n 1 1 n ? ? 7 ? ? 5 3 2 1 8 1 1 F. Matherellina walcotti n n n n n n n n n n 1 1 n ? ? 7 ? 7 5 2 2 1 8 1 1 E. Hypseloconus elongatus n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 1 1 1 D. Knightoconus antarcticus n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 1 1 1 C. Sinuites sowerbyi n n n n n n n n n n 1 1 n 1 n n n n 3 ? 2 1 1 1 1 B. Cloudia buttsi n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 A. Owenella antiquata n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 I. Dirhachopea normalis n n n n n n n n n n 1 1 n 2 i i i 2 3 2 2 1 2 1 1 2. Dirhachopea subrotunda n n n n n n n n n n 1 1 n 2 i i i 2 3 2 2 1 2 1 1 3. Schizopea typica n n n n n n n n n n 1 1 n 2 i i i 2 4 2 2 1 2 1 1 4. Sinuopea sweeti n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 I 1 5. Taeniospira emminencis n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 6. Ceratopea canadensis n n n n n n n n n n 1 2 2 2 i i i 2 4 3 2 1 4 1 1 7. Gasconadia putilla n n n n n n n n n n 1 1 n 1 n n n n 3 2 3 1 2 1 1 8. Jarlopsis conicus n n n n n n n n n n 1 1 n 2 i i i 2 6 2 2 1 3 1 1 9. Ophileta supraplana n n n n n n n n n n 1 1 n 2 i 2 i 2 4 3 2 1 4 1 1 10. Rhombella umbilicata n n n n n n n ii n n 1 1 n 2 i 1 i 2 6 2 2 1 3 1 1 11. Prohelicotoma uniangulata n n n n n n n n n n 1 1 n 2 i 1 i 2 4 1 2 1 6 1 1 12. Sinuopea basiplanata n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 13. Taeniospira 1st. clairi n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 14. Bridgeites Idisjuncta n n n n n n n n n n 1 2 2 2 i 3 1 2 3 2 2 1 7 1 1 15. Bridgeites planodorsalis n n n n n n n n n n 1 2 2 2 i 1 1 2 5 3 2 I 6 1 1 16. Bridgeites supraconvexa n n n n n n n n n n 1 2 2 2 i 3 1 2 5 3 2 1 7 1 1 17. Euconia etna n n n n n n n n n n 1 1 n 1 n n n n 6 2 2 1 3 1 1 18. Ceratopea llaurentia n n n n n n n n n n 1 2 2 2 2 3 i 2 5 3 2 1 4 1 1 19. Ceratopea pygmaea n n n n n n n n n n 1 2 2 2 2 3 i 2 5 3 2 1 4 1 1 20. Orospira bigranosa n n n n n n n n n n 1 2 2 2 3 2 i 1 4 3 2 1 2 1 2 21. Macluritella stantoni n n n n n n n n n n 1 1 n 2 1 1 ? 7 4 1 2 1 7 1 1 22. Teiichispira odenvillensis n n n n n n n n n n 2 1 n 2 1 1 1 2 4 1 2 1 7 1 1 23. Teiichispira loceana n n n n n n n n n n 2 1 n 2 1 1 1 2 4 1 2 1 7 1 1 24. Palliseria robusta n n n n n n n n n n 2 3 2 1 n n n 2 3 1 1 1 9 1 1 25. Mitrospira longwelli n n n n n n n n n n 2 3 3 1 n n n 2 3 1 1 1 9 1 1 26. Teiichispira kobayashi n n n n n n n n n n 2 1 n 2 1 1 1 2 3 2 2 1 5 1 1 27. Teiichispira sylpha n n n n n n n n n n 2 2 n 2 1 1 1 2 3 2 2 1 8 1 1 28. Monitorella auricula n n n n n n n n n n 2 1 n 2 1 1 1 2 3 2 2 1 7 1 1 29. Maclurites magna n n n n n n n n n n 1 1 n 2 1 1 1 2 3 3 2 1 7 1 1 30. “ Eccyliopterus ornatus ” n n n n n n n n n n 1 1 n I n n n n 3 2 2 1 6 1 1 31. Maclurites bigsbyi n n n n n n n n n n 1 1 n 2 1 1 i 2 4 3 2 1 7 1 1 NUMBER 88 105 Appendix 2—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 AC. Mollusc ? ? 4 7 ? ? 7 ? 1 ? ? ? ? 7 ? ? ? ? 7 7 ? ? 7 ? 1 AB. HelcioneUa subrugosa ? 2 4 2 1 1 1 1 1 ? 1 ? 1 7 1 2 2 1 n 1 6 2 1 1 1 AA. Lalouchella merino 1 2 4 2 3 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 1 6 2 1 1 1 Z. Pelagiella subangulata 1 1 4 1 1 1 1 1 2 2 1 7 1 7 1 2 2 1 n 1 6 2 4 1 2 Y. Coslipelagiella zazvorkai 1 1 4 1 1 1 1 1 2 3 1 7 1 7 1 2 2 1 n 1 6 2 4 1 2 X. Oelandia rugosa 1 7 4 ? 1 1 1 1 1 ? 1 ? 1 7 1 2 1 1 n 1 6 2 1 1 I w. Coreospira rugosa 1 2 4 2 1 1 1 1 1 ? 1 7 1 7 1 2 3 1 n 1 4 2 3 1 1 V. Sinuella minuata ? 1 4 7 1 1 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 2 2 2 1 1 u. Chippewaella patellitheca ? 1 3 2 1 1 1 1 1 7 1 7 1 7 1 2 5 1 n 1 7 2 1 1 1 T. Strepsodiscus major ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 3 2 3 2 1 s. Chalarostrepsis praecursor ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 3 2 3 2 1 R. Eobucania mexicana 7 1 3 1 1 1 1 1 1 7 1 ? 1 ? 1 2 5 1 n 1 3 2 4 1 1 Q Strepsodiscus paucivoluta ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 3 3 2 2 1 p. Modestospira poulseni 7 1 7 7 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 2 2 2 1 1 0. Peelerophon oehlerti 7 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 3 2 4 1 1 N. Kiringella pyramidal is 7 1 4 2 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 7 2 1 1 1 M. Cyrtolites sp. ? 2 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 5 1 n 1 5 2 2 1 1 L. Macluritellal walcotti ? 1 4 ? 1 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 2 1 2 0 1 2 K. Euomphalopsis involuta 7 1 4 ? 1 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 1 1 2 1 1 2 J. “Maclurites ” thomsoni 7 1 ? ? 2 1 1 1 1 7 1 ? 1 ? 2 1 n 2 2 1 3 2 5 1 2 I. Kobayashiella circe ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 2 1 n 2 1 1 2 2 5 1 2 H. Scaevogyra swezeyi 7 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 2 1 n 2 1 1 3 2 5 1 2 G. Matherella saratogensis 7 1 4 1 1 1 1 1 1 7 1 7 1 ? 2 1 n 2 1 1 2 2 5 1 2 F. Matherellina walcotti ? 1 4 1 1 1 1 1 1 7 1 7 1 7 2 1 n 2 1 1 2 2 5 1 2 E. Hypseloconus elongatus ? 1 4 2 1 1 1 1 1 ? 1 ? 1 7 1 2 5 1 n 1 7 2 3 1 1 D. Knightoconus antarcticus ? 1 4 2 1 1 1 1 1 ? 1 ? 1 7 1 2 5 1 n 1 7 2 3 2 1 C. Sinuites sowerbyi ? I 3 2 2 1 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 3 1 1 B. Cloudia buttsi ? 1 4 7 1 1 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 2 2 5 1 1 A. Owenella antiquata ? 1 4 7 1 1 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 2 2 5 1 1 1. Dirhachopea normalis 7 1 4 1 1 1 1 1 1 7 1 ? 1 7 1 2 3 1 n 1 3 2 4 2 2 2. Dirhachopea subrotunda ? I 4 1 1 1 1 1 1 ? 1 ? 1 7 1 2 3 1 n 1 3 2 4 1 2 3. Schizopea typica ? 1 4 1 1 1 1 1 1 7 1 ? 1 7 1 2 3 1 n 1 3 2 3 2 2 4. Sinuopea sweeti ? 1 4 1 1 1 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 2 2 5 1 2 5. Taeniospira emminencis 7 1 4 1 1 1 1 1 1 7 1 7 1 ? 1 1 n 1 n 1 4 2 4 1 2 6. Ceratopea canadensis ? 2 3 1 1 1 1 1 1 7 1 ? 1 ? 1 2 2 1 n 3 3 2 4 1 2 7. Gasconadia putilla ? 1 4 1 3 2 1 2 2 3 1 7 1 ? 1 2 1 1 n 1 2 2 5 1 2 8. Jarlopsis conicus 7 1 3 1 1 1 1 1 2 2 1 ? 1 ? 1 2 4 1 n 1 3 2 4 1 2 9. Ophileta supraplana ? ? 3 1 1 1 1 1 1 7 1 ? 1 ? 1 2 2 1 n 2 1 2 3 1 2 10. Rhombella umbilicata ? 1 3 1 1 1 1 1 2 2 1 ? 1 ? 1 2 3 1 n 1 3 2 4 1 2 11. Prohelicotoma uniangulata ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 3 1 n 2 4 2 3 2 2 12. Sinuopea basiplanata ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 4 1 2 13. Taeniospira 1st. clairi 7 1 3 1 1 2 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 3 2 5 1 2 14. Bridgeites Idisjuncta ? 1 3 1 1 1 1 1 1 ? 1 7 1 ? 1 2 3 1 n 3 3 2 3 2 2 15. Bridgeites planodorsalis 7 2 4 1 1 1 1 1 1 7 1 7 1 ? 1 2 3 1 n 3 3 2 3 2 2 16. Bridgeites supraconvexa ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 3 1 n 3 3 2 3 2 2 17. Euconia etna 7 1 3 1 1 1 1 1 2 2 1 ? 1 ? 1 2 3 1 n 1 2 2 5 1 2rt 18. Ceratopea llaurentia 7 2 2&3 1 1 1 1 1 1 ? 1 7 1 ? 1 2 3 1 n 3 4 2 4 1 2 19. Ceratopea pygmaea 7 2 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 3 1 n 3 4 2 4 1 2 20. Orospira bigranosa 1 1 3 1 1 1 1 1 1 7 1 ? 1 7 1 2 2 1 n 2 2 2 4 1 2 21. MacluriteUa stantoni ? 1 4 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 3 1 n 2 5 2 2 1 2 22. Teiichispira odenvillensis ? 1 3 1 1 1 1 1 1 ? 1 ? 1 7 1 2 1 1 n 1 4 3 4 1 2 23. Teiichispira 1 oceana ? 1 1 1 1 1 1 1 1 7 1 ? 1 7 1 2 3 1 n 2 5 3 3 1 2 24. Palliseria robusta ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 1 3 2 2 1 2 25. Mitrospira longwelli ? 1 4 ? 1 1 1 1 1 7 1 7 1 ? 1 2 1 1 n 1 3 2 2 1 2 26. Teiichispira kobayashi 7 1 3 7 1 1 1 1 1 ? 1 ? 1 7 1 2 1 1 n 1 6 3 3 1 2 27. Teiichispira sylpha ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 1 4 3 2 1 2 28. Monitorella auricula 7 1 3 1 1 1 1 1 1 7 1 7 1 ? 1 2 1 1 n 1 4 3 4 1 2 29. Maclurites magna 7 1 4 1 1 1 1 1 1 ? 1 7 1 7 1 2 3 1 n 1 5 3 3 1 2 30. “Eccyliopterus omatus ” ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 2 1 1 n 1 5 3 4 1 2 31. Maclurites bigsbyi ? 1 4 1 1 1 1 1 1 7 1 ? 1 7 1 2 3 1 n 1 5 2 3 1 2 106 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. T - 1 1 1 1 1 1 1 1 1 1 1 1 1 i 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 AC. Mollusc 3 1 7 7 7 7 7 7 7 7 7 7 1 7 7 7 7 7 AB. Helcionella subrugosa 3 1 2 1 n n n 1 n n n n 1 n 1 0 1 n AA. Latouchella merino 3 1 2 1 n n n 1 n n n n 1 n 1 0 1 n Z. Pelagiella subangulata 3 1 1 2 3 1 1 2 3 1 1 1 1 n 1 0 1 n Y. Costipelagiella zazvorkai 3 1 1 1 n n n 1 n n n n 1 n 1 0 1 1 X. Oelandia rugosa 3 1 2 1 n n n 1 n n n n 1 n 1 0 7 n w. Coreospira rugosa 3 1 7 1 n n n 1 n n n n 1 n 1 1 7 n V. Sinuella minuata 3 1 1 1 n n n 1 n n n n 1 n 1 1 7 n u. Chippewaella patellitheca 3 1 ? 1 n n n 1 n n n n 1 n 1 2 7 n T. Strepsodiscus major 3 1 2 1 n n n 1 n n n n 1 1 1 2 7 n S. Chalarostrepsis praecursor 3 1 7 1 n n n 1 n n n n 1 1 1 2 7 n R. Eobucania mexicana 3 1 7 1 n n n 1 n n n n 1 1 1 7 7 n Q Strepsodiscus paucivoluta 3 1 2 1 n n n 1 n n n n 1 1 1 2 7 n p. Modestospira poulseni 3 1 7 1 n n n 1 n n n n 1 n 1 2 7 n 0. Peelerophon oehlerti 3 1 1 I n n n 1 n n n n 1 n 1 2 7 n N. Kiringella pyramidalis 3 1 7 1 n n n 1 n n n n 1 n 1 2 7 n M. Cyrtolites sp. 3 1 1 1 n n n 1 n n n n 1 n 1 2 7 n L. Macluritellal walcotti 3 1 2 2 2 2 1 2 2 i i 1 1 1 1 1 7 n K. Euomphalopsis involuta 3 1 2 1 n n n 1 n n n n 1 n 1 2 7 n J. “ Maclurites ” thomsoni 1 1 1 1 n n n 1 Tl n n n 1 1 1 2 1 n I. Kobayashiella circe 0 1 7 1 n n n 1 n n n n 1 1 1 0 7 n H. Scaevogyra swezeyi 1 1 1 1 n n n 1 n n n n 1 1 1 2 7 n G. Matherella saratogensis 0 1 1 1 n n n 1 n n n n 1 1 1 2 7 n F. Matherellina walcotti 0 1 1 1 n n n 1 n n n n 1 1 1 2 7 n E. Hypseloconus elongatus 3 1 2 1 n n n 1 n n n n 1 1 1 3 7 n D. Knightoconus antarcticus 3 1 2 1 n n n 1 n n n n 1 1 1 3 7 n C. Sinuites sowerbyi 3 1 1 1 n n n I n n n n 1 1 1 2 7 n B. Cloudia buttsi 3 1 1 1 n n n 1 n n n n 1 n 1 1 7 n A. Owenella antiquata 3 1 1 1 n n n 1 n n n n 1 n 1 1 7 n 1. Dirhachopea normalis 4 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 2. Dirhachopea subrotunda 4 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 3. Schizopea typica 4 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 4. Sinuopea sweeti 6 1 1 1 n n n 1 n n n n 1 n 1 1 7 n 5. Taeniospira emminencis 6 1 I 1 n n n 1 n n n n 1 i 1 2 7 n 6. Ceratopea canadensis 4 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 7. Gasconadia putilla 6 1 7 1 n n n 1 n n n n 1 i 1 1 7 n 8. Jarlopsis conicus 5 1 2 1 n n n 1 n n n n 1 i 1 3 7 n 9. Ophileta supraplana 4 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 10. Rhombella umbilicata 5 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 11. Prohelicotoma uniangulata 4 1 2 1 n 11 n 1 n n n n 1 i 1 2 7 n 12. Sinuopea basiplanata 6 1 1 1 n n n 1 n n n n 1 i 1 2 7 n 13. Taeniospira 1st. clairi 7 2 1 1 n n n 1 n n n n 1 i 1 2 7 n 14. Bridgeites Idisjuncta 3 1 7 1 n n n 1 n Tl n n 1 i 1 3 7 n 15. Bridgeites planodorsalis 3 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 16. Bridgeites supraconvexa 3 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 17. Euconia etna 5 1 7 1 n n n 1 n n n n 1 i 1 2 7 n 18. Ceratopea llaurentia 4 ? 2 1 n n n 1 n n n n 1 i 1 2 7 n 19. Ceratopea pygmaea 4 2 2 1 n n n 1 n n n n 1 i 1 2 7 n 20. Orospira bigranosa 4 1 2 2 3 i i 2 3 1 i 1 1 i 1 2 7 n 21. Macluritella stantoni 2 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 22. Teiichispira odenvillensis 2 1 7 1 n n n 1 n n n n 1 i 1 2 7 n 23. Teiichispira loceana 3 1 7 1 n n n 1 n n n n 1 i 1 2 7 n 24. Palliseria robusta 1 1 3 1 n n n 1 n n n n 1 i 1 3 7 n 25. Milrospira longwelli 1 1 3 1 n n n 1 n n n n 1 i 1 4 7 n 26. Teiichispira kobayashi 2 1 7 1 n n n 1 n n n n 1 i 1 2 7 n 27. Teiichispira sylpha 2 1 3 1 n n n 1 n n n n 1 i 1 2 7 n 28. Monitorella auricula 2 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 29. Maclurites magna 2 1 3 1 n n n 1 n n n n 1 i 1 4 7 n 30. “Eccyliopterus ornatus ” 2 I 2 1 n n n 1 n n n n 1 i 1 2 7 n 31. Maclurites bigsbyi 2 1 3 3 3 3 1 1 n n n n 1 i 1 4 7 n NUMBER 88 107 Appendix 2— Continued. Number Species 1 2 3 4 5 6 7 8 9 I 0 1 1 1 2 1 3 1 4 i 5 1 6 1 7 1 8 1 9 2 0 2 1 2 2 2 3 2 4 2 5 32. Maclurina logani 2 2 2 2 2 1 1 2 2 2 1 1 1 n 4 1 1 2 2 1 1 n n n n 33. Maclurina manitobensis 2 2 2 2 2 1 1 2 2 2 1 1 1 n 2 1 1 1 2 1 1 n n n n 34. Maclurites sedgewicki 2 2 2 2 2 1 1 2 2 2 1 1 1 n 2 1 1 1 2 I 1 n n n n 35. Maclurites expansa 2 2 2 2 2 1 1 2 2 2 1 1 1 n 3 1 1 1 2 1 1 n n n n 36. Ophileta complanata 2 2 6 6 2 3 3 2 4 4 1 1 1 n 3 1 1 7 2 3 1 n n n n 37. Lecanospira compact a 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 1 1 2 2 2 1 n n n n 38. Lecanospira nereine 2 2 6 6 2 2 2 2 4 4 1 1 1 n 4 1 1 7 2 2 1 n n n n 39. Barnesella llecanospiroides 2 2 4 4 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 2 1 n n n n 40. Malayaspira hintzei 2 3 4 6 2 3 3 2 4 4 I 1 1 n 4 2 2 2 2 2 1 n n n n 41. Malayaspira rugosa 2 2 5 5 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 2 1 n n n n 42. Barnesella measuresae 2 2 4 4 2 3 3 2 3 3 I 1 1 n 4 2 1 2 2 2 1 n 11 n n 43. Lytospira angel ini 2 2 4 4 2 3 3 2 3 3 1 1 1 n 4 1 1 2 2 1 1 n n n n 44. Lytospira yochelsoni 2 2 4 4 2 3 3 2 3 3 1 1 1 n 4 2 1 2 2 2 1 n n n n 45. Maclurina lannulata 2 3 5 6 2 3 3 2 4 4 1 1 1 n 4 2 2 2 2 2 1 n n n n 46. Rossospira harrisae 2 3 4 6 2 3 3 2 4 4 1 1 1 n 4 2 2 1 2 2 1 n n n n 47. Ecculiomphalus bucklandi 2 3 2 6 2 3 3 2 4 4 1 1 1 n 4 7 2 1 2 2 1 n n n n 48. Lytospira gerrula 2 3 3 5 2 3 3 l 2 3 1 1 1 n 3 2 1 2 2 1 1 n n n n 49. Lytospira Inorx’egica 2 3 3 5 1 3 2 1 2 3 1 1 1 n 3 7 1 2 2 2 1 n n n n 50. Ophiletina cf. 0. sublaxa 2 2 1 1 2 3 3 2 2 2 1 1 1 n 2 2 1 2 2 1 1 n n n n 51. Lytospira subrotunda 2 2 4 4 2 3 3 2 1 1 1 1 1 n 3 1 1 2 1 n n n n n n 52. Pararaphistoma lemoni 2 2 6 6 2 3 3 2 4 4 1 1 1 n 4 2 I 2 2 i i n n n n 53. Climacoraphistoma vaginati 2 1 5 4 1 3 2 2 4 4 1 i 1 n 4 2 1 2 2 2 i n n n 11 54. Lesueurilla bipatellare 2 1 6 5 1 3 2 2 4 4 1 1 1 n 3 1 1 2 2 2 i n n n n 55. Lesueurilla marginal is 2 1 5 3 1 3 2 2 4 4 1 1 1 n 4 2 1 2 2 2 i n n n n 56. Lesueurilla prima 2 1 5 4 1 3 2 2 4 4 1 1 1 n 4 2 1 2 2 2 i n n n n 57. Palaeomphalus giganteus 2 2 5 5 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 3 i n n n n 58. Climacoraphistoma damesi 2 1 5 4 1 3 2 2 4 4 1 1 1 n 3 2 1 2 2 2 i n n n n 59. Eccyliopterus alatus 2 2 4 4 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 1 i n n n n 60. Eccyliopterus Iprinceps 2 2 3 3 1 3 2 2 3 3 1 1 1 n 4 1 1 2 2 2 i n n n n 61. Eccyliopterus regularis 2 2 4 4 2 3 3 2 4 4 1 1 1 i 4 2 1 2 2 2 i n n n n 62. Lesueurilla infundibula 2 1 5 3 1 3 2 2 4 4 1 1 1 n 4 2 1 2 2 2 i n n n n 63. Eccyliopterus louderbacki 2 1 5 4 1 3 3 2 4 4 1 1 1 n 4 2 1 2 2 2 i n n n n 64. Lesueurilla declivis 2 2 5 5 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 2 i n n n n 65. Pararaphistoma qualteriata 2 1 5 4 1 3 2 2 4 4 1 1 1 n 4 2 1 1 2 2 i n n n n 66. Pararaphistoma schmidti 2 1 6 4 1 3 2 2 4 4 1 1 1 n 4 1 1 1 2 1 i n n n n 67. Helicotoma gubanovi 2 2 4 4 2 3 3 2 1 1 1 1 1 n 4 7 1 ? 2 2 2 i 2 i nrt 68. Scalites katoi 2 2 5 5 2 3 3 2 4 4 1&2 1 1 n 2 2 1 1 2 2 1 n n n n 69. Helicotoma medfraensis 2 2 5 5 2 3 3 2 4 4 1 1 1 n 4 2 1 2 2 1 1 n n n n 70. Lesueurilla scotica 2 3 4 3 1 3 2 2 4 4 1 1 1 n 3 1 1 1 2 2 1 n n n n 71. Pachystrophia devexa 2 2 3 3 2 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 72. Raphistoma striata 2 2 4 4 2 3 3 2 1 1 2 1 1 n 4 2 1 7 2 2 2 i i i n 73. Raphistomina lapicida 2 2 2 2 2 3 3 2 1 1 1 1 1 n 4 1 1 1 2 2 1 n n n n 74. Scalites angulatus 2 2 4 4 2 3 3 2 4 4 1 1 1 n 4 ? 1 ? 2 2 1 n n n n 75. Holopea insignis 1 ? ? 7 ? ? ? ? ? 7 7 7 1 n 3 1 1 1 1 n n n n n n 76. Eccyliopterus beloitensis 2 1 4 3 1 3 3 2 3 3 l i 1 n 4 7 1 2 2 i i n n n n 77. Holopea rotunda 1 ? ? ? ? ? 7 ? 7 7 7 7 1 n 3 1 1 1 1 n n n n n n 78. Pachystrophia contigua 2 2 3 3 2 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 79. Pachystrophia spiralis 2 2 3 3 2 3 3 1 1 3 1 1 1 n 3 1 1 1 1 n n n n n n 80. Raphistomina aperta 2 2 2 2 2 3 3 2 1 1 1 1 1 n 4 1 1 1 2 2 i n n n n 81. Raphistomina fissurata 2 2 2 2 2 3 3 2 1 1 1 1 1 n 3 1 1 1 2 2 i n n n n 82. Eccyliopterus owenanus 2 2 3 3 2 3 3 2 4 4 1 1 1 n 4 ? 1 2 2 1 i n n n n 83. Holopea ampla 1 ? ? ? 7 ? ? ? 7 ? 7 1 1 n 3 1 1 1 1 n n n n n n 84. Holopea pyrene 1 ? ? ? 7 7 ? 7 7 ? ? ? 1 n 3 1 4 1 1 n n n n n n 85. Holopea symmetrica 1 ? 7 ? ? ? ? 2 ? ? 1 1 1 i 3 1 1 1 1 n n n n n n 86. Raphistoma peracuta 2 2 4 4 2 3 3 2 1 1 2 1 1 n 4 1 1 1 2 2 1 n n n n 87. Raphistomina rugata 2 2 2 2 2 3 3 2 1 1 1 1 1 n 5 1 1 1 2 2 1 n n n n 88. Raphistoma tellerensis 2 2 3 3 2 3 3 2 4 4 2 1 1 n 3 2 1 1 2 2 1 n n n n 89. Sinutropis lesthetica 2 2 3 3 2 3 3 1 1 3 1 1 1 n 3 1 1 1 1 n n n n n n 90. Pachystrophia gotlandica 2 2 3 3 2 3 3 1 1 3 1 1 1 n 3 1 1 1 1 n n n n n n 91. Lytospira triquestra 2 2 2 2 2 2 2 1 1 3 1 1 1 n 2 1 1 1 1 n n n n 11 n 108 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 32. Maclurina logani n 2 1 2 1 1 1 1 1 n n 1 1 2 1 n 1 2 1 1 n 1 9 1 n 33. Maclurina manitobensis n 2 1 1 1 1 1 1 1 n n 1 1 2 1 n 1 2 1 1 n 1 9 1 n 34. Maclurites sedgewicki n 2 1 1 1 1 1 1 1 n n 1 1 2 1 n 1 2 1 1 n 1 9 1 n 35. Maclurites expansa n 2 1 3 I 1 1 1 1 n n 1 1 2 1 n 1 2 1 1 n 1 9 1 n 36. Ophileta complanata n 2 3 4 I 1 1 1 1 n n 1 1 2 l n 2 2 1 2 1 1 5 1 n 37. Lecanospira comp acta n 2 2 4 I 1 1 1 1 n n 1 ] 1 1 n 2 2 1 2 I 1 7 1 n 38. Lecanospira nereine n 2 2 4 1 1 1 1 1 n n 1 1 1 1 n 2 2 1 ? n 1 7 1 n 39. Barnesella llecanospiroides n 2 2 4 1 1 1 1 1 n n 1 4 1 1 n 2 2 I 1 n 1 7 1 n 40. Malayaspira hintzei n 2 2 4 1 1 1 1 1 n n 1 4 3 1 n 2 2 1 1 n 1 8 1 n 4 1. Malayaspira rugosa n 2 2 3 1 1 1 I 1 n n 1 4 3 1 n 2 2 1 1 n 1 8 2 n 42. Barnesella measuresae n 2 2 4 1 1 1 1 1 n n 1 4 1 1 n 2 2 1 1 n 1 7 1 n 43. Lytospira angelini n 2 2 3 1 1 1 1 1 n n 1 1 n 1 ii 1 2 1 2 i 1 7 1 n 44. Lytospira yochelsoni n 2 2 4 1 1 1 1 1 n n 1 4 1 1 n 2 2 1 I n I 7 1 n 45. Maclurina lannulata n 2 2 3 1 1 1 1 1 n n 1 4 3 1 n 2 2 1 1 n 1 2 1 n 46. Rossospira harrisae n 2 2 4 1 1 1 I 1 n n 1 4 3 1 n 2 2 1 1 n 1 6 1 n 47. Ecculiomphalus bucklandi n 2 2 3 1 1 1 1 1 n n 1 4 2 1 n 2 2 1 ? n 1 6 1 n 48. Lytospira gerrula n 2 2 4 1 1 1 1 1 n n 1 4 1 1 n 1 2 1 2 i I 7 1 n 49. Lytospira Inorvegica n 2 3 4 1 1 1 1 1 n n 1 4 1 1 n 1 2 n 2 i 1 6 ? n 50. Ophiletina cf. 0 . sublaxa n 2 2 2 1 I I 1 1 n n 1 4 1 1 n 1 2 i 2 i 1 7 1 n 51 . Lytospira subrotunda n it n n n n n n 1 n n n 1 2 1 n n n n l n 1 C 1 n 52. Pararaphistoma lemoni n 2 3 4 i 1 I 1 1 n n i 1 2 1 n 2 2 i 2 i 1 5 2 n 53. Climacoraphistoma vaginati n 2 3 4 i 1 I 1 2 2 1 i 3 1 1 n 2 3 i 2 i 1 7 3 2 54. Lesueurilla bipatellare n 2 3 5 i 1 1 I 2 2 1 i 3 1 1 n 2 3 i 1 n 1 9 3 1 55. Lesueurilla marginalis n 2 3 5 i 1 1 1 2 2 1 i 3 1 1 n 2 3 i 2 i 1 9 3 1 56. Lesueurilla prima n 2 3 4 i 1 1 1 2 2 1 i 3 1 1 n 2 3 i 2 i 1 7 2 1 57. Palaeomphalus giganteus n 2 3 4 i 1 1 1 1 n n i 3 2 1 n 2 2 i 2 i 1 5 2 1 58. Climacoraphistoma damesi n 2 3 3 i 1 1 1 2 2 i i 3 3 I n 2 3 i 1 n 1 8 3 2 59. Eccyliopterus alatus n 2 3 6 i 1 1 1 1 n n i 3 2 1 n 2 2 i 2 i 1 8 1 n 60. Eccyliopterus Iprinceps n 2 3 5 i 1 1 1 1 n n i 3 2 I n 2 3 n 1 n 1 7 7 n 6 1 . Eccyliopterus regularis n 2 3 5 i 1 1 1 1 n n i 3 2 I n 2 2 i 2 i 1 5 1 n 62. Lesueurilla infundibula n 2 3 4 i 1 1 2 2 2 1 i 3 1 l n 2 3 i 2 i 1 6 3 i 63. Eccyliopterus louderbacki n 2 3 4 i I 1 2 2 2 1 i 3 1 1 n 2 3 i 2 i 1 7 1 n 64. Lesueurilla declivis n 2 3 4 i 1 1 1 I n n i 3 2 1 n 2 2 i 2 i 1 7 2 i 65. Pararaphistoma qualteriata n 2 3 4 i 1 1 1 2 2 i i 3 1 1 n 2 3 i 2 i 1 8 3 2 66. Pararaphistoma schmidti n 2 3 4 i 1 1 1 2 2 i i 3 1 1 n 2 3 i 1 n 1 7 3 2 67. Helicotoma gubanovi 1 2 4 2 i 1 1 2 2 n n i 1 1 1 n 2 2 i 1 n 1 B 2 n 68. Scalites katoi n 2 3 5 i 1 1 1 1 n n i 3 1 1 n 2 2 i 2 1 1 B 2 n 69. Helicotoma medfraensis n 2 3 2 2 1 1 1 1 n n i 1 2 2 n 2 2 i 2 I 1 7 2 n 70. Lesueurilla scotica n 2 3 3 1 1 1 1 2 2 i i 3 1 1 n 2 3 i l n 1 8 3 i 1 1 . Pachystrophia devexa n n n n n n n n 1 n n n 1 2 n n n n n 1 n I 7 1 n 72. Raphistoma striata 1 2 4 5 i i i 2 2 2 i 1 1 1 i n 2 2 i 1 n 1 B 3 i 73. Raphistomina lapicida n 2 3 5 i i i 1 1 n n 1 1 2 i n 2 2 i 2 1 1 6 1 n 74. Scalites angulatus n 2 4 5 i i i 1 7 n n 1 n n i n 2 2 i 1 n 1 A 2 n 75. Holopea insignis n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n 76. Eccyliopterus beloitensis n 2 3 4 1 i i 2 2 n i i n 1 1 n 2 3 i 2 1 1 6 1 n 77. Holopea rotunda n n n n n n n n 1 n n n n n n n n n n 1 n 1 B 1 n 78. Pachystrophia contigua n n n n n n n n 1 n n n i 2 1 n n n n 1 n 1 9 1 n 79. Pachystrophia spiralis n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 C 1 n 80. Raphistomina aperta n 2 3 5 i i i i ? n n i i 2 i n 2 2 1 2 1 1 5 1 n 8 1. Raphistomina fissurata n 2 3 3 i i i i ? n n i i 2 i n 1 2 1 2 1 1 7 1 n 82. Eccyliopterus owenanus n 2 3 6 i i i i 1 n n i 3 2 i n 2 2 1 2 1 1 8 1 n 83. Holopea ampla n n n n n n n n 1 n n n 1 2 i n n n n 1 n 1 9 1 n 84. Holopea pyrene n n n n n n n n 1 n n n n n n n n n n 1 n 1 7 1 n 85. Holopea symmetrica n n n n n i n n 1 n 11 n n n n n n n 1 1 n 1 D 1 n 86. Raphistoma peracuta n 2 3 5 1 i 1 1 1 n n 1 i 2 1 n 1 2 1 2 1 1 8 1 n 87. Raphistomina rugata n 2 3 5 1 i 1 2 1 n n 1 i 2 1 n 1 2 1 1 n 1 8 1 n 88. Raphistoma tellerensis n 2 3 5 1 i I 1 1 n n 1 i 1 2 n 2 2 1 1 n 1 B 1 n 89. Sinutropis lesthetica n n n n n n n n 1 n 11 n i 2 n n n n n 1 n 1 7 1 n 90. Pachystrophia gotlandica n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 C 1 n 91. Lytospira triquestra n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 7 1 n NUMBER 88 109 Appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 6 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 32. Maclurina logani 3 3 3 3 3 7 1 3 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 33. Maclurina manitobensis 3 3 3 3 4 7 1 3 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 34. Maclurites sedgewicki 3 3 3 3 4 7 1 4 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 35. Maclurites expansa 3 4 3 3 5 7 2 2 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 36. Ophileta complanata 3 3 2 2 5 5 2 2 2 2 2 3 1 1 n n n n 1 n 1 3 1 n 1 37. Lecanospira compacta 3 4 2 2 5 5 2 1 1 2 1 3 1 1 n n n n 1 n 1 3 1 n 1 38. Lecanospira nereine 3 4 2 2 5 5 2 1 1 1 n n 1 1 n n n n 1 n 1 3 1 n 1 39. Barnesella llecanospiroides 3 4 2 1 5 3 2 1 1 2 i 2 1 1 n n n n 1 n 1 3 1 n 1 40. Malayaspira hintzei 3 2 3 1 5 4 3 3 4 2 3 2 2 2 2 4 i i 1 n 1 3 1 n 2 41. Malayaspira rugosa 3 2 3 1 5 4 2 3 3 2 1 2 2 1 n n n n 1 n 1 3 1 n 1 42. Barnesella measuresae 3 3 2 1 5 3 2 1 1 2 3 2 1 1 n n n n 1 n 1 3 I n 1 43. Lytospira angel ini 3 2 1 2 5 5 2 2 2 2 1 3 1 1 n n n n 1 n 1 2 1 n 1 44. Lytospira yochelsoni 3 3 2 1 5 4 2 2 2 2 1 3 1 1 n n n n ? n 1 3 1 n 1 45. Maclurina lannulata 3 2 3 1 5 4 3 2 2 2 2 2 2 2 2 2 2 i 1 n 1 3 1 n 2 46. Rossospira harrisae 3 2 3 1 5 4 3 3 4 2 3 1 2 2 2 4 1 i 1 n 1 3 1 n 2 47. Ecculiomphalus bucklandi 3 3 2 1 5 4 3 3 4 2 1 1 2 1 n n n n 1 n 1 3 1 n 2 48. Lytospira gerrula 3 3 2 1 5 4 2 1 1 2 1 3 1 1 n n n n ? n 1 2 1 n 1 49. Lytospira Inorvegica 3 3 7 1 6 4 2 1 1 2 1 n 1 1 n n n n ? n 1 2 1 n 1 50. Ophiletina cf. 0. sublaxa 2 3 2 1 5 3 2 4 4 1 n n 1 2 i 4 i i 1 n 1 3 1 n 1 51. Lytospira subrotunda 3 2 2 2 5 5 1 3 3 2 i 3 1 1 n n n n 7 n 1 2 1 n 1 52. Pararaphisloma lemoni 1 2 2 1 5 5 2 1 1 2 i 2 1 1 n n n n 1 n 1 3 1 n 1 53. Climacoraphistoma vaginati 3 3 3 2 5 5 2 1 1 1 n n 1 1 n n n n 1 n 1 3 1 n 1 54. Lesueurilla bipatellare 3 2 2 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 3 1 n 1 55. Lesueurilla marginalis 3 3 2 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 56. Lesueurilla prima 3 3 2 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 57. Palaeomphalus giganteus 1 2&3 1 1 6 6 2 2 2 1 n n 1 1 n n n n 1 n 1 3 1 n 1 58. Climacoraphistoma damesi 3 3 3 1 6 4 2 2 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 59. Eccyliopterus alatus 3 3 2 1 6 6 1 1 1 l n n I 1 n n n n 1 n 1 2 1 n 1 60. Eccyliopterus Iprinceps 3 4 ? 1 6 5 2 1 1 2 1 n 1 1 n n n n ? n 1 3 1 n 1 61. Eccyliopterus regularis 3 2 2 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 62. Lesueurilla infundibula 3 4 1 1 5 3 2 2 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 63. Eccyliopterus louderbacki 3 2 1 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 64. Lesueurilla declivis 3 3 2 2 6 6 2 1 1 1 n n 1 1 n n n n 1 n 1 2 1 n 1 65. Pararaphisloma qualteriata 2 3 3 1 6 5 2 1 1 1 n n 1 1 n n n n 1 n 1 3 1 n 1 66. Pararaphisloma schmidti 2 3 3 1 6 4 3 1 2 1 n n 1 1 n n n n 1 n 1 3 1 n 1 67. Helicotoma gubanovi 2 2 2 3 2 5 1 4 1 1 n n 1 1 n n n n ? n 1 2 1 n 1 68. Scalites katoi 2 3 2 3 2 5 1 3 1 1 n n 1 1 n n n n i n 1 3 1 n 1 69. Helicotoma medfraensis 3 2 2 1 4 4 2 3 3 2 3 2 1 1 n n n n i n 1 2 1 n 1 70. Lesueurilla scotica 3 4 1 1 6 5 2 1 1 I n n 1 1 n n n n i n 1 3 1 n 1 71. Pachystrophia devexa 1 2 2 2 4 4 2 2 2 1 n n 1 1 n n n n i n 1 2 1 n 1 72. Raphistoma striata 2 2 2 3 2 5 1 3 1 1 n n 1 2 i 1&3 3 i i n 1 2 1 n 1 73. Raphistomina lapicida 1 2 3 2 4 4 2 1 1 1 n n 1 2 i 1 2 i i n 1 3 1 n 1 74. Scalites angulatus 2 3 2 3 2 5 1 4 1 1 n n 1 1 n n n n i n 1 3 1 n 1 75. Holopea irtsignis 1 2 2 3 3 4 3 3 6 1 n n 1 1 n n n n i n 1 2 1 n 1 76. Eccyliopterus beloitensis 3 2 1 2 6 6 2 1 1 1 n n 1 1 n n n n i n 1 2 1 n 1 77. Holopea rotunda 2 1 2 3 2 4 3 4 5 1 n n 1 1 n n n n 2 i 1 1 1 n 1 78. Pachystrophia contigua 1 2 2 2 4 4 3 3 4 1 n n 1 1 n n n n 1 n 1 2 1 n 1 79. Pachystrophia spiralis 3 2 2 3 4 5 2 4 4 1 n n 1 1 n n n n 1 n 1 1 1 n 1 80. Raphistomina aperta 1 2 2 2 4 4 2 1 1 1 n n 1 2 i 1 2 1 1 n 1 3 1 n 1 81. Raphistomina fissurata 1 2 2 2 4 4 2 1 1 l n n 1 2 i 1 2 1 1 n 1 3 1 n 1 82. Eccyliopterus owenanus 3 3 2 1 6 6 1 1 1 1 n n 1 i n n n n 1 n 1 2 1 n 1 83. Holopea ampla 1 2 2 3 3 4 3 3 5 1 n n 1 i n n n n 1 n 1 2 1 n 1 84. Holopea pyrene 1 2 2 3 3 4 3 3 6 1 n n 1 i n n n n 1 n 1 2 1 n 1 85. Holopea symmetrica 2 1 2 3 0 4 3 4 5 1 n n 1 i n n n n 1 n 1 1 1 n 1 86. Raphistoma peracuta 2 3 2 3 4 5 2 1 1 1 n n 1 2 2 4 2 i 1 n 1 3 1 n 1 87. Raphistomina rugata 2 3 1 2 4 4 2 1 1 1 n n 1 2 1 1 1 i 1 n 1 3 1 n 1 88. Raphistoma tellerensis 3 3 2 3 3 4 2 2 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 89. Sinutropis lesthetica 3 2 2 2 5 5 1 3 2 1 n n 1 1 n n n n 1 n 1 2 1 n 1 90. Pachystrophia gotlandica 3 2 2 2 5 5 1 3 2 1 n n 1 1 n n n n 1 n 1 1 1 n 1 91. Lytospira triquestra 3 2 2 2 4 4 1 4 3 1 n n 1 1 n n n n 1 n 1 2 1 n 1 110 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 1 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 I 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 32. Maclurina logani n n n n n n n n n n 1 1 n 2 1 1 1 2 4 3 2 1 7 1 1 33. Maclurina manitobensis n n n n n n n n n n 1 1 n 2 1 1 1 2 4 3 2 1 7 1 1 34. Maclurites sedgewicki n n n n n n n n n n 1 1 n 2 1 1 1 2 4 2 2 1 7 1 1 35. Maclurites expansa n n n n n n n n n n 1 1 n 2 1 1 1 2 3 3 2 1 7 1 1 36. Ophileta complanata n n n n n n n n n n 1 I n 2 1 2 1 2 4 3 2 1 4 1 1 37. Lecanospira compact a n n n n n n n n n n 1 1 n 2 1 2 1 2 4 3 2 1 7 1 1 38. Lecanospira nereine n n n n n n n n n n 1 ? ? ? ? 7 ? ? 4 3 ? 1 7 1 1 39. Barnesella llecanospiroides n n n n n n n n n n 2 1 n 2 1 2 1 2 3 3 2 1 7 1 1 40. Malayaspira hintzei 2 2 i n n n n n n 1 2 1 n 1 n n n n 4 2 2 2 6 1 1 41. Malayaspira rugosa n n n n n n n n n n 2 1 n 1 n n n n 4 2 2 2 6 1 1 42. Barnesella measuresae n n n n n n n n n i 2 1 n 2 5 3 1 3 3 3 2 1 7 1 1 43. Lytospira angel ini n n n n n n n n n n ? 1 n 1 n n n n 4 3 2 1 8 1 1 44. Lytospira yochelsoni n n n n n n n n n n ? 1 n 2 5 3 1 3 4 3 2 1 7 1 1 45. Maclurina lannulata 2 2 i n n n n n n i 1 1 n 1 n n n n 4 1&2 2 2 6 1 1 46. Rossospira harrisae 2 2 i n n n n n n i ? 1 n I n n n n 4 2 2 2 7 1 1 47. Ecculiomphalus bucklandi 2 2 i n n n n n n i 1 1 n 1 n n n n 5 2 2 2 7 1 1 48. Lytospira gerrula n n n n n n n n n n ? 1 n 1 n n n n 4 3 2 1 7 1 1 49. Lytospira Inorvegica n n n n n n n n n n 1 1 n 1 n n n n 5 2 3 1 4 1 2 50. Ophiletina cf. 0 . sublaxa n n n n n n n n n n 1 1 n 2 5 3 3 n 5 2 2 1 6 1 1 51. Lytospira subrotunda n n n n n n n n n n 1 1 n I n n n n 4 2 2 1 4 1 1 52. Pararaphistoma lemoni n n n n n n n n n n 1 1 n 2 2 1 1 2 5 3 1 1 4 1 1 53. Climacoraphistoma vaginati n n n n n T1 n n 11 n 1 1 n 2 2 3 1 2 4 2 2 1 4 1 1 54. Lesueurilla bipatellare n n n n n n n n n n 1 1 n 2 3 3 1 2 3 2 2 1 4 1 1 55. Lesueurilla marginalis n n n n n n n n n n 1 1 n 2 3 3 1 2 3 2 2 I 4 1 1 56. Lesueurilla prima n n n n n n n n n n 1 1 n 2 2 3 1 2 4 2 2 1 5 1 1 57. Palaeomphalus giganteus n n n n n n n n n n 1 2 2 2 2 3 2 2 5 2 2 1 4 1 1 58. Climacoraphistoma damesi n n n n n n n n n n 1 1 n 2 2 1 1 2 3 3 2 1 4 1 1 59. Eccyliopterus alatus n n n n n n n n n n 1 2 2 2 2 3 1 2 4 3 2 1 4 1 1 60. Eccyliopterus Iprinceps n n n n n n n n n n 1 1 n 2 2 2 1 2 5 2 3 1 6 1 2 61. Eccyliopterus regularis n n n n n n n n n n 1 1 n 2 2 3 1 2 4 3 2 1 5 1 1 62. Lesueurilla infundibula n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 6 1 1 63. Eccyliopterus louderbacki n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 5 1 1 64. Lesueurilla declivis n n n n n n n n n n 1 1 n 2 2 3 2 2 4 2 2 1 5 1 1 65. Pararaphistoma qualteriata n n n n n n n n n n 1 1 n 2 2 2 1 2 4 2 2 1 3 1 1 66. Pararaphistoma schmidti n n n n n n n n n n 1 1 n 2 2 1 1 2 5 2 2 1 3 1 1 67. Helicotoma gubanovi n n n n n n n n n n 1 2 2 2 2 2 1 2 1 2 2 1 2 1 1 68. Scalites katoi n n n n n n n n n n 1 3 2 2 2 3 2 2 1 2 2 1 2 1 1 69. Helicotoma medfraensis n n n n n n n n n n I 1 n 2 1 1 1 2 4 2 2 1 3 1 1 70. Lesueurilla scotica n n n n n n n n n n 1 1 n 2 2 1 1 2 4 2 2 1 4 1 1 71 . Pachystrophia devexa n n n n n n n n n n I I n 1 n n n n 3 2 2 1 3 1 1 72. Raphistoma striata n n n n n n n n n n 1 3 2 2 2 2 2 2 1 2 2 1 2 1 1 73. Raphistomina lapicida n n n n n n n n n n 1 I n 2 1 1 2 2 5 2 2 1 3 1 1 74. Scalites angulatus n n n n n n n n n n 1 2 2 ? ? ? ? ? 1 2 2 1 2 1 1 75. Holopea insignis n n n n n n n n n n 1 1 n 2 1 1 1 2 I 2 2 1 3 1 1 76. Eccyliopterus beloitensis n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 5 1 1 77. Holopea rotunda n n n n n n n n n ii 1 1 n 1 n n n n 2 2 2 1 2 1 1 78. Pachystrophia contigua n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 2 1 1 79. Pachystrophia spiralis n n n n n n n n n n 1 I n 1 n n n n 3 2 2 1 2 1 1 80. Raphistomina aperta n n n n n n n n n n 1 1 n 2 2 2 2 2 3 2 2 1 3 1 1 81. Raphistomina fissurata n n n n n n n n n n 1 1 n 2 1 I 2 2 5 2 2 1 2 1 1 82. Eccyliopterus owenanus n n n n n n n n n n 1 2 2 2 2 3 1 2 4 3 2 1 4 1 1 83. Holopea ampla n n n n n n n n n n 1 1 n I n n n n 2 2 2 1 2 1 1 84. Holopea pyrene n n n n n ii n n n n 1 1 n 2 1 i i 2 1 2 2 1 3 1 1 85. Holopea symmetrica n n n n n n n n n n 1 1 n 1 n n n n 2 2 2 1 2 1 1 86. Raphistoma peracuta n n n n n 11 n n n n 1 2 2 2 2 2 2 2 3 2 2 1 2 1 1 87. Raphistomina rugata n n n n n n n n n n 1 2 3 2 I 1 1 2 5 2 2 1 2 1 1 88. Raphistoma tellerensis n n n n n n n n n n 1 3 3 2 2 3 1 2 1 2 2 1 2 1 l 89. Sinutropis lesthetica n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 5 1 1 90. Pachystrophia gotlandica n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 2 I 1 91. Lytospira triquestra n n n n n n n n n n I 1 n 1 n n n n 3 2 2 1 7 1 1 NUMBER 88 111 Number Species 32. Maclurina logani 33. Maclurina manitobensis 34. Maclurites sedgewicki 35. Maclurites expansa 36. Ophileta complanata 37. Lecanospira compact a 38. Lecanospira nereine 39. Barnesella 'llecanospiroides 40. Malayaspira hintzei 41. Malayaspira rugosa 42. Barnesella measuresae 43. Lytospira angel ini 44. Lytospira yochelsoni 45. Maclurina lannulata 46. Rossospira harrisae 47. Ecculiomphalus bucklandi 48. Lytospira gerrula 49. Lytospira Inorvegica 50. Ophiletina cf. O sublaxa 51. Lytospira subrotunda 52. Pararaphistoma lemoni 53. Climacoraphistoma vaginati 54. Lesueurilla bipatellare 55. Lesueurilla marginalis 56. Lesueurilla prima 57. Palaeomphalus giganteus 58. Climacoraphistoma damesi 59. Eccyliopterus alatus 60. Eccyliopterus Iprinceps 61. Eccyliopterus regularis 62. Lesueurilla infundibula 63. Eccyliopterus louderbacki 64. Lesueurilla declivis 65. Pararaphistoma qualteriata 66. Pararaphistoma schmidti 67. Helicotoma gubanovi 68. Scalites katoi 69. Helicotoma medfraensis 70. Lesueurilla scotica 71. Pachystrophia devexa 72. Raphistoma striata 73. Raphistomina lapicida 74. Scalites angulatus 75. Holopea ins ignis 76. Eccyliopterus beloitensis 77. Holopea rotunda 78. Pachystrophia contigua 79. Pachystrophia spiralis 80. Raphistomina aperta 81. Raphistomina fissurata 82. Eccyliopterus owenanus 83. Holopea ampla 84. Holopea pyrene 85. Holopea symmetrica 86. Raphistoma peracuta 87. Raphistomina rugata 88. Raphistoma tellerensis 89. Sinutropis 'lesthetica 90. Pachystrophia gotlandica 9 1. Lytospira triquestra Appendix 2—Continued. 1 1 1 l 1 1 0 0 0 0 0 0 1 2 3 4 5 6 7 i 4 i i 7 14 11 ? 1 ? ? 1 7 13 11 7 13 11 7 14 11 7 17 7 1 7 14 11 7 14 11 7 13 11 7 14 11 7 2 4 11 7 2 4 11 7 14 11 7 14 11 7 14 11 7 2 4 11 114 11 7 14 11 7 2 4 11 7 2 3 11 7 13 11 7 13 11 7 17 7 1 7 13 11 7 2 3 1 1 7 14 11 7 2 4 1 1 12 4 11 7 2 4 1 1 7 13 11 7 13 11 7 2 3 1 1 7 13 11 7 13 11 7 13 11 7 14 11 7 13 11 7 13 11 7 14 11 7 1111 7 12 11 7 14 11 7 14 11 7 13 11 7 13 11 7 14 11 7 14 11 7 13 11 7 13 11 7 2 4 11 7 14 11 7 14 11 7 13 11 7 12 11 7 1 1&2 1 1 7 13 11 7 14 11 7 2 4 1 1 7 2 4 11 111111 0 0 0 1 1 1 7 8 9 0 12 1 I i 7 i 7 1117 1 ? 1117 1? I 1 1 7 1 7 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1117 1? 1 1 1 7 1 7 1 1 1 7 1 7 1117 1? 1117 1? 1 1 1 7 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 2 2 1 7 1 1 2 1 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 2 2 2 1 7 1 1 1 7 1 7 1 2 2 2 1 7 1 1 2 2 1 7 1 1 1 7 1 7 1 1 2 3 1 7 1 1 2 2 1 7 1 1 2 3 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 1 7 I 7 1 1 2 2 1 7 1 1 2 2 1 7 1 I 2 3 1 7 1 1 1 7 1 7 1 1 1 7 1 7 1 1 2 1 1 7 1 1 1 7 1 7 1 1 2 2 1 7 1 1 1 7 1 7 1 1 1 1 1 I 111111 4 5 6 7 8 9 7 i 2 1 i n~ 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 I 2 2 1 n 7 I 2 2 1 n 7 1 2 2 1 n 7 1 2 3 1 n 7 1 2 2 1 n 7 1 2 3 1 n 7 1 1 n 2 2 7 1 2 1 1 n 7 1 2 2 1 n 7 1 2 3 1 n 7 1 1 n 2 2 7 1 1 n 2 3 7 1 1 n 2 2 7 1 2 4 1 n 7 1 2 3 1 n 7 1 2 2 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 4 1 n 7 1 2 3 1 n 7 1 1 n 2 3 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 1 1 n 7 1 2 3 1 n 7 12 3 1" 7 1 2 3 1 n 7 1 7 n 1 n 7 1 2 3 1 n 7 1 7 n 7 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 1 1 n 7 1 2 3 1 n 7 1 2 2 1 n 7 12 3 1" 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 3 1 n 7 1 2 1 1 n 7 1 2 3 1 n 7 1 2 2 1 n 7 1 2 1 1 n 7 1 2 2 1 n 7 1 2 2 1 n 7 1 2 1 1 n 111111 2 2 2 2 2 2 0 12 3 4 5 15 2 3 12 15 2 3 12 15 2 3 1 2 1 5 3 3 1 2 2 2 2 3 12 2 2 2 3 12 7 2 2 3 12 2 3 2 3 1 2 2 4 2 3 1 2 2 2 3 3 12 2 3 2 3 12 2 2 2 12 2 2 12 0 12 2 4 2 4 12 2 2 2 1 1 2 2 2 2 1 12 2 12 0 12 2 17 0 12 2 3 2 3 2 2 12 2 12? 3 4 2 3 1 2 3 4 2 3 2 2 3 4 2 4 1 2 3 4 2 4 12 3 4 2 3 2 2 3 4 2 4 12 3 4 2 4 1 2 3 3 2 1 2 2 3 1 7 0 7 2 3 3 2 1 2 2 2 5 2 4 12 2 6 2 4 1 2 3 3 2 3&4 2 2 14 2 4 12 1 4 2 4 12 1 5 2 5 2 2 3 4 2 5 1 2 1 4 2 3 2 2 3 4 2 4 1 2 2 3 2 3 12 1 6 2 5 1 2 2 3 2 4 1 2 1 5 2 5 1 2 2 5 13 12 2 6 2 4 1 2 1 4 2 4 1 2 2 5 2 4 1 2 2 6 2 3 12 2 3 2 3 1 2 2 3 2 5 1 2 3 3 2 2 7 2 2 5 2 4 1 2 2 5 13 12 1 3 2 4 1 2 3 3 2 3 1 2 2 4 2 3 1 2 3 4 2 5 1 2 2 5 2 3 1 2 2 5 2 3 1 2 2 5 2 112 112 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2— Continued. 1 1 1 1 1 1 1 1 1 1 1 1 i 1 i 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 32. Maclurina logani 2 1 3 3 3 3 1 1 n n n n 1 1 1 4 ? n 33. Maclurina manitobensis 2 1 3 2 3 3 1 1 n n n n 1 1 1 4 ? n 34. Maclurites sedgewicki 2 1 3 2 3 3 1 1 n n n n 1 1 1 4 ? n 35. Maclurites expansa 2 1 3 1 n n n 1 n n n n 1 1 1 4 ? n 36. Ophileta complanata 4 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 37. Lecanospira compacta 2 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 38. Lecanospira nereine 2 1 2 1 n n n I n n n n 1 1 1 2 ? n 39. Barnesella llecanospiroides 2 1 ? 1 n n n 1 n n n n 1 1 I 2 ? n 40. Malayaspira hintzei 3 1 ? 1 n n n 1 n n n n 1 1 1 1 ? n 41. Malayaspira rugosa 3 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 42. Barnesella measuresae 3 1 ? 1 n n n 1 n n ii n 1 1 1 2 ? n 43. Lytospira angelini 2 1 2 1 n n n 1 n n n n 1 1 2 2 ? n 44. Lytospira yochelsoni 2 ? 2 1 n n n 1 n n n n 1 1 2 3 ? n 45. Maclurina lannulata 2 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 46. Rossospira harrisae 3 1 2 1 n n n 1 n n n n 1 1 1 1 ? n 47. Ecculiomphalus bucklandi 3 1 2 1 n n n 1 n n n n 1 1 1 3 ? n 48. Lytospira gerrula 2 ? ? 1 n n n 1 n n n n 1 1 2 3 ? n 49. Lytospira Inorvegica 2 ? 2 1 n n n 1 n n n n 1 1 2 2 ? n 50. Ophiletina cf. 0. sublaxa 3 1 ? 1 n n n 1 n n n n 1 I 1 1 ? n 51. Lytospira subrotunda 2 ? 2 1 n n n 1 n n n n 1 1 2 3 7 n 52. Pararaphistoma lemoni 4 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 53. Climacoraphistoma vaginati 3 2 2 1 n n n 1 n n u n 1 1 1 2 ? n 54. Lesueurilla bipatellare 3 1 ? 1 n n n 1 n n n n 1 1 1 1 ? n 55. Lesueurilla marginalis 3 1 ? 1 n n n 1 n n n n 1 1 1 1 ? n 56. Lesueurilla prima 3 2 2 1 n n n 1 n n n n 1 1 1 2 ? n 57. Palaeomphalus giganteus 4 3 2 1 n n n 1 n n n n 1 1 1 2 7 n 58. Climacoraphistoma damesi 3 2 2 1 n n n I n n n n 1 1 1 2 ? n 59. Eccyliopterus alatus 3 2 2 1 n n n 1 n n n n 1 1 I 2 7 3 60. Eccyliopterus Iprinceps 3 ? 2 1 n n n 1 n n n n 1 1 1 2 ? n 61. Eccyliopterus regularis 3 2 2 1 n n n 1 n n n n 1 1 1 3 ? n 62. Lesueurilla infundibula 3 2 2 1 n n n I n n n n 1 1 1 3 ? n 63. Eccyliopterus louderbacki 3 1 ? 1 n n n 1 n n n n 1 1 1 3 ? n 64. Lesueurilla declivis 3 2 2 1 n n n 1 n n n n 1 1 1 2 ? n 65. Pararaphistoma qualteriata 4 2 2 1 n n n 1 n n n n 1 1 1 3 ? 3 66. Pararaphistoma schmidti 4 2 2 1 n n n 1 n n n ii 1 1 1 2 ? n 67. Helicotoma gubanovi 5 2 ? 2 2 3 1 1 n n n n 1 1 1 2 7 n 68. Scalites katoi 5 2 ? 1 n n n 1 n n n n 1 1 1 3 ? n 69. Helicotoma medfraensis 4 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 70. Lesueurilla scotica 3 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 1 1. Pachystrophia devexa 4 1 2 1 n n n 1 n n n n 1 n 1 2 7 n 72. Raphistoma striata 5 3 1 1 n n n 1 n n n n 1 i 1 3 7 n 73. Raphistomina lapicida 4 I 2 I n n n 1 n n n n 1 i 1 2 ? n 74. Scalites angulatus 5 2 ? 1 n n n 1 n n n n 1 i 1 3 ? n 75. Holopea ins ignis 5 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 76. Eccyliopterus beloitensis 3 1 ? 1 n n n 1 n n n n 1 i 1 3 ? n 77. Holopea rotunda 6 2 1 I n n n 1 n n n n 1 i 1 2 ? n 78. Pachystrophia contigua 4 1 ? 1 n n n 1 n n n n 1 i 1 2 ? n 79. Pachystrophia spiralis 4 1 2 2 4 i i 2 1 1 i 1 1 i 1 2 ? n 80. Raphistomina aperta 5 1 2 1 n n n 1 n n n n 1 i 1 2 ? n 81. Raphistomina fissurata 5 1 2 1 n n n 1 n n n n 1 i 1 2 ? n 82. Eccyliopterus owenanus 3 ? ? 1 n n n 1 n n n n 1 i I 2 ? n 83. Holopea ampla 5 1 ? 1 n n n 1 n n n n 1 i 1 2 ? n 84. Holopea pyrene 5 1 2 1 n n n 1 n n n n 1 i 1 2 7 n 85. Holopea symmetrica 6 2 1 1 n n n 1 n n n n 1 i 1 1 ? n 86. Raphistoma peracuta 4 1 ? 1 n n n 1 n n n n I i 1 2 ? n 87. Raphistomina rugata 4 1 1 1 n n n 1 n n n n 1 i 1 1 ? n 88. Raphistoma tellerensis 5 2 2 2 3 i I 2 3 1 1 i 1 i 1 3 ? n 89. Sinutropis lesthetica 4 1 2 2 4 i 1 2 4 1 1 i I n 1 2 ? n 90. Pachystrophia gotlandica 4 1 2 3 1 2 1 1 n n n n I n 1 2 ? n 91. Lytospira triquestra 2 1 2 1 n n n l n n n n 1 n I 3 ? 3 NUMBER 88 113 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 92. Euomphalus tubus 2 2 2 2 2 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 93. Lytospira subuloides 2 2 2 2 2 2 2 1 1 3 1 1 1 n 2 1 1 1 1 n n n n n n 94. Ceratopea unguis 2 2 5 5 2 3 3 2 4 4 1 1 1 n 3 1 1 2 2 3 1 n n n n 95. Boucotspira aff. B. fimbriata 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 2 1 n n n n 96. Lophonema peccatonica 2 2 5 5 2 3 3 2 4 4 1 1 2 i 3 1 1 1 2 2 1 n n n n 97. Polehemia taneyensis 2 2 5 5 2 3 3 2 3 3 1 1 2 l 3 1 1 1 2 2 1 n n n n 98. Walcottoma frydai 2 2 5 5 2 3 3 2 3 3 1 1 2 i 3 1 1 1 2 2 1 n n n n 99. Helicotoma planulata 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 3 1 n n n n 100. Helicotoma tennesseensis 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 3 1 n n n n 101. Ophiletina sublaxa 2 2 3 3 2 2 2 2 3 3 1 1 2 i 4 1 1 1 2 3 2 i 2 1 n 102. Ophiletina angularis 2 2 3 3 2 2 2 2 3 3 1 1 2 i 4 1 1 1 2 3 2 i 2 1 n 103. Oriostoma bromidensis 2 2 2 2 2 1 1 2 2 2 1 1 2 i 4 1 1 1 2 2 1 n n n n 104. Euomphalopterus lordovicius 2 2 2 2 2 3 3 2 2 2 1 1 2 i 3 1 1 1 2 1 1 n n n n 105. Euomphalopterus aff. E. ordovicius 2 2 2 2 2 3 3 2 2 2 1 1 2 i 3 1 1 1 2 1 1 n n n n 106. Euomphalopterus cariniferus 2 2 2 2 2 3 3 2 2 2 1 1 2 i 2 1 1 1 2 1 1 n n n n 107. Palaeomphalus Igradatus 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 1 1 n n n n 108. Trochomphalus Idimidiatus 2 2 3 3 2 3 3 2 1 1 1 1 2 i 3 1 1 1 2 1 1 n n n n 109. Helicotoma blodgetti 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 3 1 n n n n 110. Helicotoma robinsoni 2 2 3 3 2 3 3 2 3 3 1 1 2 i 2 1 1 1 2 3 1 n n n n 111. Helicotomal Girvan sp. 2 2 4 4 2 3 3 2 3 3 1 1 2 i 3 1 1 1 2 1 1 n n n n 112. Straporillina cf. S. circe 2 2 3 3 2 3 3 2 2 2 1 1 2 i 2 1 1 1 2 1 1 n n n n 113. Euomphalopterus alatus 2 2 1 1 2 3 3 2 1 1 1 1 2 i 4 1 2 1 1&2 1 1 n n n n 114. Euomphalopterus frenatus 1 2 ? ? 2 3 3 2 2 2 1 1 2 i 4 1 2 1 2 1 1 n n n n 115. Euomphalopterus praetextus 2 2 1 1 2 3 3 2 1 1 1 1 2 i 3 1 1 1 1 11 n n n n n 116. Euomphalopterus subcarinatus 2 2 1 1 2 3 3 2 2 2 1 1 2 i 4 1 2 1 2 1 i n n n n 117. Euomphalopterus togatus 1 2 ? ? 2 3 3 2 2 2 1 1 2 i 4 1 2 1 2 1 i n n n n 118. Euomphalopterus undulans 2 2 1 1 2 2 2 2 1 1 1 1 2 i 3 1 1 1 1 n n n n n n 119. Grantlandispira christei 2 2 3 3 3 2 3 2 1 1 1 1 2 i 3 1 1 1 1 n n n n n n 120. Poleumita discors 2 2 0 0 2 3 3 2 2 2 1 2 2 2 4 1 2 1 2 1 i n n n n 121. Pycnomphalus acutus 2 2 1 1 2 3 3 2 1 1 1 1 2 1 3 1 1 1 1 n n n n n n 122. Pycnomphalus obesus 2 2 1 1 2 3 3 2 1 1 1 1 2 1 3 1 1 1 1 n n n n n n 123. Discordichilus dalli 1 n n n n n n n n n n 1 2 2 3 I 1 1 1 n n n n n n 124. Discordichilus mollis 1 n n n n n n n n n n 1 2 2 3 1 1 1 1 n n n n n n 125. Discordichilus kolmodini 1 n n n n n n n n n n 1 2 2 4 1 1 1 1 n n n n n n 126. Poleumita alata 2 2 0 0 2 3 3 2 2 2 i 2 2 2 4 1 1 1 2 i i n n n n 127. Poleumita octavia 2 2 0 0 2 3 3 2 2 2 i 2 2 1 5 1 4 1 2 i i n n n n 128. Poleumita rugosa 2 2 0 0 2 3 3 2 2 2 i 2 2 2 5 1 4 1 2 i i n n n n 129. Pseudophorus profundus 1 ? 7 ? ? ? ? 7 ? ? ? ? 2 2 4 1 1 2 1 n n n n n n 130. Pseudophorus stuxbergi ? ? ? ? ? ? ? ? ? 7 ? 1 2 2 4 1 1 1 1 n n n n n n 131. Siluriphorus gotlandicus ? ? 7 ? ? 7 ? ? 7 ? ? 1 2 2 4 1 1 1 1 n n n n n n 132. Siluriphorus undulans ? ? 7 ? 7 ? ? 7 7 7 ? 2 2 2 5 1 1 1 1 n n n n n n 133. Streptotrochus incisus 1 ? ? ? ? 7 ? 2 7 ? ? 1 2 1 2 1 1 1 2 i i n n n n 134. Streptotrochus aff. S. incisus 1 ? 7 7 ? ? 7 2 ? ? ? 1 2 1 2 1 1 1 1 n n n n n n 135. Streptotrochus lamellosa 1 ? 7 ? ? ? ? ? ? 7 7 1 2 1 4 1 1 1 2 i i n n n n 136. Streptotrochus lundgreni 1 ? ? ? ? ? ? 2 ? 7 ? 1 2 1 2 1 1 1 2 i i n n n n 137. Streptotrochus ? visbeyensis 2 2 1 1 2 1 1 2 2 2 1 1 2 1 4 1 1 1 2 i i n n n n 138. Hystricoceras astraciformis 1 ? ? ? ? 7 ? ? 7 7 ? 1 2 1 2 1 1 2 2 i i n n n n 139. Poleumita granulosa 2 2 0 0 2 3 3 2 2 2 1 2 2 2 4 1 2 1 2 2 i n n n n 140. Euomphalus walmstedti 2 2 0 0 2 3 3 2 2 2 1 2 2 2 4 1 1 1 2 1 i n n n n 141. Centrifugus planorbis 2 1 5 3 2 3 1 2 4 4 1 2 2 2 4 1 2 1 2 1 2 1 i 1 n 142. Spinicharybdis wilsoni 1 ? ? ? ? ? 7 ? ? 7 ? 1 2 1 4 1 2 1 ? n n n n n n 143. Turbocheilus immaturum 2 1 0 0 2 3 3 2 1 1 1 1 2 1 3 1 1 1 1 n n n n n n 144. Pseudotectus comes 1 ? 7 ? ? ? ? ? ? ? ? 1 2 2 4 1 1 1 1 n n n n n n 145. Straparollus bohemicus 1 ? ? ? ? ? 7 ? ? ? ? ? ? 2 4 1 1 1 1 n n n n n n 146. Hormotoma artemesia 2 3 4 6 3 3 4 1 3 4 1 1 1 n 3 1 1 1 2 5 2 1 2 1 n 147. Hormotoma confusa 2 3 4 6 3 3 3 2 3 3 1 1 1 n 2 1 1 1 2 5 2 1 2 1 n 148. Hormotoma Idubia 2 2 4 4 2 2 2 2 3 3 1 1 1 n 2 1 1 1 2 3 2 1 2 1 n 149. Hormotoma Isimulatrix 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 5 2 1 2 1 n 150. Ectomaria adelina 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 1 n 151. “Hormotoma" “ cassina ” 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 4 2 2 3 1 n 114 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 92. Euomphalus tubus n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 A 1 n 93. Lytospira subuloides n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 7 1 n 94. Ceratopea unguis n 2 1 3 1 1 1 1 1 n n 1 1 2 1 n 2 2 1 2 1 1 5 3 1 95. Boucotspira aff. B. fimbriata n 2 2 3 1 1 1 1 1 n n 1 4 2 1 n 1 2 1 2 2 1 7 1 n 96. Lophonema peccatonica n 2 1 3 1 1 1 1 1 n n 1 1 1 1 n 1 2 1 2 2 1 6 3 1 97. Polehemia taneyensis n 2 1 3 1 1 1 1 1 n n 1 I 1 1 n 1 2 1 2 2 1 6 3 1 98. Walcottoma frydai n 2 2 3 1 1 1 1 1 n n 1 1 1 1 n 1 2 l 2 2 1 8 1 n 99. Helicotoma planulata n 2 3 4 1 1 1 1 1 n n 1 4 2 1 n 2 2 1 2 1 1 8 1 n 100. Helicotoma tennesseensis n 2 3 4 1 1 1 1 1 n n 1 4 2 1 n 2 2 1 2 1 1 8 1 n 101. Ophiletina sublaxa n 2 3 3 1 1 1 1 1 n n 1 1 3 1 n 2 2 1 2 1 1 7 1 n 102. Ophiletina angularis ii 2 3 3 1 1 1 1 2 2 i 1 1 3 1 n 2 2 1 2 1 1 7 1 n 103. Oriostoma bromidensis n 2 2 4 1 1 1 1 1 n n 1 4 2 I n 1 2 1 2 1 1 A 1 n 104. Euomphalopterus lordovicius n 2 2 4 1 1 1 1 1 n n 1 4 2 1 n 1 2 1 2 2 1 6 1 n 105. Euomphalopterus aff. E. ordovicius n 2 2 4 1 1 1 1 1 n n 1 4 2 1 n 1 2 1 2 2 1 6 1 n 106. Euomphalopterus cariniferus n 2 2 3 1 1 1 1 1 n n 1 4 2 1 n 1 2 1 2 2 1 8 1 n 107. Palaeomphalus Igradatus n 2 1 3 1 1 1 1 1 n n 1 4 2 1 n 1 2 1 2 1 1 8 1 n 108. Trochomphalus Idimidiatus n 2 2 2 1 1 1 1 1 n n 1 1 2 1 n 1 2 1 ? n 1 9 1 n 109. Helicotoma blodgetti n 2 3 4 1 1 1 1 1 n n 1 n 2 1 n 2 2 1 1 i 1 9 1 n 110. Helicotoma robinsoni n 2 3 4 1 1 1 1 1 n n 1 4 2 1 n 2 2 1 1 n 1 6 1 n 111. Helicotomal Girvan sp. n 2 3 4 1 1 1 1 1 n n 1 4 2 1 n 2 2 1 2 i 1 7 1 n 112. Straporillina cf. S'. circe n 2 1 1 1 1 1 1 1 n n 1 1 2 1 n 1 2 1 2 i 1 9 1 n 113. Euomphalopterus alatus n 2 2 1&3 2 1 1 1 1 n n 1 1&4 2 1 n 1 2 1 2 2 1 9 1 n 114. Euomphalopterus frenatus n 2 5 1 2 1 1 1 1 n n 1 4 2 1 n 1 2 1 1 n 1 A 1 n 115. Euomphalopterus praetextus n n n n n n n n 1 n n 1 1 2 n n n n n 1 n 1 9 1 n 116. Euomphalopterus subcarinatus n 2 2 3 2 i i i 1 n n 1 4 2 i n 1 2 1 2 2 1 8 1 n 117. Euomphalopterus togatus n 2 5 3 2 i i i 1 n n 1 4 2 i n 1 2 1 2 1 1 C 1 n 118. Euomphalopterus undulans n n n n n n n n 1 n n 1 1 2 n n n n n 1 n 1 9 1 n 119. Grantlandispira christei n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 A 1 n 120. Poleumita discors n 2 2 3 i 1 2 1 1 n n i 4 2 i n i 2 i 2 1 1 C 1 n 121. Pycnomphalus acutus n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 A 1 n 122. Pycnomphalus obesus n n n n n n n n 1 n n n 1 2 11 n n n n 1 n 1 A 1 n 123. Discordichilus dalli n n n n n n n n 1 n n i 5 2 n n n n n 1 n 1 C 1 n 124. Discordichilus mollis n n n n n n n n 1 n n i 5 2 n n n n n 1 n 1 C 1 n 125. Discordichilus kolmodini n n n n n n n n 1 n n i 5 2 n n n n n 1 n 1 D 1 n 126. Poleumita alata n 2 2 3 1 i i i 1 n n i 4 2 i n i 2 i 2 2 1 9 1 n 127. Poleumita octavia n 2 2 3 1 i 2 i 1 n n i 4 2 i n i 2 i 2 1 1 C 1 n 128. Poleumita rugosa n 2 2 3 1 i 2 i 1 n n i 4 2 i n i 2 i 2 1 1 c 1 n 129. Pseudophorus profundus n n n n n n n n 1 n n i 5 n n n n n n 1 n 1 B 1 n 130. Pseudophorus stuxbergi n n n n n n n n 1 n n i 5 2 n n n n n 1 n 1 9 1 n 131. Siluriphorus gotlandicus n n n n n n n n 1 n n i 5 2 n n n n n 1 n 1 C 1 n 132. Siluriphorus undulans n i n n n i i 1 1 n n i 5 2 i n i 2 i 1 n 1 c 1 n 133. Streptotrochus incisus n 2 5 1 2 i i 1 1 n n i 6 2 i n i 2 i 2 i 1 c 1 n 134. Streptotrochus aff. S. incisus n n n n n n n n 1 n n i 6 2 n n n n n 1 n 1 c 1 n 135. Streptotrochus lamellosa n 2 2 4 i i i i 1 n n i 4 2 i n 1 2 1 2 2 1 8 1 n 136. Streptotrochus lundgreni n 2 5 1 2 i i i 1 n n i 6 2 n n n n n 1 n 1 B 1 n 137. Streptotrochus ? visbeyensis n 2 2 3 1 i i i 1 n n i 4 2 i n 1 2 i 2 2 1 A 1 n 138. Hystricoceras astraciformis n 2 5 1 2 i i i 1 n n i 6 2 i n 1 2 i 1 n 1 B 1 n 139. Poleumita granulosa n 2 2 4 1 i 2 i 1 n n i 4 2 i n 1 2 i 2 i 1 C 1 n 140. Euomphalus walmstedti n 2 2 2 1 i 1 i 1 n n i 4 2 i n 1 2 i 2 i 1 B 1 n 141. Centrifuges planorbis n 1 n n n n n n 2 2 i i 1 1 i i 1 2 i 1 n 1 F 1 n 142. Spinicharybdis wilsoni n n n n n n n n 1 n n i n n n n n n n ? n 1 9 1 n 143. Turbocheilus immaturum n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 A 1 n 144. Pseudotectus comes n n n n n n n n 1 n n 1 5 2 n n n n n 1 n 1 C 1 n 145. Straparollus bohemicus n n n n n n n n 1 n n 1 5 2 n n n n n ? n l A 1 n 146. Hormotoma artemesia 1 1 n n n n n 2 1 n n 1 1 2 1 1 1 2 1 1 n 1 8 1 n 147. Hormotoma confusa 1 n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 8 1 n 148. Hormotoma Idubia 1 2 1 2 n 1 1 3 1 n n 1 1 2 2 2 1 2 1 2 i 1 7 1 n 149. Hormotoma Isimulatrix 1 1 n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 7 1 n 150. Ectomaria adelina 1 1 n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 9 1 n 151. “ Hormotoma ” “ cassina ” 1 1 n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 8 1 n NUMBER 88 115 Appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 6 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 92. Euomphalus tubus 3 2 2 3 2 7 1 4 3 1 n n 1 1 n n n n 1 n 1 1 1 n 1 93. Lytospira subuloides 3 3 2 2 4 4 1 5 4 1 n n 1 1 n n n n 7 n 1 2 1 n 1 94. Ceratopea unguis 1 2?3 2 3 2?3 3 1 1-3 2 2 2 3 1 1 n n n n 1 n 1 3 1 n 1 95. Boucotspira aff. B. fimbriata 2 3 2 3 2 3 2 4 4 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 96. Lophonema peccatonica 2 3 2 3 3 4 1 4 3 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 97. Polehemia taneyensis 2 3 2 3 3 4 1 4 3 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 98. Walcottoma fiydai 2 3 2 3 2 5 1 5 3 1 n n 1 2 1 2 1 n 1 n 1 3 1 n 2 99. Helicotoma planulata 2 3 2 3 3 4 1 4 2 1 n n 1 2 1 3 2 2 1 n 1 3 2 1 2 100. Helicotoma tennesseensis 2 3 2 3 3 4 2 4 4 1 n n 1 2 1 3 2 2 1 n 1 3 2 1 2 101. Ophiletina sublaxa 2 3 2 3 4 8 1 4 3 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 102. Ophiletina angularis 2 3 2 3 4 8 1 4 3 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 103. Oriostoma bromidensis 2 3 2 3 3 4 2 3&4 3 1 n n 1 2 3 3 2 2 1 n 1 3 1 n 2 104. Euomphalopterus lordovicius 2 3 2 3 1 2 2 5 5 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 105. Euomphalopterus aff. E. ordovicius 2 3 2 3 1 2 2 5 5 1 n n 1 2 3 4 1 2 1 n 1 3 I n 2 106. Euomphalopterus carini/erus 2 3 2 3 2 4 2 5 5 1 n n 1 2 3 4 1 2 1 n 1 3 1 n 2 107. Palaeomphalus Igradatus 2 3 2 3 3 5 2 4 4 1 n n 1 1 n n n n 1 n 1 3 2 1 2 108. Trochomphalus Idimidiatus 2 2 2 3 1 4 2 5 5 1 n n 1 2 2 4 i i 1 n 1 3 1 n 2 109. Helicotoma blodgetti 2 3 2 1 4 2 1 3 2 1 n n 1 2 2 3 2 2 1 n 1 3 2 i 2 110. Helicotoma robinsoni 2 3 2 1 3 2 1 4 2 1 n n 1 2 2 3 2 2 1 n 1 3 2 i 2 111. Helicotoma ? Girvan sp. 2 3 2 3 3 4 2 4 4 1 n n 1 2 1 3 2 2 1 n 1 3 2 i 2 112. Straporillina cf. S. circe 2 2 2 3 1 4 2 5 5 1 n n 1 1 n n n n 1 n 1 2 1 n 2 113. Euomphalopterus alatus 2 1 3 3 3 5 2 5 5 1 n n 1 2 1&2 4 2 2 1 n 1 1 1 n 2 114. Euomphalopterus frenatus 2 1 7 3 1 4 1 7 4 7 n n 1 1 n n n n l n 1 1 1 n 2 115. Euomphalopterus praetextus 2 1 1 3 3 4 1 5 5 1 n n 1 2 i i i 2 1 n ] 3 1 n 2 116. Euomphalopterus subcarinatus 2 3 3 3 3 4 1 5 4 1 n n 1 2 3 4 2 2 1 n 1 3 1 n 2 117. Euomphalopterus togatus 2 3 3 3 2 4 1 6 5 1 n n 1 2 2 4 2 2 1 n 1 3 1 n 2 118. Euomphalopterus undulans 2 1 2 3 1 4 1 6 4 1 n n 1 1 n n n n 1 n 1 1 1 n 2 119. Grantlandispira christei 2 2 2 3 0 5 2 6 6 1 n n 1 1 n n n n 1 n 1 2 1 n 2 120. Poleumita discors 2 3 2 3 2 5 2 5 5 1 n n 1 2 i 4 i 2 1 n 1 3 1 n 2 121. Pycnomphalus acutus 2 2 2 3 0 5 2 6 6 1 n n 1 1 n n n n 1 n 1 2 1 n 2 122. Pycnomphalus obesus 2 2 2 3 0 5 2 6 6 1 n ■ i 1 1 n n n n 1 n 1 2 1 n 1 123. Discordichilus dalli 2 2 2 3 1 5 1 6 5 1 n n 1 1 n n n n 1 n 1 2 1 n 2 124. Discordichilus mollis 2 2 2 3 1 5 1 6 4 1 n n 1 1 n n n n 2 1 2 2 1 n 2 125. Discordichilus kolmodini 2 2 2 3 1 7 1 8 6 2 n n 1 1 n n n n 2 1 2 2 1 n 2 126. Poleumita alata 2 3 2 3 2 4 2 5 5 1 n n 1 2 3 4 1 2 1 n ] 3 1 n 2 127. Poleumita octavia 2 2 2 3 2 4 2 5 5 1 n n 1 2 1 1 1 1 1 n 2 1 n 1 128. Poleumita rugosa 2 2 2 3 2 5 2 5 5 1 n n 1 2 1 1 1 2 1 n 1 2 1 n 2 129. Pseudophorus profundus 7 7 7 3 0 6 2 5 5 1 n n 1 1 n n n n 2 1 4 1 n 2 130. Pseudophorus stuxbergi 2 3 3 3 2 4 2 5 5 1 n n 1 1 n n n n 2 1 3 ? n 2 131. Siluriphorus gotlandicus 2 3 2 3 2 4 1 7 5 1 n n 1 1 n n n n 2 1 3 1 n 2 132. Siluriphorus undulans 2 3 2 3 2 4 1 7 5 1 n n 1 1 n n n n 2 1 3 1 n 2 133. Streptotrochus incisus 2 3 3 3 2 4 1 6 5 1 n n 1 1 n n n n 1 n 3 1 n 2 134. Streptotrochus aff. S. incisus 2 3 2 3 2 4 1 6 5 1 n n 1 1 n n n n 1 n 3 1 n 2 135. Streptotrochus lamellosa 2 3 3 2 3 3 2 6 6 1 n n 1 2 3 4 i 2 1 n 3 1 n 2 136. Streptotrochus lundgreni 2 2 3 3 2 4 1 7 5 1 n n 1 1 n n n n 1 n 2 1 n 2 137. Streptotrochus ? visbeyensis 2 3 3 3 2 3 2 5 5 1 n n 1 2 3 4 1 2 1 n 3 1 n 2 138. Hyslricoceras astraciformis 2 2 3 3 2 5 1 7 5 1 n n 1 1 n n n n 1 n 2 1 n 2 139. Poleumita granulosa 2 3 2 3 3 5 1 5 6 1 n n 1 2 1 4 i 2 1 n 4 1 n 2 140. Euomphalus yvalmstedti 2 3 2 3 2 5 2 5 5 1 n n 1 2 1 4 i 2 1 n 1 2 1 n 1 141. Centrifugus planorbis 2 3 2 3 0 9 1 7 1 1 n n 1 1 n n n n 1 n 1 3 1 n 2 142. Spinicharybdis wilsoni 2 1 1 3 1 4 1 7 4 1 n n 1 1 n n n n 1 n 1 1 1 n 2 143. Turbocheilus immaturum 2 2 2 3 0 5 2 6 6 1 n n 1 1 n n n n 1 n 1 1 1 n 1 144. Pseudotectus comes 2 2 2 3 1 6 1 8 6 1 n n 1 1 n n n n 2 1 2 2 1 n 2 145. Straparollus bohemicus 2 2 2 3 2 4 2 5 5 1 n n 1 2 1 1 1 2 1 n 1 2 1 n 1 146. Hormotoma artemesia 3 2 2 1 5 4 1 5 5 2 2 3 1 1 n n n n 1 n 1 1 2 2 1 147. Hormotoma confusa 2 2 2 1 5 4 2 4&5 5 2 1 n 1 1 n n n n 1 n 1 2 2 1 1 148. Hormotoma Idubia 3 3 2 2 5 5 2 5 5 2 1 3 1 1 n n n n 1 n 1 3 2 1 1 149. Hormotoma 'Isimulatrix 2 2 2 1 5 4 2 5 5 2 1 3 1 1 n n n n 1 n 1 2 2 1 1 150. Ectomaria adelina 2 3 2 3 4 5 1 4 2 1 n n 1 2 1 1 1 1 1 n 1 3 1 n 2 151. “ Hormotoma '’ “ cassina ” 2 3 2 3 5 5 1 4 3 1 n n 1 2 1 1 1 1 1 n 1 3 l n 2 116 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 7 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 1 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 92. Euomphalus tubus n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 3 1 1 93. Lytospira subuloides n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 7 1 1 94. Ceratopea unguis n n n n n n n n n n 1 2 2 2 3 1&3 2 i 4 2 2 1 4 1 1 95. Boucotspira aff. B. fimbriata 2 2 2 n n n n n n 1 1 1 n 2 3 1 1 2 3 2 1 1 4 1 1 96. Lophonema peccatonica 2 2 2 n n n n n n 1 1 1 n 2 3 1 1 2 3 2 2 1 4 1 1 97. Polehemia taneyensis 2 2 2 n n n n n n 1 1 1 n 2 3 1 1 2 3 2 2 1 4 1 1 98. Walcottoma frydai 2 2 2 n n n n n n 1 1 1 n 2 3 1 1 2 3 2 2 1 3 1 1 99. Helicotoma planulata 1 1 2 n n n n n n 1 1 2 3 2 1 1 1 2 3 2 2 1 4 1 1 100. Helicotoma tennesseensis 1 1 2 n n n n n n 1 1 1 n 2 1 1 2 2 3 2 2 1 4 1 1 101. Ophiletina sublaxa 4 3 2 1 i 1 i 2 1 1 1 2 2 2 3 3 1 3 5 2 2 1 4 1 1 102. Ophiletina angularis 4 3 2 1 i 1 i 2 1 1 1 2 2 2 3 3 1 3 5 2 2 1 4 I 1 103. Oriostoma bromidensis 1 2 2 n n n n n n 1 1 1 n 2 3 3 3 2 3 2 2 1 3 1 1 104. Euomphalopterus lordovicius 3 3 2 i 1 i i 3 i 1 1 3 2 2 4 1 2 2 5 1 1 1 4 1 1 105. Euomphalopterus aff. E. ordovicius 3 3 2 i 1 i i 3 i 1 1 3 2 2 4 1 2 2 5 1 1 1 4 1 1 106. Euomphalopterus cariniferus 3 3 2 i 1 i i 3 i 1 1 1 n 2 4 1 2 2 5 2 2 1 4 1 1 107. Palaeomphalus Igradatus 1 1 2 n n n n n n 1 1 1 n 1 n n n n 3 1 2 1 4 1 1 108. Trochomphalus Idimidiatus 2 3 1 n n n n n n n 1 2 2 1 n n n n 3 2 1 1 3 1 2 109. Helicotoma blodgetti 4 3 2 n n n n n n i 1 2 3 2 i 1 i 2 3 2 2 I 3 1 1 110. Helicotoma robinsoni 4 3 2 n 11 n n n n i 1 2 3 2 i 1 i 2 3 2 2 1 5 1 1 111. Helicotomal Girvan sp. 1 2 2 n n n n n n i 1 1 n 1 n n n n 3 1 2 1 4 1 1 112. Straporillina cf. S. circe 1 1 n n n n n n n n 1 2 2 1 n n n n 3 2 1 1 3 1 2 113. Euomphalopterus alatus 3 4 2 2 1 i 2 3 i i 1 1 n 2 1&4 i 2 2 4 2 2 1 3&4 1 1 114. Euomphalopterus frenatus 3 4 2 2 2 2 2 3 i 2 1 1 n 1 n n n n 4 2 2 1 3 1 1 115. Euomphalopterus praetextus 3 4 2 2 2 2 2 3 i 1 1 1 n 2 i i 2 i 4 1 2 1 3 1 1 116. Euomphalopterus subcarinatus 3 4 2 2 1 1 2 3 i 1 1 1 n 2 4 i 2 2 5 2 2 1 4 1 1 117. Euomphalopterus togatus 3 4 2 2 2 1 2 3 i 2 1 1 n 2 1 i 2 1 5 2 2 1 2 1 1 118. Euomphalopterus undulans 3 4 2 2 1 2 2 3 i 1 1 1 n 1 n n n n 3 1 2 1 3 1 1 119. Grantlandispira christei 2 3 1 n n n n n n 1 1 3 2 1 n n n n 3 2 1 1 3 1 2 120. Poleumita discors 1 3 2 n n n n n n 1 1 2 2 1 n n n n 3 3 1 1 3 1 1 121. Pycnomphalus acutus 3 3 1 1 i 1 i 3 i 1 1 3 2 1 n n n n 3 1 1 1 3 I 2 122. Pycnomphalus obesus n n n n n n n n n n 1 3 2 1 n n n n 3 1 1 1 3 1 2 123. Discordichilus dalli i 2 2 n n n n n n n 1 2 2 1 n n n n 5 2 3 1 2 1 1 124. Discordichilus mollis i 1 2 n n n n n n n 1 2 1 1 n n n n 6 2 3 1 2 1 1 125. Discordichilus kolmodini i 1 2 n n n n n n n 1 2 1 1 n n n n 6 3 1 1 1 1 1 126. Poleumita alata 3 3 2 1 1 1 i 3 i 1 1 1 n 2 4 1 2 2 4 3 1 1 3 1 1 127. Poleumita octavia n n n n n n n n n n 1 2 2 1 n n n n 3 3 1 1 1 1 1 128. Poleumita rugosa i 3 2 n n n n n n n 1 2 2 1 n n n n 3 3 1 1 3 1 1 129. Pseudophorus profundus 3 4 2 2 1 i i 3 2 i 1 2 3 2 3 i 2 1 5 2 2 1 3 1 1 1 30. Pseudophorus stuxbergi 3 4 2 2 1 i i 3 2 i 1 2 3 2 3 i 2 1 5 2 2 1 3 1 1 131. Siluriphorus gotlandicus 4 3 2 1 1 i i 3 1 i 1 2 2 2 3 i 2 1 5 2 2 1 2 1 1 132. Siluriphorus undulans 4 3 1 n n n n n n i 1 2 2 1 n n n n 4 2 2 1 2 1 1 133. Streptotrochus incisus 2 2 2 n n n n n n i 1 2 2 1 n n n n 5 2 3 1 2 1 1 134. Streptotrochus aff. S. incisus 2 2 2 n n n n n n i 1 2 2 1 n 11 n n 5 2 2 1 2 1 1 135. Streptotrochus lamellosa 2 3 2 i i i i 3 i i 1 2 2 1 n n n n 4 2 2 1 2 1 1 136. Streptotrochus lundgreni 2 4 2 n n n n n n i 1 2 2 1 n n n n 5 2 3 1 2 1 1 137. Streptotrochus ? visbeyensis 2 2 2 n n n n n n i 1 2 2 1 n n n n 5 2 2 1 2 1 1 138. Hystricoceras astraciformis 2 4 2 n n n n i n 2 1 2 2 1 n n n n 5 2 3 1 2 1 1 139. Poleumita granulosa 1 3 2 n n n n n n n 1 2 2 1 n n n n 3 3 1 1 3 1 1 140. Euomphalus walmstedti n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 4 1 1 141. Centrifugus planorbis i 3 2 n n n n n n 1 1 2 2 1 n n n n 3 3 2 1 3 1 1 142. Spinicharybdis wilsoni 3 4 2 2 1 n 2 3 i 2 1 1 n 1 n n n n 5 2 2 1 4 1 1 143. Turbocheilus immaturum n n n n n n n n n n 1 2 2 1 n n n n 3 1 1 1 3 1 ? 1 44. Pseudotectus comes i 1 2 n n n n n n n 1 2 1 1 n n n n 6 2 1 1 2 1 1 145. Straparollus bohemicus n n n n n n n n n n 1 2 2 1 n n n n 3 1 2 1 4 1 I 146. Hormotoma artemesia n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 2 1 147. Hormotoma confusa n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 I 1 1 148. Hormotoma Idubia n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 149. Hormotoma 'Isimulatrix n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 1 50. Ectomaria adelina 1 1 1 n n n n n n 1 1 2 3 1 n n n n 3 3 2 1 1 1 1 151. “ Hormotoma" “ cassina ’’ 1 1 1 n n n n n n 1 1 2 3 1 n n n n 4 3 2 1 1 1 1 NUMBER 88 117 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 1 1 i 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 92. Euomphalus tubus ? 1 4 1 1 1 1 1 2 2 1 7 1 ? 1 1 n 2 2 1 4 2 4 1 2 93. Lytospira subuloides ? ? 4 1 1 1 1 1 1 7 1 7 1 7 1 2 1 1 n 2 3 1 1 1 2 94. Ceratopea unguis ? 2 4 1 1 1 1 1 1 ? 1 7 1 7 1 2 2 1 n 2 4 2 4 1 2 95. Boucotspira aff. B. fimbriata ? 1 3 1 1 1 1 1 1 ? 1 7 1 ? 1 1 n 2 2 1 3 2 4 1 2 96. Lophonema peccatonica ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 2 1 2 3 2 5 1 2 97. Polehemia taneyensis 7 1 3 1 1 1 1 1 1 ? 1 7 1 ? 1 1 n 2 1 2 3 2 5 1 2 98. Walcottoma frydai ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 2 1 2 3 2 5 1 2 99. Helicotoma planulata 7 1 3 1 1 1 1 1 1 7 1 7 1 ? 1 1 n 2 2 1 4 2 3 1 2 100. Helicotoma tennesseensis ? 1 3 1 1 1 1 1 1 ? 1 ? 1 7 1 1 n 2 2 1 4 2 3 1 2 101. Ophiletina sublaxa 9 1 3 1 1 1 1 1 2 2 1 ? 1 7 1 1 2 2 1 1 4 2 3 2 2 102. Ophiletina angularis 7 1 3 1 1 1 1 1 2 2 1 7 1 9 1 1 2 2 1 1 4 2 3 1 2 103. Oriostoma bromidensis ? 1 4 1 1 1 1 1 1 ? 1 ? 1 7 1 1 n 2 1 1 3 2 7 1 2 104. Euomphalopterus lordovicius ? 1 3 1 1 1 1 1 1 ? 1 7 l 7 1 1 n 2 3 2 3 2 5 1 2 105. Euomphalopterus aff. E. ordovicius 7 1 3 1 1 1 2 2 1 7 1 7 1 9 1 1 n 2 3 2 3 2 5 1 2 106. Euomphalopterus cariniferus 7 1 4 1 1 1 1 1 1 7 1 7 1 9 1 1 n 2 2 2 3 2 4 1 2 107. Palaeomphalus Igradatus ? 1 3 1 1 1 1 1 1 7 1 ? 1 7 1 1 n 2 1 1 4 2 3 1 2 108. Trochomphalus Idimidiatus 1 1 4 9 1 1 1 1 1 7 1 7 1 9 1 1 n 7 n 7 3 2 4 1 2 109. Helicotoma blodgetti ? 1 3 l 1 1 1 1 1 ? 1 ? 1 ? 1 1 2 2 2 1 4 2 3 1 2 110. Helicotoma robinsoni ? 1 3 i 1 1 1 1 1 ? 1 7 1 9 1 1 n 2 2 1 4 2 3 1 2 111. Helicotoma ? Girvan sp. ? 1 3 i 1 1 1 1 1 ? 1 9 1 ? 1 1 n 2 2 1 4 2 4 1 2 112. Straporillina cf. S. circe 1 1 4 l 1 1 1 1 1 ? 1 7 1 7 1 1 n 2 2 2 3 2 4 1 2 113. Euomphalopterus alatus ? 1 4 i 1 1 1 1 1 ? 1 ? 1 ? 1 I n 2 2 2 2 2 5 1 2 114. Euomphalopterus frenatus ? 1 3 i 1 1 1 1 1 ? 1 7 1 ? 1 1 n 2 2 2 2 2 5 1 2 115. Euomphalopterus praetextus 9 1 4 l 1 1 1 1 1 7 1 7 1 7 1 1 n 2 2 2 2 2 5 1 2 116. Euomphalopterus subcarinatus ? 1 4 i 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 2 2 2 2 2 5 1 2 117. Euomphalopterus togatus 9 1 4 i 1 1 1 1 1 7 1 ? 1 ? 1 1 n 2 2 2 2 2 5 1 2 118. Euomphalopterus undulans ? 1 3 l 1 1 1 1 1 7 1 7 1 ? 1 1 n 2 3 2 2 2 4 1 2 119. Grantlandispira christei 2 1 3 ? 7 1 1 1 1 ? 1 ? 1 ? 1 1 n 2 2 2 3 2 4 1 2 120. Poleumita discors ? 1 5 3 1 1 1 1 1 ? 1 ? 1 7 1 2 2 1 n 2 3 2 4 1 2 121. Pycnomphalus acutus 2 1 2 2 1 1 1 1 1 9 1 7 1 7 1 1 n 2 2 2 3 2 4 1 2 122. Pycnomphalus obesus 2 1 2 2 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 2 2 2 3 2 4 1 2 123. Discordichilus dalli 9 1 3 1 1 1 1 1 2 4 1 7 1 ? 1 1 n 2 5 2 2 2 5 1 2 124. Discordichilus mollis 7 1 3 2 1 1 2 1 2 5 1 ? 1 7 1 1 n 2 3 2 3 2 4 1 2 125. Discordichilus kolmodini ? 1 2 2 1 1 2 1 2 5 1 ? 1 ? 1 1 n 2 5 2 3 2 4 1 2 126. Poleumita alata ? 1 5 3 1 1 1 1 1 7 1 ? 1 7 1 2 2 1 n 2 3 2 4 1 2 127. Poleumita octavia 7 1 5 3 1 1 1 1 1 7 1 ? 1 7 1 1 n 1 n 2 4 2 5 1 2 128. Poleumita rugosa ? 1 5 3 1 1 1 1 1 7 1 ? 1 ? 1 2 2 1 n 2 4 2 4 1 2 129. Pseudophorus profundus 7 1 2 2 1 1 2 1 2 3 1 ? 2 2 1 1 n 2 5 2 2 2 4 1 2 130. Pseudophorus stuxbergi 7 1 2 2 1 1 2 1 2 3 1 ? 2 2 1 1 n 2 5 2 273 2 4 1 2 131. Siluriphorus gotlandicus ? 1 2 2 1 1 2 1 2 2 1 7 2 3 1 1 n 2 5 2 1 2 4 1 2 132. Siluriphorus undulans ? 1 2 2 1 1 2 1 2 2 1 ? 2 3 1 1 n 2 5 2 1 2 4 1 2 133. Streptotrochus incisus ? 1 3 1 1 2 1 1 2 3 1 ? 1 ? 1 1 n 2 3 2 2 3 5 1 2 134. Streptotrochus aff. S. incisus ? 1 2 1 1 1 1 1 2 2 1 ? 2 1 1 1 n 2 2 2 2 2 5 1 2 135. Streptotrochus lamellosa ? 1 2 1 1 1 1 1 2 2 1 ? 1 ? 1 1 n 2 3 1 2 3 5 1 2 136. Streptotrochus lundgreni ? 1 3 1 1 2 1 1 2 4 1 7 1 ? 1 1 n 2 2 2 2 3 5 1 2 137. Streptotrochus! visbeyensis 7 1 2 1 1 1 1 1 2 3 1 ? 1 7 1 1 n 2 3 2 3 2 5 1 2 138. Hystricoceras astraciformis ? 1 3 1 1 2 1 1 2 3 1 ? 1 ? 1 1 n 2 5 2 1 3 6 1 2 139. Poleumita granulosa ? 1 5 3 1 1 1 1 1 7 1 ? 1 ? 1 2 i 1 n 2 3 2 4 1 2 140. Euomphalus walmstedti ? 1 5 3 1 1 1 1 1 7 1 ? 1 ? 1 2 i 1 n 1 3 2 4 1 2 141. Centrifugus planorbis ? 1 5 3 1 1 1 1 1 ? 1 ? 1 7 1 2 2 1 n 2 3 2 3 1 2 142. Spinicharybdis wilsoni ? 1 3 1 1 1 1 1 1 7 1 ? 1 ? 1 1 n 2 i 2 2 2 5 1 2 143. Turbocheilus immaturum ? 1 2 7 ? 1 2 1 1 7 1 7 1 ? 1 1 n 2 2 2 3 2 4 1 2 144. Pseudotectus comes ? 1 2 2 1 1 2 1 2 5 1 ? 1 ? 1 1 n 2 3 2 3 1 4 1 2 145. Straparollus bohemicus ? 1 5 2 1 1 1 1 1 7 1 ? 1 ? 1 2 1 1 n 1 3 2 4 1 2 146. Hormotoma artemesia ? 1 3 1 1 2 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 147. Hormotoma confusa ? 1 3 1 1 2 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 148. Hormotoma Idubia ? 1 3 1 1 2 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 149. Hormotoma Isimulatrix ? 1 3 1 1 2 1 1 1 ? 1 ? 1 9 1 1 n 1 n 1 1 2 6 1 2 150. Ectomaria adelina ? 1 3 1 1 2 1 1 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 151. “ Hormotoma ” “ cassina ” 7 1 3 1 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 1 2 6 1 2 118 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 92. Euomphalus tubus 5 1 2 2 4 1 1 1 4 1 1 1 1 1 1 1 ? n 93. Lytospira subuloides 2 1 2 1 n n n 1 n n n n 1 n 1 ? ? n 94. Ceratopea unguis 4 2 2 1 n n n 1 n n n n 1 1 1 3 ? n 95. Boucotspira aff. B.fimbriata 5 1 ? 1 n n n 1 n n n n 1 1 1 2 ? n 96. Lophonema peccatonica 5 2 ? 1 n n n 1 n n n n 1 1 1 2 ? n 97. Pole hernia taneyens is 5 2 ? 2 i 3 2 1 n n n n 1 1 1 2 ? n 98. Walcottoma frydai 5 1 ? 2 i 3 2 2 n n n n 1 1 1 ? ? n 99. Helicotoma planulata 4 2 1 3 3 3 2 1 n n n n 1 1 1 2 ? n 100. Helicotoma tennesseensis 4 1 1 1 n n n 1 n n n n 1 1 1 2 ? n 101. Ophiletina sublaxa 4 1 ? 1 n n n 1 n n n n 1 1 1 1 ? n 102. Ophiletina angularis 4 1 7 1 n n n 1 n n n n 1 1 1 1 ? n 103. Oriostoma bromidensis ? 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 104. Euomphalopterus lordovicius 5 1 ? 1 n n n I n n n n 1 1 1 2 ? n 105. Euomphalopterus aff. E. ordovicius 5 1 ? 1 n n n 1 n n n n 1 1 1 2 ? n 106. Euomphalopterus cariniferus 5 1 2 1 n n n 1 n n n n 1 1 1 2 ? n 107. Palaeomphalus Igradatus 4 1 ? 1 n n n 1 n n n n 1 1 1 2 ? n 108. Trochomphalus Idimidiatus 4 1 ? 1 n n n 1 n n n n 1 1 1 ? ? n 109. Helicotoma blodgetti 4 2 1 3 i 1 2 1 n n n n 1 1 1 2 ? n 110. Helicotoma robinsoni 4 2 ? 3 3 3 2 1 n n n n 1 1 1 2 ? n 111. Helicotoma ? Girvan sp. 5 1 ? 1 n n n 1 n n n n 1 1 1 2 ? n 112. Straporillina cf. S. circe 4 1 ? 1 n n n 1 n n n n 1 1 1 2 ? n 113. Euomphalopterus alatus 5 1 2 1 n n n 1 n n n n ? n 1 3 ? n 114. Euomphalopterus frenatus 6 1 2 1 n n n 1 n n n n ? n 1 2 ? n 115. Euomphalopterus praetextus 5 2 2 1 n n n 1 n n n n ? n 1 2 ? n 116. Euomphalopterus subcarinatus 5 1 2 1 n n n 1 n n n n 1 i 1 2 ? n 117. Euomphalopterus togatus 5 1 2 1 n n n 1 n n n n ? i 1 2 ? n 118. Euomphalopterus undulans 5 1 2 1 n n n 1 n n n n ? n 1 2 ? n 119. Grantlandispira christei 5 1 ? 2 3 i 1 ? n n n n 7 i 1 3 ? n 120. Poleumita discors 4 3 2 2 2 3 1 2 i 2 i i 1 i 1 3 ? n 121. Pycnomphalus acutus 5 1 2 1 n n n 1 n n n n ? n 1 2 ? n 122. Pycnomphalus obesus 5 1 2 1 n n n 1 n n n n 7 n 1 2 ? n 123. Discordichilus dalli 6 1 ? 1 u n n 1 n n n n 1 n 1 1 ? n 124. Discordichilus mollis 6 1 ? 1 n n n 1 n n n n 1 n 1 1 7 n 125. Discordichilus kolmodini 6 1 ? 1 n n n 1 n n n n 1 n 1 2 ? n 126. Poleumita alata 5 3 2 2 2 3 i 2 1 2 1 1 1 i 1 2 7 n 127. Poleumita octavia 6 1 2 2 3 3 i 2 3 2 1 1 1 i 1 2 ? n 128. Poleumita rugosa 4 3 2 2 3 3 i 2 2 1 1 1 1 i 1 3 ? n 129. Pseudophorus profundus 5 1 2 1 n n n 1 n n n n ? n 1 2 ? n 130. Pseudophorus stuxbergi 5 1 ? 1 n n n 1 n n n n 7 n 1 2 ? n 131. Siluriphorus gollandicus 6 1 2 1 11 n n 1 n n n n 7 n 1 2 ? n 132. Siluriphorus undulans 6 1 2 1 n n n 1 n n n n ? n 1 2 ? n 133. Streptotrochus incisus 6 2 ? 1 n n n 1 n n n n 7 n 1 1 ? n 134. Streptotrochus aff. S. incisus 6 1 ? 1 n n n 1 n n n n ? n 1 1 ? n 135. Streptotrochus lamellosa 6 1 ? 1 n n n 1 n n n n ? n 1 2 ? n 136. Streptotrochus lundgreni 6 2 2 1 n n n 1 n n n n ? n 1 2 ? n 137. Streptotmchusl visbeyensis 6 1 ? 1 n n n 1 n n n n ? n 1 1 ? n 138. Hystricoceras astraciformis 6 2 2 1 n n n 1 n n n n ? n 1 2 ? n 139. Poleumita granulosa 4 3 2 2 2 i i 2 1 2 i i 1 1 1 3 ? n 140. Euomphalus walmstedti 4 1 2 1 n n n 1 n n n n 7 1 1 3 ? n 141. Centrifugus planorbis 3 1 2 3 2 3 1 2 2 2 i i 1 1 1 3 ? n 142. Spinicharybdis wilsoni 6 1 ? 1 n n n 1 n n n n 1 n 1 2 ? n 143. Turbocheilus immaturum 5 1 ? 1 n n n 1 n n n n 7 i 1 2 7 n 144. Pseudotectus comes 6 1 ? 1 n n n 1 n n n n 1 i 1 2 ? n 145. Straparollus bohemicus 5 1 2 1 n n n 1 n n n n ? n 1 2 ? n 146. Hormotoma artemesia 8 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 147. Hormotoma confusa 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 148. Hormotoma Idubia 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 149. Hormotoma Isimulatrix 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 150. Ectomaria adelina 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 151. “ Hormotoma ” “ cassina ” 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n NUMBER 88 119 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 152. Fusispira Smithville Fm. sp. 1 ? ? 2 ? ? 7 7 ? 3 1 1 1 n 1 1 1 1 1 n n n n n n 153. Hormotoma augustina 2 3 4 7 3 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 1 2 1 n 154. Hormotoma zelleri 2 3 3 6 3 3 4 1 3 4 1 1 1 n 2 1 1 I 2 5 2 1 2 1 n 155. Lophospira perangulata 2 2 4 4 2 3 3 2 3 3 1 I 1 n 2 1 ? 1 2 3 2 2 3 1 n 156. Subulitid El Paso Fm. sp. 1 ? ? 2 7 ? 7 7 ? 3 1 1 1 n 2 1 1 1 1 n n n n n n 157. Pagodospira cicelia 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 ? 1 2 3 2 2 3 1 n 158. Plethospira cannonensis 2 2 3 3 2 2 2 2 3 3 1 1 1 n 2 1 1 1 2 4 2 1 2 1 n 159. Plethospira cassina 2 2 3 3 2 2 2 2 3 3 1 1 1 n 2 1 1 1 2 4 2 1 2 1 n 160. Seelya ventricosa 2 2 3 3 2 2 2 2 3 3 1 1 1 n 2 1 1 1 2 4 2 1 2 1 n 161. Lophospira grandis 2 2 3 3 2 2 2 2 3 3 1 1 1 n 2 1 1 1 2 3 2 1 2 1 n 162. Straparollina pelagica 2 2 1 1 2 1 1 2 3 3 I 1 I n 2 1 1 1 2 2 1 n n n n 163. Plethospira ? turgida 2 2 4 4 2 1 1 2 3 3 1 1 1 n 4 1 1 1 2 3 2 i 2 i n 164. Turritoma acrea 2 3 4 5 1 3 3 2 3 3 1 1 1 n 3 1 1 1 2 5&6 2 2 3 i n 165. Turritoma Cotter Fm. ornate sp. 2 2 4 4 2 3 3 2 3 3 1 1 1 n 3 1 1 1 2 5 2 2 3 i n 166. Turritoma cf. T. acrea 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 I 1 2 5 2 1 2 i n 167. Hormotoma Setul Fm. sp. 2 3 2&3 7 3 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 1 2 i n 168. Turritoma lanna 2 3 3 5 3 3 4 2 3 3 1 1 1 n 2 1 1 1 2 4 2 1 2 i n 169. Murchisortia callahanensis 2 2 3 3 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 170. Ectomaria prisca 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 171. Hormotoma gracilis 2 3 3 7 3 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 1 2 i n 172. Daidia cerithioides 1 ? ? ? ? ? 7 ? ? ? ? 1 1 n 4 1 1 1 2 1 1 n n n n 173. Ectomaria pagoda 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 1 n 174. Haplospira 1 nereis 1 ? ? ? 7 7 ? ? ? ? ? 1 1 n 3 1 1 1 2 1 1 n n n n 175. Hormotoma bellicincta 2 3 4 6 3 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 i 2 i n 176. Hormotoma salteri 2 3 3 7 3 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 i 2 i n 177. Hormotoma trentonensis 2 2 4 4 3 3 4 2 3 3 1 1 1 n 2 1 1 1 2 4 2 i 2 i n 178. Loxonema murrayana 2 2 4 4 3 3 3 2 3 3 1 1 1 n 2 1 1 1 1 n 11 n n n n 179. Omospira alexandra 2 2 2 2 3 3 4 2 3 3 1 1 1 n 2 1 1 1 2 4 2 i 2 i n 180. Omospira laticincta 2 2 2 2 3 3 3 2 1 1 1 1 1 n 2 1 1 1 2 4 2 i 2 i n 181. Straparollina circe 2 2 1 1 2 2 2 2 3 3 1 1 1 n 3 1 1 1 1 n n n n n n 182. Straparollina erigione 2 2 1 1 2 2 2 2 3 3 1 1 1 n 3 1 1 1 1 n n n n n n 183. Girvania excavata 2 2 2 2 2 3 3 2 1 1 1 1 1 n 2 1 1 1 1 n n n n n n 184. Murchisonia Pt. Clarence Fm. sp. 2 2 1 1 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 185. Rhabdostropha primitiva 2 2 2 2 3 3 3 2 3 3 1 1 1 n 2 1 1 1 1 n n n n n n 186. Spiroecus girvanensis 2 2 2 2 2 3 3 2 1 1 1 1 1 n 2 1 1 1 2 1 1 n n n n 187. Daidia aff. D. cerithioides 1 ? 7 ? ? ? ? 7 7 7 ? 1 1 n 4 1 1 1 2 1 1 n n n n 188. Ectomaria cf. E. pagoda 2 2 2 2 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 189. Ectomaria cf. E. prisca 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 190. Ectomaria laticarinata 2 2 3 3 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 191. Ectomaria nieszkowskii 2 2 4 4 2 3 3 2 3 3 1 1 1 n 2 1 1 1 2 6 2 2 5 i n 192. Hormotoma insignis 2 3 3 5 1 3 4 1 3 4 1 1 1 n 3 1 1 1 2 4 2 1 2 i n 193. Holopella regularis 2 2 1 1 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 194. Hormotoma centervillensis 2 3 4 7 3 3 4 1 3 4 1 1 1 n 4 1 1 1 2 4 2 i 3 i n 195. Hormotoma cingulata 2 3 3 7 1 3 4 1 3 4 1 1 1 n 2 1 1 1 2 4 2 i 3 i n 196. Kjerulfonema cancellata 2 2 0 0 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 197. Kjerulfonema quinquecincta 2 2 0 0 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 198. Cyrtostropha coralli 2 3 4 7 3 3 4 1 3 4 1 1 1 n 4 1 1 1 2 4 2 i 3 i n 199. Goniostropha cava 2 3 2 5 1 3 4 2 3 3 1 1 1 n 3 1 1 1 2 4 2 2 4 i n 200. Hormotoma subplicata 2 3 4 4&5 2 3 4 2 3 3 1 1 1 n 2 1 1 1 2 5 2 2 2 i n 201. Hormotoma monoliniformis 2 3 3 5 3 3 4 1 3 4 1 1 1 n 3 1 1 1 2 3 2 1 3 i n 202. Hormotoma attenuata 2 3 3 5 3 3 4 1 3 4 1 1 1 n 3 1 1 1 2 2 1 n n n n 203. Loxonema ? attenuata 2 3 3 5 3 3 4 1 3 4 1 1 1 n 3 1 1 1 2 1 2 2 i 1 n 204. Macrochilus fenestratus 2 2 0 0 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 205. Rhabdostropha grindrodii 2 2 1 1 3 3 3 2 1 1 1 1 1 n 3 1 1 1 1 n n n n n n 206. Loxonema crossmanni 2 2 4 4 3 3 3 2 3 3 1 1 1 n 3 1 1 1 1 n n n n n n 207. Loxonema sinuosa 2 2 3 3 3 3 3 2 3 3 1 1 1 n 3 1 1 1 1 n n n n n n 208. Auriptygma fortior 1 ? 7 ? ? 7 7 ? ? ? 7 1 1 n 3 1 1 1 1 n n n n n n 209. Catazone allevata 2 3 2 7 1 3 4 1 3 4 1 1 1 n 2 1 1 1 2 3 2 i 3 i n 210. Catazone argolis 2 3 2 7 1 2 4 1 3 4 1 1 1 n 3 1 1 1 2 3 2 2 3 i n 211. Catazone cunea 2 3 2 7 1 3 4 1 3 4 1 1 1 n 2 1 1 1 2 3 2 1 3 i n 120 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 152. Fusispira Smithville Fm. sp. n n n n n n n n 1 n n 1 n n n n n n n ? n 1 E 1 n 153. Hormotoma augustina 1 i n n n n n 2 1 n n 1 1 2 1 1 1 2 1 2 1 1 9 1 n 154. Hormotoma zelleri 1 i n n n n n 2 1 n n 1 1 2 2 1 1 2 1 1 n 1 8 1 n 155. Lophospira perangulata 1 2 2 3 n 1 1 1 1 n n 1 1 2 1 1 1 2 1 2 1 1 A 1 n 156. Subulitid El Paso Fm. sp. n n n n n n n n 1 n n 1 n n n n n n n ? n 1 E 1 n 157. Pagodospira cicelia 1 2 2 3 n i i i 1 n n 1 1 2 1 i 1 2 1 2 1 1 9 1 n 158. Plethospira cannonensis 1 1 n n n n n i 1 n n 1 1 2 1 i 1 2 1 1 n 1 9 1 n 159. Plethospira cassina 1 1 n n n n n i 1 n n 1 1 2 1 i 1 2 1 1 n 1 8 1 n 160. Seelya ventricosa 1 1 n n n n n i 1 n n 1 1 2 1 i 1 2 1 ? n 1 9 1 n 161. Lophospira grandis 1 1 1 2 n 1 i 3 1 n n 1 1 2 2 2 1 2 1 2 i 1 A 1 n 162. Straparollina pelagica n 2 1 2 n 1 n 3 1 n n 1 4 2 2 n 1 2 1 2 i 1 C 1 n 163. Plethospira? turgida i 2 1 2 n 1 i 3 1 n n 1 1 2 2 2 1 2 1 1 n 1 7 1 n 164. Turritoma acrea i 1 n n n n n 1 1 n n 1 1 2 1 1 1 3 1 1 n 1 7 1 n 165. Turritoma Cotter Fm. ornate sp. i 1 n n n n n 1 1 n n 1 1 2 1 1 1 2&3 2 1 n 1 7 1 n 166. Turritoma cf. T. acrea i 2 i 2 n i i 2 1 n n 1 1 2 2 2 1 2 1 1 n 1 7 1 n 167. Hormotoma Setul Fm. sp. i 1 n n n n n 2 1 n n 1 n n 1 1 1 2 1 2 i 1 9 1 n 168. Turritoma lanna i 1 n n n n n 1 1 n n 1 i i 1 1 1 2 1 1 n 1 8 1 n 169. Murchisonia callahartensis i 1 n n n n n 1 1 n n 1 i 2 1 1 1 2 1 1 n 1 9 1 n 170. Ectomaria prisca i 1 n n n n n 1 1 n n 1 i 2 1 1 1 2 1 1 n 1 9 1 n 171. Hormotoma gracilis i 1 n n n n n 1 1 n n 1 i 1 1 1 1 2 1 1 n 1 9 1 n 172. Daidia cerithioides n 2 i 2 i i i 1 1 n n 1 i 2 1 n 1 2 1 2 1 1 A 1 n 173. Ectomaria pagoda 1 1 n n n n n 1 1 n n 1 i 2 1 i 1 2 1 1 n 1 9 1 n 174. Haplospira ?nereis n 2 1 2 i i i 1 1 n n 1 n 2 1 n 1 2 1 2 i 1 A 1 n 175. Hormotoma bellicincta 1 1 n n n n n 1 1 n n 1 i 2 1 i 1 2 1 1 n 1 A 1 n 176. Hormotoma salteri 1 2 i i 2 i i 1 1 n n 1 i 1 1 i 1 2 1 1 n 1 9 1 n 177. Hormotoma trentonensis 1 1 n n n n n 1 1 n n 1 i 2 1 i 1 2 1 1 n 1 B 1 n 178. Loxonema murrayana n n n n n n n n 1 n n n i 2 n n n n 1 1 n 1 B 1 n 179. Omospira alexandra i i n n n n n i 1 n n 1 i 2 1 1 1 2 1 1 n 1 A 1 n 180. Omospira laticincta i i n n n n n i 1 n n 1 i 2 1 1 1 2 1 1 n 1 B 1 n 181. Straparollina circe n n n n n n n n 1 n n n i 2 1 n 1 2 1 1 n ? 9 1 n 182. Straparollina erigione n n n n n n n n 1 n n n i 2 1 n 1 2 1 1 n ? A 1 n 183. Girvania excavata n n n n n n n n 1 n n n i 2 1 1 1 2 1 1 n 1 B 1 n 184. Murchisonia Pt. Clarence Fm. sp. n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 B 1 n 185. Rhabdostropha primitiva n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 B 1 n 186. Spiroecus girvanensis 1 2 i 2 i i n 2 1 n n i i 2 1 1 1 2 i 1 n 1 A 1 n 187. Daidia aff. D. cerithioides n 2 2 2 i i i 1 1 n n i i 2 1 n 1 2 i 2 1 1 A 1 n 188. Ectomaria cf. E. pagoda i 1 n n n n n 1 1 n n i i 2 1 i 1 2 i 1 n 1 7 1 n 189. Ectomaria cf. E. prisca i 1 n n n n n 1 1 n n i i 2 1 i 1 2 i 1 n 1 9 1 n 190. Ectomaria laticarinata i 1 n n n n n 1 1 n n i i 2 1 i 1 2 i 1 n 1 9 1 n 191. Ectomaria nieszkowskii i 1 n n n n n 1 1 n n i i 2 1 i 1 2 i 1 n 1 9 1 n 192. Hormotoma insignis i 1 n n n n n 1 1 n n i i 1 1 i 1 2 i 1 n 1 6 1 n 193. Holopella regularis n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 9 1 n 194. Hormotoma centervillensis 1 1 n n n n n i 1 n n i i 1 i i i 2 i 1 n 1 9 1 n 195. Hormotoma cingulata 1 1 n n n n n i 2 2 2 i i 1 i i i 2 i 1 n 1 7 1 n 196. Kjerulfonema cancellata n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 9 1 n 197. Kjerulfonema quinquecincta n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 B 1 n 198. Cyrtostropha coralli i i n n n n n 1 1 n n i i 1 i 1 i 2 i 1 n 1 9 1 n 199. Goniostropha cava i i n n n n n 1 1 n n i i 1 i 1 i 2 i 1 n 1 8 1 n 200. Hormotoma subplicata i 2 1 1 1 1 1 1 1 n n i i 1 i 1 i 2 i 1 n 1 9 1 n 201. Hormotoma monoliniformis i 1 n n n n n 1 1 n n i i 1 i 1 i 2 i 1 n 1 9 1 n 202. Hormotoma attenuata i 2 1 2 i i i 1 1 n n i i 1 i 1 i 2 i 1 n 1 A 1 n 203. Loxonema ? attenuata n 1 n n n n n n 2 2 2 i i 1 i 1 i 2 i 1 n 1 B 1 n 204. Macrochilus fenestratus n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 E 1 n 205. Rhabdostropha grindrodii n n n n n n n n 1 n n n i 2 i 1 1 2 1 1 n 1 B 1 n 206. Loxonema crossmanni n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 A 1 n 207. Loxonema sinuosa n n n n n n n n 1 n n n i 2 n n n n n 1 n 1 A 1 n 208. Auriptygma forlior n n n n n n n n 1 n n n n n n n n n n 1 n 1 C 1 n 209. Catazone allevata n 1 n n n n n 1 2 2 2 i 1 1 i 1 1 2 1 1 n 1 2 1 n 210. Catazone argolis n 1 n n n n n 1 2 2 2 i 1 2 i 1 1 2 1 1 n 1 4 1 n 211. Catazone cunea n 1 n n n n n 1 2 2 2 i 1 2 i 1 1 2 1 1 n 1 4 1 n NUMBER 88 121 Appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 6 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 2 3 4 5 152. Fusispira Smithville Fm. sp. 3 ? ? 3 ? 6 ? ? 5 ? n n 1 ? n n n n 2 1 1 2 2 1 153. Hormotoma augustina 3 2 2 1 5 5 1 5 2&3 2 1 3 1 1 n n n n 1 n 2 2 1 1 154. Hormotoma zelleri 3 2 2 2 5 5 1 6 5 2 2 3 1 1 n n n n 2 i 1 2 2 1 155. Lophospira perangulata 2 3 3 3 3 5 1 4 2 1 n n 1 2 i i i i 1 n 3 1 n 2 156. Subulitid El Paso Fm. sp. 3 ? ? 3 7 6 7 7 5 7 n n 1 ? n n n n 2 1 1 2 2 1 157. Pagodospira cicelia 2 3 3 1 4 5 1 4 2 1 n n 1 2 1 1 i i 1 n 3 1 n 2 158. Plethospira cannonensis 2 2 2 2 5 5 1 6 4 2 i 3 1 I n n n n 1 n 2 2 1 1 159. Plethospira cassina 2 2 2 2 5 6 1 5 4 1 n n 1 1 n n n n 1 n 2 1 n 1 160. Seelya ventricosa 2 2 2 1 5 5 1 5 4 1 n n 1 1 n n n n 1 n 2 1 n 1 161. Lophospira grandis 3 3 2 2 2&3 5 1 6 3 1 n n 1 I n n n n 1 n 2 1 n 2 162. Straparollina pelagica 3 3 2 3 0 6 1 6 4 1 n n 1 1 n n n n 1 n 2 1 n 2 163. Plethospirdl turgida 3 2 2 3 6 5 3 4 5 2 3 3 1 1 n n n n 1 n 2 2 3 1 164. Turritoma acre a 2 3 2 1 7 2 2 5 5 1 n n 1 1 n n n n 1 n 3 1 n 1 165. Turritoma Cotter Fm. ornate sp. 2 2 2 1 5 4&5 1 5 4 2 i 3 1 1 n n n n 1 n 2 1 n 1 166. Turritoma cf. T. acrea 3 2 2 2 4 4 2 5 5 2 i 3 1 1 n n n n 1 n 2 2 i 1 167. Hormotoma Setul Fm. sp. 3 2 2 1 5 4 1 5 2&3 2 i 3 1 1 n n n n 1 n 2 2 i 1 168. Turritoma lanna 2 2 2 2 5 4&5 1 4 3 1 n n 1 1 n n n n 1 n 2 1 n 1 169. Murchisonia callahanensis 2 4 1 3 4 5 1 5 3 1 n n 1 2 2 4 2 1 1 n 4 1 n 2 170. Ectomaria prisca 2 3 2 3 3 5 1 4 3 1 n n 1 1 n n n n 1 n 3 1 n 2 171. Hormotoma gracilis 2 2 2 3 3 4&5 1 4&f 3 1 n n 1 1 n n n n 1 n 2 1 n 1 172. Daidia cerithioides 3 4 2 3 1 7 1 8 2 1 n n 1 2 3 3 1 1 2 i 2 1 n 2 173. Ectomaria pagoda 2 3 2 3 3 5 1 4 3 1 n n 1 2 1 2 1 1 1 n 3 1 n 2 174. Haplospira 1 nereis 3 3 7 3 1 6 1 6 3 1 n n 1 1 n n n n 1 n 2 1 n 1 175. Hormotoma bellicincta 2 2 2 2 3 4 1 5 2&3 1 n n 1 1 n n n n 1 n 2 1 n 1 176. Hormotoma salteri 3 3 2 3 3&4 5 1 6 2&3 1 n n 1 1 n n n n 2 i 2 1 n 1 177. Hormotoma trentonensis 2 2 2 2 3 3 1 4 2&3 1 n n 1 1 n n n n 1 n 2 1 n 1 178. Loxonema murrayana 2 2 2 3 2 6 1 6 4 1 n n 1 1 n n n n 2 i 2 1 n 1 179. Omospira alexandra 2 2 2 3 1 5 1 7 1&2 1 n n 1 1 n n n n 2 i 2 1 n 1 180. Omospira laticincta 2 2 2 3 1 5 1 6 3 1 n n 1 1 n n n n 2 i 2 1 n 1 181. Straparollina circe 3 2 2 3 0 6 1 6 4 1 n n 1 1 n n n n 1 n 2 1 n 1 182. Straparollina erigione 3 2 2 3 0 6 1 6 4 1 n n 1 1 n n n n 1 n 2 1 n 1 183. Girvania excavata 2 2 2 3 2 4 1 7 3 1 n n 1 1 n n n n 2 i 2 1 n 1 184. Murchisonia Pt. Clarence Fm. sp. 2 2 2 3 2 5 1 8 4 1 n n 1 1 n n n n 2 i 2 1 n 1 185. Rhabdostropha primitiva 2 2 2 3 3 4 2 4 4 1 n n 1 1 n n n n 2 i 2 1 n 1 186. Spiroecus girvanensis 3 4 1 3 2 3 1 7 5 1 n n 1 2 2 3 i 2 2 i 3 1 n 1 187. Daidia aff. D. cerithioides 3 4 2 3 1 7 1 8 2 1 n n 1 2 3 3 i 1 2 i 2 1 n 2 188. Ectomaria cf. E. pagoda 2 3 2 2 4 3 1 4 3 1 n n 1 2 3 4 i 1 1 n 3 1 n 2 189. Ectomaria cf. E. prisca 2 3 2 2 4 4 2 4 4 1 n n 1 1 n n n n 1 n 3 1 n 2 190. Ectomaria laticarinata 2 4 1 3 4 5 1 5 2 1 n n 1 1 n n n n 1 n 4 1 n 2 191. Ectomaria nieszkowskii 2 3 2 2 4 4 1 4 2 1 n n 1 1 n n n n 1 n 3 1 n 2 192. Hormotoma insignis 3 2 2 1 5 4 2 5 5 1 n n 1 1 n n n n 1 n 1 1 n 1 193. Holopella regularis 2 2 2 3 2 6 1 7 4 1 n n 1 1 n n n n 2 i 2 1 n 1 194. Hormotoma centervillensis 2 2 2 3 3 4 1 5 4 1 n n 1 2 2 3 1 1 1 n 2 1 n 1 195. Hormotoma cingulata 3 2 2 1 5 3 2 6 5 1 n n 1 1 n n n n 1 n 1 1 n 1 196. Kjerulfonema cancellata 2 2 2 3 2 6 1 7 4 I n n 1 1 n n n n 2 i 2 1 n 1 197. Kjerulfonema quinquecincta 2 2 2 3 2 6 1 7 4 1 n n 1 1 n n n n 2 2 1 2 1 n 1 198. Cyrtostropha coralli 2 2 2 3 3 4 1 5 4 1 n n 1 2 3 4 2 n 1 n 1 2 1 n 2 199. Goniostropha cava 2 3 2 3 2 4 1 5 4 1 n n 1 1 n n n n 2 2 1 2 1 n 1 200. Hormotoma subplicata 3 3 2 3 3 4 2 5 5 1 n n 1 1 n n n n 2 2 1 3 1 n 1 201. Hormotoma monoliniformis 2 2 2 2 4 4 2 4 4 1 n n 1 1 n n n n 1 n 1 2 1 n 1 202. Hormotoma attenuata 3 4 2 3 3 6 1 6 1&2 1 n n 1 1 n n n n 2 2 1 3 1 n 1 203. Loxonema ? attenuata 3 4 2 3 1 6 1 8 3 1 n n 1 1 n n n n 2 1 1 2 1 n 2 204. Macrochilus fenestratus 2 2 2 3 0 6 1 7 I 1 n n 1 1 n n n n 2 1 1 2 1 n 1 205. Rhabdostropha grindrodii 2 2 2 3 1 4 1 8 4 1 n n 1 1 n n n n 2 1 1 2 1 n 1 206. Loxonema crossmanni 2 2 2 3 2 6 1 7 4 1 n n 1 1 n n n n 2 1 1 2 1 n 1 207. Loxonema sinuosa 2 2 2 3 2 6 1 7 4 1 n n 1 1 n n n n 2 1 1 2 1 n 1 208. Auriptygma fortior 2 2 7 3 0 6 1 8 1 1 n n 1 1 n n n n 2 1 2 2 1 n 1 209. Catazone allevata 3 3 2 1 6 2 2 5 5 1 n n 1 1 n n n n 2 1 1 3 1 n 2 210. Catazone argolis 1 2 2 1 7 2 2 4 4 1 n n 1 1 n n n n 2 1 1 3 1 n 2 211. Catazone cunea 3 3 2 1 6 2 2 5 5 1 n n 1 1 n n n n 2 1 1 3 1 n 2 122 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 7 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 1 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 152. Fusispira Smithville Fm. sp. n n n n n n n n n n 1 1 n 1 n n n n 0 3 2 1 1 3 1 153. Hormotoma augustina n n n n n n n n n n 1 2 2 1 n n n n 2 3 2 1 1 2 1 154. Hormotoma zelleri n n n n n n n n n n 1 2 2 1 n n n n 2 3 2 1 1 2 1 155. Lophospira perangulata i 1 i n n n n n n i 1 2 3 1 n n n n 3 3 2 1 1 1 1 156. Subulitid El Paso Fm. sp. n n n n n n n n n n 1 2 2 1 n n n n 0 3 2 1 1 3 1 157. Pagodospira cicelia i i 1 n n n n n n 1 1 2 3 1 n n n n 3 3 2 1 1 1 1 158. Plethospira cannonensis n n n n n n n n n n 1 2 2 1 n n n n 2 2 2 1 1 2 1 159. Plethospira cassina n n n n n n n n n n 1 2 2 1 n n n n 2 2 2 1 1 2 1 160. Seelya ventricosa n n n n n n n n n n 1 2 2 1 n n n n 2 2 2 1 1 2 1 161. Lophospira grandis i i i n n n n n n 1 1 2 2 1 n n n n 4 2 2 1 1 1 1 162. Straparoll ina pelagica i i i n n n n n n 1 1 2 2 2 3 2 1 1 4 2 2 1 1 1 1 163. Plethospira ? turgida n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 164. Turritoma acrea n n n n it n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 165. Turritoma Cotter Fm. ornate sp. n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 I 1 1 166. Turritoma cf. T. acrea n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 167. Hormotoma Setul Fm. sp. n n n T1 n n n n n n 1 2 2 1 n n n n 2 3 2 1 1 2 1 168. Turritoma ?anna n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 169. Murchisonia callahanensis 2 i 1 n n n n n n i 1 2 3 1 n n n n 3 3 2 I 1 1 1 170. Ectomaria prisca 2 2 1 n n n n n n i 1 2 3 1 n n n n 3 3 2 1 1 1 1 171. Hormotoma gracilis n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 172. Daidia cerithioides 2 i i n n n n n n i 1 2 4 ? ? 7 ? ? 3 1 1 1 1 1 1 173. Ectomaria pagoda 2 2 i n n n n n n i 1 2 3 1 n n n n 3 3 2 1 1 1 1 174. Haplospira ?nereis n n n n n n n n n n 1 2 4 ? ? 7 ? 7 3 1 1 1 1 1 1 175. Hormotoma bellicincta n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 2 1 176. Hormotoma salleri n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 177. Hormotoma trentonensis n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 178. Loxonema murrayana n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 179. Omospira alexandra n n n n n n n n n n 1 2 3 1 n n n n 3 2 2 1 1 1 1 180. Omospira laticincta n n n n n n n n n n 1 1 II 1 n n n n 3 2 2 1 1 1 1 181. Straparollina circe n n n n n n n n n n 1 2 4 2 3 1 i 1 4 1 1 1 2 1 1 182. Straparollina erigiorte n n n n n n n n n n 1 2 4 2 3 2 i 1 4 1 1 1 1 1 1 183. Girvan ia excavata n n n n n n n n n n 1 1 n 1 n n n n 2 2 2 I 1 1 1 184. Murchisonia Pt. Clarence Fm. sp. n n n n n n n n n n 1 1 n 1 n n n n 2 2 1 1 1 1 1 185. Rhabdostropha primitiva n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 1 1 1 186. Spiroecus girvanensis n n n n n n n n n 1 1 1 n 1 n n n n 2 2 2 1 1 1 1 187. Daidia aff. D. cerithioides 2 i 1 n n n n n n 1 1 2 4 ? ? ? ? ? 3 1 1 1 1 1 1 188. Ectomaria cf. E. pagoda 2 2 1 n n n n n n 1 1 2 3 1 n n n n 3 1 2 1 1 1 1 189. Ectomaria cf. E. prisca 2 3 1 n n n n n n 1 1 2 3 2 3 2 1 i 3 3 2 1 1 1 1 190. Ectomaria laticarinata 2 1 1 n n n n n n 1 1 2 3 1 n n n n 3 3 2 1 1 1 1 191. Ectomaria nieszkowskii 2 2 1 n n n n n n 1 1 2 3 1 n n n n 3 3 2 1 1 1 1 192. Hormotoma ins ignis 2 1 n n n n n n n n 1 2 2 1 n n n n 3 3 3 1 2 1 1 193. Holopella regularis n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 2 1 1 194. Hormotoma centervillensis n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 195. Hormotoma cingulata n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 196. Kjerulfonema cancellata n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 197. Kjerulfonema quinquecincta n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 198. Cyrtostropha coralli 2 2 n n n n n n n 1 l 2 2 1 n n n n 3 3 2 1 1 1 1 199. Goniostropha cava n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 200. Hormotoma subplicata n n n n n n n n n n 1 2 2 1 n n n n 2 2 2 1 1 1 1 201. Hormotoma monoliniformis n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 202. Hormotoma attenuata n n n n n n n n n n 1 2 2 1 n n n n 2 3 3 1 1 1 1 203. Loxonemaj attenuata 2 2 2 n n n n n n n 1 2 2 1 n n n n 2 3 2 1 1 1 1 204. Macrochilus fenestratus n n n n n n n n n n 1 1 n 1 n n n n 3 2 ? 1 1 1 1 205. Rhabdostropha grindrodii n n n n n n n n n n 1 1 n 1 n n n n 2 2 2 1 1 1 1 206. Loxonema crossmanni n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 2 1 1 207. Loxonema sinuosa n n n n n n n n n n 1 1 n 1 n n n n 3 2 2 1 2 1 1 208. Auriptygma fortior n n n n n n n n n n 1 1 n ? ? ? ? ? 1 3 2 1 1 1 1 209. Catazone allevata 0 1 n n n n n n n i 1 2 2 1 n n n n 4 2 1 1 3 1 1 210. Catazone argolis 0 1 n n n n n n n i 1 2 2 1 n n n n 4 3 2 1 2 1 1 211. Catazone cunea 0 1 n n n n n n n i 1 2 2 1 n n n n 3 2 1 1 2 1 1 NUMBER 88 123 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 I 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 152. Fusispira Smithville Fm. sp. ? 1 2 1 1 1 1 1 ? ? 1 ? 1 ? 1 1 n 1 n 1 2 2 7 1 2 153. Hormotoma augustina ? 1 3 1 2 2 1 1 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 154. Hormoloma zelleri ? 1 3 1 1 9 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 155. Lophospira perangulata ? 1 3 1 1 2 1 1 1 7 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 156. Subulitid El Paso Fm. sp. ? 1 2 1 I 1 1 1 1 7 1 7 1 ? 1 1 n 1 n 1 1 2 7 1 2 157. Pagodospira cicelia ? 1 3 1 1 2 1 1 1 ? 1 7 I ? 1 1 n 1 n 1 1 2 6 1 2 158. Plethospira cannonensis ? 1 3 1 1 1 1 2 1 ? 1 7 1 7 1 1 n 1 n 1 3 2 5 I 2 159. Plethospira cassina 9 1 3 1 1 1 1 2 2 2 1 ? 1 7 1 1 n 1 n 1 3 2 6 1 2 160. Seelya ventricosa ? 1 3 1 1 1 1 2 2 2 1 7 1 ? 1 1 n 1 n 1 3 2 6 1 2 161. Lophospira grandis 9 1 3 1 1 1 1 1 1 1 1 ? 1 ? 1 2 n 2 2 1 1 2 5 1 2 162. Straparollina pelagica ? 1 3 1 1 1 1 1 2 1 1 ? 1 7 1 2 n 2 2 1 1 2 5 1 2 163. Plethospira ? turgida ? 1 4 1 1 2 1 1 2 1 1 ? 1 7 1 1 n 1 n 1 4 2 6 1 2 164. Turritoma acrea ? 1 3 7 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 1 2 6 1 2 165. Turritoma Cotter Fm. ornate sp. ? 1 3 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 166. Turritoma cf. T. acrea ? 1 3 1 1 2 1 1 1 7 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 167. Hormotoma Setul Fm. sp. ? 1 2 1 2 2 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 168. Turritoma lartna ? 1 3 1 1 2 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 169. Murchisonia callahanensis ? 1 3 1 1 2 1 1 1 7 1 9 1 ? 1 1 n 1 n 1 2 3 5 1 2 170. Ectomaria prisca ? 1 3 1 1 2 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 1 171. Hormotoma gracilis 7 1 3 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 172. Daidia cerithioides 7 1 1 ? 1 1 1 3 2 1 1 ? 1 7 1 1 n 1 n 1 2 2 7 1 2 173. Ectomaria pagoda ? 1 3 1 1 2 1 1 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 174. Haplospira 1 nereis 7 1 1 ? 1 1 1 3 2 2 1 7 1 7 1 I n 1 n 1 2 2 7 1 2 175. Hormotoma bellicincta ? 1 3 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 176. Hormotoma salteri ? 1 3 1 1 2 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 177. Hormotoma trentonensis ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 6 1 2 178. Loxonema murrayana ? 1 4 1 2 1 1 1 1 7 1 7 1 7 1 1 n 1 n 1 2 2 6 1 2 179. Omospira alexandra ? 1 3 1 1 1 1 1 1 ? 1 7 1 ? 1 1 n 1 n 1 2 2 6 1 2 180. Omospira laticincta ? 1 3 1 1 1 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 6 1 2 181. Straparollina circe ? 1 1 I 1 1 1 1 2 1 1 7 1 ? 1 1 n 1 2 1 1 2 2 1 2 182. Straparollina erigione ? 1 1 1 1 1 1 1 2 1 1 ? 1 ? 1 1 n 1 2 1 1 2 5 1 2 183. Girvania excavata ? 1 1 7 1 1 1 3 1 7 1 ? 1 ? 1 1 n 1 n 1 1 1 6 1 2 184. Murchisonia Pt. Clarence Fm. sp. 7 1 7 ? 1 1 1 3 1 ? 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 185. Rhabdostropha primitiva ? 1 1 7 1 1 1 3 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 186. Spiroecus girvanensis ? 1 1 ? 1 1 1 3 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 6 1 2 187. Daidia aff. D. cerithioides ? 1 1 7 1 1 1 3 2 1 1 ? 1 7 1 1 n 1 n 1 2 2 7 1 2 188. Ectomaria cf. E. pagoda ? 1 4 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 1 6 1 2 189. Ectomaria cf. E. prisca ? 1 3 1 1 2 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 1 1 6 1 2 190. Ectomaria laticarinata 9 1 3 1 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 2 3 5 1 2 191. Ectomaria nieszkowskii ? 1 2 1 1 2 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 1 2 6 I 2 192. Hormotoma insignis ? 1 3 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 193. Holopella regularis ? 1 4 1 1 1 1 3 1 ? 1 ? 1 7 1 1 n 1 n 1 2 2 6 1 2 194. Hormotoma centervillensis ? 1 3 1 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 1 2 6 1 2 195. Hormotoma cingulata 7 1 2 1 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 2 2 6 1 2 196. Kjerulfonema cancellata 7 1 4 1 1 1 1 3 1 7 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 197. Kjerulfonema quinquecincta 7 1 3 1 1 1 1 3 1 7 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 198. Cyrtostropha coralli ? 1 3 1 1 2 1 1 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 199. Goniostropha cava ? 1 2 1 1 2 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 200. Hormotoma subplicata ? 1 3 1 1 2 1 1 1 ? 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 201. Hormotoma monoliniformis ? 1 1 1 1 2 1 1 1 ? 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 202. Hormotoma attenuata ? 1 3 1 1 2 1 1 1 7 1 7 1 7 1 1 n 1 n 1 1 2 6 1 2 203. Loxonema ? attenuata ? 1 3 1 1 2 1 2 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 204. Macrochilus fenestratus ? 1 1 ? 1 1 1 3 2 1 1 ? 1 ? 1 1 n 1 n 1 2 2 6 1 2 205. Rhabdostropha grindrodii ? 1 1 ? 1 1 1 3 1 ? 1 7 1 ? 1 1 n 1 n 1 1 1 6 1 2 206. Loxonema crossmanni ? 1 4 1 2 1 1 1 1 7 1 7 1 ? 1 1 n 1 n 1 2 2 6 1 2 207. Loxonema sinuosa ? 1 4 1 2 1 1 2 1 ? 1 7 1 ? 1 1 n 1 n 1 2 2 6 1 2 208. Auriptygma fortior ? 1 1 1 2 2 1 1 1 7 1 ? 1 ? 1 1 n 1 n 1 2 2 6 1 2 209. Catazone allevata 7 1 2 1 1 1 1 3 1 ? 1 ? 1 ? 1 1 n 1 n 1 2 2 5 1 2 210. Catazone argolis ? 1 2 ? 1 1 1 3 1 ? 1 ? 1 7 1 1 n 1 n 1 2 2 6 1 2 211. Catazone cunea ? 1 2 1 1 1 1 3 1 ? 1 ? 1 7 1 1 n 1 n 1 2 2 6 1 2 124 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 152. Fusispira Smithville Fm. sp. 7 2 ? 1 n n n 1 n n n n 1 1 1 3 ? n 153. Hormotoma augustina 7 1 1 1 n n n 1 n n n n 1 1 1 3 ? n 154. Hormotoma zelleri 8 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 155. Lophospira perangulata 6 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 156. Subulitid El Paso Fm. sp. 8 2 ? 1 n n n 1 n n n n 1 1 1 2 ? n 157. Pagodospira cicelia 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 158. Plethospira cannonensis 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 159. Plethospira cassina 7 2 1 1 n n n 1 n n n n 1 1 1 3 ? n 160. Seely a ventricosa 6 2 1 2 2 2 1 2 2 2 1 1 1 1 1 3 ? n 161. Lophospira grandis 6 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 162. Straparollina pelagica 6 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 163. Plethospira! turgida 7 2 1 1 n n n 1 n n n 11 1 1 1 2 ? n 164. Turritoma acrea 7 2 1 ? n n n 1 n n n n 1 1 1 2 ? n 165. Turritoma Cotter Fm. ornate sp. 7 2 1 2 3 2 i 1 3 2 i i 1 1 1 1 ? n 166. Turritoma cf. T. acrea 7 2 ? 1 n n n 1 n n n n 1 1 1 1 ? n 167. Hormotoma Setul Fm. sp. 7 1 ? 1 n n n 1 n n n n 1 1 1 3 ? n 168. Turritoma! anna 8 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 169. Murchisonia callahanensis 7 3 1 1 n n n 1 n n n n 1 1 1 2 7 n 170. Ectomaria prisca 8 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 171. Hormotoma gracilis 8 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 172. Daidia cerithioides 7 1 1 1 n n n 1 n n n n 1 1 1 2 ? n 173. Ectomaria pagoda 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 174. Haplospira !nereis 5 1 l 1 n n n 1 n n n n 1 1 1 ? ? n 175. Hormotoma bellicincta 7 2 1 1 n n n 1 n n n n 1 1 1 3 7 n 176. Hormotoma salteri 8 I 1 1 n n n 1 n n n n 1 1 1 1 ? n 177. Hormotoma trentonensis 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 178. Loxonema murrayana 7 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 179. Omospira alexandra 7 2 1 1 n n n 1 n n n n 1 1 1 3 ? n 180. Omospira laticincta 7 2 1 1 n n n 1 n n n n 1 1 1 3 ? n 181. Straparollina circe 4 1 1 1 n n n 1 n n n n 1 1 1 ? ? n 182. Straparollina erigione 5 1 1 1 n n n 1 n n n n 1 1 1 ? 7 n 183. Girvania excavata 7 2 1 2 3 1 1 2 i i i i 1 1 I 2 ? n 184. Murchisonia Pt. Clarence Fm. sp. 7 1 1 1 n n n 2 i i i i 1 1 1 2 ? n 185. Rhabdostropha primitiva 7 1 1 2 2 i i 2 i i i i 1 1 1 2 ? n 186. Spiroecus girvanensis 7 1 1 2 3 i i 2 i i i i 1 1 1 2 ? n 187. Daidia aff. D. cerithioides 7 1 1 1 n n n 1 n n n n 1 1 1 2 ? n 188. Ectomaria cf. E. pagoda 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 189. Ectomaria cf. E. prisca 7 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 190. Ectomaria laticarinata 6 3 1 1 n n n 1 n n n n 1 1 1 1 ? n 191. Ectomaria nieszkowskii 7 1 1 1 n n n 1 n n n n 1 1 1 2 ? n 192. Hormotoma ins ignis 7 1 1 1 n n n 1 n n ii n 1 1 1 1 2 2 193. Holopella regularis 7 1 1 1 n n n 1 n n n n I n 1 2 ? n 194. Hormotoma centervillensis 8 1 1 1 n n n 1 n n n n 1 i 1 2 ? n 195. Hormotoma cingulata 7 2 1 1 n n n 1 n n n n 1 i 1 2 ? n 196. Kjerulfonema cancellata 7 1 1 2 i 3 i 2 2 3 l i 1 n 1 2 ? n 197. Kjerulfonema quinquecincta 7 1 1 2 i 3 i 2 2 3 l i 1 n 1 2 ? n 198. Cyrtostropha coralli 8 1 1 1 n n n 1 n n n n 1 i 1 2 ? n 199. Goniostropha cava 8 1 1 1 n n n 1 n n n n 1 i 1 1 ? n 200. Hormotoma subplicata 8 1 1 1 n n n 1 n n n n 1 i 1 1 7 n 201. Hormotoma monoliniformis 7 1 1 1 n n n 1 n n n n 1 i 1 2 ? n 202. Hormotoma attenuata 8 1 1 1 n n n 1 n n n n 1 i 1 2 7 n 203. Loxonema! attenuata 8 1 1 1 n n 11 1 n n n n 1 i 1 2 ? n 204. Macrochilus fenestratus 8 1 1 2 4 2 1 1 n n n n 1 n 1 2 7 n 205. Rhabdostropha grindrodii 7 1 1 2 1 2 1 2 2 2 i 1 1 1 1 ? 7 n 206. Loxonema crossmanni 7 1 1 1 n n n 1 n n n n 1 n 1 2 ? n 207. Loxonema sinuosa 7 1 1 1 n n n 1 n n n n 1 n 1 2 1 2 208. Auriptygma fortior 7 1 1 1 n n n 1 n n n n 1 1 1 1 7 n 209. Catazone allevata 8 2 1 1 n n n 1 n n n n 1 1 1 2 ? n 210. Catazone argo!is 8 2 1 2 3 1 1 2 3 1 1 1 1 2 1 2 ? n 211. Catazone cunea 8 2 1 1 n n n 1 n n n n 1 1 1 2 ? n NUMBER 88 125 Appendix 2.—Continued. Number Species 212. Diplozone crispa 213. Donaldiella declivis 214. Donaldiella morinensis 215. Goniostropha sculpta 216. Loxonema beraultensis 217. Coelocaulus concinnus 218. Macrochilus buliminus 219. Macrochilus cancellatus 220. Macrochilina reciicosia 221. Murchisonia paradoxa 222. Sinuspira tenera 223. Stylonema mater 224. Stylonema potens 225. Clathrospira Smithville Fm. sp. 226. Clathrospira Iglindmeveri 227. Clathrospira elliptica 228. Clathrospira euconica 229. Clathrospira in/lata 230. Mourlonia mjoela 231. Clathrospira Itrochiformis 232. Clathrospira convexa 233. Clathrospira conica 234. Clathrospira subconica 235. Eotomaria canalifera 236. Eotomaria dryope 237. Eotomaria labrosa 238. Liospira larvata 239. Paraliospira mundula 240. Eotomaria supracingulata 241. Liospira angustata 242. Liospira decipens 243. Liospira subconcava 244. Euryzone kiari 245. Eotomaria elevata 246. Liospira micula 247. Liospira progne 248. Paraliospira angulata 249. Brachytomaria baltica 250. Paraliospira aff. P. angulata 251. Paraliospira rugata 252. Eotomaria notablis 253. Lophospira kindlei 254. Brachytomaria papillosa 255. Brachytomaria semele 256. Brachytomaria striata 257. Cataschisma exquisita 258. Clathrospira thraivensis 259. “ Bembexia ” globosa 260. Eotomaria rupestris 261. Crenilunula limata 262. Clathrospira biformis 263. Phanerotrema jugosa 264. Phanerotrema lindstroemi 265. Oriostoma angulifer 266. Stenoloron shelvensis 267. “ Seelya ” lloydi 268. Ulrichospira similis 269. Eocryptaulina helcinia 270. Conotoma claustrata 271. Crenilunula hallei 1 2 ~ 2 2 2 2 2 1 1 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 1 1 2345678901 2 4 4 2 2 2 2 3 3 1 3 3 5 3 3 4 13 4 1 3 4 4 3 3 4 2 3 3 1 3 3 4 3 3 3 2 3 3 1 2 4 4 2 3 3 2 3 3 1 3 3&4 6 2 2 3 1 3 4 1 ?????????? 9999999999 2 0 0 3 3 3 3 5 2 3 3 5 7 3 3 2 112 3 2 112 3 3 4 5 1 3 2 4 5 1 1 3 4 6 13 2 3 5 1 3 3 4 6 1 3 3 4 6 13 3 4 5 13 3 3 4 13 3 4 5 1 3 3 4 5 1 3 3 4 5 13 3 4 5 13 3 4 5 1 3 3 4 5 13 3 4 5 13 3 4 6 13 3 4 5 2 3 3 4 5 2 3 3 4 5 2 3 3 2 5 1 3 3 3&4 4 3 3 3 3 5 2 3 3 4 5 2 3 3 3 5 2 2 2 112 1 3 2 5 2 2 3 3 5 2 2 3 4 5 13 3 2 4 2 3 2 2 2 2 2 2 2 2 2 3 2 2 2 2 2 3 4 2 1 2 3 4 4 3 3 3 2 4 3 3 3 4 4 12 3 14 11 3 2 4 2 3 2 2 2 2 1 2 2 2 2 1 13 2 11 13 2 12 2 3 3 2 3 2 2 2 2 2 2 2 4 1 2 3 2 5 1 1 3 14 11 3 2 111 3 13 4 1 4 13 4 1 3 2 111 3 2 111 3 2 3 3 1 12 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 13 3 1 3 13 3 1 3 13 3 1 3 13 4 1 3 13 4 1 3 13 4 1 3 2 3 3 1 3 13 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 13 4 1 1 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 13 4 1 3 2 3 3 1 2 2 3 3 1 3 2 3 3 1 2 2 3 3 1 2 2 3 3 1 3 2 3 3 1 3 2 3 3 1 3 13 4 1 1 2 3 3 1 1 2 3 3 1 12 3 3 1 1 2 3 3 1 1 7 3 3 1 2 2 3 3 1 3 2 3 3 1 2 2 3 3 1 2 2 3 3 1 1 2 3 3 1 1 2 3 3 1 1 1 2 3 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 2 1 1 1 1 1 1 1 1 1111 4 5 6 7 n 3 i n 4 1 n 3 1 n 4 1 n 3 1 n 3 I n 3 1 n 3 1 n 4 1 n 3 1 n 3 1 n 3 1 n 3 1 n 3 1 n 4 1 n 3 1 n 3 1 n 3 1 n 3 n 3 n 3 1 1 n 3 1 n 3 1 n 3 1 n 3 1 n 3 1 n 4 1 n 4 1 n 4 1 n 2 1 n 2 1 n 2 1 n 3 1 n 3 1 n 2 1 n 2 1 n 4 1 n 5 1 n 4 1 n 4 2 3 n 4 1 1 n 4 1 1 n 5 1 I n 4 1 1 n 5 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 4 1 1 n 3 1 1 n 4 1 1 n 4 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 3 1 1 n 4 1 1 112 2 8 9 0 1 linn 12 2 2 12 2 2 12 4 2 linn 12 4 2 linn linn linn 12 2 2 12 11 linn linn 12 3 2 1 2 4&5 2 12 3 2 12 3 2 12 3 2 12 5 2 12 3 2 12 4 2 12 3 2 12 3 2 12 3 2 12 4 2 12 2 2 12 3 2 12 4 2 12 3 2 12 2 2 12 2 2 12 2 2 12 4 2 12 3 2 12 2 2 12 2 2 12 3 2 12 4 2 12 4 2 12 4 2 12 3 2 12 4 2 12 4 2 12 4 2 12 4 2 12 4 2 12 3 2 12 3 2 12 2 2 12 2 2 1 2 3&4 2 12 2 2 12 2 2 12 12 12 12 12 3 2 12 3 2 12 2 2 12 3 2 12 3 2 2 2 2 2 2 3 4 5 n n n n 2 2 1 n 2 3&4 I n 2 4 1 n n n n n 2 2 1 n n n n n n n n n n n n n 2 1 1 n n n n n n n n n n n n n 2 3 1 n 2 4 1 n 2 3 1 n 2 3 1 n 2 3 1 n 2 3 1 n 2 4 1 n 1 2 1 n 3 4 1 n 3 4 1 n 3 4 1 n 3 4 1 n 2 3 1 n 3 4 1 n 3 4 1 n 3 5 1 n 2 1 1 n 2 3 1 n 2 1 1 n 2 4 l n 2 3 1 n 2 1 1 n 2 1 1 n 3 4 1 n 2 5 1 n 3 5 1 n 3 5 1 n 3 3 1 n 2 4 1 n 2 5 1 n 2 5 1 n 2 5 1 n 1 2 1 n 2 3 1 n 2 3 1 n 2 2 1 n 2 5 2 2 2 3 1 n 2 5 1 n 2 5 1 n 2 3 1 n 2 3 1 n 2 3 1 n 2 3 1 n 2 1 1 n 2 5 2 1 2 5 2 2 126 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 I 2 3 4 5 6 7 8 9 0 212. Diplozone crispa n n n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 B 1 n 213. Donaldiella declivis 1 1 n n n n n 1 2 2 2 1 1 I 1 1 1 2 1 1 n 1 9 1 n 214. Donaldiella morinensis 1 1 n n n n n 1 1 n n 1 1 2 3 1 I 2 1 1 n 1 7 1 n 215. Goniostropha sculpta 1 1 n n n n n 3 2 2 2 1 1 1 1 1 1 2 1 1 n 1 7 1 n 216. Loxonema beraultensis n n n n n n n n 1 n n n I 2 n n n n n 1 n 1 8 1 n 217. Coelocaulus concinnus 1 1 n n n n n 1 2 2 2 1 1 2 1 1 1 2 1 1 n 1 7 1 n 218. Macrochilus buliminus n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 D 1 n 219. Macrochilus cancellatus n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 D 1 n 220. Macrochilina recticosta n n n n n n n n 1 n n n 1 2 n n n n n 1 n 1 C 1 n 221. Murchisonia paradoxa n 1 n n n n n n 2 2 2 1 1 1 1 1 1 2 1 1 n 1 8 1 n 222. Sinuspira tenera n 2 1 1 1 1 1 1 1 n n n 1 1 1 1 1 2 I 1 n 1 8 1 n 223. Stylonema mater i n n n n n n n 1 n n n n 2 n n n n n 1 n 1 8 1 n 224. Stylonema potens i n n n n n n n 1 n n n n 2 n n n n n 1 n 1 8 1 n 225. Clathrospira Smithville Fm. sp. i i n n n n n 2 1 n n 1 1 2 1 1 1 3 1 2 1 1 7 1 n 226. Clathrospira Iglindmeyeri i i n n n n n 3 1 n n 1 1 2 1 1 1 2 1 1 n 1 8 1 n 227. Clathrospira elliptica i 2 2 i i i 1 I 1 n n 1 1 2 1 1 1 3 1 2 1 1 7 1 n 228. Clathrospira euconica i 1 n n n n n 2 1 n n 1 1 2 1 1 1 3 1 1 n 1 7 1 n 229. Clathrospira inflata i 1 n n n n n 1 1 n n 1 1 2 1 1 1 3 1 2 i 1 7 1 n 230. Mourlonia mjoela i 1 n n n n n 1 1 n n 1 1 2 1 1 1 3 1 1 n 1 8 1 n 231. Clathrospira Itrochiformis i 1 n n n n n 2 1 n n 1 1 2 1 1 1 3 1 2 i 1 8 1 n 232. Clathrospira convexa i 1 n n n n n 1 1 n n 1 1 2 1 1 1 2 1 1 n 1 6 1 n 233. Clathrospira conica i 1 n n n n n 2 1 n n 1 1 2 1 1 1 3 1 2 n 1 7 1 n 234. Clathrospira subconica i 1 n n n n n 2 2 i 2 1 1 2 1 1 1 2 1 2 n 1 8 1 n 235. Eotomaria canalifera i 1 n n n n n 2 1 n n 1 1 2 1 1 1 4 1 2 l 1 4 1 n 236. Eotomaria dryope i 1 n n n n n 2 1 n n 1 1 2 1 1 1 4 1 2 1 1 7 1 n 237. Eotomaria labrosa i 1 n n n n n 2 1 n n 1 1 2 1 I 1 4 1 2 1 1 4 1 n 238. Liospira larvata i 1 n n n n n 2 1 n n 1 1 2 1 1 1 4 1 2 2 1 6 1 n 239. Paraliospira mundula i 1 n n n n n 2 1 n n 1 1 2 1 1 1 4 1 2 2 1 7 1 n 240. Eotomaria supracingulata i 1 n n n n n 2 1 n n 1 1 2 1 1 1 4 1 2 2 1 6 1 n 241. Liospira angustata i 1 n n n n n 2 ? n n 1 n n 1 1 1 3 1 1 n 1 4 1 n 242. Liospira decipens i 1 n n n n n 2 1 n n 1 i 2 1 1 1 3 1 1 n 1 4 1 n 243. Liospira subconcava i 1 n n n n n 2 1 n n 1 i 2 1 n 1 3 1 1 n 1 4 1 n 244. Euryzone kiari i 1 n n n n n 2 1 n n 1 i 2 1 i 1 2 1 1 n 1 9 1 n 245. Eotomaria elevata i 1 n n n n n 1 1 n n 1 i 2 1 i 1 4 1 1 n 1 A 1 n 246. Liospira micula i 1 n n n n n 2 2 2 2 1 i 1 1 i 1 3 1 1 n 1 5 1 n 247. Liospira progne i 1 n n n n n 2 2 2 2 1 i 1 1 i 1 3 1 2 i 1 4 1 n 248. Paraliospira angulata i 1 n n n n n 2 1 n n 1 i 2 1 i 1 4 1 2 2 1 9 1 n 249. Brachytomaria baltica i 1 n n n n n 2 1 n n 1 i 3 1 i 1 2 1 1 n 1 8 1 n 250. Paraliospira aff. P angulata i 1 n n n n n 2 1 n n 1 i 2 1 i 1 4 1 2 2 1 6 3 n 251. Paraliospira rugata i 1 n n n n n 2 1 n n 1 i 2 1 i 1 4 1 2 2 1 9 1 n 252. Eotomaria notablis i 1 n n ii n n 2 1 n n 1 i 2 1 i 1 4 1 2 1 1 7 1 n 253. Lophospira kindlei i 1 n n n n n 2 1 n r» 1 i 2 1 i 1 2 1 1 n 1 9 1 n 254. Brachytomaria papillosa i 1 n n n n n 2 1 n n 1 i 3 1 i 1 2 1 1 n 1 B 1 n 255. Brachytomaria semele i 1 n n n n n 2 1 n n 1 i 3 1 i 1 2 1 1 n 1 9 1 n 256. Brachytomaria striata i 1 n n n n n 2 1 n n 1 i 3 1 i 1 2 1 1 n 1 B 1 n 257. Cataschisma exquisita i 1 n n n n n 1 2 2 2 1 i 1 1 i 1 2 1 1 n 1 4 1 n 258. Clathrospira thraivensis i 1 n n n n n 1 1 n n 1 i 2 1 i 1 4 1 1 n 1 9 1 n 259. “ Bembexia” globosa i 1 n n n n n 2 1 n n 1 i 2 1 i 1 4 1 1 n 1 A 1 n 260. Eotomaria rupestris i 1 n n n n n 1 2 2 2 1 i 2 1 i 1 4 1 2 2 1 A 3 n 261. Crenilunula limata 2 1 n n n n n 2 2 1 2 1 2 3 1 i 1 1 1 1 n 1 2 1 n 262. Clathrospira biformis 1 1 n n n n n 2 1 n n 1 1 2 1 i 1 4 1 1 n 1 8 1 n 263. Phanerotrema jugosa 1 1 n n n n n 2 ? n n 1 1 3 1 i 1 3 1 1 n 1 8 1 n 264. Phanerotrema lindstroemi 1 1 n n n n n 2 2 2 i 1 1 3 1 i 1 3 1 1 n 1 8 1 n 265. Oriostoma angulifer 1 1 n n n n n 1 2 2 2 1 1 1 1 i 1 4 1 2 1 1 8 l n 266. Stenoloron shelvensis 1 1 n n n n n 1 2 2 2 1 1 1 1 i 1 4 1 2 1 1 5 1 n 267. “ Seelya ” lloydi 1 1 n n n n n 2 2 2 2 1 1 2 1 i 1 2 1 1 n 1 B 1 n 268. Ulrichospira similis 1 1 n n n n n 2 ? n n 1 1 2 1 i 1 2 1 1 n 1 B 1 n 269. Eocryptaulina helcinia 1 1 n n n n n 1 2 2 2 1 1 1 1 i 1 4 1 1 n 1 A 1 n 270. Conotoma claustrata 2 1 n n n n n 1 1 n n 1 1 3 1 i 1 1 1 1 n 1 6 1 n 271. Crenilunula hallei 2 1 n n n n n 2 2 1 2 1 2 3 1 i 2 1 1 1 n 1 2 1 n NUMBER 88 127 appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 6 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 I 2 3 4 5 6 7 8 9 0 1 2 3 4 5 212. Diplozone crispa 3 3 2 3 1 4 1 6 4 1 n n 1 1 n n n n 2 2 2 2 1 n 1 213. Donaldiella declivis 2 3 2 3 3 6 1 7 3 1 n n 1 1 n n n n 2 1 1 3 1 n 2 214. Donaldiella morinensis 2 3 1 1 4 5 1 5 4 1 n n 1 1 n n n n 2 2 1 3 1 n 1 215. Goniostropha sculpta 2 3 2?3 3 4 5 1 6 4 1 n n 1 1 n n n n 2 n 1 3 1 n 1 216. Loxonema beraultensis 2 2 2 3 3 4 1 7 4 1 n n 1 1 n n n n 2 2 1 2 1 n 1 217. Coelocaulus concinnus 3 2 2 I 5 2&3 2 5 4&5 1 n n 1 1 n n n n 1 n 1 1 1 n 1 218. Macrochilus buliminus 2 2 2 3 0 6 1 7 1 1 n n 1 1 n n n n 2 i 1 2 1 n 1 219. Macrochilus cancellatus 2 2 2 3 0 6 1 7 1 1 n n 1 1 n n n n 2 i 1 2 1 n 1 220. Macrochilina reclicosta 2 2 2 3 1 4 1 8 3 1 n n 1 1 n n n n 2 i I 2 1 n 1 221. Murchisonia paradoxa 2 2 2 1 5 4 1 6 5 1 n n 1 1 n n n n 2 i 1 2 1 n 2 222. Sinuspira tenera 2 2 2 2 3 3 1 6 3 1 n n 1 1 n n n n 2 2 1 2 1 n 1 223. Stylonema mater 2 2 2 3 2 5 1 6 5 2 n n 1 1 n n n n 2 1 1 2 2 i 1 224. Stylonema potens 2 2 2 3 2 5 1 6 5 2 n n 1 1 n n n n 2 1 1 2 2 ■ 1 225. Clathrospira Smithville Fm. sp. 2 3 2 1 6 2 2 4 4 1 n n 1 1 n n n n 1 n 1 3 1 n 1 226. Clathrospira Iglindmeyeri 2 3?4 2 1 7 4 1 5 2 1 n n 1 2 1 1 1 1 1 n 1 3 1 n 2 227. Clathrospira elliptica 2 2&3 2 1 5 2 1 4 2 1 n n 1 1 n n n n 1 n 1 2&3 1 n 1 228. Clathrospira euconica 2 3 2 l 3 3 1 5 2 1 n n 1 2 i 3 i i 1 n 1 2&3 1 n 1 229. Clathrospira in/lata 2 2&3 2 1 5 2 1 4 2 1 n n 1 1 n n n n 1 n 1 2&3 1 n 1 230. Mourlonia mjoela 2 3 2 1 5 2 1 5 1 1 n n 1 1 n n n n 1 n 1 2&3 1 n 1 231. Clathrospira Itrochiformis 2 3 2 1 6 3 2 4 4 1 n n 1 1 n n n n 1 n 1 3 1 n 1 232. Clathrospira convexa 2 1 2 1 6 2 2 3 3 1 n n 1 1 n n n n 1 n 1 1 1 n 1 233. Clathrospira conica 2 3 2 1 5 3 2 4 4 1 n n 1 1 n n n n 1 n 1 3 1 n 1 234. Clathrospira subconica 2 3 2 1 5 3 2 4 4 1 n n 1 1 n n n n 1 n 1 2&3 1 n 1 235. Eotomaria canalifera 2 3 2 1 4 4 1 4 4 1 n n 1 2 2 1 2 1 1 n 1 3 1 n 1 236. Eotomaria dryope 2 3 2 1 4 3 1 5 2 1 n n 1 2 1 3 2 1 1 n 1 3 1 n 2 237. Eotomaria labrosa 3 3 2 1 4 4 1 4 3 1 n n 1 2 2 1 2 1 1 n 1 2 1 n 1 238. Liospira larvata 2 3 2 1 4 3 3 3 4 1 n n 1 2 2 1 2 1 1 n 1 3 1 n 1 239. Paraliospira mundula 3 3 1 2 4 2 3 3 4 1 n n 1 2 3 3 1 2 1 n 1 3 1 n 2 240. Eotomaria supracingulata 2 3 2 1 4 3 3 3 4 1 n n 1 2 2 1 2 1 1 n 1 3 1 n 1 241. Liospira angustata 3 3 2 1 4 2 1 2 1 1 n n 1 2 1 1 1 1 1 n 1 2 1 n 1 242. Liospira decipens 3 3 2 1 6 5 1 2 1 1 n n 1 2 1 1 1 1 1 n 1 2 1 n 1 243. Liospira subconcava 3 4 2 1 6 5 1 2 1 1 n n 1 2 1 1 1 1 1 n 1 3 1 n 1 244. Euryzone kiari 2 2 2 1 5 4 1 5 4 1 n n 1 2 1 3 1 1 1 n 1 2 1 n 1 245. Eotomaria elevata 3 4 2 3 2 4 1 6 2 1 n n 1 2 3 4 1 1 2 i 1 4 1 n 2 246. Liospira micula 3 3 2 1 6 5 2 1 1 1 n n 1 2 1 1 1 1 1 n 1 3 1 n 1 247. Liospira progne 3 3 2 1 4 2 2 2 2 1 n n 1 2 1 1 1 1 1 n 1 2 1 n 1 248. Paraliospira angulata 2 3 1 1 4 2 3 3 4 1 n n 1 2 2 3 1 n 1 n 1 3 1 n 2 249. Brachytomaria baltica 2 2 2 2 5 2 1 5 4 1 n n 1 1 n n n n 1 n 1 3 1 n 1 250. Paraliospira aflf. P. angulata 2 2 1 1 5 2 3 3 4 1 n n 1 2 2 3 1 2 1 n 1 2 1 n 2 251. Paraliospira rugata 2 3 1 1 4 2 3 3 4 1 n n 1 2 3 3 1 2 1 n 1 2 1 n 2 252. Eotomaria notablis 2 2 2 1 4 4 1 3 1 1 n n 1 2 1 3 2 1 1 n 1 2 1 n 2 253. Lophospira kindlei 2 2 2 2 4 4 1 5 4 1 n n 1 2 2 4 1 1 1 n 1 2 1 n 2 254. Brachytomaria papillosa 2 2 2 3 3 4 1 6 4 1 n n 1 2 2 4 1 1 1 n 1 3 1 n 2 255. Brachytomaria semele 2 2 2 2 4 4 1 5 4 1 n n 1 2 2 4 1 1 1 n 1 2 1 n 2 256. Brachytomaria striata 2 2 2 3 3 4 1 6 4 1 n n 1 2 1 2 1 1 1 n 1 3 1 n 2 257. Cataschisma exquisita 2 1 2 1 7 2 3 4 5 1 n n 1 1 n n n n 1 n 1 1 1 n 1 258. Clathrospira thraivensis 3 4 2 3 2 4 1 5 3 1 n n 1 2 I 4 1 1 2 1 1 3 1 n 2 259. “ Bembexia ” globosa 3 3 2 3 2 4 1 5 4 1 n n 1 2 3 3 1 1 1 n 1 3 1 n 1 260. Eotomaria rupestris 2 2 2 1 4 2 3 3 4 1 n n 1 2 1 3 1 n 1 n 1 2 1 n 1 261. Crenilunula limata 1 2 2 1 8 2 3 2 3 1 n n 1 1 n n 1 1 1 n 1 3 1 n 1 262. Clathrospira biformis 3 3 2 3 1 2 1 6 4 1 n n 1 2 3 3 1 1 1 n 1 3 1 n 1 263. Phanerotrema jugosa 1 2 2 3 3 4 1 6 4 1 n n 1 2 2 2 1 1 2 i 1 3 1 n 1 264. Phanerotrema lindstroemi 1 2 1 3 3 4 1 5 4 1 n n 1 2 1 2 1 1 2 i 1 3 1 n 1 265. Oriostoma angulifer 3 3 2 1 5 2 2 4&5 4 1 n n 1 2 2 3 1 1 1 n 1 3 1 n 1&2 266. Stenoloron shelvensis 3 2 2 2 4 4 2 4 4 I n n 1 2 2 1 1 1 1 n 1 1 1 n 1 267. “ Seelya ” lloydi 2 2 2 3 3 4 1 6 3 1 n n 1 2 1 2 1 1 1 n 1 2 1 n 2 268. Ulrichospira similis 2 3 2 3 3 4 1 6 3 1 n n 1 2 1 4 1 1 1 n 1 3 1 n 2 269. Eocryptaulina helcinia 2 2 2 1 3 2 2 3 5 1 n n 1 1 n n n n 1 n 1 2 1 n 1 270. Conotoma claustrata 1 2 2 1 7 2 2 2 2 1 n n 1 1 n n n n 1 n 1 3 1 n 1 271. Crenilunula hallei 1 2 2 1 8 1 3 2 3 1 n n 1 1 n n n n 1 n 1 3 1 n 1 128 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 1 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 1 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 212. Diplozone crispa n n n n n n n n n n 1 1 n 1 n n n n 3 1 ? 1 2 1 1 213. Donaldiella declivis 2 2 2 n n n n n n n 1 2 2 1 n n n n 2 2 2 1 1 1 1 214. Donaldiella morinensis n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 1 1 1 215. Goniostropha sculpta n n n n n n n n n n 1 2 2 1 n n n n 4 3 1 1 1 1 1 2 16. Loxonema beraultensis n n n n n n n n n n 1 ? ? 1 n n n n 3 1 2 1 2 1 1 217. Coelocaulus concinnus n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 1 1 1 218. Macrochilus buliminus n n n a n n n n n n 1 1 n 1 n n n n 1 3 1 1 1 1 1 219. Macrochilus cancellatus n n n n n n n n n n 1 1 n 1 n n n n 1 3 1 1 1 1 1 220. Macrochilina recticosta n n n n n n n n n n 1 ? n 1 n n n n 2 1 1 1 2 1 1 221. Murchisonia paradoxa 2 3 1 n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 222. Sinuspira tenera n n n n n n n n n n 1 2 2 1 n n n n 2 3 2 1 1 1 ? 223. Stylonema mater n n n n n n n n n n 1 ? ? 1 n n n n 3 2 2 1 2 1 ? 224. Stylonema potens n n n n n n r» n n n 1 ? ? 1 n n n n 3 2 2 1 2 1 ? 225. Clathrospira Smithville Fm. sp. n n n n n n n n n n I 2 2 1 n n n n 4 3 2 1 1 1 1 226. Clathrospira Iglindmeveri i i i n n n n n n 1 1 1 n 1 n n n n 3 3 2 1 2 1 1 227. Clathrospira elliptica n n n n n n n n n n 1 2 2 1 n n n n 4 3 2 1 1 1 1 228. Clathrospira euconica n n n n n n n n n n 1 2 3 1 n n n n 4 3 2 1 2 1 1 229. Clathrospira inflata n n n n n n n n n n 1 2 2 1 n n n n 4 3 2 1 1 1 1 230. Mourlonia mjoela n n n n n n n n n n 1 2 2 1 n n n n 4 3 2 1 1 1 1 231. Clathrospira Itrochiformis n n n n n n n n n n 1 2 3 1 n n n n 4 3 2 1 1 1 1 232. Clathrospira convexa n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 233. Clathrospira conica n n n n n n n n n i 1 2 3 1 n n n n 4 3 2 1 2 1 1 234. Clathrospira subconica n n n n n n n n n i 1 2 3 1 n n n n 4 3 2 1 1 1 1 235. Eotomaria canalifera n n n n ■ i n n n n n 1 2 3 1 n n n n 4 3 2 1 4 1 1 236. Eotomaria dryope i i 1 n n n n n n i 1 2 2 1 n n n n 4 3 2 1 2 1 1 237. Eotomaria labrosa n n n n n n n n n n 1 3 2 1 n n n n 4 2 1 1 4 1 1 238. Liospira larvata n n n n n n n n n n 1 2 3 1 n n n n 5 2 3 1 4 1 1 239. Paraliospira mundula 1 1 2 n n n n n n n 1 3 3 2 3 2 i i 5 2 3 1 3 1 1 240. Eotomaria supracingulata n n n n n n n n n n 1 2 3 1 n n n n 5 2 3 1 4 1 1 241. Liospira angustata n n n n n n n n n n 1 1 n 1 n n n n 5 2 1 1 4 1 1 242. Liospira decipens n n n n n n n n n n 1 3 3 1 n n n n 5 2 1 1 4 1 1 243. Liospira subconcava n n n n n n n n n n 1 3 2 1 n n n n 5 2 1 1 4 1 1 244. Euryzone kiari n n n n n n n n n n 1 2 2 1 n n n n 4 2 2 1 1 1 1 245. Eotomaria elevata 2 i i n n n n n n 1 1 2 3 1 n n n n 2 3 2 1 1 1 1 246. Liospira micula n n n n n n n n n n 1 2 2 1 n n n n 5 2 1 1 3 1 1 247. Liospira progne ii n n n n n n n n n 1 2 3 1 n n n n 5 2 1 1 4 1 1 248. Paraliospira angulata i i 2 n n n n n n n 1 2 3 2 3 2 1 1 5 2 3 1 2 1 2 249. Brachytomaria baltica n n n n n n n n n n 1 3 3 1 n n n n 3 3 2 1 1 1 1 250. Paraliospira aff. P. angulata i i 2 n n n n n n n 1 2 3 2 3 2 i i 5 2 3 1 4 1 2 25 1 . Paraliospira rugata i i 2 n n n n n n n 1 3 3 ? ? ? ? ? 5 2 3 1 2 1 ? 252. Eotomaria notablis i i 2 n n n n n n i 1 2 3 1 n n n n 4 2 2 1 2 1 1 253. Lophospira kindlei 2 2 1 n n n n n n i 1 2 3 1 n n n n 4 3 2 1 1 1 1 254. Brachytomaria papillosa 2 2 1 n n n n n n i 1 3 3 1 n n n n 3 3 2 1 1 1 1 255. Brachytomaria semele 2 2 1 n n n n n n i 1 2 3 1 n n n n 4 3 2 1 1 1 1 256. Brachytomaria striata 1 1 1 n n n n n n i 1 3 3 1 n n n n 3 3 2 1 1 1 1 257. Cataschisma exquisita n n n n n n n n n n 1 1 n 1 n n n n 3 3 2 1 2 1 1 258. Clathrospira thraivensis 2 i 1 n n n n n n i 1 2 3 1 n n n n 3 2 2 1 2 1 1 259. “ Bembexia ” globosa n n n n n n n n n n 1 2 3 1 n n n n 4 2 2 1 1 1 1 260. Eotomaria rupestris n n n n n n n n n n 1 2 2 2 3 2 1 1 3 2 2 1 2 1 2 26 1 . Crenilunula limata n n n n n n n n n n 1 2 2 1 11 n n n 5 2 2 1 4 1 1 262. Clathrospira biformis n n n n n n n n n n 1 2 3 2 4 i i i 4 2 1 1 3 1 1 263. Phanerotrema jugosa n n n n n n n n n n 1 3 3 1 n n n n 3 3 1 1 3 1 1 264. Phanerotrema lindstroemi n n n n n n n n n n 1 3 3 1 n n n n 3 2 1 1 3 1 1 265. Oriostoma angulifer 2 2 n n n n n n n n 1 1 n 2 4 2 i 2 4 1 2 1 2 1 2 266. Stenoloron shelvensis n n n n n n n n n n 1 1 n 1 n n n n 4 1 2 1 3 1 1 267. "'Seelya" lloydi 2 i i n n n n n n 1 1 2 2 1 n n n n 3 3 2 1 1 1 1 268. Ulrichospira similis 2 i i n n n n n n 1 1 2 2 1 n n n n 3 3 2 1 1 1 1 269. Eocryptaulina helcinia n n n n n n n n n n 1 1 n 1 n n n n 4 2 2 1 2 1 2 270. Conotoma claustrata n n n n n n n n n n 1 2 2 1 n n n n 5 2 2 1 1 1 1 271. Crenilunula hallei n n n n n n n n n n 1 2 2 1 n n n n 5 2 2 1 3 1 1 NUMBER 88 129 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 I 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 212. Diplozone crispa ? 1 1 ? 2 1 1 3 2 1 1 7 1 7 1 1 n 1 n 1 2 2 6 1 2 213. Donaldiella declivis ? 1 3 ? 7 ? 1 2 1 ? 1 7 1 ? 1 1 n 1 n 1 1 2 6 1 2 214. Donaldiella morinensis ? 1 2 2 2 2 1 2 1 ? 1 7 1 ? 1 1 n 1 n 1 1 1 6 1 2 215. Goniostropha sculpta ? 1 1 7 1 2 1 2 1 7 2 7 1 7 1 1 n 1 n 1 1 2 6 1 2 216. Loxonema beraultensis ? 1 4 1 2 1 1 3 2 1 1 7 1 ? 1 1 n 1 n 1 2 2 6 1 2 217. Coelocaulus concirmus ? 1 4 1 1 2 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 1 2 6 1 2 218. Macrochilus buliminus 7 1 1 7 1 2 1 1 1 7 1 ? 1 7 1 1 n 1 n 1 2 2 6 1 2 219. Macrochilus cancellatus ? 1 1 ? 1 2 1 1 1 7 1 7 1 7 1 1 n 1 n 1 2 2 6 1 2 220. Macrochilina recticosta ? 1 1 ? 2 1 1 3 1 7 1 ? I 7 1 1 n I n 1 2 1 6 1 2 221. Murchisonia paradoxa 7 1 3 ? ? ? 1 2 1 7 1 ? 1 ? 1 1 n 1 n 1 1 2 6 1 2 222. Sinuspira tenera 7 1 3 2 1 2 1 2 1 ? 1 7 1 7 1 1 n 1 n 1 1 2 6 1 2 223. Stylonema mater ? 1 2 ? ? 1 1 3 1 ? 1 7 1 ? 1 1 n 1 n 2 2 2 6 1 2 224. Stylonema potens 7 1 2 ? 7 1 1 3 1 ? 1 7 1 7 1 1 n 1 n 2 1 2 7 1 2 225. Clathrospira Smithville Fm. sp. 7 1 2 1 7 2 1 1 7 ? 2 273 1 7 1 1 n 2 2 1 3 2 4 1 2 226. Clathrospira Iglindmeveri 7 1 3 1 1 1 1 1 7 ? 2 2 1 7 1 1 n 2 2 1 3 2 4 1 2 227. Clathrospira elliptica 7 1 2 1 1 2 1 1 ? 7 2 3 1 7 1 1 n 2 2 1 3 2 4 1 2 228. Clathrospira euconica 7 1 3 1 1 1 1 1 7 7 2 1 1 9 1 1 n 2 1 1 3 2 4 1 2 229. Clathrospira inflata ? 1 2 1 1 2 1 1 ? ? 2 3 1 7 1 1 n 2 2 1 3 2 4 1 2 230. Mourlonia mjoela 7 1 2 1 1 2 1 1 7 7 2 2 1 ? 1 1 n 2 2 1 3 2 5 1 2 231. Clathrospira 'Itrochiformis ? 1 2 1 1 1 1 1 7 7 2 2 1 7 1 1 n 2 2 1 3 2 5 1 2 232. Clathrospira convexa ? 1 ? ? 1 1 1 1 1 ? 2 2 2 2 1 1 n 2 1 1 4 2 5 1 2 233. Clathrospira conica ? 1 3 1 1 1 1 1 1 ? 2 2 1 ? 1 1 n 2 2 1 3 2 5 1 2 234. Clathrospira subconica ? 1 3 1 1 1 1 1 1 ? 2 2 1 ? 1 1 n 2 2 1 3 2 5 1 2 235. Eotomaria canal i/era ? 1 3 1 1 1 1 1 1 ? 2 2 1 ? 1 1 n 2 2 1 3 2 5 1 2 236. Eotomaria dryope ? 1 3 1 1 1 1 1 1 ? 2 1 1 ? 1 1 n 2 1 1 3 2 4 1 2 237. Eotomaria labrosa 7 1 3 1 1 1 1 4 1 ? 2 2 1 7 1 1 n 2 3 1 4 2 5 1 2 238. Liospira larvata ? 1 3 7 1 1 1 1 1 7 2 3 1 ? 1 1 n 2 4 1 3 2 4 1 2 239. Paraliospira mundula ? 1 4 1 I 1 1 4 1 7 2 3 1 ? 1 1 n 2 5 1 3 2 4 1 2 240. Eotomaria supracingulata ? 1 1 1 1 1 1 1 1 ? 2 3 1 ? 1 1 n 2 5 1 3 2 4 1 2 241. Liospira angustata 7 1 3 7 1 1 2 1 1 7 1 7 1 7 1 2 1 1 n 1 4 2 4 1 2 242. Liospira decipens ? 1 3 1 1 1 1 4 1 7 2 1 1 7 1 1 n 2 3 1 4 2 4 1 2 243. Liospira subconcava 7 1 3 2 1 1 2 4 1 7 2 1 1 ? 1 1 n 2 5 1 4 2 4 1 2 244. Euryzone kiari ? 1 3 ? 1 1 1 1 1 7 2 2 2 1 1 1 n 2 1 1 3 2 5 1 2 245. Eotomaria elevata ? 1 1 1 1 1 1 2 1 ? 2 2 2 1 1 1 n 2 1 1 4 2 5 1 2 246. Liospira micula ? 1 3 4 1 1 2 ? 1 ? 2 2 1 7 1 1 n 2 5 2 4 2 4 1 2 247. Liospira progne ? 1 3 4 1 1 2 ? 1 7 2 1 1 7 1 1 n 2 5 1 4 2 4 1 2 248. Paraliospira angulata 1 1 3 1 1 1 1 1 1 ? 2 3 1 7 1 2 i 1 n 1 3 2 4 1 2 249. Brachytomaria baltica ? 1 3 1 1 1 1 1 1 7 1 ? 7 7 1 1 n 1 n 1 2 2 5 1 2 250. Paraliospira aff. P angulata 1 1 4 1 1 1 1 1 1 7 2 1 1 7 1 2 i 1 n 1 3 2 4 1 2 251. Paraliospira rugata ? 1 3 1 1 1 1 4 1 ? 2 3 1 7 1 1 n 2 5 1 3 2 4 1 2 252. Eotomaria notablis ? 1 3 1 1 1 1 1 1 ? 2 1 1 ? 1 2 i 1 n 1 3 2 4 1 2 253. Lophospira kindlei ? 1 3 1 1 1 1 1 1 ? 2 2 2 1 1 1 n 2 i 1 3 2 5 1 2 254. Brachytomaria papillosa ? 1 3 1 1 1 1 1 1 7 2 ? 1 7 1 1 n 1 n 1 3 2 5 1 2 255. Brachytomaria semele ? 1 3 1 1 1 1 1 1 7 2 ? 1 7 1 1 n 1 n 1 3 2 5 1 2 256. Brachytomaria striata 7 1 3 ? 1 1 1 1 1 7 1 7 1 7 1 1 n 1 n 1 3 2 5 1 2 257. Cataschisma exquisita ? 1 1 1 1 1 1 1 1 ? 2 3 2 1 1 1 n 1 n 1 4 2 5 1 2 258. Clathrospira thraivensis 7 1 1 1 1 1 1 2 1 ? 2 2 2 1 1 1 n 2 1 1 3 2 5 1 2 259. “ Bembexia ” globosa ? 1 3 1 1 1 1 1 1 7 2 2 2 1 1 1 n 2 1 1 4 2 5 1 2 260. Eotomaria rupestris 1 1 3 1 1 1 1 1 1 7 2 1 1 ? 1 2 1 1 n 1 3 2 4 1 2 261. Crenilunula limata ? 1 2 1 1 1 1 1 1 7 2 2 2 3 1 1 n 2 3 1 3 2 4 1 2 262. Clathrospira biformis ? 1 3 ? 1 1 1 1 1 ? 2 2 2 1 1 1 n 2 2 1 2 2 5 1 2 263. Phanerotrema jugosa ? 1 4 1 1 1 1 1 1 ? 2 2 1 7 1 1 n 1 n 1 4 2 5 1 2 264. Phanerotrema lindstroemi 7 1 4 1 1 1 1 2 1 ? 2 2 1 7 1 1 n 1 n 1 4 2 4 1 2 265. Oriostoma angulifer 1 1 4 1 1 1 1 1 1 7 2 1 2 7 1 2 1 2 2 1 3 2 ? 1 2 266. Stenoloron shelvensis 7 1 4 1 1 1 1 1 1 ? 2 1 1 ? 1 2 1 2 2 1 3 2 4 1 2 267. “Seelya" lloydi 7 1 3 1 1 2 1 1 1 ? 1 7 1 7 1 1 n 1 n 1 2 2 5 1 2 268. Ulrichospira similis ? 1 3 1 1 7 ? ? 1 ? 1 ? 1 7 1 ? n ? n 1 2 2 5 1 2 269. Eocryptaulina helcinia 1 1 2 1 1 1 1 1 1 ? 2 1 1 7 1 1 n 2 i 1 3 2 4 1 2 270. Conotoma claustrata ? 1 2 1 1 1 1 1 1 ? 2 2 1 7 1 1 n 2 i 1 2 2 5 1 2 271. Crenilunula hallei ? 1 2 1 1 1 1 1 1 ? 2 2 2 3 1 1 n 2 2 1 3 2 4 1 2 130 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2—Continued. 1 1 1 1 1 1 1 1 1 1 1 i i 1 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 212. Diplozone crispa 7 1 1 1 n n n 1 n n n n 1 1 1 1 7 n 213. Donaldiella declivis 8 1 1 1 n n n 1 n n n n 1 1 1 2 2 2 214. Donaldiella morinensis 8 1 1 1 n n n 1 n n n n 1 1 1 2 7 n 215. Gonioslropha sculpla 8 1 1 1 n n n 1 n n n n 1 1 1 2 7 n 216. Loxonema beraultensis 7 1 1 1 n n n 1 n n n n 1 n 1 2 7 n 217. Coelocaulus concinnus 7 1 1 1 n T1 n 1 n n n n 1 1 1 3 7 n 2 18. Macrochilus buliminus 7 I 1 1 n n n 1 n n n n 1 n 1 2 7 n 219. Macrochilus cancellatus 7 1 1 2 4 2 i 2 4 i 1 I 1 n 1 2 7 n 220. Macrochilina recticosta 7 1 1 1 n n n 1 n n n n 1 1 1 7 7 n 22 1. Murchisonia paradoxa 8 1 1 1 n n n 1 n n n n 1 1 1 2 7 n 222. Sinuspira tenera 8 1 1 1 n n n 1 n n n n 1 1 1 2 2 2 223. Stylonema mater 7 1 1 2 3 i i I 3 i i i 1 n 1 2 7 n 224. Stylonema potens 7 1 1 2 3 i i 1 3 i i i 1 n 1 2 7 n 225. Clalhrospira Smithville Fm. sp. 6 2 ? 1 n n n 1 n n n n 1 1 1 3 7 n 226. Clathrospira Iglindmeyeri 6 2 1 1 n n n 1 n n n n 1 1 1 3 7 n 227. Clathrospira elliptica 6 2 1 1 n n n 2 2 i 1 1 1 1 1 3 7 n 228. Clathrospira euconica 6 7 1 1 n n n 1 n n n n 1 1 1 3 7 n 229. Clathrospira in/lata 6 2 1 1 n n n 1 n n n n 1 1 1 3 7 n 230. Mourlonia mjoela 6 2 1 1 n n n 1 n n n n 1 1 1 3 7 n 231. Clathrospira Itrochiformis 6 2 1 1 n n n 1 n n n n 1 1 1 3 7 n 232. Clathrosp ira con vexa 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 233. Clathrospira conica 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 234. Clathrospira subconica 6 2 1 2 3 1 i 2 4 i i i 1 2 1 2 7 n 235. Eotomaria canalifera 5 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 236. Eotomaria dryope 5 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 231. Eotomaria labrosa 5 3 1 1 n n n 1 n n n n 1 1 1 2 7 n 238. Liospira larvata 4 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 239. Paraliospira mundula 4 1 1 1 n n n I n n n n 1 1 1 1 7 n 240. Eotomaria supracingulata 4 1 1 1 n n n 1 n n n n 1 1 1 3 7 n 241. Liospira angustata 5 3 1 1 n n n 1 n n n n 1 1 1 2 7 n 242. Liospira decipens 5 3 1 1 n n n 1 n n n n 1 1 1 1 7 n 243. Liospira subconcava 5 3 1 1 n n n 1 n n n n 1 1 1 1 7 n 244. Euryzone kiari 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 245. Eotomaria elevata 6 2 1 1 n n n 1 n n n n 1 1 1 1 7 n 246. Liospira micula 5 1 1 2 3 1 1 2 4 1 1 1 1 1 1 1 1 11 247. Liospira progne 5 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 248. Paraliospira angulata 4 1 1 1 n n n 1 n n n n 1 1 1 1 7 n 249. Brachytomaria baltica 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 T1 250. Paraliospira aff. P. angulata 4 1 1 1 n n n 1 n n n n 1 1 1 3 7 n 251. Paraliospira rugata 4 1 1 1 n n n 1 n n n n 1 1 1 1 7 n 252. Eotomaria notablis 5 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 253. Lophospira kindlei 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 254. Brachytomaria papillosa 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 255. Brachytomaria semele 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 256. Brachytomaria striata 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 251. Cataschisma exquisita 6 1 1 2 4 i i 2 4 1 1 1 2 2 1 2 7 n 258. Clathrospira thraivensis 6 2 1 1 n n n 1 n n n n 1 1 I 1 7 n 259. “ Bembexia ” globosa 6 2 1 1 n n n 1 n n n n 1 1 1 2 7 n 260. Eotomaria rupestris 5 1 ? 1 n n n 1 n n n n 1 1 1 1 7 n 261. Crenilunula limata 5 2 1 2 3 3 2 2 3 3 i i 1 1 1 ? ? n 262. Clathrospira biformis ? 2 1 2 3 1 1 1 n n n n 1 1 1 2 7 n 263. Phanerotrema jugosa 6 2 1 2 3 2 1 1 n n n n 1 1 1 2 7 A 264. Phanerotrema lindstroemi 5 2 1 2 3 2 1 1 n n n n 1 1 1 4 7 n 265. Oriostoma angulifer 7 1 1 3 1 3 2 1 n n n n 1 1 1 2 ? n 266. Stenoloron shelvensis 5 1 1 1 n n n 1 n n n n 1 1 1 2 1 1 267. “Seelya" lloydi 6 1 1 2 2 2 i 2 2 2 i 1 1 1 1 7 ? n 268. Ulrichospira similis 6 1 1 1 n n n 1 n n n n 1 1 1 7 7 n 269. Eocryptaulina helcinia 5 1 1 1 n n n 1 n n n n 1 n 1 2 7 n 270. Conotoma claustrata 6 2 1 2 3 1 1 2 3 i i i 1 2 I 2 7 n 271. Crenilunula hallei 6 2 1 2 2 3 1 2 2 3 i i 1 2 1 3 ? n NUMBER 88 131 Appendix 2.—Continued. 1 1 1 1 1 i 1 1 1 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 272. Oehlertia gradata 2 3 1 5 1 1 1 2 3 3 1 1 1 n 4 1 3 1 2 4 2 2 3 1 n 273. Oehlertia scutulata 2 3 1 5 1 1 1 2 3 3 1 1 1 n 3 1 1 1 2 4 2 2 3 1 n 274. Pleurorima wisbeyensis 2 3 2 4 1 2 2 2 3 3 1 1 1 n 3 1 1 1 2 3 2 1 2 1 n 275. Promourlonia aff. P. furcata 2 3 2 4 2 2 2 2 3 3 1 1 1 n 4 1 1 1 2 2 2 2 3 1 n 276. “ Longstajfia ” “ laquetta ” 2 3 2 4 2 2 2 2 3 3 1 1 1 n 4 1 1 1 2 2 2 2 3 1 n 277. Phanerotrema loccidens 2 2 2 2 2 1 1 2 3 3 1 2 1 n 4 1 1 i 2 2 2 2 5 1 n 278. Stenoloron aequilatera 2 1 5 3 1 2 1 2 3 3 1 1 1 n 4 l 1 1 2 1 2 2 4 1 n 279. Oriostoma dispar 2 1 3 2 3 1 3 3 3 1 1 1 1 n 3 1 1 1 2 1 2 2 3 1 n 280. Murchisonia othemensis 2 2 2 2 2 2 2 2 3 3 1 1 1 n 3 1 1 1 2 3 2 2 3 1 n 281. “Seelya" Ivitellia 2 2 2 2 2 1 1 2 3 3 1 1 I n 4 1 1 1 2 2 2 2 3 1 n 282. Coelozone verna 2 3 2 4 1 1 2 2 3 3 1 1 1 n 4 1 1 1 2 5 2 1 3 1 n 283. Conotoma glandiform is 2 3 1 3 2 1 1 2 3 3 1 1 1 n 3 1 1 1 2 2 2 2 4 2 i 284. Euryzone connulastus 2 3 2 3 1 1 2 2 3 3 1 1 1 n 4 1 2 1 2 5 2 1 3 I n 285. Globispira prima 2 2 1 3 1 2 2 2 3 3 1 1 1 n 3 1 2 1 2 3 2 1 2 1 n 286. Oehlertia cancellata 2 3 1 4&5 1 1 1 2 3 3 1 1 1 n 4 1 3 1 2 3 2 2 3 1 n 287. Prosolarium procerum 2 3 1 4 1 1 1 2 3 3 1 1 1 n 4 1 1 1 2 2 2 2 5 2 2 288. Pleurorima migrans 2 3 2 3 1 1 2 2 3 3 1 1 1 n 3 1 2 1 2 5 2 1 2 1 n 289. Pleurorima aptychia 2 3 2 3 1 2 2 2 3 3 1 1 1 n 3 1 2 1 2 5 2 1 3 1 n 290. Phanerotrema dolia 2 2 3 3 2 1 1 2 3 3 1 2 1 n 4 1 1 1 2 5 2 2 3 1 n 291. Spiroraphe bohemica 2 1 5 3 1 2 3 2 3 3 1 1 1 n 4 1 1 1 2 1 2 2 4 1 n 292. Stenoloron pollens 2 1 4 2 1 2 1 2 3 3 1 1 1 n 4 1 1 1 2 1 2 2 3 1 n 293. Stenoloron voluta 2 1 3 2 3 2 3 3 3 1 1 1 1 n 3 1 1 1 2 1 2 2 3 1 n 294. Umbotropis albicans 2 2 2 2 2 1 2 2 3 3 1 1 1 n 2 1 1 1 2 2 2 2 1 1 n 295. Seelya movdartensis 2 3 2 3 1 1 2 2 3 3 1 1 1 n 3 1 2 1 2 5 2 1 2 1 n Appendix 2.—Continued. 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 5 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 272. Oehlertia gradata 1 1 n n n n 11 2 2 2 3 2 4 2 l 1 1 2 1 1 n 2 8 1 n 273. Oehlertia scutulata 1 1 n n n n n 2 2 2 3 2 4 2 1 1 1 2 1 1 n 2 9 1 n 21 A. Pleurorima wisbeyensis 1 2 5 1 i i i 1 2 2 2 1 1 1 1 1 1 2 1 1 n 1 4 1 n 275. Promourlonia aff. P. furcata 1 1 n n n n n 2 1 n n 1 1 2 l 1 1 2 1 1 n 1 A 1 n 276. “ Longstajfia ” “ laquetta'' 1 I n n n n n 2 1 n n 1 1 2 1 1 1 2 1 1 n 1 A 1 n 277. Phanerotrema loccidens 1 1 n n n n n 2 ? n n 1 1 3 1 1 1 3 1 1 n 1 9 1 n 278. Stenoloron aequilatera 1 1 n n n n n 2 2 2 2 1 1 2 1 1 1 4 1 2 i 1 5 1 n 279. Oriostoma dispar 1 1 n n n n n 1 2 2 2 1 n n 1 2 1 4 1 2 i 1 9 1 n 280. Murchisonia othemensis 1 1 n n n n n 2 2 2 2 1 i 2 1 1 1 2 1 1 n 1 9 1 n 28 1 . “ Seelya '’ Ivitellia 1 1 n n n n n n 1 n n 1 i 2 1 1 1 3 1 1 n 1 A 1 n 282. Coelozone verna 1 1 n n n n n i 2 2 2 1 i 2 1 1 1 3 1 1 n 1 5 1 n 283. Conotoma glandiformis 2 1 n n n n n i 1 n n 1 i 2 1 1 1 1 1 1 n 1 5 1 n 284. Euryzone connulastus 1 1 n n n n n i 2 2 2 1 i 1 1 1 1 3 1 1 n 1 5 1 n 285. Globispira prima I 2 5 1 1 1 1 i 2 2 2 1 i 1 1 1 1 2 1 1 n 1 6 1 li 286. Oehlertia cancellata 1 1 n n n n n i 2 2 3 2 4 2 1 1 1 2 1 1 n 2 8 1 n 287. Prosolarium procerum 2 1 n n n n n 2 2 1 2 1 2 3 1 1 1 1 1 1 n 1 1 1 n 288. Pleurorima migrans 1 2 5 1 1 1 1 1 2 2 2 1 1 1 1 1 1 2 1 1 n 1 4 1 n 289. Pleurorima aptychia 1 1 n n n n n 1 2 2 2 1 1 1 1 1 1 2 1 1 n 1 5 1 n 290. Phanerotrema dolia 1 1 n n n n n 1 ? n n 1 1 3 1 1 1 3 n 1 n 1 A 1 n 29 1 . Spiroraphe bohemica 1 1 n n n n n 2 2 2 2 1 1 2 1 1 2 4 i 2 1 1 7 1 n 292. Stenoloron pollens 1 1 n n n n n 2 2 2 2 1 1 1 1 1 1 4 i 2 1 1 6 1 n 293. Stenoloron voluta 1 1 n n n n n 1 2 2 2 1 n n 1 2 1 4 i 2 1 1 8 1 n 294. Umbotropis albicans 1 1 n n n n n 2 2 2 2 1 1 1 1 1 1 4 i 1 n 1 9 1 n 295. Seelya moydartensis 1 2 5 1 1 1 1 1 2 2 2 1 1 1 1 1 1 2 i 1 n 1 4 1 n 132 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Appendix 2.—Continued. 5 5 5 5 5 5 5 5 5 6 6 6 S 6 6 6 6 6 6 7 7 7 7 7 7 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 4 5 6 7 8 9 0 1 2 3 4 5 272. Oehlertia gradata 2 3 2 1 8 1 3 4 6 1 n n ? n n n n 1 n 1 3 1 n 1 273. Oehlertia scutulata 2 2 2 1 8 1 3 5 6 1 n n ? n n n n 1 n 1 2 I n 1 274. Pleurorima wisbeyensis 1 2 2 1 7 2 2 3 4 2 1 n 1 n n n n 1 n 1 2 1 n 1 275. Promourlonia aff. P. furcata 3 2 2 3 2 3 1 6 5 1 n n 2 3 4 2 1 1 n 1 2 1 n 2 276. “ Longstaffia ” “ laquetta" 3 2 2 3 2 3 1 6 5 1 n n 2 3 4 1 1 1 n 1 2 1 n 2 277. Phanerotrema loccidens 1 2 2 3 3 4 1 5 4 1 n n 2 2 2 1 1 I 1 1 3 1 n 1 278. Stenoloron aequilatera 1 1 2 1 7 3 3 2 1 1 n n 1 n n n n 1 n 1 2 1 n 1 279. Oriostoma dispar 2 2 2 2 6 6 2 3 3 1 n n 1 n n n n 1 n 1 2 1 n 1 280. Murchisonia othemensis 2 3 2 3 5 4 1 5 3 1 n n 1 n n n n 1 n 1 3 1 n 1 281. “ Seelya ” Ivitellia 1 2 2 3 3 6 I 5 3 1 n n 1 n n n n 1 n 1 2 1 n 1 282. Coelozone vema 1 1 2 1 7 2 2 2 2 2 3 n 1 n n n n 1 n 1 3 1 n 1 283. Conotoma glandiformis I 2 2 1 7 2 2 3 3 1 n n 1 n n n n 1 n I 2 1 n 1 284. Euryzone connulastus 1 1 2 1 7 2 3 2 4 2 3 n 1 n n n n 1 n 1 2 1 n 1 285. Globispira prima 1 2 2 1 6 2 2 4 4 2 3 n 1 n n n n 1 n 1 2 1 n 1 286. Oehlertia cancellata 2 3 2 1 4&5 2 3 5 6 1 n n ? n n n n 1 n 1 3 1 n 1 287. Prosolarium procerum 1 2 2 1 8 1 3 2 2 1 n n 1 n n n n 1 n 1 3 1 n 1 288. Pleurorima migrans 1 2 2 1 7 2 3 3 5 2 1 n 1 n n n n 1 n 1 2 1 n 1 289. Pleurorima aptychia 1 2 2 1 7 2 3 3 4 2 3 n 1 n n n n 1 n 1 2 1 n 1 290. Phanerotrema dolia 1 2 2 3 4 5 1 4&5 4 1 n n 2 3 2 i i 2 i 1 3 1 n 1 291. Spiroraphe bohemica 1 1 2 1 6 2 3 4 1 1 n n 1 n n n n 1 n 1 2 1 n 1 292. Stenoloron pollens 3 1 2 2 4 4 3 4 4 1 n n 2 2 1 1 1 1 n 1 1 I n 1 293. Stenoloron voluta 3 2 2 2 4 4 1 4 4 1 n n 2 2 3 1 1 1 n 1 1 1 n 1 294. Umbotropis albicans 2 2 2 1 5 2 1 4 2 1 n n 1 n n n n 1 n 1 2 1 n 1 295. Seelya moydartensis 1 2 2 1 7 2 3 3 5 2 i n 1 n n n n 1 n 1 2 1 n 1 Appendix 2— Continued. 7 7 7 7 8 8 8 8 8 8 8 8 8 8 9 9 9 9 9 9 9 9 9 9 1 0 Number Species 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 272. Oehlertia gradata n n n n n n n n n n 1 2 3 2 2 1 1 1 3 2 1 1 1 1 1 273. Oehlertia scutulata n n n n n n n n n 1 I 2 3 2 2 1 1 1 3 2 1 1 1 1 1 21 A. Pleurorima wisbeyensis n n n n n n n n n n 1 I n 1 n n n n 3 2 2 1 2 1 1 275. Promourlonia aff. P. furcata 2 3 n n n n n n n i 1 2 3 1 n n n n 4 2 2 1 2 1 1 276. “ Longstaffia '’ “ laquetta ” 2 3 n n n n n n n i 1 2 3 1 n n n n 3 3 2 1 1 1 1 277. Phanerotrema loccidens n n n n n n n n n n 1 3 3 I n n n n 3 3 1 1 1 1 1 278. Stenoloron aequilatera n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 3 1 1 279. Oriostoma dispar n n n n n n n n n n 1 ? ? 2 n n n n 4 2 2 1 4 1 1 280. Murchisonia othemensis n n n n n n n n n n ? 2 2 1 n n n n 3 3 2 1 1 1 1 281. “Seelya” Ivitellia n n n n n n n n n n 1 2 3 1 n n n n 3 2 2 1 1 1 1 282. Coelozone verna n n n n n n n n n n 1 2 1 1 n n n n 5 2 2 1 2 1 1 283. Conotoma glandiformis n n n n n n n n n n 1 2 2 1 n n n n 3 3 2 1 1 1 1 284. Euryzone connulastus n n n n n n n n n n 1 2 1 1 n n n n 5 2 2 1 2 1 1 285. Globispira prima n n n n n n n n n n 1 1 n I n n n n 3 2 2 1 2 1 1 286. Oehlertia cancellata n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 1 1 1 287. Prosolarium procerum n n n n n n n n n n 1 2 2 1 n n n n 6 2 2 1 4 1 1 288. Pleurorima migrans n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 289. Pleurorima aptychia n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 290. Phanerotrema dolia n n n n n n n n n n 1 1 2 1 n n n n 3 3 2 1 2 1 1 291. Spiroraphe bohemica n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 3 1 1 292. Stenoloron pollens n n n n n n n n n n 1 1 n I n n n n 5 l 2 1 3 1 1 293. Stenoloron voluta n n n n n n n n n n 1 ? ? 1 n n n n 4 2 2 1 3 1 1 294. Umbotropis albicans n n n n n n n n n n 1 1 n 1 n n n n 5 2 2 1 2 1 1 295. Seelya moydartens is n n n n n n n n n n 1 2 2 1 n n n n 3 2 2 1 2 1 1 NUMBER 88 133 Appendix 2.—Continued. 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 i 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 1 1 i 1 2 2 2 2 2 2 Number Species 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 6 7 8 9 0 1 2 3 4 5 272. Oehlerlia gradata ? 1 1 ? 1 1 l 1 1 ? 2 2 2 2 1 1 n 2 n 1 3 2 6 1 2 273. Oehlerlia scutulata ? 1 1 ? 1 1 1 1 1 ? 2 2 2 2 1 1 n 2 i 1 3 2 5 2 274. Pleurorima wisbeyensis 7 1 1 1 1 1 1 1 1 7 2 3 2 2 1 1 n 2 2 1 5 2 5 2 275. Promourlortia aff. P. furcata 7 1 3 1 1 1 1 1 1 ? 2 3 2 2 1 1 n 2 2 1 4 2 4&5 2 276. “ Longstaffia ” “ laquetta" ? 1 3 1 1 I 1 1 1 7 2 2 2 1 1 1 n 2 1 1 4 2 5 2 277. Phanerotrema loccidens 7 1 4 1 1 1 1 1 1 ? 2 2 1 ? 1 1 n 2 1 1 3 2 5 2 278. Stenoloron aequilatera ? 1 4 7 1 1 1 1 1 ? 2 0 1 ? 1 2 1 1 n 2 3 2 4 2 279. Oriostoma dispar ? 1 4 1 1 1 1 1 1 ? I 7 1 ? 1 2 2 2 i 1 3 2 3 2 280. Murchisonia othemensis ? 1 3 1 1 ? ? ? 1 ? 1 7 1 7 1 ? n ? n 1 2 2 5 2 281. “ Seelya ” Ivitellia 7 1 3 1 1 1 1 1 1 ? 2 2 2 1 1 1 n 2 i 1 4 2 5 2 282. Coelozone verna ? 1 4 1 1 1 1 1 1 ? 2 4 1 ? 1 1 n 2 3 1 5 2 3 2 283. Conotoma glandiformis 7 1 1 1 1 1 2 1 1 ? 1 ? 2 I 1 1 n 2 2 1 1 2 6 2 284. Euryzone connulastus ? 1 4 1 1 1 1 1 1 ? 2 3 1 ? 1 1 n 2 2 1 5 2 3 2 285. Globispira prima ? 1 3 1 1 1 1 1 1 ? 2 3 2 2 1 1 n 2 2 1 5 2 5 2 286. Oehlerlia cancellata 7 1 2 1 1 1 1 1 1 ? 2 3 1 7 1 1 n 2 1 1 1 2 6 2 287. Prosolarium procerum ? 1 2 1 1 1 1 1 7 ? 2 4 2 3 1 1 n 2 2 1 3 2 4 2 288. Pleurorima migrans ? 1 4 1 1 1 1 1 1 7 2 3 2 1 1 1 n 2 2 1 5 2 5 2 289. Pleurorima aptychia ? 1 4 1 1 1 1 1 1 ? 2 3 1 7 I 1 n 2 2 1 5 2 4 2 290. Phanerotrema dolia ? 1 4 1 ? 2 1 1 1 7 2 3 1 ? 1 1 n 1 n 1 4 2 5 2 291. Spiro raphe bohemica ? 1 2 7 1 1 1 1 1 ? 2 0 1 ? 1 2 1 1 n 2 3 1 4 2 292. Stenoloron pollens 7 1 2 2 1 1 1 1 1 ? 2 0 1 ? 1 2 1 1 n 2 3 2 4 2 293. Stenoloron voluta ? 1 4 1 1 1 1 1 1 7 1 ? 1 ? 1 2 1 2 2 1 3 2 4 2 294. Umbotropis albicans ? 1 4 ? 1 1 1 1 1 ? 2 1 1 ? 1 1 n 2 2 1 3 2 3 2 295. Seelya moydartensis ? 1 4 1 1 1 1 1 1 ? 2 3 2 1 1 1 n 2 2 1 5 2 5 2 Appendix 2 —Continued. 1 1 1 1 1 1 1 i 1 1 i i i 1 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 4 4 4 4 Number Species 6 7 8 9 0 I 2 3 4 5 6 7 8 9 0 1 2 3 272. Oehlerlia gradata 6 2 1 2 4 1 1 2 4 1 1 1 1 2 1 2 ? n 273. Oehlerlia scutulata 6 2 1 2 4 1 1 2 4 1 1 1 1 2 1 2 ? n 274. Pleurorima wisbeyensis 6 1 1 2 4 1 1 2 4 1 1 1 1 2 1 2 ? n 275. Promourlonia aff. f! furcata 5 3 1 1 n n n 1 n n n n 1 1 1 3 ? n 276. “Longstaffia" “laquetta" 6 2 1 1 n n n 1 n n n n 1 1 1 3 ? n 277 . Phanerotrema loccidens 6 2 1 2 3 2 i 1 n n n n 1 1 1 2 ? n 278. Stenoloron aequilatera 5 1 1 1 n n n 1 n n n n 1 n 1 3 ? n 279. Oriostoma dispar 3&4 3 1 1 n n n 1 n n n n 1 i 1 2 ? n 280. Murchisonia othemensis 6 1 1 1 n n n 1 n n n n 1 i 1 ? ? n 281. “Seelya" Ivitellia 6 2 1 1 n n n 1 n n n n 1 i 1 3 ? u 282. Coelozone verna 4 1 1 1 n n n 1 n n n n 1 i 1 3 ? n 283. Conotoma glandiformis 6 2 1 2 3 i i 2 3 1 1 1 1 2 1 2 7 n 284. Euryzone connulastus 5 1 1 1 n n n 1 n n n n 1 1 1 2 7 n 285. Globispira prima 6 1 1 2 4 1 1 2 4 i i 1 1 2 1 2 ? n 286. Oehlerlia cancellata 7 ? 1 2 4 2 1 2 4 2 i 1 1 2 1 1 ? n 287. Prosolarium procerum 6 2 1 2 3 3 1 2 n 3 i 1 2 2 1 3 ? n 288. Pleurorima migrans 6 1 1 2 4 1 1 2 4 1 i 1 1 2 1 3 ? n 289. Pleurorima aptychia 6 1 1 2 4 1 1 2 4 1 i 1 1 2 1 3 ? n 290. Phanerotrema dolia 7 1 1 2 2 3 1 1 n n n n 1 1 1 1 ? n 291. Spiroraphe bohemica 5 1 1 1 n n n 1 n n n n 1 n 1 3 1 i 292. Stenoloron pollens 5 1 1 1 n n n 1 n n n n 1 n 1 2 1 i 293. Stenoloron voluta 5 1 1 2 i i 1 1 n n n n 1 1 1 2 ? n 294. Umbotropis albicans 4 1 1 1 n n n 1 n n n n 1 1 1 1 ? n 295. Seelya moydartensis 6 1 1 2 2 3 i 2 4 i i i 1 2 1 3 7 n Appendix 3. Stratigraphic Data Species are arranged within clades or paraclades of similar species (e.g., “Hormotomoids” include all high-spired “murchisoniinae”). Stratigraphic “time” scales for each clade or paraclade precede the data giving first appearance (FKA), last appearance (LKA), lower and upper 95% confidence intervals (LB and UB), and number of sampled horizons (H). Biogeographic time scales reflect the numbers of sampling opportunities within the four main Ordovician/Silurian provinces (see Wagner, 1995a, for details). Generic names re¬ flect the names used prior to the revisions suggested by this study. Early “Archaeogastropods” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian 1-10 - - - Early Tremadoc 11-48 - - - Late Tremadoc 49-64 - - - Early Arenig 65-111 - - - Middle Arenig - - - - Finds and Ranges Species Province H FKA LB LKA UB 1. Dirhachopea normalis Laur 10 1 -4.0 10 15.0 2. Dirhachopea subrotunda Laur 8 1 -5.8 10 16.8 3. Schizopea typica Laur 13 1 -13.4 40 54.4 4. Sinuopea sweeti Laur 5 11 -33.1 40 84.1 5. Taeniospira emminencis Laur 8 1 -26.2 40 67.2 6. Ceratopea canadensis Laur 9 41 12.0 90 119.0 7. Gasconadia putilla Laur 21 11 1.0 60 70.0 8. Jarlopsis conicus Laur 16 11 -3.8 63 77.8 9. Ophileta supraplana Laur 19 11 0.0 60 71.0 10. Rhombella umbilicata Laur 5 11 -66.9 63 140.9 11. Prohelicotoma uniangulata Laur 7 11 -30.5 60 101.5 12. Sinuopea basiplanata Laur 8 11 -9.4 40 60.4 13. Taeniospira ?st. clairi Laur 3 11 -133.9 40 184.9 14. Bridgeites ^.disjuncta Laur 4 49 14.5 63 97.5 15. Bridgeitesplanodorsalis Laur 17 49 32.6 111 127.4 16. Bridgeites supraconvexa Laur 8 49 20.4 90 118.6 17. Euconia etna Laur 8 49 6.2 111 153.8 18. Ceratopea llaurentina Laur 4 65 44.3 73 93.7 19. Ceratopea pygmaea Laur 22 65 56.1 111 120.2 20. Orospira bigranosa Laur 10 65 52.0 90 103.0 134 NUMBER 88 135 i. “Euomphalinaes”: I.l. “Ophiletoids” and 1.2. “Macluritoids” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian - - - - Early Tremadoc 1-21 - - - Late Tremadoc 22-36 - - - Early Arenig 37-53 - - - Middle Arenig - - - - Late Arenig 54 54-79 55-60 - Llanvim 55-69 80-85 61 - Llandeilo 70-77 96 62-63 - Early Caradoc 78-90 100 64-65 - Middle Caradoc 91-114 - - - Late Caradoc 115-122 - 66 - Ashgill 123-125 - 67-72 - Finds and Ranges Species Province H FKA LB LKA UB 9. Ophileta supraplana Laur 19 1 -6.0 32 39.0 11. Prohelicotoma uniangulata Laur 7 1 -25.6 32 58.6 21. Macluritella stantoni Laur 2 22 -295.1 32 349.1 22. Teiichispira odenvillensis Laur 8 40 30.5 53 62.5 23. Teiichispira 1oceana Laur 7 33 15.6 53 70.4 24. Palliseria robusta Laur 4 46 51.7 54 56.3 25. Mitrospira longwelli ToqTab 5 60 21.8 85 123.2 26. Teiichispira kobayashi ToqTab 11 55 39.6 89 104.4 27. Teiichispira sylpha ToqTab 10 46 26.0 85 105.0 28. Monitorella auricula ToqTab 3 60 -65.6 85 210.6 29. Maclurites magna Laur 25 55 47.9 96 103.2 30. “ Eccyliopterus ornatus ” Laur 6 82 72.5 90 99.5 31. Maclurites bigsbyi Laur 6 82 72.5 90 99.5 32. Maclurina logani Laur 24 78 71.7 114 120.3 33. Maclurina manitobensis Laur 15 82 68.7 125 138.2 34. Maclurites sedgewicki ToqTab 3 90 56.2 96 129.8 35. Maclurites expans a Laur 1 123 undef 125 undef Maclurites expansa Balt 1 67 undef 72 undef 36. Ophileta complanata Laur 8 1 -26.9 41 68.9 37. Lecanospira compacta Laur 7 22 12.9 32 41.1 38. Lecanospira nereine Laur 4 22 -3.3 32 57.3 39. Bamesella llecanospiroides Laur 2 42 -73.3 45 160.3 Barnesella llecanospiroides ToqTab 2 55 -89.1 59 203.2 40. Malayaspira hintzei ToqTab 8 55 26.4 96 124.6 41. Malayaspira rugosa ToqTab 10 54 36.5 88 105.5 42. Barnesella measuresae ToqTab 7 60 38.4 85 106.6 43. Lytospira angelini Balt 9 55 48.0 66 73.0 44. Lytospira yochelsoni ToqTab 2 60 -1122.0 100 1282.0 45. Maclurina 2 annul at a ToqTab 5 60 21.8 85 123.2 46. Rossospira harrisae ToqTab 1 60 undef 85 undef 47. Ecculiomphalus bucklandi ToqTab 3 90 36.9 100 153.1 Ecculiomphalus bucklandi Laur 4 55 2.1 77 129.9 48. Lytospira gerrula ToqTab 1 90 undef 96 undef 49. Lytospira Inorvegica Balt 3 62 37.9 66 90.2 50. Ophiletina cf. O. sublaxa ToqTab 2 97 -18.3 100 215.3 Ophiletina cf. O. sublaxa Laur 1 123 undef 125 undef 51. Lytospira subrotunda Laur 2 102 -589.9 125 816.9 136 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY I. “Euomphalinaes”: 1.3.1 “Raphistomatids” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian - - - - Early Tremadoc - - - - Late Tremadoc - - - - Early Arenig 1 - - - Middle Arenig - 2-5 2 2 Late Arenig - 6-21 3-26 3-21 Llanvim 2-19 22-34 27-46 22 Llandeilo 20-30 25—40 47-48 23-34 Early Caradoc 31-84 - 49-66 35-40 Middle Caradoc 85-103 41-42 67-73 - Late Caradoc 104-117 41-42 74-94 - Ashgill 118-121 41-42 95-97 - Early Llandovery 122-123 - - - Late Llandovery 124-125 - - - Early Wenlock 126-132 - - - Late Wenlock 133-135 - - - Early Ludlow - - - 136 Late Ludlow - - 137 Pridoli - - - 138 Finds and Ranges Species Province H FKA LB LKA UB 52. Pararaphistoma lemoni Laur 1 1 undef 1 undef Pararaphistoma lemoni ToqTab 8 2 -20.4 34 56.4 53. Climacoraphistoma vaginati Gond 4 3 -6.2 6 15.2 Climacoraphistoma vaginati Balt 10 5 -17.0 48 70.0 54. Lesueurilla bipatellare Balt 8 2 -30.0 48 80.0 55. Lesueurilla marginalis Balt 36 2 -8.5 96 106.5 Lesueurilla marginalis ToqTab 1 36 undef 40 undef 56. Lesueurilla prima Gond 4 2 -16.4 9 27.4 57. Palaeomphalus giganteus ToqTab 11 6 -6.8 34 46.8 58. Climacoraphistoma damesi Balt 1 5 undef 16 undef 59. Eccyliopterus alatus Balt 6 5 -41.6 48 94.6 60. Eccyliopterus Iprinceps Balt 18 24 12.0 73 85.0 61 . Eccyliopterus regularis Balt 20 5 -4.2 48 57.2 62. Lesueurilla infundibula Balt 9 5 -6.0 23 34.0 63. Eccyliopterus louderbacki ToqTab 2 3 -544.8 21 568.8 64. Lesueurilla declivis Balt 5 5 -12.6 16 33.6 Lesueurilla declivis ToqTab 2 3 -544.8 21 568.8 65. Pararaphistoma qualteriata Balt 15 5 -22.0 94 121.0 Pararaphistoma qualteriata ToqTab 6 3 -37.3 40 80.3 66. Pararaphistoma schmidti Balt 8 5 -24.9 48 77.9 Pararaphistoma schmidti ToqTab 1 36 undef 40 undef 67. Helicotoma gubanovi ToqTab 1 6 undef 21 undef 68. Scalites katoi ToqTab 14 6 -3.1 40 49.1 69. Helicotoma medfraensis ToqTab 1 22 undef 34 undef 70. Lesueurilla scotica ToqTab 2 22 -352.8 34 408.8 1 1. Pachystrophia devexa Balt 22 27 13.5 97 111.1 72. Raphistoma striata Laur 29 2 -1.8 30 33.8 73. Raphistomina lapicida ToqTab 2 6 -455.3 21 482.3 Raphistomina lapicida Laur 7 11 -57.1 92 160.1 74. Scalites angulatus Laur 5 2 -40.6 30 72.6 75. Holopea insignis Laur 14 31 2.3 117 145.7 76. Eccyliopterus beloitensis Laur 5 31 -48.4 84 163.4 NUMBER 88 137 Species Province H FKA LB LKA UB 77. Holopea rotunda Laur 4 65 -22.4 102 189.4 78. Pachystrophia contigua Laur 1 31 undef 32 undef 79. Pachystrophia spiralis ToqTab 5 6 -31.1 40 77.1 80. Raphistomina aperta Laur 41 33 26.0 102 109.0 81. Raphistomina fissurata Laur 1 65 undef 84 undef 82. Eccyliopterus owenanus Laur 13 93 84.7 115 123.3 83. Holopea ampla Laur 8 32 -3.9 84 112.1 84. Holopea pyrene Laur 4 85 43.6 102 143.4 85. Holopea symmetrica Laur 5 67 29.4 94 127.6 86. Raphistoma peracuta Laur 5 85 46.4 117 130.6 87. Raphistomina rugata Laur 2 93 -743.1 121 957.1 88. Raphistoma tellerensis Laur 2 118 2.7 121 236.3 89. Sinutropis lesthetica Laur 3 118 98.7 121 140.3 90. Pachystrophia gotlandica Laur 8 122 114.5 132 139.5 91. Lytospira triquestra Laur 6 125 105.9 142 161.1 92. Euomphalus tubus Laur 1 127 undef 132 undef 93. Lytospira subuloides Gond 3 136 121.5 138 142.5 I. “Euomphalinaes”: “Helicotomids” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian - - - Early Tremadoc - - - - Late Tremadoc I - - Early Arenig 1 —44 - - Middle Arenig - - - Late Arenig - 45-48 - - Llanvim - 45-48 - Llandeilo - - 49-50 - Early Caradoc 45-69 49-59 - - Middle Caradoc 70-83 49-59 - - Late Caradoc 84-86 49-59 - Ashgill 87-96 - 51-60 - Early Llandovery 97-103 - - - Late Llandovery 104-110 - - - Early Wenlock 111-142 - - - Late Wenlock 143-158 - - - Early Ludlow 159-180 - 159 Late Ludlow 181-189 - - 160-171 Pridoli - - - 172 Finds and Ranges Species 19. Ceratopea pygmaea 94. Ceratopea unguis 95. Boucotspira aff. B.fimbriata 96. Lophonema peccatonica 97. Polehemia taneyensis 98. Walcottomafrydai 99. Helicotomaplanulata 100. Helicotoma tennesseensis 101. Ophiletina sublaxa 102. Ophiletina angularis 103. Oriostoma bromidensis Province H FKA Laur 22 2 Laur 16 1 ToqTab 8 45 Laur 12 2 Laur 15 2 ToqTab 2 45 Laur 26 45 Laur 11 45 Laur 6 45 Laur 1 70 Laur 1 45 LB LKA UB -6.2 44 52.4 -11.3 44 56.3 34.8 59 69.2 -14.8 44 60.8 -10.9 44 56.9 -70.3 48 163.3 36.7 96 104.4 34.0 69 80.0 4.7 82 122.3 undef 79 undef undef 46 undef 138 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Species Province H FKA LB LKA UB 104. Euomphalopterus lordovicius ToqTab 1 49 undef 59 undef Euomphalopterus lordovicius Laur 2 87 -201.3 96 384.3 105. Euomphalopterus aff. E. ordovicius ToqTab 1 49 undef 59 undef 106. Euomphalopterus cariniferus Balt 5 49 -145.0 180 374.0 107. Palaeomphalus Igradatus Balt 7 46 33.6 60 72.5 108. Trochomphalus Idimidiatus Balt 3 46 -26.5 60 132.5 109. Helicotoma blodgetti Laur 2 87 -201.3 96 384.3 110. Helicotoma robinsoni Laur 1 87 undef 96 undef 111. Helicotomal Girvan sp. Laur 1 87 undef 96 undef 112. Straparollina cf. S. circe Laur 3 87 38.7 96 144.3 113. Euomphalopterus alatus Laur 35 101 91.4 187 196.6 Euomphalopterus alatus Gond 5 159 122.3 183 219.8 114. Euomphalopterusfrenatus Laur 1 101 undef 103 undef 115. Euomphalopterus praetextus Laur 1 101 undef 103 undef 116. Euomphalopterus subcarinatus Laur 11 97 57.0 187 227.0 117. Euomphalopterus togatus Laur 3 101 -285.4 180 566.4 118. Euomphalopterus undulans Laur 2 101 14.5 103 189.5 119. Grantlandispira christei Laur 1 97 undef 103 undef 120. Poleumita discors Laur 34 101 91.2 189 198.8 121. Pycnomphalus acutus Laur 9 101 54.6 180 226.4 122. Pycnomphalus obesus Laur 6 101 16.2 180 264.8 Pycnomphalus obesus Gond 3 159 101.0 170 228.0 123. Discordichilus dalli Laur 2 107 -1392.2 158 1657.2 124. Discordichilus mollis Laur 5 127 47.6 180 259.4 125. Discordichilus kolmodini Laur 2 127 -334.3 142 603.3 126. Poleumita alata Laur 3 104 -161.7 158 423.7 127. Poleumita octavia Laur 7 128 81.5 183 229.5 128. Poleumita rugosa Laur 24 107 94.4 180 192.9 129. Pseudophorus profundus Laur 1 132 undef 142 undef 130. Pseudophorus stuxbergi Laur 4 107 -10.3 157 274.3 131. Siluriphorus gotlandicus Laur 10 104 84.5 142 161.5 132. Siluriphorus undulans Laur 4 132 72.2 157 216.8 133. Streptotrochus incisus Laur 6 107 68.8 142 180.2 134. Streptotrochus aff. S. incisus Laur 4 107 24.2 142 224.8 135. Streptotrochus lamellosus Laur 2 112 -781.7 142 1035.7 136. Streptotrochus lundgreni Laur 3 112 -37.7 142 291.7 137. Streptotrochus ? visbeyensis Laur 4 111 37.4 142 215.6 138. Hystricoceras astraciformis Laur 2 143 -289.5 157 589.5 139. Poleumita granulosa Laur 5 143 126.8 153 169.2 140. Euomphalus walmstedti Laur 3 143 -40.5 180 363.5 141. Centrifugus planorbis Laur 11 159 149.3 180 189.7 142. Spinicharybdis wilsoni Laur 1 181 undef 189 undef 143. Turbocheilus immaturum Gond 4 158 128.1 170 199.9 144. Pseudotectus comes Gond 1 159 -244.6 170 575.6 145. Straparollus bohemicus Gond 3 159 101.0 170 228.0 NUMBER 88 139 II. Murchisoniinaes”: II.3. “Hormotomoids” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian - - - — Early Tremadoc 1-3 - - - Late Tremadoc 4-12 - - — Early Arenig 13-87 - - - Middle Arenig - - - - Late Arenig 88-89 88-110 - - Llanvim 90 111-122 - - Llandeilo 91 123-126 — Early Caradoc 92-134 127-128 — — Middle Caradoc 135-216 127-128 — — Late Caradoc 217-295 127-128 288-290 - Ashgill 296-308 - 291-300 - Llandovery 309-324 - - - Late Llandovery 325-330 - - - Early Wenlock 331-338 - - - Late Wenlock 339-344 - - — Early Ludlow 345-360 - - 345 Late Ludlow 361-364 - - 346-357 Pridoli 365-367 - - 358-359 Finds and Ranges Species 13. Taemospira 1st. clairi 146. Hormotoma artemesia 147. Hormotoma confusa 148. Hormotoma Idubia 149. Hormotoma Isimulatrix 150. Ectomaria adelina 151. “ Hormotoma ” “ cassina ” 152. Fusispira Smithville Fm. sp. 153. Hormotoma augustina Hormotoma augustina 154. Hormotoma zelleri 155. Lophospira perangulata 156. Subulitid El Paso Fm. sp. 157. Pagodospira cicelia Pagodospira cicelia 158. Plethospira cannonensis 159. Plethospira cassina 160. Seelya ventricosa 161. Lophospira grandis 162. Straparollina pelagica 163. Plethospira ? turgida 164. Turritoma acrea 165. Turritoma Cotter Fm. ornate sp 166. Turritoma cf. T. acrea 167. Hormotoma Setul Fm. sp. 168. Turritoma ?anna 169. Murchisonia callahanensis 170. Ectomaria prisca 171. Hormotoma gracilis Hormotoma gracilis 172. Daidia cerithioides 173. Ectomaria pagoda Province H FKA Laur 3 1 Laur 15 4 Laur 8 4 Laur 3 4 Laur 37 4 Laur 6 13 Laur 8 36 Laur 5 36 Laur 7 30 ToqTab 20 91 Laur 3 20 Laur 85 36 Laur 2 36 Laur 7 36 ToqTab 7 91 Laur 6 16 Laur 4 20 Laur 1 20 Laur 23 16 Laur 7 13 Laur 5 20 Laur 1 13 Laur 2 13 Laur 3 36 ToqTab 6 91 ToqTab 7 91 ToqTab 4 88 Laur 11 92 Laur 80 91 Balt 3 288 Laur 4 93 Laur 11 93 LB LKA UB -13.5 3 17.5 -21.2 87 112.2 -53.1 87 144.1 -121.6 29 154.6 -5.2 87 96.2 -107.8 126 246.8 0.6 87 122.4 -40.4 87 163.4 -18.1 87 135.1 84.3 122 128.7 -28.3 29 77.3 25.1 308 318.1 -1463.2 87 1586.2 -7.2 87 130.2 61.1 126 155.9 -60.3 87 163.3 -136.4 87 243.4 undef 29 undef -37.5 312 368.8 -51.7 90 154.7 -80.0 87 187.0 undef 15 undef -188.8 19 220.8 -215.2 87 338.2 57.1 122 155.9 64.4 122 148.6 -6.3 128 222.3 -3.5 308 403.5 80.1 308 319.3 -923.3 300 1314.3 -102.5 177 372.5 -2.0 308 403.0 140 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Species Province H FKA LB LKA UB 174. Haplospira 1 nereis Laur 8 93 35.2 177 234.8 175. Hormotoma bellicincta Laur 57 93 75.7 308 325.3 176. Hormotoma salteri Laur 36 93 69.2 308 331.8 177. Hormotoma trentonensis Laur 93 93 85.2 287 295.0 178. Loxonema murrvana Laur 6 93 2.9 177 267.1 179. Omospira alexandra Laur 9 93 43.7 177 226.3 180. Omospira laticincta Laur 1 93 undef 134 undef 181. Straparollina circe Laur 3 93 -317.6 177 587.6 182. Straparollina erigione Laur 2 135 89.4 177 222.6 183. Gin’ania excavata Laur 1 288 undef 295 undef 184. Murchisonia Pt. Clarence Fm. sp. Laur 2 288 -317.4 308 913.4 185. Rhabdostropha primitiva Laur 4 288 239.7 308 356.3 186. Spiro ecus gir\>anensis Laur 2 288 -317.4 308 913.4 187. Daidia aff. D. cerithioides Laur 1 298 undef 308 undef 188. Ectomaria cf. E. pagoda Laur 2 298 244.9 308 361.1 189. Ectomaria cf. E. prisca Laur 3 298 -19.1 308 625.1 190. Ectomaria laticarinata Laur 1 298 undef 308 undef 191. Ectomaria nieszkowskii Balt 3 291 242.7 300 348.3 192. Hormotoma ins ignis Balt 8 288 281.2 297 303.8 193. Holopella regularis Laur 4 300 159.7 360 500.3 194. Hormotoma cenlervillensis Laur 2 309 251.3 310 367.7 195. Hormotoma cingulata Laur 6 313 262.1 360 410.9 Hormotoma cingulata Gond I 300 undef 312 undef 196. Kjerulfonema cancel lata Laur 8 313 294.4 323 362.6 197. Kjerulfonema quinquecincta Laur 2 311 -63.8 323 697.8 198. Cyrtostropha coralli Laur 2 328 -623.4 360 1311.4 199. Goniostropha cava Laur 7 327 312.1 344 358.9 200. Hormotoma subplicata Laur 2 325 36.7 334 622.3 201. Hormotoma monoliniformis Laur 2 327 -19.0 338 684.0 202. Hormotoma attenuata Laur 10 325 307.0 360 378.0 203. Loxonema ? attenuata Laur 1 330 undef 338 undef 204. Macrochilusfenestratus Laur 1 330 undef 338 undef 205. Rhabdostropha grindrodii Laur 2 328 -623.4 360 1311.4 206. Loxonema crossmanni Laur 1 328 undef 332 undef 207. Loxonema sinuosa Laur 4 339 272.3 367 433.7 208. Auriptygmafortior Gond 4 346 270.1 357 341.9 209. Catazone allevata Gond 4 346 270.1 357 341.9 210. Catazone argo/is Gond 2 346 -74.8 357 686.8 211. Catazone cunea Gond 2 346 -74.8 357 686.8 212. Diplozone crispa Laur 3 345 267.7 360 437.3 213. Donaldiella declivis Gond 6 346 286.2 312 325.8 214. Donaldiella morinensis Laur 2 345 -116.3 357 821.3 215. Goniostropha sculpta Gond 2 346 -74.8 357 686.8 216. Loxonema beraultensis Gond 3 346 237.2 357 374.8 217. Coelocaulus concinnus Gond 1 346 undef 358 undef 218. Macrochilus buliminus Laur 2 345 -116.3 360 821.3 219. Macrochilus cancellatus Laur 3 345 267.7 360 437.3 Macrochilus cancellatus Gond 1 346 undef 357 undef 220. Macrochilina recticosta Gond 3 346 232.4 359 380.6 221. Murchisonia paradoxa Laur 1 345 undef 360 undef 222. Sinuspira tenera Gond 4 346 270.1 357 341.9 223. Stylonema mater Gond 1 346 undef 357 undef 224. Stylonema potens Gond 3 346 237.2 359 374.8 NUMBER 88 141 II. “Murchisoniinaes”: II.4. “Eotomarioids” “Time” Scales Stage/Province Laurentia Toquima- Table Head Baltica Gondwana Dolgellian - - — — Early Tremadoc - - - — Late Tremadoc - - _ Early Arenig 1-2 - - Middle Arenig 3-4 - — Late Arenig 5 5-8 8-17 — Llanvim 6 9-14 18-33 — Llandeilo 7 15-21 34 — Early Caradoc 8-61 21-28 35-59 — Middle Caradoc 62-126 21-28 60-66 - Late Caradoc 127-174 21-28 67-71 — Ashgill 175-192 - 72-83 - Early Llandovery 193-212 - - - Late Llandovery 213-219 - - Early Wenlock 220-241 - - Late Wenlock 242-248 - - Early Ludlow 249-263 - - 249-256 Late Ludlow 264-272 - - 257-273 Pridoli 273-275 - - 274 Finds and Ranges Species 225. Clathrospira Smithville Fm. 226. Clathrospira Iglindmeyeri 227. Clathrospira elliptica 228. Clathrospira euconica 229. Clathrospira injlata 230. Mourlonia mjoela 231. Clathrospira 'hrochiformis 232. Clathrospira convexa 233. Clathrospira conica 234. Clathrospira suhconica 235. Eotomaria canalifera 236. Eotomaria dryope 237. Eotomaria labrosa 238. Liospira lar\>ata 239. Paraliospira murtdula 240. Eotomaria supracingulata 241. Liospira angustata 242. Liospira decipens 243. Liospira subconcava 244. Euryzone kiari 245. Eotomaria elevata 246. Liospira micula 247. Liospira progne 248. Paraliospira angulata 249. Brachytomaria baltica 250. Paraliospira aff. P. angulata 25 1. Paraliospira rugata 252. Eotomaria notablis 253. Lophospira kindlei Lophospira kindlei 254. Brachytomaria papillosa Brachytomaria papillosa 255. Brachytomaria semele Province H FKA Laur 8 1 ToqTab 3 5 Balt 45 5 Laur 1 5 Balt 24 12 Balt 9 11 Laur 3 7 Laur 4 20 Laur 17 8 Laur 30 8 Laur 4 8 Laur 15 8 Laur 1 8 Laur 11 10 Laur 7 8 Laur 11 20 Laur 4 20 Laur 12 8 Laur 19 10 Balt 7 60 Laur 1 62 Laur 35 102 Laur 76 25 Laur 6 25 Balt 3 69 Laur 2 177 Laur 3 175 Balt 16 35 Laur 1 177 Balt 1 82 Laur 2 177 Balt 1 72 Balt 4 72 LB LKA UB -58.2 87 146.2 0.0 15 0.0 -1.0 71 77.0 undef 6 undef 4.9 53 60.2 -24.4 71 106.4 -89.6 26 122.6 -377.9 192 589.9 -40.1 192 240.1 -3.1 92 103.1 -111.6 59 178.6 -22.6 109 139.6 undef 9 undef -34.0 109 153.0 -83.3 117 208.3 -19.6 109 148.6 -187.0 109 316.0 -31.8 109 148.8 -12.0 109 131.0 41.7 81 99.3 undef 126 undef 92.0 192 202.0 16.0 174 183.4 -153.1 192 370.1 6.2 81 143.8 -284.3 192 653.3 146.0 180 209.0 15.9 81 101.1 undef 192 undef undef 83 undef -284.3 192 653.3 undef 81 undef -169.5 176 417.5 142 SMITHSONIAN CONTRIBUTIONS TO PALEOBIOLOGY Species Province H FKA LB LKA UB Brachytomaria semele Laur 2 177 61.7 180 295.3 256. Brachytomaria striata Laur 1 177 undef 192 undef 257. Cataschisma exquisita Laur 3 177 -151.4 244 572.4 258. Clathrospira thraivensis Laur 2 177 -284.3 192 653.3 259. “Bembexia” globosa Laur 1 177 undef 192 undef 260. Eotomaria rupestris Balt 3 72 14.0 83 141.0 261. Crenilunula limata Laur 19 210 198.1 263 274.9 262. Clathrospira biformis Laur 3 210 41.0 244 413.1 263. Phanerotrema jugosa Laur 5 193 168.0 209 234.0 264. Phanerotrema lindstroemi Laur 6 210 176.1 263 274.9 265. Oriostoma angulifer Laur 4 210 163.0 241 288.0 266. Stenoloron shelvensis Laur 9 199 174.1 241 265.9 261. “ Seelya ” lloydi Gond 17 210 193.1 274 290.9 268. Ulrichospira similis Laur 2 199 -118.1 209 526.1 269. Eocryptaulina helcinia Laur 2 210 -799.1 244 1253.1 270. Conotoma claustrata Laur 5 213 170.4 241 283.6 271. Crenilunula hallei Laur 11 213 190.6 263 285.4 Crenilunula hallei Gond 7 257 242.1 274 288.9 272. Oehlertia gradata Laur 6 213 158.9 263 317.1 273. Oehlertia scutulata Laur 4 216 156.2 241 300.8 274. Pleurorima wisbeyensis Laur 1 229 undef 241 undef 275. Promourlonia aff. P. furcata Laur 5 213 124.8 272 360.2 276. “ Longstaffia ” “ laquetta ” Laur 1 213 undef 215 undef 277. Phanerotrema loccidens Laur 6 227 187.8 263 302.2 Phanerotrema loccidens Gond 2 227 -205.5 241 673.5 278. Stenoloron aequilatera Laur 9 219 201.6 248 265.4 279. Oriostoma dispar Laur 1 229 undef 241 undef 280. Murchisonia othemensis Laur 1 227 undef 241 undef 281. “Seelya” Ivitellia Laur 2 242 -392.3 263 897.3 282. Coelozone vema Gond 3 257 170.1 274 360.9 283. Conotoma glandiformis Laur 2 249 -183.5 263 695.5 284. Eurvzone connulastus Gond 4 257 215.6 274 315.4 285. Globispira prima Gond 3 250 129.3 274 394.8 286. Oehlertia cancellata Laur 3 249 176.6 263 335.5 287. Prosolarium procerum Gond 1 257 undef 274 undef Prosolarium procerum Laur 1 273 undef 275 undef 288. Pleurorima migrans Gond 16 250 243.0 274 281.0 289. 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