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Gelidiales (rhodophyta, red algae) in British Columbia and Northern Washington : taxonomy, morphology,… Renfrew, Dawn Elizabeth 1988

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GELIDIALES  (RHODOPHYTA,  NORTHERN WASHINGTON:  R E D ALGAE) IN BRITISH COLUMBIA A N D TAXONOMY, MORPHOLOGY,  AND LIFE  DEVELOPMENT  HISTORY  by DAWN ELIZABETH RENFREW MSc, Acadia University, 1983 BSc, The University of Toronto, 1980 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY  in  THE FACULTY OF GRADUATE STUDIES Botany We accept this thesis as conforming to the required standard  THE UNIVERSITY OF BRITISH COLUMBIA August 1988 c  Dawn Elizabeth Renfrew, 1988  In  presenting this  degree at the  thesis  in  University of  partial  fulfilment  of  of  department  this thesis for or  by  his  or  requirements  British Columbia, I agree that the  freely available for reference and study. 1 further copying  the  representatives.  an advanced  Library shall make  it  agree that permission for extensive  scholarly purposes may be her  for  It  is  granted  by the  understood  that  head of copying  my or  publication of this thesis for financial gain shall not be allowed without my written permission.  Department  of  P)(  The University of British Columbia Vancouver, Canada  DE-6 (2/88)  ABSTRACT  A  study  of the  red  algal  order  Gelidiales  in  British  Washington was conducted. Gelidiales are represented  Gelidium coulteri,  species,  caloglossoides,  Columbia  in the  G. purpurascens,  and  northern  study area by four  G. vagum  and a key to identify the taxa is provided. Earlier reports of  crinale, G. pusillum, G. robustum  G. sinicola  and  from  British  northern Washington have been shown to be misidentifications of the other and  are  excluded from  Columbian flora  Pterocladia  and  the  flora.  The occurrence  is significant because  of  G. vagum  it appears to have  been  Columbia  and  taxa  in the  introduced  G.  British from  the northwest Pacific and now has a disjunct distribution, occurring in Japan and China  and on two islands in the  vagum sister  and  the  Gulf  of California  species. They share the  genus)  and  including  a  Strait of Georgia, British species,  unique character  morphological resemblance,  arrangement  of  G.  cortical  but  cells,  johnstonii,  of monoecy  differ  shape  and  Gelidium  Columbia. are  considered  tetrasporangial  cortex, presence/absence stichidia  and  be  (i.e. unique in the  in several other flattening  of  characters,  tetrasporangial  stichidia, shape of stichidial apices and position of the apical initial with to the adjacent  to  respect  of an apical furrow and sterile margin in  presence/absence  of intact  third  order  filaments  in  mature cystocarps.  The life  history of  Gelidium triphasic and  species  Gelidium vagum for which  Polysiphonia-type  tetrasporophytes,  and  this has  was  been  completed in culture, only accomplished.  life history with  carposporophytes  ii  the  second  Gelidium vagum  isomorphic (monoecious)  growing on gametophytes.  has  gametophytes The  rare  a  occurrence  (G. purpurascens, G. coulteri)  or absence  (Pterocladia caloglossoid.  gametophytes in the field and the lack of successful completion of a life history in culture suggests that, whereas these species may  Polysiphonia-type  life history  in situ,  occasionally complete a  vegetative or apomictic mechanisms are more  important in maintaining field populations. Chromosome counts of n= 14-15 were made on undivided tetrasporangia of  G. vagum.  The pattern of spore germination in  vagum  Gelidium coulteri, G. purpurascens  and  is similar for carpospores and tetraspores. A single germ tube grows from  the spore, the entire cytoplasmic contents of the spore evacuate into the germ tube and a wall cuts the germ tube off from the empty spore. The germ tube divides unequally to form a concave and a fusiform cell. The primary attachment rhizoid forms from a derivative of the concave cell and the sporeling apical cell forms from a fusion cell derivative. As apical organization is established, the subapical cell produces two lateral periaxial cells.  Carpogonium  and  purpurascens  carposporophj'te and  development  G. vagum. The  was  functional  followed carpogonium  in  Gelidium  is intercalary.  Non-functional carpogonia divide transversely with a concave wall and become sessile and degenerate. The fertilized carpogonium consistenty forms a fusion cell by fusing with adjacent cortical cells. Fusion cell lobes  cut off gonimoblast  filaments that send out processes that fuse with haploid gametophyte cells of nutritive  chains.  Carposporangia  are produced  terminally  and  laterally  by  gonimoblast cells and protrude into two locules that have formed as the cortex is pushed away from the plate of second order filaments. Carpospores are released iii  G.  through a single ostiole in each locule.  Spermatogenesis,  tetrasporogenesis  and  rhizine  morphology  were  studied  ultrastructurally for the first time in Gelidiales. In spermatangial development, apical chloroplasts in the spermatangial mother cell are cut off by a furrow, a mechanism previously not reported in red algae for exclusion of chloroplasts from spermatangia. As the spermatangium matures, spermatangial vacuoles form from the  coalescence  of  vesiculated  endoplasmic  reticulum  and  dictyosome-derived  vesicles. Spermatangial vacuoles are discharged from the spermatangium just prior to the release of the spermatium.  Stages of tetrasporangium development are marked b}' wall development. Prior to karyokinesis, the  young  tetrasporangium  by  is dominated  Following karyokinesis, chloroplasts, starch grains and abundance, a wall layer is deposited, and  a  large nucleus.  dictyosomes  dictyosomes  increase in  undergo a series of  morphological changes from small and flat, to large and flat producing fibrillar vesicles, to hemispherical and producing cored vesicles. At cytokinesis an electron dense wall layer is deposited around the tetrasporangium and in the cleavage furrow, and cored vesicles secrete a mucilage-like material inside this wall layer.  iv  TABLE OF CONTENTS Abstract  ii  Table of Contents  v  List of Figures  ~  x  1  1  List of Tables  V  Acknowledgements  1  xii  Chapter I. Introduction A . Taxonomic History B. Characterization of Gelidiales C.  1 1 5  Differentiating Gelidium and Pterocladia  ....  D. Need for Additional Information Chapter II. Materials  7  9 10  Chapter III. Taxonomy and Nomenclature A . Introduction B. Methods C. Taxa Present in British Columbia and Northern Washington 1. Gelidium coulteri Harvey  a. b. c. d. e.  Habitat and Habit Vegetative Anatomy Reproductive Anatomy Discussion Seasonality and Distribution  2. Gelidium purpurascens  a. b. c. d. e.  Gardner  Habitat and Habit Vegetative Anatomy Reproductive Anatomy Discussion Seasonality and Distribution  12 12 15 17 18  18 19 20 22 24 25  26 27 30 34 37  3. Gelidium vagum Okamura  38  a. Habitat and Habit b. Vegetative Anatonvy c. Reproductive Anatomy d. Discussion e. Seasonality and Distribution 4. Pterocladia caloglossoides (Howe) Dawson a. Habitat and Habit b. Vegetative Anatom}' c. Reproductive Anatonvy d. Discussion e. Seasonality and Distribution  39 40 42 46 52 53 53 55 56 57 62  v  D. Previously Reported Taxa Excluded from the Revised Flora 63 1. Gelidium robustum (Gardner) Hollenberg et Abbott . 64 2. Gelidium crinale (Turner) Lamouroux 67 3. Gelidium pusillum (Stackhouse) Le Jolis 69 4. Gelidium sinicola Gardner  70  Chapter IV. Life History of Gelidium in Culture A. Introduction B. Methods 1. Culturing 2. Chromosome Counts C. Spore Germination and Early Development D. Reproduction in Cultured and Field Plants 1. Culture Results 2. Field Observations of Reproduction E. Discussion 1. Spore Germination 2. Life Histories 3. Chromosome Counts  73 73 75 75 78 78 80 80 84 85 85 89 91  Chapter V. Carpogonium and Carposporophyte Development A. Introduction B. Methods C. Carpogonium Development in Gelidium D. Carposporophyte and Cystocarp Development E. Discussion Chapter VI. Spermatiogenesis in Gelidium A. Introduction B. Methods C. Spermatiogenesis D. Comparison of Spermatiogenesis in Gelidium Florideophycean Genera Chapter VII. Ultrastructure of Tetrasporogenesis in Gelidium A. Introduction B. Methods C. Ultrastructural Observations 1. Pre-meiotic Tetrasporangium 2. Post-meiotic Tetrasporangium 3. Tetrasporangium at Cytokinesis D. Discussion  93 93 96 96 98 100  and Other  105 105 107 108 112 122 122 123 123 124 126 127 128  Chapter VIII. General Discussion and Conclusion  137  References  142  Figures  163 vi  Tables  262  Appendix 1. Herbarium specimens examined  275  Appendix 2. Procedure for Embedding Material in JB4 Methacrylate  288  Appendix 3. Preparation of Tetrasporangial and Spermatangial Material for Transmission Electron Microscopy  290  vii  List of Tables  Table 1. Historical summary of genera included in Gelidiaceae  262  Table 2. Sites and dates of field collections of Gelidiales from the study area  264  Table 3. Herbaria from which loans were obtained and their abbreviations .... 267 Table 4. Species of Gelidiales recorded from the northeast, northwest and southeast Pacific Ocean areas  268  Table 5. Differences between Gelidium purpurascens and Gelidium  robustum  .... 273  Table 6. Anatomical differences between Gelidium johnstonii and Gelidium  vagum  274  viii  List of Figures  Abbreviations used in figures  163  Fig. 1. Map of study area  165  Fig. 2. Barkley Sound collection sites  167  Fig. 3. Southern Vancouver Island, Gulf Islands and San Juan Islands collection sites  169  Fig. 4. Strait of Georgia collection sites  171  Fig. 5. Key to Gelidiales in British Columbia and northern Washington  173  Figs. 6-11. Gelidium coulteri vegetative morphology  175  Figs. 12-17. Gelidium coulteri tetrasporophytes  177  Figs. 18-20. Gelidium coulteri male gametophytes  179  Figs. 21-24. Gelidium coulteri habit  181  Figs. 25-29. Gelidium purpurascens habit and vegetative morphology  183  Figs. 30-34. Gelidium purpurascens vegetative morphology  185  Figs. 35-40. Ultrastructure of rhizines in Gelidium purpurascens and G. vagum 187 Figs. 41-44. Gelidium purpurascens tetrasporoph3'tes  189  Figs. 45-48. Gelidium purpurascens female gametophytes  191  Figs. 49-51. Gelidium purpurascens carposporophytes  193  Figs. 52-56. Gelidium purpurascens male gametophytes  195  Figs. 57-63. Gelidium purpurascens apical and lateral initials, historical and type collections  197  Figs. 64-68. Gelidium vagum habit and vegetative morphology  199  Figs. 69-76. Gelidium vagum vegetative and tetrasporophyte morphology  201  Figs. 77-83. Gelidium vagum reproductive morphology  203  Figs. 84-89. Gelidium vagum gametophytes  205  x Figs. 90-95. Gelidium vagum reproductive' i morphology  207  Figs. 96-100. Gelidium vagum Japanese material  209  Figs. 101-106. Gelidium johnstonii habit, vegetative and reproductive morphology 211 Figs. 107-111 Gelidium johnstonii reproductive morphology and G. vagum habit  213  Figs. 112-118. Pterocladia caloglossoides habit and vegetative morphology  215  Figs. 119-121. Pterocladia caloglossoides vegetative morphology  217  Figs. 122-128. Pterocladia caloglossoides reproductive morphology  219  Figs. 129-133. Pterocladia caloglossoides reproductive morphology  221  Figs. 135-140. Historical collections and type specimens  223  Fig. 141. Polysiphonia-type life history  225  Figs. 142-153. Sporeling development of Gelidium vagum in culture  227  Figs. 154-162. Sporeling development of Gelidium vagum in culture  229  Figs. 163-168. Reproduction of Gelidium vagum in culture  231  Figs. 169-173. Reproduction of Gelidium purpurascens and G. coulteri in culture 233 Figs. 174-177. Chromosomes of Gelidium vagum  235  Figs. 178-183. Carpogonium and carposporophyte development in Gelidium purpurascens and G. vagum  237  Figs. 184-188. Carposporophyte development in Gelidium purpurascens and G. vagum  239  Figs. 189-192. Ultrastructure of spermatiogenesis in Gelidium vagum  241  Figs. 193-196. Ultrastructure of spermatiogenesis in Gelidium vagum  243  Figs. 197-200. Ultrastructure of spermatiogenesis in Gelidium purpurascens and  G. vagum  245  Figs. 201-204. Ultrastructure of spermatiogenesis in Gelidium purpurascens and  G. vagum  247  Figs. 205-208. Ultrastructure of spermatiogenesis in Gelidium purpurascens and  G. vagum  249  Fig. 209. Summary of spermatiogenesis in Gelidium  251  Figs. 210-215. Ultrastructure of tetrasporogenesis in Gelidium purpurascens  253  Figs. 216-220. Ultrastructure of tetrasporogenesis in Gelidium purpurascens  255  Figs. 221-226. Ultrastructure of tetrasporogenesis in Gelidium purpurascens  257  Figs. 227-232. Ultrastructure of tetrasporogenesis in Gelidium coulteri and G. purpurascens  259  Acknowledgements  I have received am  very  assistance and suppport from  grateful.  Many  concern,  thanks to my Dr.  and  project,  to Dr. R. F . Scagel who kindly took over  another  position  to  am  grateful  Dr.  to Ira  G. I.  collections  for  who  their  got  generous all-round  me  when  to whom I  interested  Dr. Garbary  gave support from  in  this  left  for  beginning  to  to Dr. P. W. Gabrielson who always showed interest  much of his time for discussion, advice and as my diving buddy. I Borgmann,  Hawkes, Cindy Durance to  Garbary  Dr. K . M . Cole who  end. I am deeply indebted and offered  D. J .  supervisors  support  and  to  a number of people,  and  and  collected  field  who  very  important  gave much appreciated  Dr. B. R. Oates, Dr. M . W.  Gary Kendrick who also were diving buddies,  Hansen  were  Dr. W. Wheeler,  to  material  my  work.  intertidally  for  Dr. B . Hunt  me.  and  These  Don DeReussi  hospitality and access to dive sites at Whalebone Bay and  Georgina Point. I thank Dr. J . P. van der Meer, Dr. P. C. Silva, Dr. M . H . Hommersand  and  Dr.  I.  Akatsuka  for  valuable  discussions.  I  appreciate  the  assistance of Dr. V . L. Anderson (at A H F H ) , Dr. S. Honey (at BM), Dr. P. B . Hamilton (at C A N A), Dr. G. J . Cacavio (at F); Dr. D. Duggins (at  F H L ) , Dr.  M . J . Wynne (at MICH), Dr. J . Tanaka (at TNS), Julie Oliveira (at U B C ) , Dr. P. C. Silva (at UC), Dr. J . N . Norris (at US) and Dr. M . F. Denton (at WTU) for  sending  herbarium  loans  or  looking  for  possible  Pterocladia caloglossoides  isot3pes. Dr. J . W. Waaland and Dr. M . J . Wynne graciously loaned r  from their private collections.  xii  specimens  Funding for this research was provided by the National Science and Engineering Research Council of Canada grants to Dr. Cole (0645), Dr. Scagel (A4471) and Dr. Garbary (U0014), an H. R. Macmillan Family Scholarship, a Kit Malkin Scholarship and a British Columbia Post Secondary Scholarship.  To all those acknowledged here, to Frank and Dancer who helped me through the hardest part and to my friends who helped keep it all in perspective, thank you — you are greatly appreciated.  xiii  C H A P T E R I. INTRODUCTION  A.  TAXONOMIC  Gelidiales  Kylin  HISTORY  (1923:  132)  currently  encompass  the  single  family  Gelidiaceae  Kutzing (1843: 390) which contains 11 genera: Acanthopeltis Okamura (in Yatabe, 1892:  157), Beckerella Kylin  529),  Gelidium  (1956:  Lamouroux  Porphyroglossum  Kutzing  139),  (1813:  (1847a:  Gelidiella Feldmann et Hamel (1934:  128),  775),  Onikusa  Pterocladia  J.  Akatsuka G. Agardh  Pterocladiastrum Akatsuka (1986c: 55), Ptilophora Kutzing  (1986a:  63),  (1852:  482),  (1847b: 25), Suhria J .  G. Agardh ex Endlicher (1843: 41) and Yatabella Okamura (1900: 1). Historically, a heterogeneous assemblage to the  family  (Kutzing,  of genera  (now included in four orders) was ascribed  1849; Schmitz, 1889). A historical summary  of genera  included in Gelidiaceae in seven major classifications is presented in Table 1.  The family vide Silva  Gelidiaceae was erected 1980: 83) to encompass  fibrous  internal  cruciate  tetrasporangia  comprised  structure,  Acrocarpus  by Kutzing  (1843: 390, 405 as "Gelidieae";  cartilaginous, pinnately branched  exserted  cystocarps,  on distinct tetrasporangial Kutzing  (1843:  405),  small  globose  branches. Ctenodus  plants,  with  spermatia  and  The family Kutzing  originally  (1843:  407),  Echinocaulon Kutzing (1843: 405) and Gelidium, with the later addition (Kutzing, 1849)  of  Polycladia  Montagne  (1847:  (1846:  25), Delisea Lamouroux (1819:  378),  Thysanocladia  (Endlicher)  41), Chondrodon Kutzing  (1847b:  Lindley 5) and  Phacelocarpus Endlicher et Diesing (1845: 289), and the substitution of Euctenodus Kutzing  (1847b:  5) for Ctenodus. J .  G. Agardh's  1  (1851)  concept  of Gelidiaceae  Introduction / 2 (as Gelidieae) was narrower, including only the genera Gelidium [composed of the subgroups  Acrocarpus  ( — Gelidium  in  Gelidium], Pterocladia (newly erected), belonging to the family.  part),  Echinocaulon  Suhria and Ptilophora,  Wurdemannia  Harvey  (1853:  ( = Gelidiella)  all genera  and  currently  245) was added  (with a  query) because of a similarity in thallus construction to Gelidium (J. G. Agardh, 1876).  Schmitz  construction Gelidiaceae  and  devised  cystocarp  14 genera  heterogeneous, Binderella  (1889)  a  classification  construction  in five tribes.  and all genera  Schmitz (in Engler  and  family  of the tribes,  and Prantl,  based  development  Schmitz's  in four  scheme  and  concept  on  thallus  recognized  in  was broad and  i.e. Binderelleae [including  1897: 342) and Choreocolax Reinsch  (1874-1875: 61)], Harveyelleae [including only Harveyella Schmitz et Reinke (1889: 28)], Wrangelieae [including Wrangelia C. A . Agardh (1828: M.  136), Atractophora H .  Crouan et P. L . Crouan (1848: 371) and Naccaria Endlicher (1836: 6)] and  Caulacantheae eventually  [including only Caulacanthus Kiitzing (1843: 395)] were  to other  gelidiaceous  plants,  families. Schmitz did recognize, and  he  placed  Gelidium,  however,  transferred  the similarities of  Pterocladia, Suhria,  Porphyroglossum,  Ptilophora, Acropeltis Montagne (1837: 355) and Schottmullera Grunow (in Schmitz 1889: 440; nomen nudum) in the tribe Gelidieae and excluded Wurdemannia from Gelidiaceae. 1892)  Schottmullera  and, along  with  was  renamed  Spencerella  Acanthopeltis  Darbishire  (1896:  by  Okamura  199), it  (in Yatabe,  was  added  to  Naccaria  to  Gelidiaceae (Schmitz and Hauptfieish, 1897).  Oltmanns  (1904)  Wrangeliaceae removal  removed  Harveyella,  Wrangelia,  Atractophora  (Gigartinales). De Toni (1924) apparently  of Wrangeliaceae,  because  he added  and  disagreed with Oltmanns'  Haliacantha  J.  G. Agardh  (1899:  Introduction / 3 109)  and  Gulsonia  Gelidiaceae.  He  Choreocolax  was  Caulacanthus  also  Harvey  (1855:  added  Yatabella  removed  added  Gelidiella,  to  and  the  tribe  Spencerella  Cryptonemiales  to  Gelidiaceae.  Wrangelieae  to  (Sturch,  were transferred to Gigartinales (Sturch,  Hamel, 1934, respectively) was  to  334)  the  tribe  1926), 1926  in  the  Gelidieae.  Harveyella  and  and Feldmann and  and the new genus Gelidiocolax Gardner (1927b: 340) Feldmann and Hamel (1934) renamed  because the name Echinocaulon  Echinocaulon  was a later homonym for a genus of  Polygonaceae. Kylin (1956) revised the contents of Gelidiaceae as it had developed from  Schmitz, transferring Binderella  Spencerella  to  Ceramiales,  removed Gelidiocolax  and  to  adding  Gigartinales, Haliacantha, Beckerella.  Fan  and  Gulsonia  Papenfuss  to Cryptonemiales. Fan (1961) placed Gelidiella  and  (1959)  in its own  family, Gelidiellaceae, based on the absence of rhizines and apparent lack of a sexual generation.  Recently, Santelices and Montalva (1983) merged the monotypic genus, Acropeltis, characterized by shield-shaped tetrasporangial stichidia, with Gelidium,  stating that  this character  schemes of  classification  "has no taxonomic  significance  in presentty  accepted  of the Gelidiaceae." As a result of studies of vegetative characters  that emphasized surface cell morphology, Akatsuka (1986a) erected Onikusa, on  Gelidium  using  pristoides (Turner) Kiitzing  morphological  Akatsuka  (1986c)  characters erected  with  an  Pterocladiastrum,  (1849:  786)  emphasis based  on on  from  South  cortical the  Africa.  based Also  cell  morphology  "Robust"  and "Poor  Knights" forms of Pterocladia lucida from New Zealand.  The position of Gelidiaceae has fluctuated between inclusion in Nemaliales Schmitz  Introduction / 4 in Engler (1892) and recognition as an independent order, Gelidiales Kylin (1923). When  Gelidiaceae  was  erected,  Kiitzing  Periblasteae. Schmitz (1883) placed  (1843) included  Gelidiaceae  it in the order  in its own  order, but later  (Schmitz, 1889) included it in Nemaliales (as "Nemalioninae"), where it remained until Kylin (1923) raised it to ordinal rank, coordinate  with  Nemaliales (as  "Nemalionales"; see Nicolson and Norris, 1983 for correct spelling and use of Nemaliales), C^ptonemiales Schmitz in Engler  (1892), Gigartinales Schmitz in  Engler (1892), Rhodymeniales Schmitz in Engler (1892) and Ceramiales Oltmanns (1904). Kylin saw Nemaliales and Gelidiales as similar in that the fertilized carpogonium in both was the starting point for the gonimoblast. According to Kylin (1923), auxiliary cells were absent in Nemaliales but present in Gelidiales, functioning in a nutritive capacity. Kylin noted that Gelidiales were believed to be diplobiontic, in contrast to the supposed haplobiontic Nemaliales, although he specifically refrained from using life history characters to separate the orders.  However, misinterpreting that Kylin (1923) had elevated Gelidiaceae to ordinal rank on the basis of life history differences (diplobiontic in Gelidiales versus haplobiontic in Nemaliales), Dixon (1961) returned Gelidiaceae to the Nemaliales (as  "Nemalionales").  Polysiphonia-type species, based  He  cited  the  life histor}' (Dixon, on the rare  questionable  existence  1961) in some Gelidium  occurrence  or absence  of a  and Pterocladia  of generations,  gametophytes, and the lack of reports of sexual plants for Gelidiella an  arguement against regarding  pointed  out that  Bonnemaisonia)  some  regular  usually  species, as  Gelidiaceae as strictly diplobiontic. Dixon also  members  of Nemaliales  (as Nemalionales) (e.g.  appeared to be diplobiontic. Hence life history type no longer  Introduction / 5 could be used however,  to separate Gelidiales from Nemaliales. Papenfuss  that  characterized  ordinal  status  uniquely by  cells and Gelidium-type  Gelidiales  contained  for  Gelidiales  was  (1966)  warranted  as  argued,  they  were  nutritive chains, the production of only two pericentral  spore germination.  the  single  family  Gelidiaceae  until  Fan  (1961)  erected  a  second family, Gelidiellaceae, to contain species of Gelidiella (discussed above). The report in  of rhizines in Gelidiella calcicola (Maggs and Guiry,  a  Gelidiella  species  Maggs and Guiry,  from India  (Sreenivasa  1987) contradict Fan's  Rao and  1987) and  Trivedi,  cystocarps  1986, cited in  (1961) characters for separation  of the  family and Maggs and Guiry (1987) merged Gelidiellaceae with Gelidiaceae.  B. CHARACTERIZATION  At  present,  Garbary  OF  Gelidiales is  et al.,  1982;  GELIDIALES  recognized  as  Gabrielson and  an  independent  Garbary,  1986,  order  (Santelices,  1987;  Hommersand  Fredericq, 1988). The order appears monophyletic (Gabrielson and Garbary, 1987),  having  Gelidiales are cap  several  upright  and  1937) axes  Cole,  with  without  1982),  germling an  spore  thick-walled,  germination  and  and  that  show  development  intervening  produce two periaxial cells at elongate,  characters  uniaxial, pseudoparenchymatous.  (Pueschel  (Chemin,  unique  set have  it  apart  from  other  1974; and 1986, orders.  pit plugs with a single plug  "Gelidium-type" proceeding  discoid stage  spore  directly  (Boillot,  germination  to  prostrate  1963).  Axial  or cells  180° (Fan, 1961). and inner cortical cells produce  unicellular  rhizines  sporeling development  that  grow  (Papenfuss,  basipetally. 1966)  and  Gelidium-type rhizines  are  Introduction / 6 features unique to Gelidiales (Fan, 1961; Santelices, 1974).  Rhizines, also referred to as rhizoids and hyphae (Feldmann and Hamel, 1936; Dixon, 1958; Fan, 1961), are unique anatomical features of the vegetative thallus of Gelidiales. Present in all genera of Gelidiales, rhizines are found throughtout the thallus in all except Gelidiella where they are reported to occur only near attachment pads(Maggs and Guiry, 1987). Rhizines differ from multicellular rhizoid filaments that grow between medullary cells of some other genera of red algae, in  that  gelidioid  rhizines  are  unicellular.  This  distinguishes  rhizines sensu  Feldmann and Hamel, 1936) from external attachment rhizoids.  Spermatangia are produced by the transverse division of spermatangial mother cells (Tazawa,  1975), a character unique to Gelidiales at the ordinal rank  (Gabrielson and Garbary, 1987). In a recent study of pre- and post-fertilization events in Gelidium pteridifolium  Hommersand and Fredericq (1988) revised some  previous interpretations. According to Hommersand and Fredericq the carpogonium is intercalary and has associated chains of nutritive cells. An auxiliary cell sensu Drew (1954) is absent, because as the fertilized carpogonium enlarges, it makes non-obligatory fusions with adjacent chains of nutritive cells to form a fusion cell. The nutritive cells fused with are not "specified" (sensu Drew), and thus cannot be  interpreted  as  auxiliary  carposporangium-producing  cells.  gonimoblast  The  within  cruciate, irregular  cruciate or tetrahedral  presumed  of the Polysiphonia-type  to be  fusion  completed for only two species of Gelidium  a  cell  initiates  pericarp.  Tetrasporangia  (Guiry, 1978). The (Santelices,  a filamentous  life  1974), but  are  history is has  been  (Macler and West, 1987; van der  Introduction / 7 Meer, pers. comm.). The life history of the type species of Gelidium  still needs  to be followed in culture.  C. DIFFERENTIATING  GELIDIUM AND  PTEROCLADIA  The most speciose and geographically widespread genera of Gelidiaceae, Gelidium and Pterocladia  (Santelices, 1974), are both represented in the British Columbia  flora. Gelidium and Pterocladia are similar morphologically, and their differentiation has long been problematic (see e.g. Santelices, 1974, unpubl. mscr.; Stewart, 1976; Akatsuka, 1986a; Rodriguez and Santelices, 1987, unpubl. mscr.). Rodrigeuz and  Santelices  (unpubl. mscr.) reviewed  characters previously  proposed for  separating these genera. Currently, only the number of cjstocarp locules (two in r  Gelidium  and one in Pterocladia) can be used consistently to separate the genera.  Okamura (1934) had suggested that rhizine position and medullary cell form are useful  differentiating  characters,  generalizing  that  in Gelidium  rhizines  are  distributed in the cortex and outer medulla and medullary cells are rounded and loosely packed, whereas in Pterocladia medulla  and  medullary  cells  rhizines are distributed only in the inner  are angular and  closely  packed.  Despite this  distinction, Okamura noted some exceptions to these generic rhizine patterns. Others (i.e. Santelices, 1974; Stewart, 1976) have found these characters to be variable and inconsistent, and thus incapable of discriminating between  Gelidium  and Pterocladia  (Rodriguez and Santelices, unpubl. mscr.). Stewart (1976) studying  the  flora, proposed  California  the presence  or absence  of basally incurved  (geniculate) branches as a character to separate these two genera with geniculate branches present in Gelidium  and absent in Pterocladia.  However, this character  Introduction / 8 was  found to be inconsistent when tested on more species of both genera  (Rodriguez and Santelices, unpubl. mscr.).  Recently, Akatsuka  (1981,  1986a, 1986c) proposed  that  surface cortical cell  morphology, arrangement and orientation, in surface view and longitudinal section, are  capable of differentiating  Japanese  species of Gelidium  and  Pterocladia.  Akatsuka's criteria were found only to be consistent for the most basal parts of (upright) axes (Rodriguez and Santelices, unpubl. mscr.). Rodriguez and Santelices (1987, unpubl. mscr.) have proposed apical architecture characters that almost consistently separate Gelidium other Pterocladia  from Pterocladia  (except P.  but  and possibly  species with acute apices). They suggested that apical and  lateral initials are morphologically similar in Pterocladia Gelidium  bulbosa  and that lateral initials  ma3  are never indented in Gelidium.  ?  but are different in  be in cortical indentations in Pterocladia These  criteria were tested  further on  gelidioid species in British Columbia and northern Washington.  Rodriguez and Santelices (1987, unpubl. mscr.) implied that other features of apical architecture (lateral initials close to or far from the apical initial, apical initial protruding beyond or indented between adjacent cortical lobes, and lateral initials protruding from or indented between cortical lobes) also might be used for generic segregation of Gelidium  and Pterocladia.  While discussing the states of  these characters in small groups of species of Gelidium and Pterocladia Rodriguez and Santelices (unpubl. mscr.) do not demonstrate how used to vegetatively discriminate Gelidium and Pterocladia.  the characters may  be  Introduction / 9 D. NEED  FOR ADDITIONAL  INFORMATION  Given the economic importance of Gelidiales as a high quality source of agars (Santelices,  1974, 1986), surprisingly  little  is known  about the biology of  members of the order. Manj' taxa have been described, but the morphological plasticity of species has caused great taxonomic confusion (e.g. Dixon, 1958), and few  careful, comparative,  monographic  studies have  been  made. Gelidiaceae  occupied a pivotal position in the definition of an auxiliary cell, yet there are still many aspects of pre- and post-fertilization development to be resolved, with few  species  having  spermatiogenesis better understand  been  studied  in  detail.  Developmental  studies of  and tetrasporogenesis, particularly ultrastructural, are needed to developmental patterns that may help establish the position of  Gelidiales with respect to other orders. The life history for members of the order has been presumed Polysiphonia-type  and was recently determined from culture  studies for only two species (Macler  and West, 1987; van der Meer, pers.  comm.); further confirmation is needed in other species, particular^' the generic types from  their type  locality. The seaweed  flora of British  Columbia and  northern Washington is being revised (Garbary et al., 1980, 1982; Hawkes and Scagel, 1986a, 1986b; Lindstrom Gelidiales contributes to this goal.  and Scagel,  1987), and this  research on  C H A P T E R  Collections  of  Gelidium  coulteri,  G.  II.  MATERIALS  purpurascens,  G.  vagum  and  Pterocladia  caloglossoides were made on the southern part of British Columbia and some of the San  Juan  Islands  (Washington). In addition, herbarium  specimens were  examined from previous collections throughout the study area. Most of this study was  carried out on freshly collected or 5%  ethanohglacial acetic acid) liquid  Formalin or acetic alcohol (3:1  preserved plants. Herbarium  collections were  relied on for type specimens, for representatives from a taxon's range, and for comparisons of local species with specimens which do not occur in the study area.  A total of 56 field collections were made from 27 sites between Nov. 1983 and Aug. 1987  (Table 2). Collections were made intertidally and from the shallow  subtidal using SCUBA. Some sites were selected based on previous reports of gelidiaceous algae (e.g. South, 1968; Scagel, 1973; Garbary et al., 1984), or from herbarium records. Other sites were selected on the basis of a suitable habitat and substratum (e.g. bedrock, boulders). All collection sites (with the exception of the Barkley Sound locations) were chosen because of their relative accessibility to shore diving; a boat usually was not available.  Of the collection sites (Figs. 1-4), eleven sites were in Barkley Sound, west coast of Vancouver Island (Fig. 2), seven in the Strait of Juan de Fuca and Gulf of Georgia  (Fig. 3), five on Hornby  and  Denman Islands, mid-way along the  western side of the Strait of Georgia (Fig. 4), and four mid-waj' along the 10  Materials / 11 eastern side of the Strait of Georgia on the mainland (Fig. 4).  Frequent collections were made at Orlebar Point, Gabriola Island (Table 2) in the Strait of Georgia, a site chosen for its accessibility (suitable for a day trip), good diving conditions (safe entry, some protection from winter  storms and  availability of accurate marine weather reports) and the abundance of three gelidiaceous approximately  species.  Gelidium  and  Pterocladia  populations  were  studied  ever3' 6-8 weeks throughout the year. Collections were qualitative  with presence/absence, growing condition, and reproductive state of the plants noted.  Herbarium specimens were obtained from AHFH, CANA, FHL, MICH, TCD, UC, US, USM and WTU  (see Table 3 for abbreviations; Holmgren et al, 1978). After  the AHFH loans used in this study were obtained, the AHFH collection was moved to LAM. A detailed list of all herbarium specimens studied is given in Appendix 1.  C H A P T E R III. TAXONOMY AND N O M E N C L A T U R E  A. INTRODUCTION  Since the Gardner,  first  as  have  Gelidium  subsequent  100). Scagel (1957:  area  (Setchell and  reported.  Setchell and  (Collins,  1913:  142) interpreted  114; Kylin,  1925:  all of these reports  13 and to be  (Turner) Lamouroux (1825: 191). [Kylin's (1925) report of G. amansii of Setchell and Gardner's G.  amansii  in his  [by  Scagel (1957:  Island (CANA  A  G.  crinale  is a repeat  some  G.  "G.  records  amansii"  can  Gardner (1927a: 275), a taxon recorded  G. purpurascens  141, as  early  see  from  amansii]  the  southern  part  of Vancouver  Gelidium  cartilagineum  (L.) Gaillon (1828:  15) was reported by  (1925) from San Juan Island, Washington. Gardner (1927a: 280) described  cartilagineum  referred  to  records of by  G.  3843, C A N A 4346, C A N A 4349).  second species,  Kylin  Connell,  (1903) record, because he stated that he did not  collections]. A t least  probably be attributed to  Gardner  Lamouroux (1813: 41) from Vancouver Island  amansii  reports  been  study  Lamouroux (1813: 128) and  Gelidium  J . G. Agardh (1852: 482) have reported  1928:  of Gelidiaceae in the  1903), a total of nine species of  Pterocladia  (1903)  published records  G. G.  var.  robustum  cartilagineum. cartilagineum  to  encompass  the  Californian  plants  previously  Previous British Columbian and northern Washington were considered to be  Scagel (1957). These large, robust  G.  cartilagineum  var.  robustum  plants were later reported by Hollenberg  and Abbott (1965: 1179) as a distinct species, G .  12  robustum.  Taxonomy and Nomenclature / 13 Gelidium  coulteri Harvey (1853: 117) was collected first from the southern part  of Vancouver Island (as G. crinale in 1908, CANA  3474, Scagel, 1957). The  identification of this species in the study area as G. coulteri first was made by Harlin (1969: 258) and Abbott (1970: 1) from Washington, and by Scagel (1973: 138) from British Columbia. Scagel (1957) first reported G. crinale in British Columbia and Washington  (CAN 207, now CANA  3474, and V  1456 (V =  herbarium of the British Columbia Provincial Museum). Gelidium sinicola Gardner (1927a: 278) was mentioned first in descriptive ecological works (Stephenson and Stephenson 1961a: 15 and 1961b: 229) and later as a taxonomic record (Norris and  West, 1966: 176). Gelidium  pulchrum  Gardner  (1927a: 279) has been  recorded only once in an ecological study of zonation in the Strait of Georgia (Stephenson and Stephenson, 1961b: 229). Norris and Wynne (1968) reported G. pusillum  Le Jolis (1863: 139) from Washington, but there are few (Widdowson,  1974; Scagel et al., 1986) there are few subsequent reports of this taxon in the study area. Similarly, there are few records of Pterocladia caloglossoides Dawson (1953: 76), since it was reported by Norris and Hollenberg (1969: 116). The most recent purpurascens  A  addition  to the gelidiaceous flora  of British  Columbia was G.  Gardner (1927a: 279) (Scagel, 1973; Waaland, 1973).  larger number of gelidiaceous taxa occur in adjacent regions of the Pacific  (Table 4). To the south, 11 species of Gelidium, Feldmann  six species of Gelidiella  et Hamel (1934: 529) and four species of Pterocladia are present in  California, Pacific Mexico and the Gulf of California  (Dawson, 1944, 1953;  Stewart 1976; Stewart and Norris, 1981; Santelices and Stewart, 1985). From Pacific Panama, two species of Gelidium  and one species of Gelidiella  were  Taxonomy and Nomenclature / 14 reported in an ecological study (Lubchenco et al., 1984). Nine species of Gelidium, one Gelidiella and four Pterocladia are recorded from Pacific Colombia, Peru and Chile (Santelices and Stewart, 1985; Santelices and Montalva, 1983; Schnetter and Bula Meyer, 1982; Dawson et al, Gelidium, 1977;  four Gelidiella and four Pterocladia  Santelices  Pterocladia,  Yatabella Gelidium  1964). From Hawaii, four species of  plus  and  Stewart,  species  have been  1985).  Beckerella,  of  reported  species, two Gelidiella  from  are currently recognized (Santelices,  Numerous  species  Acanthopeltis,  of Gelidium  Gelidiella,  Onikusa  the northwest Pacific. From  species, four Pterocladia  species and a single species each of Onikusa,  species, two  Acanthopeltis  reported (Akatsuka, 1986b). For China, eight Gelidium  Japan,  and  24  Beckerella  and Yatabella  species, one  and  are  Gelidiella  species and two Pterocladia species are recorded (Bangmei et al., 1983; Santelices and  Stewart, 1985) some in common with Japan. Hommersand  (1972) noted  some floristic elements in common between northeast and northwest Pacific areas, suggesting possible floristic affinities of some Gelidiaceae between British Columbia and the northwestern Pacific.  At  present, the  distinction  between  Gelidium  and  Pterocladia  can  only  be  definitively determined by the number of locules in the cystocarp (Santelices, unpubl. mscr.). Several other vegetative features have been suggested for generic separation. Characters such as rhizine distribution (subcortical or outer medullary in Gelidium versus inner medullary in Pterocladia by Okamura, 1934), morphology of emergence of branches from the main axis (branches basally bent in Gelidium and unbent in Pterocladia,  Stewart, 1968) and shape and orientation of cortical  cells (Akatsuka, 1981, 1986a) have been found inconsistent and unreliable when  Taxonomy and Nomenclature / 15 tested on larger numbers of taxa (Rodriguez and Recently, Rodriguez and  Santelices, unpubl. mscr.).  Santelices (1987, unpubl. mscr.) proposed  shape and  position of the apical and lateral initials to distinguish these two genera. They proposed that in Pterocladia  apical and lateral initials are indistinguishable in  shape and always found depressed below the level of outer cortical cells, whereas in Gelidium,  the lateral initials are different in shape from apical initials and are  never present in cortical depressions (or could be, in a few cases, in indistinct depressions). These characters were consistent for a large number of Pacific species, with the exception of Pterocladia  bulbosa Loomis (1960: 7) (Rodriguez and  Santelices, unpubl. mscr.). All of the features were tested in the species of Gelidium  B.  and Pterocladia in this study and are discussed under each taxon.  METHODS  Material was  hand-sectioned, or sectioned to 10-20  um  on a Bailey Instruments  Tissue-Freez freezing microtome, thick enough to determine relationships between cells as indicated by pit-connections and to trace files of cells. Transverse and longitudinal sections were stained using 1% fixed with 10%  aqueous aniline blue for 15-45 s,  HC1, washed with distilled water and mounted in 40%  Karo  clear syrup to which a small amount of Formalin and aniline blue stain had been  added.  Nuclei  were  observed  by  staining  sections  with  Wittmann's  hematoxylin (Wittmann, 1965) for one hour, followed by fixing with 45% acetic acid, and washing and mounting in either distilled water or in 40% Karo (with a few drops of Formalin added).  In  some  specimens,  apices,  tetrasporangial  stichidia,  male  or  female  Taxonomy and Nomenclature / 16 gamete-producing  branchlets and/or cystocarps were excised. Tissue was fixed in  2.5% glutaraldehyde in Sorensen's phosphate buffer (pH 7.2), washed in buffer, dehydrated through a graded methanol series, infiltrated through a graded series of  JB4 catatysed solution  A  concentrations and embedded  in activated JB4  methacrylate (Ruddell, 1967; Appendix 2). The JB4 blocks were sectioned on a Sorvall JB4 microtome. Other material was embedded epoxy  resin  transmission sectioned  (Spurr, 1969) and fixed electron  with  approximately  a  microscope  glass  2.5 M m .  knife  following  the recipe  (TEM) fixation on a  (Appendix  Reichert OM  Sections of embedded  in Spurr's low-viscositj'  U3  described under 3). Blocks were  ultramicrotome at  material were then stained with  toluidine blue (pH 4.4) for 20 s, rinsed in tap water and mounted in 40% Karo clear syrup with Formalin and aniline blue stain added.  For  each taxon, measurements giving maxima or minima (e.g. maximum height)  are taken from all specimens studied. Axis width and branch angle measurements were made from 10 plants, selected haphazardly. For cortical cell, medullar}' cell and rhizine sizes, 20-50 measurements were made on 4-5 plants of each taxon from  different localities for which fresh or Formalin-preserved specimens were  available. Carpogonial size was taken from  4-5 measurements on 4-5 plants;  cystocarp diameter was from 10 measurements on 1-2 plants; carposporangial size was from 20 measurements on 4-5 plants. Twent} measurements of spermatium 7  diameter were made on 2 plants. Tetrasporangial stichidium length and diameter of  tetraspore release holes was based  on 16-20 measurements of 4 plants;  tetrasporangial size was taken from 30-40 measurements of 3-4 plants, except for Pterocladia  caloglossoides where sporangial size was from 15 measurements on 3  Taxonomy and Nomenclature / 17 plants.  The "!" notation, given with the information on type specimen for the species in this study, indicates that I have  seen the  type(s), and n.v.  (non vide) indicates  that I have not seen the specimen(s).  Rhizine  ultrastructure  Island)  and  was  observed  in  thalli  (from  spermatangial  sterile  (from  male  Orlebar  Point,  gametophytes  Gabriola  cultured  from  tetraspores of whalebone Bay, Gabriola Island) of Gelidium purpurascens,  and in  the  Hornbj'  spermatangial  part  of G.  vagum gametophytes  Island. Longitudinal sections of axes  from  1 mm long, from  branch apex (for young rhizines) and from  Tribune Bay,  3-5 mm proximal to the  2-3 cm proximal to the branch apex  (for older rhizines), were prepared for T E M study.  C.  TAXA  PRESENT  IN  BRITISH  COLUMBIA  AND  NORTHERN  WASHINGTON  Four  gelidiaceous  taxa  in  the  study  area  are  morphologically distinct  and  reproductive characters confirm their identities (Fig. 5). The Gelidiales in the local flora  are  Pterocladia crinale,  Gelidium  coulteri, G. purpurascens,  caloglossoides.  Specimens  G. sinicola and G. pusillum  upon  G. vagum Okamura (1934: 58) and which  were based  records  of  G.  robustum,  G.  were re-examined and found to  be misidentifications of the four taxa reported herein. The report of G. Gardner (1927a: 279) (Stephenson and Stephenson, 1961b) is  pulchrum  unsubstantiated.  Taxonomy and Nomenclature / 18 1. Gelidium coulteri Harvey  1853: 117  Synonym: G. undulatum  Loomis, 1960: 4, pi. 6, fig. 1-4, n.v.  Lectotype: TCD, collected by Dr. Coulter. Selection of the lectotype was by an unknown person, and it is not known if the designation was ever published  (Parnell, pers. comm. 1988). Harvey  tetrasporangial  plant,  so  the  lectotype  (1853) described a  specimen  should  be a  tetrasporophyte, n.v. Type locality: California, U.S.A. Isotypes: TCD!, AHFH 53929!  a. Habitat and Habit  This is the only  local Gelidium  species that is restricted  to the intertidal,  occurring epilithically at relatively protected sites from the mid-littoral to the 0 m tidal level (Canadian chart datum). Plants are soft and non-cartilaginous, although short plants can be slightly stiff. They are brown to reddish-brown when fresh and dry to brown-black. The growth form is a turf, small thalli (to 2 cm tall) are dense, whereas protected  sites  tend  taller thalli (to 5 cm to form  taller,  looser  tall) are less dense. Plants from turfs. Upright  axes  arise from  branched, creeping axes, and may divide to form 2-3 main axes, up to 5 cm tall, but more commonly 1-3 cm tall.  Branching is sparse and usually only to two orders, though occasionally some  Taxonomy and Nomenclature / 19 third order branches may be present (Fig. 6). Branch angles are wide, 45-60°, on widety spaced, irregularly to suboppositely positioned branches. Many . branches are  short and determinate, giving axes a narrow outline. Some short plants  appear densely branched but to a higher order, since branches are more closely spaced together with relatively fewer short proliferous-like branches. Axes are terete and usually narrow, to 0.5 mm up to a maximum of 1 mm less common. Each Branches  wide although occasionally axes may be  and compressed. This wider, compressed form was  branch order is narrower  than the one it arises from.  taper abruptly at the apex, but branch bases are untapered and  unconstricted.  Creeping, terete axes of smaller diameter than uprights, that produce occasional, scattered, pads colourless, formed  attach  the thalli  to the substratum.  Attachment  pads are  of elongated cortical and medullary cell extensions amassed  together, and somewhat spreading on contact with the substratum  (Fig.  8).  Attachment pads are absent in upright axes.  b. Vegetative Anatomy  Thallus construction is uniaxial, with a domed apical cell (Fig. 7) producing a subapical cell that cuts off two periaxial cells. In transverse section, cortical and medullary layers are apparent (Figs. 9, 10). The outer cortex is two to three cell layers and is composed of cells of approximate^' the same size. Outer cortical cells of uprights are isodiametric, 3-8 uffi long and 3-8 um wide, those of the creeping axes are larger and more oval, 6-16 um long and 4-9 um wide.  Taxonomy and Nomenclature / 20 In  surface  view, cortical cells  approximately  of both  upright  and creeping  axes (viewed  halfway between plant apex and base) are randomly oriented and  equidistant, showing no pattern or groups (e.g. pairs or tetrads) (Fig. 11). Secondary pit-connections are absent between the outermost cortical cells, but are present between second and third cortical cell layers of the cortex. The inner cortex, of 1-2 cells, grade to the medulla (Figs. 9, 10), and become progressively larger and more elongate inward. The medulla occupies approximately  75% of the  axis radius (Fig. 9), with elongate, thick walled cells, 28-228 Mm long and 10-21 Mm wide, that are widely spaced, and have abundant secondary pit-connections to adjacent cells (Fig. 10). Intercellular spaces are packed with rhizines (Figs. 9, 10), which are also present in the inner cortex, being sparse in young axes but abundant in older ones. Unicellular and thick walled, rhizines are small diameter (3-4 Mm) and are very elongate (at least 500-700 Mm  long) (Fig. 10), and also  present in creeping axes.  c. Reproductive Anatomy  Tetrasporangia  are restricted usually to determinate fertile branches (stichidia).  Stichidia in Gelidium  coulteri are elongate, (proportionately longer and narrower  than in other local Gelidium  species), 0.7-1.9 mm  long and 0.2-0.4 mm  wide,  and general^ appear somewhat like proliferous branchlets (Fig. 12). The stichidia maj'  themselves bear a few short branchlets (Fig. 13), arranged  irregularly  alternate to subopposite on higher branch orders or on distal parts of major axes (Fig. 12). Tetrasporangia are scattered irregularly on stichidia (Fig. 14), with a tendency for older tetrasporangia to be in the more basal parts of stichidia.  Taxonomy and Nomenclature / 21 Older stichidia produce additional tetrasporangia following release of those formed first (Fig. 16). Developing tetrasporangia first are recognizable 90-300 nm behind the apex, are cut off from a cortical cell 3-4  cells below the surface. The  pit-connection to the bearing cell is basal initially and becomes lateral as the sporangium  increases  its  size  with  much  inward  expansion  (Fig.  15).  Tetrasporangia divide successively [i.e. cytokinesis of the first (transverse) division is completed before the second cytokinesis is initiated], to form four cruciately arranged  tetraspores (Figs. 15, 17). At maturity, tetrasporangia are oval and  deeply embedded in the cortex, but with their outer ends close to the thallus surface (Fig. 17). Mature tetrasporangia are 26-41  um  long and 16-31  Mm  wide,  with adjacent cortical cells usually elongated and curved around them.  Gametophj'tes of Gelidium coulteri have not been collected from the field in the study area, but male gametophytes have been grown in culture from released tetraspores. Female gametophytes were not detected in cultures, possibly because of low  numbers of germinated  sporelings. Gametophytes are  assumed to be  dioecious because male gametophytes only produced male gametes.  Spermatangia  are  borne in superficial sori near the bases of indeterminate  branches. Strongly  fertile  branch  a  to  within  few  male gametophytes produce spermatangia along the millimeters  of  the  apex. At  low  magnification,  spermatangial areas appear pale pink (vs. brownish red where not fertile) because spermatangia are not pigmented. The fertile area appears slightly thicker because the wall is ruptured by spermatangial release, and an abundance of mucilage is present (Fig. 18). Spermatangial mother cells differentiate from cortical cells and  Taxonomy and Nomenclature / 22 are half the diameter of vegetative cortical cells. Each spermatangial mother cell cuts  off a single spermatangium by a transverse  20).  Spermatia  released  from  spermatangia  are  (periclinal) division small,  2.1-3.1  (Figs.  Mm at  19,  widest  diameter, and hemispherical to spherical in shape.  d. Discussion  The  intertidal  spindle-shaped distinguish  few  tetrasporangial  Gelidium  Particularly British  habitat,  the  plants  of  stichidia,  coulteri from  diagnostic are  Columbia  orders  of  turf-like  other  narrow  growth  gelidiaceous  abundant, G.  branches,  short,  coulteri  branch  habit  taxa  in  and  dark  the  study  area.  I found  that  simple branchlets.  correspond  well  with  outline,  G.  colour  coulteri  in  California (Figs. 21-24). A comparison of type material (lectotype and isotype in TCD,  Figs.  California localities plants  22,  with are  are  24; British  isotype  Columbia  Californian  and  plants  contributing to their bushy  of this  species  23) that  from plants  branching patterns,  bushier.  also adds  Fig.  indicates  similar  appear  The fertilitj'  another  Monterey Bay, from but  the  two  Californian  (tetrasporangial) of  order of (stichidial) branches,  appearance.  Cystocarps of Gelidium coulteri are placement  53929,  plants  conspecific. They have  more branched  some of the  AHFH  known from  in Gelidium  California, but evidence for the  is supported  additional^' by  features  of  apical and lateral branch initials. Apical initials are large and domed, protruding beyond  adjacent  initials  differ  cortical  from  cells  apical  and  initials,  form being  acute  branch  inverted  apices  conical,  but  (Fig. 7). are  Lateral  similar  in  Taxonomy and Nomenclature / 23 position to apical cells as both initials are not in depressions of the cortex. These observations agree with Rodriguez and Santelices' (1987, unpubl. mscr,) proposed use of vegetative features to characterize Gelidium.  disagreement with  their Pterocladia characters.  A number of reports pertaining to collections of Gelidium  coulteri from British  Columbia and several misidentified herbarium specimens of G. coulteri (in UBC), show that this species has been confused with other taxa. At times G. coulteri has been referred to erroneously as "G. crinale" 3474 (formerly as CAN A40469, UBC  pusillum"  207); Scagel 1973 - UBC A40467, UBC A40468, UBC  A40470, UBC  A53975, UBC  A31363, UBC  A60236, UBC  (e.g. UBC  A60254, UBC  UBC  "Caulacanthus  A28645, ustulatus"  A12298, UBC  UBC (UBC  A13259, UBC  These  other  taxa  A37402,  UBC  A64934), as "G.  A60268, UBC  A36545), as "G. robustum" (e.g. South, 1968 A29575,  UBC  A31363) and  A36198, as  UBC  "Endocladia  A36199), as muricata"  (UBC  A19739). In a few cases (UBC A30401, UBC  A312511, UBC A37605), Pterocladia coulteri".  A31426, UBC  A28351; FHL 3055), as "G. sinicola" (e.g.  A24837, UBC  UBC A1449, UBC A14403, UBC vouchers  (e.g. Scagel, 1957 - CANA  (G.  caloglossoides has been misidentified as "G. crinale,  G.  pusillum,  G.  sinicola  and  P.  caloglossoides) are generally poorly understood, and probably accounting for their confusion with G. coulteri.  The report of Gelidium sinicola by Stephenson and Stephenson (1961b) from the area of Nanaimo, British Columbia may represent a mistaken identification of G. coulteri. Unfortunately, the authors gave no voucher numbers, description or  Taxonomy and Nomenclature / 24 figures of the  reported  taxon, and  therefore the  plants' identity cannot be  confirmed. Collections from the area (Orlebar Point, False Narrows, Lock  Bay  and Davisons Beach, all on Gabriola Island, near Nanaimo), however, show that G.  coulteri is common. Stephenson and  infralittoral zone, and  Stephenson were listing plants in the  they referred to G. sinicola  They also reported the presence of G. pulchrum, texture  (probably  G. purpurascens,  as a short, moss-like alga. a larger plant with a wiry  see discussion later in this chapter). This  allusion to a turf-like growth habit and short stature suggests that G. coulteri is probably the  taxon that Stephenson and  Stephenson (1961b) were calling  G.  sinicola.  e. Seasonality and  Gelidium  Distribution  coulteri is perennial, persisting as short uprights when not growing  actively. Year round observations  of G. coulteri were made at Orlebar Point,  Gabriola Island (Fig. 3). Plants began showing growth from regenerated apices in early spring (March) and continued growing throughout the summer. They became fertile (tetrasporic) in August, remained fertile until November, but those few plants that were fertile did not produce tetrasporangia abundantly. Growth slowed in November and  December, indicated by  increasing overgrowth of apices  by  diatoms. Plants ceased growth in January and axes were eroded back to shorter lengths. Apices could not be detected and branch ends appeared truncated  and  broken.  Gelidium coulteri is common throughout most of the study area and was  recorded  Taxonomy and Nomenclature / 25 from almost all of my collection sites that had Gelidium or Pterocladia. Its range extends  north  on both the open  Pacific  and Strait of Georgia shores of  Vancouver Island, and on the British Columbia mainland to the Queen Charlotte Islands. The most northerly collection is from Pincer Island, British Columbia (52°11'N latitude). Gelidium and  coulteri ranges south, through Oregon (Doty, 1947)  California to Punta Pequena, (Pacific) Baja California del Norte, Mexico  (Dawson, 1953).  2. Gelidium purpurascens  Gardner  1927a: 275  Synonyms: Gelidium contortum Loomis, 1960: 4, n.v. Gelidium densum Gardner, 1927a: 278, n.v. Gelidium distichum Loomis, 1949: 2, n.v. Gelidium gardneri Loomis, 1960: 5, n.v. Gelidium papenfussii Loomis, 1949: 1, n.v.  Gelidium polystichum Gelidium pulchrum  Gardner, 1927a: 276, n.v. Gardner, 1927a: 279, n.v.  Gelidium ramuliferum  Gardner, 1927a: 279, n.v.  Gelidium setchellii Gardner, 1927a: 275, n.v. Holotype: UC 93572 cystocarpic! Type locality: Moss Beach, San Mateo County, California, U.S.A.  Taxonomy and Nomenclature / 26 a. Habitat and  Habit  Gelidium purpurascens is one of the more conspicuous gelidiaceous species in the study area. It occurs epilithically in protected sites in the shallow sublittoral from 0-12  m  depths, but is most abundant and vigorous at 0-5 m  purpurascens may  depths. Gelidium  be present at localities where there is much siltation, but it is  absent from kelp and seagrass beds. This species is often one of the dominant seaweeds where it occurs.  Plants are robust, cartilaginous, stiff (they are stiff enough to support their branches  when  out  of the  water), and  dark  red  to maroon  and  dry to  blackish-red. Plants grow as individuals or in small clumps and do not form a turf. Individuals may  be much-branched and  "brush-like" if older, or pyramidal  and flat when younger (Fig. 25). Most of the biomass is in upright axes, which are attached to the substratum by a smaller system of creeping axes. Usually a single upright axis arises from creeping axes, but the upright may  divide to  produce 2-4 main axes that can grow to 15 cm tall.  Plants are well-branched with up  to four orders of branches (Figs. 25, 27).  Branches are distichous to irregularly arranged, being subopposite to alternate in different  parts  indeterminate,  of the  plant or  even  along  the  same axis. Branches are  such that older branches are longer  and  more branched than  younger ones. This gives a pyramidal aspect to at least well-branched sectors of the  plant. A  divergent  branching  angle of 30-45°  also contributes  to the  triangular outline of plants. Upper portions of plants appear best developed,  Taxonomy and Nomenclature / 27 whereas lower portions often are less well-branched, possibly because of shading or damage from grazers or abrasion. Proliferous branches may develop, but these are not common or abundant. Axes are terete to compressed. Main axes may be 1  mm  wide, and progressively higher  orders  of branches  are successively  narrower with the highest order branches to 250 (im wide. Branch apices are acute to rounded, with bases that are not tapered or constricted.  Creeping axes are terete, narrower than major and second order upright axes, less  deeply  pigmented  and  sparsely  and  irregularly  branched  (Fig.  27).  Attachment of creeping axes to the substratum is by occasional attachment pads (Fig. 33) that are colourless and formed from elongated cortical cells amassed together into a solid structure (Fig. 34). They are found only on creeping axes (there is no secondary attachment of upright axes).  b. Vegetative Anatomy  The thallus is uniaxial with a domed apical cell, not obvious at low magnification but easily visible in longitudinal or saggital sections (Fig. 28). Each subapical cell cuts off two, laterally opposite, periaxial cells. In transverse sections a pigmented cortex  and unpigmented  medulla can be distinguished with  grading into the medulla without  the inner cortex  an abrupt transition (Figs. 26, 29). Surface  cortical cells are smallest (4-9 Mm long and 2-7 Mm wide), and inner cortical cells are progressively larger (Figs. 29, 30). Outer cortical cells are isodiametric to oval with their longer axes, if detectable, randomly oriented (Fig. 31). Inner cortical cells are oval to short cylindrical, their length being approximately  twice  Taxonomy and Nomenclature / 28 their width  (Fig. 30). Secondary pit-connections are  abundant between inner  cortical cells but are absent from the outer two cortical layers. Medullary cells um wide with innermost cells longest  are cylindrical, 18-107 jum long and 10-25  (Fig. 30), and have abundant secondary pit-connections (Fig. 32).  Rhizines are abundant in the inner cortex and outer medulla, but in decreased abundance in the central medulla (Fig. 26). They are present in creeping axes in the same position and abundance as in uprights. Rhizines are unicellular, narrow (3-4 um  wide), very thick walled and unbranched (Figs 29, 30). They are cut  off from the inner proximal develop proximal  to the  corner of inner cortical cells 280-325 jum  apex as  a  and  small protuberance, growing basipetally  between inner cortical and medulla^ cells to at least 2-3 times the length of the longer medullary cells (i.e. at least 300 um long).  The  cell wall is the rhizine's most conspicuous feature. Its radius (1000-1500  nm) is greater than or equal to the diameter of the protoplast, and the wall is thicker in older cells (Fig. 36) than in younger ones (Fig. 35). Fibrils of the wall are interwoven, but generalty run parallel to the long axis of the cell (Fig. 39) and  are loosest near the plasma membrane (Fig. 35). The  ground matrix  between fibrils is electron transparent. The metabolism of young rhizines appears to be directed toward wall deposition. The profile forming  (Fig. 35). Mitochondria  plasma membrane has a convoluted  are prominent and  closely associated with the  face of dictyosomes (Fig. 39). Dictyosomes are abundant and large,  500-1000 nm  wide and  600-800 nm  tall  (Fig. 39), their  mature vesicles  containing fibrillar material similar in diameter and electron densit}*- to the cell  Taxonomy and Nomenclature / 29 wall fibrils (Fig. 39). However, there is no evidence of vesicles with fibrillar contents secreting to the wall, even in cells showing numerous secretory vesicles. Another form of vesicle (Figs. 37, 39), containing spherical and tubular bodies and resembling multivesicular bodies or lomasomes (Scott and Dixon, 1973b), is present. The origin of these vesicles is unclear, but they do not appear to be directly dictyosome-derived. There is evidence of many of these vesicles fusing with the plasma membrane and releasing their contents to the wall (Figs. 37, 39). Outside and  immediately  adjacent to the plasma membrane are many  tubular body fragments that are not present in older walls, farther away from the cytoplasm. The tubular bodies must either degenerate or become dispersed, forming  new  wall. The  maturation of the fibrillar  abundance  of both  types of vesicles  contents into tubular bodies and  suggests the  their  subsequent  release to the cell wall. In young rhizines, near their point of initiation from a subcortical cell, proplastids are present (Fig. 38, arrow) with onty an inner encircling thj'lakoid, a few  plastoglobuli and areas of DNA.  These proplastids,  included in the rhizine cytoplasm at the time of division of the subcortical cell, do not replicate; they are seen only occasionally and are absent from older or more distant parts of the rhizine.  There is much less secretion and wall deposition activity in old rhizines. The cytoplasm is vesiculate and vacuolate (Fig. 40), many of the vesicle membranes look degenerate, mitochoindria are few and the cytoplasm is thin and sparse. The plasma membrane profile is smooth (Fig. 36) and not convoluted as in more active cells.  Taxonomy and Nomenclature / 30 c. Reproductive Anatomy  Tetrasporangia (Fig.  are located in determinate, stichidial branchlets of upright axes  41) that are short and clavate, compressed, and approximately twice as  wide as vegetative branchlets of the same order or one order lower. Stichidia usually form on distal parts of plants or axes and impose an additional order of branching  over the vegetative  branching  pattern. Thus tetrasporangial plants  appear denser and more branched.  Tetrasporangia are scattered over the stichidial surface, but are absent from the narrow sterile margin (Fig. 42). Sporangia are not oriented in rows, and different ages are interspersed due to the continuous production of tetrasporangia (Fig. 42) after mature ones are released. Similar ages of sporangia equidistant. Tetrasporangia  are approximately  are first distinguishable 30-60 um proximal  to the  apex (Fig. 43). The pit-connection to the subtending cortical cell is initially basal (arrowheads in Fig. 43), but later becomes lateral expands deeper into the branch. Tetrasporangia  as the tetrasporangium  are cruciately divided with the  first division periclinal and the second (in each half) anticlinal (Fig. 44). At maturity tetrasporangia are 40-81 am long and 20-49 um wide. Tetraspores are pyramidal on release because of mutual compression in the sporangium, but soon round up. The cortex of the tetrasporangial part of the stichidium is thicker than the vegetative axes and occupies  2/3 of the axis radius. Inner cortical cells  adjacent to the widest part of the tetrasporangium are elongated.  Gelidium purpurascens is dioecious, and male and female gametophytes occur in  Taxonomy and Nomenclature / 31 low  numbers  in  field  collections.  Gametophytes  are  isomorphic  with  tetrasporophytes. Carpogonia and cystocarps occur terminally on highest order branches or on short, side branchlets of upright axes (Figs. 45, 48). These branchlets have the capacity to continue growing, but do not if reproduction is successful. Occasionally a second fertile area occurs when apical growth continues beyond the first set of carpogonia  (Fig. 48). The apex of the carpogonial  stichidium of G. purpurascens is acute, with the apical cell not recessed into an apical  notch  as  reported  by  Hommersand  and  Fredericq  (1988) for G.  pteridifolium from South Africa.  Numerous (20-32) carpogonia develop in the fertile area, and are initiated five to seven axial cells proximal to the apex (Fig. 47). The apex continues to grow, and later in development the cystocarp appears 1-3 mm  proximal to the apex.  Carpogonia are 13-29 um long and 3.5-5.5 um wide. Sizes are approximate as measurements did not account for the curvature of cells. Carpogonia differentiate in third order filaments above and below the second order plate (i.e. a plate of second  order filaments connected  by  secondary  pit-connections), and  their  trichogynes project to both surfaces of the branchlet (Fig. 47). Carpogonia are intercalary in vegetative filaments and have two pit-connections (Fig. 46). The carpogonium is the second cell in a third order filament cut off from a cell of the second order plate, close to the axial filament. The upper pit-connection, of the carpogonium is to a vegetative cell (Fig. 46) that produces higher order vegetative  branches. In non-fertile  axes, the cell  in the position  of the  carpogonium has a second upper pit-connection (three pit-connections in total) to another branched vegetative chain. Thus, the carpogonium replaces a vegetative  Taxonomy and Nomenclature / 32 chain whose outermost cells contribute to a small patch of thallus surface. The absence of surface cortical cells, which have been replaced by the carpogonium, leaves a small gap or "hole" in the thallus surface. The confluence of adjoining "holes", resulting from a double row of carpogonia, creates a medial furrow in the  upper and lower surface of the female branchlet. Trichogynes  protrude  through this gap.  Carpogonia farther back from the apex are sessile, cut off from the subtending cell by a distinctive concave wall. It was determined (see Chapter 5) that intercalary carpogonia are functional and may be fertilized. When they are past fertility, the concave wall cuts off the distal part of the carpogonium, which then degenerates while the "supporting" cell remains functional.  Upon fertilization, nutritive filaments composed of short, curving chains of small, densely  staining, globular cells, proliferate  from  basal  cells  of third  order  filaments and curve towards and around second order plate cells. The fertilized carpogonium forms a large, multilobed, multinucleate fusion cell by fusing with surrounding cortical cells. The fusion cell cuts off gonimoblast initials that form a branching, filamentous gonimoblast  and that produce uninucleate  carposporangia  terminally and laterally (Fig. 51). A more detailed description of post-fertilization development is given later in Chapter 5.  Mature  cystocarps  of Gelidium  purpurascens,  420-560  um  in diameter, are  bilocular with one ostiole per locule (Fig. 50). They are spherical, protruding from both surfaces of the branch and are wider than the sterile portion of the branch  Taxonomy and Nomenclature / 33 (Fig. 50). The  carposporophyte is centered around the second order plate (Fig.  50) of gametophyte cells. The  cystocarp cortex is the same thickness as the  vegetative cortex, but it is composed of more cell layers. Cystocarp cortical cells are smaller and  somewhat more tightly packed than in the vegetative cortex.  Mature cystocarps of G. purpurascens have characteristic filaments between second order plate cells and the cortex (Figs. 49, 50). These filaments are third order files of cells that arise from cells of the second order plate, contribute to the cortex and  extend to the surface. As  the cj^stocarp cavity expands and  the  cortex is pushed away from the second order plate, third order filaments stretch but do not break and  are thus intact in the mature cystocarp. There is no  evidence to suggest a nutritive function for these stretched filaments; they are simply part of the vegetative construction of the thallus over which cystocarp morphology is imposed. Filaments of a similar appearance have been reported as "nutritive filaments" in other genera, e.g. Gracilaria are not homologous to "nutritive filaments" of  Carposporangia are 20-39 um  long and  (Dawson, 1949), but they  Gracilaria.  12-22  um  wide, ovate to obpyriform,  uninucleate, pigmented, have a large, stellate chloroplast and are formed singly, i.e. not  in chains  (Fig. 51). Thej' have  a  thick coating of mucilage or  polysaccharide that fills the cystocarp cavit3'. Carpospores are released through an ostiole in the pericarp wall.  Spermatangia are produced in superficial sori on short stichidial branchlets of upright  axes  (Fig. 54),  similar  in position and  arrangement  (Fig. 52) to  tetrasporangial stichidia. In living plants at low magnification the fertile area has  Taxonomy and Nomenclature / 34 a colourless halo (Fig. 54) that corresponds to the spermatangial mother cell wall ruptured by gamete release, partially empty of spermatia and surrounded with large amounts of mucilage. Surface cortical cells 200-300 um  proximal to the  apex are transformed by one to two  longitudinal divisions into two  spermatangial  a  mother  cells  replacing  single  outer  cortical  to four  cell.  Each  spermatangial mother cell cuts off a single spermatangium by a characteristic transverse (periclinal) division enlarging  (Fig. 56). As  spermatangial vacuole  magnification. A  appears  the spermatangium  basally  as  a  single, conical spermatium, 3.1-5.2 um  clear  develops, an area  at high  diameter, is released  through a narrow channel in the thickened wall (Figs. 55, 56), and rounds up into a spherical or short oval shape (Fig. 53).  d.  Discussion  The bilocular cystocarp of this species dictates generic placement in Gelidium, opposed to Pterocladia. that Gelidium which  two  Rodriguez and Santelices' (1987, unpubl. mscr.) proposed  could be separated from Pterocladia have  been  as  investigated  morphologically different in Gelidium,  using vegetative characters, of  here: 1) apical  and  lateral  initials are  whereas they are similar in Pterocladia, and  2) lateral initials never are found in indentations of the cortex in Gelidium whereas they are in Pterocladia. from the study area and  Apical architecture of G. purpurascens  plants  from California agree with this suggested pattern:  lateral initials are not in depressions and they are inverted conical in shape (Fig. 58),  whereas the apical initial is domed and hemispherical (Fig. 57). Rodriguez  and Santelices (unpubl. mscr.) also observed that the cortical cell pattern of the  Taxonomy and Nomenclature / 35 basal 1-2 cm and Pterocladia,  of upright axes could be used to discriminate between Gelidium but the cortical cell pattern was  not consistent elsewhere on the  axes as proposed by Akatsuka (1981, 1986a). In surface view, basal cortical cells of G. purpurascens  are isodiametric, equidistant and  are not aligned parallel to  the axis (Fig. 60), in agreement with Rodriguez and characterization of Gelidium.  Santelices' (unpubl. mscr.)  Localized in small patches at the very base of  uprights, elongate, elliptical, cortical cells have their long axes perpendicular to the branch axis (Fig. 61). This is unlike the pattern noted for both Gelidium (isodiametric and no alignment) and Pterocladia  (elliptical and parallel to branch  axis) (Rodriguez and Santelices, unpubl. mscr.), but this pattern is not universal or widespread on a branch.  The  British Columbian entity called Gelidium purpurascens corresponds well with  the holotype of G. purpurascens (UC 93572) (Figs. 62, 63) and other Californian material belonging to this species. Diagnostic characters given by Gardner (1927a) are that G. purpurascens (Figs. 62, 63) is tall, but not as large and coarse as G. robustum (Hollenberg and Abbott, 1965); well branched, including lower parts of axes; has compressed, basally constricted and geniculate pinnate branches to four or five orders; produces initially single cystocarps on fertile branchlets and later exhibits proximal, lateral, cystocarpic branchlets. Stewart (1976) modified the description  to  include  geniculate  abundant basal branching diagnostic  character.  and  Plants  or  non-geniculate  plants, with  or  without  did not specifiy basal branch constriction as a collected in  the  study  area fit the  redefined  description, particularly as branches of G. purpurascens are not basally constricted when compared to the marked basal constriction of higher order branches of G.  Taxonomy and Nomenclature / 36 robustum.  Stewart (1976) generally described Gelidium purpurascens as large and similar in branching pattern to G. robustum, but not as coarse or tall as the latter species. A comparison of the holotype (UC 93572, Figs. 62, 63) and of specimens of G. purpurascens from the study site, with the holotype of G. robustum (UC 294572) (Fig.  138)  showed  that  G.  purpurascens  G.  and  robustum  are different  morphologically. Although both species show the same number of branch orders, G.  purpurascens  purpurascens  is more  there  proliferous  are often  several  and  more  dominant  densely  branched. In  axes that  may  G.  divide into  equivalent axes, whereas G. robustum has a single main axis obvious throughout the length of the plant. Transverse purpurascens  sections of vegetative branches of G.  show that the subcortex may  be thick but that ordered rows of  cells are not evident; similar sections of G. robustum branches also show a thick subcortex, but it is ordered into pallisade-like rows. Tetrasporangial stichidia also differ between these species; stichidia of G. purpurascens have a narrow, almost inconspicuous, sterile margin, whereas stichidia of G.  robustum  sterile  branching  margin.  purpurascens  Thus,  despite  some  can be distinguished  similarity  morphologically  in and  have a wide pattern,  anatomically  from  G. G.  robustum (Table 5).  Some of the earliest Gelidium purpurascens CANA (Scagel,  (CANA  4349, UBC  collections from British  3473, CANA  3740 (Fig. 59), CANA  Columbia are of G. 3843, CANA  A1952). These were called G. cartilagineum  1957), and later  G.  robustum  when Hollenberg  and  4346,  var. robustum Abbott (1965)  Taxonomy and Nomenclature / 37 elevated the variety to species status. Wherever possible, subsequent reports of G. purpurascens in the study area (Scagel, 1973; Foreman, 1977; Lindstrom and Foreman, 1978; Pueschel and Cole, 1982) have been confirmed. Publications that reported "G. robustum" in the study area (Scagel, 1967; South, 1968, record from  Twin Beaches, Gabriola Island; Scagel, 1973), cited voucher  herbarium  specimens or illustrations, correspond not to G. robustum, but to G. purpurascens.  e. Seasonality and  Distribution  Gelidium purpurascens is perennial, growing from April through October, but only persists as uprights with little growth (demonstrated by heavy diatom epiphytism) over winter months. Tetrasporophytes are fertile from February to December, most abundantly in May  to October. Female gametophytes bear carpogonia in  August and November collections, and male gametophytes are fertile in June, Jul}' and November.  Gelidium purpursacens is widespread throughout the study area and found along the  British  Columbia  mainland, Queen  Charlotte Islands, Hornby  Island and  Denman Island, Strait of Georgia, Gulf Islands, Strait of Juan de Fuca and along the length of the west coast of Vancouver Island. It is abundant at sites where it occurs and is present at all of m}' collection sites where any other species of Gelidium  and/or Pterocladia are found.  Tetrasporophytes are fertile throughout the study area but fertile gametophytes are  rare. Female gametophytes have been collected onty from Diana Island and  Taxonomy and Nomenclature / 38 Geer. Islets, both in Barkley Sound (Fig. 2), and male plants have been identified only from Nootka Sound, Vancouver Island (Fig. 52), Esteban Point, Vancouver Island and Barkley Sound.  Gelidium  purpurascens  is distributed from Baja California del Norte (Stewart,  1976), to southeast Alaska (UBC A69432, UBC A69433). Its morphology is plastic, although in British Columbia it is distinctly different from other local species. In some environments plants are much less robust, paler and branches become long and flexuous, especially near apices, appearing similar to G. vagum Okamura (1934). The two species are still distinguishable by the cartilaginous texture of G. purpurascens  (vs. lax and soft in G. vagum),  compressed axes of G. purpurascens  the terete to  (vs. compressed to flattened in G. vagum),  the narrow sterile margin of tetrasporangial stichidia in G. purpurascens (a sterile margin is absent in G. vagum), dioecious gametophytes of G. purpurascens (vs. monoecious gametophytes in G. vagum), and the presence of stretched third order filaments in mature cystocarps of G. purpurascens (these filaments are broken in G. vagum cystocarps).  3. Gelidium vagum O k a m u r a 1934: 58  Type: A holotype was not designated by Okamura (1934) in the protologue. Syntype specimens are in MAK (Akatsuka, pers. comm. 1987) but not available for loan. It is not known if Okamura designated a holotype, but no lectotype has been published.  Taxonomy and Nomenclature / 39 Type locality: Not specified, although the distribution in Japan (around Honshu and southern .Hokkaido) was described (Okamura, 1934).  a. Habitat and * Habit  Gelidium  vagum usually grows in the shallow subtidal from 0-13 m  depths as  well as in very low intertidal pools. Wherever plants or parts of plants have been exposed to air they are bleached and dead. This species is most abundant in the shallow subtidal, 0-4 m, where it may  be a dominant, but it is not a  dominant in populations of deeper water. Gelidium  vagum grows epilithically on  vertical walls and tops and sides of boulders, occuring in somewhat silted (e.g. Tribune Bay, Hornby Island) to protected localities (e.g. Denman Island and Ford Cove, Hornby Island) with a moderate amount of water motion. Plants are soft and lax and do not remain upright when exposed in air as does the more rigid G. purpurascens.  They range from red to yellowish-red (deeper plants), and dry  to brownish red, and grow as individuals, not as a turf, even though several upright axes may  arise from the same creeping basal axis. Plants may  be up to  10 cm tall, but are usually shorter (Figs. 64, 65), and can be fertile (tetrasporic or gametophytic) in the field at 1 cm tall.  Gelidium thin  and  vagum shows considerable morphological variation in growth form, from sparsely  branched, to wide, robust and  well branched. Generally,  deeper-growing plants are thinner, lighter in colour, smaller and less branched. Plants are branched to the fourth order, and occasionally well-developed plants have fifth order branches (Fig. 64). Branching is divergent with a branch angle  Taxonomy and Nomenclature / 40 of 45-60° for all ranks of branching. One to three main axes usually are distinguishable, although in some plants the main axis divides subdichotomously distally, and neither  branch  can be recognized  as derivative. Branching is  distichous and irregularly alternate to subopposite, and successive branches are well separated (approximately 1-2 mm  apart). Shallow-growing plants may be  quite bushy, whereas deeper plants are sparsely branched. Axes vary from compressed to flattened, with the most robust plants being conspicuously flattened, especially at branch points, and smaller, less robust plants terete to compressed. Main axes and lower order branches vary in width from 0.2-2.5 mm.  Branch  bases often are narrowest (but never constricted), with the branch axes widening gradually to the widest point, about 1-2 cm behind the apex. Higher order branches are usually progressively narrower and constricted basally, but may taper abruptly to acute apices, the distal 1 mm unpigmented  of which appears whitish or  in fresh material. Occasionally, ultimate branches become long and  whip-like (Fig. 66). Higher order axes branch in the same way as main axes.  Creeping axes are pigmented, irregularly branched, terete, narrower than most upright branches and bear attachment pads. Attachment pads are unpigmented, scattered irregularly along prostrate axes formed from the confluence of elongated cortical cells, and spread out at the point of contact with the substratum.  b. Vegetative Anatomy  Thallus construction is uniaxial, with a conspicuous, domed apical cell domed (Fig. 68). Cells of the axial row each cut off laterally two periaxial cells (Fig. 68). In transverse section cortical and medullary layers are distinctive (Figs. 67,  71),  Taxonomy and Nomenclature / 41 with a smooth and  rapid transition between inner cortex and  outer medulla.  Outer cortical cells are isodiametric to slightly elongate, 4.0-11.3 /xm  long and  3.0-6.2 jim wide, with their longest axes irregularly oriented in the periclinal plane and equidistant (i.e. not associated into clusters) (Figs. 69, 70). However, near the apex, where cell divisions are occurring, the longest axes of outer cortical cells are in the anticlinal plane (Fig. 68). Secondary pit- connections are absent between cells of the outer two  cortical cell layers, but are abundant  between inner cortical cells. Inner cortical cells are larger and (16-60 ixm  long, and  12-29  Mm  wide) than  outer  cylindrical with their long axis at approximately  more elongate  cortical, and  short  and  30-45° to the branch axis.  Medullary filaments are oriented periclinally and cortical filaments are anticlinal, so that the transition part of filaments of the inner cortex and outer medulla are oriented at oblique angles as noted by Akatsuka for G.  vagum  in Japan  (1981, 1986a).  Medullary  cells are larger (75-290 Mm long and  10-29  Mm wide) than inner  cortical cells, with the longest cells deepest in the medulla (Fig. 72). The}' are cylindrical, unpigmented, thick-walled and have abundant secondary pit-connections (Fig. 68).  Rhizines are intercalary between inner cortical and medullar}' cells (Figs. 67, 71) and increase in abundance farther behind the apex. They are cut off from the proximal end of inner cortical and outer medullary cells (Fig. 73) approximately nine axial cells behind the apex. Rhizines are abundant in the inner cortex and outer medulla (Fig. 71), and later grow between medullary cells (Fig. 67). They  Taxonomy and Nomenclature / 42 are unicellular, unpigmented, refractive, elongate (measured to at least 970 um),  um  and most probably  are longer), have a narrow diameter (3-4  very thick  walls (1.1-1.5 Mm),  and do not form secondary pit-connections with adjacent cells.  c. Reproductive Anatomy  Tetrasporangial stichidia are produced on upright axes along with the vegetative branches (Fig. 76). Unbranched or once branched, 1.5 mm stichidia  are  usually  determinate  and  arranged  long and 1 mm  wide,  distichously, subopposite to  alternate, cylindrical to compressed, ovate to lanceolate with a blunt-rounded to blunt-tapered  apex (Fig. 75). Occasionally the stichidial apex continues to grow  bej'ond a fertile area and produces another tetrasporangial area a short distance beyond the original patch (Fig. 74), however, usually the apex does not continue vegetative  growth but  produces one  to three  short stichidial branchlets just  proximal to the original fertile area. Tetrasporangial stichidia may narrow or wide plants over one  cm  tall and may  be produced on  be present on any branch  order, but usually are not present on the main axis if there is more than one branch order.  Tetrasporangia  are scattered randomly, i.e. not arranged in rows or V pattern,  over the stichidial surface. There are generally more mature tetrasporangia in the basal part of the stichidium as these differentiate first, but, after release of the first-formed sporangia, secondary production of tetrasporangia results in a mixture of sporangia of different ages (Figs. 79, 81). Tetrasporangia  occur around the  entire diameter of the axis, and a sterile stichidial margin is lacking (Fig. 75).  Taxonomy and Nomenclature / 43 Six to seven axial cells proximal  to the  apex (Fig. 77) tetrasporangia are  initiated from inner cortical cells (Fig. 79). They are terminal on one of the two branches borne by a cortical cell. The (Fig.  79),  but  becomes lateral  sporangial pit-connection is initially basal  (Fig. 83)  by  the  basal  expansion  of the  sporangium below the pit-connection. The first division of the tetrasporangium is transverse (Fig. 81), and  the second division produces four cruciately arranged  tetraspores (Fig. 78), 46-89 um  long and  25-46 /um  wide. Spores are released  through small holes, smaller than spore widths, (diameter 5.1-10.0 um)  in the  sporangial wall (Fig. 80). Either spores must be squeezed during release, or holes are stretched. The space remaining after spore release contains a substance that stains weakly with aniline blue and appears to have been partially drawn or squeezed out of the cavitj' (Fig. 83). Based on similar observations  of other  workers studjdng tetrasporogenesis of red algae (Peyriere, 1970; Pueschel, 1982), this substance is probably  mucilage or polysaccharide  from  sporangium wall  breakdown or a spore secretion.  Gelidium vagum is monoecious, with male and female gametes consistently formed in adjacent sections of apical parts of gametophyte branches (Fig. 85); no fertile gametophytes have been seen that produce only one sex of gametes. Gametangia are borne on short, determinate stichidia of upright axes (Figs. 82, 84), but occasionally  stichidia  occasionally may  bear  two  more  stichidial  branchlets.  Stichidial  apices  continue growth after a cystocarp has been produced and give  rise to another patch of spermatangia and carpogonia 1.5-3  mm  distal to the  first gametangia. Gametangial stichidia are distichous and irregularly alternate to subopposite on second or third order branches (except on very small, 1-2 cm tall  Taxonomy and Nomenclature / 44 plants where they are on the main axis). Spermatangia are located just proximal to the area that produces carpogonia (9-10 axial cells or 70-250 um behind the apex). At low magnification fertile male areas can be recognized as areas with a somewhat  raised  and  thickened  cuticle  that  corresponds  to  the  layer  of  unpigmented spermatangia (Fig. 85).  Carpogonia are produced three to six axial cells (50-220 um) proximal to the apex, and  the region of carpogonial production extends proximally along the  length of 4-5 axial cells (100-300 um). Carpogonia are elongate (10-25 nm long, and 3-6  um  wide at the widest, basal part), curve forward and outward and  taper gradually to a trichogyne that extends directly to the thallus surface (Fig. 90). The carpogonium  is intercalary, the second to basal cell of a third order  filament, and has two pit-connections, one to the next distal cell and one to the next proximal cell of the filament (Fig. 90). All other cells of the filament appear unmodified. A double row of carpogonia (from both lateral sides of the axial row) are directed toward the upper branch surface and another double row are  directed to the lower branch surface because third order filaments extend  both above and below the second order plate (Fig. 89). In the vegetative branch, cells in the same position as the carpogonium  have one proximal pit-connection  (to the basal cell of the file) and two distal pit-connections (to a distal cell in the same file and to a higher order file), but in the gametangial branch the carpogonium replaces one file and its subsequent branches, creating a gap in the branch surface. The alignment of gaps from adjacent carpogonia forms an axial furrow through which trichogynes protrude.  Taxonomy and Nomenclature / 45 Associated with carpogonia and young carposporophytes  are short, curved chains  of two to four isodiametric cells, referred to as "nutritive filaments" (Fig. 88) (Hommersand and Fredericq, 1988). These are cut off from basal cells of third order filaments and curve toward the second order plate. A lobed, multinucleate fusion cell develops after fertilization, from the fusion of the carpogonium and cortical and  subcortical cells (Fig. 92). Fusion cell lobes cut off gonimoblast  initials that develop into gonimoblast filaments composed of small, elongate cells that do not stain darkly with aniline blue. Gonimoblast cells fuse with apical cells of nutritive filaments. The  gonimoblast  between  order  cells  of  carposporangia,  the  second  35-75 um  long and  branches extensively around  plate,  15-25  um  producing  single,  wide in one  and  obpyriform  of two  locules  formed between the second order plate and cortex (Fig. 93). A large quantity of polysaccharide  or mucilage,  staining  faintly  with  aniline blue, surrounds  the  carpospores, much of it remaining in the locules after carpospore release. A more detailed description of carpogonium and carposporophyte  development is given in  Chapter 5.  As  the cystocarp swells, third order filaments between the cortex and second  order plate (Fig. 91) stretch and are broken, in contrast to Gelidium purpurascens cystocarps where filaments stretch but wider, 300-550 um  remain intact. Mature cystocarps are  in diameter, than the vegetative part of the bearing branch,  bilocular, and locules are separated by the plate of second order filaments (Fig. 93). The carposporoplryte is restricted to the area around the second order plate. The cystocarp is domed at the ostiole, but not apiculate, and has one ostiole per locule (Fig. 93).  Taxonomy and Nomenclature / 46 Spermatangia are cut off from spermatangial stichidial surface for 150-500 um  mother cells over most of the  immediately  proximal  (9-10 axial cells or  70-250 /um proximal to the apex) (Fig. 85) to the carpogonial area. Stichidia may include some small patches of vegetative cortical cells. At low magnification, fertile  male  areas  can be recognized  by their  raised  and thickened wall,  corresponding to the layer of unpigmented spermatangia and mucilage (Fig. 85). Spermatangial mother cells are narrow and elongate (Fig. 86), and each produces a single spermatangium by a periclinal (transverse) division. A clear, rounded body corresponding spermatangia. Each  to a spermatangial spermatangium  vacuole can be seen basally in some  releases  a  single  spermatium  (Fig. 87)  1.5-2.5 um in diameter. Spermatia are hemispherical to blunt-conical at release, later becoming spherical. From  the first-formed (most proximal)  spermatangial  mother cells to the youngest (most distal) along the length of male area of the stichidium  can  be  seen  a  progression  and  intermixing  of  spermatangial  developmental stages, suggesting percurrent production of spermatangia.  d. Discussion  The bilocular cystocarp of this species indicates placement in Gelidium.  Supporting  this generic placement, apical and lateral initials are morphologically different and neither is in a cortical indentation. The monoecious nature of gametoplrytes of this species does not correspond to any Gelidium species previously reported from British Columbia. Gelidium purpurascens is morphologically distinct and dioecious. I have seen separate male and female gametophytes of G. purpurascens (male: UC 305373; female: UC 93572, UC 276633, UC 296689, UC 305364, UC 305371),  Taxonomy and Nomenclature / 47 and G. contortion  Loomis, G. distichum  considered synonyms of G. purpurascens, Gelidium  coulteri  is dioecious and  Loomis and G.  irregulare  Loomis, all  are known to be dioecious (Silva, 1978).  morphologically  distinct  (Macler  and West,  1987), and Pterocladia caloglossoides also is morphologically distinct.  Monoecj' has not been reported previously in Gelidium, but has been observed in Acanthopeltis feature  japonica  of G.  (Gelidiaceae) (Kaneko,  vagum  from  British  1968). Monoecy  Columbia  and  gametes  is a consistent are abundantly  functional (judging from the large number of fertile cystocarps), eliminating the possibility  of the observed condition being  a genetic abberation. There are,  however, a large number of Gelidium species for which males are unknown. Two such species, G. vagum from the northwestwern Pacific (Japan and China) and G.  johnstonii  from  the  Gulf  of California,  resemblance to the British Columbian monoecious  Illustrations  of Gelidium  show  a  striking  morphological  Gelidium.  vagum from China (Bangmei  et al.,  1983; Santelices,  unpubl. mscr.) and Japan (Okamura, 1934; Segawa, 1959) are morphologically similar  to the monoecious  Gelidium  from British Columbia. In particular, the  original description and figures of G. vagum (Okamura, 1934 pi. 25) described a morphology  completely  within  the range  of form  displayed by  the British  Columbian monoecious species. British Columbian plants also agree with Santelices' (unpubl. mscr.) description of Chinese G. vagum. To date, Okamura has provided the only illustrations of a cystocarpic plant and a cystocarp, but the diagnosis lacks illustrations or mention of internal cystocarp anatomy.  Taxonomy and Nomenclature / 48 Because the type of Gelidium Metropolitan University (MAK; other  Akatsuka, pers. comm.), was  specimens of Japanese G.  Museum, Tokyo [TNS TNS  vagum, in the Makino Herbarium of the Tokyo  25847] and  25817, TNS  vagum, borrowed from the National 25823 (Fig. 94), TNS  a single sheet with two  (A56807), were examined. Two  unavailable for loan,  25824, TNS  Science 25825,  to four Japanese plants in  plants are cystocarpic [TNS  25824 (Fig 95),  UBC TNS  25825], and when sectioned were found to be monoecious (Fig. 96). Spermatangia in these plants are like those in the British Columbian monoecious species, occurring just proximal  to the  female gametangial area, and  later at, and  extending below, the cystocarp base. Cystocarp anatomy also is similar in British Columbian and  Japanese plants (Fig. 96). Occasional^, however, cystocarps of  Japanese plants have an intact third order filament between the second order plate and cortex (arrowhead in Fig. 97), whereas equivalent filaments in British Columbian material are broken during cystocarp expansion.  As in the British Columbian material, tetrasporangial stichidia of G. vagum lack a sterile margin, and have tetrasporangia scattered irregularly over the stichidial surface (Fig. 98). Tetrasporangial stichidial anatomy is similar in both entities, although in Japanese plants, tetrasporangia are embedded somewhat more deeply in the thallus with one or two  cortical layers lying outside the outer end of  sporangia (Figs. 99, 100).  In light of the similarity of other characters, i.e. the morphology of thalli with regard  to branching  pattern, branch size, shape and  degree of constriction,  presence of male and female gametangia not only on the same thallus (monoecy)  Taxonomy and Nomenclature / 49 but in the same arrangement, the absence of a sterile margin in tetrasporangial stichidia  and the arrangement  of tetrasporangia  in stichidia, the difference  observed in breakage of sterile filaments linking cells of the second order plate and cortex, and in the degree of tetrasporangia embedding are considered minor. Thus, this monoecious British Columbian Gelidium  is considered to be conspecific  with G. vagum.  Spermatangia also were unknown for Gelidium  johnstonii  Setchell et Gardner  (1924: 742), when it was described from the Gulf of California, and illustrations 1  (Setchell and Gardner, 1924 pis. 46a, 72, 73) showed a strong morphological resemblance between this species and G. vagum. Akatsuka (1986b) suggested that G. vagum was related closeh' to G. johnstonii. Examination of the holotype (Fig. 101), isotypes and paratype, as well as specimens identified as G. johnstonii collected by Dawson (AHFH AHFH  4192, AHFH  2211, AHFH  4150, AHFH  4156, AHFH 4179,  4193, AHFH 4194, AHFH 50267, AHFH 50268, AHFH  50299, LAM 52684 in AHFH, LAM 52894 in AHFH), revealed that small, to 5 cm tall, plants resemble G. vagum morphologically, but that larger plants are more robust and coarse, lacking the delicate appearance of G. vagum.  Gelidium  johnstonii and G. vagum differ anatomically in several vegetative and reproductive features listed below and summarized in (Table 6). Contrary Gardner (1924) the medulla of the holotype  of G. johnstonii  to Setchell and is not sparse,  however the outer cortex is three to four cells as they had reported (Figs. 102, Holotype: (Johnston 27) CAS 1343 in UC! Isotypes: (Johnston 27) CAS 484385 in UC!, CAS 484386 in UC!, CAS 484388 in UC! Paratype: (Johnston 13) CAS 484390 in UC! Type locality: Bahia San Francisquito,Gulf of California, Baja California del Norte, Mexico 1  Taxonomy and Nomenclature / 50 103). In contrast, the outer cortical cells of G. vagum are unaligned.  Tetrasporangial stichidia of Gelidium johnstonii are elongate, to 3 mm,  spatulate  (Fig. 106), flattened, and have a blunt apex (Fig. 104) and a sterile margin of variable width (Fig. 104) unlike the shorter, ovate, terete to compressed stichidia with more pointed apices of G. domed and  vagum. The  apical initial of G. johnstonii is  protrudes slightly beyond adjacent cortical cells (Fig. 105), but in  tetrasporangial and gametangial stichidia the apical cell is level with the adjacent cortex or recessed slightly between adjacent cortical lobes (Fig. 109). In G. vagum, the apical initial of both vegetative branches and fertile stichidia protrudes beyond adjacent cortical cells (Figs. 68, 75). In G. johnstonii a furrow develops immediately  behind the apical cell in gametangial  and  tetrasporangial stichidia  (Fig. 105, 109) which is not filled in by cells of higher order filaments for 250-350 Mm  behind  the  apex. An  apical  furrow  is absent  in G.  vagum  tetrasporangial stichidia (Fig. 75), but is present in gametangial stichidia, as a result  of cortical  cells being  replaced by  carpogonia  (Fig. 89, development  described earlier). The furrow in G. vagum gametangial stichidia is not analogous developmentally to the furrow in G. johnstonii tetrasporangial stichidia.  The mature cystocarp of Gelidium johnstonii  (Fig. 107) has stretched but intact  third order filaments (Fig. 108), as in G. purpurascens,  and  differs from  G.  vagum where third order filaments are broken. Significantly though, G. johnstonii is monoecious, male and female gametes appearing in the same position as in G. vagum (Fig. 111).  Taxonomy and Nomenclature / 51 Despite the morphological similarity of smaller thalli of Gelidium vagum and G. johnstonii,  and that both species are unique in being moneocious, these taxa are  regarded  as  separate  species  because  of several  consistent  and  significant  character differences (Table 6). Since they are both moneocious, a condition that is  otherwise  unknown  in  Gelidiales,  and  because  of  the  uniformity  of  spermatangial position with respect to carpogonia, it is reasonable to propose that G. johnstonii  and G. vagum are closely related sister species.  A similar pattern of warm-temperate to sub-tropical, East Pacific—West Pacific disjunct  distributions  Okamura, Tinocladia Prionitis and  has  been  observed for Pachydictyon  crassa (Suringar) Kylin, Endarachne  coriaceum  binghamiae  (Holmes)  J. Agardh,  cornea (Okamura) Dawson, Ishige sinicola (Setchell et Gardner) Chihara  Lomentaria  catenata  Harvey, and  for the  species  pairs  Eisenia  arborea  Areschoug and E. bicyclis (Kjellmann) Setchell, and Carpopeltis divaricata Okamura and  Binghamia  California  J. G.  Agardh  (Hommersand,  1972). In addition,  numerous other species pairs have been suggested on the basis of preliminary observations (Hommersand, 1972), and Gelidium johnstonii  and G. tenue Okamura  (1934: 56) are found as a pair of taxa. The description and illustrations of G. tenue (Okamura, 1934, pis. 23, 31 fig. 8-10) are not strikingly reminescent of G. vagum, and a similarity between G. tenue and G. vagum has not been noted in discussions  of Japanese  Santelices, unpubl.  or Chinese species  mscr.). No  evidence has  of Gelidium  (Akatsuka,  1986b;  been presented that supports a sister  taxa relationship between G. tenue and G. vagum.  The combination of the 1) narrow range of distribution in the Pacific Northwest,  Taxonomy and Nomenclature / 52 restricted to British Columbia of Gelidium  vagum (see below), and 2) its great  abundance and vigour within its distribution, suggests that this species may have been a recent introduction rather than a relict of a population of a once wider distributional range in the northwest Pacific. Many oyster farms are present in the  Hornby Island-Denman Island area. As such, the importation of Japanese  oyster spat is a likely vehicle for the introduction of G. vagum into British Columbia. Scagel (pers. comm.) has suggested that another species, Lomentaria hakodatensis, of the British Columbian algal flora may have also been introduced accidental^, probably in oyster spat. In view of the rapid colonization of Hornby Island and Denman  Island, future range expansion of G. vagum in British  Columbia might be expected. To date no other west coast of North America populations are known, although a single sterile specimen of G. vagum (UBC A64965) (Fig. 110) appeared in the drift at Ladysmith, farther south in the Strait of Georgia.  e. Seasonality and  Gelidium  Distribution  vagum is perennial in British Columbia. Fertile tetrasporophytes  and  cystocarpic plants have been collected in the fall (August, September, October), whereas plants collected in April were infertile. Japanese specimens (TNS; UBC) were tetrasporangial in May and July and gametangial in Jul}'.  In British Columbia, Gelidium vagum is restricted to Hornby Island and Denman Island (Strait of Georgia).  Taxonomy and Nomenclature / 53 In addition to the Strait of Georgia in the area of Hornby Island and Denman Island  in British  Columbia, G. vagum is distributed  in the Huanghai Sea  (Bangmei et al, 1983), and the Yellow Sea of China (Akatsuka, 1987; Santelices, unpubl. mscr.), the Korea Strait and Sea of Japan, (Tokida, 1954; Akatsuka, 1987; Kajimura, 1987), and the Pacific coasts of Honshu and southern Hokkaido (Okamura, 1934; Akatsuka, 1987).  4. Pterocladia  caloglossoides (Howe) Dawson  1953: 76  Basionym: Gelidium caloglossoides Howe 1914: 96, fig. 7, pis. 34, 35 Synonym: Pterocladia parva Dawson 1953: 77 fig. 2, pl. 6, n.v. Holotype: Coker 59 (NY); lost on loan (Thiers, pers. comm. 1987), if no other specimens are found, a neotype should be made. Type locality: Island of San Lorenzo, Peru  a. Habitat and Habit  Pterocladia  caloglossoides  grows  in the shallow  subtidal,  1-7 m  depths, in  moderately exposed to protected locations. It is epilithic on tops and sides of boulders, on platforms or on walls, epiphytic on crustose corallines and also occurs on calcareous shells. Plants are soft, but not lax, and are dark red, drying to blackish red. They form a dense, low turf of upright axes that arise from an extensive system of prostrate axes (Fig. 112). Many uprights develop along the length of a single prostrate axis. Although the most vigorous plants  Taxonomy and Nomenclature / 54 may reach 2 cm tall, the turf usually is not more than 1 cm tall.  Branching in Pterocladia caloglossoides ranges from very sparse to abundant; the number of branch orders is low; uprights may  be unbranched or may  bear a  maximum of two branch orders, though second order branches were uncommon in the  study area. Branches are distichous, irregularly subopposite, and the branch  angle is almost 90° (Fig. 113). Axes are compressed  to flattened at branch  bases, especially where branches are regenerating from cut ends. Upright axes vary greatly in width, and a single branch may  range from 240-690 Mm. Axes  taper gradually at their bases and abruptlj' at their apices to an acute tip.  Prostrate  and  upright  axes  are  similar,  except  that  prostrate  axes  have  attachment pads at frequent intervals whereas pads are lacking on upright axes. Occasionally becoming  an  upright  axis  bends  down, developing attachment  and  prostrate. Branching of prostrate axes is subopposite distichous, but  when producing uprights, one to five axes may Uprights  pads  are  perpendicular  attachment pad (230-260 Mm  to  prostrate  arise from  branches  and  a single node.  occur  opposite  an  (Figs. 114, 116). Prostrate axes are less variable in width  diameter) than upright axes and are approximately the same width  as narrower uprights. Prostrate axes do not taper at branch points, but have acute apices.  Attachment of prostrate axes to the substratum is by numerous attachment pads formed medially on the ventral surface of creeping axes opposite the upright branches (Fig. 116) and formed on the lower surface of an upright branch that  Taxonomy and Nomenclature / 55 is becoming prostrate. Cortical cells elongate, or produce elongate, unpigmented, rhizoid-like lobes that adhere to form attachment pads (Figs. 120, 121). Pads are to 1.0 mm  long and are 0.2-0.5 mm  wide and flare out distally, as the distal  end of cells incorporated in the pad are swollen and bulbous.  b. Vegetative Anatomy  Thallus construction is uniaxial with a conspicuous, domed apical cell and two periaxial cells derived from each subapical cell (Fig. 115). Second order filaments grow laterally outward at a 45-60° angle to the axial filament. These are evident just behind apices where cells of second order filaments are in diagonal rows from the axial filament (Fig. 115).  Transverse  sections of Pterocladia caloglossoides reveal a cortex of only 1-2 cell  layers (Fig. 117) and a medulla. The outer wall of superficial cortical cells is as wide as the protoplast diameter. Cortical cells in upright axes are pigmented, oval or short cylindrical, 7-10 um long and 5-10 um wide, and in surface view are equidistant, but unordered and irregularty arranged  (Fig. 119). Cortical cells  of creeping axes are slightly larger, but are otherwise  like cortical cells of  upright axes. Outer cortical cells lack secondary pit-connections, whereas the inner cortical  cells  (in specimens  where  two rows  are present)  have secondary  pit-connections.  The  medulla is 3-5 cells thick and in upright axes composed of unpigmented,  cylindrical  cells, 12-27 urn long  and 7-13 nm  wide, that  have abundant  Taxonomy and Nomenclature / 56 secondary pit-connections (Figs. 117, 118). Medullary cells of prostrate axes are slightly larger. Cells of the axial filament are conspicuous in longitudinal section (Fig. 118). Rhizines, present in the medulla  (Fig. 117), are very abundant in  upright axes but are more sparse in creeping axes. They are initiated from the inner proximal corner of medullary or inner cortical cells, at a position 6-7 axial cells behind the apex.  c. Reproductive Anatomy  Tetrasporophytes are the only reproductive thalli collected from the study area. Gametophytes  are  unknown  in British  Columbia,  northern  Washington  and  southeast Alaska.  Tetrasporangia form only on main axes and  short lateral branches of upright  axes (Fig. 113), occuring in sori but are not in stichidia; the branch apex grows vegetativery beyond the sorus. Short, fertile, first order branches narrow rapidty at the base, and taper abruptly to an acute apex, similar in shape of similarly positioned vegetative branchlets. One  of the most striking features of Pterocladia  caloglossoides is the characteristic "V" arrangement of tetrasporangia in the sorus (Fig. 122). The more proximal Vs radiating laterally at 45°  are oldest. Sporangia occur in pairs of lines  to the axial filament. The  margin of the flattened  branch is sterile, with tetrasporangia borne on both faces of branches which are recognizable approximately five axial cells behind the apex. They are cut off from medullary cells adjacent to the second order plate (Fig. 123) and have a lateral  pit connection  (Fig. 124)  to  their  bearing  cell. Tetrasporangia  are  Taxonomy and Nomenclature / 57 cruciately divided (Fig. 125), sometimes irregularly, and 20-45 um 15-40  Aim wide. Only rarely do young  released  sporangia,  indicating  sporangia  that percurrent  long and  form between cavities of  production  of tetrasporangia is  uncommon in P. caloglossoid.es.  d. Discussion  Pterocladia  caloglossoides originally was described by Howe (1914) as Gelidium  caloglossoides from the Island of San Lorenzo, Peru. It was found on shells dredged from 2.5 fathoms (5 m depth) by Dr. Robert E. Coker. The holotype (Coker 59 in NY) was lost while on loan (Thiers, pers. comm. 1987) and it is unknown if isotypes exist. I have written to AHFH, BM, FH, MICH, NY and US, but their curators have not located isotypes of G. caloglossoides. Coker 59 was the only material mentioned by Howe in the diagnosis. Questions may be raised of the connection between the taxon Howe described as G. caloglossoides from Peru and Dawson's identification of material from Pacific Mexico as P. caloglossoides  that  was  used  in comparison  with  British  Columbian  P.  caloglossoides. In Howe's original diagnosis, G. caloglossoides was characterized by it's creeping habit, small size, radiate clusters of 2-5 upright branches opposite attachment pegs, flattened axes, surface cells in distinct oblique rows at branch apices,  narrow  Gametophytic  medulla,  plants  either  and were  tetrasporangia not found  similarity of G. caloglossoides to G. pusillum was  in "distinct  oblique  or described. Howe  lines".  noted the  but believed that G. caloglossoides  sufficiently distinct on a number of criteria to warrant recognition as a  separate species.  Taxonomy and Nomenclature / 58 Hollenberg (1942) reported Gelidium  caloglossoides  from southern California and  Monterey, central California., which was included by Smith (1944) in the marine flora of the Monterey Peninsula. Diagnostic features used to recognize this taxon were its prostrate habit, small size, flattening of the branches, branch pattern and arrangement of tetrasporangia in Vs. Smith (1944) also noted that in the Monterey  specimens the upright axes have few orders of branching and that  branches are almost perpendicular to main axes. Dawson (1953) transferred G. caloglossoides to Pterocladia based on unilocular cystocarps found in material from Pacific Baja California del Norte, observing that the tetrasporangial plants from Baja California were the same as the material described from Peru by Howe (1914).  I  have compared  British  Columbian  Pterocladia  caloglossoides  with Dawson's  material from Pacific Mexico (AHFH Dawson 8593, liquid-preserved and specimens on microscope slides 1176, 1177 from Guadalupe Island, off Baja California, and AHFH Dawson 8733 liquid-preserved 'and microscope slides 1325, 1326,  1327  from Barra de Navidad, Jalisco on the mainland of Mexico). British Columbian P. caloglossoides resembles to both these collections, although it is most similar to the Guadalupe Is. plants. Specimens from Guadalupe Is. were a maximum of 1 cm  tall, with unbranched  uprights, the tallest of which were undamaged by  grazing or abrasion. Liquid-preserved Guadalupe 1176  and  three  of four  tetrasporangia clearly in Vs  plants  on  slide  (Fig. 126). On  Is. specimens, those on slide  1177  are tetrasporophytes, with  the creeping axes are numerous  attachment pads opposite from which two to four upright branches arise (Fig. 114). These  Pacific  Mexico  plants fit easily  within  the description  of P.  Taxonomy and Nomenclature / 59 caloglossoides  given and illustrated by Howe (1914). Baja California plants are  less well developed than those from Peru, but British Columbian  plants from  Orlebar Point, Gabriola Island, where frequent collections were made throughout the year, showed much variation in the development of upright axes from their almost complete absence to unbranched axes, and to well branched axes. British Columbian P. caloglossoides also corresponds well with all other illustrations and descriptions given for central California, and Pacific Mexico and Gulf of California P. caloglossoides  (Smith, 1944; Stewart, 1976; Stewart and Norris, 1981), and  thus is determined to occur in the study area.  One plant of Dawson's slide 1177 from Guadalupe Is. (Fig. 127) is cystocarpic. Its morphology matches that of tetrasporophytes, and it is reasonable to assume that the specimen is representative of Pterocladia  caloglossoides.  From what can  be seen from a whole mount of a cj^stocarpic plant (giving a "top" view of a cystocarp), an ostiole appeared on only one face of the cystocarp (Fig. 129), indicating a single locule. Sections of a cystocarp from liquid-preserved material (AHFH Dawson 3733) show a single locule (Fig. 128). Thus the species as known from Pacific Mexico does indeed belong in Pterocladia, and Dawson (1953) was  justified  in transferring Gelidium  caloglossoides  admits that absolute confirmation of placement obtained interesting  by  observing cystocarps in type  that  Dawson  later  used  to Pterocladia,  in Pterocladia  though he  could  only be  localitj' Peruvian material. It is  the combination  G.  caloglossoides  when  describing specimens from the Peruvian flora (Dawson et al., 1964).  Added evidence for the generic placement of this taxon in Pterocladia comes from  Taxonomy and Nomenclature / 60 apical and lateral initial morphology observed in British Columbian material. Both types of initials are domed hemispherical and protruding, and neither occur in cortical depressions. This finding is in agreement with Rodriguez and Santelices' (unpubl. mscr.) proposal that apical and lateral initials are similar in shape in Pterocladia  and dissimilar in Gelidium,  but it contradicts their statement that  either the apical or lateral initials are depressed Pterocladia. other  in cortical indentations in  The "indentation of initials" character while apparently reliable for  taxa  of Pterocladia  was  not corroborated  in British  Columbia  P.  caloglossoides.  Pterocladia caloglossoides as identified by E. Y. Dawson from Barra de Navidad, Jalisco, Mexico (AHFH Dawson 8733) are heavily damaged from abrasion or grazing, with truncated uprights that only reach a maximum of 0.5 mm Some of the truncated axes have regenerated  tall.  apical cells, but little regrowth  occurred before the plants were collected and preserved. Two plants (slide 1327, AHFH) have many attachment pads and several uprights opposite the pads. Sections of vegetative plants (Fig. 132; slide 1326, AHFH) show a cortex and medulla that is well developed  compared to British Columbian P. caloglossoides.  Dawson's material does correspond to the illustrations and description given by Howe for Gelidium branching  caloglossoides,  particularly  in the distinctive  features of  of the prostrate axis, attachment pad position and arrangement of  tetrasporangia in Vs. Thus I agree with  Dawson  (1953) that the Mexican  material with a single cystocarp locule is the same as G. caloglossoides (Howe, 1914) and it belongs in Pterocladia.  Taxonomy and Nomenclature / 61 Dawson (1953) noted that "antheridia" were unknown in Pterocladia caloglossoides, however, specimens of his (slide 1325, A H F H ) are a whole mount of one female and two male thalli. The fertile apices of these gametophtyes (Fig. are  129), whereas in clusters  replaces  a  vegetative  apices are more acute and tapering.  of two to four  vegetative  are blunt rounded  cortical  in sori behind cell  (Fig.  apices  131),  and  Spermatangia  (Fig. 130).  Each  spermatangia  cluster  are  cut  off  transversely and singly from spermatangial mother cells (Fig. 133).  When  Dawson  described  a  new  gametophyte strongly  made  the  species,  new  combination  P. parva  Dawson  Pterocladia (1953:  77)  caloglossoides, based  1  he  on only  also female  material. Dawson's illustration (1953, pi. 6 fig. 2) of P. parva  resembles  P.  caloglossoides,  an  observation  supported  by  is  Dawson's  comment that the illustration of P. caloglossoides in Smith (1944, pi. 44 figs. 3, 4) is more  similar to P. parva  than to P. caloglossoides. Pterocladia parva  later  was condsidered a synonym of P. caloglossoides by Stewart and Norris (1981).  The  similarity  Stewart  (1976)  turf-like.  In  of Gelidium and  pusillum  Stewart  California,  and  however,  characteristic of the genus Gelidium. reservation  (Stewart,  to  Pterocladia  Norris (1981), G.  caloglossoides because  pusillum  has  The name G. pusillum  both  was are  noted small  bilocular  G. sinicola, P. parva  California  material,  although  and P. caloglossoides have  more  definite  thorough comparison with type material of G.  determinations  and  cystocarps,  was used with some  1976) for material from California. Other species names,  latifolium,  by  been  applied to  necessarily  pusillum.  h o l o t y p e : Dawson 425 on sheet 4181, plus slides 1178-1181 in A H F H Type locality: San Felipe, Baja Californiadel Norte, Gulf of California  await  G. this a  Taxonomy and Nomenclature / 62 e. Seasonality and Distribution  Pterocladia  caloglossoides is perennial, persisting for most of the year as prostrate  axes with few  uprights that become  more abundant and form  a turf in the  summer months, reaching their maximum development in August and September. Tetrasporangial sori are most abundant at this time. A collection was made in February  1987  at  Georgina Point, Mayne  Island (Fig. 3),  however,  in which  tetrasporangial sori were present but sparse.  Pterocladia  caloglossoides is relatively common and widespread in the study area,  but it likely has been overlooked and not reported from many sites due to its small size and creeping habit. This species has been collected from Orcas and San Juan Islands in northern Puget Sound (Fig. 3), Mayne and Gabriola Islands in the Gulf of Georgia (Fig. 3) and Hornby and Denman Islands in the Strait of Georgia (Fig. 4), Sooke (Juan de Fuca Strait), Barkley Sound (Fig. 2) and Nootka Island, on the west coast of Vancouver Island, and the Queen Charlotte Islands (Fig. 1).  This study extends the northern limit of distribution of Pterocladia caloglossoides from northern Washington (Norris and Hollenberg, 1969) to British Columbia (e.g. UBC  A12295,  UBC A19667,  UBC A28937,  UBC A37605,  UBC A64648,  for  others see Appendix 1) and southeast Alaska (UBC A69423, UBC A69424, UBC A69425, A69430,  UBC A69426, UBC A69431).  (Santelices,  1977), the  UBC A69427, Pterocladia  UBC A69428,  caloglossoides also  Great Barrier Reef,  Australia  is  UBC A69529, reported from  (Cribb,  UBC Hawaii  1983), the Indian  Taxonomy and Nomenclature / 63 Ocean south of Sumatra (Weber van Bosse,  1921), and Natal, South Africa  (Norris, 1987).  D.  PREVIOUSLY  REPORTED  TAXA  EXCLUDED  FROM  THE  REVISED  FLORA  Gelidium  crinale, G. pusillum,  G. robustum and G. sinicola have been reported in  the local flora (Widdowson, 1974; Scagel et al., 1986), but these are based on taxonomic misidentifications as none of these taxa actually occur in the study area.  With the  exception  of G. robustum,  these taxa  are  generally poorly  understood and their species limits poorly defined.  In the first two decades of this century, Gelidium amansii (type locality "in mari Indico  ad ins.  Franciae, Madagascar, Indiae  orientalis",  Kiitzing,  1849)  was  reported from southern British Columbia and northern Washington (Setchell and Gardner, 1903; Collins, 1913; Kylin, 1925; Connell, 1928), and reported plants were likened to P. B.-A., No. 585 (distributed as G. amansii,  Collins et al.,  1903, but in fact are now recognized to be G. robustum collected in California) by Setchell  and Gardner (1903). Collins (1913) noted that British Columbian  plants based on collections by Macoun from Ucluelet, Departure Bay and Victoria and by Tilden from Port Renfrew (Minnesota Seaside Station) he referred to G. amansii were much smaller than Californian specimens, Without exact collection data confirmation of Collins' detemination cannot be made but UBC and CANA have Macoun and Tilden collections from these locations (Macoun: UBC A1952, CANA 3843, CANA 4346, CANA 4349 (Fig. 135); Tilden: UBC A5195). All of  Taxonomy and Nomenclature / 64 these specimens were determined to be G. purpurascens. amansii early  Kylin (1925) included G.  in the Friday Harbour flora but had not seen any specimens. These records  of  G.  amansii  have  been  treated  by  later  authors as  misidentifications (Scagel, 1957).  There is a single report of Gelidium  pulchrum  from the area of Nanaimo  (Stephenson and Stephenson, 1961b). Vouchers of this record do not exist and the Nanaimo area has been recollected without finding any specimens that are this taxon. This record has been included as misidentified G. purpurascens  by  Scagel et al. (1986) in the most recent floristic treatment of British Columbia, northern Washington and southeast Alaska.  In the following discussion, I treat those taxa more recently reported for the local  flora  but  that  are, in  fact,  absent, based  mostly  on  taxonomic  misidentifications.  1.  Gelidium robustum (Gardner) Hollenberg et Abbott  1965: 1179  Basionym: Gelidium cartilagineum var. robustum Gardner 1927a: 280, pl. 54 Holotype: UC 294572! Type locality: near Ensenada, Baja California del Norte, Pacific Mexico  Gelidium robustum is a well defined species ranging from central Baja California, Mexico (Stewart, 1976) to central California (Silva, pers. comm.). First  was  Taxonomy and Nomenclature / 65 described by Gardner (1927a) as a variety of G. cartilagineum  (L.) Gaillon, it  was later elevated to species rank by Hollenberg and Abbott (1965). Based on an examination of the holotype from near Ensenada, Baja California del Norte (UC 294572, Fig. 138) and additional representative material (UC 395419 from San  Pedro, California  collected  by  Gardner; UC  California collected by Dawson; and UC 756503, and  UC  940173 from  collected by Dawson; and UC  647822, from  756464, UC  756469, UC  White Pt., 756470, UC  various localities in Baja California, Mexico  1451987 from Portuguese Bend, California collected  by Loomis), it was concluded that none of the British Columbian plants (with the possibility of one exception) correspond to this taxon.  There is an interesting specimen in UBC  (UBC A7861) of a plant that is tall,  robust, tetrasporangial and clearly is Gelidium  robustum (Fig. 136). It poses  somewhat of a mystery. The annotation label states the collection locality as "Shoal B. Victoria" from 1917 by an unknown collector. There is no "Shoal B." near Victoria, British Columbia  (Canadian Coast Pilot) although it is unclear  whether the reference is to Shoal "B", or whether B is an abbreviation for bay, bight, beach etc.. However, there are no such locales near Victoria, B.C., but there are some in the San Juan Islands a few km  to the east of Victoria  (Canadian Coast Pilot). A considerable amount of seaweed collecting was done in southern Vancouver  Island as early as 1917, and it is unlikely that if G.  robustum existed at Victoria in 1917, such a large, conspicuous plant would have been collected only once and never at an3' time between central California and British Columbia. Admittedly, a small population could have been in Victoria (perhaps introduced from California in the early 1900's), that has subsequently  Taxonomy and Nomenclature / 66 disappeared. It is also possible that  collection  data  were confused between  samples from a variety of locations along the coast. As it stands, even if G. robustum had been present in British Columbia, it does not exist here now, and G. robustum must be excluded from the flora.  In the study area, several specimens of Gelidium purpurascens as G. robustum.  Differences between  were misidentified  these taxa already have been discussed  (Table 4). Several of the literature records alluding to the presence of G. robustum in the study area lacked voucher specimens or illustrations, and those records could not be investigated. None of the reports for which there are vouchers  identified  as "G. robustum"  were  G. robustum,  in fact  but are  misidentifications. Scagel (1957) cited five specimens from Departure Bay and Victoria  on Vancouver  3473 = CAN  Island  209, CANA  (as G. cartilagineum  3740 = CAN  74, CANA  var. robustum) 3843 = CAN  (CANA  353, CANA  4346 = CAN 208, CANA 4349 = CAN 310), and all were re-examined and found to be G. purpurascens,  and Scagel's (1967) illustration is of G. purpurascens.  South's (1968) records of "G. robustum" of taxa: specimens Pterocladia  from  Orlebar  from Gabriola Island were of a variety  Point  (UBC A28937, UBC  A29575) are  caloglossoides and G. coulteri respectively, specimens from Lock Bay  (UBC A36.199) and False Narrows (UBC A36198) are G. coulteri and specimens from  Twin  purpurascens  Beaches  (UBC A28645, UBC  A28646) are G. coulteri  respectively. All records of "G. robustum"  from  Barkley  (Scagel, 1973) that could be located were found to be G. purpurascens.  and G. Sound  Taxonomy and Nomenclature / 67  2. Gelidium crinale  (Turner) L a m o u r o u x  1825: 191  Basionym: Fucus crinalis Turner 1819: 4 Synonyms: Acrocarpus crinalis (Turner) Kutzing 1868: 11, pl. 33 figs, a-c, n.v. Acrocarpus spinescens Kutzing 1868: 12, pl. 33 figs, d-e, n.v. Acrocarpus corymbosus Kutzing 1868: 13, pl. 36 figs, a-c, n.v. Lectotype: Specimen annotated by Setchell as "This is Turner's idea of G. crinale and may  be taken as type. f. W.A.S." (from Turner Herbarium in  BM) (Dixon and Irvine, 1977b) Isotype: AHFH 55234, n.v. Type locality: Kilmouth, England  Descriptions of Gelidium  crinale (Gardner, 1927a; Dawson, 1944, 1953; Taylor,  1957; Santelices, 1977) demonstrate the lack of well defined diagnostic characters for this species. Santelices (unpubl. mscr.) especially about the species limits and  identifications  has noted the confusion  of this taxon, and the resulting  misapplications of the name. Collins (1913) first recorded G. crinale in the study area from Victoria, B.C. and cited P. B.-A. No. 195 (Collins, et al., 1896, Fasc. IV) from Port Jefferson, Long Island, New  York, G. crinale  as a reference  specimen, but this specimen is different from any species present in British Columbia or northern Washington. Collins' record for G. crinale is based on a Macoun collection from Victoria (Collins, 1913), which is probably CANA 3474 (formerly CAN 1908  207) (Figs. 134, 137) from Beacon Hill, Victoria, collected June  and labelled  as G. crinale.  Re-examination of the plants on this sheet  Taxonomy and Nomenclature / 68 revealed them to be G. coulteri.  The inclusion of Gelidium crinale in the British Columbia flora has continued with the  repeated citation of CANA  Departure Bay, B.C. (as V  3474  (as CAN  207); for the record from  1456) (Scagel, 1957), the specimen could not be  located. A specimen (UBC A66778) collected by Scagel from Departure Bay as "G. crinale" probablj' corresponds to the missing V  1456 and is stamped "on  loan from the B.C. Provincial Museum". This specimen is now recognized to be G. purpurascens. crinale" UBC  In UBC  there are numerous specimens misidentified  that were determined to be G. coulteri  A37402, UBC  A53975, UBC  A60254, UBC  A64934), and one that is Pterocladia caloglossoides  (UBC A31363, UBC A60268, UBC  as "G. A31426,  A60460, UBC  (UBC A19667). Many other  specimens originally labelled as "G. crinale" have been annotated as other species, e.g. G.  coulteri,  G.  purpurascens,  P.  caloglossoides,  reflecting  the  taxonomic  confusion of G. crinale, and the assumption of its presence in the local flora, time of the collection.  Gelidium crinale first was recorded from California by Collins (1903 in Collins et al., P. B.-A., Fasc. XXIII, No. 1138), and it was the basis for a new variety, G. crinale var. luxurians  Collins (1906: 111). Later authors continued to record  the presence of G. crinale var. luxurians  from California (Gardner, 1927a) and  Baja California del Norte, Mexico (Dawson, 1953). Dawson (1944, 1953, 1966) listed both G. crinale (var. crinale) and G. crinale var. luxurians Mexico and the Gulf of California.  from Pacific  Taxonomy and Nomenclature / 69 Stewart (1974) considered the Californian Gelidium  crinale var. luxurians  (i.e. P.  B.-A., Fasc. XXIII, No. 1138; and the material mentioned in Gardner, 1927a) to be Pterocladia media Dawson (1958: 68). The remaining specimens of "G. crinale var. luxurians"  Dawson and G. crinale var. crinale sensu Dawson (1944, 1953)  were considered G. pusillum  by Stewart and Norris (1981). Thus G. crinale var.  crinale and G. crinale var. luxurians  do not occur in the Gulf of California or in  the northeast Pacific.  3. Gelidium pusillum  (Stackhouse) Le Jolis  1863: 139  Basionym: Fucus pusillus  Stackhouse 1801: 6  Synonyms: Fucus caespitosus Stackhouse 1801, pi. 12 n.v. ACrocarpus pusillus  (Stackhouse) Kutzing 1849: 762 n.v.  Lectotype: in BM (Dixon and Irvine, 1977b) Type locality: Sidmouth, England  There is no evidence to support the presence of Gelidium pusillum  in the study  area. Of the reports of "G. pusillum"  and  from British  Columbia  northern  Washington (given in Scagel et al., 1986), only Norris and Wynne (1968) cited voucher  specimens  herbarium pusillum  and Garbary  et al. (1984) did not specify  which UBC  specimens they used. Norris and Wynne (1968) first reported G.  in the study area from San Juan Island, Washington, but the specimen  they reported (WTU, Norris 5723) could not be located. Collections other than those of Norris and Wynne (1968) of gelidioid plants from San Juan Is. belong  Taxonomy and Nomenclature / 70 to G. purpurascens  (UBC A4402, UBC A4403; FHL 2849; private herbarium of  W.R. Waaland #1583) and to Pterocladia caloglossoides (WTU 248018). The Scagel et al. (1986) report of "G. pusillum"  from Barkley Sound, based on a specimen  (UBC A61245), is now identified as P. caloglossoides. Other herbarium specimens from the study area labelled as "G. pusillum"  are mis-identifications, and actually  G. coulteri (UBC A24837, UBC A28351, FHL 3055) and P. caloglossoides (UBC A33645, UBC A64648).  Gelidium  pusillum  circumscribed  has a worldwide distribution and, like G. crinale,  is poorly  (Santelices, unpubl. mscr.). Different workers have used varying  species concepts that have incorporated numerous other species (Dixon and Irvine, 1977a; Stewart, 1976; Stewart and Norris, 1981), or they have recognized many varieties (Dawson, 1944, 1953; Santelices, 1977; Schnetter and Bula Meyer, 1982).  The name  seems  to have  been  applied  commonly  to any small,  turf-forming, compressed to flattened gelidiaceous plants. This taxon clearly needs revision on a global scale, beginning with a thorough study of the type material.  4. Gelidium  sinicola  Gardner  1927a: 278  Holotype: (Gardner 2615) as UC 276620! Type locality: Point Cavallo, San Francisco Bay, California  Records of "Gelidium  sinicola" in the study area result from confusion as to the  definition of the taxon. Gelidium  sinicola  Gardner  (1927a) was described as  Taxonomy and Nomenclature / 71 "sparse" (i.e. not abundant) and "of limited distribution", and based only on the type locality collection from Point Cavallo, San Francisco Bay, California (Gardner 2615 as UC 276620) (Fig. 139). Even Gardner was apparently confused as to the identity of this species. His later collections from the same tidepool as the type collection (Gardner 7179, UC 494898) (Fig. 21), were noted in his field book "G. sinicola  ? at least from the same pool as type came" (Silva, pers.  comm. 1985), and on the herbarium "Gelidium".  sheet he identified the plant only as  Added later, but not in Gardner's handwriting (possibly by E. Y.  Dawson, fide Silva, pers. comm. 1985), is "coulteri Harv. f.".  Both the type specimen and Gardner 7179 have a morphology within the limits of Gelidium  coulteri. Gelidium  that it is cylindrical rather than flattened (Gardner,  1927a).  Stewart  (1976) noted  that could fit  sinicola differs from G. coulteri in  (Fig. 140), and it is narrower  that  G. coulteri  varies  widely in  compression of branches, and I also have seen much variation of axis width and compression in G. coulteri from the study area. Silva (pers. comm., 1985) has visited the type locality of G. sinicola  several times and has made extensive  collections around San Francisco Bay, and writes of seeing plants he considers to be morphological^' variable G. coulteri, but never any cylindrical plants that could correspond to G. sinicola.  Because the type material is infertile, there are few  characters for species comparison. Based on a comparison of height, apical and basal axis diameter, branching pattern and rhizine abundance and position, in Pterocladia media, G. sinicola (holotype and Gardner's diagnosis), and Californian G. crinale var. luxurians,  Stewart (1974) suggested that G. sinicola might be a  synonym of P. media. Generic placement cannot be confirmed for G. sinicola  Taxonomy and Nomenclature / 72 because no cystocarpic plants are known to exist.  Despite the problems of the actual identity of Gelidium  sinicola,  records of it in  the study area are based on misidentifications of other gelidioids. Specimens in UBC  labelled originally as "G. sinicola" belong to G. coulteri (UBC  A14403) or Pterocladia  caloglossoides  reports of "G. sinicola"  in the study area (given in Scagel et al.,  (UBC  A29576, UBC  A1449, UBC  A39127). Of the 1986), only  Norris and West (1966) cite vouchers. The voucher cited by Norris and West (Norris 5087) could not be located. Thus G. sinicola does not appear to occur in the study area.  C H A P T E R IV. LIFE HISTORY OF GELIDIUM IN  A.  CULTURE  INTRODUCTION  Until recently, the life history had not been completed successfully in culture for anj'  species of Gelidiales. This is somewhat surprising considering the economic  potential and interest in the group. Most commercial uses of Gelidiaceae have been restricted  to wild harvest. It was  assumed from  field collections that  Gelidiaceae have a Polysiphonia-type life history (Fig. 141) (Dixon, 1961, 196.3) with a sequence of (haploid) gametophyte, (diploid) carposporophyte  and (diploid)  tetrasporophyte  phases, where  phases  free-living  isomorphic,  and  and  gametophyte the  and  tetrasporophyte  carposporophyte  develops  on  are  the female  gametophyte. In order to confirm this, I attempted to culture the local Gelidium species. According to recent studies on the life histories of G. coulteri (Macler and  West, 1987)  and  G.  vagum (van der Meer, pers. comm.), both species  require a short time to complete the life history (two months for G. coulteri, Macler and West, 1987).  In the current study it was phases of Gelidium distribution  (see  noted that both gametophytic and tetrasporophytic  vagum are abundant enough within the species' restricted Chapter  3)  to  suggest  a  Polysiphonia-type  life  history.  Gametophytes of the other local species, however, are rare or absent in the study area, suggesting that a Polysiphonia-type  life history may  occur in the studj' area, even though the species may  not commonty  demonstrate it under  different environmental conditions or geographical areas. Instead, populations 73  may  Life History of Gelidium in Culture / 74 be propagating or persisting vegetatively, undergoing  an asexual life history or  repeating tetrasporophytes. In the study area, gametophytes of G. purpurascens rarely are found and have been collected from only a few restricted sites. Male gametophytes are difficult to differentiate in the field, although nature  of spermatangial  the stichidial  branchlets imposes an additional order of branching  making male plants detectable with some practice. It would be expected that if cystocarpic plants are collected, male gametophytes also might be present. Despite the  paucity of G. purpurascens  gametophytes, the species is abundant and  widespread in the study area, and in late summer and early fall tetrasporangial plants are common. This raises the question of the kind of life history that predominates  in most  G. purpurascens  populations  in British  Columbia and  northern Washington. Are tetraspores viable, and do they produce gametophytes and  ultimate^ cycle sexually, or do populations cycle asexually or persist  vegetatively?  Gametophytes of G. coulteri never  have been collected  although they are known from farther south  in the study  area,  (e.g. California, Dawson, 1953).  Tetrasporophytes are not often fertile, observations suggest a relatively restricted time period (August to November), and fertile plants are probably overlooked. Detection is more difficult when tetrasporophytes are only weakly fertile because tetrasporophytic purpurascens  branchlets of G.  and  G.  vagum.  coulteri Detection  are not as distinctive of  tetrasporangia  on  as in G. Pterocladia  caloglossoides plants is difficult because the small size of these plants (never taller than 2 cm) makes low magnification necessary for detection of fertility. When P. caloglossoides tetrasporophytes are fertile, tetrasporangia are abundant,  Life History of Gelidium in Culture / 75  and most upright axes bear fertile sori.  B.  METHODS  1. Culturing Cystocarpic and tetrasporic Gelidium vagum was and  Galleon  Point,  Hornby  Island  (Fig. 4)  collected from Denman Island in  late  September,  1986.  Tetrasporangial G. purpurascens was collected from Geer Islets and Dixon Island, both on the west coast of Vancouver Island (Fig. 2) in August 1984, Georgina Point, Mayne Island (Fig. 3) in November, 1985 and February, 1986, Orlebar Point, Gabriola Island (Fig. 3) in October and December, 1985, August and October, 1986, and from Whalebone Bay, Gabriola Island (Fig. 3) in October and November, 1986. Gelidium coulteri tetrasporophytes  were collected from Denman  Island in September, 1986 and from Orlebar Point, Gabriola Island in October, 1986. Tetrasporangial Pterocladia caloglossoides was collected from Georgina Point, Mayne Island, Orlebar Point, Gabriola Island and Galleon Point, Hornby Island in September, 1986.  Field collected plants were brushed to remove detritus and as many epiphytes as possible before  culturing. Fertile  tetrasporangial or cystocarpic stichidia were  excised and placed into 2X6 cm plastic petri plates containing culture medium. In the case of infertile plants, apices were excised and placed into petri dishes. For each isolate and set of growth conditions at least two dishes were prepared. The  culture  medium  used  was  half strength,  modified  Provasoli's  enriched  Life History of Gelidium in Culture / 76 seawater medium (PES) (McLachlan, 1973). Initially, full strength PES but it later was  found that plants grew as well and with fewer contamination  problems in 1/2 strength PES. This was  a particular benefit in the first stages  of isolation when unialgal cultures were being seawater medium (SWM, was  was used,  tried. Growth was  started. McLachlan's enriched  McLachlan, 1973), without soil and liver extracts, also similar to that of 1/2 PES, but algal contaminants also  seemed to thrive. Enrichment nutrients were filter sterilized with a 0.45  um  millipore filter prior to mixing.  Two  walk-in  environmental growth chambers, one  photoperiod) and 16.42 at  jiEm" s~ 2  20.15  uEm~ s~ 2  1  and  one  at 20°C, 16:8  at 10°C,  8:16  (light:dark  photoperiod  and  were used, along with a smaller Percival 1-35-L incubator set  1  24°C, 12:12  photoperiod and  7.91  uEm" s" . Each isolate was 2  1  cultured  under all three sets of conditions. Growth conditions were maintained as constant as  possible but  were subject to chamber breakdowns and  disturbances  (e.g.  opening doors during dark periods). Several growth conditions were utilized to improve the chance of completion of a gelidioid life history. Conditions were not intended  to  environmental  test responses factors,  as  of  plant  growth  temperature,  and  daylength  reproduction and  to  irradiance  differing varied  simultaneously between growth conditions. As the chambers were a group facility, they were set at temperatures and daylengths that were most amenable to all users, Gelidium  conditions  that  would  not  necessarily  have been  chosen  for  optimal  growth or life history studies. Aeration did not promote tetraspore or  gamete production and made it harder to control contamination.  Life History of Gelidium in Culture / 77 After incubation of spore producing tissue for 2-5 days (depending on chamber conditions), released spores and germinated sporelings were removed with a finely drawn out pipette operated by light suction and placed into petri plates of fresh medium. The  medium  was  changed every  one  contamination and, after the first one to two  to two  weeks depending on  months, once every two months.  For isolates used in life history studies the medium was changed monthly.  Tetraspores or carpospores released from isolates were removed to new and the above process was the dishes  were stirred  medium,  repeated. When monoecious isolates produced gametes, and  swirled each day  to encourage fertilization. In  dioecious cultures, fertile male and female gametophytes were placed together into a dish and stirred and swirled daify.  Tetraspores  of  Gelidium  purpurascens  and  G.  vagum,  and  carpospores of  G.  vagum were released readily from plants under the culture conditions. Germination was  most successful from the tetraspore and carpospore-bearing plants that were  collected in September. At other times, spores released readily but most failed to germinate. Gelidium  coulteri tetraspores released and  germinated less readily so  that few sporelings were obtained. Several techniques with fertile tetrasporophytes of P.  caloglossoides such as drying the plants prior to immersion for release,  cutting the spore-bearing branches and putting cheese-cloth covers over dishes to reduce light intensity, were unsuccessful at releasing tetraspores. For all species apical fragments also were cultured.  Life History of 2.  Chromosome  Gelidium  in Culture / 78  Counts  Chromosome counts were attempted  Gelidium vagum  on nuclei of undivided tetrasporangia of  collected from Hornby Island. Squashes were made by first  softening excised tetrasporangial stichidia in 4% KOH  for 10 minutes. Tissue was  stained with Wittmann's hematoxylin (Wittmann, 1965) for one hour, the cover slip added and tissue partly squashed. New  stain was  added at intervals to  prevent drying out. Material was destained by drawing 45% acetic acid under the coverslip for 30-60 seconds, then rinsed with distilled water and mounted  in 40%  Karo,  and  tissue  was  firmly  squashed  permanently  to spread cells  thoroughly.  C.  SPORE  GERMINATION  AND  EARLY  DEVELOPMENT  The pattern of spore germination, and early development of the sporeling to the stage of apical cell organization, are similar in and  G. vagum  Gelidium coulteri, G. purpurascens  for both tetraspores and carpospores. The following account is of  spore germination events in  G. vagum.  The released spore is spherical with a large, stellate chloroplast and prominent nucleus (Fig. 142). It germinates by producing a small hyaline protuberance (Fig. 143), which expands (Fig. 144) into a germ tube, slightly wider and longer than the spore. The spore contents evacuate into this tube, and a wall forms cutting off the cytoplasm from the spore (Fig. 145). In most cases the evacuated spore appears completely empty, but occasionally granular debris (Figs. 146, 147) or  Life History of Gelidium in Culture / 79 parts of the chloroplast remain. This pattern of spore germination, referred to as "Gelidium-type"  germination  (Chemin,  1937),  is characteristic of the order  Gelidiales (Papenfuss, 1966; Santelices, 1974). In some spores a germ tube did not form, and the cytoplasm divided several times within the spore wall. Later germination stages were not seen, and it is unlikely that the sporeling survived.  The sporeling divides longitudinally and unequally with a curved wall (Fig. 147), to form "concave" and "fusiform" daughter cells (after Boillot, 1963). The larger, concave daughter cell, divides several times transversely and longitudinally (to the axis established by the germ tube and first division), forming 10-14 isodiametric cells (Figs. 148,  149, 150). The  most distal cell of this concave cell-derived  group, opposite the spore, elongates, forming a primary attachment rhizoid (Figs. 151, 152). The rhizoid is unpigmented, thick-walled and most of the cytoplasm is contained in the tip. Adjacent distal cells may  subsequently similarly elongate,  forming additional rhizoids (Fig. 153) that attach the sporeling to the substratum. Similar to other workers (e.g. Macler and West, 1987) I have found that if attached sporelings are removed from the substratum they do not reattach later. Up to four primary rhizoids may  form on a single sporeling.  The smaller, fusiform daughter cell of the young sporeling divides several times transversely and longitudinally to form small, isodiametric cells (Figs. 154, 155). Two  longitudinally oriented clusters of cells comprise the sporeling, each cluster  originating from the concave or fusiform daughter cell and separated by the first division plane of the germ tube (Figs. 155, 156). When the entire sporeling is approximately  20-40 cells  large,  the  most  proximal  cell  of the  fusiform  Life History of Gelidium in Culture / 80 cell-derived group cuts off a shallow, hemispherical apical cell (Figs. 157, adjacent  to the  original spore wall. Up  158),  to this point in development, the  sporeling increases very little in size beyond that of the germ tube.  The uniaxial construction typical for the genus develops following sporeling apical cell formation. The  sporeling increases in size, growing in the opposite direction  to the primary rhizoid and past the original spore wall (Fig. 160). Organization is uniaxial and shows a characteristic pattern. The domed, hemispherical, apical cell divides transversely (Figs. 159,  162). The  subapical cell divides obliquely  twice to form two lateral, opposite periaxial cells. Each periaxial cell cuts off a lateral second order filament that grows outwards contributing to thallus width (Figs. 68, 159). Periaxial cells also cut off third order filaments above and below the lateral plane that determine the thickness of the thallus. As the sporeling increases in length, branch initials are formed from modified cortical cells (Fig. 159), and attachment pads develop from the outward elongation and adhesion of cortical cells (Fig. 161).  D.  REPRODUCTION  1.  Culture  IN  CULTURED  AND  FIELD  PLANTS  Results  Reproductive structures appeared on cultured plants of G. were one to two  cm  vagum, when they  tall, approximately 5-6 weeks after spore germination in  isolates grown at 20°C. Cultured tetrasporophytes, with the same morphology as tetrasporophytes  from the field, produced tetrasporangia on stichidia (Figs. 163,  Life History of Gelidium in Culture / 81 166). Plants grew faster at 20 °C and high light (long day and high irradiance) and  were narrower and less flattened than field collected plants. At 24°C and  low  irradiance, plants  Tetrasporophytes  were  narrow  grown at 10 °C  and  weak  grew more  and  slowly  only  sparsely  and took much  fertile. longer  (several months to one year longer) to become fertile, if they did so at all. Plants grown at 10°C more often appear similar to field material, with wider, flatter branches than  plants  grown  at 20 °C.  Tetrasporangial  stichidia were  determinate, cylindrical, with rounded or acute apices and produced tetrasporangia around the entire stichidium in an unordered manner. Tetraspores were released and germinated into gametophytes (Fig. 167).  Gelidium  vagum gametophytes produced carpogonia and spermatangia on short  branchlets (Fig. 164), as in field collected material, with an axis producing a succession  of fertile  branchlets  so that  progressively  older  carpogonia and  post-fertilization stages were observed in more proximal parts of the plant (Fig. 165). Developmental sequences of male and female gametes and carposporophytes were  similar to those  observed  in field collected gametophytes. Fertilization  occurred in culture, although not as extensively as in field material, evidenced by the lower frequency of cystocarps per cultured plant. Greater water motion in the  field  than  in the occasional^'  swirled,  unaerated  cultures  may  bring  non-motile spermatia in contact with trichogynes more effectively. After three to five weeks, carpospores were released from bilocular cystocarps. Carpospores were obpyriform large,  when in the cystocarp, but rounded-up on release. They contained a  central, stellate  previously described.  chloroplast  and  germinated  into  tetrasporophytes,  as  Life History of Gelidium in Culture / 82 Gelidium  vagum isolates originally  initiated  produced fertile gametophytes and released ' carpospores,  and  from  tetraspores and  tetrasporophytes  tetraspores, and  gametophytes. These tetrasporophytes  carpospores,  (respectively), that in turn  germinated into tetrasporophytes  and  (Fig. 168) and gametophytes became fertile  completing the life history (simultaneously from two different starting points).  Life histories could not be completed in culture for the other local gelidiaceous species, but some insights into the life histories of Gelidium purpurascens and  G.  coulteri in the study area were provided by culturing. Tetrasporangial stichidia of Pterocladia  caloglossoides were placed into culture several times, but tetraspores  were not released under anj' of the experimental  culture conditions, and  the  vegetative fragments grew well at 10°C and 20°C but did not become fertile.  Cultures  of  purpurascens  G.  field-collected tetrasporophytes  were  started  (Fig. 169). The  from  the cultures grown at 20 °C.  released  from  resulting gametophyte sporelings  were dioecious; both carpogonial and spermatangial in  tetraspores  plants were readily identifiable  Fertile male gametophytes developed in four  isolates (Geer Islets, Dixon Island, Majme Island and Whalebone Bay on Gabriola Island, Fig. 170). Female gametophj'tes were identified only in Whalebone Bay cultures (Fig. 171). Carpogonia and  spermatangia in cultured plants appeared  mature but evidence of fertilization was  not observed, possibly due to inadequate  water motion in culture dishes, so that non-motile spermatia could not contact trichogynes. incompatability  Alternatively, or  fertilization  inviability,  fertilization undetectable.  or  early  might  have  carposporophyte  occurred abortion  but could  gamete make  Life History of Gelidium in Culture / 83 Tetraspores did not release as readily from stichidia of Gelidium germination was  coulteri, and  less successful than for G. purpurascens and G. vagum, but a  few isolates of gametophytes were maintained. Fertile gametophytes (found only in 20°C cultures) were male (Figs. 19, 172,  173); female gametophytes were  undetected (under all growth conditions), and it is unknown if they existed in the cultures, or if all non-fertile plants from tetraspores were male gametophytes or simply non-reproductive.  Undivided tetrasporangia were used for chromosome counts rather than other cell types (e.g. apical cells, spermatangia  or spermatia) because they had the largest  nuclei and most visible chromosomes. Even so, nuclei were still small (3-7ym diameter), and  it was  difficult to obtain the resolution necessary for reliable  chromosome counts. Counts were made only on nuclei squashed flat enough so that chromosomes were in a single focal plane. Counts of 14 bivalents and a single, additional small body were made on the best view of a dividing nucleus (Figs. 174, 175). The bivalents consisted of closely paired chromosomes at the most contracted stage of prophase I. It was unclear whether the small body was a separate chromosome, small bivalent or an extraneous body. If it was a small chromosome, then it should be noted that there are size differences between some chromosomes.  Counts of 14 and at least 12 and 13 (Fig. 176) bivalents were obtained from other dividing tetrasporangial nuclei. Nuclei in which counts bivalents  were  obtained  contained  a  large  deeply  staining  of 12  and  13  body probablj'  comprising two or more bivalents. Thus the chromosome number obtained from  Life History of Gelidium in Culture / 84 all of these counts is n= 14-15. In some nuclei, five large bodies were observed (Fig. 177).  2. Field  Observations  of  Reproduction  Tetrasporophytes and gametophytes of Gelidium vagum are equally abundant in the field when reproductive plants were found (August to October). Gametophytes of G. purpurascens  were rarely collected in the study area. Of 18 sites in the  study area from which I collected G. purpurascens,  female gametophytes were  found at only two sites (Kirby Point, Diana Island and Geer Islets, both in Barkley Sound). Six specimens of G. purpurascens  female gametophytes were  found from a total of several hundred records of this species (UBC A41842, Fleming Island, Barkley Sound; UBC UBC  A39126, Lawn Point, Vancouver Island;  A41586, Wizard Island, Barkley Sound; UBC  A36876, Amos Island, west  coast of Vancouver Island; UBC A41946, Clarke Island, Barklej' Sound; and UBC A46026, Bamfield, Barkley Sound; UBC  A67446, Kirby Point, Diana Island).  Male gametophytes of G. purpurascens rarely are collected from the study area, and the only specimens known are three records in UBC Sound, Vancouver Island; UBC  (UBC A53657, Nootka  A10813, Esteban Point, Vancouver Island; and  UBC A41415, Tzartus Island, Barkley Sound).  Tetrasporophytes  of Gelidium  gametophytes in my purpurascens  own  purpurascens  are much  more  abundant  than  collections and in herbarium specimens. Whereas G.  gametophytes were found only in the southern half of the study  area, while tetrasporophytes were found fertile throughout British Columbia and  Life History of Gelidium to its northern range limit in southeast Alaska (UBC  in Culture / 85  A69432). Where female  gametophytes occur, they bear many fully developed cystocarps, suggesting that successful fertilization occurs in the field. Tetrasporangia are abundant and appear deeply pigmented.  Fertile gametophytes of Gelidium  coulteri are unknown in the study area, but  occur farther south in California. Although tetrasporangial specimens have been collected in British Columbia, they are not often abundantly fertile. Gametophytes also are unknown for Pterocladia  caloglossoides  from the study area, but are  reported for this species from California (Dawson, 1953). Pterocladia  caloglossoides  is tetrasporophytic throughout British Columbia, but infertile in southeast Alaska collections (UBC A69427, UBC  E.  1.  A69423, UBC  A69428, UBC  A69424, UBC  A69429, UBC  A69425, UBC  A69426,  UBC  A69430, UBC A69431).  DISCUSSION  S p o r e G e r m n ia o t i n  Spore germination of Gelidium. vagum, G. purpurascens and  G. coulteri is uniform  for all tj'pes of spores and the species. In this study, it was differentiate species or spore types using any  impossible to  aspect of spore morphology or  germination pattern. The similarity between tetraspore and carpospore germination also was noted by Yamasaki (1960). Furthermore, spore germination in Gelidium (Chemin, 1937; Katada, 1949, 1966;  Guzman-del  Proo  et  al.,  1955; Yamasaki, 1960; Boillot, 1963; Kaneko, 1972;  this  study), Pterocladia,  Acanthopeltis  Life History of Gelidium in Culture / 86 (Katada, 1955) and Gelidiella  (Chihara and Kamura, 1963; Sreenivasa Rao, 1971)  showed the same general pattern.  Differences in reports of spore germination are minor and, in at least some instances, appear to result from the interpretation of observations. For several hours after release, and before attachment, carpospores of some Gelidium species reportedly showed amoeboid motion (Boillot, 1963; Guzman-del Proo et al., 1972). Chemin (1937) in particular was fascinated by carpospore  mobility, reporting  paths and rates for spores of several red algal species. Germination observations of the local species of Gelidium  were started  after spore  attachment, thus  amoeboid motion, if present, was not observed in the spores. Several authors observed one to three (mitotic) nuclear divisions before spore evacuation in G. vagum latifolium,  (Kaneko,  1966), Gelidiella  G. pulchellum  acerosa  and G. pusillum  (Sreenivasa  Rao, 1971),  Gelidium  (Boillot, 1963), but onlj' a single  nucleus entered the germ tube.  Empty spore germination (i.e. evacuation of the entire spore cytoplasm into the germ tube) occurs in other higher rhodophyte groups, but only in Gelidiales is it followed by direct development of the upright plants, without an intervening discoid phase. Chemin (1937) designated all species with emptj' spore germination, as having "Gelidium-type"  germination regardless of subsequent developments, and  Boillot (1963) later demonstrated the unique nature of this pattern in Gelidium.  The first division of the germ tube after being cut off from the spore, is highly significant, as the plane of this division and the distinctive concave and fusiform  Life History of Gelidium in Culture / 87 daughter cells  determine the polarity  of the sporeling. Observations of the  sporeling from different orientations can be misleading. For example, Sreenivasa Rao (1971) noted that the first division of the germ tube in Gelidiella acerosa is sometimes transverse. This interpretation probably resulted from a face view of the fusiform or concave cell (analogous to a valve view of a diatom), where the curved wall separating them is not seen, and the first divisions of fusiform and concave cells is transverse. There appears to be general agreement  that the  concave cell is the larger of the two daughter cells and gives rise to the primary rhizoid (Boillot, 1963; Chihara and Kamura, 1963; this work). However, there is confusion in the literature as to the developmental role played by the fusiform and concave cells. In Gelidiella acerosa, Sreenivasa Rao (1971) reported that the rhizoid originated from the fusiform cell group, whereas Chihara and Kamura (1963) observed the opposite, a concave cell group origin of the rhizoid. It is most likely that the concave cell produces the primary rhizoid in Gelidiella as in the other members of the order. In a series of manipulations of the environment of germinating spores, Katada  (1949, 1955) determined that the  specific gravity of the water affected the length and number of rhizoids produced, but the pattern of development was invariant.  Interpretation  of the origin  of the apical  cell  also  has  caused confusion.  Observations made here indicate that the fusiform cell group is the site of apical cell production. Specifically, the cell of the fusiform cell group closest to the spore wall, differentiates as the apical  cell. This  illustrated or stated by some workers in Gelidium japonicum,  G. pacificum,  G. pusillum,  agrees with the pattern amansii,  G. divaricatum,  Pterocladia. tenuis and Acanthopeltis  G.  japonica  Life History of Gelidium in Culture / 88 (Katada, 1955), G. amansii (Yamasaki, 1960), and Gelidiella acerosa (Chihara and Kamura, 1963) but is contrary to observations of Boillot (1963). Boillot (1963) reported the formation of two apical initials in Gelidium latifolium, and G. pusillum, the  G.  pulchellum'  one contributed by the fusiform cell group and another from  concave cell group. It was  unclear which initial takes over, but Boillot  believed that it is the apical cell from the concave cell group. Yamasaki (1960) illustrated a developmental sequence like that observed in the local  Gelidium  species, but he referred indistinctly to "upper" and "lower" daughter cells. He stated that the upper daughter cell gave rise to the creeping part of the plant, and the lower daughter cell produced the primary rhizoid and upright or creeping branches. This is unlike  any  other gelidioid  developmental pattern; it maj'  correspond to the dual apical cells mentioned by Boillot. According to Yamasaki's (1960) illustrations, however, spore development patterns appear similar to those of other gelidioids. Some authors do not indicate the origin of the apical cell, and their illustrations are sometimes too confusing to determine where this occurs (e.g. Kaneko, 1966; Sreenivasa Rao, 1971).  In conclusion, possibly the most notable feature of spore germination and early sporeling development in Gelidiales is the uniformity of the developmental pattern in all species, genera and spore types studied to date, and the distinctiveness this pattern with respect to other red algal groups.  Life History of Gelidium  in Culture / 89  2. Life Histories  The completion of the Gelidium  vagum life history in culture shows that the  British Columbian plants have a Polysiphonia-type life history and, based on the common occurrence of tetrasporophytes and gametophytes in the field, this seems to be an accurate description of the life history in the study area.  The scarcity with which Gelidium  purpurascens gametophytes are collected in the  study area suggests that gametophytes either fail to survive in the field (for example, if environmental conditions are unsuitable for spore or sporeling survival or full development) or gametophytes are commonfy produced but rarely become fertile (and hence are not recognized as gametophytes). Low abundance of fertile male  and female  gametophytes in the field  would  decrease  the chance of  fertilization, accounting for the rarity of cystocarpic G. purpurascens in the study area. In British Columbia, G. purpurascens is probably capable of completing the Polysiphonia-type life history, but factors such as low tetraspore viability and low abundance of gametophytes or unfavourable environmental conditions could prevent the common occurrence of this sexual life history in situ. Occasional periods of favourable environmental Polysiphonia-type  life  conditions could permit the completion of a sexual  history  and range  expansion, whereas more commonly,  populations persist vegetatively or reproduce asexually.  The British Columbian populations of Gelidium purpurascens persist by perennating as upright fronds or creeping axes and may environmental  conditions are suitable  recruit new  for gametophytic  individuals when  sporeling  survival or  Life History of Gelidium in Culture / 90 gametophyte fertility. Drift plants of G. purpurascens appear to be whole plants torn from the substratum rather than fragments of plants. Drift plants and attached plants in the field do not exhibit the abundant production of attachment pads that can be seen in cultured plant fragments. These observations suggest that propagation operating  by vegetative fragmentation  here, but  further  field  confirmation. Although there was  does not appear be a mechanism  experimentation  would  be  neccessary  for  no evidence in cultures, tetraspores also could  be recycling apomictically as alternate dispersal agents. However, further work would be  required  to determine if such  Columbia G. purpurascens. simply  maintaining  Information  a  process  is operating  in British  on whether populations are expanding or  their current extent would also be  needed to assess the  existence and importance of the above mechanisms.  The  occurrence of fertile male Gelidium  coulteri plants in 20°C culture  significant because, although none were recorded herbarium  records  or my  own  from  the  study area  was (from  collections), their presence indicates that the  capacity for the production of male gametophytes exists. Male plants could be overlooked  in the field or they could be absent if environmental conditions are  not suitable for their development. A seawater temperature of 20°C is uncommon in British Columbia but is occasionally reached, particularly in sheltered locations. Gelidium  coulteri probably propagates vegetative^ by creeping axes and possibly  apomictically by  tetraspores in British Columbia, at the northern  limit of its  distribution.  In  culture, Pterocladia  caloglossoides failed  to release (or germinate in  situ)  Life History of Gelidium in Culture / 91 tetraspores,  although  they  were  abundant  on  stichidia.  It is unknown if  tetraspores function at all as propagules (meiotically or apomictically). Clearly vegetative propagation  and perennation  by creeping axes must be important in  the maintenance or expansion of this species' local populations.  Gelidium purpurascens, G. coulteri and Pterocladia  caloglossoides display the pattern  of latitudinal variation in life history expression proposed by Dixon, (1965), where progressively more northern populations of a species show reduced fertility of gametophytes and, even more northerly populations show reduced tetrasporophyte fertility. This trend ultimately results in only vegetative propagation northern  range limits. Gelidium  purpurascens  variesin  part from  at species' this pattern  because some of the most northerly collections of this species, from southeast Alaska, are tetrasporangial (UBC A69432).  3. Chromosome Counts  There have been few  chromosome counts made on  any  (1966) observed n=7-10 in tetraspore germlings of G. obtained  n= 4  and  5 for cells of the cortex and  Gelidiaceae. Kaneko vagum,  Dixon (1954)  nutritive tissue of female  gametophytes of G. corneum, and Kaneko (1968) counted n=15  for tetrasporangia  of Acanthopeltis japonica. A demonstration of the uncertainty often associated with such counts is offered in the observations made on G. latifolium. obtained "4, 5, 9 and obtained n=18  10 chromosomes" for carposporophytes,  Dixon (1954)  but Boillot (1963)  for young gametophyte sporelings, and Magne (1964) counted n or  2n = 25-30 for vegetative cells of G. latifolium var. luxurians.  It is possible that  Life History of Gelidium in Culture / 92 some varieties could represent polyploid groups (e.g. is G. latifolium var. luxurians a polyploid of G. latifolium  var. latifolium!)  or that polyploids could be included  with diploids in the same, or other, species. Counts of n= 14-15 obtained from undivided tetrasporangia of G. vagum in this study, are closest to counts made by  Boillot (1963) for G. latifolium  japonica. not  and by Kaneko (1968) for Acanthopeltis  The G. vagum counts of n= 14-15 made here are not similar to and do  appear to be multiples of the only  other  counts made for G. vagum  (n = 7-10) by Kaneko (1966). The importance of good preparations and accurate information about cell type and generation counted must be stressed. Too few reliable studies have been made and corroborated to confidently state chromosome numbers of species, genera or base numbers for the family.  C H A P T E R V. CARPOGONIUM AND  A.  As  CARPOSPOROPHYTE D E V E L O P M E N T  INTRODUCTION  with  spermatiogenesis  and  tetrasporogenesis, there have been few detailed  studies on pre- and post- fertilization development in Gelidium. investigations were those of Dixon (1959) on G. latifolium Fan  (1961) on G.  robustum (as G. cartilagineum  and Fredericq (1988) on G. pteridifolium. Gelidiales, Gelidium,  Pterocladia,  Suhria,  The most recent  and  G.  var. robustum) and  pulchellum, Hommersand  Additional selected species of genera of Beckerella,  and  Acanthopeltis,  have been  observed for comparative purposes (Fan, 1961; Hommersand and Fredericq, 1988) or briefly touched on in other studies (Kraft, 1976). The interpretations made by Hommersand  and  Fredericq  (1988) were significantly  different  from  previous  authors, most notably as to the nature of the carpogonium, and their findings were corroborated in the current investigation. In view of the paucity of studies on these developmental processes, additional descriptive observations based on local Gelidium species were warranted.  The  lack of studies of pre- and  post- fertilization development in members of  Gelidiales is surprising given the healthy debate that has taken place historically over the ordinal recognition of the group. Gelidiales now universally at the ordinal level by reproductive Gelidium-type  and  are recognized almost  a suite of characters other than female  carposporophyte characters, such as  a single pit plug cap,  spore germination, the presence of rhizines, the production of two  distichous periaxial cells, and the transverse division of the spermatangial mother 93  Carpogonium and Carposporophyte Development / 94 cell in forming  a  spermatangium  (see  Chapter  1, herein; Santelices,  1974;  Gabrielson and Garbary, 1986, 1987; Hommersand and Fredericq, 1988).  Kylin  (1923) elevated  the  Gelidiaceae,  a  family  in  the  Nemaliales  (as  "Nemalionales"), to ordinal status based on the unchanged, fertilized carpogonium that was  the starting point for the gonimoblast, and  auxiliary cells that are  present but function as nurse cells in nutrition of the carposporophyte. Kylin's (1923) system of ordinal classification, was female  gametophytic  and  based entirely on features of the  carposporophytic  reproductive  apparatus,  namely  characters of the carpogonial branch, auxiliary cells, connecting filaments and the gonimoblast. It must be noted that contrary to Dixon's (1961) belief that life history  differences  (Kylin, 1923  believed  Nemaliales  were  haplobiontic  and  Gelidiales diplobiontic) were also a criterion in the elevation of Gelidiaceae to an order, Kylin (1923) discussed these differences but stated that the life history character should not be used systematically at that time. Both Nemaliales and Gelidiales lacked a "typical" auxiliarj' cell, i.e. any  cell that the carpogonium  fused with and from which the gonimoblast developed (Kylin, 1928, thus  were  set  Rhodymeniales and  apart  from  the  other  orders  1956), and  (Cryptonemiales, Gigartinales,  Ceramilales) which possessed a "typical" auxiliary cell. Kylin  (1928) studied pre- and post-fertilization events in detail in Gelidiaceae, and the family was  important in his formulation of the distinction between "generative"  ("typical") and "nutritive" auxiliary cells. Even after his addition of the criterion that a typical auxiliary cell also not be a carpogonial branch cell (Kylin, 1935, 1937), he still maintained that a "typical" auxiliary cell was  absent in Gelidiales.  Kylin (1928) believed that the gonimoblast in Gelidiales developed directly from  Carpogonium and Carposporophyte Development / 95 the unchanged, fertilized carpogonium. When this was found to be erroneous and that the carpogonium  underwent non-obligate fusions with adjacent cells (Dixon,  1959), the presence of generative auxiliary cells became open to interpretation and dependent on the definition of the auxiliary cell. Drew (1954) gave the most useful definition, as a cell of specified position in the thallus with which the carpogonium fuses prior to gonimoblast formation. Accordingly, Gelidiales lack an auxiliary cell. Details of the auxiliary cell debate are given by Santelices (1974) and Hommersand and Fredericq (1988). The ordinal position of Gelidiales has been questioned extensively (Dixon, 1959, 1961; Papenfuss, 1966), being largely dependent Dixon's  on the perceived presence or absence of an auxiliary cell and, in (1961) view,  the lack  of life  history  differences,  i.e.  haplobiontic  Nemaliales (as Nemalionales) and diplobiontic Gelidiales. Dixon (1961) argued for returning  Gelidiales to Nemaliales as Kylin's  (1923,  1928, 1956) characters  separating the orders were not valid. On the other hand, Papenfuss (1966) argued that Gelidiales was supported at ordinal rank by the presence of unique chains or nutritive auxiliary cells, by the presence of only two periaxial cells, and by the unique pattern of spore germination. It is important that carpogonium and carposporophyte development be studied in more gelidioids to assess variation at the genus and species levels and to determine features of the developmental processes that are common to all members of the order. The present study contributes to a more complete understanding of carpogonium and carposporophyte development in the order.  Carpogonium and Carposporophyte Development / 96  B. METHODS  Of the local gelidiaceous species, fertile (female) gametophytes of only vagum  and  G.  purpurascens  occurred  carposporophyte development followed  in the study  area.  Gelidium  Carpogonium and  the same pattern in both species. The  process also was observed, in less detail, in sections from herbarium specimens of  G. robustum  (UC 395419, 647822, 756470, 940173; UBC A62199).  Observations were made on dried and Formalin preserved material following the light microscope methods described in Chapter 3. Hematoxylin stained material also was used, prepared as in Chapter 4. The only modification for the purpose at  hand (as opposed to chromosome counts) was to make light squashes of  apices and cystocarps, enough to spread out clusters offilaments,but not enough to break the continuity of filaments. Hand made razor blade sections were stained with hematoxylin for 1 h, destained with 45% acetic acid for 15-30 s, and rinsed with water.  C. CARPOGONIUM  DEVELOPMENT  The  Gelidium  carpogonium of  IN GELIDIUM  differentiates from the second to basal cell of a  third order filament that is a normal part of the vegetative thallus. In the vegetative  condition, the second  to basal cell of a third  order filament is  intercalary with three pit-connections: one to the basal cell of the filament, one to the third cell of the filament, and one to another distal cell basal in a fourth order filament. When differentiated as a carpogonium, the second to basal cell of  Carpogonium and Carposporophyte Development / 97 a third order filament bears only two pit-connections (Fig. 182), one to the basal cell and  one to the third cell of the third order filament. The  vegetative filament and any  fourth order  of its higher order branches, is replaced by the  gradually tapering, arched trichogyne of the carpogonium. Carpogonia are produced sequentially along the axis in the fertile area (Fig. 47). Two  rows of carpogonia  are produced lateral to the axial row, on third order filaments, extending to both surfaces of the compressed to flattened axis. Thus four rows of carpogonia are formed, with  two  rows visible  in either  a  longitudinal or saggital section.  Replacement of a cluster of vegetative cortical cells by  the trichogyne of each  carpogonium results in the production of a medial gap or furrow on both thallus surfaces (Fig. 89). Trichogjmes protrude to the thallus surface through this gap.  The  carpogonium is initially intercalary. Farther back in the fertile zone are  sessile carpogonia, cut off from the supporting cell by a distinctive concave wall (Fig.  179). These carpogonia  supporting cell, and  have  a  single pit-connection to an  intercalary  a second to basal cell in a third order filament. The  question arises whether intercalary carpogonia are immature, and require a final division to become mature and  sessile, or whether intercalary carpogonia are  mature and functional, and undergo a division that discards the trichogyne after being past receptivity to spermatia. Many sessile carpogonia were observed in a degenerating  condition, whereas  the  supporting  cell  appeared  healthy (i.e.  cytoplasm dense as in adjacent cells) (Fig. 180). An intercalary carpogonium, just fertilized,  was  observed  cortical cells through  at an early stage of fusion cell initiation, fusing to  two  expanded pit-connections (Fig. 181). Thus functional  carpogonia are intercalary in Gelidium.  Carpogonium and Carposporophyte Development / 98 D.  CARPOSPOROPHYTE  AND  CYSTOCARP  DEVELOPMENT  Following fertilization, the carpogonium expands and fuses with several adjacent cells  through  widened  pit-connections,  forming  a  large,  irregularly  lobed,  multinucleate fusion cell (Figs. 182, 183). Occasionally two fusion cells can be seen in a single apex, suggesting that two carpogonia  may be fertilized and  develop a genetically heterogenous carposporophyte in a single cystocarp. It is difficult to be certain of this, however, as fusion cells may have very long, narrow lobes. When the fusion cells are widely spaced it seems likely that the carposporophytes are discrete.  Concomitant with fertilization, short chains of small, isodiametric cells, referred to as "nutritive filaments" (Hommersand and Fredericq, 1988), are cut off from the bases of cells of third order filaments (Figs. 183, 184). In Gelidium fusion cells always are present in apices where nutritive suggesting that nutritive filaments are initiated nutritive  filament formation  may  be slightly  vagum,  filaments are seen,  at fertilization. The timing of earlier  in G. purpurascens, as  nutritive filaments are present in apices that appeared to lack a fusion cell. Nutritive filaments curve in, towards and around the plate of second order cells (Fig. 183). They reach a maximum length of six cells, the apical cell of which is slightty larger and round, other cells being short and cylindrical (Fig. 184). All nutritive filament cells have prominent nuclei.  Fusion cell lobes cut off weakly staining, uninucleate gonimoblast cells, which form branching  chains winding around and between cells of the second order  Carpogonium and Carposporophyte Development / 99 plate (Fig. 185). Gonimoblast cells cut off elongated processes that contact and fuse with apical cells of nutritive filaments (Fig. 186). This fusion is between the diploid gonimoblast (carposporophyte) cells and haploid nutritive filament (female gametophyte) cells.  The gonimoblast produces single carposporangia laterally and terminally (Fig. 188), which project into one of the two locules created between the second order plate and cortex at cystocarp expansion (Fig. 187). Uninucleate carposporangia expand to become ovate to obpyriform with a darkly staining cytoplasm and stellate chloroplast. Carposporangia are produced continually by the gonimoblast, and a variety of ages is seen in a cystocarp. The growing carposporophyte pushes the cortex away from the second order plate, creating locules and causing third order filament cells to stretch. In Gelidium  vagum  these filaments break  as the  cystocarp matures (Fig. 187), whereas they remain intact in G. purpurascens (Fig. 50). The cystocarp cortex does not increase in thickness over the vegetative cortex, although some cells stretch laterally.  At  maturity the cystocarp has expanded beyond the width of the vegetative  branch. Locules are filled with carposporangia and faintly staining mucilage (Fig. 187).  Nutritive filaments no  longer  are distinguishable,  and  the entire  carposporophyte (except the sporangia) appears vacuolate, staining poorly (with aniline blue). One ostiole per locule forms in the cystocarp cortex, due to the failure of part of the cortex to fill in. It is a simple, round opening and is not beaked or protruding. There is no evidence that the ostiole forms from tearing of the cortex. Carpospores  are released, leaving behind  some  mucilage  in the  Carpogonium and Carposporophyte Development / 100 cystocarp. There is no evidence of percurrent production by carposporangia.  E. DISCUSSION  The  most significant observation made concerns the intercalary carpogonium in  Gelidium.  Dixon (1959) and Fan (1961) both noticed intercalary carpogonia but  their interpretations differed. Intercalary carpogonia were interpreted developmental  stages by Fan  and  as early  as aberrant gametes by Dixon. They both  agreed that intercalary carpogonia are non-functional. They saw sessile carpogonia as mature, but no basis for this assumption was provided, on the other hand, Hommersand and Fredericq (1988) regarded intercalary carpogonia as functional and  sessile carpogonia as non-functional; while the}' examined the material no  illustrations were provided.  In this study on Gelidium,  an intercalary carpogonium was observed fusing with  cortical cells through two widened pit-connections at a verj' ear^' stage of fusion cell formation. Sessile carpogonia never formed fusion cells and often were seen in stages of degeneration, suggesting a non-functional condition. Reports of sessile carpogonia fusing with the supporting cell following fertilization (Dixon, 1959; Fan, 1961), stem carpogonium  from  observations of fusion  is visible, and  cells  where  the  outline  of the  where the fusion cell narrows somewhat before  expanding at what is believed to be the supporting cell. Earl}' stages of this supposed  fusion  purpurascens,  are  however,  not  reported  intercalary  or  illustrated.  carpogonia  are  In  G.  expanded  vagum at  and the  G. base,  particular!}' near pit-connections. Hommersand and Fredericq (1988) reported the  Carpogonium and Carposporophyte Development / 101 retention of part of the carpogonium outline in the fusion cell, and this could explain  the  earlier reports of Fan  (1961) and  Dixon  (1959), if the basal  carpogonium outline is retained as well as the trichogyne lobe. It should be noted that fusion cells in G. vagum and G. purpurascens showed no carpogonial outline, but this observation probably carpogonia of Gelidium  depends on fusion cell age. Thus the functional  are intercalary, as noted by Hommersand and Fredericq  (1988). If unfertilized, the carpogonium can cut off the trichogyne lobe, and the basal portion can then revert to a vegetative cortical cell.  An  is reported in female branchlets of Gelidium  apical notch  1961) and G. pteridifolium British Columbia  robustum (Fan,  (Hommersand and Fredericq, 1988), but is absent in  G. purpurascens  and  G.  vagum. Hommersand  and Fredericq  (1988) speculated that retarded growth of the apical cell and axial row, evidenced by overgrowth of the apical cell by adjacent cortical lobes, was  responsible for  development of the axial furrow. Growth of third and higher order filaments near the axial row  was  outpaced by equivalent filaments lateral and distal to the  axial row. Since there does not appear to be retarded axial row development in species lacking an apical notch, the axial furrow was  interpreted as originating  from the absence of cortical cells that belonged to filaments replaced by  the  carpogonium.  Timing of the formation of nutritive filaments varied between studies and be a taxonomically useful character. In Gelidium pteridifolium are  initiated  as  the  carpogonium  differentiates,  (Hommersand and Fredericq, 1988). In G. latifolium,  but  may  nutritive filaments before  fertilization  and G. pulchellum  (Dixon,  Carpogonium and Carposporophyte Development / 102 1959), G. robustum (Fan, 1961) and G. vagum (herein), fertilization is probably the stimulus for nutritive filament production. It is difficult to be certain from reports by Dixon  and  Fan, as both believed the functional carpogonium  intercalary. Thus when Dixon after the carpogonium  was  (1959) observed  that nutritive filaments formed  mature, the observation corresponds  forming after the carpogonium became sessile and functional  carpogonia  probably  were being  was  to filaments  non-functional, while other  fertilized. Fan  (1961) noted that  nutritive filaments form during carpogonium development, but the exact timing is not known with respect to functional carpogonia or fusion cell formation.  While  there  is general  agreement  that, upon  fertilization,  the carpogonium  enlarges and becomes lobed, there is no agreement on the nature of fusions with surrounding cortical cells. Fan (1961) reported no fusions between the fusion cell and cortical cells, whereas Dixon (1959) accepted that fusions may  occur but are  not obligate. In this study and in all fusion cells seen by Hommersand and Fredericq (1988) there were fusions with cortical cells. My findings agree with Hommersand and Fredericq (1988) that fusions of the carpogonium with cortical cells are a regular feature of earty carposporophyte development in Gelidium, that the fusions may  but  not be with cells in a specified position with respect to the  carpogonium.  Fusions regularly occur between cells of nutritive filaments and gonimoblast cells. Nutritive filament cells clearly function as nutritive auxiliary initially  dense  cytoplasm  becomes  sparse  and  vacuolate  cells, as their  after  fusions  and  gonimoblast growth. I suspect that the reported lack of fusions between nutritive  Carpogonium and Carposporophyte Development / 103 filaments and gonimoblast  (Dixon, 1959) resulted simply from failure to observe  them and not their absence. Confirmation should be made, as substantiation of Dixon's (1959) observation could be important systematically.  Gelidium  latifolium  and  G. pulchellum  differ from G. purpurascens  in carposporangial initiation. In the former two  and  G. vagum  species, clusters of uninucleate  initials are cut off from gonimoblast cells that develop into carposporangia (Dixon, 1959). In the British Columbian Gelidium  species, carposporangia  are produced  singly, terminal^ and laterally on gonimoblast cells, rather than in clusters. In G.  latifolium  vegetative  and  cortex  G. and  pulchellum, ostioles  the  do  cystocarp  not  form;  cortex  is thicker than  carpospores  are  released  the by  degeneration of the pericarp (Dixon, 1959). This is different from G. purpurascens and G. vagum where the cystocarp cortex is not thickened over the vegetative cortex and one ostiole is present per locule. In G. pteridifolium ostiolar  regions" that  develop  have  plugs  that  later  break  the "potential down allowing  carpospore release (Hommersand and Fredericq, 1988).  Returning briefly to the question of the presence of an (generative) auxiliary cell in Gelidiales, discussion of this problem is given by Hommersand and Fredericq (1988). It is clear that the gonimoblast and  not from  an  develops from lobes of the fusion cell  unchanged carpogonium  as Kylin  (1928) proposed. Having  shown in this work that the functional carpogonium is intercalary  and  not  sessile, observations interpreted as carpogonia fusing with their supporting cells are erroneous and are actually of the unequal enlargement of some parts of the carpogonium (e.g. base and pit-connection region) relative to others. Using Drew's  Carpogonium and Carposporophyte Development / 104 (1954) definition of an auxiliary cell, the possibility that the supporting cell functions as an auxiliary cell can be eliminated, as a supporting cell does not exist in Gelidium.  Given that fusions with cortical cells occur, the question of the  nature of the auxiliary cell is reduced to whether any particular cortical cell is always the cell with which the carpogonium fuses and whether it functions in initiation of the gonimoblast or is strictly nutritive. I concur with Hommersand and  Fredericq (1988) that there is, as yet, too little information available on  cortical cell specificity or function. Recent observations  do not suggest that a  specified cortical cell is involved, and gonimoblast was not seen developing from the part of the fusion cell that fused with cortical cells. In light of Hommersand and  Fredericq's  (1988)  findings,  and  because  some  previously  reported  developmental patterns have been generalized as "typical for the Gelidiales" (e.g. Kraft, 1976), re-investigations of previously studied species would be in order to clarify debated aspects.  CHAPTER VI. SPERMATOGENESIS IN  A.  GELIDIUM  INTRODUCTION  In 1925,  Grubb wrote the following concerning the state of knowledge of male  organs in the Florideae (Grubb, 1925). Although nearly 150 years have passed since the first record of spermatia in the red algae occurred in print, our knowledge of these minute bodies which play so important a part in the reproductive processes of Rhodophyceae is surprisingly inadequate and limited. Records of the observation of antheridia in more than 120 European species of the Florideae are to be found scattered in algal literature, but of these the vast majority are simply notes to the effect that male plants have been seen and recognized. A certain number give a short description detailing the position of the antheridia, and some include a slight account of their structure, but owing to the nature of the material, modern cytological methods of investigation have only been brought to bear on a few forms. A similar situation still exists in 1988, even though spermatangia are known for many more species, and their development at the light microscope level has been studied to some extent. Few  observations of the development of red algal male  gametes have been made using electron microscope techniques.  The  male gamete and associated structures generally do not play an important  role in red algal taxonomy. The presence of spermatangia and their position on the thallus are noted, but spermatangial morphology usually is not used as a taxonomic character. Part of the reason for this is the often cryptic nature of male gametangia and the resulting rare collections of male gametophytes or their recognition as such in collections. The importance of male characters in taxonomy 105  Spermatiogenesis in Gelidium I 106 also depends on the taxonomic rank at which the characters can be applied. Gabrielson and Garbary (1987) recently have used spermatangial characters at the ordinal rank in the construction of phylogenetic trees. For example, the transverse (as opposed to oblique) division of the spermatangial mother cell to form a spermatangium  is characteristic of Gelidiales (Gabrielson and Garbary,  1987). At the generic rank, spermatangial position has been used to separate Gracilaria  into  Gracilaria  and Polycavernosa  subgenera  (Yamamoto, 1975)  and  to differentiate  the genera  (Bangmei and Abbott, 1985). As has been discussed  earlier in this work, the character of spermatangia occurring on the same plants as carpogonia (i.e. monoecy) instead of on separate male gametophytes (i.e. dioecy) can be used at the specific rank to recognize Gelidium  vagum and G.  johnstonii.  Spermatangia are known for appproximately 30-40% of gelidiaceous species, but their development seldom has been followed (Dixon, 1959; Akatsuka, 1970, 1973; Tazawa, 1975). This is understandable given the small size of spermatangial mother cells, spermatangia and spermatia.  Among the nine electron microscope studies of male gamete development in higher rhodophytes, none has been carried out on a member of the Gelidiales. Electron microscopj' of male gametes was genera  (Peyriere,  Levringiella densa;  gardneri  Kugrens,  1971,  Griffahsia  and  Erythrocystis  1974,  Janczewskia  performed primarily on ceramialean  flosculosa;  Kugrens  saccata; Scott and gardneri;  Scott  denudata and P. harveyi; Kugrens, • 1980, Polysiphonia  et  and  West,  1972b,  Dixon, 1973a, Ptilota al.,  1980,  hendryi), and  Polysiphonia Bonnemaisonia  Spermatiogenesis in Gelidium I 107 hamifera (Bonnemaisoniales) also was studied (Simon-Bichard-Breaud, 1971, 1972a, 1972b).  In  addition, there are  rhodophytes  (Peyriere,  1974,  Nitophyllum,  and Furcellaria).  some fragmentary  Polysiphonia,  reports  Rhodomela,  on  Laurencia,  other higher Polyneura,  In this study spermatiogenesis in two of the four  gelidioids was followed using light and electron microscopy, as well as in a third species using only light microscopy.  B.  METHODS  The species used in this study were Gelidium  purpurascens,  G. vagum and G.  coulteri (light microscopy only). Light microscopy methods are included in Chapter 3. Both field and  cultured  material  of G.  vagum  was  fixed  for electron  microscopy. Gelidium purpurascens male gametophytes were not seen in the field, consequently material for this study on spermatiogenesis was obtained from male gametophytes grown to maturity in culture from tetraspores.  Freshly collected field material was acclimated to a constant photoperiodic regime (12:8 LD  in 20° C) in order to sjmchronize cell divisions with light and dark  periods. Material was fixed 1 to 2 h after chamber lights came on to maximize the  chance of seeing cell division (van der Meer, pers. comm.). Fixation  was  carried out in the refrigerator for 7 h in 2.5% glutaraldehyde, a 1:1 mixture of 5% glutaraldehyde and Sorensen's phosphate buffer (pH 7.2). For rapid fixation and to prevent blockage of intercellular spaces by air bubbles, tissue was cut from plants in a drop of fixative. Fixed tissue was  postfixed in 1% osmium  tetroxide in the refrigerator for 16 h and then dehydrated. Dehydration  was  Spermatiogenesis in Gelidium I 108 carried out in a graded methanol series. Material was oxide, followed by  infiltrated in propylene  embedding in a graded series of (10-100%) Spurr's epoxy  resin. In the case of overnight steps, tissue was  placed on the rotator at room  temperature for the first and last hours, and in the refrigerator (not rotating) for  the  time  in between. A  sample  protocol of the  transmission electron  microscope fixation procedure for spermatangia is provided in Appendix 3.  Gold-purple to silver sections (170-75 um thick) of embedded material were cut on a Reichert OM  U3  ultramicrotome  using a Dupont diamond knife. Grids were  stained in saturated uranyl acetate in 70% methanol for 45 min, and then with Reynolds' lead citrate (Reynolds,  1963)  for 5.5 min. Sections were viewed and  photographed in Zeiss EM10A and EM10C transmission electron microscopes.  C.  SPERMATIOGENESIS  Spermatangial development is the same in all Gelidium  species in this study.  Spermatangia are borne in a superficial layer (Fig. 189)  on  short, ultimate  branchlets of gametophytes. In monoecious G. vagum, they occur proximal to the female area of branchlets, a short distance behind the apex. Spermatangia are initiated  and  mature at the same time as  the more distal carpogonia. In  dioecious G. coulteri and G. purpurascens, spermatangia extend practically to the apex of the branchlet, and the size of the fertile area expands as the branchlet lengthens. Spermatangia are conspicuous by being colourless and associated with a "halo" of mucilage.  Spermatiogenesis in Gelidium  I 109  Vegetative cortical cells are pigmented; their dominant cytological components are a few large chloroplasts located peripherally (Fig. 191). The nucleus is usually basal, and the central part of the cell is occupied by a vacuole (Fig. 191). In surface view, cells in male areas are paler and smaller in diameter  than  vegetative cortical cells, and grouped in two's or four's (Fig. 190). These smaller diameter cells are spermatangial mother cells produced by the anticlinal halving of a surface cortical cell and the subsequent division of these cells in half (Fig. 192). Spermatangial  mother cells  thus  have  pit-connections to a subtending  cortical cell and/or to another spermatangial mother cell (Fig. 192).  Elongated  spermatangial  mother  cells  contain  apical  plastids  ranging  from  proplastids to fully developed chloroplasts, typical of red algae, with an inner encircling  thylakoid  and aligned  single  thylakoids (Figs.  phycobilisomes. Starch grains accumulate  193, 203) bearing  in the basal half of spermatangial  mother cells (Figs. 192, 193), and the nucleus also is basal, elongating into a central position (Fig. 193) prior to nuclear division (Fig. 194). Although a large number of cells were observed, and plants were fixed at times when, according to previous reports, the possibility of seeing dividing cells was highest (Scott et al.,  1980; Scott, pers. comm.; Davis and Scott, 1986; van der Meer, pers.  comm.), mitotic  stages were not seen. After  karyokinesis, the spermatangial  mother cell nucleus returns to a basal position (Fig. 194). Cytokinesis occurs in a transverse plane characteristic of the Gelidiales, dividing the spermatangial mother  cell  approximately  in half  (Fig. 204)  and  producing  a  distal  spermatangium. In longitudinal sections of fertile male branches, the transverse divisions  of spermatangial  mother  cells  help  distinguish  spermatangial  from  Spermatiogenesis in Gelidium I 110 vegetative areas where cortical cells divide somewhat obliquely.  The spermatangium contains a prominent nucleus with condensed chromatin (Figs. 197, 201), endoplasmic reticulum and numerous mitochondria sometimes associated with  dictyosomes  (Figs.  195), but  lacks chloroplasts and  proplastids. If all  chloroplasts are not successfully excluded prior to cytokinesis of the spermatangial mother cell, they must degenerate very soon after the spermatangium is cut off as extrusion of membranes is sometimes observed in the young spermatangium (Fig. 198). In some views, spermatangial mother cells appear to be expelling an entire chloroplast by either a furrow or by reforming the plasma membrane proximal to the chloroplast (Fig. 193). Soon after the spermatangium is cut off, many cored vesicles appear in the cytoplasm. Their origin is unclear, although similar cored vesicles were dictyosome-derived  in developing tetrasporangia (see  Chapter 7). Cored vesicles fuse with the plasmalemma (Fig. 199), contributing to a  finely  fibrillar,  mucilage-like  layer  surrounding  the  spermatangium.  The  spermatangium is surrounded by the spermatangial mother cell wall (Fig. 205). One  or two  large spermatangial  spermatangium  (Fig.  dictyosome-derived Spermatangial  199),  vesicles  apparently  (Fig. 195),  form  from  in the basal part of the the  vesiculated  coalescence ER  of  (Fig. 196),  vacuoles can be discerned in the light microscope  areas at the base of spermatangia with whorls  vacuoles  uncored, or both.  as unstained  (Fig. 189). Spermatangial vcauoles are filled  of fibrillar material (Figs. 197,  200)  and  enlarge to a volume  approximately equal to that of the remainder of the spermatangial cytoplasm.  Release of the spermatangial vacuole along with adjacent cytoplasm (Fig. 203) is  Spermatiogenesis in Gelidium I 111 followed by release of a single spermatium from the spermatangium. In early stages of spermatium release, the discarded pit plug (originally between the spermatangial mother cell and spermatangium) is seen between the spermatangial vacuole and spermatangial mother cell. The new spermatangial mother cell wall is thicker and interspersed with vesicles near spermatangial  mother cell  adjacent to this  the pit plug (Fig. 205).  area  The  of thickened wall contains  additional vesicles, some of which are released to the wall. The space between the spermatangial vacuole and spermatangial mother cell often contains many vesicle remnants. (Figs. 204, 205) that could function in gamete release. The spermatium being released is squeezed through a weakened and ruptured area of the thallus wall (Fig. 202). It is conical in shape (Fig. 206), rounding up when free of the gametophyte. The spermatium is wall-less and contains a prominent nucleus  with  much  condensed  chromatin  (Fig. 206), several  large, oval  mitochondria and abundant cored vesicles that continue to be released to the cell exterior (Fig. 206). Chloroplasts or proplastids are absent.  Percurrent production of spermatangia occurs when the spermatangial mother cell expands into the space vacated by the previous spermatangium and cuts off another spermatangium (Figs. 205, 207). Occasionally, two spermatia are seen near the spermatangial mother cell that produced  them (Fig. 205), suggesting  rapid spermatium production. Loose pit plugs and successive old spermatangial mother cell wall layers indicate that at least four to five spermatangia can be produced percurrently from a single spermatangial mother cell (Figs. 207, 208).  Spermatiogenesis in Gelidium I 112 D.  COMPARISON  OF  FLORIDEOPHYCEAN  SPERMATIOGENESIS  IN  GELIDIUM  AND  OTHER  GENERA  Detailed ultrastructural studies of florideophyte spermatiogenesis have been made on Griffithsia  flosculosa (Peyriere, 1971), Levringiella  (Kugrens and West, 1972b), Ptilota  densa (Scott and Dixon, 1973a), Janczewskia  gardneri (Kugrens, 1974), Polysiphonia P.  hendryi  1971, similar  (Kugrens, 1980)  and  gardneri, Erythrocystis saccata  harveyi, P. denudata (Scott et al., 1980),  Bonnemaisonia  hamifera (Simon-Bichard-Breaud,  1972a, 1972b) and the general pattern of male gamete development is in all, although  there  are  some differences in certain details. For  clarification, it should be noted that Dixon (1959) used a terminology that is contrary to other authors. His "spermatangial mother cell", which expands giving rise to "spermatangia" producing two  by  oblique divisions, is equivalent to a cortical cell  to four spermatangial mother cells (Dixon, 1959). Likewise, his  "spermatangium", which divides transversely, is equal to the the spermatangial mother cell dividing to produce spermatangia (Dixon, 1959).  Kugrens' (1980) interpretation of spermatangium and spermatium in  Polysiphonia  hendryi generated terminology that also is at variance with that of other authors. He  interpreted the spermatangial mother cell as producing spermatia, the wall  matrix surrounding the spermatia representing the spermatangium. It is unclear whether the spermatangium also surrounds the "spermatial" mother cell, but if so, this spermatangium cannot be homologous to the spermatangium of other genera  which excludes the spermatangial mother cell. Clearly the outer wall  matrix, surrounding the spermatangial mother cell and developing male gametes,  Spermatiogenesis in Gelidium I 113 was produced by the periaxial cell before production of spermatangial mother cells and is equivalent to the wall produced by other vegetative periaxial cells. Thus I have retained the usage of other authors for (e.g. Scott and Dixon, 1973a) for "spermatangial mother cell" which produces a "spermatangium" and releases a "spermatium".  I  do  agree  with  Kugrens' (1980) designation  spermatangial  "vacuole", as opposed to "vesicle" and have followed his interpretation.  Spermatangial  mother cells differ from  vegetative cortical cells in that their  chloroplasts are less well developed (Kugrens, 1974; Peyriere, 1974), or, if well developed (i.e. with many single parallel thylakoids), are fewer and less dominant components of the cell (Kugrens and  West, 1972b), resulting in the reduced  pigmentation seen at the light level. The latter is true for the Gelidium species studied here. Spermatangial mother cells also are less vacuolate than cortical cells (Peyriere, 1971; smaller  and  Scott and  more  basal  Dixon, vacuoles,  1973a; Kugrens, 1974; except  in  Polysiphonia  this study), with hendryi  where  spermatangial mother cells have differentiated directly from pericentral cells and are thus vacuolate (Kugrens, 1980). The transverse division of the spermatangial mother cell to produce a spermatangium is characteristic of Gelidiales and of the genera  Gracilaria  and  Hypnea  (Tazawa, 1975). Spermatangia are cut off by  oblique divisions in other genera (Tazawa, 1975).  In Gelidium  each spermatangial mother cell produces a single spermatangium at  any one time, and over time sequential production of spermatangia some species, however, (e.g. Levringiella spermatangia  occurs. In  gardneri and Erythrocystis saccata) several  are produced simultaneously by a single spermatangial mother cell  Spermatiogenesis in Gelidium I 114 (Kugrens and West, 1972b), and percurrent production is not mentioned or visible in  illustrations. In  Polysiphonia  other  hendryi)  spermatangial  mother  species  (e.g. Ptilota  multiple, percurrent cell  (Scott  and  densa, Janczewskia  spermatangia  Dixon,  gardneri  and  are cut off from  1973a; Kugrens,  1974,  a  1980  respectively).  Chloroplasts in varying forms may (e.g.  Gelidium).  be present within spermatangia or are absent  If present, they most often are rare (Ptilota densa: Scott and  Dixon, 1973a), degenerating (Levringiella proplastids (Erythrocystis  gardneri:  Kugrens  and  West, 1972b),  saccata: Kugrens and West, 1972b; Polysiphonia  hendryi:  Kugrens, 1980), or have fewer single thylakoids than vegetative cells (Griffithsia flosculosa:  Peyriere, 1971). Organelles such as vacuoles, starch grains and the  spermatangial mother cell nucleus in Griffithsia  flosculosa  (Peyriere, 1971) are  positioned basally in the dividing spermatangial mother cell and excluded from the spermatangium. But in Gelidium,  are readity  well-developed chloroplasts  occur apically in spermatangial mother cells. These chloroplasts appear to be released prior to spermatangial formation and then degenerate outside the cell. Evidence of expelled membranous material was also occurs in Levringiella  gardneri,  seen in Gelidium  Erythrocystis  saccata  and probably  (Kugrens  and West,  1972b) and Ptilota densa (Scott and Dixon, 1973a).  Spermatangia show abundant signs of metabolic activity. In Gelidium Griffithsia 1972a,  (Peyriere, 1972b),  1971,  1974)  mitochondria are  and  Bonnemaisonia  closely  associated  vesicles were more abundant in Gelidium  (this study),  (Simon-Bichard-Breaud, with  dictyosomes. Cored  compared to Ptilota (Scott and Dixon,  Spermatiogenesis in Gelidium I 115 1973a) and Janczewskia  (Kugrens, 1974) and are probably dictyosome-derived.  There is evidence that their contents are secreted extracellularly, contributing to wall development. Although in Griffithsia  (Peyriere, 1971, 1974), Bonnemaisonia  (Simon-Bichard-Breaud, 1972a), and Janczewskia  (Kugrens, 1974), small amounts  of cytoplasmic starch are present early in spermatangial development, none was seen  in  spermatangia  of  Gelidium.  Because  spermatangial mother cells, it is unlikely  starch  is always  to be included  basal in  in newly formed  spermatangia. Since plastids are absent, starch could not be formed de novo in spermatangia.  The layers surrounding spermatangia in Gelidium differ from the three layered wall  described  (Kugrens and  in Janczewskia^  (Kugrens, 1974), Erythrocystis  West, 1972b). The  outermost and  and  thickest layer  Levringiella in Gelidium  consists of very loose, fibrillar, mucilage-like material (similar in appearance to the  contents of the spermatangial vacuole). Late in spermatangial development a  thin, granular, inner wall layer appears. From the description given by Kugrens (1974), the second layer in Janczewskia thick mucilage-like layer of Gelidium.  spermatangia may The  correspond to the  spermatangial wall of Polysiphonia  hendryi (Kugrens, 1980) does not appear to have any layers analogous to those in Gelidium. In P. hendryi, a dark line of compressed fibrils forms a "separating layer" at the beginning of spermatium enlargement (Kugrens, 1980, Figs. 7, 12). In  Kugrens' terminology, a spermatial wall (equivalent to spermatangial wall)  with a distinctive reticulate fibrillar construction forms around the spermatangium and is gelatinous in nature (Kugrens, 1980, Fig. 12). Both a separating layer and a reticulate spermatangial wall are absent in Gelidium,  but the reticulate  Spermatiogenesis in Gelidium wall  occupies  the  same  histochemical comparison  position  as  the  mucilage  of the reticulate wall and  layer  in  mucilage  I 116  Gelidium.  A  layer would be  valuable.  The  spermatangial nucleus in Gelidium  contains large amounts of condensed  chromatin, but the components of the mitotic apparatus were not seen. It was proposed for Griffithsia  flosculosa and Ptilota densa in which condensed chromatin  also occurs that the spermatangial nucleus is arrested at prophase (Peyriere, 1974;  Scott and  Dixon,  Kugrens (1974, 1980) Polysiphonia  Other  noted  chromatin. Contrarily,  that in released spermatia of Janczewskia  and  hendryi the chromatin was dispersed.  electron  reported  1973a), hence the condensed  a  microscope  predominance  studies of spermatiogenesis of dictyosomes  or ER,  or  in florideophytes have a  temporally  changing  dominance of both in spermatangia, the amount and kind of activity varying with species (Pe3riere, 1974). Dictyosomes and ER vesicles are the source of secreted r  mucilages,  wall  material  and  the  spermatangial  vacuole.  Gelidium,  In  spermatangial vacuoles were large, indicating much secretion. The source of this large volume of material in Gelidium  is not obvious. Dictyosomes occasionally are  seen but are not abundant, although cored vesicles believed to be derived from dictyosomes  are abundant, suggesting rapid  production of vesicles  from  few  dictyosomes. Occasionally in the early stages of spermatangial vacuole formation, vesiculated ER  is abundant basalry, seemingly  spermatangial vacuole. However, ER  involved in formation of the  is not apparent in spermatangia  possessing small spermatangial vacuoles.  alreadj'  Spermatiogenesis in Gelidium I 117 Spermatangial  vacuoles  are  the  most  prominent  developmental  feature  of  spermatangia. They are usually one to three in number and at maturity occupy up to one half the spermatangial volume. Their origin is debated; an endoplasmic reticulum (ER) origin is suggested in Levringiella Janczewskia in  (Kugrens, 1974) and Polysiphonia  Erythrocystis  (Kugrens  (Simon-Bichard-Breaud, Gelidium, and  ER  and  hendryi (Kugrens, 1980), whereas  West,  1972b)  and  Bonnemaisonia  1972a) the evidence indicates a dictyosome origin. In  as in Ptilota (Scott and Dixon, 1973a), both dictyosome-derived vesicles contribute  histochemical  tests,  to spermatangial vacuole the  spermatangial  (Simon-Bichard-Breaud, 1972a) and Polysiphonia polysaccharides. In Gelidium but  (Kugrens and West, 1972b),  formation. As vacuoles  in  determined  by  Bonnemaisonia  hendryi (Kugrens, 1980) contain  the internal structure consists of whorls of fibrils,  in Ptilota it also appears as bands of granular material (Scott and Dixon,  1973a). The spermatangial vacuolar contents of Polysiphonia  hendryi initially are  granular and later develop into fibrillar material (Kugrens, 1980), and a change in chemical reactivity or change in internal appearance occurs at maturity in Bonnemaisonia  (Simon-Bichard-Breaud,  1972a).  The  vacuoles also varies between taxa. In Bonnemaisonia are  position  of spermatangial  two spermatangial vacuoles  produced, one of which is basal and the other apical (Simon-Bichard-Breaud,  1971, 1972a, 1972b). They mature and are released in these positions without migration. To  date, Bonnemaisonia  is the  only  species  studied  where all  spermatangial vacuoles are not basal.  The role of the spermatangial vacuole in the spermatangium In  Gelidium  has been debated.  the spermatangial vacuole appears to function in release of the  Spermatiogenesis in Gelidium I 118 spermatium. The vacuole always is released from the spermatangium before the gamete is released, and it remains approximately the same size and configuration after release. Thus it does not seem to contribute to the mucilage or adhesive coating of the spermatium, as has been suggested for Griffithsia Levringiella  and Erythrocystis  Dixon, 1973a). The  (Peyriere, 1971),  (Kugrens and West, 1972b) and Ptilota  spermatium mucilage  probably is produced  (Scott and  primarily from  cored vesicles. The fact that the spermatangial vacuole never is retained within the Gelidium In  spermatium also suggests that it functions in spermatium release.  Levringiella  West, 1972b) and Polysiphonia  (Kugrens and  hendryi  (Kugrens,  1980), spermatia commonly retain their spermatangial vacuoles. No other workers have reported retention of spermatangial vacuoles in spermatia. It could reflect a real generic difference between taxa, or it could be due to accidental release of immature spermatia during electron microscopy  preparation. Kugrens and West  (1972b) proposed that osmotic pressure generated by the polysaccharides contained in  the spermatangial  vacuole forced the pit plug between the spermatangial  mother cell and spermatangium to break, enabling spermatial release. However, in Polysiphonia deposition  hendryi (Kugrens, 1980) the pit plug ruptures by spermatangial wall and  spermatangium  enlargement  and  the  separating layer breaks,  facilitating release of the spermatium. In several cases the pit plug in Gelidium was. unattached  on both faces, and  the spermatangial mother cell showed a  plasma membrane profile suggesting secretion of wall material under the plug that effectively breaks any solid connection between the spermatangium and the spermatangial mother cell. Although spermatium release could not occur with an intact pit-connection, the breaking of the pit-connection may sufficient,  for  spermatium  release. It  may  be  be necessary, but not  difficult  to  separate  the  Spermatiogenesis in Gelidium consequences of pit-connection breakage and  I 119  spermatangial vacuole release, as  expulsion of the spermatangial vacuole and adjacent cytoplasm effectively breaks the pit-connection. A spermatia  detailed ultrastructural study of mature but unreleased  [sometimes  seen  as  a  second  spermatangial mother cell (Fig. 205)] may  spermatium  in  series  the spermatangial mother cell, also released a  microscopy  a  resolve this question.  Dixon (1959) suggested that the lower cell of his "spermatangium" in i.e.  over  Gelidium,  spermatangium. Electron  clearly shows that this does not occur, at least in the  Gelidium  species studied here.  In the Gelidiales there are morphological variations in male gametophytes or the manner  in which  the  spermatangia  exception of Acanthopeltis japonica  are  borne on  gametophytes. With the  (Kaneko, 1968), Gelidium  johnstonii  and  G.  vagum, all gelidioids where males are known are dioecious. Spermatangia in these plants form close behind the apex of fertile branchlets, and as the apex grows the  male  area  lengthens. In monoecious plants, however, spermatangia  are  prevented from forming immediately behind the fertile branchlet apex, as this area differentiates carpogonia and nutritive filaments. In local G. purpurascens, male gametophytes are isomorphic with female gametophytes and tetrasporophytes and  are not smaller than female gametophytes as reported for G.  latifolium  (Dixon, 1959).  Male plants or male segments of monoecious gametophytes are paler in colour than  vegetative regions (Fan,  1961;  Akatsuka,  1970,  1973,  1979). Electron  Spermatic-genesis in Gelidium I 120 microscopy has shown that spermatangia lack chloroplasts and thus should be colourless.  Spermatangial  mother  cells  contain  fewer  and  less  prominent  chloroplasts than vegetative cortical cells and might be expected to be somewhat pigmented. Male areas also may  be recognized at low magnification by the  thickened mucilage layer over spermatangia (Dixon, 1959; Akatsuka, 1970), or by the  dilated and flatter apices of fertile branches  character  applies  only  (Tazawa, 1975). The latter  species (Gelidium  to certain  amansii,  G.  pacificum,  Pterocladia nana, Tazawa, 1975; P. caloglossoides, this work) and was not evident in  coulteri, G. johnstonii,  G.  G.  purpurascens  or  G.  vagum.  In  transverse  or  longitudinal sections, spermatangia are distinguished easily from vegetative cortical cells  by  the  transverse  walls  between  spermatangial  mother  cells  and  spermatangia, and the narrower width of spermatangial mother cells. In contrast, vegetative cortical cells have oblique division planes.  Despite  the  gametophytes,  differences the  in  pattern  how  and  of their  where  spermatangia  development  are  in florideophytes  borne  on  including  Gelidium is remarkably uniform. In addition, two species of Gelidium studied here could not be differentiated at the ultrastructural level.  Too few ultrastructural studies of spermatiogenesis have been carried out to know yet  if there are variations on the basic pattern that may  significance. The  mechanism  of exclusion  be of taxonomic  or elimination of chloroplasts from  spermatangia, the mode of formation of spermatangial vacuoles, the number and position of spermatangial vacuoles and the presence or absence of percurrent production of spermatangia vary in studies to date and may  have some potential  Spermatiogenesis in Gelidium I 121 for use in taxonomic studies. The mechanism of spermatium release also needs clarification.  C H A P T E R VII.  U L T R A S T R U C T U R E OF TETRASPOROGENESIS  IN  GELIDIUM  A.  INTRODUCTION  Tetrasporogenesis  in  red  algae  has  been  studied  more  extensively  than  spermatiogenesis at both light microscopy and ultrastructural levels. The origin of tetrasporangia and production of tetraspores have been noted in many gelidiaceous species. Fan (1961) remarked on the uniformity of tetrasporangial development in Gelidiales. Tetrasporangial characters (excluding arrangement on fertile branches) have not been used as taxonomic characters in Gelidiales, with the exception of the  production of bispores in Suhria  vittata and  Onikusa  pristoides (as  Gelidium  pristoides) (Fan, 1961). Tetrasporangial initials are borne laterally on subcortical cells, or "terminal in position on a lateral filament" as often stated (e.g. Fan, 1961). The  young sporangium has a single pit-connection that is initially basal  and becomes lower lateral as the sporangium expands. The  tetrasporangium is  deeply pigmented and divides into four cruciate or irregular cruciate, occasionally tetrahedral (Dixon, 1959) tetraspores. Release occurs through the degraded apical sporangial wall (Dixon, 1959). Additional sporangia are produced by the expansion of other initials after release of earlier formed sporangia.  Ultrastructurally, tetrasporogenesis has been studied in the following Ceramiales, Griffithsia  flosculosa  (Peyriere, 1969,  West, 1972c), Ceramium (Scott  and  Dixon,  1970), Levringiella  sp. (Chamberlain and  1973b), Callithamnion 122  gardneri  (Kugrens  Evans, 1973), Ptilota  roseum  (Konrad  and  hypnoides  Hawkins, 1974b),  Ultrastructure of Tetrasporogenesis Polysiphonia  denudata  Erythrocystis Corallina 1984),  two  1982),  and  Thomas,  1975;  montagnei (Santisi and De Masi,  officinalis  (Pueschel,  (Scott  (Peel et al.,  species 1982),  and  ultrastructural  of  species  1973), Haliptilon  the  studs'  of  cuvieri  Rhodymeniales,  Palmariales,  Palmaria  of tetrasporangial  in order to determine  and  Scott,  1977),  1981), two members of Corallinales,  of Hildenbrandiales, Hildenbrandia one  one  Alley  in Gelidium I 123  (Vesk and  rubra  Gastroclonium  palmata  development  and  H.  Borowitzka, occidentalis  clavatum  (Pueschel,  in Gelidium,  (Gori,  1979).  was  This  undertaken  their patterns and compare them with previous studies of  other red algae.  B.  METHODS  Fertile tetrasporophytes collected study 6.  from  the  of Gelidium coulteri, G. purpurascens  field  or  obtained  area. Tissue was prepared  However,  fixation  and  temperature  walls.  and  Material  postfixed  culture  isolates  for electron microscopy as  osmication  temperature to allow penetration tetrasporangial  from  times  were  of glutaraldehyde was  fixed  in  in osmium for  originating from  the  outlined in Chapter  longer  or  at  a  warmer  and osmium through the thick  glutaraldehyde  18-19  and G. vagum were  h.  at  for  room  4  h.  at  room  temperature.  The  tissue preparation protocol that was used is given in Appendix 3.  C. ULTRASTRUCTURAL  The  process  purpurascens  of and  tetraspore  OBSERVATIONS  production  G. vagum is  similar  and at  maturation both  light  in and  Gelidium electron  coulteri,  G.  microscope  Ultrastructure of Tetrasporogenesis in Gelidium  I 124  levels, although there are differences between these species regarding the position of sporangia on stichidia. The account given here is applicable to all three local Gelidium  species. Structures that exhibited the greatest amount of change from  their appearance in vegetative cells were the nucleus, dictyosomes and wall.  1. Pre-meiotie Tetrasporangium  The  presence  marker  or absence of three  for  events  tetrasporangial  during  initial cell.  development  Vegetative  which tetrasporangia differentiate, grains,  and chloroplasts  distinctive  wall layers  of  mature  cortical cells  have  that are few  serves as tetraspores  (similar  a central vacuole, in number, large  to  a reliable from  Fig.  191)  basal nucleus,  the from  starch  and peripheral. Young  sporangial initials contain the remnants of a vacuole (Fig. 210), and the vacuolar contents appear to be incorporated into vesicles and secreted extracellularly (Fig. 212).  Occasionally, whorls of membranes are extruded from the cytoplasm (Fig.  211). In tetrasporangial initials, chloroplasts divide producing man3 smaller, ovate r  chloroplasts. Chloroplasts are unaligned and generally peripheral, but also may be scattered  throughout  dictyosomes cells,  are absent,  and  starch  tetrasporangia fibrillar,  the  is  electron  cytoplasm  (Fig.  214).  In  very  young  tetrasporangia,  or are so few that they were not seen in sections of  grains  are  absent.  The  convoluted  (Figs.  210,  216).  transparent  wall  (layer  1)  plasma Vegetative (Fig.  212)  membrane cells that  of  have is  a  young loose,  thickened  to  100-250 nm wide during differentiation and early development of the sporangium (Fig.  215).  Dictyosomes  and vesicles  that  might be contributing wall material  (Fig. 216) are not abundant, thus the source of wall components is unclear.  Ultrastructure of Tetrasporogenesis in Gelidium I 125 In young tetrasporangia the interphase nucleus is enlarged and central (Figs. 214, 215) and contains a large nucleolus with an uneven granular composition (Figs. 213, 219). Nucleolar vacuoles are evident as areas of the nucleolus with the same electron density as the nucleoplasm (large arrows, Fig. 213), and have been associated with RNA synthesis (Peel et al., 1973). Pores are abundant in the nuclear envelope (arrowheads, Fig. 213) and perinuclear endoplasmic reticulum (PER)  is  present  Mitochondria  are  (small  arrows,  concentrated  in  Fig. the  213,  Fig.  immediate  218,  arrows,  vicinity  of  Fig. the  219).  nucleus.  Dictyosomes appear just prior to karyokinesis and are small, flattened and up to 12 cisternae thick, with ends of distal cisternae inflated and saccate (Fig. 221). Larger cisternae contain an elongate core of moderately electron dense, fibrillar material (Fig. 221). The cisternal peripherj', as well as the entire contents of young cisternae, are electron transparent and sparsely fibrillar. There is a close association between mitochondria and the forming face of dictyosomes (Fig. 221). The narrow space between these organelles is unlike the surrounding cytoplasm in that it is free of ribosomes. Small, narrow, lenticular starch grains also appear prior to karyokinesis (Figs. 214, 215, 218, 219), first located centrally in the  vicinity  of  the  nucleus  and  between  chloroplasts,  and  later  scattered  throughout the tetrasporangium.  Actual meiotic division stages or synaptonemal complexes were not observed in these preparations. Several elongated and spindle shaped nuclei with depressions at their ends (Figs. 218, 219) and one with a narrow waist and chromatin (Fig. 217) probably were preparing for division. Somewhat older tetrasporangia with two nuclei but had not yet undergone cytokinesis (Fig. 220).  Ultrastructure of Tetrasporogenesis in Gelidium I 126 2. Post-meiotic  Tetrasporangium  Following karyokinesis, as the tetrasporangium enlarges, starch grains, chloroplasts and  dictyosomes become more abundant (Fig 220). Starch grains change from  lenticular to ovate in shape (Figs. 220, 224). Chloroplasts divide by pinching into two  and become somewhat more elongate. Thej' are well developed, with an  encircling thylakoid interior to the double chloroplast membrane, many parallel, single thylakoids, plastoglobuli and areas of DNA  (Figs. 221, 225). Dictyosomes  become more prominent, particularly around the periphery of the sporangium. At low magnification they are noticable as clusters of sacs (Fig. 223) that still are flat but composed of more cisternae with larger inflated sacs. They are abundant in  the tetrasporangium  at all stages  of development. Mitochondria  are small  relative to chloroplasts and are closely associated with the forming face of all dictyosomes  (Fig. 221). Endoplasmic  reticulum  (ER) also  is present in  tetrasporangia, although often difficult to distinguish in the dense cytoplasm (Figs. 221, 222).  A second tetrasporangial wall layer (layer 2) (Fig. 222) is deposited inside the first  layer. This  occurs  when  abundant in the cytoplasm  starch  grains  and chloroplasts have become  (Figs. 220, 224) during  karyokinesis  but before  cytokinesis. Layer 2 (arrow, Fig. 227) is narrower (20-25 nm wide) and more densely fibrillar than layer 1 and of medium electron density, appearing  as a  grey band between white (layer 1) and black (layer 3) bands (Figs. 227, 232). It is deposited by abundant dictyosome- (Fig. 225) and ER-derived vesicles (Figs. 222, 225).  Ultrastructure of Tetrasporogenesis in Gelidium I 3. T e t r a s p o r a n g i u m at  The  127  Cytokinesis  tetrasporangium undergoes cytokinesis at the two-nucleate stage, observations  supported by light microscope studies on hematoxylin-stained material. The second meiotic  division then occurs in both cells followed  perpendicular uninucleate  by  another cytokinesis in  planes, producing a cruciately divided tetrasporangium with four tetraspores  (Fig.  78).  Each  tetraspore  nucleus  is  small  and  inconspicuous, obscured by the great abundance of other organelles and inclusions. Chloroplasts and  starch grains are very abundant (Fig. 226). In some mature  tetraspores, chloroplasts are oriented radially around the nucleus (Fig. 229), but this arrangement does not seem to be universal. Starch  grains have reached  their maximum size in the tetrasporangium after cytokinesis (tetraspores) (Fig. 226). They are electron transparent, substructure dimensional  although and  dark,  angular  non-membrane bound and  shadow-like  (Fig. 228).  marks  make  Endoplasmic  appear to lack  them  reticulum  appear occurs  three in  the  peripheral cytoplasm of older tetrasporangia contributing vesicles to wall formation (Fig. 231).  In  tetrasporangia  that  are  undergoing  the  second  cytokinesis,  and  shortly  thereafter, dictyosomes are abundant throughout the cytoplasm (Fig. 226), but not as prominent as in earlier stages. They are small and obscured by an abundance of chloroplasts and  starch grains. Dictyosomes also have undergone a striking  morphological change. The3' are distinctly concave; all of the approximately  9-12  cisternae per stack are strongly curved (Figs. 228,  230). There is a line of  electron dense material inside the closely associated  cisternae of the younger  Ultrastructure of Tetrasporogenesis in Gelidium. I 128 (forming face) half of the dictyosome (arrowhead, Fig. 230) that is progressively more restricted in extent in the central region of more mature cisternae (Fig. 230). The dark material occurs in closely appressed, narrowest parts of cisternae and is absent from the saccate ends. Cisternae that are innermost in the cup that is formed are oldest. Their inflated sacs separate and are released as spherical vesicles (cored vesicles) which have an electron dense core and a medium density fibrous periphery (Fig. 230). Cored vesicles release their contents to the cell wall, contributing to its formation (Fig. 231). Mature tetraspores have few cored vesicles and visible dictyosomes.  The third and final wall layer of the tetrasporangium (layer 3) is produced at cytokinesis (Figs. 226, 227, 232). It is 25-55 nm  wide, stains strongly with  osmium and is densely fibrillar, with parallel fibrils much more clearly aligned than in other layers (Fig. 227). Soon after layer 3 is completed, a layer of whorled fibrillar material, similar to the mucilage produced by spermatangia and contained in spermatangial vacuoles, appears internal to layer 3 and surrounds the tetraspores (Fig. 232). As in spermatangia,  this material represents the  contents of dictyosome-derived cored vesicles (Figs. 231, 232).  D.  DISCUSSION  General  trends of developmental  changes during tetrasporogenesis in Gelidium  agree with patterns observed in other florideophytes. An extensive comparison and discussion of tetrasporogenesis was presented by Vesk and Borowitzka  (1984).  Some differences in details do occur between the species, and these may be  Ultrastructure of Tetrasporogenesis in Gelidium I 129 taxonomically  important. The tetrasporangial wall differs from the wall of the  vegetative cell in most species (e.g. Ptilota  hypnoides, Scott and Dixon, 1973b;  Levringiella  gardneri,  Borokitzka,  1984), and a proteinaceous (Pueschel,  Hildenbrandia  Kugrens  rubra, H.  1979), Callithamnion  and  West, 1972c; Haliptilon  cuvieri,  Vesk and  1979) cuticle forms [e.g. in  occidentalis (Pueschel, 1982), Palmaria  palmata  (Pueschel,  roseum (Konrad Hawkins, 1974b) and Ptilota hypnoides (Scott  and Dixon, 1973b)]. It has been proposed that the cuticle and tetrasporangial wall function vegetative  to isolate the tetrasporangial cytoplasm from  cells  and  sporangia  Borowitzka, 1984). In Gelidium,  at  other  developmental  the influence of  stages  (Vesk  and  a cuticle was not apparent and neither was a  unique wall formed early in tetrasporangial development; the initial wall thickens and a second wall layer is not deposited until karyokinesis.  Division figures and Synaptonemal  synaptonemal  complexes  were not observed in  complexes are definitive proof of meiosis, but have been seen in  few studies (Kugrens and West, 1972a, Janczewskia Gonimophyllum  palmata;  Gelidium.  skottsbergii  and  Polycoryne  gardneri, Levringiella  gardneri;  Broadwater, Scott and Pobiner, 1986a, Dasya  Pueschel,  1979,  baillouviana).  gardneri, Palmaria  They occur  in few sections because of their small size and thus are less likely to be seen than larger, more conspicuous indicators of nuclear  division, such as spindle  shaped nuclei and division furrows.  Increase in chloroplast number throughout tetrasporangial development occurs by pinching of single chloroplasts into two. This has been observed in Gelidium and reported  in Griffithsia  flosculosa  (Peyriere, 1969), Ptilota  hypnoides (Scott and  in Gelidium 1 130  Ultrastructure of Tetrasporogenesis Dixon, 1973b), Palmaria 1982)  and  palmata  Haliptilon  cuvieri  (Pueschel, 1979), Gastroclonium (Vesk  and  Borowitzka,  clavatum (Gori,  1984). Early  in  the  development of P. palmata tetrasporangia, rapidly replicating chloroplasts have the appearance of proplastids with few  or no  single parallel thylakoids  (Pueschel,  1979), while later in development plastids are differentiated with several parallel thylakoids. Chloroplasts in early stages of tetrasporogenesis in H. cuvieri had few thylakoids, but  later  became fully  developed  (Vesk  and  Borowitzka, 1984).  Chloroplasts in Gelidium and Erythrocystis montagnei (Santisi and De Masi, are  well  developed  at  all stages of tetrasporogenesis;  1981)  proplastids were not  observed in this study.  The importance of dictyosomes and ER  in secretory activity is recognized in all  ultrastructural studies of tetrasporogenesis  (e.g. Peyriere, 1970;  Chamberlain and  Evans, 1973; Konrad Hawkins, 1974b; Alley and Scott, 1977). During periods of wall deposition, dictyosomes and dictyosome-derived vesicles become more abundant, particularly  in the  peripheral cytoplasm, and  becoming more dilated and reduced  (in mature  hypertrophied.  cisternae  After wall and  tetraspores), dictyosomes  decrease  increase  in number,  mucilage secretion is in  dilation,  size  and  number.  Many observers of tetrasporogenesis report or illustrate close associations between mitochondria and the forming face of dictyosomes (Peyriere, 1969, 1970; Kugrens and  West, 1972c; Chamberlain and Evans, 1973;  Peel et al,  Dixon, 1973b; Konrad Hawkins, 1974b; Scott and Scott, 1977; Pueschel, 1979,  1982;  1973;  Scott and  Thomas, 1975;  Alley and  Santisi and DeMasi, 1981; Gori, 1982; Vesk  Ultrastructure of Tetrasporogenesis in Gelidium I 131 and  Borowitzka,  1984;  this  study). Dark  layers  occur  between adjacent,  closely-appressed cisternae in the younger half of the hemispherical dictyosome. These have been likened to cementing layers (Konrad Hawkins, 1974b), but their nature is not known, and they have not been reported in other plants and animals (Alley and Scott, 1977). Products of dictyosomes also change through time, evidenced by Without  the variety of vesicles reported to be dictyosome-derived.  histological information it is difficult to determine  the homology of  vesicles reported as fibrillar (Kugrens and West, 1972c, Levringiella granular  (Kugrens  and  West, 1972c, Levringiella  gardneri;  Evans, 1973, Ceramium sp.), dense (Peyriere, 1969, Griffithsia  gardneri),  Chamberlain  and  flosculosa), with a  fibrillar core (Chamberlain and Evans, 1973, Ceramium sp.; Santisi and De Masi, Erythrocystis  1981,  montagnei),  and  granular  with  a  fibrillar  outer region  (Chamberlain and Evans, 1973, Ceramium sp.). Late in cytokinesis, dictyosomes of Gelidium  (this study), Levringiella  sp. (Chamberlain  and  1974b), Erythrocystis  Evans,  gardneri (Kugrens and West, 1972c), Ceramium 1973), Callithamnion  montagnei (Santisi and De  (Pueschel, 1979), Gastroclonium  clavatum  roseum (Konrad  Masi, 1981), Palmaria  (Gori, 1982)  and  Callithamnion  Hawkins, palmata roseum  carpospores (Konrad Hawkins, 1974a) have a distinctive hemispherical morpholog3' and  produce  cored vesicles. Peyriere (1970) reported positive tests for acid  mucopolj'saccharides within cored vesicles formed stages in Griffithsia vesicles  of  flosculosa,  carpospores  (similar in appearance  at two different developmental  and Tripodi and De Masi (1975) found that cored  in Pterocladia  to Gelidium  capillacea  and  Polysiphonia  sertularioides  cored vesicles in tetrasporangia) gave a  positive reaction to Thiery's test for polysaccharides. These cored vesicles are sometimes close to the plasma membrane, suggesting their contents are being  Ultrastructure of Tetrasporogenesis in Gelidium I 132 released to the wall (Tripodi and De Masi, 1975). The contents of cored vesicles in Gelidium  (this study) and Erythrocystis montagnei (Santisi and De Masi, 1981)  are secreted to the inner tetrasporangium wall or mucilage layer and are absent from released tetraspores (Santisi and De Masi, 1981). Similar appearing vesicles are believed to function in spore adhesion in other species (Vesk and Borowitzka, 1984, Haliptilon  cuvieri).  Dictyosome cisternae undergo a maturing process as they move through the stack (by addition and attrition of other cisternae) from the proximal (forming) face to the distal face. Alley and Scott (1977) proposed that they arise at the forming face from the fusion of small, ER-derived vesicles, and vesicles in transition between ER and the forming face of dictyosomes were seen in Erythrocystis montagnei (Santisi and De Masi, 1981). In Gelidium,  ER occurs near the forming  face, but vesicles never were seen in the space between mitochondria and the youngest  cisternae. It is possible that  small  contributing vesicles were not  sectioned or that thej' are added laterally and not centrally. Konrad Hawkins (1974b) stated that ER vesicles from chloroplast blebs fuse to form cisternae, and also that at least some cisternae contain chloroplast enzjanes capable of starch degradation, as cisternae were seen near eroding starch grains. In Gelidium, chloroplasts in the general vicinity of dictyosomes showed no evidence of special positioning near dictyosomes, or of blebbing, and starch grains were not in an eroding or eroded state. Vesicle production from dictyosomes is by release of entire cisternae (Peyriere, 1969, 1970; Alley and Scott, 1977) or release of dilated ends of cisternae as vesicles (Konrad Hawkins, 1974b). In  Gelidium,  dictyosome-derived vesicles are smaller than a whole cisternum, indicating that  Ultrastructure of Tetrasporogenesis in Gelidium I 133 vesicles probably are derived from cisternal ends.  Endoplasmic reticulum also is known to produce striated vesicles (Kugrens and West, 1972c) and  mucilage  vesicles  (Pueschel, 1979). In Gelidium,  vesicles  containing membranes or smaller vesicles, similar to multivesicular bodies or lomasomes (Scott and Dixon, 1973b; Chamberlain and Evans, 1973), release their contents to the wall.  Endoplasmic reticulum is associated with the nucleus at meiosis as PER. was not abundant in Gelidium,  but is extensive, oriented parallel to the nuclear  envelope or radially at different times in Corallina officinalis (Peep et al., and Haliptilon  PER  1973)  cuvieri (Vesk and Borowitzka, 1984), and believed to be related to  high metabolic and synthetic activity of the nucleus prior to, and throughout, meiosis. Extensive, radially-oriented PER  may  be characteristic of Corallinales; it  has only been reported from this order, although it is absent in some corallines (e.g. Jania  rubens (Linnaeus) Lamouroux, Peel et al.,  Gray and Fosliella  1973; Lithothrix  sp., Vesk and Borowitzka, 1984). Endoplasmic reticulum also  is important in inner tetrasporangium wall formation in Haliptilon and  aspergillum  cuvieri (Vesk  Borowitzka, 1984) and is present but not conspicuous at the cytoplasm  periphery in Gelidium.  Vesicles in Gelidium the plasma officinalis  tetrasporangia release contents to the wall by fusing with  membrane. No  vacuoles, similar  to those reported  (Chamberlain and Evans, 1973), Ptilota  1973b), Polysiphonia  denudata  (Alley and  in  Corallina  hypnoides (Scott and Dixon,  Scott, 1977), Erythrocystis  montagnei  Ultrastructure of Tetrasporogenesis  in Gelidium I  134  (Santisi and De Masi, 1981) and Griffithsia flosculosa (Peyriere, 1970) were seen. Mucilage sacs, present  in Palmaria  palmata  (Pueschel,  1979), also were not  observed in Gelidium.  Deposition of a series of cell wall layers results from the cytoplasmic secretory activity described above. Unlike all other species where tetrasporogenesis has been studied (Vesk and Borowitzka, 1984), Gelidium does not deposit a tetrasporangial initial wall early in differentiation that is different from the vegetative wall. A histochemical comparisons  study of tetrasporangium wall layers in Gelidium with  vegetative  walls, as  the  vegetative  wall  would enable  of G.  pacificum  Okamura has an outer sulfated polysaccharide layer and an inner cellulosic layer (Akatsuka and  Iwamoto, 1979). Based on  formation,  outer, thick, loosely fibrillar  the  position, appearance and wall  (layer  1) of the  time of Gelidium  tetrasporangium is probably equivalent to the tetrasporangial mother cell wall in Levringiella  gardneri (Kugrens and West, 1972c) and Ptilota hypnoides (Scott and  Dixon, 1973b), layer three in Callithamnion  roseum (Konrad Hawkins, 1974b), and  the inner tetrasporangial wall in Palmaria palmata (Pueschel, 1979), Hildenbrandia rubra and H. occidentalis (Pueschel, 1982). The medium density, grey wall (layer 2) in Gelidium  is different  from  layer  1  and  is similar in position  appearance to the tetrasporangial initial wall in Ptilota  hypnoides (Scott  and and  Dixon, 1973b). The last wall layer laid down in Gelidium (layer 3) is continuous with the dark layer separating tetraspores at cytokinesis, and a similar layer is recognizable in other genera. A mucilage layer also cleaves the tetrasporangium in other florideophyte genera (e.g. Hildenbrandia,  Pueschel, 1982; Haliptilon,  Vesk  and Borowitzka, 1984). This layer is recognizable by its position with respect to  Ultrastructure of Tetrasporogenesis in Gelidium  I 135  the dark, inner wall and its swirled fibrillar appearance. Peyriere (1970) found in  Griffithsia  flosculosa  that the last type of tetrasporangial secretion, which  appears similar to the mucilage-like material in Gelidium,  reacted positively for  mucopolysaccharides in histochemical tests.  In  all ceramialean and  corallinalean species studied ultrastructurally to date,  tetraspores are tetrahedrally or zonately (respectively) arranged in the sporangium. Both  meiotic divisions of the nucleus are completed  before cytokinesis, and  cleavage is simultaneous (Kugrens and West, 1972c, Levringiella and Dixon, 1973b, Ptilota  hypnoides; Chamberlain  sp.; Alley and Scott, 1977, Polysiphonia Haliptilon  cuvieri; Broadwater,  except in Erythrocystis  and Evans,  gardneri; Scott Ceramium  1973,  denudata; Vesk and Borowitzka, 1984,  Scott and  Pobiner, 1986b, Dasya  baillouviana)  montagnei where the cleavage furrow is half completed  before karyokinesis is over (Santisi and De Masi, 1981). Cytokinesis is initially sequential, and later simultaneous in Palmaria  palmata  tetrasporangia.  zonately  tetrasporangia  (Pueschel, 1979) of Hildenbrandia  and rubra  in the and  H.  with cruciately divided to  occidentalis  cruciate^  divided  (Pueschel, 1982).  Pueschel (1979, 1982) observed the first furrow forming after meiosis and not being completed until after the second perpendicular furrows had been initiated. Gelidium  also has  observed  ultrastructurally.  cruciately  divided  Light  tetrasporangia, but cytokinesis was  microscope  studies  not  with hematoxylin-stained  tetrasporangia suggested that division is completely sequential and that the second meiotic division occurs after the first cytokinesis.  In  summary,  whereas  tetrasporogenesis follows  a  similar  pattern  in the  Ultrastructure of Tetrasporogenesis in Gelidium florideophytes  studied  to date, some  differences  between  genera  I 136  have been  recorded such as extent and orientation of PER, types, functions and sources of vesicles, and timing of cleavage furrow development. As in spermatiogenesis, there are not yet enough studies on several members of a taxon (i.e. genus, family, order) to determine whether these differences are phylogenetically significant or whether differences merely represent variations among individuals.  C H A P T E R VIII. G E N E R A L DISCUSSION AND  In  spite of the great confusion confounding  Pterocladia  CONCLUSION  the taxonomy of Gelidium  and  species (e.g. Dixon, 1958), it is possible to determine species, if the  range of morphological and anatomical variability within a species is studied and determined for each taxon with in a restricted geographic area. In this study, conducted in British Columbia and northern Washington, species were assigned to genera on the basis of cystocarp morphology. Where cystocarps were not known for  local species (i.e. G. coulteri, P.  caloglossoides), ecological, vegetative and  tetrasporangial characters were used to assign names to taxa. Although six taxa were recorded originallj' from the area, it was found in this study that only four species of Gelidiales are  present: Gelidium  coulteri, G.  purpurascens,  G.  vagum  and P. caloglossoides. A key was  constructed (Fig. 5) to enable identification of  these  biogeographic  species. An  emphasis  on  and  developmental patterns of  reproductive structures as well as vegetative features, and on ecological characters may  prove useful in further taxonomic determinations of gelidioid algae.  To date, the only character that can be used reliably to distinguish between Gelidium the  and Pterocladia  apical  is the number of cystocarp locules. However, some of  architecture characters noted  by  Rodriguez and  Santelices (1987,  unpubl. mscr.) show potential for separating some of the species of these genera vegetatively. Two  of these characters, similarit.y/dissimilarity of apical and lateral  initials  indentation of lateral  Gelidium,  and G.  the  coulteri, G.  purpurascens  initials, were examined  and  G.  vagum,  and  the  in the three one  Pterocladia  species, P. caloglossoides, from the study area. Apical and lateral initials of the 137  General Discussion and Conclusion / 138 local gelidiaceous species were found to conform initials are similar and domed in Pterocladia,  to the pattern: both types of  but apical initials are domed and  lateral initials are dissimilar, inverted conical in Gelidium.  The second character  given by Rodriguez and Santelices (1987, unpubl. mscr.) was P.  caloglossoides  although  exceptions in some Pterocladia character  of  Pterocladia  from Gelidium  the}' suggested  that  this  not consistent for  character might  have  species, including P. caloglossoides. Thus this second  indentation of initials  is incapable  of  differentiating  between  in the study area. In the present study, neither apical  nor lateral initials are indented between cortical lobes in P. caloglossoides. Thus the similarity in shape of apical and lateral initials in Pterocladia the dissimilarity of these initials in Gelidium cystocarp locules, are characters that may  species, and  species, in addition to number of be used to separate Gelidium  and  Pterocladia.  Gelidium  vagum undergoes a regular Polysiphonia-type life history in culture, and  the common occurrence of gametophytes and tetrasporophytes suggests the same is operating in the field. According to culture evidence and field observations G. purpurascens  in the study  area is capable of, but probably  rarely actually  completes, a Polysiphonia-type life history in situ. Fertile male gametophytes grew from tetraspores of G. coulteri in culture, but there are no field collections of fertile gametophytes. The complete Polysiphonia-type  life history probably occurs  only occasionally, if at all, in G. coulteri from the study area and vegetative or apomictic Pterocladia  mechanisms  probably  sustain  the  population  most  caloglossoides populations also persist predominantly  apomictically. Chromosome counts  of n= 14-15  of the  time.  vegetatively or  were obtained from undivided  General Discussion and Conclusion / 139 tetrasporangia of G. vagum.  The pattern of spore germination was found to be similar for both carpospores and tetraspores. A single germ tube grows from the spore, the entire cytoplasmic contents of the spore evacuate into the germ tube and a wall cuts off the germ tube from the empty spore. The germ tube divides unequally to form a concave and a fusiform cell. The primary attachment rhizoid forms from a derivative of the concave  cell, and  the sporeling apical cell forms from  a fusiform cell  derivative. As apical organization is established the subapical cell produces two lateral periaxial cells.  Studies of developmental  patterns in local gelidiaceous species revealed some  previously unknown or unsubstantiated aspects. Hommersand and Fredericq (1988) recently proposed that carpogonia in Gelidiales are intercalary and that sessile carpogonia  reported  by  previous  workers  (Dixon,  1959;  Fan,  1961)  are  non-functional, but provided no data. The present study showed that the fusion cell is initiated from an intercalary carpogonium. In light of Hommersand and Fredericq's (1988) observations and  the corroborative findings of this stud}',  female gamete and carposporoprryte development needs to be followed in detail for a wide range of other gelidiaceous species. Such an expanded study is predicted to reveal developmental variations that may  be important as ordinal characters.  Ultrastructure of spermatiogenesis in Gelidiales was documented for the first time in this study. Exclusion of apical chloroplasts from the spermatangial mother cell is accomplished by the previously unreported mechanism of an ingrowing furrow  General Discussion and Conclusion / 140 that cuts off the cytoplasm containing the chloroplast. The spermatangium cut off from  the spermatangial mother cell is achloroplastic. A spermatangial vacuole  develops  in the spermatangium  from  both  dictyosome-derived  vesicles and  vesiculated endoplasmic reticulum. The spermatangial vacuole is released from the spermatangium prior to spermatium discharge. The percurrent production of at least four spermatangia  from  a single  spermatangial mother cell  occurs in  Gelidium.  Ultrastructure of tetrasporogenesis in a member of the Gelidiales was documented here for the first time. The stage of wall development can serve as a marker for developmental events in the tetrasporangium. Prior to meiosis, the nucleus is large,  dominating  the cytoplasm,  and  the wall  is thickened. Following  karyokinesis, a second distinctive wall layer is deposited and chloroplasts and starch grains increase in number. Dictyosomes become more abundant and the number of cisternae increase and cisternae become more inflated. At the second cytokinesis a dark wall layer is deposited around the tetrasporangium and in the cleavage  furrow.  Starch  grains  Dictyosomes are strongly curved  and chloroplasts  dominate  the cytoplasm.  and young cisternae contain a dark-staining  material where thej' are closely appressed. Cored vesicles are produced by the dictyosomes  and subsequently release their contents forming a whorled fibrillar  layer inside the dark tetrasporangium wall at the second cytokinesis.  This study is presented as a contribution to the taxonomy  and biology of  Gelidiales with a focus on representatives of the order in British Columbia and northern Washington. 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St. John  Plant Science Laboratory, University  of Hawaii,  Honolulu, Hawaii, 111 pp. Santelices, B. 1977. A  taxonomic review of Hawaiian Gelidiales (Rhodophyta).  Pacific Science 31:61-84. Santelices, B.  The  1986.  wild  species of Gelidium  harvest and  culture of  in Chile. FAO  the  economically important  Fisheries Technical Paper N. 281.  pp. 165-192. Santelices, B. unpubl. mscr. Taxonomic studies on Chinese Gelidiales (Rhodophyta). Santelices,  B.  &  Montalva,  S.  1983.  Taxonomic  studies  on  Gelidiaceae  (Rhodophyta) from central Chile. Phycologia 22:185-196. Santelices, B. & Stewart, J.G. 1985. Pacific species of Gelidium other  Gelidiales  (Rhodophyta), with  keys  and  Lamouroux and  descriptions  to the  common or economically important species. In: Taxonomy of Economic Seaweeds, with reference to some Pacific and  Caribbean species. (Ed. by  LA. Abbott and J.N. Norris) pp. 17-31. California Sea Grant College Program, University of California, La Jolla. 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The  genus Polycavernosa  (Gracilariaceae, Rhodophyta): a comparison with Gracilaria  Chang et Xia Grew, and a  key to the species. In: Taxonomy of Economic Seaweeds with Reference to Some Pacific and  Caribbean Species, (Ed. by LA. Abbott and J.N.  Norris), pp. 157-162. California  Sea Grant  Program, University of  California, La Jolla. Xia, B.M., Engzhan, X. & Junfu, Z. 1983. Gelidiaceae. In: Common Seaweeds of China. (Ed. by C.K. Tseng) pp. 66-68. Science Press, Beijing. Yamamoto, H. 1975. The relationship between Gracilariopsis Japan. Bull. Fac. Fish. Hokkaido  and Gracilaria  from  Univ. 26:217-222.  Yamasaki, H. 1960. Studies on the propagation of gelidiaceous algae VI. On the early development and morphogeny in Gelidium Jap. Soc. Sci. Fish. 26:116-122. Yatabe, B. 1892. Iconographia florae japonicae.  Vol. 1, Tokyo.  amansii Lmx.  Bull.  FIGURES Abbreviations Used in Figures.  AP  apical cell  C  carposporophyte  CL  chloroplast  COR  cortex  CPG  carpogonium  CS  carposporangium  D  dictyosome  ER  endoplasmic reticulum  FC  fusion cell  G  gonimoblast  LO  locule  N  nucleus  NC  nutritive chain  NO  nucleolus  OS  ostiole  P  attachment pad  PA  periaxial cell  PC  pit connection  PL  second order plate  PR  prostrate axis  R  rhizine  SMC  spermatangial mother cell 163  / 164 ST  starch grain  SV  spermatangial vacuole  UP  upright axis  W  wall  1°  cell of first order filament (axial file)  2°  cell of second order filament  / 165 Fig. 1. Map of study area and area of enlargement maps in Figs. 2, 3, 4.  166  / 167 Fig. 2. Barkley Sound collection sites, numbered as follows: 1. Brady's Beach, 2. Diplock Is., 3. Dixon Is., 4. Dodger Channel, 5. Geer Islets, 6. Haines Is., 7. Kirby Pt., 8. Meade Islets, 9. Roquefeuil Bay, 10. Ross Islets.  168  / 169 Fig. 3. Southern Vancouver Island, Gulf Islands and San Juan Islands collection sites, numbered  as follows: 11. Madrona Pt., 12. Georgina Pt., 13. False  Narrows, 14. Orlebar Pt., 15. Whalebone Bay.  170  / 171 Fig. 4. Strait of Georgia collection sites, numbered  as follows: 16. Earls Cove,  17. Finn Cove, 18. Frolander Bay, 19. Halfmoon Bay, 20. Denman Is., 21. Ford Cove, 22. Galleon Pt., 23. Helliwell Park, 24. Tribune Bay.  1.72  / 173 Fig. 5. Key to Gelidiales in British Columbia and northern Washington.  la. Plant dark red; mostly of branched, flattened, creeping axes, frequently forming attachment pads at branch nodes; uprights present or absent, never more than 2 cm tall, sparse branches approximately 90° to axis; tetrasporangia in V's...  Pterocladia caloglossoides  lb. Plant red to yellowish-red, dark red to maroon or brown to brownish-red; creeping axes branched, terete, with attachment pads usually not formed at branch nodes; uprights dominant, usuallj' more than 2 cm tall, well branched, branches 30-60° to axis; tetrasporangia not in V's  2  2a. Plant intertidal, brown to brownish-red; forms a turf; usually two orders of branches and many short second order branches; gametophytes not known from British Columbia or northern Washington  Gelidium coulteri  /  2b. Plant shallow subtidal (0-13 m); red to yellowish-red or dark red to maroon; does not form a turf; branching to three to four orders with abundant second, third and fourth order branches; gametophytes rare or common in British Columbia and northern Washington  3  3a. Plant robust, cartilaginous; portions of plant with pyramidal aspect; axes terete to compressed; tetrasporangial stichidia with narrow sterile margin; dioecious; third order filaments of mature cystocarp (between second order plate and pericarp) intact  Gelidium purpurascens  3b. Plant soft and lax; not triangular in aspect; axes compressed to flattened; tetrasporangial stichidia lack sterile margin; monoecious; lacks intact third order filaments in mature cystocarp  Gelidium vagum  / 175 Figs. 6-11. Gelidium  coulteri  plant, scale bar = 5 mm;  vegetative  morphology.  Fig. 6.  habit of infertile  Fig. 7. Apex, apical cell domed and protruding, whole  mount, scale bar = 20 um; Fig. 8. Attachment pads from prostrate axis, whole mount, scale bar = 50 um; Fig. 9. Vegetative upright axis showing cortex, medulla and  rhizines  (arrowhead),  transverse  section,  scale  bar = 50  nm;  Fig. 10.  Vegetative, upright axis, secondary pit-connections (arrows) between inner cortical and medullary cells, rhizines, longitudinal section, scale bar = 20 um; Fig.  11.  Outer cortical cells unoriented, isodiametric, whole mount, scale bar = 20 Mm. Figs. 6, 11 Kirby Pt., Diana Is.; Fig. 7 east side Denman Is.; Figs. 8-10 Haines Is.  \  1  76  / 177 Figs. 12-17. Gelidium coulteri tetrasporophytes. Fig. 12. Habit with tetrasporangial stichidia (arrows), scale bar = 5 mm; whole mount, scale bar = 300 embedded  tetrasporangia  Undivided  tetrasporangium  Fig. 13. Branched tetrasporangial stichidium,  M m ; Fig. 14. Tetrasporangial  (arrows),  whole mount, scale bar = 50  stichidium  with  M m ; Fig.  15.  with lateral pit connection (arrowhead), longitudinal  section, scale bar = 20 M m ; Fig. 16. Various ages of tetrasporangia, longitudinal section, scale bar = 50 transverse  M m ; Fig. 17. Mature, cruciately divided tetrasporangium,  section, scale bar = 20  M m ; Fig. 12 Kirby Pt., Diana Is.; Fig.  Meade Islets; Figs. 14-17 Galleon Pt., Hornby Is.  13  / 179 Figs. 18-20. Gelidium  coulteri male gametophytes. Fig. 18. Spermatangial (arrow)  and  of branch, whole mount, scale bar = 20  vegetative  Spermatangial spermatia  parts mother  cells  (arrowheads),  (large  longitudinal  arrows), spermatangia section,  scale  urn; Fig. 19.  (small  bar=10  um;  arrows) and Fig. 20.  Spermatangial mother cells (arrow) with periclinal wall cutting off spermatangium (arrowhead), transverse  section, scale bar =10  from tetraspores, source: east side Denman Is.  um.  Figs. 18-20  Plants cultured  180  / 181 Figs. 21-24. Gelidium  coulteri habit. Fig. 21. Herbarium specimens UC 494898,  tetrasporophyte, Gardner's collection (#7179) from same tidepool as G. sinicola holotype, Point Cavalo, Marin Co., California, scale bar = 20 um; Fig. 22. Isotype in TCD, tetrasporophyte, scale bar = 20 scale bar=10 mm; scale bar=10  mm.  mm;  Fig. 23. Isotype, AHFH 53929,  Fig. 24. Reproduction of photograph of lectotype in TCD,  182  / 183 Figs. 25-29. Gelidium  purpurascens  habit and vegetative morphology.  Habit of part of tetrasporophyte, scale bar=10 mm;  Fig. 25.  Fig. 26. Vegetative branch  showing cortex, medulla and rhizines (arrows), transverse section, scale bar = 50 Mm; Fig. 27. Habit of basal part of tetrasporophyte showing prostrate axes and tetrasporangial stichidia (arrows), scale bar =10  mm;  Fig. 28. Apex of female  gametophyte branch, apical cell domed, longitudinal section, scale bar =10 Fig.  29. Vegetative  branch, rhizines (arrows) in inner  cortex/outer  Mm;  medulla  transverse section, scale bar = 20 Mm. Figs. 25, 27, 29 Meade Islets, Fig. 26 Orlebar Pt., Gabriola Is., Fig. 28 Kirby Pt., Diana Is.  184  / 185 Figs. 30-34. Gelidium  purpurascens  branch with cortex, medulla and bar = 20  um;  morphology. Fig. 30. Vegetative  rhizines (arrows), longitudinal section, scale  Fig. 31. Surface cortical cells isodiametric, unoriented, equidistant,  whole mount, scale bar = 20 interconnected  vegetative  um;  Fig. 32. Medullary cells of second order plate  by secondary pit connections (arrows), squashed longitudinal section,  scale bar = 30  um;  Fig. 33. Attachment pad  on prostrate axis, whole mount,  scale bar =100  um;  Fig. 34. Attachment pad formed from elongate cortical cells,  longitudinal section of prostrate axis, scale bar = 50  um.  Fig. 30  Orlebar Pt.,  Gabriola Is., Figs. 31, 32 east side Denman Is., Figs. 33, 34 Whalebone Bay, Gabriola Is.  1  86  / 187 Figs. 35-40. Ultrastructure of rhizines in Gelidium purpurascens Figs. 35, 36, 39, 40. Gelidium  purpurascens.  and G. vagum.  Fig. 35. Young rhizines with  convoluted plasma membrane, large vesicles with fibrillar contents (arrows), thick wall, transverse  section, scale bar = 500  nm; Fig. 36. Old rhizine with smooth  plasma membrane profile, cytoplasm vesiculate, membranes thickened, very thick wall, transverse section, scale bar = 500  nm; Figs. 37, 38. Gelidium  vagum. Fig.  37. Young rhizine with vesicles containing vesicular and tubular contents (arrows), some tubular  bodies in wall outside  longitudinal section, scale bar = 200  indented plasma membrane (arrowheads),  nm; Fig. 38. Part of young rhizine with  nucleus and chloroplast (arrow) with inner encircling thylakoid but lacking inner parallel thylakoids, longitudinal section, scale bar = 500 nm; Fig. 39. Young rhizine with dictyosome (arrow) producing vesicles with fibrillar contents and releasing contents to wall (at arrowheads), elongate mitochondrion (M), longitudinal section, scale  bar = 400  nm;  Fig. 40. Old rhizine, cytoplasm  vesiculate, membranes  thickened, plasma membrane profile smooth, longitudinal section, scale bar = 400 nm; Figs. 35, 39 Orlebar Pt., Gabriola  Is., Figs. 36, 40 culture source:  Whalebone Bay, Gabriola Is., Figs. 37, 38 culture source: Tribune Bay, Hornby Is.  188  / 189 Figs.  41-44.  Gelidium  purpurascens  stichidia, habit, scale bar = 500  tetrasporophytes.  Fig. 41. Tetrasporangial  M m ; Fig. 42. Tetrasporangial stichidium with  narrow sterile margin (arrow), whole mount, scale bar = 50 M m ; Fig. 43. Stichidial branch apex, young tetrasporangia with basal pit connections  (arrowheads) and  maturing tetrasporangia with one periclinal division (arrows), longitudinal section, scale bar = 50 M m ; Fig. 44. Cruciately divided tetrasporangium, transverse section, scale bar = 20 M m . Figs. 41, 44 Orlebar Pt., Gabriola Is., Fig. 42 Meade Islets, Fig. 43 Cape Suspiro, Alaska.  1  90  / 191 Figs. 45-48. Gelidium purpurascens stichidium, habit, scale bar =100  female  gametophytes. Fig. 45. Carpogonial  um; Fig. 46. Intercalary carpogonium, with two  pit connections (arrowheads), squashed longitudinal section, scale bar = 5 um; Fig. 47. Apex  with intercalary carpogonia  (all arrows), most proximal carpogonia  sessile (large arrows), longitudinal section, scale bar = 400 Mm; Fig. 48. Cystocarps single and in series on branch, habit, scale bar = 2 mm. Fig. 45 Whalebone Bay, Gabriola Is., Fig. 46 Geer Islets, Figs. 47, 48 Kirby Pt., Diana Is.  / 193 Figs. 49-51. Gelidium  purpurascens  order filament between second section,  scale  carposporophyte  bar =10  um;  concentrated  carposporophytes.  order plate Fig. 50. around  and  Mature second  Fig. 49. Stretched third  cystocarp cortex, longitudinal cystocarp order  with  plate  two  and  locules, producing  carposporangia, longitudinal section through ostiole of upper locule, scale bar = 50 um;  Fig. 51. Detail of carposporophyte,  young carposporangia with  coating (arrows), longitudinal section, scale bar = 25 umFig. 50 Kirby Pt., Diana Is.  mucilage  Figs. 49, 51 Geer Islets,  1 94  / 195 Figs. 52-56. Gelidium  purpurascens  mount, scale bar = 5 mm;  male gametophytes.  Fig. 52. Habit, whole  Fig. 53. Spermatia, whole mount, scale bar = 20 jum;  Fig. 54. Spermatangial stichidium with thickened and ruptured thallus wall over spermatangia,  whole  mount, scale  bar=100  um;  Fig. 55.  Holes  in wall  (arrowheads) through which spermatia have been released, whole mount, scale bar =10  iim; Fig. 56. Spermatangial mother cells (large arrows) and spermatangia  (small arrows) cut off by periclinal wall, longitudinal section, scale bar =10 Fig. 52 Nootka Sound UBC  A53657, Figs. 53, 54, 56 cultured from tetraspores,  source: Whalebone Bay, Gabriola Is., Fig. 55 Esteban Pt., Vancouver A10813.  um.  Is. UBC  196  / 197 Figs. 57-63. Gelidium purpurascens apical and lateral initials, historical and type collections. Fig. 57. Apical cell domed, protruding, whole mount, scale bar =15 um;  Fig. 58. Lateral initial conical (arrow), whole mount, scale bar =15  59. Macoun collection from Departure Bay, scale bar=15 um;  B.C.,  CANA  3740 (was  um; Fig. CAN  74),  Fig. 60. Isodiametric, equidistant, unoriented, surface cortical  cells near base of upright axis, cells branch axis runs left/right in figure, whole mount, scale bar = 50 um;  Fig. 61. Patch of surface cortical cells near base of  upright axis where cells are elliptical and oriented perpendicular  to branch axis,  branch axis runs left/right in figure, whole mount, scale bar = 40 um; Holotype with herbarium labels, UC Mateo Co.,  California, scale bar = 40  93572, cystocarpic, from Moss Beach, San um;  93572, cystocarpic, from Moss Beach, San Um.  Fig. 62.  Fig. 63.  One  holotype plant,  UC  Mateo Co., California, scale bar = 20  Figs. 57, 58, 60, 61 Orlebar Pt., Gabriola Is.  198  / 199 Figs. 64-68. Gelidium  vagum habit and vegetative morphology. Fig. 64. Habit of  gametophyte, scale bar=10 mm; mm;  Fig. 66. Habit  bar =10  mm;  Fig. 65. Habit of tetrasporophyte, scale bar=10  of gametophyte with  Fig. 67. Vegetative  whip-like ultimate branches, scale  branch with  (arrows), transverse section, scale bar = 25  um;  cortex, medulla and  rhizines  Fig. 68. Apical organization in  vegetative branch, apical cell cuts off cells of axial file/first order filament, periaxial cells cut off second order filament, rhizines (arrow) cut off inner cortical cells, longitudinal section in plane of second order plate, scale bar = 50 um. Figs. 64-68 Tribune Bay, Hornby Is.  200  / 201 Figs. 69-76. Gelidium  vagum vegetative and tetrasporophyte morphology. Fig. 69.  Surface cortical cells isodiametric, equidistant, unoriented, from base of upright axis, whole mount, scale bar = 50 um;  Fig. 70. Surface cortical cells isodiametric,  equidistant, unoriented, from upright axis midway between base and apex, whole mount, scale bar = 50 um;  Fig. 71. Vegetative branch with cortex, medulla and  rhizines (arrows) transverse section, scale bar = 30 ym;  Fig. 72. Vegetative branch  showing cortex and medulla (M), longitudinal section, scale bar = 30 urn; Fig. 73. Rhizine (arrow) cut off inner cortical cell, longitudinal section, scale bar =10 Fig. 74. Tetrasporangial bar =100  um;  stichidium with two  Fig. 75. Tetrasporangial  mount, scale bar = 30 um; scale bar = 5 mm.  fertile areas, whole mount, scale  stichidium, lacks sterile margin, whole  Fig 76. Habit of tetrasporophyte with stichidia (arrow),  Figs. 69, 72, 73  Helliwell Park, Hornby Is., Fig. 70 Ford  Cove, Hornby Is., Figs. 71, 75, 76 Tribune Bay, Pt., Hornby Is.  um;  Hornby Is., Fig. 74  Galleon  202  / 203 Figs. 77-83.  Gelidium vagum  reproductive morphology. Fig. 77. Stichidium apex  with young tetrasporangia (arrows), longitudinal section, scale bar = 30 um; Fig. 78. Mature tetrasporangium, cruciately divided, tetraspore nuclei visible, squash preparation, scale bar = 20  um;  Fig. 79. Young tetrasporangium with basal pit  connection (arrowhead), longitudinal section, scale bar = 20 um;  Fig. 80. Holes in  thallus wall (arrows) where tetraspores have been released, whole mount, scale bar = 20  um;  Fig. 81. Undivided  tetrasporangium  with  lateral  pit connection  (arrowhead) and mature divided tetrasporangia, transverse section, scale bar = 30 um;  Fig. 82. Gametangial stichidia, whole mount, scale bar=150 Mm;  Spaces vacated  Fig. 83.  (arrows) by released tetrasporangia, filled with weakly staining  substance, discarded pit plug in lateral position (arrowhead), longitudinal section, scale bar = 20 Mm.  Figs. 77, 81, 82 Galleon Pt., Hornby Is., Figs. 78, 79, 80  Tribune Bay, Hornby Is., Fig. 83 Ford Cove, Hornby Is.  204  / 205 Figs. 84-89. Gelidium scale bar = 4 mm;  vagum gametophytes. Fig. 84. Habit of cystocarpic plant,  Fig. 85. Gametangial area with male gametes (arrowheads)  proximal and young carposporophyte derived from fertilized female gametes distal (arrow), longitudinal section, scale bar = 30 um;  Fig. 86. Spermatangium cut off  spermatangial mother cell by periclinal wall (arrow), transverse section, scale bar =10  um;  Fig. 87. Release of spermatium  transverse section, scale bar =10  um;  (arrow) from  spermatangium,  Fig. 88. Part of young carposporophyte  with nutritive chains (arrows), longitudinal section, scale bar =15  um;  Fig. 89.  Young carposporophyte with furrows created by replacement of cortical cells by carpogonia  (arrowheads),  through  which  carpogonial  trichogynes  protrude,  transverse section, scale bar = 40 um. Figs. 84-89 Tribune Bay, Hornby Is.  206  / 207 Figs.  90-95. Gelidium  vagum  reproductive  morphology. Fig.  90.  Intercalary  carpogonium, lower pit connection out of focal plane, longitudinal section, scale bar=10 Mm; Fig. 91. Third order filaments (arrowheads) stretched between second order plate and pericarp in young, expanding cystocarp, longitudinal section, scale bar=15 Mm; Fig. 92. Fusion cell with two section, scale bar = 75 filaments  Mm; Fig. 93.  absent, longitudinal  section,  narrow lobes (arrows), longitudinal  Mature cystocarp, scale  bar =100  tetrasporophyte from Muroran, Hokkaido, Japan, UBC  stretched Mm;  Fig.  third order 94.  A56807, scale bar=10  Fig. 95. Habit, tetrasporophyte from Hideshima, Iwate-ken, Japan, TNS scale bar=10 mm. Denman Is.  Figs. 90-92 Tribune Bay,  Habit,  Hornby Is., Fig. 87  mm;  25823, east side  / 209 Figs. 96-100. Gelidium vagum Japanese material. Fig. 96. Mature cystocarp and adjacent male gametangial area (arrows, to right), longitudinal section, scale bar = 50  um;  Fig. 97. Mature cystocarp, stretched third order filaments absent  from central part of cystocarp but occasionally present (arrow) at less expanded ends, longitudinal section, scale bar = 50 um; Fig. 98. Tetrasporangial stichidium, sterile margin lacking, whole  mount, scale bar = 50  p.m; Fig. 99. Immature  (arrowheads) and mature (arrow) tetrasporangia, longitudinal section, scale bar = 20 um; Fig. 100. Tetrasporangial stichidium, note embedding of sporangia under two rows  of cortical  cells,  transverse  Ta-no-hama, Iwate-ken, Japan, TNS Japan, UBC A56807.  section,  scale  bar = 50  um.  Figs. 96-98  25824, Figs. 99, 100 Muroran, Hokkaido,  210  / 211 Figs. 101-106. Gelidium johnstonii habit, vegetative and reproductive morphology. Fig. 101. Habit of holotype, CAS  1343  in UC,  tetrasporophytes, Bahia  San  Francisquito, Gulf of California, Baja California del Norte, Mexico, scale bar = 40 mm;  Fig. 102. Vegetative branch with cortical cells in anticlinal rows, medulla  and  rhizines  (arrowheads), transverse  section, scale bar = 50  nm;  Cortical cells in anticlinal rows, rhizines between inner cortical and cells, transverse section, scale bar =100  Fig. 103. medullary  jim; Fig. 104. Tetrasporangial stichidium  with sterile margin, whole mount, scale bar = 20 nm;  Fig. 105. Apical cell and  furrow (arrowheads) of tetrasporophyte branch, whole mount, scale bar = 30 Fig. 106. Habit of tetrasporophyte with stichidia, scale bar=10 mm.  um;  Figs. 104,  105 Puerto Escondido, Gulf of California, Baja California del Sur, Mexico, Fig. 106 Punta Perico, Gulf of California, Mexico.  212  / 213 Figs. 107-109, 111. Gelidium johnstonii  reproductive morphology. Fig. 110.  vagum habit. Fig. 107. Mature cystocarp, longitudinal section, scale bar = 50  G. um;  Fig. 108. Stretched third order filament in cystocarp between second order plate and  pericarp, longitudinal  gametangial  section,  stichidium, note  apical  scale  bar = 40  furrow  and  iim; Fig. 109. apical  cell  Apex  of  slightly recessed  between cortical lobes, whole mount, scale bar = 50 um; Fig. 110. Gelidium vagum habit of drift plant from Ladysmith, B.C., UBC  A64965, scale bar = 40 um; Fig.  111. Mature cystocarp (top) with ostiole (arrow) and adjacent proximal (bottom) spermatangial area (arrowheads), whole mount, scale bar=100 um.  Fig. 107  Puerto Escondido, Gulf of California Baja California del Sur, Mexico, AHFH 50299, Fig. 108 Punta Escondido, Gulf of California, Baja California del Sur, Mexico, Dawson #7170, Fig. 109 Ensenada Bocochibampo, Sinaloa, Mexico, AHFH 4179, Fig. I l l Bahia Aqua Verde, Gulf of California, Baja California del Sur, Mexico, AHFH 502677.  214  / 215 Figs.  112-118. Pterocladia caloglossoides habit and vegetative morphology. Fig.  112.  Habit,  scale bar=10  mm; Fig. 113.  Habit of upright axes, branches  perpendicular to main axis, tetrasporangial, scale bar =150  um; Fig. 114. Habit  of prostrate axes, several upright branches and attachment pad at a node, scale bar = 700 um; Fig. 115. Apex of vegetative branch, apical initial and lateral initial (upper left corner) domed and protruding, files of cells in V's, whole mount, scale bar=100 um; Fig. 116. Bottom view of flared attachment pad on prostrate axis and four radiating upright branches, whole mount, scale bar =100 um; Fig. 117. Upright axis with cortex, medulla and rhizines (arrows), transverse section, scale bar = 30 um; Fig. 118. Vegetative axis with narrow cortex, medulla and rhizines (arrows), transverse section, scale bar = 30 um. Fig. 112 Sea Otter Sound, Alaska, Fig. 113 Orlebar Pt., Gabriola Is., Fig. 114 Barra de Navidad, Jalisco, Mexico, Dawson #3733 slide #1327 (in AHFH) Figs. 115-118 Georgina Pt., Mayne Is.  21 6  / 217 Figs. 119-121.  Pterocladia  caloglossoides  vegetative morphology. Fig. 119. Surface  cortical cells oval, equidistant, unoriented, whole mount, scale bar = 50 um; Fig. 120. Attachment pad on prostrate whole mount, scale bar = 30  Mm;  axis, composed  of elongated cortical cells,  Fig. 121. Attachment pad on prostrate axis  composed of elongated cortical cells, longitudinal section, scale bar = 40 Mm. Figs. 119-121 Georgina Pt., Mayne Is.  21 8  / 219 Figs.  122-128. Pterocladia  Tetrasporangial  caloglossoides  stichidium, tetrasporangia  reproductive  morphology.  Fig. 122.  in Vs, whole mount, scale bar =100  M m ; Fig. 123. Tetrasporangial stichidium, transverse section, scale bar = 40 M m ; Fig. 124. Tetrasporangium with lateral pit connection, squash preparation, scale bar = 20  am;  preparation, material  Fig. 125. Mature,  scale bar = 20  cruciately  divided  tetrasporangium, squash  M m ; Fig. 126. Tetrasporangia  in Vs in Dawson's  (Dawson #8593 slide 1177) from Guadalupe Is., Baja California del  Norte, Mexico, whole  mount, scale  Dawson's  material  (Dawson #8593  California  del Norte, Mexico, whole  bar=100 M m ; Fig. 127. Cystocarp in slide  1177) from  mount, scale  Guadalupe  bar = 40  Is., Baja  M m ; Fig. 128.  Cystocarp with single locule, Dawson's material from Barra de Navidad, Jalisco, Mexico, Dawson #3733  (wet preserved  in AHFH), longitudinal section, scale  bar =30 M m . Figs. 122-125 Georgina Pt., Mayne Is.  220  / 221 Figs.  129-133. Pterocladia  Gelidium  caloglossoides  reproductive morphology.  Fig. 134.  coulteri. Fig. 129. Cystocarp with single ostiole (arrowhead), Dawson  #8593 slide 1177, whole mount, scale bar=100 urn; Fig. 130. Spermatangial area, Dawson #3733 slide 1325, whole mount, scale bar = 20  ym; Fig.  131.  Spermatangia, Dawson #3733 slide 1325, whole mount, scale bar = 40 um; Fig. 132.  Vegetative branch  with  narrow  medulla,  Dawson  #3733  transverse section, scale bar = 20 ym; Fig. 133. Spermatangia  slide  1326,  (arrow), Dawson  #3733 slide 1325, optical section of whole mount, scale bar = 30 ym; Fig. 134. Habit of Macoun's collection of "Gelidium  crinale"  CANA 3474 (was CAN 207), scale bar=10 mm.  from Beacon Hill, Vicotria  Fig. 129 Guadalupe Is., Baja  California del Norte, Figs. 130-133 Barra de Navidad, Jalisco, Mexico.  222  / 223 Figs. 135-140. Historical collections purpurascens,  and type specimens. Fig. 135. Gelidium  Macoun's collection as "G. amansii"  from Departure Bay, CANA  4349 (was CAN 310), scale bar = 20 mm; Fig. 136. G. robustum, collection from "Shoal B. Victoria", UBC  A7861, scale bar = 50  mm;  Fig. 137. G. coulteri,  Macoun's collection as "G. crinale" from Beacon Hill, Victoria, CANA 3474 (was CAN  207), scale bar = 50 mm; Fig. 138. G. robustum holotype, UC 294572, near  Ensenada, Baja California del Norte, Mexico, scale bar = 50 mm;  Fig. 139. G.  sinicola holotype, UC 276620, Point Cavalo, Marin Co., California, scale bar=50 mm;  Fig. 140. G. sinicola  holotype (UC 276620), vegetative  transverse section, scale bar = 30  mm.  branch, terete,  224  / 225 Fig.  141.  Polysiphonia-type  life  gametophyte and carposporophyte and  gametophyte,  history.  Triphasic  (with  tetrasporophyte,  phases) life history, isomorphic tetrasporophyte  carposporophyte  developing  in  female  gametophyte  axis,  magnified cystocarp shown in longitudinal section; solid (black) structures diploid, hollow (white) structures haploid; not drawn to scale.  226  male gametophyte N  / 227 Figs.  142-153. Gelidium  Ungerminated  vagum  sporeling  development  in culture. Fig. 142.  tetraspore; Fig. 143. Germ tube (arrow) initiation,  gametophyte;  Fig. 144. Germ tube enlargement, gametophyte; Figs. 145, 146. Evacuation of cytoplasm  from  spore  into  germ  tube  and  wall  cutting  off germ  tube,  tetrasporophytes; Fig. 147. First unequal division of germ tube into concave (larger) and fusiform (smaller) cells, tetrasporophytes; Fig. 148. Division of germ tube  into  concave  (right) and  fusiform  (left) cells, gametophyte;  Fig. 149.  Transversely divided concave cell, most distal cell of group initiating primary rhizoid, tetrasporophyte; Fig. 150. Variety of developmental stages, original spores all  from  same cystocarp, tetrasporophytes; Fig. 151. Elongation  of primary  attachment rhizoid, originated from cell in concave group, tetrasporophyte; Fig. 152. Elongated rhizoid, gametophyte; Fig. 153. Second attachment rhizoid produced (before apical cell initiation), gametophyte. Figs. 142-153 scale bar = 30  jum. Figs.  142, 143 culture source: Tribune Bay, Hornby Is., Figs. 144, 148, 149, 153 culture source: Tribune Bay, Hornby Is., Figs. 145-147, 150, 151 culture source: east side Denman Is., Fig. 152 culture source: Ford Cove, Hornby Is.  / 229 Figs. 154-162. Gelidium vagum sporeling development in culture. Fig. 154. First division of fusiform cell (arrow), tetrasporophyte; Fig. 155. Fusiform and concave cell groups distinct, tetrasporophyte; Fig. 156. Fusiform (arrowhead) and concave (arrow) cell groups distinct, tetrasporophytes; Fig. 157. Apical cell (arrowhead) formation  opposite  rhizoid, tetrasporophyte;  Fig. 158  Apical cell  (arrowhead)  formation from fusiform group cell, tetrasporophyte; Fig. 159. Sporeling increased in  size,  with  apical  organization, tetrasporophyte;  Fig. 160.  sporelings  with  apical  organization; Fig. 161. Attachment  pad  Gametophyte formed from  adhesion of elongated cortical cells, tetrasporophyte; Fig. 162. Branch initiation (at arrow), gametophyte. Figs. 154, 155, 157-161 scale bar = 30 um, Figs. 156, 162 scale bar = 40 um.  Figs. 154, 155 culture source: east side Denman Is., Figs.  156-159 culture source: Galleon Pt., Hornby Is., Figs. 160-162 culture source: Tribune Bay, Hornbj' Is.  / 231 Figs. 163-168. Gelidium vagum reproduction in culture. Fig. 163. Tetrasporangial stichidia  on tetrasporophyte  from carpospores, scale bar = 400  um;  Fig. 164.  Gametangial stichidium on gametophyte grown from a tetraspore, scale bar =100 um; Fig. 165. Gametangial stichidia on gametophyte from a tetraspore, scale bar = 700  um; Fig. 166. Tetraspores being released from tetrasporophyte grown  from a carpospore, scale bar = 500  um;  Fig. 167. Germination  of tetraspores  released from tetrasporophyte grown from carpospore, scale bar = 300  um; Fig.  168. Fertile stichidia on tetrasporophyte grown from carpospore, originally grown from tetraspore, scale bar = 500  um. Figs. 163, 166, 167 culture source: east  side Denman Is., Fig. 164, 165 culture source: Galleon Pt., Hornby Is., Fig. 168 culture source: Tribune Bay, Hornby Is.  232  / 233 Figs. 169-173. Reproduction of Gelidium purpurascens  and G. coulteri in culture.  169. Gelidium purpurascens,  tetrasporangial stichidia from field, excised for  tetraspore release, scale bar = 200  um; Fig. 170. G. purpurascens, habit of fertile  Fig.  male gametophyte grown from tetraspore, scale bar = 500 purpurascens, bar = 500  um;  um;  Fig. 171. G.  habit of fertile female gametophyte grown from tetraspore, scale Fig. 172. G. coulteri, spermatangial area of male gametophyte  grown from tetraspore, scale bar = 200 um; Fig. 173. G. coulteri, habit of fertile male gametophyte grown from tetraspore, scale bar = 500 um. Fig. 174 Orlebar Pt., Gabriola Is., Figs. 170, 171 culture source: Whalebone Bay, Gabriola Is., Figs. 172, 173 culture source: east side Denman Is.  / 235 Figs. 174-177. Chromosomes of Gelidium vagum. Figs. 174, 176, 177. Prophase I of undivided tetrasporangia, hematoxylin stained, Tribune Bay, Hornby Is.; Figs. 174-176 n= 14-15, Fig. 175. drawing of nucleus in Fig. 174; Fig. 177. five large chromosomal bodies; scale bar = 50 um for all figures.  / 237 Figs.  178-183.  purpurascens  Carpogonium  and  carposporophyte development  and G. vagum. Figs. 178. Gelidium  in  vagum, intercalary carpogonium,  longitudinal section, scale bar=10 um; Figs. 179-183 G. purpurascens. Sessile carpogonium  longitudinal section, scale bar =10 with cortical  Fig. 179.  (arrow) with curved basal wall, longitudinal section, scale  bar=10 um; Fig. 180. Sessile carpogonium  fusing  Gelidium  (arrow) vacuolate and degenerate,  um; Fig. 181. Fertilized carpogonium (arrow)  cells through two pit connections (arrowheads), showing  functional carpogonium is intercalary, squash of longitudinal section, scale bar =10 um; Fig. 182. Multilobed, multinucleate fusion cells, squash of longitudinal section, scale bar=10 um; Fig. 183. Young carposporophyte, clusters of nutritive chains around second order plate cells and fusion cell (arrow), longitudinal section, scale bar = 300 um. Fig. 178 Ta-no-hama, Iwate-ken, Japan, TNS 25824, Figs. 179, 180, 183 Kirby Pt., Diana Is., Figs. 181, 182 Geer Islets.  238  / 239 Figs. 184-188. Carposporophyte vagum.  Figs.  184,  185.  Gelidium  purpurascens.  longitudinal section, scale bar=10 um; filaments vagum.  (arrows), longitudinal Fig. 186.  Lobe  in Gelidium  development  purpurascens  Fig. 184.  One  locule  of  Fig. 185. Fusion cell and gonimoblast  section, scale  of gonimoblast  mature  G.  Nutritive chains,  cell  bar = 20  ym;  Fig. 186-188. G.  (arrow) fused with  (arrowhead) of (upper) nutritive chain, longitudinal section, scale bar =10 187.  and  cystocarp  showing  apical cell um; Fig.  carposporophyte  producing  carposporangia, longitudinal sections through ostiole, scale bar = 50 um;  Fig. 188.  Gonimoblast cells producing carposporangia terminally section, scale bar=10 um.  and  laterally, longitudinal  Figs. 184, 185 Geer Islets, Figs. 186, 188 Tribune  Bay, Hornby Is., Fig. 187 Ta-no-hama, Iwate-ken, Japan, TNS 25824.  240  / 241 Figs. 189-192. Ultrastructure of spermatiogenesis in Gelidium Spermatangial  mother cells (arrow) and  section, scale bar=10 um;  spermatangia  Fig. 190. Spermatangia  vagum. Fig. 189.  (arrowhead), longitudinal  (small cells) and vegetative  cortical cells (larger), whole mount, scale bar = 30 um; Fig. 191. Ultrastructure of vegetative cortical cell with large plastids, basal nucleus (arrowhead) and central vacuole, scale bar=1000 nm;  Fig. 192.  Ultrastructure  of two  spermatangial  mother cells each with a pit connection to base of a third spermatangial mother cell, which has a pit connection to a cortical cell, scale bar = 1000  nm.  Figs.  189, 192 Tribune Bay, Hornby Is., Fig. 190 Galleon Pt., Fig. 191 east side Denman Is.  242  / 243 Figs. 193-196. Ultrastructure of spermatiogenesis in Gelidium  vagum. Fig. 193.  Spermatangial mother cell with elongate, central nucleus and furrow (arrowheads) possibly excluding apical chloroplast; Fig. 194. Spermatangial mother cell with two nuclei;  Fig. 195. Spermatangium  with  two  prominent  dictyosomes  (arrows)  contributing vesicles to young spermatangial vacuole, mitochondrion (M); Fig. 196. Spermatangium with basal pit connection and young spermatangial vacuole, cored vesicles (arrowheads), large nucleus; scale bar = 500 193-196 Tribune Bay, Hornby Is.  nm  for all figures. Figs.  244  / 245 Figs. 197-200. Ultrastructure of spermatiogenesis in Gelidium purpurascens and G. vagum. Figs. 197,  198. Gelidium  vagum. Fig. 197. Spermatangium with  2-3  spermatangial vacuoles and large nucleus with condensed chromatin (dark areas), scale  bar = 300  condensed  nm;  chromatin  Fig. 198. (dark  purpurascens. Fig. 199. Two  Spermatangial  areas),  scale  vacuole  bar = 300  nm;  release, Figs.  nucleus 199,  200  with G.  spermatangial vacuoles released from spermatangium,  spermatangium with many cored vesicles (arrows), scale bar = 800 nm;  Fig. 200.  Spermatangial vacuole, contents whorled fibrillar, scale bar = 300 nm.  Figs. 197,  198 Tribune Bay, Hornbyls., Figs. 199, 200 cultured Whalebone Bay, Gabriola Is. material.  2 46  / 247 Figs. 201-204. Ultrastructure of spermatiogenesis in Gelidium purpurascens vagum. many  Figs.  201,  cored  204.  vesicles  Gelidium  (arrows),  purpurascens.  mitochondria  Fig. 201.  (M)  and  and G.  Spermatangium  nucleus  with  with  condensed  chromatin (dark areas), scale bar = 40 nm; Figs.  202, 203 G. vagum. Fig. 202.  Spermatangium  starting  spermatangial  released,  cored vesicles  many  spermatium,  scale  release  bar = 400  (arrows), spermatangial Spermatium  to  being  spermatium,  (arrows)  nm;  Fig. 203.  vacuoles already  released  and  and  thick mucilage coating at Spermatangium  released,  wall  vacuoles  with  already apex of  cored vesicles  scale bar = 500 nm; Fig. 204.  material  (arrowhead)  secreted  by  spermatangial mother cell, spermatangial mother cell on right at cytokinesis, scale bar = 700  nm.  Figs.  201,  204  cultured  Figs. 202, 203 east side Denman Is.  Whalebone  Bay, Gabriola  Is.  material,  248  / 249 Figs. 205-208. Ultrastructure of spermatiogenesis in Gelidium purpurascens and G. vagum.  purpurascens.  Figs. 205-207.Gelidium  Fig. 205. Release  spermatangia, spermatangial mother cell secreting new previous  wall  layer  (arrows)  spermatangium, scale bar = 800  surrounding nm;  of percurrent  wall (arrowheads), note  spermatangial  mother  cell  and  Fig. 206. Release of conical spermatium,  large nucleus and many cored vesicles (arrows), scale bar = 500 nm; Fig. 207. Spermatangial  mother  cells  and  old wall  layers  (arrowheads) of percurrent  spermatangia, scale bar = 2000 nm; Fig. 208. G. vagum spermatangial mother cell with  old wall  spermatangia,  layers scale  and  discarded  bar=1000  nm.  pit plugs  Figs.  (arrowheads) of percurrent  205-207 cultured  Gabriola Is. material, Fig. 208 Tribune Ba3', Hornby Is.  Whalebone  Bay,  250  / 251 Fig. 209. Summary of spermatiogenesis in Gelidium.  a-c. Longitudinal division of  an outer cortical cell to form 4 spermatangial mother cells; d-j. Summary of ultrastructural development of the spermatangium, d. Spermatangial mother cell (smc) with basal nucleus (n), starch grains (st) and chloroplasts (cl) throughout cytoplasm; e. Apical chloroplast (cl) of spermatangial mother cell (smc) being cut off  from  rest  of cell  transversely  divided  dictyosome  (arrowhead)  Spermatangium  by  furrows  to form  a spermatangium  and  (sp) with  (arrows); f. Spermatangial  vesiculated 2  (sp), spermatangium  endoplasmic  spermatangial  mother cell  reticulum  vacuoles  with a  (arrow); g.  (sv) developed  from  dictyosome-derived vesicles and endoplasmic reticulum, nucleus (n) with condensed chromatin; h. Spermatangial  vacuoles (sv) released  from  the spermatangium,  spermatangium with cored vesicles (cv) and an external layer of mucilage-like material (m); i. Spermatium (s) released from spermatangium, spermatium with cored vesicles, a layer of mucilage-like material and nucleus with chromatin;  j. Spermatangial  mother  cell  expanded  into  space  spermatium to form another spermatangium; drawings not to scale.  condensed  vacated by  252  / 253 Figs. 210-215. Ultrastructure of tetrasporogenesis in Gelidium  purpurascens. Fig.  210. Remains of vacuole (V) in very young tetrasporangium (before expansion of nucleus  and  Membranous  thickening of vegetative wall), scale bar = 2000 nm; material  (arrows)  extruded  from  pre-meiotic  Fig. 211.  tetrasporangium,  mitochondrion (M), scale bar = 400 nm; Fig. 212. Different section of very young tetrasporangium of Fig. 210, material from vacuole (V) incorporated in vesicles (arrows) and  released to wall, scale  bar = 400  nm;  Fig. 213.  Nucleus  of  pre-meiotic tetrasporangium with perinuclear endoplasmic reticulum (small arrows), many nuclear pores (arrowheads), large nucleolus, large nucleolar vacuoles (large arrows), scale bar=500 nm;  Fig. 214. Pre-meiotic tetrasporangium with central  nucleus, large nucleolus, lenticular starch grains (arrowheads) central around the nucleus, scale bar=2000 nm;  Fig. 215. Pre-meiotic tetrasporangium, nucleus and  nucleolus large, central, chloroplasts mostly  peripheral, lenticular starch grains  (arrows) around nucleus, scale bar=1000 nm; Figs. 210, 212, 213 Georgina Pt., Mayne Is., Figs. 211, 214, 215 Tribune Bay, Hornby Is. culture.  254  / 255 in Gelidium purpurascens. Fig.  Figs. 216-220. Ultrastructure of tetrasporogenesis 216.  Convoluted plasma membrane (arrowhead) of pre-meiotic  small transparent  vesicles (arrows) of unknown origin release their contents to  wall, several mitochondria (M), scale bar = 200 of  young  tetrasporangium,  tetrasporangium, central  nm; Fig. 217. Transverse section  nucleus  (arrow) dumbell  shaped, possibly  undergoing meiosis, many chloroplasts, lenticular starch grains (arrowheads), scale bar = 2000  nm;  Fig. 218. Spindle-shaped,  tetrasporangium with large nucleolus, small  single  nucleus  of  pre-meiotic  amount of perinuclear endoplasmic  reticulum (arrowheads), lenticular starch grains, mitochondrion (M), scale bar = 400 nm;  Fig. 219. Elongate  nucleolus,  perinuclear  endoplasmic  (arrowheads), scale bar = 500 first meiotic  division  nucleus  of pre-meiotic reticulum  tetrasporangium  (arrows),  lenticular  with  starch  large grains  nm; Fig. 220. Two nuclei in tetrasporangium after  and before cytokinesis, many  ovate chloroplasts, many  lenticular to ovate starch grains, grey wall layer (layer 2) present (arrow) inside layer 1 wall (W), scale bar = 3000 nm; Fig. 216 Georgina Pt., Mayne Is., Figs. 217-220 Tribune Bay, Hornby Is., culture.  256  / 257 Figs. 221-226 Ultrastructure of tetrasporogenesis  in Gelidium  purpurascens. Fig.  221. Flat dictyosome associated with mitochondrion (M) in tetrasporangium about the  time  of meiosis,  bar = 200  nm;  post-meiotic present  dictyosome  cisternae  Fig. 222. Vesiculate  endoplasmic  tetrasporangium, vesicles contain  (W2), scale  bar = 400  nm;  contains  fibrillar  reticulum  fibrillar  material, scale at periphery  material,  Fig. 223. Part  of  wall a  of  layer 2  post-meiotic  tetrasporangium with many chloroplasts and starch grains, peripheral dictyosomes flat with many cisternae per stack, mitochondrion (M), scale bar = 800  nm; Fig.  224. Tetrasporangium at about the time of karyokinesis, many chloroplasts, many lenticular to ovate starch grains, wall layer 2 (W2) present, scale bar = 3000 nm; Fig. 225. Peripheral cytoplasm of post-meiotic tetrasporangium, many dictyosomes and layer  associated mitochondria (M), vesiculate endoplasmic reticulum 2, scale  bar = 400  nm;  Fig. 226. Tetrasporangium  (arrows), wall  at the time of  cytokinesis, wall layer 3 (arrow) present interior to layer 2 (arrowhead), many ovate  starch  grains, many  chloroplasts, scale bar =4000 nm; Figs. 221-226  Tribune Bay, Hornby Is., culture.  258  / 259 in Gelidium  Figs. 227-232. Ultrastructure of tetrasporogenesis purpurascens.  Figs.  227,  228,  230-232  Gelidium  coulteri and G.  purpurascens.  Fig. 227.  Tetrasporangial wall layers at cytokinesis, layer 1 (Wl), layer 2'(arrow), layer 3 (arrowhead), scale bar = 200 nm; Fig. 228. Cytoplasm cytokinesis, starch  grains  ovate, large dictyosome  mitochondrion (M), scale bar = 400  of tetrasporangium after  curved  nm; Fig. 229. Gelidium  and associated  with  coulteri, chloroplasts  radiating around nucleus, starch grains between chloroplasts, scale bar =1000 nm; Fig.  230. Strongly  post-cytokinesis  curved  dictyosome  tetrasporangium,  dark  associated material  with  mitochondrion  (M) in  (arrowhead) between appressed  younger cisternae, cored vesicles (arrows) produced at dictyosome maturing face, scale bar = 200  nm;  Fig. 231. Cored  vesicles (arrows) releasing contents to  cleavage furrow (F) of tetrasporangium, scale bar = 400 nm; Fig. 232. Wall of mature tetrasporangium (after cytokinesis), layer 1 (Wl), layer 2 (black arrow), layer 3 (arrowhead), mucilage-like  layer (white arrow), scale bar = 60 nm; Figs.  227, 228, 230-232 Tribune Bay, Hornby Is., culture, Fig. 229 Brady's Beach, Bamfield.  260  / 261  TABLES  Table  1.  Historical  summary  of  genera  included  in  Gelidiaceae  (horizontal  continuation of table on next page).  Kiitzing, 1843  Kiitzing, 1849  J. G. Agardh,  Schmitz, 1889  1876 Acrocarpus  Acrocarpus  Gelidium  Acropeltis  Ctenodus  Chondrodon  Pterocladia  Atractophora  Echinocaulon  Delisea  Ptilophora  Binderella  Gelidium  Echinocaulon  Suhria  Caulacanthus  Euctenodus  Wurdemannia  Choreocolax  Gelidium  Gelidium  •  Phacelocarpus  Harveyella  Polycladia  Naccaria  Thysanocladia  Porphyroglossum Pterocladia Ptilophora Schottmullera Suhria Wrangelia  262  Kylin, 1956  Fan, 1961  current, 1988  Acanthopeltis  Acanthopeltis  Acanthopeltis  Acropeltis  Acropeltis  Beckerella  Beckerella  Beckerella  Gelidiella  Gelidiella  Gelidium  Gelidium  Gelidiocolax  Porphyroglossum  Onikusa  Gelidium  Pterocladia  Porphyrogloss um  Porphyroglossum  Ptilophora  Pterocladia  Pterocladia  Suhria  Pterocladias trum  Ptilophora  Yatabella  Ptilophora  Suhria  Suhria  Yatabella  Yatabella  / 264 Table 2. Sites and dates of field collections of Gelidiales from the study area.  Site  Latitude  Longitude  Dates collected  Brady's Beach,  48°50'N  125°09'W  24/vi/86  48°56'N  125° 07'W  28/viii/84  48°51'N  125°07'W  27/viii/84  48°51'N  125°12'W  29/viii/84  48°56'N  125°07'W  28/viii/84  48°50'N  125° 12'W  29/viii/84  48°51'N  125°13'W  29/viii/84, 27/viii/85,  Barkley Sound Diplock Island (NE corner), Barkley Sound Dixon Island (SE corner), Barkley Sound Dodger Channel, Barkley Sound Geer Islets, Barkley Sound Haines Island, Barkley Sound Kirby Point, Diana  23/vi/86  Island, Barkley Sound Meade Islets, Barkley  48°56'N  125°07'W  28/viii/84  48°52'N  125°07'W  27/viii/84  Sound Roquefeuil Bay, Barklej' Sound  /  Ross Islets, Barkley  48°52'N  125°10'W  28/viii/86  48°41.5'N  122°54'W  24M/85  48°52'N  123°17'W  16/xi/85, 14/ii/86,  Sound Madrona Point, Orcas Island, Gulf of Georgia Georgina Point,  21/ix/86, 13/viii/87  Mayne Island, Gulf of Georgia False Narrows,  49°08'N  123°49'W  8/x/85  49°12'N  123"49'W  8/x/85, ll/xii/85,  Gabriola Island, Strait of Georgia Orlebar Point, Gabriola Island, Strait  28/i/86, 3/iii/86,  of Georgia  7/v/86, 4/vi/86, 21M/86, 29/viii/86, 28/x/86, 17/U/87, 5/iii/87, 5/viii/87  Whalebone Bay,  49°11'N  123°48'W  28/x/86, 27M/86, 5/viii/87  Gabriola Island, Strait of Georgia Earls Cove, Strait of  49°45'N  124°00'W  6/iii/85  49°59'N  124°46'W  6/xi/83  Georgia Finn Cove, Strait of Georgia  / 266 Frolander Bay, Strait  49°45'N  124°17'W  5/xi/83  49°30'N  123°56'W  6/iii/85  49°30.5'N  124°44'W  29/ix/86, 26/viii/87  of Georgia Halfmoon Bay, Strait of Georgia Denman Island (E side), Strait of Georgia Ford Cove, Hornby  49°30'N  Island, Strait of Georgia 124°41'W Galleon Point, Hornby  l/x/86  49°33'N  124°40'W  30/ix/86  49°31'N  124°35'W  23/iv/87  49°32'N  124°38'W  Island, Strait of Georgia Helliwell Park, Hornby Island, Strait of Georgia Tribune Bay, Hornby Island, Strait of Georgia  15/x/84, 30/ix/86  / 267 Table  3.  Herbaria  from which  loans were obtained  and their abbreviations  (Holmgren et al, 1981).  Herbarium  Abbreviation  Allan Hancock Foundation, collection housed in LAM  AHFH  National Museum of Canada, Ottawa  CANA  California Academy of Sciences, collection housed in UC  CAS  Friday Harbor Laboratories, Friday Harbor  FHL  Los Angeles Countj' Museum, Los Angeles  LAM  School of Botany, Trinity College, Dublin  TCD  National Science Museum, Tokyo  TNS  University of British Columbia, Vancouver  UBC  University of California, Berkeley  UC  University of Washington, Seattle  WTU  / 268 Table  4. Species of Gelidiales  recorded from  southeast Pacific Ocean areas. Numbered  the northeast, northwest and  superscripts refer to references as  follows: 1. Akatsuka, 1986b 2. Dawson, 1944 3. Dawson, 1953 4. Dawson et al., 1964 5. Santelices, 1977 6. Santelices and Montalva, 1983 7. Santelices and Stewart, 1985 8. Schnetter and Bula Meyer, 1982 9. Stewart, 1976 10. Stewart and Norris, 1981 11. Bangmei et al., 1983  Taxon  Japan  China  Hawaii  Mexico  Central  & Calif-  and  ornia  South America  Acanthopeltis japonica Okamura  •  Beckerella irregularis Akatsuka et Masaki Beckerella subcostata (Okamura) Kylin Gelidiella  acerosa (Forsskal)  Feldmann et Hamel Gelidiella  adnata Dawson  Gelidiella  hancockii Dawson  Gelidiella  ligulata Dawson  Gelidiella  machristiana  Gelidiella  mexicana Dawson  Gelidiella  myrioclada Borgesen  Gelidiella  ramellosa (Kiitzing)  Dawson  Feldmann et Hamel Gelidiella  refugiensis Dawson  Gelidiella  stichidiospora  Gelidium  amamiense Tanaka  Gelidium  amansii (Lamouroux)  Dawson  Lamouroux Gelidium  arborescens Gardner  Gelidium  bulae Schnetter  Gelidium  cartilagineum (L.)  Greville Gelidium  chilense (Montalva)  Santelices et Montalva  Gelidium  corneum (Hudson)  Lamouroux Gelidium coronadense Dawson Gelidium  coulteri Harvey  Gelidium  crinale (Turner)  Lamouroux Gelidium  deciduum Dawson  Gelidium  decumbensum Okamura  Gelidium  divaricatum  Gelidium  isabelae Taylor  Gelidium japonicum  vonMartens  (Harvey)  Okamura Gelidium johnstonii et  Setchell  Gardner  Gelidium  kintaroi (Okamura)  Yamada Gelidium  latifolium  (Greville)  Bornet Gelidium  lingulatum  Gelidium  linoides Kutzing  Gelidium  nanum Inagaki  Gelidium  nudifrons Gardner  Gelidium pacificum Gelidium planisculum  Kutzing  Okamura Okamura  Gelidium pluma Loomis  Gelidium polycladum  Kutzing  Gelidium polystichum Gardner Gelidium pristoides Turner Gelidium  pseudointricatum  Skottsberg et Levring Gelidium pulchrum  Gardner  Gelidium purpurascens Gardner Gelidium pusillum  (Stackhouse)  LeJolis Gelidium  reediae Loomis  Gelidium  rex Santelices et Abbott  Gelidium  rigens Greville  Gelidium  robustum (Gardner)  Hollenberg et Abbott Gelidium  sclerophyllum Taylor  Gelidium  subfastigiatum Okamura  Gelidium  sinicola Gardner  Gelidium. vagum Okamura Gelidium yamadae (Okamura) Fan Onikusa japonica  (Harvey)  Akatsuka Pterocladia  bulbosa Loomis  Pterocladia  caerulescens (Kutzing)  Santelices  Pterocladia caloglossoides (Howe) Dawson Pterocladia capillacea (Gmelin) Bornet et Thuret Pterocladia densa Okamura Pterocladia mcnabbiana Dawson Pterocladia media Dawson Pterocladia nana Okamura Pterocladia tenuis Okamura Yatabella hirsuta Okamura  /  Table 5. Differences between Gelidium purpurascens and Gelidium  robustum  Character  Gelidium purpurascens  Gelidium  robustum  height  usually not > 15 cm  commonly >15 cm  main axis  1 - several  1  branching  dense, branches close  coarse, loose, branches  together, separated by  separated by at least  1-2 mm  mm  higher order branch bases  not constricted  constricted  inner cortex  cells unordered  cells ordered in pallisade-like rows  tetrasporangial stichidium  narrow sterile margin  wide sterile margin  / 274  Table 6. Anatomical differences between Gelidium johnstonii and Gelidium vagum  Characters  Gelidium johnstonii  Gelidium  vagum  vegetative: outer cortical  in anticlinal files  not in anticlinal files  shape  spatulate  ovate, tapered apically  apex  blunt  pointed  sterile margin  present  absent  apical cell  even or recessed  protruding  apical furrow  present  absent  flattened  yes  no - at most compressed  intact, stretched  not intact, broken  cells tetrasporangial stichidia:  cystocarp: third order filaments  APPENDIX  1. HERBARIUM SPECIMENS  gametophyte,  G  =  gametophyte,  M  EXAMINED  F  =  female  =  male  gametophyte,  T  =  tetrasporophyte, specimens without a letter designation are vegetative  Gelidium coulteri  ISOTYPE  AHFH 53929  California  CANA 3474  Beacon Hill, Victoria  FHL 3055  Cape Alava, Washington  TCD  California  photograph of LECTOTYPE  TCD  California  ISOTYPE  UBC A906  Monterey, California  T  UBC A907  Monterey, California  UBC A1449  Moss Beach, California  UBC A2793  Amos Is., Kyuquot, Vancouver Is.  UBC A4890  Mukkaw Bay, Washington  UBC A10812  Mills Peninsula, Vancouver Is.  UBC A10814  Esteban Point, Vancouver Is.  UBC  A11940  McLean Is.. Vancouver Is.  UBC A11947  Spring Is., Kyuquot, Vancouver Is.  UBC A12073  Miracle Beach, Vancouver Is.  UBC A12294  Spring Is., Kyuquot, Vancouver Is. 275  UBC A12298  McLean Is., Vancouver Is.  UBC A12299  McLean Is., Vancouver Is.  UBC A12300  McLean Is., Vancouver Is.  UBC A13259  Perez Rocks, Vancouver Is.  UBC A13478  Walters Cove, Vancouver Is.  UBC A13658  La Jolla, California  UBC A14403  Mukkaw Bay, Washington  UBC A15173  Malaspina Narrows, Queen Charlotte Strait  UBC A19739  Brooks Peninsula, Vancouver Is.  UBC A24527  Pescadero Point, San Mateo Co., California  UBC A24837  Mukkaw Bay, Washington  UBC A28351  Orlebar Point, Gabriola Is., B.C.  UBC A31363  Decanso Bay, Gabriola Is., B.C.  UBC A31426  Davison's Beach, Gabriola Is., B.C.  UBC A36197  Brooks Peninsula, Vancouver Is., B.C.  UBC A36198  False Narrows, Gabriola Is., B.C.  UBC A36199  Lock Bay, Gabriola Is., B.C.  UBC A36372  Jackobson Point, Brooks Peninsula, Vancouver Is., B.C.  UBC A36373  Bunsby Is., Vancouver Is., B.C.  UBC A36374  Bunsby Is., Vancouver Is., B.C.  UBC A36545  Amos Is., Vancouver Is., B.C.  UBC A36711  Bunsby Is., Vancouver Is., B.C.  UBC A36712  Bunsby Island, Vancouver Is., B.C.  UBC A36713  Bunsby Is., Vancouver Is., B.C.  / 277 UBC A37077  Fossil Beach, Grassy Is., B.C.  UBC A37078  Fossil Beach, Grassy Is., B.C.  UBC A37079  Grassy Is., B.C.  UBC A37219  Yellow Bluff, Vancouver Is., B.C.  UBC A37220  Yellow Bluff, Vancouver Is., B.C.  UBC A37221  Fossil Beach, Grassy Is., B.C.  UBC A37223  Yellow Bluff, Vancouver Is., B.C.  UBC A37402  Pincer Is., Vancouver Is., B.C.  T  UBC A37403  Pincer Is., Vancouver Is., B.C.  T  UBC A37404  Nootka Is., Vancouver Is., B.C.  T  UBC A38472  Lawn Point, Vancouver Is., B.C.  UBC A38473  Lawn Point, Vancouver Is., B.C.  UBC A38474  Lawn Point, Vancouver Is., B.C.  UBC A39128  Arab Cove, Vancouver Is., B.C.  UBC A40404  Mukkaw Bay, Washington  UBC A40405  Mukkaw Bay, Washington  UBC A40415  Crescent Beach, Clallam Co., Washington  UBC A40467  Diana Is., Barkley Sound, B.C.  UBC A40468  Diana Is., Barkley Sound, B.C.  UBC A40469  Diana Is., Barklej' Sound, B.C.  UBC A40470  Diana Is., Barkley Sound, B.C.  UBC A42814  Lawn Point, Vancouver Is., B.C.  UBC A43519  Fleming Is., Barkley Sound, B.C.  UBC A43520  Reeks Is., Barkley Sound, B.C.  UBC A43521  Aguilar House Beach, Bamfield, B.C.  T  T  / 278 UBC A43522  Aguilar House Beach, Bamfield, B.C.  UBC A43523  Effingham Is., Barkley Sound, B.C.  UBC A47159  S. California (L.A. Co.)  UBC A50505  Mussel Point, Monterey, California  UBC A53814  Kirby Point, Diana Is., Barkley Sound, B.C  UBC A53975  Brooks Peninsula, Vancouver Is., B.C.  UBC A60236  Aguilar House Beach, Bamfield, B.C.  UBC A60254  Diana Is., Barkley Sound, B.C.  UBC A60268  Execution Rock, Barkley Sound, B.C.  UBC A60460  Aguilar House Beach, Bamfield, B.C.  UBC A61984  Pigeon Point, California  UBC A62016  Montana de Oro, California  UBC A64934  Ladysmith, B.C.  UC 305332  Moss Beach, Pacific Grove, California  WTU  246345  Steamboat Is., Thurston Co., Washington  WTU  246351  Mukkaw Bay, Washington  Gelidium johnstonii  AHFH 69  San Francisquito Bay, Baja California, Mexico  T, ISOTYPE  AHFH 2211  Pond Is., off Angel de la Guardia Is.,  G  Mexico AHFH 4150  Ensenada Bocochibampo, near Guayamas, Mexico  T  / 279 AHFH 4156  Bahia Carrizal, near Cabo Circo, Sonora, Mexico  AHFH 4179  Ensenada Bocochibampo, Sonora, Mexico  AHFH 4192  Isla Jorge, Gulf of California, Mexico  AHFH 4193  Isla Patos, near Isla Tiburon, Gulf of  G  California, Mexico AHFH 4194  Isla Patos, near Isla Tiburon, Gulf of California, Mexico  AHFH 50267  Bahia Aqua Verde, Baja California, Mexico  G,T  AHFH 50268  Punta Perico, Salinas Bay, Carmen Is., Gulf  G,T  of California, Mexico AHFH 50299  Puerto Escondido, Baja California, Mexico  G,T  CAS 1343 in  San Francisquito Bay, Baja California, Mexico  G, T HOLOTYPE  UC CAS 484385  San Francisquito Bay, Baja California, Mexico  T, ISOTYPE  San Francisquito Bay, Baja California, Mexico  T, ISOTYPE  San Francisquito Bay, Baja California, Mexico  G, T ISO-  in UC CAS 484386 in UC CAS 464388  TYPE  in UC CAS 484390  San Marcos Is., Gulf of California, Mexico  T, PARATYPE  in UC LAM 52684  Puerto Escondido, Gulf of California, Mexico  T  LAM 52894  SW end of Isla Partida, Gluf of California,  T  Mexico  / 280 Gelidium  purpurascens  T  CANA 3473  Beacon Hill, Victoria, B.C.  CANA 3740  Beacon Hill, Victoria, B.C.  CANA 3843  Departure Bay, Vancouver Is., B.C.  CANA 4346  Departure Bay, Vancouver Is., B.C.  CANA 4349  Departure Bay, Vancouver Is., B.C.  FHL 1205  Santa Cruz, California  FHL 2849  Mitchell Point, San Juan Is., Washington  UBC A903  Ensenada, California  UBC A1799  East Sound, Orcas Is., Washington  UBC A1952  Ucluelet, Vancouver Is., B.C.  T  UBC A4263  Garden Is., Kyuquot, B.C.  T  UBC A4264  Garden Is., Kyuquot, B.C.  T  UBC A4265  Kains Is., B.C.  UBC A4402  American Camp Beach, San Juan Is.,  T  T  T  T  Washington UBC A4403  American Camp Beach, San Juan Is.,  T  Washington UBC A5195  Minnesota Seaside Station, Vancouver Is., B.C.  UBC A6744  Sunset Bay, Oregon  UBC A6755  Cape Arago, Oregon  UBC A10310  Cortes Is., B.C.  UBC A10514  Catala, Vancouver Is., B.C.  T  T  / 281 M  UBC A10813  Esteban Point, Vancouver Is., B.C.  UBC A11271  Ogden Point Breakwater, Victoria, B.C.  UBC A11278  Rosebush Is., Vancouver Is., B.C.  T  UBC A11329  Dorcas Point, Strait of Georgia, B.C.  T  UBC A11929  McLean Is., Vancouver Is., B.C.  T  UBC A11937  McLean Is., Vancouver Is., B.C.  T  UBC A11988  Spring Is., Vancouver Is., B.C.  T  UBC A12296  McLean Is., Vancouver Is., B.C.  T  UBC A13479  Walters Cove, Vancouver Is., B.C.  T  UBC A17771  Acous Peninsula, B.C.  T  UBC A19032  Hisnit Is., B.C.  UBC A28349  Orlebar Point, Gabriola Is., B.C.  UBC A28646  Twin Beaches, Gabriola Is., B.C.  UBC A31414  Lock Bay, Gabriola Is., B.C.  T  UBC A35158  Arab Cove, Vancouver Is., B.C.  T  UBC A35175  Arab Cove, Vancouver Is., B.C.  UBC A36371  Brooks Peninsula, Vancouver Is., B.C.  T  UBC A36710  Bunsby Is., Vancouver Is., B.C.  T  UBC A36875  Union Is., Vancouver Is., B.C.  T  UBC A36876  Amos Is., Vancouver Is., B.C.  F  UBC A37603  Blight Is., Vancouver Is., B.C.  T  UBC A38471  Lippy Point, Vancouver Is., B.C.  T  UBC A38475  Arab Cove, Vancouver Is., B.C.  T  UBC A38734  Arab Cove, Vancouver Is., B.C.  T  UBC A38908  Arab Cove, Vancouver Is., B.C.  T  F  / 282 UBC A39126  Lawn Point, Vancouver Is., B.C.  F  UBC A39622  Arab Cove, Vancouver Is., B.C.  T  UBC A40192  Amphitrite Point, B.C.  T  UBC A40743  Haines Is., Barkley Sound, B.C.  T  UBC A40793  Helby Is., Barkley Sound, B.C.  T  UBC A41214  Meade Islets, Barkley Sound, B.C.  T  UBC A41384  Helby Is., Barkley Sound, B.C.  T  UBC A41415  Fleming Is., Barkley Sound, B.C.  M  UBC A41491  Grappler Inlet, Bamfield, B.C.  T  UBC A41586  Wizard Islet., Barkley Sound, B.C>  F  UBC A41606  Effingham Is., Barkley Sound, B.C.  T  UBC A41607  Effingham Is., Barkle}' Sound, B.C.  T  UBC A41696  Roquefeuil Baj', Barkley Sound, B.C.  T  UBC A41842  Fleming Is., Barkley Sound, B.C.  F,T  UBC A41843  Fleming Is., Barkley Sound, B.C.  UBC A41844  Fleming Is., Barkley Sound, B.C.  T  UBC A41946  Clarke Is., Barkley Sound, B.C.  F,T  UBC A42162  Reeks Is., Barkley Sound, B.C.  T  UBC A42436  Fleming Is., Barkley Sound, B.C.  T  UBC A42437  Fleming Is., Barkley Sound, B.C.  T  UBC A42438  Fleming Is., Barkley Sound, B.C.  T  UBC A42439  Fleming Is., Barkley Sound, B.C.  T  UBC A42733  Arab Cove, Vancouver Is., B.C.  UBC A43525  Diana Is., Barkley Sound, B.C.  T  UBC A43526  Bamfield, B.C.  T  / 283 UBC A45812  Bamfield, B.C.  T  UBC A46026  Bamfield, B.C.  F,T  UBC A46960  Sear Is., B.C.  UBC A53291  Kyuquot Channel, B.C.  T  UBC A53297  Kyuquot Bay, Kyuquot Sound, B.C.  T  UBC A53396  Saanedra Is., Nootka Sound, B.C.  UBC A53447  Henderson Point, Saanich Inlet, Vancouver Is., B.C.  UBC A53657  Friendly Cove, Nootka Sound, B.C.  UBC A54172  Hotham Inlet, Harmony Is., Jervis Inlet,  M  B.C. UBC A55706  Lasqueti Is., B.C.  UBC A57371  Helby Is., Barkley Sound, B.C.  UBC A57429  Moorsam Bluff, Jervis Inlet, B.C.  UBC A57442  east of Brittain Rock, Jervis Inlet, B.C.  UBC A57598  mouth of Glacial Creek, Jervis Inlet, B.C.  UBC A58562  Station Is., B.C.  UBC A58672  island W of Fox Is., B.C.  UBC A60374  Diana Is., Barkely Sound, B.C.  T  UBC A60485  Brady's Beach, Bamfield, B.C.  T  UBC A64402  head of Pennell Sound, Queen Charlotte Is.,  T  B.C. UBC A64426  heaqd of Pennell Sound, Queen Charlotte Is., B.C.  UBC A66660  Tasu Sound, Queen Charlotte Is., B.C.  T  / 284 UBC A66682  Horn Rocks, Tasu Sound, Queen Charlotte Is., B.C.  UBC A66778  Departure Bay, Vancouver Is., B.C.  UBC A67220  Cunningham Is., B.C.  UBC A67249  Bachelor Bay, B.C.  UBC A67446  Kirby Point, Diana Is., Barkley Sound, B.C.  F, T  UBC A69432  Cape Suspiro, Alaska  T  UBC A69433  Cape Suspiro, Alaska  UBC A69434  Kanaka Bay, San Juan Is., Washington  T  UC 93572  Moss Beach, San Mateo Co., California  F HOLOTYPE  UC 276633  Moss Beach, San Mateo Co., California  F  UC 296689  Moss Beach, San Mateo Co., California  F, T  UC 305332  Moss Beach, San Mateo Co., California  F  UC 305364  Pebble Beach, Monterey, California  F  UC 305371  Pescadero, California  F  UC 305373  Pescadero, California  M  UC 1452112  Moss Beach, California  F  private collection of J.R. Waaland: 1583  Gelidium  Kanaka Point, San Juan Is., Washington  robustum  / 285 UBC A1448  Moss Beach, California  T  UBC A7861  Shoal B, Victoria  T  UBC A37790  Asilomar Point, California  UBC A47163  Palos Verdes, California  UBC A47164  Santa Catalina Is., California  UBC A50638  Point Pinos, Monterey Peninsula, California  UBC A58591  Santa Cruz Is., California  UBC A60676  Pedros Blancas, California  UBC A62199  Carpinteria Reef, California  UBC A63553  Monterey, California  UC 294572  near Ensenada, California  T  F  T HOLOTYPE  UC 395419  San Pedro, California  F  UC 647822  White's Point, San Pedro, California  F  UC 756464  Punta Santa Rosalia, Baja California, Mexico  UC 756469  Punta Santa Rosalia, Baja California, Mexico  UC 756470  Rosario, Baja California, Mexico  UC 940173  Natividad Is., Baja California, Mexico  F  UC 1451987  Portuguese Bend, San Pedro, California  F  Point Cavalo, Marin Co., California  HOLOTYPE  Gelidium  sinicola  UC 276620  / 286 Gelidium vagum  TNS 25817  Harutachi, Hidaka, Japan  T  TNS 25823  Hideshima, Iwate-ken, Japan  T  TNS 25824  Ta-no-hama, Iwate-ken, Japan  F  TNS 25825  Ta-no-hama, Iwate-ken, Japan  F  TNS 25847  Same, Aomori-ken, Japan  T  UBC A56807  Muroran, Hokkaido, Japan  T  UBC A64965  Ladysmith, Vancouver Is., B.C.  T  Pterocladia caloglossoides  UBC A12295  McLean Is., Vancouver Is., B.C.  UBC A19667  Bunsby Is., Vancouver Is., B.C.  UBC A28937  Orlebar Point, Gabriola Is., B.C.  T  UBC A29576  Orlebar Point, Gabriola Is., B.C.  T  UBC A30401  Whiffen Spit, Sooke, Vancouver Is., B.C.  UBC A31251  Whiffen Spit, Sooke, Vancouver Is., B.C.  UBC A33545  Arab Bay, Vancouver Is., B.C.  UBC A37605  Inner Bajo Reef, Nootka Is., Vancouver Is., B.C.  UBC A39126  Lawn Point, Vancouver Is., B.C.  UBC A39127  Lawn Point, Vancouver Is., B.C.  UBC A61245  Diana Is., Barkley Sound, B.C.  UBC A64648  head of Rennell Sound, Queen Charlotte Is., B.C.  UBC A69423  Cape Suspiro, Alaska  UBC A69424  Cape Suspiro, Alaska  UBC A69425  Cape Suspiro, Alaska  UBC A69426  San Clemente Is., Alaska  UBC A69427  Sea Otter Sound, Alaska  UBC A69428  Sea Otter Sound, Alaska  UBC A69429  Sea Otter Sound, Alaska  UBC A69430  Sea Otter Sound, Alaska  UBC A69431  SeaOtter Sound, Alaska  WTU  Mitchell Bay, San Jaun Is., Washington  248018  APPENDIX 2. PROCEDURE FOR EMBEDDING MATERIAL  IN JB4  METHACRYLATE Fixation 2.5% glutaraldehyde in Sorensen's phosphate buffer (pH 7.2) (1:1) 4 h, room temperature 4-  3 10 min washes in Sorensen's buffer 4-  Dehydration 10%, 25%, 40%, 50%, 70%, 80%, 90%, 100%, 100%, 100% MeOH each step 10 min 10% - 80% in refrigerator, 90%, 100% at room temperature 4-  Infiltration 1 part catalyzed solution A : 3 parts 100% MeOH 1h 4-  1 part catalyzed solution A : 1 part 100% MeOH 4h 4-  3 parts catalyzed solution A : 1 part 100% MeOH 8 h or overnight, refrigerator 4-  100% catalyzed solution A overnight, refrigerator 288  / 289 4-  Embedding 1 part solution B (activator) : 25 parts catalyzed solution A poured into mold , tissue arranged in mold, capped with a stub, hardened in refrigerator, overnight  APPENDIX 3. P R E P A R A T I O N OF TETRASPORANGIAL SPERMATANGIAL  AND  M A T E R I A L FOR TRANSMISSION E L E C T R O N MICROSCOPY  Fixation 2.5% glutaraldehyde in Sorensen's phosphate buffer (pH 7.2) (1:1) 4 h, room temperature/spermatangial: 7 h, refrigerator  tetrasporangial:  4-  3 10 min washes in Sorensen's buffer 4-  Osmication 1% OsO„ in buffer (1:1) tetrasporangial:  18-19 h, refrigerator/spermatangial: 16 h, refrigerator 4-  3 10 min washes in buffer 4-  Dehydration 10%, 25%, 40%, 50%, 70%, 80%, 90%, 100%, 100%, 100% MeOH each step 10 min 10% - 80% in refrigerator, 90%, 100% at room temperature 4-  Infiltration 25%, 50%, 75%, 100% propylene oxide in MeOH 20 min, room temperature 4-  290  / 291 Embedding 10% Spurr's epoxy resin in propylene oxide overnight, refrigerator, on rotator 4-  25%, 40%, 55%, 70%, 80%, 90%, 100%, 100%, 100% Spurr's in propylene oxide each step \ day at room temperature on rotator, or overnight in refrigerator 4-  Polymerization in Spurr's epoxy resin under 18 psi vaccuum \ day, then 600 min, 70°C, 18 psi  

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