A Review of the Prymnesiophyta, Emphasizing the Morphology and Systematics of Hymenomonas Stein (1878) and Pleurochrysis Pringsheim (1955)

A REVIEW OF THE PRYMNESIOPHYTA, EMPHASIZING THE MORPHOLOGY AND SYSTEMATICS OF HYMENOMONAS STEIN (1878) AND PLEUROCHRYSIS PRINGSHEIM (1955)

Cory Dashiell

A Thesis Submitted to the University of North Carolina Wilmington in Partial Fulfillment of the Requirements for the Degree of Master of Science

Department of Biology and Marine Biology

University of North Carolina Wilmington

2010

Approved by

Advisory Committee

Dr. Wilson Freshwater Dr. Richard Dillaman

Dr. Alison Taylor Dr. J. Craig Bailey Chair

Accepted by

Dr. Roer Dean, Graduate School

TABLE OF CONTENTS

ABSTRACT...... iii

ACKNOWLEDGMENTS ...... iv

DEDICATION...... v

LIST OF TABLES...... vi

LIST OF FIGURES ...... vii

CHAPTER 1 ...... 1

Background Information ...... 1

CHAPTER 2 - PHYLOGENY...... 11

Introduction...... 11

Methods ...... 14

Results ...... 18

Discussion...... 23

CHAPTER 3 – CONFOCAL MICROSCOPY...... 32

Introduction...... 32

Methods ...... 33

Results ...... 35

Discussion...... 37

LITERATURE CITED ...... 39

FIGURE LEGENDS ...... 75

ABSTRACT

The systematics of several isolates belonging to the Hymenomonadaceae and

Pleurochrysidaceae (Prymnesiophyceae) were newly examined or reexamined. A new species,

Pleurochrysis dimidius, is described on the basis of light- and electron microscopic observations and comparative analyses of nuclear 18S rRNA and plastid rbcL gene sequences. Results confirm that the Hymenomonadaceae and Pleurochrysidaceae are sister taxa within the

Coccolithales. Two isolates identified as Hymenomonas spp are re-classified as Pleurochrysis elongata and P. pringsheimii comb. nov. An updated classification for these two families of coastal coccolithophorids is presented.

iii

ACKNOWLEDGMENTS

I would like to acknowledge the hard work of my advisor, Dr. J. Craig Bailey. Without him this thesis would not have been possible. I would like to thank my committee members, Dr.

Dillaman, Dr. Freshwater, and Dr. Taylor, for their time, advice, and valuable suggestions, Mark

Gay for all of his help with my microscopy, and my friends and family for their support throughout my time at UNC Wilmington.

This work was supported by NSF grant 0328316 awarded to JCB. The culture of

Pleurochrysis dimidius was a gift from the late Dr. Paul Krugrens.

DEDICATION

I would like to dedicate this thesis to my parents. They have always been there for me and encouraged me to do my best. Without their love and encouragement I wouldn’t have been able to become the person that I am today.

LIST OF TABLES

Table Page

1. GenBank accession numbers for the 18S RNA species of haptophytes included in the

study...... 56

2. Revised classification for the Hymenomonadaceae and Pleurochrysidaceae based on

information obtained in this study ...... 62

3. Staining specificity of the lectins used in this study for the three Pleurochrysis spp.

...... 63

LIST OF FIGURES

Figure Page

1. Diagrammatic representation of cell structures of coccolithophores...... 66

2. Line drawings of coccoliths in selected Pleurochrysidaceae and

Hymenomonadaceae...... 67

3. Light micrographs of isolates examined in this study ...... 68

4. TEM micrographs of Pleurochrysis dimidius sp. nov ...... 69

5. Strict consensus tree depicting relationships among haptophyte species based

upon 18S rRNA gene sequences...... 70

6. Maximum likelihood tree depicting relationships among haptophyte species based

upon rbcL gene sequences...... 71

7. Maximum likelihood tree depicting relationships among species belonging to the

Hymenomonadaceae and Pleurochrysidaceae based upon 18S rRNA gene

sequences ...... 72

8. DIC images of nonmotile diploid and haploid life history phases of Pleurochrysis

species...... 73

9. Four patterns of binding specificity seen in the species of Pleurochrysis...... 74

vii

CHAPTER 1 - BACKGROUND INFORMATION ON THE PRYMNESIOPHYTA

Prymnesiophyceae

The phylum Prymnesiophyta is divided into two classes, the Pavlovophyceae and the Prymnesiophyceae (which includes the coccolithophorids) and encompasses organisms colloquially known as haptophytes (Hibberd 1972, Hibberd 1976, Nicholls

2002). There are over 200 marine species of haptophytes but very few freshwater representatives (Jordan and Green 1994, Jordan and Chaimberland 1997). Less than 12 freshwater species are known and are placed in seven genera (Hymenomonas,

Chrysochromulina, Acanthoica, Anacanthoica, Pavlova, Diacronema, and

Exanthemachrysis) (Nicholls 2002). Hymenomonas roseola is the only freshwater coccolithophorid reported from North America (Lackey 1939, Meyer and Brook 1968,

Smith 1950, Stoermer and Sicko-Goad 1977).

Haptophytes possess chloroplasts that lack a girdle lamellum and most contain chlorophylls a and c1/c2, β-carotene, diadinoxanthin, and diatoxanthin. Important fucoxanthin derivatives (19’-hexanoxyfucoxanthin and 19’-butanoyloxyfucoxanthin) and chlorophyll c3 divide the phylum into four main subgroups (Jeffrey and Wright 1994,

Jordan et al. 1995). Haptophytes possess flagellae that may be equal or unequal in length and lack mastigonemes, except in the Pavlovophycidae where fibrous hairs and knobscales are present on the longer flagellum (Jordan and Chamberlain 1997). Eyespots are lacking in most haptophytes but are present in the Pavlovophycidae. Most species have fibrillar, unmineralized body scales, and many species also possess calcified scales known as coccoliths (Jordan and Chamberlain 1997).

As the name of the group implies, many haptophytes posses a haptonema that is emergent or nonemergent (i.e., reduced). The haptonema is a structure unique to haptophytes and Christensen (1962) used the haptonema and features of the cell covering to segregate the haptophytes from the Chrysophyceae. The haptonema comprises a protrusion supported by microtubules covered by plasmallema scales and is inserted at the anterior end of the cell between the flagella. The number of microtubules making up the haptonema varies from three to eight among species (Billard and Inouye 2004). The presence of a haptonema is plesiomorphic for haptophytes and has been adapted for several uses. Haptonema are known to be used to adhere to substrata and in some species of Chrysochromulina the haptonema has been implicated in prey capture. The haptonema is also sometimes used in collision avoidance responses and may be chemotactic (Graham and Wilcox 2000, Kawachi and Inouye 1995, Kawachi et al. 1991).

Haptonema vary in length, with some species possessing a long haptonema capable of coiling or uncoiling, whereas others possess a reduced (short) bulbous haptonema

(Billard and Inouye 2004). Although some haptonema are capable of coiling and uncoiling, the structure does not beat like a flagellum (Graham and Wilcox 2000).

Species that lack haptonema and have lost them secondarily typically possess intracellular microtubules indicating its former presence (Jordan and Chamberlain 1997).

The haptonema is positioned close to the left basal body. The microtubules of the haptonema form an arc (often described at C- or U-shaped) oriented with the concave portion positioned towards the left basal body. The left basal body corresponds to the mature (longer) flagellum and the right basal body, located farther from the haptonemal

base, corresponds with the immature (shorter) flagellum (Figure 1. Billard and Inouye

2004).

Coccolithophores

The order Coccolithales encompasses a group of haptophyte algae belonging to the Prymnesiophyceae. All haptophytes possessing calcified scales belong to the

Coccolithales, although the Prymnesiophyceae includes families that are non-calcifying

(i.e. Isochrysidaceae) or have non-calcifying stages in their life histories (i.e.

Noelaerhabdaceae, Hymenomonadaceae, and the Pleurochrysidaceae) (Young et al.

2005). In these taxa, the cell covering is composed only of nonmineralized, organic, fibrillar body scales (Billard and Inouye 2004). The coccolithophores are often considered the most ecologically important of the haptophytes because of the importance of their calcified scales in fossil studies and marine biogeochemistry (Holligan 1993,

Jordan and Chamberlain 1997).

Coccolithophores reproduce asexually by binary fission, although syngamy has been observed in a few species (Schwarz 1932, Gayral and Fresnel 1983). After cell division, coccoliths are redistributed between the daughter cells (Billard and Inouye

2004).

The Golgi apparatus in haptophytes is always positioned on the anterior side of the nucleus, and lies more-or-less between the nucleus and the proximal ends of the basal body/haptonemal complex (Graham and Wilcox 2000, Nicholls 2002). It is responsible for scale production in species such as Hymenomonas carterae (Pienaar 1969) and many

others. Coccoliths formed in Golgi derived vesicles are extruded onto the cell surface and form the coccosphere (Graham and Wilcox 2000).

Organic Body Scales

Organic body scales are non-calcifying circular or elliptical fibrillar structures that contain cellulose. These scales are found underneath the coccoliths in some species of coccolithophorids and are the only form of cell wall in families that produce a non- calcifying haploid phase (i.e. Noelaerhabdaceae, Pleurochrysidaceae,

Hymenomonadaceae) (Billard and Inouye 2004, Graham and Wilcox 2000). The organic body scales are produced in the cisternae of the Golgi apparatus and then secreted over the plasmalemma (Billard and Inouye 2004, Brown et al. 1970, Graham and Wilcox

2000, Pienaar 1976b).

Organic body scales are of three types. Type 1 scales are circular in shape and have concentric and radial fibers ornamenting both sides. They are generally rimmed.

Type 2 scales are rimless elliptical organic scales with a pattern of concentric fibrils on the distal face and radiating fibrils in four quadrants on the proximal face. Type 3 scales, while less common than type 1 or 2, are found in varying shapes (circular to elliptical) and possess a pattern of radiating fibrils that do not meet at a center point and are arranged in four segments (Billard and Inouye 2004).

Coccoliths

Coccoliths are external body scales formed from calcium carbonate often found associated with proximal organic plate scales (i.e., proximal to base plate scales) (Green

and Jordan 2002). Coccoliths of some species can be very elaborate in design and vary greatly. Morphology of coccoliths has been used in the past to differentiate among different coccolithophorid species (Green and Jordan 2002).

There are several different types of coccoliths but they are divided into two basic types that differ in the calcium carbonate crystal-units and how they are produced.

Heterococcoliths are composed of two crystal-units (V and R) that differ by their chirality, whether the coccolith morphotypes can be described as right or left handed

(Young et. al 1999). Types of heterococcoliths include cricoliths, helicoliths, and pappoliths (Billard and Inouye 2004). Heterococcoliths are produced internally and mineralize in the Golgi derived vesicles (Nicholls 2002, Young et al. 1999).

Holococcoliths are composed of one type of crystal, crystallites that are smaller than the crystal-units of heterococcoliths. They are termed calyptoliths, crystalloliths, and laminoliths (Billard and Inouye 2004). It has been suggested that holococcoliths are produced in the Golgi apparatus and are mineralized extracellularly at the plasma membrane, but evidence is lacking (Graham and Wilcox 2000, Nicholls 2002, Manton and Leedale 1969, Young et al. 1999).

Haptophyte life cycles

Many species of coccolithophores (and other prymnesiophytes) have been found to possess an alternating life cycle including diploid and haploid phases. In heteromorphic species the phases are often characterized by different cell coverings and different morphologies. The organism may alternate between motile and non-motile stages and forms may bear scales of different kinds (Billard and Inouye 2004). For

example, Coccolithus pelagicus has been found to alternate between heterococcolith and holococcolith life-cycle phases (Billard 1994, Young et al. 1999). It has been suggested that the haplo-diploid phases of the coccolithophores’ life cycle are an adaptation to a seasonally variable environment or a way to exploit two different niches in an environment (Houdan et al. 2004).

The relatively recent discovery that holococcolithophorids are often phases in the alternate life cycles of heterococcolithophorids has greatly affected the taxonomy and nomenclature of many described species (Billard 1994, Billard and Inouye 2004, Noël et al. 2004, Young et al. 2000). Organisms previously classified as separate species on the basis of coccolith morphology are now recognized as different phases of the life history of a single biological species. These discoveries have caused some authors to suggest that DNA sequences should be used in subsequent taxonomic classifications (Saéz et al.

2008).

Previous studies suggest that diploid life stages commonly possess heterococcoliths and that holococcoliths are indicative of the haploid stage. Some families of coccolithophorids, including the Noelaerhabdaceae, Hymenomonadaceae, and

Pleurochrysidaceae, possess haploid stages that are non-calcifying. These non-calcifying stages have only organic body scales for a cell covering. The cell wall, or periplast, of these organisms is comprised of multiple layers of organic scales anchored by fibrillar or columnar material (Leadbeater 1994, Billard and Inouye 2004).

Pleurochrysidaceae and Hymenomonadaceae

The families Hymenomonadaceae and Pleurochrysidaceae have been recognized as sister clades within the Coccolithales (Saéz et al. 2004). Billard and Inouye (2004) and others have classified Pleurochrysis (Pringsheim 1955) within the family

Pleurochrysidaceae (Young and Brown 1997, Edvardsen et al. 2000, Kleijne et al. 2001).

The type species for the genus is Pleurochrysis scherffelii (Pringsheim 1955).

Hymenomonas roseola, the type species of Hymenomonas, was originally described from brackish waters by Stein (1878).

Species assigned to Hymenomonas Stein (1878) and Pleurochrysis Pringsheim

(1955) are very similar. Most Pleurochrysis species possess a reduced bulbous haptonema as do Hymenomonas spp. Hymenomonas spp. still possess haptonemal scales, which are reduced body scales located near the flagellar pole, suggesting that the haptonema has been recently lost (Billard and Inouye 2004). Pleurochrysis species can be either motile or non-motile in nature as can those placed in Hymenomonas (Green and

Jordan 2002). Both genera also include species that have plastids (usually two) with bulging pyrenoids that project from the periphery of the plastid toward the cells’ interior.

Pyrenoids in most other species of coccolithophores are of the immersed type (Billard and Inouye 2004, Green and Jordan 2002, Manton and Peterfi 1969).

Hymenomonas and Pleurochrysis are mainly separated on the basis of scale morphology. Hymenomonas species possess monomorphic crown-shaped coccoliths

(tremaliths) that overlay organic scales and the species also sometimes possesses smaller elliptical scales on diploid cells. In contrast Pleurochyrsis has cricoliths on diploid cells.

The cricoliths are monomorphic and composed of two narrow shields or elements.

Pleurochrysis possesses a haploid, pseudofilamentous ‘Apistonema-like’ stage formed from the meiospores of the coccolith-bearing cells (Leadbeater 1970, von Stosch 1967).

It is believed that the families Pleurochrysidaceae and Hymenomonadaceae have secondarily abandoned production of calcified coccoliths in the haploid (holococcolith phase) (Houdan et al. 2004). Pleurochrysis possesses type 2 body scales while

Hymenomonas produces two types of scales, with the proximal layer being type 2 scales identical to Pleurochrysis, and the distal layer being composed of scales that are rimmed and appear homologous to un-calcified holococcoliths (Billard and Inouye 2004). Recent studies indicate that coccoliths produced during different phases of a species’ life cycle often differ (Billard and Inouye 2004) therefore, it may be necessary to re-examine the systematics of the genera Hymenomonas and Pleurochrysis.

Ecological significance

Haptophytes, especially coccolithophorids, are of extreme ecological and biogeochemical importance in marine environments. Haptophytes are a significant constituent of sedimentary rock and coccoliths compose a large proportion of marine calcareous deposits (Black and Barnes 1961, Bramlette 1958, Jordan and Chamberlain

1997). Coccolithophorids play a significant role in the cycling of carbon in the environment. The calcium-carbonate scales are responsible for between 20 and 40% of the total amount of vertical transport of carbon to the deep ocean each year (Brand 1994,

Broecker and Peng 1982, Graham and Wilcox 2000, Henderiks and Pagani 2007).

Coccolithophores may be in danger due to increasing ocean acidity. Increases of carbon dioxide (CO2) in the atmosphere since the Industrial Revolution have resulted in a

decrease in the pH of the ocean that can interfere with both calcifying and non-calcifying species. These species will likely be affected by the changes in the composition of seawater, especially the effects on calcium carbonate (Doney et al. 2009). The ocean is expected to become even more acidic over the next century (0.3-0.4 pH units), increasing at a rate higher than any seen in the past 650,000 years (Caldeira and Wickett 2003,

Caldeira et al. 2007, Guinotte and Fabry 2008, Lueker et al. 2000, Mehrbach et al. 1973,

Siegenthaler et al. 2005). Few studies have been performed to examine the effects that this may have on marine coccolithophores, but species such as Emiliania huxleyi and

Geophyrocaps oceanica have been found to experience decreased calcification rates under more acidic conditions while Coccolithus pelagicus and Calcadiscus leptoporus have been shown to not experience a decrease in calcification rates and the ability to adapt to changing acidity (Langer et al. 2006). It will be important to study the way increasing ocean acidification may act with other stressors such as increasing ocean temperature to fully understand the effect that this may have on these organisms.

Calcified scales also serve roles in stratigraphic and palaeoceanographic studies because they are an important microfossil group commonly used to determine relative ages for marine sediments (Jordan and Chamberlain 1997). The calcified coccoliths are well preserved in ocean sediments and have been dated as far back as the Carboniferous

(Siesser 1994) or Late Triassic periods (c. 200 mya) (Perch-Nielsen 1985, Green et al.

1990, Young et al. 1994). Calcified coccoliths are responsible for the naming of the

Cretaceous period (c. 125 mya), which was named for chalk deposits composed almost entirely of coccoliths (Graham and Wilcox 2000). In the petroleum industry, fossils of coccoliths are used as bioindicators (Young et al. 1994).

Blooms are common for a number of haptophyte species (e.g., Chrysochromulina,

Phaeocystis, Prymnesium and Emiliania) and the products of these blooms can have tremendous impacts on the environment (Jordan and Chamberlain 1997). During the final stages of a bloom, when the cells become stressed, are dying, or are lyzed, they sometimes release extracellular compounds that are harmful to other species.

Phaeocystis is responsible for producing large amounts of dissolved organic carbon

(DOC) and foam that can suffocate near shore animals whereas mucilage is responsible for fouling commercial fishing nets. Emiliania huxleyi and Phaeocystis blooms release dimethylsulphoniopropionate (DMSP) that hydrolyzes into dimethyl sulphide (DMS) and acrylic acid (Jordan and Chamberlain 1997, Keller et al. 1989, Liss et al. 1994, Malin and

Steinke 2004, Malin et al. 1993). In addition to DMS production being important in the sulphur cycle and the global climate, DMS is also responsible for increasing acid rain

(Graham and Wilcox 2000, Jordan and Chamberlain 1997, Malin and Steinke 2004).

Blooms of calcified species affect the carbon cycle and provide a sink for excess carbon by forming chalk deposits (Jordan and Chamberlain 1997, Stanley et al. 2005).

CHAPTER 2 - SYSTEMATICS

SYSTEMATICS OF THE HYMENOMONADACEAE AND

PLEUROCHRYSIDACEAE (PRYMNESIOPHYCEAE)

INTRODUCTION

The Coccolithales Schwarz 1932 (emend. Edvardsen and Eikrem et al. 2000) includes four principal families; the Calcidiscaceae Young and Brown 1997,

Coccolithaceae Poche 1913 (emended Young and Brown 1997), Hymenomonadaceae

Senn 1900, and Pleurochrysidaceae Fresnel and Billard 1991. A controversial fifth family, the Reticulosphaeraceae, including the unusual, unmineralized amoeboflagellates

Reticulosphaera socialis and R. japonensis, is recognized by some authorities but not others (Cavalier Smith et al. 1996, Grell 1989a, 1989b, 1990, Grell et al. 1990, Jordan et al. 2004)

This study focuses on the systematics of the Hymenomonadaceae and

Pleurochrysidaceae, which are resolved as sister taxa based upon comparative morphological and DNA sequence studies (Saéz et al. 2008). The Hymenomonadaceae includes two genera, Hymenomonas Stein 1878 and Ochrosphaera Schussnig 1930. The type species of Hymenomonas (H. roseola) was described by Stein (1878) without reference to scales, and the taxon presently includes six other species found in brackish or freshwater ecosystems (Gayral and Fresnel 1979, Green and Jordan 2002, Saéz et al.

2004, 2008). Ochrosphaera is monotypic and typified by O. neapolitana (Schussnig

1930). The Pleurochrysidaceae (excluding Reticulosphaera) includes a single genus,

Pleurochrysis Pringsheim 1955 that is typified by P. scherfelii Pringsheim. Eight other species are now included in Pleurochrysis and some (e.g., P. carterae) are among the best studied of all coccolithophorids. Morphological, ultrastructural, and DNA sequence comparisons suggest that Jomonlithus Inouye and Chihara (1983) likely belongs in the

Hymenomonadaceae or Pleurochrysidaceae but to which family is presently unclear

(Inouye and Chihara 1983, Jordan et al. 2004, Saéz et al. 2004).

The Hymenomonadaceae and Pleurochrysidaceae include coastal coccolithophorids characterized by a free-swimming (motile) diploid life history phase possessing two subequal flagella and a short, bulbous, haptonema (Outka and Williams

1971, Saéz et al. 2004). Diploid cells are covered by layers of unmineralized organic body scales with different faces; microfibrils on the proximal surface are arranged radially whereas those on the distal surface of the scale are arranged in a concentric or whorled pattern (Manton and Leedale 1969, Pienaar 1969, Outka and Williams 1971).

Atop these organic body scales are base plate scales with calcified rims (coccoliths) classified, depending upon their form, as cricoliths or trematoliths (Manton and Peterfi

1969). Species assigned to the Hymenomonadaceae or Pleurochrysidaceae are primarily distinguished on the basis of coccolith morphology (Fig. 2). Diploid

Hymenomonadaceae bear crown shaped or tube shaped tremaliths composed of several identical trapezoidal sub-elements (Jordan and Kleine 1994). In contrast, diploid

Pleurochrysidaceae are characterized by more or less elliptical cricoliths that are composed of several anvil shaped components of two types, distinguished by the orientation of their crystal components; which are radial (R) or vertical (V) (Fresnel and

Billard 1991, Outka and William 1971, Saéz et al. 2004, 2005, Young et al. 1992). For

comparison, coccoliths representing Pleurochrysis, Hymenomonas, and Ochrosphaera species are depicted in Figure 2.

The haploid life history phase in species assigned to both families takes the form of benthic pseuodfilaments (sometimes falsely branched), individual cells, or of groups of several cells (Pringsheim 1955, von Stosch 1955, 1958, 1967, Parke 1961, Leadbeater

1971, Hibberd 1980). Coccoliths are unknown for haploid cells and, instead, haploid cells are covered by organic body scales only (Saéz et al. 2008). Clusters of haploid cells and the pseudofilamentous habit of haploid cells arise as a result of interactions among the overlapping and firmly compressed organic body scales of adjacent cells

(Leadbeater 1970).

This study of the Hymenomonadaceae, Pleurochrysidaceae, and related taxa was initiated by the receipt of a freshwater alga originally collected and isolated into unialgal culture by the late Dr. Paul Kugrens. Multiple kinds of data obtained for the alga

(reported herein) indicate that the isolate is the benthic, presumptively haploid, non- mineralizing life history phase of a previously unknown Pleurochrysis species. A new species is erected to accommodate the isolate, which is referred to throughout this paper as Pleurochrysis dimidius. To properly describe this alga it was necessary to (re)examine strains ascribed to Hymenomonas available for study in algal culture collections. Light microscopic and SEM observations as well as DNA sequence data were collected for three isolates identified as belonging to Hymenomonas Stein. What follows in this contribution is: (1) The first description of a new pleurochrysidalean coccolithophorid species using the phylogenetic species concept and based entirely upon data obtained from the haploid phase of the organism’s life history, (2) A phylogenetic and taxonomic

reassment of several isolates assigned to Hymenomonas, and (3) a re-evaluation of

Ochrosphaera and Jomonilithus. A revised classification for the Hymenomonadaceae and Pleurochrysidaceae is presented.

METHODS

Cultures

Pleurochrysis dimidius was collected from a freshwater stream in Colorado, USA; further collection information is unavailable. Unialgal cultures were (re)established at

UNCW using serial dilution and micropipette-picking techniques and maintained in

DYIV medium (Andersen et al. 1997) at 15oC under a 14:10 light:dark photoregime and under ambient light conditions in the laboratory (22-24oC). Cultures of Hymenomonas elongata (CCAP 961/3) and H. pringsheimii (CCAP 944/2) were obtained from the

Culture Collection of Algae and Protozoa (www.ccap.ac.uk). Hymenomonas roseola

(CCAC ASW 02009) was obtained from the Culture Collection of Algae at the

University of Cologne, Germany (www.ccac.uni-koeln.de). The three isolates were grown at room temperature (22-24oC) under ambient light conditions. Hymenomonas elongata and H. pringsheimii were maintained in f/2 medium and H. roseola was grown in DYIV medium (Andersen et al. 1997, Guillard and Ryther 1962).

Salinity tolerance was examined for Pleurochrysis dimidius. Ten different salinities (5, 8, 10, 15, 20, 22, 25, 30, 33, 36) were examined, formulated by mixing freshwater DYIV and saltwater f/2 media. The media was directly inoculated with P. dimidius cells, i.e., no attempt was made to acclimate cells from lower to higher salinities. Cultures were maintained at room temperature (22-24oC) under ambient light

conditions and growth was monitored over 60d. At the end of the experiment cells were examined using a light microscope.

Microscopy

Light microscopic observations of cultures were made using a Zeiss Axio Imager

(Z1) microscope equipped with Axio Vision 4.5 software and an Axio Cam HRm digital camera. Cell sizes were digitally measured and are reported as averages obtained from

100 arbitrarily selected cells.

Electron microscopy

For TEM nonmotile cells of Pleurochrysis dimidius were fixed on ice in 0.5M sodium cacodylate buffer (pH 6.8) with 4% paraformaldehyde and 0.25M sucrose for 90 min. The primary fixation was followed by buffer rinses (on ice) with decreasing amounts of sucrose (0.25M, 0.125M, 0.0625M, 0M), 15min each. Cells were subsequently stained overnight in 2% osmium tetroxide in 0.2M sodium cacodylate buffer at 4oC, then rinsed twice for 15 min in 0.2M sodium cacodylate buffer, followed by a 15 min rinse in dH2O. Cells were dehydrated in an ascending series of ETOH (50,

70, 95, 100, 100%) for 15min each. Cells were immersed in 100% propylene oxide for

15 min and then infiltrated using a 2:1 mixture of propylene oxide and Spurr’s epoxy resin (Spurr 1969) for 1hr, then infiltrated with a 1:1 mixture of propylene oxide and

Spurr’s resin for 1 hr on a rotator, followed by a further infiltration with 100% Spurr’s

resin overnight. Cells were then placed in fresh 100% Spurr’s epoxy resin, concentrated by centrifugation (5,000 rpm for 3 min), and cured at 60oC for 8 hrs.

Sections were cut using a Reichert-Jung Ultracut E Ultramicrotome with glass or diamond knives, collected on copper grids, and stained with 2% uranyl acetate (in 50%

ETOH) and Reynold’s lead citrate (Reynolds 1963). Sections were examined with a

Philips CM 12 transmission electron microscope operated at 80kV. Micrographs were taken using a plate camera and Kodak EM4489 3 ¼” x 4” film. The film was developed, negatives were digitized using a Microtek Scanmaker i900, and images were processed and labeled in Adobe Photoshop 7.0.

DNA extraction, PCR amplification, and gene sequencing

DNA was extracted from each isolate as described in Bailey et al. (1998). The

18S rRNA gene was amplified in 100 µL reactions containing 73.5 µL dH2O, 20 µL 5X

PCR buffer, 2 µL dNTP, 0.5 µL GoTaq (Promega, Madison, WI), and 1 µL each of primers P1 and P7 (10 µM) (Medlin et al. 1988). The PCR profile included an initial denaturing step at 94oC for 4 min, followed by 35 cycles of 94°C for 30s, 50°C for 30 s,

72°C for 90 s, and a final extension at 72°C for 7 min. A portion of the rbcL gene was amplified and sequenced from each isolate as described above using primer combination

PrL1/PrL4 (Fujiwara et al. 1994). Amplified products were purified using the GeneClean

II Kit (Qbiogene, Carlsbad, CA). PCR products were sequenced on both strands using the BigDye Terminator cycle sequencing kit (v. 3.1, Applied Biosystems [ABI], Foster

City, CA). Sequences were analyzed on an ABI 3130xl automated DNA sequencer (ABI,

Foster City, CA). Sequences were edited and assembled using Sequencher v. 4.9 (Gene

Codes Corporation, Ann Arbor, MI, USA).

Phylogenetic analyses

The global prymnesiophyte 18S rRNA alignment included sequences for 72 isolates that were first aligned using ClustalX (Thompson et al. 1997) then edited by eye in MacClade (Maddison and Maddison 2002). A region of one hundred fifty three characters that could not be confidently aligned was excluded from subsequent analyses.

The prymnesiophyte 18S rRNA trees were rooted using sequences from the diatom

Thalassiosira pseudonana (AF374481) and the cryptomonad Cryptomonas ovata

(AB240952). A second 18S rRNA data set limited to 29 taxa classified in the

Pleurochrysidaceae and Hymenomonadaceae was also analyzed. One hundred sixteen characters that could not be confidently aligned were excluded from analyses and the tree was rooted using the 18S rRNA sequence for Pavlova gyrans (U40922). The 18S rRNA sequence for Reticulosphaera japonensis reported by Cavalier-Smith et al. (1996) is controversial (see discussion) and for this reason trees were constructed including and excluding the sequence. The rbcL alignment included 39 prymnesiophytes and was rooted on the rbcL sequence for Cryptomonas ovata (AM051210).

The ‘global’ prymnesiophyte 18S rRNA alignment was analyzed using MrBayes

(Huelsenbeck and Ronquist 2001) and run for a total of 1,010,000 generations using a general time reversible (GTR) model and parameters obtained from MrModeltest

(Nylander 2004), which had a proportion of invariable sites = 0.4746 and a gamma distribution (α = 0.4668). Trees were sampled every 100 generations, producing a total

of 10,100 trees. Ninety percent of these trees were used to form the majority rule consensus tree, excluding any trees sampled prior to stabilization of the likelihood values.

Parsimony and maximum likelihood (ML) analyses of the

Pleurochrysidaceae/Hymenomonadaceae 18S rRNA sequence matrix and the prymnesiophyte rbcL data were conducted using PAUP v.4.0b10 (Swofford 2002).

Heuristic parsimony searches were conducted with 100 random sequence addition replicates using the tree bisection-reconnection (TBR) algorithm, character states unordered, and gaps treated as missing data. Parsimony bootstrap values were generated based upon 10,000 replicates using the fast-stepwise addition option. For ML a

GTR+I+G substitution model was determined as the best-fit model of substitution using

MrModeltest (v. 2.2; Nylander 2004) and a heuristic search using TBR branch swapping on a single starting tree derived from random sequence additions was ran.

RESULTS

Morphology

Diploid cells of Hymenomonas pringsheimii (CCAP 944/2) took the form of motile or nonmotile cells with mean diameters of 11.7 µm, range=5.7-20.9 µm, n=100)

(Fig. 3A and B). Motile cells possess two subequal length flagella and a short haptonema

(Fig. 3A). Nonmotile cells possessed two plastids with bulging, in-ward facing pyrenoids and often, but not always, a vacuole could be discerned (Fig. 3B). Diploid cells of

Hymenomonas elongata (CCAP 961/3) closely resembled those of H. pringsheimii and are approximately the same size (mean dia. 10.2 µm, range=6.2-19.3 µm, n=100).

Hymenomonas elongata motile cells are characterized by subequal length flagella, a short

haptonema, and two plastids with prominent pyrenoids (Fig. 3C - E). Except for the presence of flagella and a haptonema, the morphology of nonmotile cells was identical to that for motile cells in both isolates (Fig. 3A-E). Coccoliths were not observed for either species.

Morphological data for Hymenomonas roseola (ASW 02009) are unavailable because the culture could not be maintained in the laboratory.

Solitary Pleurochrysis dimidius cells are typically spherical or globose in shape, ranging in diameter from 11.5 – 18.72µm (mean=14.62µm) (Fig 3H). Cells may form sarcinoid clusters or psuedofilaments in culture in which case adpressed cells are hemispheroidal (Fig 3F, G, I-L). Individual cells typically have one or two parietal plastids with conspicuous bulging pyrenoids (Fig 3F-L). One or more light refracting granuals are usually, but not always, present. The composition of these granules is unknown. Clusters of two or more cells are surrounded by a common layer of material as evident in Fig 3 (F, G, J and L).

Pleruochrysis dimidius cells were found to be viable and reproducing asexually at all salinities investigated. No changes in morphology were seen and cells behaved the same as in freshwater.

Pleurochrysis dimidius cells did not fix well for electron microscopy. Two connected highly vacuous cells each with two plastids and at least one ‘granular body’ are depicted in Fig 4A. Cells contain a typical nucleus with scattered chromatin, multiple mitochondrial profiles, and at least one Golgi body (Fig 4B). Nucleoli were not observed. Detail of a ‘granular body’ is depicted in Fig 4B and its’ contents arguably appear crystalline in nature. Plastids contain multiple thylakoids and bulging, inward-

facing pyrenoids traversed by several thylakoids (Fig 4C). Cells are not bounded by a true cell wall. Instead, protoplasts are surrounded by numerous layers of organic scales

(Figs 4A, D-F). Multiple cells are held together by an outer layer of scales that envelopes all cells (cf. Fig 3 F-L and Fig 4 A,F). An inner layer of scales separates adjacent cells and is apparently composed of scales contributed by each cell (Figs 4A and 4F). Our best micrographs of the scale layer are shown in Figures 3D-F. Our interpretation of the image in Figure 4E is of a single sloughing and slightly twisted scale. Vesicles of different sizes and shapes were common at the periphery of cells just beneath the scale layer (Figs 4G,H). Scales were not observed in these vesicles and their function is unknown.

Phylogenetic analyses

GenBank accession numbers for the 18S rRNA and rbcL sequences determined in this study are presented in Table 1. The global 18S rRNA alignment included 72 taxa,

1690 characters, and cladistic analyses of these data yielded 35,627 equally parsimonious trees (L= 1517, CI=0.52, RI=0.08) swapped to completion. A strict consensus tree depicting relationships inferred among prymnesiophytes based on these 18S rRNA data is shown in Figure 5. The overall topology of the tree is consistent with results obtained in previous studies of prymnesiophyte phylogeny also employing 18S rRNA sequences

(Edvardsen et al. 2000, Saéz et al. 2004, 2008). Species belonging to the

Hymenomonadaceae and Pleurochrysidaceae are resolved as descendants of a common ancestor. The parsimony bootstrap value for this node is not robust (62%) but the

Bayesian posterior probability is much higher (100%) (Fig. 5). The monophyly of

Pleurochrysidaceae sensu stricto, including Pleurochrysis dimidius sp. nov., is strongly supported but this clade includes sequences for Hymenomonas elongata (CCAP 961/3) and H. pringsheimii (CCAP 944/2); the simplest explanation for this observation is that the latter two isolates are misidentified and belong in Pleurochrysis (Fig. 5). Isolates representing the Hymenomonadaceae did not form a distinct clade in the 18S rRNA tree and relationships among the hymenomonadaceaen isolates examined are largely unresolved (Fig. 5).

The global rbcL alignment included 40 prymnesiophyte taxa and cladistic analyses of these sequences yielded the maximum likelihood phylogram shown in Figure

6. It is difficult to directly compare the rbcL tree (Fig. 6) with the 18S rRNA tree (Fig. 5) because the trees include different species and the rbcL tree contains data for far fewer taxa. The Hymenomonadaceae is represented in the rbcL tree by a single taxon,

Hymenomonas roseola, which is resolved as sister to the Pleurochrysidaceae – including

Pleurochrysis dimidius – with reasonable bootstrap support (84/74). The rbcL tree also implies that Hymenomonas elongata (CCAP 961/3) and H. pringsheimii (CCAP 944/2) are misidentified and belong in Pleurochrysis (cf. Figs 5 and 6). Furthermore, this tree implies very different evolutionary histories for sequences labeled Coccolithus pelagicus

(AF196307) and Coccolithus pelagicus 2 (EU082829). These sequences are placed in very different positions in the rbcL tree and it is unlikely that they are derived from the same species (Fig. 6).

The Hymenomonadaceae+Pleurochrysidaceae 18S rRNA alignment included 29 taxa and 1757 nucleotide characters. ML analysis of this alignment placed species in one of two sister clades whose monophyly is robustly supported by parsimony bootstrap data

and weakly supported by ML bootstrap data (Fig 7). The first clade includes species or isolates assigned to Hymenomonas, Jomonlithus, Ochrosphaera, and Reticulosphaera.

The Hymenomonas species placed within this clade did not form a monophyletic subgroup. Instead these results strongly support the hypothesis that H. roseola

(ASW02009) and Jomonlithus littoralis are sister species. In fact, no differences were found between the 18S rRNA sequences for H. roseola (1680 bp) and Jomonlithus littoralis (1721 bp) over the 1680 nt that could be compared. Evidence for a close relationship between Hymenomonas coronata and H. globosa is lacking and neither of these species is allied with H. roseola. Ochrosphaera verrucosa, three Ochrosphaera sp. isolates, and Reticulosphaera socialis were resolved as a distinct clade with moderate

(71%, 81%) bootstrap support but relationships among the five isolates could not be determined.

The second clade includes species assigned to Hymenomonas or Pleurochrysis and is robustly supported (95%, 100%) by the data. Taxa assigned to Hymenomonas and

Pleurochrysis belonging to this clade are paraphyletic with respect to one another.

Hymenomonas elongata (CCAP 961/3) and Pleurochrysis elongata (HAP79) were resolved as closely related species, with identical 18S sequences. H. elongata (CCAP

961/3) and Pleurochrysis carterae (CCMP874) were resolved as sister species but this relationship is only moderately supported by the data (71%). Twenty substitutions were observed between the 18S rRNA sequences for these two species over 1744 bp.

Pleurochrysis dimidius, H. pringsheimii (CCAP 944/2) and Pleurohrysis sp.

(MBIC10549) are resolved as closely related taxa within this clade, with identical 18S sequences over the 1609 nucleotides that could be compared.

Pleurochrysis dimidius sp. nov. Dashiell et Bailey

Cells solitary and spherical or clustered forming sarcinoid packets or pseudofilamentous. Cells 11-19µm in diameter, typically with one or two plastids, conspicuous inward-facing pyrenoids, and one or more refractile bodies. Individual cells covered by layers of small organic scales; clusters bounded by scale layers enveloping multiple cells.

Holotype: CCMP 0000, Fig. 3K

Habitat: Freshwater stream.

Distribution: Colorado, USA

Entomology for the specific epithet: dimidius =half, referring to the fact that only “half” of the life history (the haploid portion) of the organism is now known.

DISCUSSION

Phylogenetic analyses of 18S rRNA and rbcL sequences are consistent with the hypothesis that the Hymenomonadaceae and Pleurochrysidaceae are sister taxa within the

Coccolithales. However, it is evident that the systematics of these taxa needs to be revised. Herein (1) a new species, Pleurochrysis dimidius, is established,

(2) the taxonomy of “Hymenomonas” isolates that have been misidentified are readdressed, (3) the circumscription of the Hymenomonadaceae is updated, and (4) a new

classification for the Hymenomonadaceae and Pleurochrysidaceae is presented (cf.

Jordan et al. 2004).

Pleurochrysis dimidius sp. nov.

A new species, Pleurochrysis dimidius, belonging in the Pleurochrysidaceae is erected in this study. Comparative morphological and ultrastructural data as well as our nuclear 18S rRNA and rbcL gene trees indicate that this alga is a natural member of the

Prymnesiophyceae belonging to the genus Pleurochrysis (Fresnel and Billard 1991,

Gayral and Fresnel 1983, Leadbeater 1971). Previous morphological and life history studies of pleurochrysidalean algae imply that the organism is haploid, alternating with a diploid, heteromorphic life history phase bearing organic body scales and Pleurochrysis- type cricoliths (von Stosch 1967, Brown 1969, Leadbeater 1970, 1971, Inouye and

Chihara 1979, Young et al. 2003). Morphological and ultrastructural features observed

(e.g., the alga’s habit, subcellular arrangement of organelles, a periplast composed of layers of organic body scales, the nature and orientation of pyrenoids, thylakoids, etc.) are entirely consistent with comparable accounts of the haploid life history phases of

Pleurochrysis spp (Leadbeater 1970, 1971, Mills 1975, Pienaar 1976). Despite numerous attempts (involving different media, calcium carbonate concentrations, vitamins, and substrates) the alga could not be induced to complete its’ life history in culture (data not presented). Swimming cells of any sort were not observed (Gayral and Fresnel 1983, von

Stosch 1967). In culture the alga reproduces only by means of simple cell division and although organic body scales are produced observations of coccoliths, flagella, and a haptonema are lacking. It is tempting to speculate that motile gametes were not observed

because our P. dimidius isolate is homothallic. On the other hand, all life history stages of Pleurochrysis placolithoides were observed by Fresnel and Billard (1991) based upon observations of clonal cultures derived from a single, diploid coccolith-bearing cell.

Analyses of rbcL sequence data imply that the alga is not conspecific with any other previously examined species. Based on these data we establish P. dimidius as a new species within the family using the phylogenetic species concept (Wheeler and Meier

2000). This is, to our knowledge, the first time that any coccolithophorid species has been established without reference to coccoliths, which historically have been the tool most often employed discriminate among coccolithophorid species. Because

Hymenomonas and Pleurochrysis species often abandon coccolith production in culture, as in this study, comparative studies of P. dimidius coccoliths could not be made

(Braarud 1955, Manton and Peterfi 1969). In a comparable precedent, Leadbeater (1971) found the coccoliths of Pleurochrysis scherffelii indistinguishable from those of P. carterae but retained the species separately based on differences between the morphologies of their benthic haploid stages. Interestingly, Stein’s (1878) original description of Hymenomonas and his diagnoses for the type species H. roseola include no information on coccoliths, of which he was almost certainly unaware (Scherffel 1927,

Schiller 1930).

Pleurochrysis dimidius was isolated from a freshwater stream and only a handful of other prymnesiophytes have been collected from freshwaters (Gayral and Fresnel

1979, Green and Jordan 2002, Saéz et al. 2004, 2008). Nevertheless, viable P. dimidius cells were observed at all salinities examined in this study (0-36) after 60 d. This demonstrates that the alga is euryhaline and agrees with the fact that the greater majority

of coccolithophorids are marine. These experiments imply that growth and reproduction in P. dimidius may not be limited to freshwater.

Pleurochrysis elongata

Three isolates bearing the specific epithet “elongata” and assigned either to

Hymenomonas or Pleurochrysis were examined. Briefly, morphological data for these isolates – if any – is incomplete and equally consistent with placing them in either genus.

To our knowledge none of these isolates now produce coccoliths in culture and for generic assignment this problem is insurmountable on the basis of morphology alone.

In contrast, DNA sequence data indicate with actionable certainty that all three isolates are natural members of Pleurochrysis and strains CCAP 961/3, CCMP874, and

ALGO HAP97, are herein, if not previously, assigned to that taxon (Saéz et al. 2004).

Following this line of reasoning, the question arises: “Are the Pleurochrysis elongata strains examined here conspecific or not?” Answering this question requires us to interweave (1) gene sequence comparisons, with (2) the culture histories of the strains,

(3) phylogenetic inferences, and (4) rules for proper recognition of taxa according to the

ICBN.

Historical analyses of records indicate that strain CCAP 961/3 is the ‘authentic’ culture for Pleurochrysis elongata (Droop) Jordan. The culture was established by

Droop in 1953 providing the basis for his description, in 1955, of Syracosphaera

‘Hymenomonas’ elongata. Droop (1955) did not examine the coccoliths of this organism in detail, which were subsequently reinvestigated by Braaud (1960) who found that the alga produced cricoliths. On the basis of this observation and observation of

Syracosphaera carterae, Braaud (1960) established Cricosphaera to include cricolith- bearing coccolithophorids lacking a known benthic stage like that previously observed in

Pleurochrysis sherffelii by Pringsheim (1955). Later, Christensen (1978) observed the alternate, benthic life form in his studies of Cricosphaera carterae. Thus, according to the principle of priority, Cricosphaera Braaud (1960) is now treated as a later synonym of Pleurochrysis Pringsheim (1955). Cricosphaera carterae and C. elongata were transferred to Pleurochrysis by Jordan (1993) but, apparently, the proper name of the alga was not updated by the CCAP.

Strain HAP79 was assigned to P. elongata by Saéz et al. (2004) apparently only on the basis of comparisons of 18S rRNA sequence information, whereas strain

CCMP874 is identified as Pleurochrysis elongata in Edvardsen et al. (2000). The 18S rRNA sequence determined for the authentic strain of Pleurochrysis elongata (CCAP

961/3) was identical to the sequence reported by Saéz et al. (2004) for strain HAP79.

This implies that the two strains may be conspecific. However, the 18S rRNA gene evolves too slowly among closely related prymnesiophytes for species level work. In contrast, Pleurochrysis elongata (CCAP 961/3) and Pleurochrysis elongata (CCMP874) are resolved as sister species in this study but the relationship between the two isolates is not necessarily a close one. Twenty substitutions were observed between the 18S rRNA sequences for Pleurochrysis elongata (CCAP 961/3) and Pleurochrysis elongata

(CCMP874). These differences arguably provide grounds for considering the two isolates as separate species and indicate that strain CCMP874 is not ‘elongata’ and must be recognized as a separate species. However, identifying strain CCMP874 as P. carterae

is unwise because (1) the strain was not examined in this study, and (2) it is certainly not conspecific with von Stosch’s (1967) strain of P. carterae (see Fig. 7).

Pleurochrysis sp. (CCAP 944/2)

Strain CCAP 944/2 identified as Hymenomonas pringsheimii was examined in this study. Available morphological as well as 18S rRNA and rbcL sequence data indicate that this alga is member of Pleurochrysis, not Hymenomonas. Hymenomonas pringsheimii Parke et Green was established with no reference to diploid or haploid cell type or coccoliths and described therin as being similar to Pleurochrysis scherffelii (Parke and Green 1976). The strain no longer produces coccoliths and additional studies are needed to determine if the alga warrants recognition as a new Pleurochrysis sp.

Systematics of Hymenomonadace (s.s.)

The Hymenomonadaceae includes two genera: Hymenomonas and Ochrosphaera

(Jordan et al. 2004, Fresnel and Probert 2005). However, our DNA sequence analyses demonstrate that the Hymenomonadaceae also includes Jomonlithus littoralis and

Reticulosphaera spp.

Cells of Hymenomonas spp and J. littoralis do not differ appreciably at the ultrastructural level (Inouye and Chihara 1983). Hymenomonas and Jomonlithus are, instead, recognized as separate based on two fundamental differences. First, an emergent haptonema has not been observed for Jomonlithus littoralis whereas a reduced (short) but visible haptonema is present in Hymenomonas spp (Inouye and Chihara 1983). Second,

Jomonlithus littoralis possesses two, if not three, distinct types of scales that are unlike

those described for Hymenomonas spp (Manton and Peterfi 1969, Mills 1975, Pienaar

1976, Inouye and Chihara 1980). The 18S rRNA sequences for Hymenomonas roseola

CCAC ASW 02009 and Jomonlithus littoralis are identical to one another.

Hymenomonas roseola CCAC ASW 02009 did not produce coccoliths in culture and for this reason there is no way to determine if tremaliths are of the Hymenomonas or

Jomonlithus type. It is conceivable that the two strains are conspecific but, in the absence of coccolith data for any true representative of Hymenomonas, it was decided to maintain

Hymenomonas and Jomonlithus as distinct genera.

The taxonomic status and phylogenetic affinities of Reticulosphaera Grell are controversial. Reticulosphaera includes two species, R. socialis (the type species) and R. japonensis (Grell 1989a, Grell 1990). Both are dimorphic with an ameboid benthic stage and heliozoan-like pelagic stage. Reticulosphaera socialis and R. japonensis exhibit similar morphologies and life forms, but slightly differ in cell size and shape. Also, the benthic stage of R. socialis is more apt to be heterotrophic whereas the benthic stage of R. japonensis is typically photosynthetic (Grell 1990). The benthic stage of R. socialis is a mixotrophic reticulopodial amoeba that consumes diatoms. When diatom prey are scarce

R. socialis will transform to autotrophic “yellow cells” with reduced reticulopodia and prominent plastids and if light limited, cells will transform into the free floating pelagic stage (Grell 1989b, Grell et al. 1990).

Pelagic R. socialis cells possess thin filapodia and two flagella that are difficult to discern at the light microscope level. Flagellar length is unknown. TEM studies confirm that the flagella are heteromorphic; one flagellum is characterized by a basal swelling positioned near the eyespot of an adjacent plastid, whereas a basal swelling is lacking on

the second flagellum. This photoreceptor apparatus is not unlike those previously reported for many photosynthetic stramenopiles (ex. chrysophytes, synurophytes, and xanthophytes). Plastids are traversed by 6-8 lamellae each composed of three adpressed thylakoids. Pelagic cells possess tubular hairs lacking bases and terminal filaments that are transported by a Golgi-derived vesicle and extruded onto the cell surface. Vesicles containing what could possibly be interpreted as scales have been observed; however, the composition, function, and destination, of purported scales are unknown (Grell et al.

1990).

Characters including the rhizopodial form, yellowish-colored plastids (suggesting a lack of fucoxanthin?), lack of a rhizoplast, heteromorphic flagella possessing a transitional helix, and tubular hairs present on the cell suggested to Grell et al. (1990) that

Reticulosphaera might belong in the Xanthophyceae. Cavalier-Smith (1993) then placed

Reticulosphaera in the Flavoretea, which he included also among the stramenopiles.

Analysis of an 18S rRNA sequence for R. socialis subsequently reduced the class to sublcass Flavoretophycidaceae within the Prymnesiophceae (Cavalier-Smith et al.

1996). In later analyses, R. socialis was resolved as closely related to Pleurochrysis and the family Reticulosphaeraceae Cavalier-Smith (1996) was placed in the order

Coccolithales (Edvarsen et al. 2000).

In our 18S rRNA analyses we used the molecular sequence data provided on

GenBank from the Cavalier-Smith et al. (1993) study for Reticulosphaera socialis. The

18S rRNA analyses for the Hymenomonadaceae+Pleurochrysidaceae were performed both with and without the R. socialis sequence and yielded identical results. Our

observations strongly support the hypothesis that Reticulosphaera is a close relative of

Ochrosphaera and belongs in the Hymenomonadaceae.

Based on the data obtained in this study a revised classification for the families

Hymenomonadaceae and Pleurochrysidaceae is presented in Table 2.

CHAPTER 3 – CONFOCAL MICROSCOPY

COMPARISON OF LECTIN-BINDING GLYCOCONJUGATES IN ALTERNATE

LIFE HISTORY PHASES OF HYMENOMONAS (PRYMNESIOPHYCEAE).

INTRODUCTION

Scale morphologies are useful for identifying haptophyte species that have similar sizes, shapes, and flagellar or haptonemal features (Rhodes and Burke 1996). One tool that can aid in the identification of cell wall components is the use of fluorescent dyes and lectins that preferentially stain glycoconjugates found within various components of cells and abound on cell surfaces (Allen et al. 1988; Ramoino 1997; Roberts et al. 2006).

Fluorescent plant derived lectins have been used in past studies to detect the presence, localization, and configurations of glycoconjugates in algal cells (Roberts et al. 2006).

Twenty lectins directly labeled with the fluorochrome fluorescein isothiocyanate

(FITC) and one fluorescent dye, Calcofluor White, were used to examine nonmotile diploid cells of Pleurochrysis pringsheimii Parke and Green, P. elongata (Droop) Parke and Green, and Pleruochrysis dimidius Dashiell and Bailey.

The flourecent dye Calcofluor White binds to cellulose, as well as other beta- linked glucans, and chitin (Galbraith 1981; Hughs and McCully 1975). Calcofluor White has been shown to bind to the unmineralized organic scales of Chrysochromulina chiton, suggesting that this dye could be used to examine scales of similar composition (Rhodes and Burke 1996).

These three species of Pleurochrysis possess organic, fibrillar scales. Although coccolithophores often produce calcified cells, some life stages produce only organic scales as in the haploid stage of Pleurochrysis (Billard and Inouye 2004). Calcifying species of coccolithophores have also been known to stop calcifying in culture (Holligan et al. 1993).

Pleurochrysis forms a psuedofilamentous chain of cells, while the closely related taxa of P. pringshemii and P. elongata form spherical and occasionally flagellate unicells in culture. The unique psuedofilamentous nature of the unknown species invites studies to attempt to unravel its biology.

METHODS

Experimental organisms

Pleurochrysis elongata (CCAP 961/3) and Pleurochrysis pringsheimii (CCAP

944/2) were obtained from the Culture Collection of Algae and Protozoa (ccap.ac.uk).

Pleurochrysis dimidius Dashiell and Bailey was collected from a freshwater stream in

Colorado, USA. The three isolates were grown at room temperature (22-24oC) under ambient light conditions. Cultures of P. elongata and P. pringsheimii were maintained in f/2 medium (Guillard and Ryther 1962); Pleurochrysis dimidius was maintained in DYIV medium (Andersen et al. 1997). Cells in log phase were collected by centrifugation for microscopic analyses.

Autofluorescence

Autofluorescence was analyzed in order to be able to distinguish the autofluorescence from lectin specific fluorescence using a lambda scan performed with an Olympus Fluoview 1000 Laser Scanning Microscope. User defined excitation wavelengths were set from 400 to 700nm with 10nm increments. Each laser was used individually to excite the specimen and detect cellular autofluorescence. Lambda scans were performed at 405nm (Blue diode laser) and 488nm. Plastids were found to autofluoresce between 650-700nm. No other cellular structures were found to autofluoresce.

DIC confocal microscopy

Light micrographs (single plane images 1µm thick) were obtained using differential interference contrast (DIC) optics on an Olympus Fluoview 1000 Laser

Scanning Confocal Microscope. Cells were viewed on a glass cover slip on an inverted microscope to avoid altering the shape of the cells.

FITC labeled lectins

Staining of the live cells with each of the twenty FITC labeled lectins (Vector

Labs) was performed by adding 20µl of 1mg/L of the different lectins to a 980µl cell suspension to achieve a final concentration of 20µg/mL. After 15min, cells were rinsed twice in DYIV culture medium (Pleurochrysis dimidius) or f/2 medium (P. elongata and

P. pringsheimii) for 5mins before imaging with the confocal microscope.

A multi channel argon laser with an excitation wavelength of 488nm was used to image cells and a 405/488/543 dichromatic mirror was used with a bandpass filter of 500-

550nm on the first detector to detect FITC emissions (emission peak at 520nm). Longer wavelengths of 650-700 were also analyzed using an additional filter to detect plastid fluorescence and a mirror was used for the second detector.

Calcofluor white

Cells of Pleurochrysis dimidius, P. elongata, and P. pringheimii were stained for

15 min with 0.33% Calcofluor White by adding 500µl of a 1% Calcofluor White solution to a 1ml cell suspension. Cells were rinsed twice in culture medium for 5 min before viewing. A blue diode laser with an excitation wavelength of 405nm was used to image the cells. A 405/488/543 dichromatic mirror was used with a bandpass filter of 425-

475nm on the first detector to detect emission from the Calcofluor White peak at 455nm.

Longer wavelengths were also analyzed with an additional filter to detect plastid fluorescence as above.

Processing Images

Microscope images were processed using Fluoview Viewer v. 1.7b and Adobe

Photoshop v. 7. Images were merged using the Fluoview Viewer to create stacks and saved as Tiff files. Tiff files were either 640x640 pixels for the lambda scan or

1024x1024 pixels for single x-y images.

RESULTS

For the lambda scan, both the 405 and 488 lasers showed excitation at ~650nm and returning to a baseline fluorescence at ~700nm. Peak fluorescence was observed at

~670nm and is attributed to endogenous chloroplast fluorescence.

Four distinct patterns of fluorescence were observed, with lectins showing specificity to droplets (Fig. 9A and B), plastids (Fig 9C and D) cell wall (Fig 9E and F), or extracellular structures (Fig 9G and H) or a combination of the structures.

Only two lectins, Doclichos biflorus agglutinin (DBA) and peanut agglutinin

(PNA), showed no binding in one the species, Pleurochrysis pringsheimmi (Table 3).

Binding specificity for the droplets was found with all 23 lectins in at least one species. The only exceptions were that Calcaflour white showed no binding affinity for droplets in Pleurochrysis dimidius and Conconavalin A showed no binding affinity for droplets in P. pringsheimii.

With the exception of Calcaflour white, the other 22 lectins showed binding specificity for the plastids in at least one of the Pleurochrysis dimidius. PSA, Con A, and

VVA did not bind to plastids in P. pringsheimmi. PSA, Con A, and LCA did not bind to plastids in P. elongata. PHA–E and ECL did not bind to plastids in Pleurochrysis dimidius. Cross specificity between droplets and plastids was seen in all three species with 8 lectins: WGA, SBA, UEA I, GSL I, PHA-L, DSL, GSL II, and STL.

Calcafluor white and four lectins showed specificity for the cell wall. Calcaflour white caused fluorescence in the cell walls of all three species. PSA and Con A caused fluorescence in only the cell walls of P. pringsheimmi and P. elongata. RCA 120 and

LCA caused fluorescence in only the cell walls of P. elongata.

Six lectins, PSA, Con A, LCA, PHA-E, ECL, Jacalin and VVA caused fluorescence in the scales of the undescribed species of Pleurochrysis.

DISCUSSION

Only chloroplast fluorescence was observed with the lambda scans. Each individual laser wavelength was able to cause fluorescence of the specimen.

Preliminary results indicate that morphological differences are present between the unknown species of Pleurochrysis and P. pringsheimii and P. elongata.

This observation may be indicative of a life history change between the unknown species and the known species that has resulted in a differing composition of the cell wall.

The different binding specificity seen in the species of Pleurochrysis may suggest that the different droplets contain different storage compounds, which is further supported by the refractive nature of some of the smaller droplets. Some haptophytes store their photosynthetic products, normally a beta-1,3-linked glucan, outside of the chloroplast and the binding specificity seen may be for a glucose storage product

(Alderkamp et al., 2007).

The chloroplasts of coccolithophores produce starch from photosynthesis, which is a granular substance composed of d-gluocose monomers, often with alpha-1,4 or beta-

1,3 glycosidic bonds (Kim and Archibald, 2008). The staining specificity seen within the chloroplasts may be indicative of sugar storage compound in the chloroplast or may even show the localization of the sugar product of photosynthesis that has not yet been extruded into a extraplastidial lipid droplet.

Although the species studied do not appear to have calcified scales, Pleurochrysis species generally contain a life history phase that contains calcified scales (coccoliths)

(Billard and Inouye, 2004). Studies of Pleurochyrsis haptonemofera demonstrate that lectins Con A, RCA I, and LCA strongly bind this species’ calcified scales (Hirokawa et al., 2005). Con A, RCA I, LCA also bound to the cell walls of Pleurochrysis elongata, possibly implying that although P. elongata was not presently producing calcified scales, it may still have similar glycoconjugates present on its plasmallema. These data suggest that there may be differences between the glycoconjugates experessed during the different life cycle phases of Pleurochrysis but further investigation is needed to make any definitive statements.

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Table 1. GenBank accession numbers for the 18S rRNA and/or rbcL sequences of isolates (with strain numbers, if available) included in the prymnesiophyte trees inferred in this investigation.

Species 18S rbcL

Braarudosphaera bigelowii isolate TP05-6-b AB250785

Calcidiscus leptoporus AB043690

Calcidiscus leptoporus isolate AS31 AJ544116

Calcidiscus quadriperforatus isolate ASM35 AJ544115

Calptrosphaera sp. MBIC10517 AB183608

Calyptrosphaera sphaeroidea AB043628

Calyptrosphaera sphaeroidea 2 D45842

Chryoschromulina parva AB043694

Chrysochromulina acantha strain T20 AJ246278

Chrysochromulina alifera AB043695

Chrysochromulina brevifilum strain MBIC10518 AB058358

Chrysochromulina campanulifera strain J10 AJ246273

Chrysochromulina hirta AB043632

Chrysochromulina hirta 2 D45846

Chrysochromulina hirta strain 1Y AJ246272

Chrysochromulina kappa strain EN3 AJ246271

Chrysochromulina parva strain FU44-40 EU024987

Chrysochromulina polylepis isolate B11 AJ004866

Chrysochromulina scutellum strain G7 AJ246274

Chrysochromulina sp. TKB8936 AB043697

Chrysochromulina spinifera AB043700

Chrysochromulina throndsenii strain K11 AJ246277

Chrysochromulina throndsenii strain L12 AJ246279

Chrysoculter rhomboideus AB158370 AB158371

Coccoid haptophyte CCMP625 U40924

Coccolithus braarudii isolate IBV73 AJ544117

Coccolithus pelagicus AF196307

Coccolithus pelagicus 2 EU082829

Coccolithus pelagicus strain PLY 182g AJ246261

Corcontochrysis noctivaga DQ207406

Cruciplacolithus neohelis AB043689

Cruciplacolithus neohelis strain CCMP 298 AJ246262

Cryptomonas ovata strain CCAC 0064 AM051210

Cryptomonas ovata strain NIES 274 AB240952

Diacronema vlkianum AF106056

Diacronema vlkianum strain HAP67 AJ515246

Emiliania huxleyi M87327

Emiliania huxleyi AB043631

Emiliania huxleyi 2 D45845

Exanthemachrysis gayraliae AB043701

Exanthemachrysis gayraliae strain AC15 DQ531625

Gephyrocapsa oceanica AB043630

Gephyrocapsa oceanica 2 D45844

Gephyrocapsa oceanica strain PLY G01 AJ246276

Helicosphaera carteri AB043692

Helladosphaera sp. MBIC10514 AB183607

Hyalolithus neolepis AB183265 AB183266

Hymenomonas coronata ALGO HAP58 bis AM490982

Hymenomonas elongata CCAP 961/3 XXXXXXX XXXXXX

Hymenomonas globosa strain ALGO HAP30 AM490981

Hymenomonas pringsheimii CCAP 944/2 XXXXXXX XXXXXX

Hymenomonas roseola CCAC ASW 02009 XXXXXXX XXXXXX

Imantonia rotunda AB043696

Imantonia rotunda strain UIO 101 AJ246267

Imantonia sp. MBIC10497 AB183605

Isochrysis galbana AJ246266

Isochrysis galbana strain UIO 102 AB043693

Isochrysis sp. SAG 927-2 AY119783

Isochrysis sp. santou DQ071574

Isochrysis sp. zhangjangensis DQ075203

Jomonlithus littoralis ALGO Je5 AM490979

Ochrosphaera sp. MBIC10788 AB183638

Ochrosphaera sp. strain MBIC10476 AB183604

Ochrosphaera sp. strain MBIC10548 AB183615

Ochrosphaera verrucosa strain ALGO HAP82 AM490980

Pavlova gyans AY119784

Pavlova gyrans U40922

Pavlova lutheri AF106053 AY119785

Pavlova pingui AB293551

Pavlova pseudogranifera strain HAP33 AJ515249

Pavlova salina AB043633

Pavlova salina L34669

Pavlova salina 2 AF106059

Pavlova salina 2 D45847

Pavlova sp. strain CCMP 1416 AJ243369

Pavlova sp. strain CCMP 1416 U40925

Pavlova virescens strain HAP16 AJ515248

Pavlova viridis DQ075201

Phaeocystis antarctica Karsten SK23 X77481

Phaeocystis cordata AF163147

Phaeocystis globosa Scherffel X77476

Phaeocystis jahnii AF163148

Phaeocystis pouchetii AB280613

Phaeocystis pouchetii (Hariot) Lagerheim X77475

Platychrysis sp. MBIC10528 AB183611

Platychrysis sp. TKB8934 AB043699

Pleurochrysis carterae D11140

Pleurochrysis carterae isolate HAP1 AJ544120

Pleurochrysis carterae strain von Stotsch AJ246263

Pleurochrysis dentata isolate HAP6 AJ544121

Pleurochrysis dimidius XXXXXXX XXXXXX

Pleurochrysis elongata strain CCMP 874 AJ246264

Pleurochrysis elongata strain HAP79 AM936924

Pleurochrysis gayraliae strain ALGO HAP10 AM490972

Pleurochrysis haptonemofera AB043688

Pleurochrysis placolithoides strain ALGO HAP59bis AM490977

Pleurochrysis pseudoroscoffensis strain ALGO HAP48 AM490973

Pleurochrysis roscoffensis strain ALGO HAP32 AM490974

Pleurochrysis scherfelii strain ALGO HAP11 AM490978

Pleurochrysis sp. ALGO Langue du chat AM490975

Pleurochrysis sp. CCMP 300 AJ246275

Pleurochrysis sp. CCMP 875 AJ246265

Pleurochrysis sp. LKM-20072 AM490976

Pleurochrysis sp. NMBjih026 EF208116

Pleurochrysis sp. strain MBIC10443 AB183596

Pleurochrysis sp. strain MBIC10549 AB183616

Prymnesium calathiferum U40923

Prymnesium nemamethecum AJ246268

Prymnesium parvum AB043698

Prymnesium parvum strain K081 AJ246269

Prymnesium patelliferum L34670

Reticulosphaera socialis X90992

Thalassiosira pseudonana AF374481

Umbilicosphaera foliosa isolate ESP6M1 AJ544119

Umbilicosphaera sibogae AB043691

Umbilicosphaera sibogae isolate ETH4728 AJ544118

Umbilicosphaera sibogae var. foliosa AB043629

Umbilicosphaera sibogae var. foliosa 2 D45843

Table 2. Revised classification for the Hymenomonadaceae and Pleurochrysidaceae based on information obtained in this study.

Taxon Family Species Synonyms, Basionyms Authority Hymenomonadaceae Hymenmonoas coronata Mills 1975 Hymenomonas globosa Magne 1954 Hymenomonas lacuna Pienaar 1976 Hymenomonas pringsheimii Parke et Green 1976 Hymenomonas roscoffensis Chadefaud et Feldmann 1949 Hymenomonas roseola H. coccolithophora Massart et Conrad 1926 Stein 1878 H. danubiensis Kamptner 1928 H. flava Stokes 1888 H. scherffelii Conrad 1928 Ochrosphaera neapolitana Schussnig 1930 Reticulosphaera japonensis Grell 1990 Reticulosphaera socialis Grell 1989 Jomonlithus littoralis Inouye et Chihara 1983 Pleurochrysidaceae Pleurochrysis carterae Braarud et Fagerland 1946 Pleurochrysis dentata Johansen et Doucette 1988 Pleurochrysis elongata Syracosphaera (Hymenomonas) elongata Droop 1955 (Droop) Jordan 1993 Cricosphaera elongata (Droop) Braarud 1960 Hymenomonas elongata Parke et Greene 1976 Pleurochrysis gayraliae Beuffe 1978 Pleurochrysis haptonemofera Inouye et Chihara 1979 Pleurochrysis placolithoides Fresnel et Billard 1991 Pleurochrysis Gayral et Fresnel 1983 pseudoroscoffensis Pleurochrysis roscoffensis Dangeard 1934 Pleurochrysis scherffelii Pringsheim 1955

Table 3. Staining specificity of the lectins used in this study for three Pleurochrysis spp.,

P. dimidius (Pd), P. pringsheimii (Pp), and P. elongata (Pe). Pleurochrysis cells studied are purportedly haploid whereas those for P. elongata and P. pringsheimii are diploid.

Positive staining is indicated by an “X”, absence of detectable staining is denoted by “-“.

Note that the presence of organic body scales could not be confirmed for P. elongata or

P. pringsheimii diploid cells at the level of the light microscope. If present, they were not detected by the stains used in these experiments. (Results denoted by “?”).

Binding Cell Lectin/Dye Source Affinity Droplets Plastids Wall Scales Pd - - X - Calcafluor Cellulose, β- Pp X - X ? White linked glucans Pe X - X ?

Pd X X - X Pisum sativum Glucose, Pp X - X ? PSA agglutinin Mannose Pe X - X ? Pd X X - X Glucose, Pp - - X ? Con A Concanavalin A Mannose Pe X - X ? Pd X X - X Lens culinaris Glucose, Pp X X - ? LCA agglutinin Mannose Pe X - X ?

Pd X X - - Wheat germ N-Acetyl Pp X X - ? WGA agglutinin Glucosamine Pe X X - ? Pd X X - - Datura N-Acetyl Pp X X - ? DSL stramonium lectin Glucosamine Pe X X - ? Pd X X - X Lycopersicon esculentum N-Acetyl Pp X X - ? LEL (Tomato) lectin Glucosamine Pe X X - ? Pd X X - - Solanum tuberosum N-Acetyl Pp X X - ? STL (Potato) lectin Glucosamine Pe X X - ?

Pd X X - - Ulex europaeus Pp X X - ? UEA I agglutinin Fucose Pe X X - ?

Pd X - - X Phaseolus vulgaris Complex Pp X X - ? PHA - E Erythroagglutinin Structures Pe X X - ? Pd X X - - Phaseolus vulgaris Complex Pp X X - ? PHA - L Leucoagglutinin Structures Pe X X - ?

Pd X X - - Pp - - - ? PNA Peanut Agglutinin Galactose Pe X X - ?

Griffonia Pd X X - - (Bandeiraea) simplicifolia Pp X X - ? GSL I Lectin I Galactose Pe X X - ? Pd X - - X Erythrina Pp X X - ? ECL cristagalli lectin Galactose Pe X X - ? Pd X X - X Artocarpus intergrifolia Pp X X - ? Jacalin (Jackfruit) seeds Galactose Pe X X - ? Pd X - - - Galactose, N- Ricinus communis Actyl Pp X X - ? RCA 120 agglutinin I Galactosamine Pe X X X ?

Pd X X - - Succinylated Succinylated Wheat Germ N-Acetyl Pp X X - ? WGA Agglutinin Galactosamine Pe X X - ? Griffonia Pd X X - - (Bandeiraea) simplicifolia N-Acetyl Pp X X - ? GSL II Lectin II Galactosamine Pe X X - ? Pd X X - X N-Acetyl Pp X - - ? VVA Vicia vilosa lectin Galactosamine Pe X X - ? Pd X X - - Dolichos biflorus N-Acetyl Pp - - - ? DBA agglutinin Galactosamine Pe X X - ? SBA Soybean N-Acetyl Pd X X - -

Pd X X - - Pp X X - ? SBA agglutinin Galactosamine Pe X X - ?

From: Billard and Inouye 2004

Fig. 1 Dashiell and Bailey

Fig 2. Dashiell and Bailey

Fig. 3 Dashiell and Bailey

Fig. 4 Dashiell and Bailey

Fig. 5 Dashiell and Bailey

Fig. 6 Dashiell and Bailey

Fig. 7 Dashiell and Bailey

Fig. 8 Dashiell and Bailey

Fig. 9 Dashiell and Bailey

FIGURE LEGENDS

Figure 1. From: Billard and Inouye (2004). Diagrammatic representation of cell structures of coccolithophores. Morphological features seen in various coccolithophores are combined in a single figure. Various types of coccoliths are drawn as silhouettes. Two types of coccolith- forming vesicles found in Pleurochrysis (top) and Emiliania (bottom) are illustrated. Pyrenoid

(P1) is typical in the coccolithophores and pyrenoid (P2) is seen in Emiliania and Gephyrocapsa.

Heteroccoliths are blotted black and holococcoliths are blotted by a lattice pattern.

Abbreviations CL: columnar deposit, Cv: coccolith forming vesicle, D: peculiar dilation of Golgi body, F: flagellum, H: haptonema, M: mitochondrial profiles, N: nucleus, P1: pyrenoid traversed by thylakoids, P2: pyrenoid traversed by tubular structures, PER: peripheral endoplasmic reticulum, Re: reticular body, SC: unmineralized organic scales.

Figure 2. Line drawings of coccoliths in selected Pleurochrysidaceae (A,B) and

Hymenomonadaceae (C-F). (A, B) Pleurochrysis carterae. (A) Anvil shaped V and R subelements composing cricoliths and (B) top-down view of entire coccolith demonstrating the interlocking arrangement of V and R subelements (redrawn from Okazaki et al. 1998). (C)

Hymenomonas coronata tremalith (after Inouye and Chihara 1980). (D) Hymenomonas lacuna tremalith (redrawn from Pienaar 1976). (E,F) Two tremalith morphologies reported for

Ochrosphaera verrucosa (redrawn from Inouye and Chihara 1980).

Figure 3. Light micrographs of isolates examined in this study (A, B) Unmineralized diploid cells of Hymenomonas pringsheimii (CCAP 944/2). (A) Motile cell bearing two flagella and a

short haptonema (white arrowhead, see inset). (B) Nonmotile cell with two peripheral plastids

(asterisks). (C-E) Unmineralized diploid cells of Hymenomonas elongata (CCAP 961/3). (C, D)

Motile cells with two flagella (black arrowheads), short haptonema (white arrowhead), and inward-facing, buldging pyrenoids (white asterisks). (E) Nonmotile cell of Hymenomonas elongata with two parietal plastids (asterisks). (F-L) Haploid unmineralized, nonmotile cells of

Pleurochrysis dimidius sp. nov. (F-I) Single, paired and ‘sarcinoid’ cell clusters with single plastid (black asterisk), pyrenoid (white asterisk), and refractile granules or droplets (white arrow). Note that clustered cells are enveloped by a common layer of mucilage or scales (black arrow) (J-L) Pleurochrysis dimidius sp. nov.; pseudofilamentous and sarcinoid habits.

Scale bars = 5 µm.

Figure 4. TEM micrographs of Pleurochrysis dimidius sp. nov. Abbreviations G: Golgi body,

GB: granular body, IL: inner layer of scales, M: mitochrondria, N: nucleus, OL: outer layer of scales, P: plastid, Py: pyrenoids, SL: organic scale layer, T: thylakoid, V: vesicles, (A) Two cells surrounded by an outer layer of scales enveloping both cells and an inner layer of scales that separates the cells. (B) Close up on a cell of Pleurochrysis dimidius sp. nov. featuring a prominent nucleus osmophyllic granular body, mitochondria, plastid, and Golgi body. (C) Close up on a chloroplast showing thylakoid running singly and a bulging inward facing pyrenoid with thylakoids traversing it. (D) Outer organic scale layer. (E) Outer layer of organic scales. A scale can be seen peeling off the outer layer (arrow). (F) Organic scale layer of two neighboring cells interlocking and causing the pseudofilamentous behavior of the cells. (G) Golgi derived vesicles depositing organic scales onto the outer scale layer. (H) Golgi derived vesicles depositing organic scales onto the outer scale layer.

Figure 5. Strict consensus tree depicting relationships among haptophyte species based upon 18S rRNA gene sequences. Numbers above branch are bootstrap values (≥70%), numbers below are

Baysian posterior probability values. Hymenomonas and Pleurochrysis strains examined in this study are shown in bold. Classification follows Edvardsen et al. (2000) and Saéz et al. (2004).

Figure 6. Maximum likelihood tree depicting relationships among haptophyte species based upon rbcL gene sequences. Numbers above breach are ML bootstrap values (≥50%; 100% denoted by asterisk), numbers below branch are parsimony bootstrap values. Hymenomonas and

Pleurochrysis strains examined in this study are shown in bold.

Figure 7. Maximum likelihood tree depicting relationships among species belonging to the

Hymenmonadaceae and Pleurochrysidaceae based upon 18S rRNA gene sequences. Numbers above breach are ML bootstrap values (≥50%; 100% denoted by asterisk), numbers below branch are parsimony bootstrap values. Hymenomonas and Pleurochrysis strains examined in this study are shown in bold. Outgroup not pictured.

Figure 8. DIC images of nonmotile diploid (A,B) and haploid (C) life history phases of

Prymnesium species. (A) Pleurochrysis pringsheimii (CCAP 944/2) nonmotile diploid cell(s).

(B) P. elongata (CCAP 961/3) nonmotile diploid cells. (C) Haploid life history stage of

Pleurochrysis dimidius. Scale = 5µm

Figure 9. Four patterns of binding specificity seen in Pleurochrysis. A) Light micrograph of

Pleurochrysis dimidius. Asterisks designate droplets. Scale bars = 10µm. B) Flourescence seen in the droplets of the same cell. Arrowheads designate droplets. C) Light micrograph of cells of the Pleurochrysis dimidius. Asterisks designate plastids. D) Micrograph of the same cell showing fluorescence within the plastids. E) Light micrograph of cells of the undescribed sp. with arrowhead pointing to a cleavage furrow in the cell wall. F) Micrograph of the same cell showing binding specificity to the cell wall, including the cleavage furrow of the dividing cell.

G) Light micrograph of cells of the undescribed species. Cells of this species cling together due to extracellular fibrillar scales (arrowhead). H) Micrograph of the same cell showing binding specificity to the fibrillar scales. Note: Only cells of the undescribed species showed this pattern of binding specificity that was often characterized by a patchy distribution of fluorescence.