ABSTRACT PARROW, MATTHEW WAYNE. Feeding, Reproduction, and Sexuality in Pfiesteria Spp. and Cryptoperidiniopsoid Estuarine Hete

ABSTRACT

PARROW, MATTHEW WAYNE. Feeding, Reproduction, and Sexuality in Pfiesteria spp. and Cryptoperidiniopsoid Estuarine Heterotrophic Dinoflagellates. (Under the direction of JoAnn M. Burkholder.)

Dinoflagellates are an ancient, adaptively diverse group of protists that function as important primary producers and/or consumers in many aquatic habitats, and sometimes as toxin producers. Most species are known only from drawings, photographs, or written descriptions of field material. Comparatively few have been cultured for experimentation because of unknown nutritional requirements. Thus, studies on the behavior and reproduction of most dinoflagellates are still at the basic level of observational science.

Pfiesteria spp. and cryptoperidiniopsoids (the latter taxon not yet formally described) are closely related thecate, omnivorous dinoflagellates in temperate-subtropical estuaries. They impact food webs through their rapacious phagotrophy, and some

Pfiesteria spp. strains are also ichthyotoxic. These species are relatively easy to culture when given prey, which has allowed knowledge of their range and potential impacts to progress quickly. This research investigated feeding, reproduction, and sexuality of these ecologically significant dinoflagellates in six studies:

(i) Grazing and growth rates were compared when Pfiesteria spp. and cryptoperidiniopsoids were fed cryptophyte microalgal prey. The strains of P. piscicida and the cryptoperidiniopsoids used showed higher population growth rates on algal prey than P. shumwayae, suggesting species-specific prey preferences and growth rates. Taxa with higher growth rates exerted lower grazing pressure, indicating reliance on mixotrophy.

Growth also varied among these taxa depending on nutritional history, indicating that sudden changes in prey type can influence growth rates.

(ii) The cellular DNA content and population DNA distributions of several

Pfiesteria spp. isolates fed fish or cryptophyte algal prey were measured by flow cytometry.

The two Pfiesteria spp. differed significantly in relative cellular DNA content, in both mean

DNA content and population DNA distributions. Intraspecific differences in DNA content also occurred among clones from different geographical regions, and there was a negative correlation between DNA content and time in culture. The data supported the premise that

Pfiesteria spp. flagellated cells were interphasic, and differences in nuclear DNA content were related to growth rates.

(iii) Asexual reproduction in Pfiesteria spp., and sexuality in P. piscicida were examined, and a technique for population synchronization was developed and tested. Cell division was described in nonmotile cysts, supported by photography and flow cytometric

DNA measurements. Sexuality – including nuclear cyclosis, a precursor to meiosis – was documented in P. piscicida clonal cultures, and lipid content was tracked. Taxonomic placement of Pfiesteria spp. within the order Peridiniales was suggested.

(iv) Aspects of the life history and nutritional ecology of cryptoperidiniopsoids were investigated. The DNA content and population DNA profiles of synchronized populations from different cryptoperidiniopsoid isolates were measured and related to their life history.

Bacteria improved feeding and growth, and there was apparent nutritional benefit from dissolved organic nutrients.

(v) The method developed in (iii-iv) was modified to obtain dense, synchronous populations of Pfiesteria spp. and cryptoperidiniopsoids. Initial synchrony typically was >

90% 1C DNA. DNA analyses on synchronized populations were improved, with minimal cytoplasmic DNA contamination. The technique will be useful in ecological, biochemical, and molecular investigations of these and similar heterotrophic dinoflagellates.

(vi) Major asexual and sexual reproductive pathways of P. shumwayae fed fish were examined. Full evidence of sexuality (fusing gametes, planozygotes, and nuclear cyclosis) was only found when clonal isolates were mixed; thus, self-sterility factors apparently influenced sexuality.

These investigations provided new information on nutrition, reproduction, and sexuality in these species. Detailed accounts of dinoflagellate reproduction are rare, especially for heterotrophic species, and observations on nuclear cyclosis and meiosis are extremely rare. This research also contributed among the first DNA measurements in support of ploidy assumptions for phagotrophic dinoflagellates. The findings are fundamentally relevant to the behavior, ecology, and phylogeny of these ecologically significant dinoflagellates.

FEEDING, REPRODUCTION, AND SEXUALITY IN PFIESTERIA SPP. AND CRYPTOPERIDINIOPSOID ESTUARINE HETEROTROPHIC DINOFLAGELLATES

Matthew Wayne Parrow

A dissertation submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the Degree of Doctor of Philosophy

BOTANY

Raleigh

2003

APPROVED BY:

______Chair of Advisory Committee Committee Member JoAnn M. Burkholder Nina Strömgren Allen

______Committee Member Committee Member Daniel L. Kamykowski

BIOGRAPHY

Matthew W. Parrow was born in Canandaigua, New York, and his appreciation for biology began as a youth exploring in the woods. One of his first elementary school science projects was a microscopic survey of protists found in a rural pond. Matthew graduated from McDowell County High School in North Carolina in 1988, and received a

Bachelor of Science in Biology with minors in Chemistry and Geology from the University of North Carolina at Chapel Hill in 1992. From 1992-1996 he worked in research at the

Department of Geology at UNC-CH, investigating the stable isotope stratigraphies and evolutionary morphometrics of Paleocene-Eocene marine calcareous micro- and nannoplankton. In 1997 Matthew undertook coursework at North Carolina State

University, and in 1998 he began a graduate research assistantship in the Department of

Botany at NCSU. For his doctorate he studied the reproductive and nutritional biology of three closely related species of estuarine heterotrophic dinoflagellates, under the direction of Dr. JoAnn M. Burkholder. This dissertation presents those studies.

ACKNOWLEDGEMEMENTS

I wish to express my sincere gratitude to my major advisor, Dr. JoAnn Burkholder, for her invaluable support and guidance. JoAnn has been a truly remarkable mentor, colleague, and friend, and contributed to my doctoral education and research in ways too numerous to list. I thank you for everything, JoAnn. I am also genuinely grateful to each member of my graduate committee, Drs. Nina Strömgren Allen, Parke Rublee, and Daniel

Kamykowski. I could not have asked for a better group of advisors for education and guidance, and I thank you each for your enthusiasm, experience, and counsel. I am also grateful to Drs. Malte Elbrächter, Jane Lewis, and Diane Stoecker for their interest, advice, and encouragement.

I would like to thank Dr. Howard Glasgow for his support and efforts, which made many of my opportunities possible, and Nora Deamer, Dr. Robert Reed, Elle Allen, Greg

Melia, and Jessica Alexander for their generous help, advice, and friendship. I would like to extend my appreciation to everyone in the NCSU Botany Department and Center for

Applied Aquatic Ecology who provided personal, educational, and research support. It has been my distinct pleasure and privilege to work with such talented, friendly people, and I thank you all.

iii

TABLE OF CONTENTS

LIST OF TABLES .………………………………………………………………….….…vii

LIST OF FIGURES .………………………………………………………………….…..viii

1. COMPARATIVE RESPONSE TO ALGAL PREY BY PFIESTERIA PISCICIDA, PFIESTERIA SHUMWAYAE SP. NOV., AND AN ESTUARINE ‘LOOKALIKE’ SPECIES ...…………………………………………….2 1.1 Abstract ...………………………………………………………………………3 1.2 Introduction .…..……..…………………………………………………………5 1.3 Materials and Methods .………………..……………………………………….8 1.4 Results .………..………………………………………………………………12 1.5 Discussion …………………………..………………………………………...19 1.6 References ………………..…………………………………………………...21

2. FLOW CYTOMETRIC DETERMINATION OF ZOOSPORE DNA CONTENT AND POPULATION DNA DISTRIBUTION IN CULTURED PFIESTERIA SPP. (PYRRHOPHYTA) ..…….…………………………………….…24 2.1 Abstract .……………..………………………………………………………..25 2.2 Introduction .…..………………………………..……………………………..26 2.3 Materials and Methods .……………………………………………………….29 2.4 Results .………………………………………………………………………..33 2.5 Discussion ………………………………………………………………….…47 2.6 References ………………………………………………………………….…52

3. VEGETATIVE AND SEXUAL REPRODUCTION IN PFIESTERIA SPP. (DINOPHYCEA) CULTURED WITH ALGAL PREY, AND INFERENCES FOR THEIR CLASSIFICATION ...………………………………...... 57 3.1 Abstract .……………………………………………………………….………58

3.2 Introduction .…..………………………………………………………………60 3.3 Materials and Methods .……………………………………………………….63 3.4 Results .…………………………………………………………..……………71 3.5 Discussion ………………………………………………………………….…98 3.6 References …………………………………………………………………...115

4. ESTUARINE HETEROTROPHIC CRYPTOPERIDINIOPSOIDS (DINOPHYCEA): LIFE CYCLE AND CULTURE STUDIES ...…………………...127 4.1 Abstract .………………………………………………………………...……128 4.2 Introduction .…..………………………………………………………...……130 4.3 Materials and Methods .………………………………………………………133 4.4 Results .……………………………………………………………….………147 4.5 Discussion …………………………………………………………………....169 4.6 References …………………………………………………………….……...183

5. A CELL CYCLE SYNCHRONIZATION AND PURIFICATION TECHNIQUE FOR HETEROTROPHIC PFIESTERIA AND CRYPTOPERIDINIOPSOID DINOFLAGELLATES ANALYZED BY FLOW CYTOMETRY ...……………………………………….………………..197 5.1 Abstract .…………………………………………………………………..…198 5.2 Introduction .…..………………………………………………..……………199 5.3 Materials and Methods .……………………………………………..……….201 5.4 Results and Discussion.…………………………………………………....…203 5.5 References …………………………………………………………………...208

6. REPRODUCTION AND SEXUALITY IN PFIESTERIA SHUMWAYAE (DINOPHYCEAE) ...………………………………………………..210 6.1 Abstract .…………………………………………………………………...... 211 6.2 Introduction .…..…………………………………………………………..…212 6.3 Materials and Methods .……………………………………………………...215

6.4 Results .…………………………………………………………….……..…222 6.5 Discussion ………………………………………………………….…….....243 6.6 References ……………………………………………………………...…...257

LIST OF TABLES

Table 1.1 Mean specific growth rates, division times, maximum yields, and ingestion rates for Pfiesteria spp. and a cryptoperidiniopsoid……………………………………………..... 13

Table 2.1 Pfiesteria piscicida and P. shumwayae clonal isolate origins, culture ages, mean 1C DNA contents, and CVs ………………………..……….. 41

Table 4.1 The cryptoperidiniopsoid isolate origins, isolation dates, PCR identification, and mean 1C DNA contents ……………………….. 136

Table 4.2 Media formulations tested for axenic growth / survival of selected cryptoperidiniopsoid isolates …………………………………... 146

Table 6.1 The Pfiesteria shumwayae isolates examined, geographical origins, and time in culture……………………………….……………...……….. 216

vii

LIST OF FIGURES

Figure 1.1 Population growth of Pfiesteria piscicida, P. shumwayae, and a cryptoperidiniopsoid …………………………………...……….. 16

Figure 1.2 Rhodomonas sp. cell production in 6-day batch trials with Pfiesteria piscicida……………………………………….……………. 17

Figure 1.3 Population dynamics between Pfiesteria piscicida and Rhodomonas sp. as prey in 6-day trials ……………………………….. 18

Figure 2.1 DIC light micrograph and scanning electron micrograph of Pfiesteria shumwayae reproductive cysts ………………...……….. 34

Figure 2.2 SYTOX Green-stained 1C DNA flagellated cells of Pfiesteria piscicida, viewed with epifluorescence microscopy……...... 38

Figure 2.3 Peak DNA-fluorescence distributions for Pfiesteria piscicida and P. shumwayae flagellated cells……………………..……………... 43

Figure 2.4 Mean 1C DNA for Pfiesteria piscicida clonal isolates, versus isolate age (time since field isolation) ………………………………… 45

Figure 2.5 2C DNA stained and sorted Pfiesteria piscicida cells, viewed with epifluorescence microscopy ………………….……….… 46

Figure 3.1 Light micrographs of Pfiesteria spp. fed algal prey ………………….. 74

Figure 3.2 Light micrographs of Pfiesteria spp. fed algal prey ………………….. 80

Figure 3.3 Light and scanning electron micrographs of Pfiesteria piscicida fed algal prey …………………………………….. 85

Figure 3.4 DNA-fluorescence distributions and forward scatter signals for Pfiesteria spp. in semi-log plots…………………...………. 90

Figure 3.5 Bivariate histograms of side-light scatter versus lipid fluorescence for Pfiesteria piscicida flagellated cells……….………… 95

Figure 3.6 DNA-fluorescence distributions from Pfiesteria piscicida cells 24 h and 72 h after synchronization ……...……………………... 96

viii

Figure 3.7 Deconvoluted DNA distributions of Pfiesteria shumwayae flagellated cells 24 h and 72 h after synchronization ………..……….. 97

Figure 4.1 Depiction of a diagnostic swimming pattern commonly observed in the cryptoperidiniopsoid isolates ………..………………. 135

Figure 4.2 Light micrographs of cryptoperidiniopsoids ………………………….. 150

Figure 4.3 Light micrographs of cryptoperidiniopsoid sexual cells ……………… 154

Figure 4.4 DNA profiles of cryptoperidiniopsoid flagellated cell populations, following a synchronized excystment …………………... 161

Figure 4.5 1C DNA content versus years in culture for the cryptoperidiniopsoid isolates examined ………………………………. 162

Figure 4.6 1C DNA comparison between the cryptoperidiniopsoid isolates and Pfiesteria spp. isolates …………………………………... 163

Figure 4.7 Time-course growth/survival of three axenic cryptoperidiniopsoid isolates in supplemented media ………………... 164

Figure 4.8 Bivariate histograms of light scatter versus red autofluorescence (chla) for the cryptoperidiniopsoids ………………... 166

Figure 4.9 Time-course phototrophic growth/survival responses of three cryptoperidiniopsoid isolates in light and dark treatments ……… 167

Figure 4.10 Schematic for the life cycle of cryptoperidiniopsoids, based on flagellar and nuclear characters ……………………………... 170

Figure 5.1 (A-B) Excystment responses of Pfiesteria spp. after synchronization and purification; (C) purified cysts ………………….. 206

Figure 5.2 DNA-fluorescence plots for released flagellated cells of Pfiesteria spp. and cryptoperidiniopsoids …………………………….. 207

Figure 6.1 Light micrographs of Pfiesteria shumwayae vegetative cells ………… 225

Figure 6.2 Diagram of the most commonly observed patterns of Pfiesteria shumwayae reproductive cyst development ……..………... 227

Figure 6.3 Light micrographs of Pfiesteria shumwayae fusing gametes and a planozygote …………………………………………... 230

Figure 6.4 Scanning electron micrographs of Pfiesteria shumwayae early fusing gametes and planozygotes ……………………………………... 231

Figure 6.5 DNA-fluorescence and forward scatter signals from Pfiesteria shumwayae flagellated cell populations……………………. 232

Figure 6.6 DNA-fluorescence profiles for Pfiesteria shumwayae flagellated cells, fixed 6 h and 24 h after synchronous excystment ………...…….. 234

Figure 6.7 Light and fluorescence micrographs of Pfiesteria shumwayae sexual and feeding cells ……………………………………………….. 240

Figure 6.8 DNA profile time series of a synchronized Pfiesteria shumwayae flagellated cell population fed larval fish ……………………………... 242

Figure 6.9 Diagram of asexual reproduction observed in the Pfiesteria shumwayae cultures ……..………………………………... 246

Figure 6.10 Diagram of the proposed sexual cycle of Pfiesteria shumwayae……... 250

1. COMPARATIVE RESPONSE TO ALGAL PREY BY PFIESTERIA PISCICIDA, PFIESTERIA SHUMWAYAE SP. NOV., AND AN ESTUARINE ‘LOOKALIKE’ SPECIES

Matthew W. Parrow, Howard B. Glasgow, JoAnn M. Burkholder, and Cheng Zhang

In: Hallegraeff, G.M., Blackburn, S.I., Bolch, C.J., Lewis, R.J. [Eds.] Harmful Algal Blooms 2000. IOC UNESCO, 101-104 (2001).

1.1 Abstract

Zoospore production was examined during grazing on algal prey (Rhodomonas sp., in single-species batch-culture trials) by Neuse Estuary clonal isolates of Pfiesteria piscicida, Pfiesteria shumwayae sp. nov., and the related taxon Cryptoperidiniopsis gen. ined. Prior to the experiments, sub-cultures of P. piscicida from the same clone had been maintained for 4 months in fish-killing mode (TOX-A functional type, actively toxic; fish death at 12- to 24-hour intervals with 5,000 toxic zoospores/mL) and, separately, in temporarily nontoxic mode (TOX-B, tested as capable of killing fish but grown on cryptomonad prey, without fish, for 3 weeks). An older clone of P. piscicida was non- inducible (NON-IND, having lost its fish-killing ability over time in culture; grown for 6 months on cryptomonad prey). From the second Pfiesteria species, TOX-A (fish death at

1- to 2-hour intervals with 800 toxic zoospores/mL) and TOX-B cultures (same clone of origin) had been maintained similarly as TOX-A and TOX-B P. piscicida, respectively. The cryptoperidiniopsoid species had been fed cryptomonads similarly as had TOX-B Pfiesteria spp. NON-IND P. piscicida and the cryptoperidiniopsoid attained the highest zoospore production, appeared to retain kleptochloroplasts for longer periods, and exerted the lowest grazing pressure on algal prey, suggesting higher reliance on photosynthetic carbon acquisition. Zoospore production was intermediate by temporarily nontoxic or TOX-B

Pfiesteria spp., which were comparable in grazing pressure to TOX-A Pfiesteria spp.

Zoospore production was significantly lower in TOX-A P. piscicida (weakly toxic, based on zoospore densities and time required to kill fish) than in TOX-B or NON-IND cultures or in the cryptoperidiniopsoid. The lowest zoospore production occurred in the highly toxic

TOX-A strain of P. shumwayae sp. nov. These data suggest that zoospore production on

algal prey differs significantly among these dinoflagellates, perhaps depending on their history of toxic activity (Pfiesteria spp.) or their proclivity for mixotrophy.

1.2 Introduction

In the presence of live fish or their fresh tissues and excreta, dinoflagellates within the toxic Pfiesteria complex are stimulated to produce toxins or bioactive compounds that cause fish stress, disease, and death [1, 2]. The two toxic Pfiesteria species known thus far include P. piscicida and the more recently discovered P. shumwayae (formerly known as

Pfiesteria species B) [3, 4]. Toxic strains of these species are strongly attracted to live fish prey, based on computerized motion analysis [1] and microcapillary tube assays [5]. These characteristics – strong attraction to live fish, toxicity stimulated by the presence of live fish, and ability to produce ichthyotoxins (defined as in [6]) that cause fish death and disease – are required for species placement within the toxic Pfiesteria complex [1, 5, 6, 7].

Pfiesteria spp. have small, cryptic peridinoid zoospores [4, 8] that closely resemble the zoospores of various other, mostly undescribed ‘gymnodinoid’-like estuarine dinoflagellate species [1, 9, 10, 11]. Some of these species, such as Cryptoperidiniopsis spp. (gen. ined.) spp. [12], may have a complex life cycle [13] similar to that previously described for P. piscicida [8, 14] and P. shumwayae sp. nov. [4], with an array of flagellated, amoeboid and cyst stages. However, all Pfiesteria ‘lookalike’ species examined thus far, such as cryptoperidiniopsoid species and a species informally referenced as

‘shepherd’s crook [12], have shown no ability to cause fish disease and death in the standardized fish bioassay process (1, 4, 6, 7, 15) through toxin production by live cells at densities within field range [6, 7, 15, 16]. These replicated, repeated fish bioassays (10-12 weeks in duration) have included multiple strains, tested separately, from estuaries in

Maryland, Virginia, North Carolina, South Carolina, and Florida of the U.S. [6].

The above information on Pfiesteria spp. refers to toxic strains [1, 5, 6, 7, 15].

Many well known, so-called “toxic” algae (here, including toxic heterotrophic dinoflagellates as in [17]; e.g., toxic cyanobacteria [18], toxic prymnesiophytes [19], the toxic Pseudo-nitzschia complex [20], and the toxic Alexandrium complex [21]), are known to have naturally occurring strains which appear to be non-inducible or incapable of producing detectable toxin. For all known toxic species, molecular controls on toxin production are too poorly known to assess whether the non-inducible strains are genetically capable of toxin production [22]. Moreover, as an apparent artifact of being grown under laboratory conditions, many toxic strains of these species lose their ability to produce toxin over time in culture [20, 21, 23]. Within Pfiesteria spp., we similarly have identified functional types within one clone (strain), and among clones or strains, that differ in degree of toxicity from highly toxic to non-inducible [6, 7, 16].

Whether toxic or not, all strains of the two known Pfiesteria spp. have a flagellated zoospore stage with proficient grazing ability, particularly on some species of algal prey [1,

14]. These zoospores typically feed myzocytotically [24] by attaching peduncle to the prey cell and suctioning the contents into a food vacuole for digestion. Cryptoperidiniopsoid species consume algal prey by the same mechanism [13, 16]. In addition, Pfiesteria spp. and cryptoperidiniopsoid species can engage in limited phototrophy through acquisition of kleptochloroplasts from algal prey [4, 25]. The kleptochloroplasts can remain active within the food vacuole of the dinoflagellate for extended periods (days), providing a source of photosynthetic carbon acquisition [1, 25]. Our observations have suggested, however, that important differences exist between and within individual species with respect to algal grazing activity.

The objective of the present study was to evaluate the zoospore production, grazing, and potential for mixotrophy of different functional types of Pfiesteria piscicida, P. shumwayae sp. nov., and the cryptoperidiniopsoid species when given cryptomonad prey.

This work represents one facet of the research being conducted in our laboratory to further understanding of the complex nutritional strategies employed by toxic Pfiesteria complex dinoflagellates and certain species of close morphological resemblance (‘pfiesteria- like’species as in [6]; note that [1] used this term in reference only to ichthyotoxic species), and to strengthen insights about trophic controls on stage abundance and toxicity.

1.3 Materials and Methods

For this research, we defined a strain as a group of organisms within a species that is characterized by some particular quality [26]. We additionally considered the following functional types for Pfiesteria spp., within and/or among strains: (1) actively toxic (TOX-A

[6, 7], referred to as toxic in [1]), which had been in fish-killing mode immediately prior to the experiment; (2) temporarily nontoxic (TOX-B [6, 7], referred to as nontoxic in [1]), capable of ichthyotoxic activity but presently in a nontoxic mode – for example, given algal prey but not live fish); and (3) non-inducible (NON-IND in [6]), that is, incapable of producing bioactive substances that cause fish distress, disease and death, based on present understanding of induction of toxicity in these dinoflagellates [7]. Within the same strain

(here, clone), the TOX-B functional type serves as the ‘seed’ population for TOX-A

Pfiesteria. Over time in culture, as an artifact of laboratory conditions [7], both TOX-A and TOX-B functional types become non-inducible (in 98% of the clones, after 8-10 months for TOX-A with live fish, and after 2-6 months for TOX-B with algal prey; the remaining 2% of the clones, after 1-3 years) [7, 13].

Neuse Estuary clonal cultures were obtained in our laboratory using flow cytometric procedures as in [4, 7]. Species were identified from scanning electron microscopy (suture- swollen cells as in [1, 4]) and polymerase chain reaction molecular probes (PCR, as in [27,

28]). Toxicity of Pfiesteria spp. was verified by fish bioassays [1, 6, 7, 15]. Pfiesteria spp. can have endosymbiont bacteria, based on observations on various isolates using transmission electron micrographs [4, 16, 25]. Moreover, these heterotrophic dinoflagellates have not been grown successfully without addition of algae, fish, or other

prey [1]. Thus, in this study we considered clonal Pfiesteria as uni-dinoflagellate species culture, with endosymbiont bacteria and algal prey (as in [6]).

The three dinoflagellate taxa used in this study were tested separately with algal prey in single species trials. Six treatments were established in triplicate with an initial cell density of 150 dinoflagellate zoospores/mL, as follows: (1) Actively toxic P. piscicida

(P.pisc TOX-A functional type, Center for Applied Aquatic Ecology [CAAE] clone #271A-

1; considered a weakly toxic isolate, causing fish death in the standardized fish bioassay process [1, 6, 15] within 12 to 24 hours at cell densities of ca. 5,000 toxic zoospores

[TZs]/mL); (2) Temporarily nontoxic P. piscicida (P.pisc TOX-B functional type, CAAE clone #271A-1; tested as capable of killing fish, then grown on cryptomonad prey

[Rhodomonas sp., cloned from commercial source CCMP757], in nontoxic mode without fish for 3 weeks); (3) Non-inducible P. piscicida (P.pisc NON-IND functional type, CAAE clone #114-1-6; an older clone of kleptochloroplastidic P. piscicida that had lost its ability to cause fish disease and death over time in culture. This clone had not been toxic to fish upon repeated testing (with the fish bioassay process as in [1, 7, 15]) over the previous two years; it was grown for 6 months on cryptomonad prey prior to these experiments); (4)

Actively toxic Pfiesteria shumwayae sp. nov. (P.shum TOX-A functional type, CAAE clone #104T; regarded as a highly toxic isolate, capable of causing fish death in the standardized fish bioassay process [1, 6, 7, 15] < 1-2 hours at densities as low as 800

TZs/mL); (5) Temporarily nontoxic P. shumwayae sp. nov., (TOX-B functional type,

CAAE clone #104T; tested as capable of killing fish, then grown on cryptomonad prey without fish for 3 weeks); and (6) the cryptoperidiniopsoid species (CAAE Neuse clone

#7B-2), a kleptochloroplastidic strain that had been fed cryptomonads for 6 months

similarly as the TOX-B Pfiesteria spp. and the NON-IND P. piscicida functional types. An isolate of the NON-IND functional type of P. shumwayae sp. nov., similar in history to the isolate of NON-IND P. piscicida, was not available for this study.

In the experiments, the dinoflagellates were given algal prey as Rhodomonas sp.

(CCMP757, Bigelow Laboratory, Boothbay Harbor, Maine, U.S.A.) in a 1:15 predator : prey ratio, in batch cultures with both the predators and the prey added initially. This prey was selected because of known preference by Pfiesteria spp. for cryptomonads among algal prey species [14, 25]. The Rhodomonas sp. was cloned following its arrival from the commercial source using flow cytometric techniques as in [4, 7]. Algal prey controls

(without dinoflagellates) and dinoflagellate controls (without algal prey) were also established (n = 3) for each species and functional type. The cultures were maintained in

15 psu, sterile-filtered f/1000 medium, at 20°C and 50 µmol m-2 s-1, on a 12 : 12 hour light : dark cycle. They were gently mixed and subsampled at 12-hour intervals for 6 days. The subsamples were preserved with acidic Lugol’s solution [29], and were analyzed for cell abundance and species using light microscopy (200x, phase contrast, Olympus BH2 microscope, Olympus Corporation, Melville, NY; Palmer chambers as in [30]). These data were corroborated with a Coulter Multisizer IIe particle analyzer, Coulter Corp., Miami, FL.

One-way ANOVA was used to test for differences between controls and treatments, and among treatment means. All probability values were considered significant at p < 0.05

[31].

Mixotrophic potential was putatively assessed for controls and each treatment at the end of the experiment (day 6) using light and epifluorescence microscopy (600x, phase contrast, Olympus BH2 microscope with fluorescence attachment) on samples that had

been preserved with 2% buffered formalin [25]. Zoospores containing chloroplasts were readily discerned from those without, since the pigmented inclusions autofluoresced reddish-orange under UV excitation [4, 25]. Starch bodies were easily distinguished as well, in phase contrast [4, 25]. The percentage of zoospores containing typical chloroplast inclusions and starch bodies indicative of kleptochloroplastidy in each treatment was determined for at least 100 cells per sample, and was inferred to be directly related to mixotrophic potential [25]. Growth and ingestion rates were estimated for the time interval when the zoospore populations exhibited logarithmic growth (days 1-5), following the equations of [32, 33].

1.4 Results

All dinoflagellate controls (without algal prey) demonstrated no detectable zoospore production during the course of the experiment, with all cells encysting or otherwise disappearing from the water column by the end of the experiment (day 6). In the experimental treatments, the three dinoflagellate species and functional types differed significantly in their response to the algal prey (Table 1.1). The ‘TOX-A’ Pfiesteria spp.

(from fish-killing cultures; term used here in reference to the functional type status at the beginning of the experiment) achieved significantly lower cell production when given algal prey than temporarily nontoxic TOX-B Pfiesteria spp. and the cryptoperidiniopsoid (Fig.

1.1). Among the treatments with actively toxic functional types, the (more weakly toxic)

TOX-A Pfiesteria piscicida had significantly higher cell production on algal prey than the highly toxic TOX-A Pfiesteria shumwayae, which showed the lowest zoospore production.

NON-IND P. piscicida attained significantly higher zoospore production on algal prey than the other dinoflagellate species and functional types tested, with the cryptoperidiniopsoid species having the next highest production by the end of the experiment. Thus, dinoflagellate zoospore production on cryptomonad algal prey appeared to be directly related to the history of toxicity (or lack thereof, in the case of the cryptoperidiniopsoid species and the NON-IND Pfiesteria in fish bioassays [7]), as follows:

NON-IND > cryptoperi- >> TOX-B >> weakly toxic > highly toxic P. piscicida diniopsoid sp. Pfiesteria spp. isolate, TOX-A isolate, TOX-A P. piscicida P. shumwayae sp. nov.

Table 1.1. Mean zoospore specific growth rates, division times, maximum yields, ingestion rates, and percentage with mixotrophic inclusions by the end of the 6-day experiment, for each species and strain tested. Specific growth and ingestion rates were calculated for the time interval of exponential zoospore production (days 1-5), excluding the influence of other life stages.

Specific Growth Mean Maximum Yield Ingestion Rate (%) with Rate ( k )1 Division ( cells • mL-1 ) (I)1 Mixotrophic Treatment -1 (divisions • day ) Time (Td) ( h ) ( prey cells • zoo- Inclusions spore-1 • day-1 ) on Day 62

P. piscicida 1.43 ± .04 16.8 ± 0.5 8543±1200 9.4 ± 0.32 95 NON-IND P. piscicida 1.42 ± .05 17.0 ± 0.61 3470 ± 446 54.5 ± 3.4 8 TOX-B P. piscicida 0.75 ± .04 32.1 ± 1.8 1233 ± 136 55.8 ± 3.9 < 1 TOX-A P. shumwayae sp. nov. TOX-B 0.84 ± .10 29.2 ± 3.2 1817 ± 339 37.3 ± 1.9 10

P. shumwayae 0.22 ± .07 127 ± 31.2 592 ± 118 8.72 ± 0 .41 < 1 sp. nov. TOX-A3 cryptoperidin. 1.15 ± .05 20.9 ± 0.85 3749 ± 777 21.0 ± 2.4 70

1 Rates were calculated following the equations of [32, 33].

2 Percentage of zoospores containing autofluorescent pigmented inclusions and localized starch bodies was determined microscopically for at least 100 cells · sample-1.

3 Values for the TOX-A functional type of P. shumwayae sp. nov. are based on cell production estimates from clone #104T only. Other clonal P. shumwayae cultures (n = 3) that were treated similarly had negligibile zoospore production during the experimental period.

By the end of the experiment, the Pfiesteria spp. functional types that achieved highest zoospore production generally had exerted intermediate grazing pressure on their algal prey (Fig. 1.2). This apparent contradiction may have resulted from an increased proclivity for mixotrophy that was suggested by the TOX-B and NON-IND functional types in comparison to the TOX-A Pfiesteria spp. Most zoospores of the NON-IND functional type appeared to have functional kleptochloroplasts with chlorophyll autofluorescence, and apparently achieved higher cell production through mixotrophic nutrition relative to the

TOX-A and TOX-B Pfiesteria spp. and the cryptoperidiniopsoid. Thus, the dinoflagellates varied in kleptochloroplast retention and apparent partial reliance on photosynthesis in their nutrition.

The cryptoperidiniopsoid zoospores also apparently engaged in mixotrophy. By the end of the experiment, about 70% of the cells may have contained viable kleptochloroplasts as indicated by chlorophyll autofluorescence. These cells were enlarged and were filled with darkly staining starch bodies. TOX-B Pfiesteria spp. zoospores appeared intermediate in kleptochloroplast retention, with ca. 8-10% of the cells having viable kleptochloroplasts as indicated by chlorophyll autofluorescence. In contrast, TOX-A Pfiesteria spp. zoospores did not retain kleptochloroplasts (proportion with kleptochloroplasts < 1%), and likely depended solely upon heterotrophy for their nutrition with no contribution from photosynthesis.

The potential for partial reliance on photosynthesis or a mixotrophic habit was also inferred from the grazing activity on cryptomonad prey among the dinoflagellate species and functional types. The highest estimated ingestion rates were demonstrated by toxic

Pfiesteria piscicida (TOX-A and TOX-B, also indicated by the depletion of the prey food

source). NON-IND P. piscicida and the cryptoperidiniopsoid showed significantly lower grazing activity. Thus, the high growth rate and cell production achieved by NON-IND P. piscicida with lower algal grazing activity, in comparison to the lower growth rate and cell production achieved by toxic Pfiesteria spp. with highest grazing activity, apparently was accomplished through more substantial reliance on photosynthesis from kleptochloroplasts.

TOX-A P. piscicida exhibited tight coupling in predator-prey oscillations relative to NON-

IND P. piscicida, indicating a higher degree of (heterotrophic) reliance on the prey population for zoospore production than did the kleptochloroplastidic (mixotrophic) NON-

IND P. piscicida [25, 34, 35] (Fig. 1.3).

P.pisc TOX-A 9 P.pisc TOX-B 8 P.pisc NON-IND P.shum TOX-A 7 P.shum TOX-B / ML 3 Cryptoperidiniopsoid 6

3 ZOOSPORES x 10

123456 TIME ( DAYS )

Figure 1.1. Response of different functional types of P. piscicida, P. shumwayae sp. nov., and the cryptoperidiniopsoid zoospores to Rhodomonas sp. prey in 6-day trials. NON-IND

P. piscicida and the cryptoperidiniopsoid attained highest zoospore production on algal prey, with temporarily nontoxic (TOX-B) and actively toxic (TOX-A) Pfiesteria spp. attaining less production, respectively.

1.E+06 Rhodomonas Only With P.pisc NON-IND With P.pisc TOX-B 1.E+05 With P.pisc TOX-A

1.E+04

1.E+03

) ML / ( CELLS RHODOMONAS

1.E+02 123456 TIME ( DAYS )

Figure 1.2. Rhodomonas sp. cell production in 6-day batch trials with different functional types of P. piscicida as predators, in comparison to controls with no predation. NON-IND

P. piscicida exerted significantly less grazing pressure on algal prey than did the TOX-B and TOX-A functional types, but attained much higher zoospore production (see Fig. 1.1).

10 9 P.pisc TOX-A (heterotrophic) P.pisc NON-IND (mixotrophic) 8 / ML 3 Algal prey in NON-IND 7 Algal prey in TOX-A 6 5 4

x 10 ZOOSPORES 3

2 1 0 123456 TIME ( DAYS )

Figure 1.3. Variable population dynamics between toxic (TOX-A) P. piscicida with

Rhodomonas sp. prey and NON-IND P. piscicida with Rhodomonas sp. prey in 6-day trials.

The TOX-A functional type demonstrated tight apparent coupling in predator-prey oscillations relative to the NON-IND P. piscicida.

1.5 Discussion

The nutritional strategies of the dinoflagellate species examined here ranged from heterotrophy (actively toxic [TOX-A] Pfiesteria spp.) to apparent increased reliance on functional mixotrophy (temporarily nontoxic [TOX-B] Pfiesteria spp., the cryptoperidiniopsoid, and non-inducible [NON-IND] Pfiesteria spp.). These differences in trophic strategy may have promoted significant differences in population dynamics among the species and strains. Even within a single species of Pfiesteria, the actively toxic, temporarily nontoxic, and non-inducible functional types exhibited distinct population dynamics with algal prey – differences that appeared to be related to the heterotrophic or mixotrophic nutrition of each.

When in recently TOX-A mode, Pfiesteria spp. maintained low growth rates on a diet of algal prey without live fish, relative to the other two functional types that had been without fish prey for an extended period (several weeks or longer). This trend also was demonstrated by [7]. Nevertheless, the most tightly coupled Lotka-Volterra fluctuations were demonstrated for the TOX-A P. piscicida zoospores and their cryptomonad prey [34].

The species and strains that were capable of kleptochloroplastidy engaged in limited photosynthesis and, thus, likely were less reliant on the available prey population to supply their carbon requirement.

Mixotrophy, via the retention of functional chloroplasts from the algal prey, apparently enabled substantial increase in growth rates and cell production for the strains and functional types of Pfiesteria spp. and cryptoperidiniopsoid species that are capable of this form of photosynthetic carbon acquisition. Although (weakly) TOX-A Pfiesteria piscicida maintained the highest grazing activity on algal prey, its growth rate and cell

production were low relative to that of other P. piscicida functional types fed the same prey, suggesting that recently toxic populations are not as adept at retaining and utilizing photosynthesis from kleptochloroplasts in supplying carbon and energy needed for cell production. The (highly) TOX-A P. shumwayae zoospores exhibited both the lowest growth and ingestion rates on algal prey of any taxa tested, and exhibited no measurable retention of kleptochloroplasts during the experiment. Whether these differences were due to the experimental treatment, or endemic to the species or clone, is under investigation.

Thus, the cytological switch from pure heterotrophy to mixotrophy may not be instantaneous. Instead, certain biochemical and/or environmental controls may regulate this process. We are currently investigating the efficiencies, timing, and cytology involved in altered reliance on heterotrophy versus mixotrophy in the nutritional ecology of the toxic

Pfiesteria complex and closely related species.

Acknowledgments

Funding for this research was provided by an anonymous foundation, the North

Carolina General Assembly, the U.S. Environmental Protection Agency, the Department of

Botany at North Carolina State University, and the National Oceanic & Atmospheric

Administration Ecohab program. We thank N. Deamer-Melia, A. Hamilton, E. Hannon, and J. Springer for their counsel and assistance.

1.6 References

1. J.M. Burkholder and H.B. Glasgow, Limnol. & Oceanogr., 42, 1052-1075 (1997).

2. E.R. Fairey, J.S. Edmunds, N.J. Deamer-Melia, H. B. Glasgow, F.M. Johnson, P.R. Moeller, J.M. Burkholder, and J.S. Ramsdell, Env. Health Perspec., 107, 711-714 (1999).

3. J.M. Burkholder, Ecol. Appl. 8(1) Suppl., S37-S62 (1998).

4. H.B. Glasgow, J.M. Burkholder, S.L. Morton, and J. Springer, Phycologia (accepted).

5. P. Cancellieri, J.M. Burkholder, N.J. Deamer-Melia, and H.B. Glasgow, J. Exp. Mar. Biol. Ecol. (accepted).

6. Pfiesteria Interagency Coordination Working Group -- Glossary of Pfiesteria-Related Terms. U.S. Environmental Protection Agency, Baltimore (available at http://www.redtide.whoi.edu/pfiesteria/documents/glossary/html (2000).

7. J.M. Burkholder, Phycologia (accepted).

8. K.A. Steidinger, J.M. Burkholder, H.B. Glasgow, E. Truby, J. Garrett, E.J. Noga, and S.A. Smith, J. Phycol., 32, 157-164 (1996).

9. K.A. Steidinger, J.H. Landsberg, E.W. Truby, and B.A. Blakesley, Nova Hedwigia, 112, 415-422 (1996).

10. J.M. Burkholder, H.B. Glasgow, and C.W. Hobbs, Mar. Ecol. Prog. Ser., 124, 43-61 (1995).

11. J.M. Burkholder, M.A. Mallin, and H.B. Glasgow, Mar. Ecol. Prog. Ser., 179, 301-310 (1999).

12. K. Steidinger and colleagues, Florida Dept. of Environmental Protection – Florida Marine Research Institute, St. Petersburg, Florida, personal communication (2000).

13. D.W. Seaborn, A. Seaborn, W. Dunston, and H.G. Marshall, Va. J. Sci. 50, 337-344 (1999).

14. J.M. Burkholder, and H.B. Glasgow, Arch. Protistenk., 145, 177-188 (1995).

15. H.G. Marshall, A.S. Gordon, D.W. Seaborn, B. Dyer, W.M. Dunstan, and A. Seaborn, J. Exp. Mar. Biol. Ecol., 255, 65-74 (2000).

16. H.B. Glasgow, J.M. Burkholder, S.L. Morton, J. Springer, M.W. Parrow, and P. Cancellieri, Fish-killing activity and nutrient stimulation of a second toxic Pfiesteria species, in this volume.

17. J.M. Burkholder, Sci. Amer., 281, 42-49 (1999).

18. W.W. Carmichael, Scientific American, 270(1), 78-86 (1994).

19. A.S. Meldahl, and B. Edvardsen et al., J. Toxicol. Env. Health, 42, 289-301 (1994).

20. S.S. Bates, M.F. Hiltz, and C. Leger, Proceedings of the Sixth Canadian Workshop on Harmful Marine Algae, no. 2261, 21-26 (1999).

21. D.M. Anderson, Toxic Marine Phytoplankton: Proceedings of the Fourth International Conference on Toxic Marine Phytoplankton, E. Graneli, B. Sundstroem, L. Edler, and D.M. Anderson, eds. (Elsevier Press, New York) pp. 41-51 (1990).

22. J.M. Burkholder and J.J. Springer, Signalling in dinoflagellates, in: Microbial Signalling and Communication, R.R. England, G. Hobbs, N.J. Bainton, and D.McL. Roberts, eds., Cambridge University Press, New York, pp. 220-238 (1999).

23. P. Gentien and G. Arzul, J. Mar. Biol. Ass. U.K., 70, 571-581 (1990).

24. E. Schnepf and M. Elbrächter, Europ. J. Protistol., 28, 3-4 (1992).

25. A.J. Lewitus, H.B. Glasgow, and J.M. Burkholder, J. Phycol., 35, 303-312 (1999).

26. Academic Press Dictionary of Science and Technology, C. Morris (ed.), Academic Press, Boston, 2432 pp. (1992).

27. P.A. Rublee, J. Kempton, E. Schaefer, J.M. Burkholder, H.B. Glasgow, and D. Oldach, Va. J. Sci., 50, 325-336 (1999).

28. D.W. Oldach, C.F. Delwiche, K.S. Jakobsen, T. Tengs, E.G. Brown, J.W. Kempton, E.F. Schaefer, H. Bowers, H.B. Glasgow, J.M. Burkholder, K.A. Steidinger, and P.A. Rublee, Proc. Nat. Acad. Sci., 97-8, 4303-4308 (2000).

29. R.A. Vollenweider (ed.), A Manual on Methods for Measuring Primary Production in Aquatic Environments, 2nd edn, International Biological Programs Handbook No. 12 (Blackwell Scientific Publications, Oxford) (1974).

30. R.G. Wetzel and G.E. Likens, Limnological Analyses, 2nd edn. (W.B. Saunders, Philadelphia) (1991).

31. SAS Institute, Inc. SAS/STAT guide for personal computers, version 6. SAS Institute, Inc., Cary, North Carolina (1987).

32. B.W. Frost, Limnol. Oceanog., 17(6), 805-815 (1972).

33. J.F. Heinbokel, Mar. Biol., 47, 177-189 (1978).

34. V. Volterra, Variations and fluctuations of the numbers of individuals in animal species living together, reprinted in 1931. In: Animal Ecology, R.N. Chapman, ed., McGraw-Hill, New York, 7.4.1 (10.2) (1926).

35. J.R. Stein (ed.), Handbook of Phycological Methods: Culture Methods and Growth Measurements, Cambridge University Press, New York (1973).

2. FLOW CYTOMETRIC DETERMINATION OF ZOOSPORE DNA CONTENT AND POPULATION DNA DISTRIBUTION IN CULTURED PFIESTERIA SPP. (PYRRHOPHYTA)

Matthew W. Parrow and JoAnn M. Burkholder

Journal of Experimental Marine Biology and Ecology 267 (2002) 35-51.

2.1 Abstract

The relative cellular DNA content from 23 different clonal cultures of Pfiesteria spp. zoospores was determined using a DNA fluorochrome and flow cytometry. Significant differences between Pfiesteria piscicida and Pfiesteria shumwayae were detected, both in mean zoospore DNA content and population cell cycle DNA distribution. Intraspecific differences in DNA content were found between clonal zoospore cultures established from different geographical regions. Long-term cultures (years) of P. piscicida were available for testing, and a negative correlation between zoospore DNA content and time in culture was observed. Zoospore cell cycle-related DNA distributions were also markedly different between the two species in these clonal cultures. In all cultures tested, P. piscicida zoospores exhibited bimodal DNA flow histograms suggesting G1-S-G2+M distributions, typical of eukaryotic asynchronously cycling cells. In contrast, cultures of P. shumwayae vegetative zoospores exhibited one DNA peak distribution, indicative of synchronized cells.

These data are consistent with the hypothesis that P. shumwayae zoospores are interphasic cells, and mitosis in zoospore cultures of this species predominantly occurs as benthic or adherent non-motile division cysts. Light microscopy observations of the nuclear condition of electrostatically sorted zoospores of each Pfiesteria species also support this hypothesis.

If highly conserved, this disparity in modes of vegetative reproduction would ramify the population dynamics of the two Pfiesteria species.

2.2 Introduction

One of the first and most widespread applications of flow cytometry in aquatic microbiology has been quantification of DNA using DNA fluorochromes (Collier, 2000).

The primary benefit offered by flow cytometry over bulk DNA determinations is that the

DNA content of each individual cell is measured discretely, which allows discrimination of variability in the DNA content of subpopulations within a sample. For aquatic unicellular organisms, quantification of cellular DNA and the cell cycle distribution of proliferating cells can yield valuable information about intrinsic differences between cells or populations

(Chisholm et al., 1986). Also, since life cycle events are usually largely correlative with cell cycle events for unicellular organisms, cell-specific DNA estimations can allow elucidation of the relationship between morphologically distinct alternating cell types

(Edvardsen and Vaulot, 1996). Flow cytometry has provided a great deal of information about DNA content, cell cycle dynamics, and aspects of life cycles in a variety of aquatic microorganisms, including planktonic bacteria, prymnesiophytes, chlorophytes, and dinoflagellates (mostly photosynthetic species – Trask et al., 1982; Olsen et al., 1983;

Yentsch et al., 1983; Whiteley et al., 1993; Edvardsen and Vaulot, 1996; Veldhuis et al.,

1997).

To date, flow cytometric DNA analyses have been published for only two heterotrophic dinoflagellates, Crypthecodinium cohnii Biecheler (e.g., Wong and Whiteley,

1996; Bhaud et al., 2000) and Oxyrrhis marina Dujardin (Whiteley et al., 1993).

Heterotrophic protists largely have been avoided as subjects of flow cytometric DNA determinations, because they generally must be cultured with prey. The prey DNA can be difficult to differentiate from that of the cells of interest, and compounded problems can

occur if the prey chlorophyll autofluorescence (e.g., algal prey) overlaps the emission spectra of the DNA fluorochrome used (Whiteley et al., 1993). Also, it is difficult to obtain cell cycle synchronized cultures of heterotrophic protists, making them somewhat unfavorable as model organisms for cell cycle studies (Wunderlich and Peyk, 1969). C. cohnii is amenable to flow cytometric DNA analyses because it can be grown on defined chemical media, and cultures can be manipulated to produce short-term synchronous populations (Wong and Whiteley, 1996). The investigation of O. marina available from the published literature was conducted on extracted nuclei, which allowed flow cytometric distinction between the dinoflagellate and prey DNA (Whiteley et al., 1993).

Here, we used flow cytometry to investigate zoospore DNA content and population

DNA distribution in 23 clonal cultures of the heterotrophic estuarine dinoflagellate species

Pfiesteria piscicida Steidinger & Burkholder and Pfiesteria shumwayae Glasgow &

Burkholder within the family Pfiesteriaceae (Dinamoebales) (Steidinger et al., 1996a;

Glasgow et al., 2001). Both species have small (diameter ca. 8 µm) heterotrophic zoospores that closely resemble the zoospores of various other "gymnodinoid"-like estuarine dinoflagellate species (Steidinger et al., 1996b). The two species thus far have demonstrated a wide, largely overlapping geographic distribution in estuarine waters and sediments of the eastern U.S. seaboard and the U.S. Gulf Coast, and recently have been found in some European and Asiatic coastal environments (Allen, 2000). Many aspects of the complex life cycles of Pfiesteria spp. have been previously described (Burkholder and

Glasgow, 1995, 1997; Burkholder et al., 2001a). The focus of the present work was the vegetatively reproducing biflagellate zoospore stages that are most commonly observed in culture. For this research, we developed a sample preparation method that permitted

analysis of fixed zoospore cells (rather than isolated nuclei), so as to conserve other useful flow cytometric measurements such as light scatter (a sizing parameter) and autofluorescence of chloroplast inclusions (ingested prey). The sample preparation method also enabled storage of cells for later analysis and yielded a high concentration of fixed cells to permit rapid measurement intervals. The efficiency of the fixation and staining methods, and the minimalization of prey DNA confounding effects, were assessed based on observations with epifluorescence microscopy, and on acceptable coefficients of variation

(CVs) in the DNA fluorescence peak distributions obtained. From observations on clonal cultures, we detected an apparent difference in the predominant mode of vegetative reproduction for zoospores of the two Pfiesteria species – namely, zoospores of P. piscicida were often found swimming as pairs and thus may undergo mitosis while still motile, whereas zoospores of P. shumwayae underwent mitosis as benthic or adhesive nonmotile division cysts. We also used flow cytometric DNA analysis to explore this distinction.

2.3 Materials and Methods

2.3.1 Cultures. Clonal cultures of the two Pfiesteria spp. were obtained by micropipette isolation and, more recently (during the past two years), by flow cytometric particle sorting (Burkholder et al., 2001b; Glasgow et al., 2001). All species were verified by scanning electron microscopy (SEM) and polymerase chain reaction molecular probes

(PCR; Steidinger et al., 1996b; Rublee et al., 1999; Burkholder et al., 2001b; Glasgow et al., 2001). Thus far, these heterotrophic dinoflagellates have not been grown to yield moderate to high cell production unless algae, fish, or other prey have been added.

Therefore, cultures were maintained on a diet of live fish (juvenile tilapia, Oreochromis mossambicus Peters; Pfiesteria Interagency Coordinative Working Group [PICWG], 2000;

Burkholder et al., 2001b) or cryptophyte algae (Rhodomonas sp. CCMP757, with the algal prey having been cloned from commercial material provided by the Culture Collection for

Marine Phytoplankton, Bigelow Laboratory, Bigelow, ME) (Burkholder and Glasgow,

1995; Lewitus et al., 1999; Marshall et al., 2000; PICWG, 2000; Burkholder et al., 2001b;

Parrow et al., 2001). Since moderate salinity effects have been reported for one of the DNA fluorochromes employed (Veldhuis et al., 1997), all cultures were maintained at a salinity of 15 (United Nations Educational, Scientific, and Cultural Organization 1985).

2.3.2 Sample Preparation. Zoospore cultures were harvested during late logarithmic cell production in algal-fed batch cultures, or when zoospore density had reached high levels in excess of 104 cells ml-1 in semi-continuous cultures growing on fish prey. Briefly,

50 ml of culture volume were collected and filtered (25-µm porosity; Fisher Scientific,

Pittsburgh, PA, 09-795AA,) to remove particulate debris. The effectiveness of glutaraldehyde versus paraformaldehyde as fixatives (in each case, 1% final concentration)

was experimentally compared. The preserved samples were stored for > 24 h in darkness at

4 ˚C. Zoospores were then concentrated ten-fold by centrifugation and returned to storage conditions. This process yielded sufficient sample volume for repeated analyses, and cell concentrations (c. 105 zoospores ml-1) suitable for DNA analysis, with minimal cell clumping. Prior to analysis, 1-ml aliquots of the stock cell suspensions were removed and treated with RNAse A (1 µg ml-1 final solution as weight to volume [w/v]; Sigma, St.

Louis, MO, R-4875) for 1 h at room temperature.

The effectiveness of propidium iodide [PI] (final solution 3, 10, and 50 µg ml-1;

® Sigma, P-4170) and SYTOX Green (final concentration 5, 10, and 15 µM; Molecular

Probes, Inc., Eugene, OR, S-7020) were experimentally compared as DNA stains for the two Pfiesteria spp. Both stains have negligible base-pair specificity, and both have been successfully used in DNA analysis of various phytoplankton species (e.g., Vaulot et al.,

1994; Green et al., 1996; Velduis et al., 1997). Stained zoospores of each Pfiesteria species were incubated in darkness for 2 h or, alternatively, overnight (12-16 h) in darkness at 4 ˚C

(as in Eschbach et al., 2001). A stock suspension of chicken red blood cells (CRBCs,

Sigma, R-0504) was prepared in PBS (Sigma, D-5652) at a salinity of 15 and used as an internal DNA standard for each analysis. For each sample, the mean DNA fluorescence for a minimum of 104 zoospores under the G1 peak was compared to the CRBC signal, and zoospore DNA values were expressed as CRBC units (Vindelov et al., 1983; Veldhuis et al., 1997; Johnston et al., 1999).

2.3.3 Flow Cytometric Techniques. Flow cytometric analysis and cell sorting were completed using a COULTER® EPICS® ALTRATM Flow Cytometer with HyPerSortTM

System (Coulter Corporation, Miami, FL) equipped with a water-cooled INNOVATM

Enterprise IITM Ion Laser (Coherent, Inc., Santa Clara, CA). The instrument was equipped with a 100 µm orifice quartz flow cell, and excitation was with 150 mW power of the 488 nm laser line, focused to an elliptical spot 6 µm x 112 µm at the point of interrogation.

Instrument optics were selected to detect fluorescence from stain-DNA complexes on photomultipliers screened with appropriate band-pass filters (525 nm, SYTOX Green; 610 nm, PI). All scatter and fluorescence signals were recorded as peak signals. Signal amplification and threshold level were optimized such that the zoospore DNA distribution fell mid-scale. Unstained samples were used as negative controls. Optical alignment and stability was monitored using 10 µm diameter fluorescent latex microspheres (Coulter

Corporation, PN6605359), which yielded CVs < 1.5 %. Linearity of the flow cytometer amplification system was carefully checked using Immuno-Brite fluorospheres (Coulter

Corp., PN6603473) (Bagwell et al., 1989). Frequency-modulated electrostatic cell sorting was performed in standard sorting mode 3 (for purity in preference of yield), and electronic sort gates were user-defined to encompass any detectable fluorescence-DNA distribution subpopulation of interest. Target cells were collected on microscope slides or in test tubes, and morphology and nuclear condition was examined with light and epifluorescence microscopy (200-600x, Olympus AX70 research microscope, Olympus Corporation,

Melville, NY). Images were captured with a DEI-750 cooled-chip CCD camera (Optronics

Engineering, Goleta, CA). Sort purity (the percentage of gated cells that fell in the original sort region after re-analysis) was periodically checked, and consistently was > 95%.

2.3.4 Data Analyses. Listmode file analysis for DNA peak numbers, means, and

CVs was determined for zoospores and CRBCs using EXPO 32TM Analysis software

(Applied Cytometry Systems, Sheffield, UK). Where applicable, DNA histogram deconvolution and analysis were conducted using MultiCycle DNA Content and Cell Cycle

Analysis Software (Phoenix Flow Systems, Inc., San Diego, California). One-way

ANOVA was used to test for differences between controls and treatments, and among treatment (sample) means (SAS Institute, Inc. 1997). Probability values were considered significant at p < 0.05.

2.4 Results

2.4.1 Vegetative Division. In these cultures, P. piscicida zoospores often occurred as pairs which were occasionally observed to separate, and thus may divide vegetatively

(asexually) while still flagellated and motile. In P. shumwayae cultures, zoospores often fed until satiated as indicated by halted feeding and a swollen, sluggish appearance, with 1-

2 spherical, prey-packed food vacuole(s) occupying the majority of the epithecal portion of the cell. These zoospores often swam to the bottom of the culture container, shed their flagella, formed rounded cells, and encysted. The cysts that were formed initially appeared as relatively smooth, occlusive-walled coccoid bodies. After an indeterminate amount of time (hours to days), a cleavage furrow commonly developed, and the cyst protoplast gradually divided (Fig. 2.1A, B). Often the two cleavage products separated completely, and asynchronous excystment of offspring zoospores was observed after a variable dormancy period of > 8 h. Some cysts did not undergo cytokinesis during a 2-week period of observation, however, and were not observed to yield flagellated products. P. shumwayae vegetative division cysts in algal-fed cultures often contained a prominent reddish inclusion, perhaps analogous to the food bodies described in the asexual division cysts of the estuarine myzocytotic heterotroph Katodinium (Gymnodinium) fungiforme

(Anissimova) Loeblich III (Spero and Morée, 1981). In these P. shumwayae division cysts, such inclusions were often observed to emit a faint red autofluorescence when viewed under epifluorescence microscopy, which suggested an algal prey origin (Lewitus et al.,

1999). It is unknown whether these inclusions bear any affinity to the "red drops" that have been described in sexual (hypnozygote) cysts of certain other dinoflagellate species (e.g.,

Bibby and Dodge, 1972; Pfiester, 1976; Loeblich and Loeblich, 1984).

Figure 2.1

Figure 2.1. (A). DIC light micrograph of division cysts from clonal P. shumwayae culture.

The uppermost cyst had undergone cytokinesis, but the two protoplasts remained inside the original cyst wall. After excystment the outer translucent cyst wall remained (arrow).

When grown on algal prey (see text), a distinct reddish inclusion was apparent in many, but not all, division cysts of this species, bar = 10 µm. (B). Scanning electron micrograph of a

P. shumwayae vegetative division cyst from a clonal culture. The cyst surface was relatively smooth, but showed structures suggestive of dinoflagellate thecal sutures. Rod- shaped bacteria readily colonized the surface of these cysts, bar = 10 µm.

These vegetative division cysts of P. shumwayae tended to aggregate around irregular surfaces such as particulate debris, to which they adhered. They could easily be missed in routine sampling of cells suspended in the culture medium. In live cultures, these cysts became conspicuous especially in late logarithmic phase and stationary phase, and they persisted if zoospore populations were allowed to become prey-limited. We documented (with video microscopy) encystment, division, and excystment of two offspring zoospores. Excystment apparently occurred via an excystment aperture through which a clear amorphous cell extruded and, within seconds, developed typical zoospore morphology. In P. shumwayae grown with either algal or fish prey, after zoospore excystment, a deflated translucent outer covering was left behind (Fig 2.1A).

2.4.2 Sample Quality and DNA Staining. Comparison of the two fixatives showed that glutaraldehyde preserved the zoospores of both Pfiesteria spp. without morphological distortion. However, this fixative induced green cytoplasmic fluorescence that led to > 20% overestimation of DNA in SYTOX Green-stained cells. Paraformaldehyde fixation preserved zoospores with comparable morphological quality, but did not cause detectable cytoplasmic fluorescence and, therefore, was selected for use in the remainder of the study.

Comparison of the two DNA stains revealed that PI treatment resulted in substantial cytoplasmic fluorescence of the zoospores of both Pfiesteria spp., regardless of staining concentration and incubation time. In algal-fed cultures, prey chlorophyll autofluorescence also overlapped the zoospore PI-DNA fluorescence distribution, which further limited the utility of this fluorochrome. In contrast, SYTOX Green fluorescence was distinctly localized within the nuclei of Pfiesteria spp. zoospores, and no significant competing

fluorescence was found, regardless of prey type. Therefore, SYTOX Green was selected over PI for this study.

The selected sample preparation method provided a high yield of cells, with negligible clumping and good preservation of zoospore morphology. Epifluorescence microscopical inspection of RNAse-treated DNA stained cells revealed a bright DNA-

SYTOX Green complex, restricted to the zoospore nuclei. The binding was sufficiently specific that individual fluorescent chromosomes could be easily discerned, with very little to no background cytoplasmic fluorescence (Fig. 2.2A, B). In algal-fed cultures, zoospores with red autofluorescent chlorophyll inclusions (remnants of undigested prey chloroplasts) occasionally were seen, but instances were relatively low (c. 0-10% for stationary phase or prey-depleted populations), most likely because batch algal-fed cultures were harvested during late-logarithmic population growth when algal prey were scarce (e.g., as in Lewitus et al., 1999). In all stained samples, occasional zoospores were observed with a diffuse green fluorescence associated with the epithecal food vacuole, which was interpreted to be from remnants of undigested prey DNA. This diffuse green fluorescence was rare, however, and dim in comparison to the brilliant green fluorescent intensity of the relatively large dinoflagellate nucleus (Burkholder et al., 2001b). Prey DNA is rapidly degraded in the digestive vacuole of these dinoflagellates (also described by Lewitus et al., 1999, for P. piscicida). Thus, intracellular prey DNA effects were minimal in these treatments, as also indicated by the relatively low cytometric CVs. Algal-fed cultures additionally contained a low percentage of intact prey cells, but the low fluorescent intensity of the relatively small cryptophyte nuclei did not overlap the zoospore DNA distributions, and was below the

DNA fluorescence peak signal threshold set for the analyses (Whiteley et al., 1993).

A B

Figure 2.2. SYTOX Green-stained G1 zoospores of Pfiesteria piscicida, viewed under epifluorescence microscopy with mercury lamp excitation (note that these cells showed no discernable cytoplasmic green fluorescence). (A) Two P. piscicida zoospores (600x, water immersion objective; bar = 5 µm. (B) P. piscicida zoospore, showing the specific dye binding to the distinct condensed dinoflagellate chromosomes (1000x, oil immersion objective; bar = 1 µm).

Therefore, the cellular DNA values obtained from this technique were assumed to predominantly indicate nuclear DNA (and note that non-nuclear DNA [mitochondrial, etc.] is considered to comprise a small percentage of the total cellular DNA in other dinoflagellate species; Holm-Hansen, 1969; Karentz, 1983; Veldhuis et al., 1997). It should be noted that the adherent nature of the P. shumwayae division cysts hindered adequate sampling of this cell type for flow cytometry. Furthermore, P. shumwayae cysts were impervious to the DNA staining method optimized here for zoospores. Therefore, these cysts were not included in the flow cytometric DNA analyses. However, application of cell permeabilization techniques (modified from Kempton, 1999) permitted microscopical visualization of stained cyst nuclei in selected preparations.

A three-fold increase in SYTOX Green concentration did not yield an increase in the DNA-fluorescence intensity measured, either for the zoospores or the CRBC DNA standards. Thus we concluded that the 5 µM concentration used was sufficient to saturate the sample DNA. Increasing the incubation time to 3 days also did not alter the zoospore

DNA peak histogram means more than 5% (data not shown). The CRBC standards were uniform in DNA-fluorescence intensity from one analysis and day to the next, whether stained as internal or external standards. SYTOX Green-stained CRBCs that had been stored in darkness at 4 ˚C for two weeks exhibited < 1% difference in DNA peak mean in comparison to CRBCs that were prepared similarly 16 h prior to analysis.

2.4.3 Cytometric Analyses. DNA-fluorescence peak CVs were acceptable (< 15%;

Boucher et al., 1991; Veldhuis et al., 1997) for the samples, and substantial differences were detected between the two Pfiesteria species, both in mean relative DNA detected and in the population distribution of the DNA signals. Much of our DNA histogram

interpretation relied on previous research indicating that Pfiesteria spp. zoospores are haploid cells, reproducing primarily through vegetative cell division (Burkholder et al.,

2001a; Glasgow et al., 2001) as has been documented for the zoospores of many dinoflagellate species (e.g., Pfiester, 1984; Triemer and Fritz, 1984; Pfiester and Anderson,

1987; Graham and Wilcox, 2000). Each P. piscicida zoospore culture showed a cell cycle distribution with two peaks that may have corresponded to G1 and G2+M phases, typical of populations of eukaryotic, asynchronously cycling cells (Gray et al., 1987; Fig. 2.3A, B).

In contrast, each P. shumwayae culture demonstrated a single DNA peak distribution, indicative of synchronous cells (Wong and Whiteley, 1996; Bhaud et al., 2000; Fig. 2.3C,

D). This peak was assumed to have been composed of G1 cells, based on observations of the mode of vegetative division that was commonly employed by these P. shumwayae strains. All P. piscicida clonal cultures had significantly lower relative mean G1 zoospore

DNA (assumed as 1C; Wong and Whitely, 1996) than the P. shumwayae clonal cultures

(Table 2.1). Some intra-specific variability was also detected for both species. When the

P. piscicida cultures were evaluated for DNA content with considering culture age, oldest clones had significantly lower mean 1C zoospore DNA than newer clones (Fig. 2.4).

Zoospore DNA did not correlate with prey type for the Pfiesteria cultures examined here.

Cell sorting by flow cytometry for microscopical observations was used to further investigate vegetative reproduction in the two Pfiesteria species. Cells that comprised different DNA peaks were sorted and then examined using light and epifluorescence microscopy. When P. piscicida sample zoospores comprising the G2+M (2C) DNA- fluorescence peak were sorted and observed, often cells had nuclear characteristics indicative of a mitotic state (Fig. 2.5). No such nuclei were observed in cells comprising

Table 2.1. List of the clonal cultures examined, the original location of the isolates from which they were cloned, clone ages (= time since original isolation from an estuarine field sample), the peak mean whole-zoospore G1 DNA contents relative to the CRBC signal mean, and the coefficients of variation (CVs) for the G1 DNA-fluorescence peaks.

P. piscicida Mean G1 DNA Clone Age G1 Peak CV Clone Source Location (CRBC units (months)a (%) Designation zoospore-1)b 101685 Vandermere Creek, NC 50 2.95 9.1 113-3-0 Neuse River, NC 47 2.78 9.2 1037C Slocum Creek, NC 15 3.33 8.9 R-113-2-0 Flanners Beach, NC 47 2.49 9.9 125-4-0 Beaufort Point, NC 45 2.43 7.3 114-1-5-0 Neuse River, NC 46 2.22 9.9 R-113-4-0 Flanners Beach, NC 47 2.47 7.5 1038C Carolina Pines, NC 37 2.72 7.1 1049C Slocum Creek, NC 15 3.02 8.2 1032C Chesapeake Bay, MD 12 4.03 7.9 1031C Chesapeake Bay, MD 12 3.31 7.7 R-97-1-0 Chicamicomico River, MD 47 2.43 8.9 1033C Chesapeake Bay, MD 12 3.10 9.2 786PC2 Chesapeake Bay, MD 12 3.39 8.8

P. shumwayae Mean G1 DNA Clone Age G1 Peak CV Clone Source Location (CRBC units (months)a (%) Designation zoospore-1)b 101698 Neuse River, NC 36 6.31 10.1 101701 Slocum Creek, NC 15 6.62 10.3 101704 Chesapeake Bay, MD 12 7.70 12.4 1116C Arnell Creek, DE 8 6.78 9.9 101702 Indian River, DE 13 7.06 10.1

101692 Oslo, Norway 14 8.46 12.0 101653 Oslo, Norway 14 6.88 12.1 101691 Oslo, Norway 14 6.82 12.6 1090C Cawthorn, New Zealand 14 6.05 9.3

a Time, in months at the point of sample fixation, since the original field sampling event. b Fluorescence signals were collected for at least 104 zoospores in the G1 peak for each zoospore G1 DNA mean determination. CRBC unit = 2.33 pg DNA cell-1 (as in Vaulot et al., 1994; Veldhuis et al., 1997).

CRBC A CRBC C 1C 1C

uenc

Fre 2C

DNA

B D

Forward Scatter

DNA

Figure 2.3. Representative peak DNA distributions for (A, B) P. piscicida and (C, D) P. shumwayae clonal flagellated cells with CRBCs present as an internal standard. CRBC singlet and doublet peaks are the two leftmost DNA peaks in histograms (A) and (C). An apparent G1-S-G2+M DNA distribution (Gray et al., 1987) (1C and 2C) occurred in the P. piscicida cells (A), while only one DNA peak (1C) was found in the P. shumwayae cells

(C). Two-parameter dot plots (B, D) showed a similar DNA distribution, with forward scatter light used as a sizing parameter (see text). Histograms each represent analysis of >

20,000 flagellated cells.

the P. piscicida G1 peak, nor were apparent mitotic nuclei found in P. shumwayae zoospores comprising the single observed DNA peak for that species. Mitotic nuclei were observed in separate P. shumwayae cyst preparations. These observations support the hypothesis that, in these clones of P. shumwayae, flagellated zoospores were predominantly interphasic cells. Additionally, in 3 (out of 14) of the P. piscicida cultures examined, rarely

(in < 2% of the total cells measured) a DNA peak suggestive of a 4C DNA content was detected. However, visual inspection (after sorting) of these cells revealed typical zoospore cellular and nuclear morphology, and a non-mitotic nuclear condition. Unfortunately, the fixation method did not leave the zoospore flagella intact, and determinations other than

"indistinguishable from 4C DNA" (e.g., whether these were aberrant cells of different ploidy, or sexual products – although sexual reproduction for these species has been reported to date in the presence of fresh fish materials) were not possible. Similarly, in 3 of the 9 P. shumwayae cultures, < 2% of the total zoospores were indistinguishable from 2C.

As in the P. piscicida clones, no mitotic figures were observed in these rare P. shumwayae zoospores, and their role could not be discerned. Finally, no measurable differences were detected in forward scatter (a sizing parameter) between 1C and 2C subpopulations for P. piscicida flagellated cells (Fig. 2.3B), and P. shumwayae zoospores also exhibited a homogenous distribution about a mean with respect to forward scatter signals (Fig. 2.3D).

Thus, the 2C zoospores of P. piscicida were not detectably larger than the 1C zoospores, and the P. shumwayae zoospores also exhibited a unimodal distribution with respect to size.

5 )

-1

4 R2 = 0.68

Mean Zoospore 1C DNA ( CRBC units cell 1 0 102030405060

Time in Culture ( months )

Figure 2.4. Mean 1C DNA for these clones of P. piscicida zoospores, versus clone age

(time since field isolation, as in Table 2.1). The 1C DNA mean (G1 peak) was determined from analysis of > 20,000 zoospores. Note that clones of similar age for P. shumwayae were not available for analysis.

Figure 2.5. Representative sorted, SYTOX Green-stained, G2+M zoospores of P. piscicida, viewed under epifluorescence microscopy with mercury lamp excitation, including (A) G2 zoospores; and (B) an apparent mitotic cell, bar = 10 µm.

2.5 Discussion

This study suggested a possible difference in the vegetative reproduction of these clonal cultures of P. piscicida and P. shumwayae. Paired P. piscicida zoospores were abundant, suggesting completion of mitosis while motile was common, an occurrence that was not observed in the P. shumwayae cultures. These observations were supported by

DNA peak distributions in flow cytometry, and by visual examination of DNA-stained and sorted zoospores. Moreover, division cysts were more apparent in these clones in P. shumwayae than in P. piscicida. However, we note that in research with other P. piscicida cultures, similar vegetative cysts and other adherent benthic forms such as cysts in palmelloid masses have been described (Burkholder and Glasgow, 1997; Burkholder et al.,

2001a).

In one sample removed from a culture of P. shumwayae with live fish, we additionally documented germination of four zoospores – two from each of the two cleavage products that resulted from one pre-cleavage cyst. Thus, sequential mitotic divisions can occur while encysted. Alternatively, the excysted zoospores of P. shumwayae in this case may have represented meiotic products from sexual reproduction. A variation of this observation was reported from cultures of actively toxic P. piscicida, wherein a hypnozygote produced four zoospores (Burkholder and Glasgow 1997, Burkholder et al.

2001a).

Non-motile vegetative cysts or cells have been documented in the life cycles of other dinoflagellate species. For example, division cysts have been described for the estuarine heterotroph Katodinium (formerly Gymnodinium) fungiforme (Spero and Morée,

1981), the marine heterotroph Crypthecodinium cohnii (Bhaud et al., 1991), an isolated

endozoic dinoflagellate species (zooxanthella; Franker, 1971), the freshwater photosynthetic dinoflagellate Glenodiniopsis steinii (Lemmermann) Woloszyńska (formerly

Gloeodinium montanum) (Kelly and Pfiester, 1989), and the marine photosynthetic dinoflagellate, Amphidinium operculatum Claparède & Lachmann (formerly A. klebsii;

Barlow and Triemer, 1988; Daugbjerg et al. 2000). In descriptions of these and other dinoflagellate life cycles, similar coccoid division cells have been called zoosporangia

(Spero and Morée, 1981), digestion cysts (a type of cyst produced after feeding; Fensome et al., 1998), vegetative cysts (Pfiester and Anderson, 1987; Taylor, 1987), and parthenospores (cysts which mimic a zygotic state, but are not the result of sexual fusion;

Pfiester, 1984). Although dinoflagellate resting cysts have often been associated with sexual reproduction (Loeblich and Loeblich, 1984), not all resting cysts can correctly be assumed to be sexual products, regardless of the period of dormancy or the presence/absence of pigmented inclusions (Pfiester and Anderson, 1987; this work).

Division cysts may be associated with asexual reproduction (Franker, 1971; Spero and

Morée, 1981; Pfiester and Anderson, 1987; this study) or with sexual reproduction

(Pfiester, 1984; one observation in this work, described above).

It is not yet known whether the P. shumwayae zoospores examined here consisted purely of G1 cells, or whether some portion of DNA synthesis (S phase) also occurred while cells were motile, as has been suggested in Crypthecodinium cohnii (Bhaud et al.,

2000). A slight broadening or skewing of P. shumwayae zoospore DNA-fluorescence peak was observed in some samples, and higher CVs were measured for the DNA peak distribution for this species relative to the G1 DNA peaks for P. piscicida. The skewed

DNA fluorescence peaks could suggest partial inclusion of S phase in some P. shumwayae

zoospores, and the application of cell cycle histogram deconvolution software supported this premise for those cultures (data not shown). If so, the inclusion of some S phase cells could have led to overestimation of the mean cellular G1 DNA for those P. shumwayae populations. Random zoospore population S phase inclusion thus could also have elevated the intra-specific variability in mean 1C DNA content that was observed among the P. shumwayae clones.

A significant difference was measured in the mean relative cellular 1C DNA content between the two Pfiesteria species, for the clones investigated here. Some intraspecific variability was also detected, comparable to the variability that has been measured for other cultured marine protists, especially when isolates from different geographic regions were included in the analysis (Vaulot et al., 1994; Edvardsen and Vaulot, 1996; Green et al.,

1996). Within the P. piscicida cultures, oldest clones (years) had significantly lower mean

G1 DNA than more recent clones. This trend is distinct from that reported for some cultured photosynthetic dinoflagellates, based on microscopic chromosome enumeration

(Loper et al., 1980; Holt and Pfiester, 1982). P. shumwayae clones of similar longevity were not available for this study, so the extent to which this phenomenon may occur in that species is not yet known. Although additional cultures should be tested, the present data are of interest since toxin-producing capability of many toxic Pfiesteria clones has been lost over time in culture (PICWG 1999, Marshall et al. 2000, Burkholder et al. 2001a,b). We hypothesize that heterotrophic dinoflagellates, which likely require a more complex diet than obligate photosynthetic dinoflagellates, may more commonly lose than gain DNA when cultured for extended periods since the artificial media probably is missing required substrates. It remains to be determined whether the decrease in DNA content of cultures

maintained for years reflects loss of genes involved in toxin production, as has been found for certain toxic fungi (e.g., Cheng et al., 1999).

In subsequent research abundant division cysts were identified in several P. piscicida isolates. Flow cytometric analysis showed that the flagellated cells of these isolates had similar 1C DNA content and bimodal DNA distribution as the previously described P. piscicida clones (Table 2.1, Fig. 2.3A, B). Zoospores of these isolates may have fully replicated their DNA while motile (G + S) and persisted for some time in G2 phase, but then settled as cysts to undergo nuclear division and cytokinesis. Thus, cultured

P. piscicida zoospores might also show intermediate behavior between the general patterns of vegetative reproduction reported here for P. piscicida and P. shumwayae. Similar reproductive plasticity within a given species has also been described in other dinoflagellates (e.g., Faust, 1993).

The data and observations reported here, if conserved among other Pfiesteria zoospore populations, may provide insights about differences that have been observed between the two species in culture with algal prey. Nuclear DNA content has been shown to positively correlate with duration of cell cycle for many organisms (Van’t Hof, 1974;

Vinogradov, 1998), including dinoflagellates (Pan and Cembella, 1998). The higher DNA content measured for P. shumwayae might help to explain lower zoospore production rates relative to P. piscicida that have been observed in some cultures with algal prey

(representative zoospore specific growth rates (k, divisions day-1, mean ± SE) for P. piscicida = 0.75 ± 0.04; for P. shumwayae = 0.22 ± 0.07, in cultures with similar treatment history – Parrow et al., 2001). The common inclusion of a non-motile mitotic intermediate stage might also slow net zoospore population growth, relative to populations that

completed mitosis without such an intermediate. It is important to note, in this as for any research with clonal cultures, that natural populations of Pfiesteria spp. may behave differently than cultures maintained under the highly artificial conditions of this study; and that other clones may show different patterns of vegetative reproduction than observed here. Alternatively, Pfiesteria field populations may reproduce vegetatively in a manner consistent with these clonal cultures. If so, then differences in modes of vegetative reproduction between P. piscicida and P. shumwayae may significantly affect the comparative bloom dynamics of the two species.

Acknowledgments

Funding for this research was provided by an anonymous foundation, the North

Carolina General Assembly, the National Science Foundation, the U.S. Environmental

Protection Agency, and the Department of Botany at North Carolina State University. We thank all members of the Center for Applied Aquatic Ecology for their support, with special thanks to N. Deamer for her assistance. We also thank N. Allen, D. Kamykowski, and P.

Rublee for their counsel.

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3. VEGETATIVE AND SEXUAL REPRODUCTION IN PFIESTERIA SPP.

(DINOPHYCEA) CULTURED WITH ALGAL PREY, AND

INFERENCES FOR THEIR CLASSIFICATION

Matthew Parrow, JoAnn M. Burkholder, Nora J. Deamer, and Cheng Zhang

Harmful Algae 1 (2002) 5-33

3.1 Abstract

Algal-fed clonal zoospore cultures of Pfiesteria piscicida and P. shumwayae enabled description of certain conserved morphological and reproductive features.

Common modes of reproduction (especially via division cysts) were documented in herbivorous Pfiesteria piscicida and P. shumwayae using cultures fed algal prey, together with supporting photography and flow cytometric DNA measurements. Other cysts were characterized such as vacuolate cysts in starved P. piscicida cultures, and temporary cysts in both species fed algal prey. This study also represents the first report of sexual reproduction in Pfiesteria spp. cultures fed algal prey rather than live fish; the first report of a technique for cell cycle synchronization for these heterotrophic dinoflagellates; and the first information on storage products of cells released from Pfiesteria reproductive cysts.

Sexual reproduction in algal-fed P. piscicida clonal cultures was evidenced by fusing gametes, cells with two longitudinal flagella, and nuclear cyclosis. Both isogamous and anisogamous fusions were observed, and resulting cells with two trailing flagella (i.e. planozygotes and planomeiocytes) sometimes comprised > 50% of the flagellated cells.

These cells continued feeding activity, and eventually (hours) lost their flagella and formed cysts. Nuclear cyclosis and a subsequent cell division were observed in some thin-walled reproductive cysts prior to release of two flagellated cells. One gamete fusion event was also documented in 1 of 20 algal-fed clones of P. shumwayae, with non-motile zygote as the product. We obtained high cell synchrony (> 90% 1C) in the tested cultures using our preferential lysis technique, and tracked the decline in lipid content of excysted zoospore populations over time. The data from this study were considered together with previous research to gain insights about relationships between Pfiesteria spp. and other heterotrophic

dinoflagellates. Pfiesteria spp. should be regarded as free-living predators rather than parasites because they are prey generalists without demonstrated “host” specificity, and their flagellated feeding stages are not morphologically distinct from swimming stages.

Although they originally were placed within the Dinamoebales because amoebae can predominate, this study as well as other published research consistently has shown that the dominant stage varies depending on culture conditions, prey type/availability, and strains.

The peridinoid plate structure of each Pfiesteria species, which thus far has been conserved across culture conditions and strains, supports placement of Pfiesteria spp. within the

Peridiniales. At the species level, plate structure (differing by only 1 precingular plate) and molecular data (18S rDNA) indicate that the two Pfiesteria spp. are closely related in comparison to species grouped within other genera.

3.2 Introduction

The heterotrophic dinoflagellates Pfiesteria piscicida Steidinger et Burkholder

(Steidinger et al., 1996a) and Pfiesteria shumwayae Glasgow et Burkholder (Glasgow et al.,

2001b) are considered to be harmful algae because their toxic strains negatively impact fish and mammals (Glasgow et al., 1995; Burkholder and Glasgow, 1997a; Levin et al., 1997,

1999; Grattan et al., 1998; Burkholder et al., 2001a,b; Kim-Brinson et al., 2001). Toxic strains are cosmopolitan in distribution (Glasgow et al., 2001a; Jacobsen et al., 2002), and they have been implicated as the primary causative agents of certain major fish kills in estuaries of the mid-Atlantic U.S. (Burkholder et al., 1995a; Glasgow et al., 1995;

Burkholder et al., 2001a; Glasgow et al., 2001a). Like various other heterotrophic protists

(but unlike most other currently recognized harmful algal species), Pfiesteria spp. flagellated cells chemically detect and rapidly gather around particulate food sources in active attack behavior (Burkholder and Glasgow, 1997a; Fenchel and Blackburn, 1999;

Cancellieri et al., 2001). The resulting dense, dynamic aggregations are capable of killing and consuming prey many times larger than an individual zoospore, including various algae, ciliates, nematodes, arthropods, mollusks, and fish (Spero and Morée, 1981;

Burkholder and Glasgow, 1995,1997a; Burkholder et al., 2001a,b; Glasgow et al., 2001b).

Pfiesteria spp. also have active as well as resting benthic stages in their life cycles

(Burkholder and Glasgow, 1995, 1997a; Burkholder et al., 2001b), and the benthic subpopulations are believed to provide a recurrent “seed” population for inoculating blooms of zoospores. Encysted and other benthic stages likely play a critical role in the distribution, abundance, and basic ecology of Pfiesteria spp. (Anderson and Wall, 1978; Anderson et al.,

1995; Burkholder and Glasgow, 1995, 1997a; Burkholder et al., 2001a,b; Parrow and

Burkholder, 2002).

The primary mechanisms by which Pfiesteria cells proliferate are of fundamental relevance to zoospore occurrence in nature. Study of cell production in any microbial species requires that the species be cultured in the laboratory, under the assumption that behaviors exhibited in culture approximate those that occur in vivo. As obligate heterotrophs (although they sometimes maintain kleptochloroplasts to augment nutrition),

Pfiesteria spp. presently must be cultured with algae, fish, or other prey to achieve ecologically relevant cell production over extended periods (days to weeks; Burkholder and

Glasgow, 1995, 1997b; Lewitus et al., 1999; Burkholder et al., 2001a,b; Glasgow et al.,

2001b). Complex trophic (prey type and availability) and environmental factors can profoundly affect stage transformations and cell production in the life cycle of Pfiesteria spp. both in vitro and in vivo (Burkholder et al., 1995b; Burkholder and Glasgow, 1995,

1997a,b; Burkholder et al., 2001a,b; Glasgow et al., 2001b).

Aspects of the range of stages and behaviors within the life cycles of Pfiesteria spp. have been described previously, primarily for actively toxic strains fed live fish prey

(Burkholder and Glasgow, 1995, 1997a; Burkholder et al., 2001a,b; Glasgow et al., 2001b).

Notable differences between the two species in typical flagellated cell population DNA distributions also have been reported (Parrow and Burkholder, 2002), and certain other characteristics of vegetative reproduction in Pfiesteria spp. have been described

(Burkholder et al., 2001b; Parrow and Burkholder, 2002). The objectives of the present work were to describe and photo-document common modes of Pfiesteria spp. reproduction in clonal cultures fed algal prey; to present supporting flow cytometric DNA measurements

in further description of asexual and sexual reproduction in the two species; to develop an efficient cell cycle synchronization method for use with algal-fed Pfiesteria zoospore cultures; and to use flow cytometry to assess population DNA distribution and the major storage product of synchronized flagellated cells released from reproductive cysts. The data from this study on modes of reproduction in Pfiesteria zoospore populations fed algal prey were also considered together with supporting information from previous research, to gain insights about relationships between Pfiesteria spp. and other heterotrophic dinoflagellates.

3.3 Materials and Methods

3.3.1 Algal-fed Cultures. Most previous characterizations of Pfiesteria spp. cell morphologies and life cycles have been conducted using populations cultured with live fish as the primary prey source (Burkholder et al., 1995b, 2001a,b; Steidinger et al., 1995,

1996a; Burkholder and Glasgow, 1995, 1997a,b; Marshall et al., 2000; Glasgow et al.,

2001b). Fish-fed cultures have certain advantages over algal-fed cultures, such as focus on actively toxic Pfiesteria stages (since toxin production appears to require the presence of live fish), increased ecological relevance, and elicitation of a wider range of behaviors

(Burkholder and Glasgow, 1995, 1997a,b; Burkholder et al., 2001a,b; Glasgow et al.,

2001b). However, the addition of fish necessarily brings with it an assemblage of other microorganisms that can sometimes confound interpretation (Burkholder et al., 2001a,c).

Moreover, fish cultures are not as readily amenable to in situ observations, particularly of certain benthic cell types (Burkholder and Glasgow, 1995).

Although unialgal-fed Pfiesteria spp. are grown in a highly artificial environment, they are more readily amenable to microscopic evaluation, and the data can be reproduced by researchers who lack biohazard III facilities required to culture actively toxic Pfiesteria spp. with live fish (Burkholder et al., 2001c). For this study, the need for controlled, contaminant-free cultures with benthic encysted and planktonic stages that could be readily observed under microscopy outweighed the increased ecological relevance possible in fish- fed, actively toxic cultures. Although caution is warranted in over-extrapolation of the importance of research conducted using Pfiesteria spp. cultures which have been maintained solely on a diet of algal prey for extended periods (weeks to months), those concerns extend primarily to issues of toxicity and nutrient pollution (Burkholder et al.,

2001a,b). The relevance of the findings reported here rests on the likely assumption that the core (common) reproductive processes exhibited by Pfiesteria spp. are largely conserved, regardless of prey type availability and toxicity status.

3.3.2 Pfiesteria Zoospore Clones. Clonal cultures of the two Pfiesteria spp. (20 for each species; each clone was initiated from a single zoospore; definition following the

Pfiesteria Interagency Coordinative Working Group [PICWG], 2001 in routine maintenance at the Center for Applied Aquatic Ecology [CAAE]) were obtained by micropipette isolation of single zoospores or in the past 2.5 yr by flow cytometric cell sorting (EPICS Altra flow cytometer/particle sorter with robotic Autoclone sorting option,

Coulter Corporation, Miami, Florida, U.S.A.; Burkholder et al., 2001a; Glasgow et al.,

2001b). Species identifications of algal-fed Pfiesteria spp. zoospores were determined by scanning electron microscopy (SEM) of suture-swollen cells (Burkholder et al., 2001a;

Glasgow et al., 2001b) and polymerase chain reaction (PCR) molecular probes (Rublee et al., 1999; Bowers et al., 2000; Oldach et al. 2000). They were cross-corroborated by Dr. H.

Marshall, Old Dominion University, Norfolk, Virginia, U.S.A. (SEM); Dr. P. Rublee,

University of North Carolina-Greensboro, Greensboro, North Carolina, U.S.A. (PCR), and/or Dr. D. Oldach, University of Maryland, Baltimore, Maryland, U.S.A. (PCR). The cultures had been isolated from estuaries of the mid-Atlantic U.S. (in Delaware, Maryland,

North Carolina, and South Carolina). Culture “age” (time since initial field isolation) ranged from 6 months to 4 yr. Forty clonal cultures (20 each of P. piscicida and P. shumwayae; potentially toxic and non-inducible strains as in Burkholder et al. 2001b) were maintained in polystyrene cell culture flasks, stored horizontally at ~20oC under ambient

overhead fluorescent lighting (50 µmol photons m-2 s-1). Replicate cultures were maintained similarly in 4.5-mL chamber microscope slides for imaging.

Zoospore cultures were fed whenever algal prey approached depletion, as in the previous > 3 months, by decanting off about two-thirds of the culture and refilling the flask with an inoculum of logarithmic-growth phase algal prey consisting of either Rhodomonas sp. CCMP757, Dunaliella sp. CCMP1320 (cloned from a commercial culture; Culture

Collection for Marine Phytoplankton, Bigelow Laboratory for Ocean Science, Bigelow,

Maine, U.S.A.), or Cryptomonas sp. HP9101 (from Dr. D. Stoecker, Horn Point

Laboratory, Cambridge, Maryland, U.S.A.). The algal prey had been grown in f/2 media

(Guillard, 1975) prepared with aged, sterile-filtered natural seawater (Gulf Stream, ca. 40 km offshore from Cape Hatteras, North Carolina, U.S.A.) at a salinity of 15 (Burkholder and Glasgow, 1995; Lewitus et al., 1999; Marshall et al., 2000; PICWG, 2001; Burkholder et al., 2001b; Parrow et al., 2001). These taxa are preferred algal prey types for Pfiesteria spp. (Burkholder and Glasgow, 1995, 1997a; Glasgow et al., 2001b). For some experiments, P. piscicida was starved by not replenishing prey for 2-3 weeks (note, in contrast, that P. shumwayae did not become prey-depleted because its grazing rates apparently were below the prey growth rates).

Upon screening of algal-fed, clonal P. piscicida zoospore cultures used in this study, we noted couplings of cells in an orientation suggestive of gamete fusion rather than vegetative cell division and cells with two trailing flagella (referred to as longitudinally biflagellate), a condition indicative of a zygotic state in dinoflagellates (i.e., planozygotes, and planomeiocytes or premeiotic excystment cells; von Stosch, 1973; Beam and Himes,

1984; Walker, 1984; Popovský and Pfiester, 1990). Sexual reproduction is common in

Pfiesteria spp. populations given live fish in standardized fish bioassays, and is believed to be stimulated by rich organic materials excreted or secreted by the prey (Burkholder and

Glasgow 1995, 1997a). In the event that the couplings of cells we had observed in the algal-fed zoospore cultures were initiating the sexual cycle, we examined clonal cultures of both P. piscicida and P. shumwayae for gamete fusion, motile zygotes, zygotic cysts, and nuclear cyclosis. For some cultures, sub-cultures of the same clone fed live fish (in biohazard III facilities under growth conditions described in Burkholder et al., 2001c) were available for comparison. The CAAE also has various non-clonal cultures identified as uni- species isolates of P. piscicida or P. shumwayae, enabling comparisons of zoospore reproduction in clonal and non-clonal cultures.

3.3.3 Sample Preparation. For microscopy, cultures were either observed live or after fixation in 0.5% (final concentration) microscopy-grade glutaraldehyde. Glycerol

(Anderson et al., 1995) or methyl cellulose (von Stosch, 1973) was added to selected subsamples to slow cell motility. For qualitative observation and imaging of nuclear condition, live subsamples were incubated for 30 min in darkness in 5 µM SYTO 11 (final concentration; Molecular Probes, Inc., Eugene, Oregon, U.S.A.). Fixed subsamples were stained in darkness for >2 h in 5 µM SYTOX Green (final concentration) (Molecular

Probes, Inc). To permeabilize the resistant wall for nuclear staining of encysted stages, subsamples were mixed at 50% v/v with a permeabilization buffer consisting of 755 mM

NaCl, 5mM EDTA, 0.01% SDS, 0.1% CTAB, 70 mM sodium citrate, 0.1% Triton X-100

(final concentration; Sigma, St. Louis, Missouri, U.S.A.) in deionized water (modified from

Adachi et al., 1996; Kempton, 1999), heated for 1 h at 60ºC, mixed and stained similarly as

other fixed samples. For scanning electron microscopy, samples were prepared following the procedures described by Burkholder and Glasgow (1995) and Glasgow et al. (2001b).

Samples were processed for flow cytometric DNA analysis as in Parrow and

Burkholder (2002). Briefly, 50 mL of culture were collected and filtered (25-µm porosity;

Fisher Scientific) to remove particulate debris and fixed with 0.5% paraformaldehyde (final concentration). The preserved samples were stored for > 24 h in darkness at 4˚C. Cells were then concentrated ten-fold by centrifugation and returned to storage conditions. Prior to staining, 1-ml aliquots of the stock cell suspensions were removed and treated with

RNAse A (1 µg ml-1 final solution as weight to volume [w/v]; Sigma) for 1 h at room temperature. Cell suspensions were stained with 5 µM SYTOX Green and returned to storage conditions for 12 –14 h. Similarly stained chicken red blood cells (CRBCs;

BioSure, Grass Valley, California, U.S.A.) and/or Immuno-Brite fluorospheres (Coulter

Corporation, Miami, Florida, U.S.A.) were used as fluorescence standards. Preparations selected for lipid staining were fixed in paraformaldehyde as above and stained with 1 µg mL-1 (final concentration, v/v) Nile Red (Molecular Probes, Inc.) from a 250 µg mL-1 stock solution prepared in acetone (Cooksey et al., 1987) for 30 min in darkness prior to microscopic or flow cytometric analyses.

3.3.4 Light Microscopy. In situ routine culture examination was performed daily using an Olympus CK40 inverted culture microscope (Olympus Corporation, Melville,

New York, U.S.A.) at 40-400x. Critical observations, imaging, and video microscopy were performed in optical glass-bottomed containers using an Olympus AX70 research microscope equipped with a water-immersion 60x objective. Mercury lamp excitation and a triple-band pass filter block were employed for simultaneous epifluorescence imaging of

lipid-stain and DNA-stain complexes (red and green emission, respectively; Parrow and

Burkholder, 2002). Image data were captured with a DEI-750 cooled-chip CCD camera

(Optronics Engineering, Goleta, California, U.S.A.) and/or video recorded using a Sony

SVD-9500MD S-video recorder.

3.3.5 Flow Cytometry. Analyses and cell sorting were performed with the flow cytometer/cell sorter using 150 mW of the 488 nm laser line focused to an elliptical point of interrogation 6 µm height x 112 µm wide; cells traversed the laser beam in a 100 µm channel quartz flow cell. Instrument optics were selected to detect fluorescence from stain complexes on photomultipliers screened with appropriate band-pass filters; 525 nm,

SYTOX Green-DNA (Parrow and Burkholder, 2002); 575 nm, Nile Red-lipid (Brzezinski et al., 1993; Reed et al., 1999). Forward-angle light scatter and side-angle light scatter were recorded as relative measurements of cell size/granularity. Signal amplification and threshold levels were optimized so that signals fell mid-scale. Integrated, peak and logarithmic fluorescence and scatter signals were recorded as appropriate on 104 to 105 cells per analysis, displayed using EXPO32 Analysis software (Applied Cytometry Systems,

Sheffield, UK), and stored in listmode format. Unstained samples were used as negative controls. In some cases CRBCs were used as internal DNA standards, and/or fluorospheres as fluorescence reference standards. Optical alignment and signal stability were monitored using 10 µm diameter fluorescent latex microspheres (Coulter Corporation), and linearity of the cytometer amplification system was checked using Immuno-Brite fluorospheres

(Bagwell et al., 1989). Frequency-modulated electrostatic cell sorting was performed in standard sorting mode 3 (for purity in preference of yield), and electronic sort regions were defined on the EXPO32 interface to encompass any detectable fluorescence-DNA

distribution subpopulation of interest. Sorted cells were collected on microscope slides, sealed under a coverslip ridged with silicone grease, and microscopically examined with a

20x high-dry objective or a 60x water objective.

The flow cytometric DNA analyses focused on motile cells and did not include encysted stages, because all known Pfiesteria spp. cyst types are impervious to the DNA staining methods optimized here for flagellated stages. In addition, the adherent and clumping nature of the cysts makes them ill-suited as subjects for flow cytometry (Parrow and Burkholder, 2002). The application of stringent cell permeabilization steps allowed stain penetration of some, but not all, cysts. Therefore, investigation of cyst DNA content was limited to qualitative observations (e.g., number of nuclei and visual estimation of nuclear size).

3.3.6 Cell Cycle Synchronization. A modification of the preferential lysis technique of Franker (1971) was used to synchronize Pfiesteria spp. cell cycles. Selected clonal cultures of P. piscicida (n = 3) were serially inoculated with a small volume (< 10 mL) of centrifuged algal prey concentrate whenever prey approached depletion. The resulting dense flagellate cultures produced abundant benthic cysts. P. shumwayae clonal cultures (n

= 3) were selected once cysts became relatively abundant, which required > 1 month of culture maintenance under the conditions used here. For each species, the culture media was decanted and the flask was refilled with ultra-distilled water (Millipore Corporation,

Bedford, Massachusetts, U.S.A.), maintained for > 2 h to ensure that remaining flagellated cells had lysed. The water was removed and sterile-filtered seawater was added (0.22 µm; salinity 15). Thus, flagellated cells excysted into media devoid of prey. After < 24 h the excysted, motile cell population was isolated from remaining cysts. Sub-samples of the

synchronized flagellate population were removed at 4-hr intervals over 48 hr, and were fixed for analysis by light microscopy and flow cytometry.

3.3.7 Data Analyses. Listmode data were analyzed using EXPO32 Analysis software and plotted as univariate signal height distributions or bivariate frequency distributions. Depending upon the analysis, peak numbers (area), means, and coefficients of variation (CVs) were computed directly by defining regions on the plotted cytograms.

Potential doublets in DNA stained cells were discerned by plotting DNA fluorescence signal peak height versus integral signal area (Wersto et al., 2001). DNA-fluorescence intensity histogram deconvolution to yield the proportions of cells occupying G1, S, and G2 phases was performed using MultiCycle DNA Content and Cell Cycle Analysis Software

(Phoenix Flow Systems, Inc., San Diego, California, U.S.A.). One-way ANOVA was used to test for differences among treatment (sample) means, with significance at p < 0.05 (SAS

Institute, Inc., 1997).

3.4 Results

3.4.1 Zoospore Production. Zoospores of both species exhibited considerable plasticity in length (ca. 7 –17 µm) and volume (largest cells representing a ca. 10-fold increase in volume), with one to three prey-replete food vacuoles per zoospore. These food vacuoles sometimes extended from the epitheca into the hypotheca. In Pfiesteria spp., flagellated cell size is primarily related to the amount of ingested prey material, not nuclear content (Burkholder et al., 2001b; Parrow and Burkholder, 2002).

The tested strains of P. piscicida and P. shumwayae zoospores fed on all algal prey types via phagocytosis (as myzocytosis; Gaines and Elbrächter, 1987). However, the two

Pfiesteria spp. differed in both grazing rate on algal prey cells (inferred from prey disappearance from the medium), and zoospore production. The P. piscicida cultures consistently grazed algal prey to negligible densities within 2-4 days, and demonstrated robust cell production in a relatively short time (ca. 105 zoospores mL-1 within 2-4 days).

Dunalliela sp. prey typically persisted longer than the cryptomonads in P. piscicida cultures. P. shumwayae, in contrast, did not graze available prey (regardless of type) to negligible concentrations during a 2-month period, and reached maximal densities of ca. 2 x

104 zoospores mL-1. These observations are consistent with the lower P. shumwayae zoospore production rates on algal prey in comparison to P. piscicida that were presented in

Parrow et al. (2001), as discussed in Parrow and Burkholder (2002).

3.4.2 Zoospore Reproduction. 3.4.2a P. piscicida. The zoospores of these P. piscicida strains underwent cytokinesis while in a coccoid, nonmotile state (called ‘division cysts’ by Franker, 1971; von Stosch, 1973; Spero and Morée, 1981; Popovský and Pfiester,

1990; Beam and Himes, 1984; Faust, 1993; called zoosporangia by von Stosch, 1973;

Drebes and Schnepf, 1988; Popovský and Pfiester, 1990); and referred to here as division cysts). Attached pairs of motile zoospore were also observed in an oblique alignment with parallel cingula (Figure 3.1a), considered to be a common cytokinetic orientation for many gymnodinoid and armored dinoflagellate species (von Stosch, 1973; Walker and Steidinger,

1979; Walker, 1984; Pfiester and Anderson, 1987). The P. piscicida cell pairs were highly motile and, therefore, difficult to follow for extended periods (hours).

We observed cytokinesis in division cysts in all P. piscicida cultures examined.

Commonly in formation of these cells (Figure 3.1b), apparent 2C (G2 cell-cycle phase zoospores, as indicated from qualitative observation of nuclear size; Parrow and

Burkholder, 2002) often appeared to begin the process of nuclear division as they lost their flagella, became coccoid, and settled to the bottom of the culture vessel. Alternatively, karyokinesis apparently occurred after the cells had completely encysted. At some point early in the encystment process, the cells became impermeable to passive nuclear stains.

They initially appeared as thin-walled, round to oval cysts (Figure 3.1c), and often developed a surface notch marking the plane of internal protoplast division (Figure 3.1d).

Nuclear staining indicated that each offspring cell contained one nucleus, similar in size to that occurring in 1C zoospores (Figure 3.1d). Excystment of the two flagellated offspring cells was typically synchronous, and the emerging zoospore pair were often temporarily conjoined by a lateral connection (Figure 3.1e). Offspring zoospores extruded amorphously through a fissure in the transparent cyst wall, to which the pair often remained attached by their longitudinal flagella for several seconds prior to breaking free. Excystment occurred in less than 6 h for the majority of cysts observed. Some primary reproductive cysts underwent a complete cyst division, resulting in two secondary cysts, each with a newly

formed cyst wall, which remained internal to the primary cyst wall unless it dissolved (i.e., eleutheroschisis, Pfiester and Anderson, 1987; Figure 3.1f). Upon excystment, secondary cysts yielded two zoospores each. The lateral connection of excysted offspring pairs was typically short lived (seconds to minutes). During logarithmic growth reproductive cysts covered the bottom of culture flasks. We additionally observed similar division cysts in representative non-clonal, algal- and fish-fed cultures that tested as P. piscicida using PCR molecular probes (Rublee et al., 1999; Bowers et al., 2000; Oldach et al., 2000).

3.4.2b P. shumwayae. Cell division was not observed in zoospores of these P. shumwayae strains while motile, nor were zoospore pairs suggestive of binary fission observed (except for temporarily connected excystment cells, below). Zoospore reproduction was observed to occur via spherical to oval division cysts as reported earlier by Parrow and Burkholder (2002). Abundant cysts of similar morphology were produced in all P. shumwayae cultures, initially appearing as round, relatively smooth, transparent or occlusive walled bodies (ca. 10-20 µm) that tended to adhere to debris or flask edges

(Figure 3.1g). The protoplast of each cyst often completely divided within the primary cyst wall to produce two secondary cysts (Figure 3.1g,h), which remained inside the primary cyst wall unless it dissolved. As in P. piscicida, excystment occurred from primary cysts or secondary cysts via a fissure in the cyst wall through which the excystment protoplasts extruded amorphously. The excystment products from primary cysts (those that did not undergo cyst division to form secondary cysts) were 2 zoospores, which were often conjoined by a temporary lateral connection (Figure 3.1i-l). Secondary cysts yielded two zoospores each, as in the primary cysts. In each case a transparent cyst husk was left behind. As in P. piscicida, the conjoinment of excysted offspring zoospores was short lived

Figure 3.1

Figure 3.1. Light micrographs of Pfiesteria spp. fed algal prey: (a) P. piscicida zoospore pair with an oblique orientation suggestive of binary cell division while motile. (b) P. piscicida with stained DNA, including a zoospore (upper cell) and an encysted cell with a dividing nucleus (lower cell). (c) P. piscicida thin-walled reproductive cyst containing a divided nucleus, a single replete food vacuole, and abundant cytoplasmic lipid droplets. (d)

P. piscicida reproductive cyst in late division, with two stained nuclei bisected by the division plane. (e) Recently excysted pair of temporarily coupled P. piscicida offspring zoospores with visible transverse and longitudinal flagella. The degree of temporary connection between excysted offspring pairs varied; this pair represents the maximum observed. (f) P. piscicida secondary cysts with stained DNA, each cyst is uninucleate and temporarily retained in the primary cyst wall. (g) P. shumwayae reproductive cysts with thin walls and abundant cytoplasmic lipid droplets; the upper left cyst had divided to produce two secondary cysts. (h) P. shumwayae reproductive cysts; the right cyst was early in division with a visible cleavage furrow, whereas the left cyst had completely divided into two secondary cysts that were still enclosed in the hyaline primary cyst wall. (i-l) P. shumwayae, series of observations including: (i) Primary cyst (uppermost cell) and secondary cysts (two lower cells). (j) Primary cyst beginning excystment; note that the transparent cyst wall had bulged outward, apparently from pressure exerted by the internal cells. (k) Offspring zoospore pair extruding amorphously through a fissure in the transparent cyst wall. (l) Excystment completed, showing two temporarily conjoined offspring zoospores that had been produced by the primary cyst, and the discarded transparent cyst wall (arrow). The pigmented inclusions (prey-replete food vacuoles) were unequally distributed between offspring cells. Scale bars = 10 µm.

(seconds to minutes). Thus, the final number of germination products produced from a single P. shumwayae primary cyst was either 2 or 4 zoospores. Cysts that did not yield flagellated products tended to darken to a reddish-brown color over time (months).

Although in this study we observed a maximum of 4 offspring flagellated cells from 1 encysted progenitor cell, in this and other research with cultures of each Pfiesteria sp. we have found reproductive cyst arrangements which suggested that secondary cysts underwent an additional division, producing 4 tertiary cysts (Figure 3.2a). If each of these presumed tertiary cysts produced 2 flagellated excystment cells, a maximum of 8 offspring cells from 1 encysted progenitor is feasible.

P. shumwayae cysts became increasingly apparent as zoospore densities increased.

In some older, relatively dense cultures, cysts represented > 50% of the total cells. The cysts tend to occur in clumps attached to irregular surfaces. In observations of the well- plate microcultures used in our cloning procedures, many culture wells that initially appeared devoid of cysts, but more careful examination revealed a dense pile of hundreds of cysts in one discrete location on the bottom surface. We additionally observed similar division cysts in representative non-clonal, algal- and fish-fed cultures that tested as P. shumwayae using PCR molecular probes (Rublee et al., 1999; Bowers et al., 2000; Oldach et al., 2000).

3.4.3 Other Cysts. Zoospores of both species produced morphologically similar temporary cysts. These cysts did not divide and were not considered reproductive, but presumably formed as a rapid response to unfavorable stress (Burkholder et al., 2001a).

Pfiesteria spp. temporary cysts were not ecdysal (Taylor, 1987) but, rather, formed when zoospores lost their flagella, secreted a thin continuous protective covering, and settled to

the bottom of the culture vessel (Figure 3.2b). The stimulus for this response was unknown. The thin covering of temporary cysts in these algal-fed cultures differed from the thick mucilaginous covering that previously was observed during temporary cyst formation by zoospores in fish-killing cultures (Burkholder and Glasgow 1997a, Burkholder et al.

2001b). These temporary cysts apparently were quiescent until favorable environmental conditions were detected (as in other dinoflagellate species described, for example, by

Burkholder, 1992; Anderson et al., 1995). Excystment began with the protoplast extruding through a fissure in the transparent cyst wall (Figure 3.2c). Emergent protoplasts rapidly acquired a zoospore profile and, as with other Pfiesteria cyst types, the outer translucent protective cyst covering was left behind (Figure 3.2d).

We documented another type of cyst, here called vacuolate cysts as the first known report, in starved P. piscicida cultures. Formation was preceded by a swelling of the zoospore epitheca, resulting in rounded, vacuolate cells that swam in tight circles (Figure

3.2e). The cells lost their flagella and settled to the bottom of the culture vessel to form spherical translucent cysts with the nucleus closely appressed to the cyst wall (Figure 3.2f).

Condensed dinokaryotic chromosomes were clearly visible throughout this process. The formation of similar translucent, vacuolate cysts is described occurring in food-depleted cultures of the estuarine heterotroph Katodinium fungiforme Anissimova (Spero and Morée,

1981). In long-term food-depleted P. piscicida cultures, these cysts were thin-walled and often difficult to discern because they were translucent. Excystment from a vacuolated cyst was observed to yield a single translucent zoospore, and an actively feeding zoospore population developed < 24 h after adding algal prey to a bed of these cysts which had been

stored for 9 months without prey. The cysts were assumed to have been quiescent and similar in function to temporary cysts.

3.4.4 Sexual Reproduction. We documented sexual reproduction, as the first known report, in clonal Pfiesteria spp. cultured on algal prey rather than live fish. Sexuality was common in the clonal P. piscicida cultures examined and appeared to be enhanced by the addition of a dense inoculum of senescent algal prey.

3.4.4a Pfiesteria piscicida. Sexual reproduction in algal-fed P. piscicida cultures was indicated by fusing gametes, planozygotes, and cysts with nuclear cyclosis (von

Stosch, 1973; Coats et al., 1984; Beam and Himes, 1984; Pfiester, 1989). Sexuality was initiated when two gametes swam together on the same tangent, touching intermittently

(Figure 3.2g). Once gamete fusion began, the swimming behavior of the mating pair became more erratic and often degenerated into a repetitive looping pattern. Isogamous pairs (equal in size) were most commonly observed, but anisogamy sometimes also occurred in the same clone. Although presumed gametes could be distinguished from zoospores based on their smaller size and lack of pigmentation (as in Burkholder and

Glasgow, 1995, 1997b), hologamous fusions between flagellate pairs containing prey- replete food vacuoles also occurred. Therefore, zoospores could only be classified unequivocally as gametes once fusion began (i.e., hologamy; Pfiester and Anderson, 1987).

Initial gamete fusion orientations varied. Early fusing cells were often oriented with sulci touching, but asymmetric orientations were also observed (Figure 3.2h), including orientations where the epitheca of one gamete was adjacent to initiated fusion with the hypotheca of the other. In some observations, completion of syngamy occurred rapidly (< 1 h), whereas in others it appeared to require several hours. The time required for completion

of plasmogamy may have been related to the initial fusion orientation, and misaligned gamete pairs may have become reoriented later in fusion (suggested for other dinoflagellate species by von Stosch, 1973; Coats et al., 1984). Karyogamy appeared to precede the completion of plasmogamy (Figure 3.2i-l).

After completion of syngamy, the small planozygotes in these algal-fed cultures appeared similar to P. piscicida zoospores in morphology and motility, except for the presence of two longitudinal flagella. In these cultures, cells with two trailing flagella ranged from 7 to17 µm in diameter (n = 50), depending on the cell size of the initiating gamete pairs and the amount of ingested material. This size range is within the published size range for P. piscicida zoospores (Steidinger et al., 1996a; Burkholder et al., 2001a), but less than previously observed sizes for P. piscicida planozygotes (from fish-killing cultures;

Burkholder et al., 2001a), and considerably smaller than the planozygote dimensions (25-60

µm) given in the naming paper for the species that was based on isolates cultured with fish prey (Steidinger et al., 1996a). Newly formed planozygotes were small (length 7-10 µm;

Figure 3.3a-c). They engaged in phagocytosis and grew in size prior to encysting (Figure

3.3d).

Longitudinally biflagellate cells were not observed to undergo cell division while motile in these cultures. In rare instances, we observed a form of extreme anisogamy wherein a cell with two trailing flagella engaged in apparent sexual fusion with a gamete

(3C product; also reported by Beam and Himes, 1984 for Crypthecodinium cohnii

Biecheler, with fate of the polyploid zygotes unknown) or with another longitudinally biflagellate cell (4C product). We were unable to determine the developmental fate of these

Figure 3.2

Figure 3.2. Light micrographs of Pfiesteria spp. fed algal prey: (a) P. shumwayae reproductive cyst cluster; two secondary cysts apparently had divided, producing four tertiary cysts (note that in dinoflagellates, the genetic term ‘tetrad’ can be appropriately applied to such division products only if the progenitor cell was zygotic, in which case tetrad would describe the products of meiosis (2, 4, or 8 in number; Beam and Himes,

1984). (b-d) P. piscicida, series of observations including: (b) Thin-walled temporary (non- reproductive) cyst (the stimuli for temporary encystment and excystment were unknown).

(c) Excystment from the temporary cyst, showing a single cell extruding through a fissure in the transparent cyst wall. (d) Excysted zoospore, which remained attached momentarily by its longitudinal flagella to the discarded hyaline temporary cyst wall (arrow). (e-l) P. piscicida, including: (e) Swollen, highly vacuolate zoospore (flagella and nucleus visible) prior to formation a vacuolate cyst; note the ‘signet ring’ profile, similar in appearance to the mature trophozoite stage of Perkinsus sp. (Coss et al., 2001). (f) Resultant thin-walled, vacuolate cyst with dinokaryotic chromosomes visible in the appressed nucleus. (g) Pairing isogametes (gametes were colorless; the green coloration was an artifact of the SYTO 11

DNA stain). (h) Asymmetrically fusing gametes (center), with peripheral Dunaliella sp. prey. (i) Fusing gamete pair with distinct nuclei (stained). (j) Gamete pair in early nuclear fusion. (k) Gamete pair in late nuclear fusion (plasmogamy not yet complete). (l)

Presumed zygote with a single, stained diploid 2C nucleus in the hypotheca. Scale bars =

10 µm.

presumed, aberrant 3C or 4C P. piscicida cells because of their speed and extended motility but some may have maintained, at least for some time, 3 longitudinal flagella (Figure 3.3e).

Cells with two trailing flagella became increasingly common as cultures aged, and in some cultures comprised >50% of the P. piscicida flagellated cells present based on qualitative observations of live samples. In fixed preparations, glutaraldehyde was not reliable in preserving flagellae intact. Also, accurate determination of flagellar complement relied heavily on cell orientation; thus we could not consistently enumerate the trailing flagella of fixed cells. In live material we observed cells with two trailing flagella cease motility and rapidly encyst via the apparent secretion of a smooth wall. Some cysts

(presumed hypnozygotes, described for other dinoflagellate species by Pfiester, 1984;

Pfiester and Anderson, 1987) could be differentiated from asexual cysts under light microscopy by a more thickened wall (Figure 3.3f), a feature considered characteristic of zygotic cysts in other dinoflagellate species (Popovský and Pfiester, 1990; Anderson et al.,

1995). As in P. shumwayae cultures, many cysts darkened to a reddish brown color with age (weeks). These cysts may have been dormant (Burkholder and Glasgow, 1995; 1997a), but long-term viability was not assessed in this study. In these cultures, however, not all zygotic cysts were thick-walled or dormant (below), and some thick-walled cysts could have been asexual parthenospores (cysts that mimic a zygotic state, but are not the result of sexual fusion; Pfiester, 1984). Thus, in these cultures the presence of (apparently dormant) cysts was not interpreted with certainty to indicate sexual reproduction.

We documented excystment of single cells with two trailing flagella from thin walled cysts (Figure 3.3g). These cells could be called planomeiocytes (according to the technical definition; von Stosch, 1973) or, alternatively, may simply have been

planozygotes that excysted from temporary cysts. We also rarely observed excystment of two cells, each with two trailing flagella, from primary division cysts. These cells may have resulted from mitotic reproduction by diploid cells, or they may have been post- meiotic offspring cells that retained two longitudinal flagella. The former explanation would indicate a diplohaplontic potential (Pfiester, 1984), whereas the latter would suggest a condition rarely reported in dinoflagellates (e.g., Coats et al., 1984). These cells could not be distinguished from planozygotes under light microscopy. Although culture-induced abnormality (Barker, 1935; von Stosch, 1973; Loper et al., 1980) cannot be ruled out as the cause, life cycles other than strictly haplontic have been previously suggested for this and other heterotrophic dinoflagellates (Zingmark, 1970; Beam and Himes, 1984; Pfiester,

1984; Burkholder et al., 2001b).

Nuclear cyclosis has been described in zygotic nuclei of some dinoflagellates (von

Stosch, 1972, 1973; Pfiester and Anderson, 1987; Barlow and Triemer, 1988; Kita et al.,

1993), and refers to a pronounced swirling and mixing of the chromosomes during the pairing of homologues prior to the first meiotic division. Nuclear cyclosis was repeatedly observed and videotaped in P. piscicida cysts. It probably occurred more commonly than was observed, since replete vacuoles, abundant refractive lipid droplets, or darkened cyst walls obscured the nuclei of many cysts. When encountered, nuclear cyclosis was already in progress and lasted at least 1 h (the duration of the swirling prior to our observations was unknown). After the motion stopped, the chromosomal mass gradually expanded in size, and within 2-4 h a plane of protoplast division formed within the cysts. We observed nuclear cyclosis and subsequent veritable meiotic division in rapidly reproducing (< 6 h) thin-walled cysts (Figure 3.3h), demonstrating that in P. piscicida, zygotic cysts do not

always have thick walls or exhibit extended dormancy. Excystment of two flagellated cells followed nuclear cyclosis and cell division in these cysts (video link, piscnc1.mov). If two meiotic divisional steps were required, then meiosis was temporally separated

(uncoordinated; Beam and Himes, 1980), as described for some photosynthetic species

(e.g., von Stosch, 1973; Kita et al., 1993). Alternatively, a one-division meiosis could have occurred (perhaps without meiotic chromosome replication), as has been suggested for C. cohnii (Beam and Himes, 1975, 1980, 1984).

3.4.4b Pfiesteria shumwayae. Cells with two trailing flagella were not observed in these clonal P. shumwayae strains. Gamete fusion was documented in only one instance, and involved fusion between slightly anisogamous gametes. As soon as fusion began, the gametes lost their flagella and settled to the bottom of the culture, so that sexual activity may have been underestimated in these cultures. Within 12 h, the larger gamete had entirely absorbed the smaller cell. The resulting zygote appeared morphologically similar to the previously described P. shumwayae reproductive cysts, and it did not alter shape or size over a 3-day observation period. Nuclear cyclosis was not observed in P. shumwayae cysts from these cultures.

3.4.5 Population DNA Distributions. Algal prey nuclei were considerably smaller than Pfiesteria nuclei, and fell below the fluorescence threshold set for the analysis

(Whiteley et al., 1993; Parrow and Burkholder, 2002). Also, SYTOX Green-DNA emission was distinct from algal chlorophyll, thereby enabling exclusion of prey autofluorescence from the DNA analysis (Parrow and Burkholder, 2002). Flagellated cells of these Pfiesteria cultures exhibited flow cytometric population DNA distributions similar to those reported by Parrow and Burkholder (2002). Logarithmic and late-logarithmic

Figure 3.3

Figure 3.3. Light micrographs and scanning electron micrographs (SEMs, where noted) of

P. piscicida fed algal prey, including: (a) Small planozygotes with two trailing flagella, with Rhodomonas sp. prey cells. (b) Planozygote with two trailing flagella. (c) SEM of a small cell (presumed planozygote) with two longitudinal flagella. (d) Considerably larger, prey-replete (pigmented inclusions) planozygote with two trailing flagella. (e) SEM of a cell believed to be an aberrant planozygote with three longitudinal flagella. (f) Thick- walled cyst, with dinokaryotic chromosomes and a single pigmented inclusion. (g) Single cell with two longitudinal flagella (F) that had just excysted from a thin-walled cyst; note the discarded hyaline cyst wall (C) below the cell. (h) Large, thin-walled zygotic cyst undergoing nuclear cyclosis. Swirling, condensed chromosomes were visible in the nucleus

(arrow) and three prey-replete vacuoles (pigmented inclusions) filled the upper cell. Two flagellated cells were released < 3 h after nuclear cyclosis ceased. (i) Measured 3C DNA

‘doublet’ cell that had been electrostatically sorted for microscopic evaluation; the cell contained both 1C and 2C stained nuclei. (j) Synchronized flagellated cells released from reproductive cysts; the lower right cell had two longitudinal flagella. (k) Synchronized flagellated cells released from reproductive cysts, stained for lipids (red from Nile red stain; measured at 575 + 15 nm in flow cytometry) and DNA (green from SYTOX stain). Nile red stain appeared localized in cytoplasmic lipid drops, but some staining of plasma membrane was evident in this emission spectrum (ca. 640 nm, determined by the epifluorescence filter cube). Scale bars = 5 µm.

growth phase P. piscicida flagellates demonstrated a bimodal DNA distribution suggestive of G1-S-and G2 phases (1C and 2C), and typical of populations of eukaryotic asynchronously cycling cells (Gray et al., 1987) (Figure 3.4a). Logarithmic growth-phase

P. shumwayae flagellates showed a unimodal DNA distribution indicative of interphasic cells limited to a cytoplasmic growth phase, here assumed as G1 (1C), with a potential for inclusion of S phase cells (DNA synthesis; Figure 3.4b) (Wong and Whiteley, 1996; Bhaud et al., 2000; Parrow and Burkholder, 2002). Occasionally a low percentage (< 5%) of cells indistinguishable from 2C were detected in some P. shumwayae cultures, of an as-yet unknown role (Parrow and Burkholder, 2002). Logarithmically growing P. piscicida populations typically presented a measured S-phase component in 2-10% of the total population, whereas dense populations of starved P. piscicida flagellated cells presented 1C and 2C sub-populations with relatively low measured S-phase subpopulations (0-2%). In addition, P. shumwayae flagellated cells exhibited, on average twice the DNA-fluorescence of 1C P. piscicida, as reported by Parrow and Burkholder (2002).

Analysis of late-log growth phase populations of sexually reproducing P. piscicida populations in these clones often revealed a substantial increase in 2C DNA cells (Figure

3.4c). Microscopic observation revealed that many cells present at the time of fixation had two trailing flagella. Diploid cells cannot be differentiated based on DNA content from G2 zoospores preparing for mitosis, particularly in an asynchronous population, as discussed in a previous microfluorometric DNA analysis of dinoflagellate sexuality (Cetta and

Anderson, 1990). However, cells with triploid (3C) and tetraploid (4C) DNA complements can be differentiated (Figure 3.4d,e). The apparent products of 2C + 1C fusions were detected as low-frequency 3C DNA-fluorescence events. Similarly, the apparent 2C + 2C

fusions (4C) were rarely observed in culture. These events appeared in the DNA histograms in low frequency (< 5%) relative to 1C and 2C cells, as in Parrow and

Burkholder (2002), and consistent with the rarity in observations of such fusions in culture.

Alternatively, 4C cells could be the result of meiotic interphase DNA replication in motile

2C zygotes, but the low frequency encountered suggests that DNA replication in planozygotes either did not occur, or occurred infrequently. It is even more unlikely that the discrete 3C DNA subpopulations measured were the result of an incomplete DNA replication event shared by all 3C cells. When electrostatically sorted and observed microscopically, 3C and 4C cells appeared as uninucleate (rarely binucleate) cells of typical zoospore morphology. Paraformaldehyde fixation often did not preserve flagella intact, so that the flagellar complement of the 3C and 4C cells could not be determined (Parrow and

Burkholder, 2002). Some of the clonal isolates with sexual reproduction did not yield 3C and 4C DNA sub-populations; thus, the phenomenon may not be a consistent feature in sexual populations of algal-fed P. piscicida.

In any cell suspension, artificial aggregation can result in apparent cell pairs or

‘doublets’, which must be discriminated in DNA analyses to prevent inaccuracy in histogram interpretation (Shapiro, 1995). Potential doublets in these P. piscicida cultures were differentiated by constructing bivariate plots of DNA fluorescence signal peak height versus integral signal area, in which DNA doublets deviated from a linear ratio of the two processed signals (Green et al., 1996; Wersto et al., 2001). In all samples, a low percentage of possible doublets was detected to the right of the singlet diagonal distribution. In the P. piscicida cultures with high proportions of sexual cells, significantly more DNA doublets were detected (Figure 3.4f), likely because early-stage fusing gametes were in effect DNA

‘doublets’. Examination of this plot demonstrates that the majority of 3C and 4C DNA events did not deviate from the singlet line and, therefore, were not considered the result of artificial cell aggregation. An obvious pattern of DNA doublet events connected the 1C -

2C and 2C - 3C DNA singlet populations (Figure 3.4f). Microscopic evaluation of sorted cells comprising the 2C – 3C patterned doublets revealed binucleate cells in obvious 1C +

2C configurations (Figures 3.3i, 3.4f). Rarely, apparent 2C + 2C configurations were observed in separate preparations. These binucleate cells likely were in the process of either plasmogamy or cytokinesis at the moment of fixation. If the binucleate cells were dividing, then cells with both a 1C and a 2C nucleus likely were undergoing unequal nuclear division, or ‘budding’. This process rarely has been described in other dinoflagellates (Triemer and Fritz, 1984; Silva and Faust, 1995).

3.4.6 Cell Cycle Synchronization and Storage Products of a Synchronized

Population. We obtained a high degree of cell synchrony (> 90% 1C) in the tested P. piscicida and P. shumwayae cultures using our preferential lysis technique, similar to that obtained using selective attachment and filtration in previous research with C. cohnii

(Wong and Whiteley, 1996). The technique eliminated many potential microbial contaminants such as algal prey, resulting in a clean suspension of highly synchronized cells. In these preliminary experiments, sterile-filtered artificial seawater was used as the excystment media and, thus, particulate food was not available for the synchronized populations. This treatment permitted determination of short-term cell cycle progressions in the absence of exogenous particulate resources, highlighting potential nutritional restrictions on cell cycle progression and possible starvation-induced reliance on endogenous lipid reserves.

A B C 1C 1C 2C

2C y Bead 1C uenc q

Fre

Relative DNA (AU)

D r E F 1C

1C 2C 3C 4C ht

2C g y ht Scatte

g uenc nal Hei q g

Fre 3C DNA Si 4C Li Forward

Relative DNA (AU) Relative DNA (AU) DNA Signal Area

Figure 3.4. DNA distributions in semi-log plots (A-E) or linear scale (F); algal prey DNA fluorescence was below the threshold set for the analysis, and prey chlorophyll autofluorescence did not overlap SYTOX Green-DNA emission, as in Parrow and

Burkholder (2002). (A) P. piscicida in logarithmic growth phase, with 1C and 2C DNA subpopulations apparent. (B) P. shumwayae in logarithmic growth phase, showing a unimodal DNA distribution (1C DNA only; see text). (C) P. piscicida sexual culture in late logarithmic growth phase, with a dominant 2C DNA subpopulation. Microscopic observation of the same population revealed a dominance of cells with two trailing flagella.

(D) P. piscicida sexual culture in late logarithmic growth phase, with 1C, 2C, 3C, and 4C

DNA subpopulations detected. (E) Sexual P. piscicida sexual culture showing forward light scatter versus DNA fluorescence, with 1C, 2C, 3C, and 4C DNA subpopulations

apparent. Note that forward scatter signals did not increase with DNA fluorescence. (F) P. piscicida sexual culture, showing DNA signal height (peak) versus area (integral) for DNA doublet discrimination; DNA singlets fell along the diagonal line, while DNA doublets fell to the right of the singlet line. Apparent polyploid (3C, 4C) DNA distributions were not representative of all sexual populations analyzed (e.g., Figure 3.4C). AU = arbitrary units.

3.4.6a P. piscicida. Using video microscopy, we observed a ~20-30% excystment rate during the first 12 h after addition of sterile artificial seawater to the flasks containing

P. piscicida reproductive cysts for the synchronized cultures. Since these cells excysted into sterile artificial seawater, it seems unlikely that the excystment was in response to some chemical signal. The excysted cells may have responded to a chemical signal (although the medium was sterile, artificial seawater) or, alternatively, they may have excysted coincidentally to the treatment, which would imply that excystment was in response to an internal clock rather than an external signal. Either one cell or a pair emerged from each cyst. After < 24 h when flagellated cell densities were 104 mL-1, we isolated them from the remaining cyst population. The excysted cells were variable in size, and some had two trailing flagella (Figure 3.3j). These likely comprised the post-synchronization 2C DNA cells detected in P. piscicida. DNA staining revealed that the excysted cells were mostly uninucleate (rarely binucleate). As mentioned, since some apparent excystment cells had two trailing flagella, similar cells in asynchronous cultures could be mistaken for pre- encystment planozygotes (as in Anderson et al., 1983).

The newly excysted, flagellated cells obtained from the synchronization contained abundant cytoplasmic lipid storage bodies, indicated by specific Nile Red stain localization and fluorescence (Figure 3.3k) (Greenspan et al., 1985; Cooksey et al., 1987). These cells exhibited about a 10-fold increase in mean lipid-fluorescence (based on Nile Red staining), relative to late-logarithmic-growth phase (prey-depleted) asynchronous zoospores of the same clone, and also produced an increased side-light scatter signature (Figure 3.5a,b).

The degree of synchrony of the excysted cells was assessed by flow cytometric measurement of the DNA content of SYTOX Green stained subsamples. At 24 h after

synchronization by our preferential lysis technique, the DNA content of 91% of the excysted cells was consistent with the DNA of 1C P. piscicida zoospores (G1 cell cycle phase). The remaining 9% possessed 2C DNA, and none of the cells contained intermediate DNA (S phase) (Figure 3.6a). Samples were analyzed at 12-h intervals thereafter, and varied from the initial measurements by < 1%. In the final sample, analyzed at 72 h post-synchronization, 1C DNA cells comprised 90% of the population and 2C DNA cells comprised the remainder, again with no intermediate DNA cells detected (Figure

3.6b). Cell density at 24 h post-synchronization was ca. 1.4 x 104 mL-1, and at 72 h was ca.

1.3 x 104 mL-1, demonstrating that the synchronized population did not increase in number during the 48 h period. Nor were cysts observed in the cultures. The synchronized cells excysted into media devoid of prey, and apparently remained in their respective cell cycle positions because prey substrates required for DNA synthesis and cell division were lacking.

By 72 h, the lipid content of the excysted cells had decreased to levels similar to those measured in late logarithmic-growth phase, asynchronous zoospores of the same clone, indicating that the endogenous lipid reserves were utilized for motility and other cellular metabolism (Reed et al., 1999), but were insufficient to support cell production.

After that final sampling, the flagellated cells persisted in culture with no prey for 10 days following the synchronization, during which time they visibly grew smaller and more translucent until they appeared to consist of little more than a motile, membrane-bound nucleus. The cells were not observed to encyst. At 10 days we fed this culture a dense innoculum of cryptomonads. The P. piscicida cells grazed the prey and grew to normal zoospore proportions; thus, they had remained capable of phagocytotic activity.

3.4.6b P. shumwayae. We observed a < 30% excystment rate from P. shumwayae reproductive cysts during the first 12 h after addition of sterile artificial seawater. Either one cell or a pair emerged from each productive cyst. Individual excysted zoospores were uninucleate with abundant cytoplasmic lipid bodies. In addition, many excysted zoospores had 1-2 apparently food-replete epithecal vacuole(s) (e.g., Figure 3.1l). Synchronized populations exhibited a single DNA peak when analyzed with flow cytometry, similar to the DNA pattern typically observed for P. shumwayae zoospores in asynchronous cultures

(e.g., Figure 3.4b). Analysis of DNA histograms 72 h after synchronization suggested that a significant proportion of the synchronized populations had entered DNA synthesis (S) phase (Figure 3.7a,b). The cultured populations were not observed to encyst or reproduce during the 48-h sampling interval, and cell densities did not decline significantly.

A B

Zoospores Germlings

Side Light Scatter

Relative Lipid (AU)

Figure 3.5. Log-log bivariate histograms of side-light scatter versus lipid fluorescence

(indicated by Nile red stain, recorded at 575 + 15 nm, selected for cytoplasmic neutral lipids rather than plasma membrane; Greenspan et al., 1985; Brzezinski et al., 1993). (a)

Late logarithmic growth phase (prey-depleted), asynchronous zoospores. (b) Newly germinated, flagellated cells released from P. piscicida reproductive cysts (same clone as in

Figure 3.5a). Excysted cells (germlings) exhibited a ~10-fold increase in mean lipid- fluorescence, relative to late-logarithmic growth phase (prey depleted) asynchronous zoospores of the same clone. The slight increase in mean side-light scatter may have been due to abundant cytoplasmic refractive lipid droplets, since the newly germinated cells were not, on average, larger than the asynchronous zoospores. Unstained cells and debris clustered along the Y-axis in both treatments. AU = arbitrary units.

A B

24 H 72 H 1C – 91% 1C – 90% 2C – 9% 2C – 10%

Frequency

Relative DNA (AU)

Figure 3.6. Semi-log DNA distributions of synchronized P. piscicida cells (a) 24 h and (b)

72 h after synchronization (leftmost peaks indicate fluorescent bead standards). No significant cell cycle progression was observed under nutritional limitation. AU = arbitrary units.

Figure 3.7. Deconvoluted linear DNA distributions of synchronized P. shumwayae zoospores, performed with Multicycle DNA cell cycle analysis software; insets (upper left corners) show raw data. Leftmost peaks in all distributions were of CRBC DNA- fluorescence standards. (a) DNA distribution 24 h after synchronization, showing a unimodal DNA peak that was slightly skewed to the right (CV = 14.5); analysis suggested that > 90% of zoospores exhibited 1C (G1) DNA. (b) DNA distribution 72 h after synchronization, showing DNA distribution skewed farther to the right. The analysis suggested that most zoospores had entered S phase. Distribution analyses were performed by least squares curve-fitting after automatic software DNA peak detection (Chi squares <

4.4). AU = arbitrary units.

3.5 Discussion

This study represents the first report of sexual reproduction in Pfiesteria spp. cultures fed algal prey rather than live fish (below); the first report of a technique for cell cycle synchronization in these heterotrophic dinoflagellates; and the first information on storage products of cells excysted from Pfiesteria reproductive cysts. Our preferential lysis technique for cell cycle synchronization proved equally amenable to P. shumwayae as P. piscicida, and we are currently using it to investigate the lipid metabolism and the DNA cell cycle for both species. The algal-fed clonal zoospore cultures also enabled description of certain reproductive behaviors that apparently were conserved for both Pfiesteria spp.

3.5.1 Cell Reproduction and Cysts. The Pfiesteria spp. cultures used in most of this study each existed as an asynchronous population of cells in different reproductive stages, some of which appeared morphologically similar but were actually developmentally distinct. In each species, the asynchronous populations consisted of motile flagellated cells and nonmotile cysts. Motile stages were phagocytotic, while most cysts were reproductive.

Both species also produced non-reproductive temporary cysts, as previously shown for cultures grown with fish (Burkholder et al., 2001b). The final flagellated excystment products of reproductive cysts numbered 2 or 4, and excysting offspring were often temporarily conjoined. Primary reproductive cysts either yielded 2 flagellated cells after an internal protoplast division, or underwent a complete division to yield 2 secondary cysts, each with a complete new cyst wall. Secondary cysts yielded two flagellated cells. Some primary cysts had more thickened cyst walls and appeared dormant or occasionally were observed to germinate. In cultures left to starve (weeks to months), flagellated cells ultimately disappeared from the cultures. The final cyst production in these treatments was

considerably lower than preceding flagellated cell numbers, suggesting that not all flagellated cells successfully encysted under nutritional limitation (Anderson et al., 1985,

1995).

P. piscicida may divide by binary fission while motile, as reported in some fish- killing cultures (Burkholder and Glasgow,1997a), and as interpreted by Parrow and

Burkholder (2002) in some algal-fed cultures. In the clones examined in this study, paired motile cells were sometimes found in an oblique alignment suggestive of binary fission.

These cells also may have been early fusing gametes or recently excysted temporarily conjoined zoospores. Collectively, observations suggest that two classical modes of dinoflagellate cell division, desmoschisis and eleutheroschisis (Pfiester and Anderson,

1987), may be possible in Pfiesteria. Although the cell division of a given dinoflagellate species has been regarded a conserved character (often used as a taxonomic trait; Smith,

1955; Beam and Himes, 1980), similar reproductive plasticity has been described for

Prorocentrum lima (Ehrenberg) Dodge (Faust, 1993). Given the complex life cycles of

Pfiesteria spp., the considerable polymorphism among and within multiple stages, and the wide array of prey species consumed by these stages, it is possible that Pfiesteria also exhibits variability in cell division. The influence of strain variability and culturing effects on this behavior merits further investigation.

Division cysts in Pfiesteria spp. have been observed in clonal cultures fed live fish

(Burkholder et al., 2001b; Parrow and Burkholder, 2002), and in clonal and non-clonal isolates fed algal prey (this study; Parrow and Burkholder, 2002). They apparently represent a conserved stage and a conserved mode of vegetative reproduction, regardless of culture conditions and differences in prey availability. Division cysts (i.e., cell division in

nonmotile cysts) also have been reported in dinoflagellates such as the estuarine or marine coastal heterotrophs, C. cohnii, Katodinium fungiforme (Annisimova) Loeblich III, and

Paulsenella chaetoceratis (Paulsen) Chatton, P. kornmannii Drebes and Schnepf, and P. vonstoschii Drebes and Schnepf (Kubai and Ris, 1969; Beam and Himes, 1974; Spero and

Morée, 1981; Drebes and Schnepf, 1988; Bhaud et al. 1991). In each of these taxa, zygotic cysts could not be distinguished by gross morphology from vegetative cysts. Similarly, our observations from algal-fed Pfiesteria cultures suggested that zygotic cysts cannot always be distinguished from vegetative cysts under light microscopy, unless the flagellar complement of the cell initiating a particular cyst was discerned, or nuclear cyclosis was observed. We hypothesize that the ultimate number of offspring cells produce by Pfiesteria spp. reproductive cysts is related to the nutritional history of the encysting progenitor cell.

The ploidy of the encysting cell may also be important in determining the number of flagellated offspring cells produced or, alternatively as for C. cohnii (Beam and Himes,

1984), ploidy may have little influence on the number of excystment products.

3.5.2 Sexual Reproduction. Little is known about the factors that trigger sexuality in the relatively few dinoflagellate species examined to date. For photosynthetic species, gamete production has been induced by nutrient depletion (especially N or P; Pfiester and

Anderson, 1987; Pfiester, 1989; Olli and Anderson, 2002). Sexual reproduction among cultured heterotrophic dinoflagellates more commonly has been described as “spontaneous”

(i.e., factor(s) unknown; Morey-Gaines and Ruse, 1980; Beam and Himes, 1984; Timpano and Pfiester, 1986; Popovský and Pfiester, 1990). In Noctiluca, an endogenous cell division clock controlled by cell density has been shown to regulate presumed sexual reproduction (Sato et al., 1998). Alternatively, certain prey, a change in prey type, or prey

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limitation has been invoked as sexual inducers for some species (Zingmark, 1970; Spero and Morée, 1981; Pfiester and Anderson, 1987; Drebes and Schnepf, 1988). Pfiesteria has been so described, with sexuality of toxic strains expressed in the presence of live fish

(Burkholder and Glasgow, 1995, 1997a,b; Burkholder et al., 2001b; Glasgow et al., 2001b).

The present research demonstrates that sexual reproduction can also occur in P. piscicida clones maintained solely on unicellular algae for extended periods (months to years) (note that Burkholder et al. 1998 also described zoospores of Pfiesteria spp. and certain other heterotrophic dinoflagellates with higher cell production and ‘swarming’ behavior when presented with senescent prey). Cell density and swarming behavior have been suggested as factors influencing sexual reproduction in other heterotrophic dinoflagellates (Beam and

Himes, 1977; Spero and Moreé, 1981; Drebes and Schnepf, 1988; Sato et al., 1998).

The potential for underestimating planozygote abundance in Pfiesteria cultures was noted by Burkholder and Glasgow (1995), who pointed out that small planozygotes which had lost their longitudinal flagella in preserved material could not be distinguished from zoospores. Thus, early enumerations of planozygotes were based on (large) cell size if the flagellar complement could not be ascertained. Size had previously been used as a primary criterion for discerning planozygotes in certain other dinoflagellate species (Anderson et al.,

1983; Faust, 1998; Probert et al., 1998; Kremp and Heiskanen, 1999; Olli and Anderson,

2002). Most previous research on Pfiesteria life cycles focused on actively toxic fish- killing strains and consistently documented periodic abundance of gametes, planozygotes and zygotic cysts with live fish prey (Burkholder and Glasgow, 1995). Sexual stages were not found in subcultures from the same Pfiesteria clones fed algal prey rather than live fish

(Burkholder and Glasgow, 1995, 1997a,b; Burkholder et al., 2001a,b; Glasgow et al.,

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2001b). Pfiesteria populations were observed for days to weeks following transfer from fish cultures to medium with algal prey, and sexual stages declined (although phagotrophy on algal prey by planozygotes was observed; Burkholder et al., 1995b; Burkholder and

Glasgow, 1995, 1997a,b). Sexual activity may not have occurred in those cultures, which represented relatively few clones (Burkholder et al. 2001a; n < 20). Alternatively, the transfer process may have temporarily suspended sexual activity, since Pfiesteria populations often require an adjustment period prior to resuming cell production when culture conditions and/or prey type are significantly altered (Burkholder et al., 2001a,b;

Parrow et al., 2001). If sexual activity did occur, small planozygotes (commonly observed in the present research with algal-fed cultures, and well below the size range given in the P. piscicida species description by Steidinger et al. 1996a) would have been underestimated or missed.

In this research, evidence of sexual reproduction was observed in both Pfiesteria species cultured on algal prey, especially in those that had been cultured on algal prey for months to years. Evidence for sexuality in algal-fed P. shumwayae cultures thus far is limited to a single apparent anisogamous fusion that resulted directly in a cyst, with observation not yet available on excystment products. The zygotic cyst was identical in morphology to P. shumwayae vegetative cysts; thus, at present sexual and vegetative cysts cannot be differentiated with certainty in this species unless the cell(s) initiating a particular cyst are observed.

In P. piscicida, sexual reproduction was confirmed by the presence of fusing gametes, planozygotes, and nuclear cyclosis in these clonal cultures, thus indicating homothally (i.e. self-fertility). These strains tended toward hologamy but were not strictly

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hologamous, as the smallest gametes could be distinguished from other cells (Pfiester and

Anderson, 1987; Burkholder et al., 1995a, 2001a,b). The details of gametogenesis are, as yet, unclear. Sexual fusion in some zoospores may be facultative, that is, zoospores may be undifferentiated, able to act as gametes and engage in sexual fusion, or to continue with vegetative division if a gametic partner is not located (Pfiester, 1975; Beam and Himes,

1980). Facultative sexual activity could be potentially advantageous in that it would eliminate the need to find another gamete for survival (Dacks and Roger, 1999). We hypothesize that the smallest fully differentiated gametes die if they do not complete syngamy (analogous to the lack of survival of small cells that do not fuse in other species such as diatoms; Graham and Wilcox, 2000), as these cells were not observed to feed and did not contain prey inclusions. Meiotic segregation has not yet been described, and would require the isolation/production and crossing of phenotypic mutants together with careful manipulation of single cells for genetic recombination analysis (Beam and Himes, 1980,

1984). The phenotypic mutants necessary for that in Pfiesteria spp. research have yet to be isolated. Suitable phenotypic mutants could also be used in mating experiments between different Pfiesteria spp. isolates, toward determining biological or sibling species compatibility (Beam and Himes, 1977, 1984; Parrow and Burkholder, 2002).

3.5.3 Cellular DNA Content and Implications About Pfiesteria spp. Life Histories.

Flow cytometric DNA analysis demonstrated a notable difference between the two

Pfiesteria spp., as represented by these cultures, in cell cycle distributions of asynchronous flagellated populations. P. piscicida exhibited both 1C and 2C relative DNA, whereas P. shumwayae exhibited primarily 1C relative DNA with apparent inclusion of S-phase cells, and 2C DNA cells rarely detected. As in Parrow and Burkholder (2002), in these P.

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shumwayae cultures, G2 + M cell cycle phases occurred primarily in non-motile cells (i.e. division cysts) which were not included in the flow cytometric DNA analyses. Moreover,

P. shumwayae contained, on average, about twice the cellular DNA content of P. piscicida, which may help explain the lower apparent growth rate often observed for this species on algal prey (Parrow and Burkholder, 2002; described for other dinoflagellates by Pan and

Cembella, 1998). If vegetatively reproducing P. piscicida zoospores complete G1 – S – G2 phases during growth, then they spend more of their cell cycle as zoospores than P. shumwayae (that is, in asynchronous populations of equal cell numbers, more P. piscicida cells would be expected to be observed as zoospores).

Starved P. piscicida cultures appeared to express nutritionally-dependant cell cycle restriction points, beyond which DNA synthesis and cell division could not occur without addition of prey. Analysis of stationary phase (prey-depleted) P. piscicida showed that measured S phase cells were rare to nonexistent, relative to that measured in logarithmically growing (prey replete) populations. The data suggest a specific cell-cycle restriction point between G1 and S, delaying DNA synthesis until sufficient nutritional reserves are accumulated (Whiteley et al., 1993). Similarly, in a dense, synchronized but starved population, 1C and 2C cells apparently remained in their respective nuclear conditions in the absence of prey, unable to replicate DNA or divide, suggesting an additional cell cycle restriction point between G2 and M, as in other eukaryotes (Nurse and Bisset, 1981;

Whiteley et al., 1993; Bhaud et al., 1994; Wong, 1996). Small P. piscicida planozygotes also appeared to require prey before forming zygotic cysts, possibly to meet the metabolic demands of reproduction while encysted and/or to form additional reserves to survive dormancy (as previously suggested for the planozygotes of photosynthetic dinoflagellates,

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e.g., Anderson et al., 1985). In contrast, these P. shumwayae synchronized zoospores appeared to demonstrate a limited capacity for DNA synthesis in the absence of an exogenous food supply. The measured progression into S-phase likely was supported by the vacuolar food reserves commonly observed in excysting cells. We hypothesize that the observed discrepancy between the two species was the result of an incidental difference in culture conditions (prey availability) prior to cyst isolation, and does not represent a conserved species-level cell cycle trait. However, the observed progression of DNA content in synchronized populations does support the premise that P. shumwayae zoospores occupy G1 through S-phase in DNA cell cycle position, as suggested by Parrow and

Burkholder (2002).

DNA cell cycle analysis of P. piscicida zoospores was complicated by the presence or, in some cases, the predominance of 2C DNA zygotic cells. We also confirmed our observations of rare zygote-gamete (3C) and zygote-zygote (4C) fusions in some cultures with supporting flow cytometric data. In some P. piscicida cultures examined here, the majority of 2C DNA flagellated cells likely were diploid, as indicated by two longitudinal flagella. Parrow and Burkholder (2002) interpreted cell cycles in algal-fed Pfiesteria cultures under the assumption that asexual reproduction was the predominant reproductive mode. An alternate premise is that most or all 2C events observed in P. piscicida asynchronous DNA distributions are planozygotes and/or planomeiocytes. For this premise to be true, sexuality would have to be expressed consistently by the cultured population.

This condition would require that measured intermediate DNA cells (presumed as S phase) were artifactual, having resulted from high CVs on the 1C and 2C DNA peaks due to cytoplasmic DNA contamination from endosymbiotic bacteria and/or algal prey DNA

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(Parrow and Burkholder, 2002). Previously, sexual reproduction had not previously been observed in Pfiesteria strains fed algal prey for shorter periods (days to weeks; Burkholder and Glasgow, 1995, 1997a), and sexuality was intermittent in fish-killing cultures

(Burkholder et al. 1995b). In contrast, continuous sexuality has been reported for the heterotrophic dinoflagellate, C. cohnii (Beam and Himes, 1984). We are continuing to examine this possibility for Pfiesteria in ongoing work with synchronized populations.

3.5.4 Insights About Pfiesteria Versus Other Heterotrophic Dinoflagellates. The modes of reproduction documented here for Pfiesteria spp., considered together with the zoospore plate structure verified in this study and other, previously described traits, provide insights toward resolving recent questions about relationships between Pfiesteria and other heterotrophic dinoflagellates, especially parasitic forms within the Blastodiniales (Litaker et al., 1999; Steidinger et al., 2001), coccoid or flagellated forms within the Peridiniales

(Fensome et al., 1999; Steidinger et al., 2001), and amoeboid forms within the

Dinamoebales (Steidinger et al., 1996a, 2001). We consider Pfiesteria within the context of each of these orders as follows.

Placement of Pfiesteria spp. within the Blastodiniales, or within an intermediate order between the Blastodiniales (Blastodinophyceae) and the Peridiniales (Dinophyceae), has been suggested (reviewed in Steidinger et al., 2001). The Blastodiniales is a poorly defined, likely polyphyletic order (Litaker et al., 1999) known primarily for parasites.

Small subunit ribosomal DNA gene sequence analysis has indicated a close relationship between P. piscicida and a Blastodiniales representative, Amyloodinium ocellatum Brown, an ectoparasite of marine and estuarine fishes (Litaker et al., 1999). However, a close genetic relatedness between Pfiesteria spp. and A. ocellatum does not, de facto, imply a

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parasitic life style for Pfiesteria spp. since species within the Blastodiniales show a range of morphologies and physiologies, from free-living to parasitic (Cachón and Cachón, 1987).

Flagellated cells of both species attach to and feed upon the dermal tissues of nonspecific marine teleost fish species (Brown, 1934; Brown and Hovasse, 1946; Burkholder et al.,

2001a,b). However, fish are one of many food sources for Pfiesteria spp. (Burkholder et al.

2001a, Glasgow et al., 2001b). In contrast, fish are regarded as an obligate food source for

A. ocellatum (Brown, 1934; Brown and Hovasse, 1946; Landsberg et al., 1994; Noga,

1987), although other metazoans may also be parasitized (Colorni, 1994).

Heterotrophic dinoflagellates fall along a predator-parasite continuum (Coats,

1999), making the trophic designation of predator or parasite somewhat arbitrary for intermediate species. Gaines and Elbrächter (1987) defined parasitic dinoflagellates as those species exhibiting morphologically different feeding and reproductive stages, and as producing numerous progeny after a single feeding act. Pfiesteria spp. zoospores have morphologically different feeding (flagellate) and reproductive (cyst) stages, and more than

2 offspring cells (perhaps up to 8, but not hundreds or more as is common in parasitic dinoflagellates) can be produced from a progenitor flagellated cell. Pfiesteria spp. are not modified for an obligate parasitic lifestyle (Litaker et al., 1999); flagellated feeding stages in Pfiesteria are not morphologically distinct from swimming stages other than by peduncle extension and attachment, and there is no known non-flagellated, morphologically distinct trophont feeding stage.

Nor do Pfiesteria spp. demonstrate host specificity. Their naturally occurring food sources range from detritus to unicellular algae, to fish, and they readily consume exotic food sources such as human erythrocytes and rat fibroblasts (Glasgow et al., 1998; our

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unpubl. data). Thus, Pfiesteria spp. occupy an ecological role as free-living, general predators (Burkholder and Glasgow, 1995, 1997a,b; Litaker et al., 1999). Pfiesteria spp. can be maintained on a diet consisting solely of unicellular algal prey (this study) wherein, unlike the parasitic dinoflagellate, Amoebophyra (Maranda 2000), it does not occupy the algal cells. Under such conditions it would seem inappropriate to refer to the free-living

Pfiesteria population as an ‘infection,’ or the algal prey as ‘hosts.’ Rather, they are free- living herbivores grazing on algae. Thus, the term dinospore (Litaker et al., 1999;

Steidinger et al., 2001; Vogelbein et al., 2001), and other terms typically reserved for parasites (Fensome et al., 1993; Coats, 1999), might be inappropriate for use in reference to

Pfiesteria zoospores.

Placement of Pfiesteria spp. within the Peridiniales has been suggested based on two considerations. The first is that a coccoid stage with closer morphological affinity to the Peridiniales may predominate in some strains under certain culture conditions (Fensome et al., 1999; Steidinger et al., 2001). Based on the present research with algal prey, flagellated stages dominated the 40 clones examined, whereas the coccoid stages were reproductive cysts. Other research with some strains and fish prey has indicated that amoeboid stages predominate although flagellated stages are the dominant toxic form

(Burkholder and Glasgow, 1995, 1997a,b) and coccoid (palmelloid or cyst) stages can occur (Burkholder et al., 2001b). The type species P. piscicida was named considering the dominance of amoeboid stages in some strains as the primary taxonomic feature (Steidinger et al., 1996a), and the naming of the second species followed that precedent (Glasgow et al.,

2001b). Yet, published research consistently has shown that the dominant stage varies depending on culture conditions and strain (e.g., Burkholder and Glasgow, 1995, 1997a,b;

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Burkholder et al., 1992, 2001a,b; Cancellieri et al., 2001; Glasgow et al., 2001b; Parrow et al., 2001; Parrow and Burkholder, 2002).

The second consideration for possible placement of Pfiesteria spp. within the

Peridiniales is the thecal plate structure (number and arrangement) of the zoospore stage, which is clearly peridinoid (Steidinger et al., 1996a; Fensome et al., 1999). Plate tabulation, although used somewhat arbitrarily to designate species within genera (e.g.,

Steidinger et al., 2001; below), is a more consistent taxonomic trait than stage dominance because, thus far, it has generally remained consistent within a species across strains, culture conditions, and prey sources (Fensome et al., 1993, 1999; Steidinger et al., 1996b;

Taylor, 1999). The plate structures assessed and cross-corroborated for P. piscicida and P. shumwayae in this study were identical to those determined for the two species in fish- killing cultures (Marshall et al., 2000; Burkholder et al. 2001a,b; Glasgow et al., 2001b).

The “gold standard” for Pfiesteria spp. identifications, including in the formal naming papers (Steidinger et al., 1996a; Glasgow et al., 2001b) is considered to be the plate structure of its zoospores, with molecular probes regarded as secondary (less reliable) evidence (Burkholder et al., 2001a,c; PICWG, 2001). We therefore suggest that a more consistent and, therefore, more taxonomically reliable approach would be to consider the plate structure of Pfiesteria flagellated stages as the primary trait in order placement

(Fensome et al., 1999). On that basis, Pfiesteria would be better placed within the

Peridiniales (also note that A. ocellatum, apparently closely related on the basis of its 18S rDNA sequence, has a similar peridinoid plate structure; Landsberg et al., 1994).

A final question, based on plate structure, concerns the relatedness of the two

Pfiesteria spp. used in this research. Steidinger et al. (2001) suggested the need for re-

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evaluation of placement of P. shumwayae within the genus Pfiesteria, based on the premise that its plate structure is considerably different from that of P. piscicida. However, the plates of the two Pfiesteria spp. are comparable, relative to the plate variability of species within other genera. P. piscicida and P. shumwayae have identical plate structures with exception of 1 plate (5-6 precingulars may be present). In contrast, according to Steidinger et al. (2001), Peridiniopsis spp. may have 3-5 apical plates, 0-1 anterior intercalary plate, 6-

7 precingular plates, and 3-5 sulcal plates; Peridinium spp. may have 2-3 anterior intercalaries, 5-6 cingulars, and 5-6 sulcal plates; Amyloodinium spp. may have 6-8 cingulars, etc. The two Pfiesteria spp. have close genetic affiliation as well, based on their

18S rDNA sequence (Oldach et al., 2000).

3.5.5 Status of the Field and Future Directions. Investigations of dinoflagellate life cycles, even with clonal cultures, are difficult (e.g., Bursa, 1970; Walker, 1982; Popovský and Pfiester, 1990; Highfill and Pfiester, 1992; Pfiester, 1984; Pfiester and Anderson, 1987;

Marasovic, 1993; Ōuchi et al., 1994; Berland et al., 1995; Horiguchi and Chihara, 1998;

Probert et al., 1998; Seo and Fritz, 2001). Morphologically distinct cell types often have been reported, but the sequence of events that created them and their ultimate fate have often remained uncertain, especially in unsynchronized heterotroph populations (e.g.,

Zingmark, 1970; Beam and Himes, 1974, 1984; Pfiester and Popovský, 1979; Morey-Gains and Ruse, 1980; Pfiester and Lynch, 1980; Spero and Morée, 1981; Popovský and Pfiester,

1982; Timpano and Pfiester, 1986; Drebes and Schnepf, 1988; Schnepf and Drebes, 1993;

Buckland-Nicks et al., 1990, 1997; Buckland-Nicks and Riemchen, 1995; Steidinger et al.,

1996a; Burkholder and Glasgow, 1997a,b; Appleton and Vickerman, 1998; Burkholder et al., 1998; Elbrächter and Qi, 1998; Sato et. al., 1998). Observations of fixed cells can yield

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information about the cell types present at a particular moment, but is obviously limited in description of life cycle transitions. In Pfiesteria, live-cell assessment of salient features such as the flagellar complement requires high magnification and resolution, and adept tracking and focus. Addition of glycerol slows the cells and therefore is useful in making dual longitudinal flagella visible, but it is ultimately lethal to the treated population. Methyl cellulose is less disruptive to the cells, but the particulate structure of the compound hinders high-resolution imaging. Flagellated cells often lose their flagella when fixatives are added, confounding interpretations about some life history stages. Additionally, reproductive patterns and rates often fluctuate considerably among clones of Pfiesteria spp., even when they are maintained similarly. In dinoflagellates, different isolates can exhibit distinctive behaviors (e.g., Beam and Himes, 1977), and within cultures patterns of reproduction can change over time (e.g., Cetta and Anderson, 1990; Olli and Anderson, 2002). The factors that cause behavioral and morphological variability among subcultures are poorly understood, particularly for heterotrophic species (Barker, 1935; Burkholder, 2000). The reproductive patterns presented in this study appeared largely conserved among the isolates examined. However, sexual stages were more consistent or abundant in some cultures than others, and varied over time within cultures.

Although the observations presented here elucidate flagellated cell production in

Pfiesteria spp., several aspects remain to be clarified. For example, it remains unclear why some reproductive cysts underwent a complete cyst division (yielding secondary cysts, and uncommonly, tertiary cysts) prior to germination, whereas others did not. Both vegetative and zygotic cysts appear to be capable of division, but the ploidy of the flagellated cell that initiated any particular cyst was often unknown. Flagellated cells moved rapidly for

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extended periods and were difficult to track, so that direct observations of encystment were rare. P. shumwayae sexuality in these cultures was even more cryptic, with only a single observation of direct production of a P. shumwayae zygotic cyst that was morphologically indistinguishable from other cysts in light microscopy. Observation of nuclear cyclosis demonstrated that in P. piscicida, zygotic cysts do not necessarily exhibit thickened walls or extended dormancy. This condition differs somewhat from the conventional depiction of dinoflagellate sexual reproduction, largely developed from studies of photosynthetic species

(Pfiester and Anderson, 1987; Pfiester, 1989; Popovský and Pfiester, 1990; Anderson et al.,

1995), but parallels certain observations in some heterotrophic species (Beam and Himes,

1984; Drebes and Schnepf, 1988) and a few photosynthetic species (Pfiester, 1977; Kita et al., 1993).

The measured differences in cultured population DNA distributions from this study suggest important cell and life cycle differences between the two Pfiesteria spp. that require further examination. Apparent nutritionally dependent cell cycle restriction points have been identified which likely influence the basic ecology of Pfiesteria spp. (as for other heterotrophic dinoflagellates; Whiteley et al., 1993; Bhaud et al., 1994; Wong, 1996).

More information is needed on the intrinsic variability in cellular DNA distributions and patterns of cell cycle progression between P. piscicida and P. shumwayae, as well as among isolates within each species. The influence of culture conditions and other extrinsic factors on population DNA distributions, cell cycle progression, and biochemistry requires further examination to validate conserved patterns, and the extent to which these patterns can be extrapolated to field populations is not yet known. From the present research, the ability to produce large numbers of highly synchronized Pfiesteria spp. cells, as well as purified

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isolations of encysted stages, will considerably facilitate research in cell and life cycle progressions, and will aid further investigations into the biochemical and molecular events associated with the growth and proliferation of these ecologically important dinoflagellate species.

It is clear from this and previous research that encysted stages are important in the ecology of both Pfiesteria spp. Pfiesteria blooms in estuaries can be transient events that apparently result from massive recruitment from the benthos in response to some environmental signal(s) (Burkholder and Glasgow, 1995, 1997a; Burkholder et al., 1992,

1995a; 1998; 2001a; Glasgow et al. 2001a). Transient blooms are consistent with the reproductive patterns of Pfiesteria spp. presented here. Once satiated, phagocytic flagellated cells often become non-motile and reproduce, which could cause an apparent lag in the bloom. The observations presented here also support the premise that enhanced algal prey production can lead to higher abundance of Pfiesteria spp. encysted stages which can act as a “seed” population for the inoculation of future blooms. We have described nonmotile resistant stages in P. piscicida and P. shumwayae that strongly influence reproduction and survival through environmental and nutritional stresses. In ongoing research we are working to more fully characterize encysted stages in Pfiesteria spp., and their overall role in genetic recombination, species dispersal, and bloom initiation and termination.

Acknowledgments

Funding for this research was provided by an anonymous foundation, the North Carolina

General Assembly, the National Science Foundation (#9912089), the U.S. Environmental

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Protection Agency (#CR 827831-01-0), and the Department of Botany at North Carolina

State University. We thank all members of the Center for Applied Aquatic Ecology for their technical support. We also are grateful to H. Glasgow, E. Allen, N. Allen, D.

Kamykowski, and P. Rublee for their counsel.

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4. ESTUARINE HETEROTROPHIC CRYPTOPERIDINIOPSOIDS

(DINOPHYCEA): LIFE CYCLE AND CULTURE STUDIES

Matthew W. Parrow and JoAnn M. Burkholder

Journal of Phycology (accepted)

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4.1 Abstract

Cryptoperidiniopsoids are an unclassified group of delicately thecate, heterotrophic dinoflagellates known to be common in eastern U.S. estuarine waters. Over the past ten years cryptoperidiniopsoids were isolated from different geographical regions and cultured with cryptophyte algal prey. In the seven clonal isolates examined, reproduction was strongly linked to the availability of prey cells. The dinoflagellates phagocytized the contents of prey cells through a tube-like peduncle, similarly as close relatives Pfiesteria spp. and several other heterotrophic species. Cell division occurred while encysted, most commonly yielding two biflagellated offspring. Abundant fusing gametes, phagotrophic planozygotes, and cysts with a pronounced nuclear cyclosis characterized persistent sexuality. Cysts with nuclear cyclosis produced two flagellated offspring cells. The resistance of reproductive cysts to antimicrobial treatments was determined, and a simple high-yield technique was developed for population synchronization while ridding the dinoflagellates of contaminating vacuolar prey DNA and external contaminants. The DNA content and population DNA profiles of synchronously excysted cryptoperidiniopsoids from different isolates were measured using flow cytometry, and were related to the life history of these and other dinoflagellates. Cryptophyte-fed cultures with versus without extracellular bacteria were compared, and bacteria apparently promoted cryptoperidiniopsoid feeding and growth. Externally bacteria-free dinoflagellates were cultured in media enriched with dissolved organic nutrients, and apparent nutritional benefit occurred in some treatments. The potential for mixotrophic nutrition from maintenance of cryptophyte chloroplasts was examined using flow cytometrically sorted cells, but evidence of kleptoplastidy was not found in these isolates under the conditions imposed. These

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studies contributed new information about the life cycle and nutrition of cryptoperidiniopsoids.

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4.2 Introduction

It is estimated that approximately half of the living species of dinoflagellates are obligate heterotrophs (Kofoid and Swezy 1921, Schnepf and Elbrächter 1992). Among them, armored (thecate) phagotrophic dinoflagellates are becoming increasingly recognized as important biota in estuarine and shallow coastal waters (Hansen and Calado 1999,

Jacobson 1999, Jeong 1999). Some thecate photosynthetic species engage in phagotrophy

(Jacobson and Anderson 1996, Schnepf and Elbrächter 1999, Stoecker 1999), and many superficially gymnodinoid-appearing phagotrophic taxa are actually thinly armored with defined plate structures (Dodge and Crawford 1970, Burkholder et al. 1992, Landsberg et al. 1994, Steidinger et al. 1996, 2001). However, relatively few thecate, obligately heterotrophic species have been brought into controlled, sustainable laboratory culture for experimental study (Lessard 1993). Among these, Crypthecodinium cohnii (Seligo)

Chatton stands out as a model free-living species complex that can be cultured using both synthetic liquid and solid media (Provasoli and Gold 1962, Tuttle and Loeblich 1975, Beam and Himes 1980), likely because of an inherent inclination for osmotrophy related to its natural occurrence amidst rotting seaweed (Pringsheim 1956, Lee 1990). However, many important investigations of free-living phagotrophic thecate species have necessarily relied upon field material or short-term laboratory maintenance of cells (Gaines and Taylor 1984,

Jacobson and Anderson 1986, 1996, Hansen 1991, Strom and Buskey 1993, Calado and

Moestrup 1997). Other thecate phagotrophic species or isolates have proven amenable to longer-term culture, increasing the potential for detailed study (Buskey 1997, Weisse and

Kirchhoff 1997, Burkholder et al. 2001). Cryptoperidiniopsoids are one such group.

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Presently, “cryptoperidiniopsoid” refers to a generic group of estuarine phagotrophic dinoflagellates including several unnamed species with thecal plate patterns within the tabulation range of the proposed namesake genus, Peridiniopsis (Steidinger et al.

2001). Cryptoperidiniopsoids, along with various other small heterotrophic dinoflagellates, have been described as pfiesteria-like organisms (PLOs) due to a resemblance under light microscopy (LM) to flagellated cells of the genus Pfiesteria (Burkholder and Glasgow

1997, Marshall 1999, Marshall et al. 1999, Burkholder et al. 2001, Steidinger et al. 2001).

Cryptoperidiniopsoids are common in eastern U.S. estuaries (Marshall et al. 1999, Rublee et al. 2001, Seaborn et al. 2001). Aspects of cryptoperidiniopsoid feeding (myzocytosis;

Gaines and Elbrächter 1987), prey preferences, and growth rates in culture have been characterized (Seaborn et al. 1999, Marshall et al. 2000, Burkholder et al. 2001, Parrow et al. 2001, Seaborn et al. 2001, Eriksen et al. 2002). The potential importance of bacteria present in cryptoperidiniopsoid cultures has begun to be examined (Alavi et al. 2001).

Genetic sequence analyses have established that cryptoperidiniopsoids share a close phylogenetic relationship with Pfiesteria, and that both are relatively ancestral among dinoflagellates (Litaker et al. 1999, Oldach et al. 2000, Saldarriaga et al. 2001).

Despite these advances, aspects of the basic life history of cryptoperidiniopsoids are poorly known. During routine culturing we consistently have observed division in nonmotile cysts, and persistent sexuality evidenced by fusing flagellated cells (gametes) and resultant zygotes with two longitudinal flagella (planozygotes; Pfiester 1984, Pfiester and Anderson 1987). In addition, we have commonly observed prominent nuclear cyclosis prior to division in many reproductive cysts. Nuclear cyclosis is defined as a distinctive and protracted swirling of the chromosomes in dinoflagellate zygotic nuclei believed to

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coincide with homologue pairing prior to meiotic division (von Stosch 1972, 1973, Beam and Himes 1980, 1984, Pfiester 1984, 1989, Pfiester and Anderson 1987, Barlow and

Triemer 1988). Four objectives were addressed in this study: first, aspects of the life history of cryptoperidiniopsoids were described based on observations of seven strains, with supporting photography and flow cytometric DNA analyses. Second, a technique was described and tested to enhance population synchrony and diminish vacuolar DNA in these phagotrophic protists, and to simultaneously obtain pure populations. Details of the life cycle and population DNA profiles were compared to closely related species and dinoflagellates in general. Third, it has been suggested that an obligate bacterial- cryptoperidiniopsoid association may occur in cryptophyte-fed culture (Alavi et al. 2001).

Externally bacteria-free cryptoperidiniopsoids were used to further investigate that premise, as well as the potential for chemoheterotrophic growth on dissolved organic compounds by these holozoic microorganisms. Fourth, the potential for photosynthetic nutrition in cryptoperidiniopsoids through maintenance of ingested cryptophyte plastids was experimentally examined. This information is contributed to advance knowledge about heterotrophic dinoflagellates, and to assist future efforts to formally classify cryptoperidiniopsoids.

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4.3 Materials and Methods

4.3.1 Cultures. The seven cryptoperidiniopsoid isolates used in this study were obtained over the past ten years (Table 4.1). Most were isolated from estuaries by our laboratory; Dr. H. Marshall, (Old Dominion University, Norfolk, VA) kindly supplied others. Our isolation procedure was as follows: estuarine water or sediment samples containing small heterotrophic dinoflagellates were enriched (ca. 1:3 v/v) with cryptomonad culture as prey (Caron 1993), incubated in darkness, and checked periodically

(Burkholder et al. 2001). This simple method often selectively favored growth of cryptoperidiniopsoids (Seaborn et al. 1999, 2001). If dinoflagellates became abundant, serial pipette isolations and subcultures in 15-ppt salinity f/2-Si (Guillard 1975) containing cryptomonad prey were used to eliminate contaminating protists by exclusion and attrition

(Pringsheim 1964, Caron 1993). The cultures were maintained for months to years.

Prior to this study, multiple single-cell isolations were obtained from each potentially multi-species culture by flow cytometric cell deposition into 96 well microculture plates preloaded with media and cryptomonads (Glasgow et al. 2001). Thus, each culture was initiated from a single flagellated cell. The single cells were allowed to proliferate into microculture populations and then were closely examined for a cryptoperidiniopsoid-like shape and swimming pattern. Under these conditions, cryptoperidiniopsoids usually had an epitheca that was larger than the hypotheca, and a well-excavated descending cingulum with pronounced displacement.

Cryptoperidiniopsoids that congregated at the bottom of the culture vessel commonly swam in tight, clockwise, ventral-facing loops in the same plane as the vessel bottom (Fig. 4.1).

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This swimming pattern was not constant for all cells, but was a consistent, repetitive behavior (Kamykowski et al. 1992) and therefore useful as a preliminary identifier.

Distinctions under light microscopy, although imprecise, can be useful in prefatory identifications (Steidinger 1979). Based on these distinctions, clonal microcultures were selected, propagated, and tested for identity at least twice with 18s rDNA based molecular probes (PCR performed by Dr. P. Rublee, University of North Carolina – Greensboro,

Greensboro, NC and Dr. D. Oldach, University of Maryland, Baltimore, MD) using sequences considered to be specific for Pfiesteria piscicida Steidinger et Burkholder,

Pfiesteria shumwayae Glasgow et Burkholder, and a cryptoperidiniopsoid ‘brodyi’ taxon that has not been formally described (primer designed for specificity to Genbank

AF080097, Litaker et al. 1999, by Dr. D. Oldach) (Rublee et al. 1999, 2001, Bowers et al.

2000, Oldach et al. 2000 ).

Two of the seven isolates used in this study did not react positively with the cryptoperidiniopsoid ‘brodyi’ PCR primer, nor with primers specific for Pfiesteria spp.

(Table 4.1). These isolates were identical in shape and behavior to the others under LM, but may have been different cryptoperidiniopsoid species. Isolates were maintained in f/2-

Si media prepared with 15-ppt salinity, aged natural seawater (collected in the Gulf Stream ca. 2 km offshore from Cape Hatteras, NC) and fed in batch cultures with cryptophyte sp.

HP 9101 provided by Dr. D. Stoecker, Horn Point Laboratory, University of Maryland,

Cambridge, MD; initial prey density ≥ 105 cells·mL-1. Cultures were grown in polystyrene cell culture flasks stored horizontally at ca. 20 ºC under overhead fluorescent lighting (50

µmol photons·m-2·s–1) on a ~16-h : 8-h light:dark (L:D) cycle. Replicate cultures were maintained similarly in chamber microscope slides and petri dishes for imaging. High

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Figure 4.1. Depiction of a diagnostic swimming pattern commonly observed in these cryptoperidiniopsoid isolates. Flagellated cells that had congregated at the vessel bottom often swam in stationary, repetitive, clockwise ventral-facing loops.

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Table 4.1. List of the cryptoperidiniopsoid isolates examined, the geographical location from which they were obtained, isolation dates, PCR molecular identification, and the mean flagellated cell 1C DNA contents determined by flow cytometry.

Isolate designation Source location Isolation datea PCR I.D.b 1C DNA content

Cr. P.p. P.s. (CRBC units·cell-1)c

Florida Bay-AC1 Florida 08/1997 + - - 4.95

Charleston-AC1 South Carolina 06/1997 - - - 4.93

Chesapeake-543A Chesapeake Bay, MD 10/1999 + - - 4.81

Virginia-1070 Chesapeake Bay, VA 10/1999 + - - 4.42

NOAA-AC1 Pamlico River, NC 06/1997 + - - 4.78

Varsity-AC1 Raleighd, NC 05/1992 + - - 5.13

1253T-AC7 Neuse River, NC 05/2001 - - - 4.36

a Date (month/year) that the isolate source material was removed from nature. b Identification by PCR molecular probes (18S rDNA), performed at least twice for each isolate. Cr. = cryptoperidiniopsoid ‘brodyi’ (Genbank AF080097, see text), P.p. = Pfiesteria piscicida, P.s. = Pfiesteria shumwayae. c Peak fluorescence signals were collected for at least 104 cells for each 1C DNA mean determination, and DNA content was calculated relative to a chicken red blood cell (CRBC) standard signal mean. CRBC unit = 2.33 pg DNA·cell-1 (Vaulot et al. 1994, Veldhuis et al. 1997). d Varsity-AC1 was isolated from an inland aquaculture facility.

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feeding activity and rapid growth (Seaborn et al. 1999) typically led to cryptoperidiniopsoid densities of ca. 104 to 105 cells·mL-1 as the cryptoperidiniopsoids reduced cryptomonads to negligible numbers within 3-5 d, depending on the inoculation density. Cultures were transferred by decanting the spent volume and refilling with new cryptophyte culture, or by inoculating aspirated cells into cryptophyte culture. Starved cultures (maintained for days- months) were produced by not providing more cryptomonads after they were depleted.

These nonaxenic cryptoperidiniopsoids, which were not known to be toxic (Burkholder et al. 2001), were deposited at the Provasoli-Guillard National Center for the Culture of

Marine Phytoplankton, Bigelow, ME. A pure cryptophyte culture (HP9101) was prepared by gently ultrasonicating a dilute suspension of planktonic cells in an attempt to remove potentially attached bacteria (Popovský and Pfiester 1990), then flow cytometrically sorting

100 cryptophyte cells into sterile f/2-Si media. Once the population was in exponential growth, subcultures were subjected to the purification procedure of Droop (1967) using graded concentrations of a mixture of penicillin, streptomycin, neomycin, and amphotericin

B. The cryptomonad subculture that survived the highest concentration of antibiotics was tested for demonstrable contaminants (below).

4.3.2 Cyst Purification Treatments. Chemical treatments to eliminate bacteria while leaving cryptoperidiniopsoid cysts unharmed were tested using microscopical assays, wherein the abrupt termination of nuclear cyclosis was equated with cyst damage. In untreated cysts, unambiguous nuclear cyclosis lasted over 3 h and ended gradually. Abrupt cessation of chromosome movement upon addition of chemical treatments was used to indicate cytotoxicity. Subcultures grown in petri dishes were selected once reproductive cysts were abundant on the dish floor, rinsed and refilled with deionized water, and

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examined microscopically. Cysts undergoing nuclear cyclosis were located and chemical treatments were added while observing the cysts.

Short-term (≤10 min) effects of the general disinfectants sodium hypochlorite

(dilution of commercial 6% bleach) and hydrogen peroxide (dilution of a commercial 3%

H2O2 solution) on cyst viability were tested over a range of final concentrations. Both disinfectants caused rapid cell damage at fairly low concentrations, consistent with previous investigations (Ichikawa et al. 1993, Azanza et al. 2001). Sodium hypochlorite damaged cysts in under 10 minutes at concentrations > 0.004%, H2O2 at > 0.10%. Other signs of cellular damage such as contracted protoplasts and bleaching of ingested pigments were observed (Ichikawa et al. 1993). The effect of various antibiotics, alone and in concert, on cyst viability was also tested. The tested antibiotics did not alter cell morphology or inhibit nuclear cyclosis and subsequent division up to 104 IU penicillin·mL-1, 10 mg streptomycin and neomycin·mL-1, 25 µg amphotericin B, and 100 mg ampicillin and polymyxin B sulfate·mL-1. Since the goal was to promote a vigorous excystment after an extended, induced arrest (below), antibiotics were chosen over disinfectants to eliminate contaminants in the population synchronization and culture purification technique.

4.3.3 Population Synchronization and Tests for Contaminants. In these cryptoperidiniopsoid isolates, reproductive cysts typically adhered to the bottom of the culture vessels. The preferential lysis method of Parrow et al. (2002) was further developed to obtain a more synchronized excystment of flagellated cells.

All culture additions and transfers were done in a sterile positive-pressure laminar flow hood using aseptic techniques. For each isolate, subcultures were initiated with a 10- mL inoculum in 250-mL cell culture flasks stored horizontally and serially fed every 1-2

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days with a small inoculum of centrifuge-concentrated cryptomonads. After 2 weeks of vigorous feeding and reproduction, the cryptoperidiniopsoid populations were dense, with abundant reproductive cysts adhered to the culture vessel floor. The culture volume was decanted and the flasks were gently rinsed with sterile deionized water (Millipore

Corporation, Bedford, MA), then completely filled with sterile deionized water containing a strong dose (Guillard and Keller 1984) of antibiotics: penicillin (150 IU·mL-1), streptomycin (0.15 mg·mL-1), neomycin (0.10 mg·mL-1), amphotericin B (0.25 µg·mL-1), ampicillin (0.10 mg·mL-1), and polymyxin B-sulfate (0.10 mg·mL-1). Since β-lactam antibiotics (penicillin, ampicillin) act on dividing bacteria, 5.0 mL of sterility test medium

(below) prepared with deionized water were included in each flask (Droop 1967). Cysts were maintained in this state for 48 h. Within the first 12 h, reproductive cysts were observed to complete cytokinesis, but offspring did not excyst and the pigmented food vacuoles contained in many offspring grew noticeably smaller and paler.

After 48 h the deionized water was decanted and each flask was filled with 100 mL of 15 ppt-salinity sterile seawater containing penicillin (50 IU·mL-1), streptomycin (0.05 mg·mL-1), and neomycin (0.01 mg·mL-1). Flasks were returned to a horizontal position and observed for excystment. Within 3 h a robust excystment response had occurred in each flask and the swimming cells were decanted from the remaining adherent cysts.

Subsamples were taken from each flagellated cell suspension, preserved, and enumerated.

Excysted populations from each isolate were: (1) tested for the demonstrable presence of viable bacteria and fungi; (2) inoculated into flasks containing presumed axenic cryptophyte sp. HP9101; (3) preserved for DNA analysis; and (4) tested (3 of 7 isolates) for axenic population growth/survival in organically enriched media.

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The presence of bacterial and fungal contaminants was determined by direct culture in nutrient broth and on agar media. Nutrient broth was composed of 15 ppt-salinity f/2-Si enriched with bacto-peptone (1 g·L-1; Becton Dickinson and Co., Sparks, MD) and glucose

(1 g·L-1; Hoshaw and Rosowski 1973), autoclaved in culture tubes. Culture samples (1 mL) were aseptically inoculated into 20-mL broth tubes in duplicate and incubated in darkness at ca. 20˚C. Liquid cultures were checked daily for cloudiness as evidence of bacterial growth; un-inoculated broth tubes served as clarity controls. Cultures were examined microscopically (phase contrast, 750x) after 7-10 days for obvious abundance of bacteria- like particles (Guillard and Keller 1984). In addition, culture samples (0.5 mL) were also plated onto enriched 0.5x marine agar composed of Difco marine agar 2216 (27.6 g·L-1;

Becton Dickinson and Co.), agar (7.5 g·L-1), bacto-peptone (5 g·L-1), and yeast extract (1 g·L-1; Fisher Scientific, Fair Lawn, NJ) in deionized water. Culture plates were incubated in darkness at ca. 20˚C and checked daily for evidence of bacterial and fungal growth.

Cultures that did not elicit observable bacterial or fungal growth in either of these media and were without abundant bacteria-like particles were considered to be without demonstrable contaminants (Guillard and Keller 1984, Guillard 1995). Contaminant-free cultures were used to investigate feeding and development of cryptoperidiniopsoids in monoxenic cultures with cryptophyte prey toward assessing influences of extracellular bacteria on cultured cryptoperidiniopsoids. Purified cryptoperidiniopsoids were also inoculated into different organically enriched media formulations to see if the nutritional needs of the dinoflagellates might be fully or partially met through chemoheterotrophic use of dissolved compounds (Droop 1970).

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4.3.4 Sample Preparation for Microscopy and Flow Cytometry. For LM, cells were photographed live or after fixation in 0.5% glutaraldehyde (final concentration). As for

Pfiesteria spp., cryptoperidiniopsoid cysts were impermeable to passive penetration by nuclear stains. An earlier procedure to stain cyst DNA in Pfiesteria spp. required permeabilization of the cyst wall (Parrow and Burkholder 2002, Parrow et al. 2002). Here, a second in vivo staining procedure was developed. For qualitative observations and imaging of cyst nuclei, subcultures were incubated in darkness for ≥ 2 days after addition of an inoculum of cryptomonads laced with the live-cell nucleic acid stain (SYTO 11,

Molecular Probes Inc., Eugene, OR) to effect a 0.5-µM final concentration. The sublethal concentration incorporated into the DNA of flagellated cells (Parrow et al. 2002) and, thus, was also present in the nuclei of subsequent encysted cells. This allowed in vivo qualitative determinations (number, relative size) of fluorescent nuclei in cysts (Bhaud et al. 1991,

1994).

For DNA measurements in flow cytometry, dinoflagellate cells were prepared as in

Parrow et al. (2002). Briefly, 50 mL culture aliquots were filtered (09-795AA filters, 25

µm porosity; Fisher Scientific, Pittsburgh, PA) to remove debris and fixed with 0.5% paraformaldehyde (final concentration) overnight at 4ºC. Cells were then concentrated ten- fold by centrifugation (2000 rpm, 10 min) and stored in darkness at 4ºC. Prior to staining, aliquots were treated with RNase A (1 µg·mL-1 final [w/v], Sigma, St. Louis, MO) for 1 h at room temperature. Cells were then stained with 5 µM SYTOX Green (Molecular Probes,

Inc.) and returned to storage conditions for 12-14 h (Eschbach et al. 2001). Chicken red blood cells (CRBCs; Biosure, Grass Valley, CA) and fluorospheres (Coulter Corporation,

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Miami, FL) were used as DNA/fluorescence standards (Vaulot et al. 1994, Veldhuis et al.

1997, Larsen and Edvardsen 1998, Johnston et al. 1999).

4.3.5 Light Microscopy. Cells and life cycle transitions were observed, photographed, and video recorded using live subcultures contained in petri dishes or chamber slides. Routine culture examination was conducted using an Olympus CK40 inverted culture microscope (Olympus Corporation, Melville, NY) at 40-400x. For population enumeration, cells were preserved with acidic Lugol’s solution (Vollenweider

1974) and enumerated using a Palmer-Maloney counting chamber (Throndsen 1995). Cysts were each counted as one cell. An Olympus AX-70 microscope equipped with a water immersion 60x 0.9 NA objective, 0.8 NA condenser and a DEI-750 cooled chip CCD camera (Optronics Engineering, Goleta, CA) was used for imaging and videotaping.

Brightfield, differential interference contrast, phase contrast, and epifluorescence microscopy with mercury lamp excitation were used to image cells and stained nuclei. A

Zeiss Axiophot (Carl Zeiss, Inc., Thornwood, NY) equipped with a oil immersion 100x 1.4

NA objective, 1.4 NA condenser, a polarization modulation differential interference contrast (PM-DIC) system (Holzwarth et al. 1997), and a Hamamatsu Argus 20 B/W camera (Hamamatsu City, Japan) and image analysis system (Allen and Allen, 1983) was also used for high resolution, high contrast videotaping of nuclear cyclosis.

4.3.6 Flow Cytometry. Analyses and cell sorting were performed with a EPICS

Altra flow cytometer (Coulter Corporation, Miami, FL) using a 15 mW argon laser (488 nm) focused to an interrogation point 6 µm in height x 112 µm in width. Green fluorescence of the SYTOX stain was detected at 525 nm, and chlorophyll at 640 nm.

Forward angle and side angle light scatter were recorded as relative measurements of cell

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size/granularity (Shapiro 1995). Signal amplification and threshold levels were optimized so that pertinent signals fell mid-scale, and then were not altered. Integrated, peak and logarithmic fluorescence and scatter signals were recorded as appropriate on 104 to 105 cells per analysis, displayed using EXPO32 Analysis software (Applied Cytometry Systems,

Sheffield, UK), and stored in listmode format. Optical alignment and signal stability were monitored using 10-µm diameter fluorospheres (CVs < 2%) and linearity of the cytometer amplification system was checked using Immuno-Brite fluorospheres (Coulter Corporation)

(Bagwell et al. 1989). Electrostatic cell sorting was performed for purity in preference of yield, and electronic sort regions were defined on the EXPO32 interface. For relative DNA measurements, CRBCs were added to selected subsamples prior to staining. For each analysis, the mean DNA fluorescence of a minimum of 104 flagellated cells under the lowest DNA peak (assumed 1C) was compared to the CRBC signal mean, and dinoflagellate DNA values were expressed in CRBC units (Vindelov et al. 1983, Veldhuis et al 1997, Parrow and Burkholder 2002; Table 4.1). Minimization of vacuolar DNA was assessed based on coefficients of variation (CVs, relative standard deviation) for the DNA fluorescence peaks obtained.

4.3.7 Nutrition. To examine the potential for chemoheterotrophic growth on dissolved compounds and the possibility for axenic propagation of these dinoflagellates, purified cells from the synchronized excystments were inoculated into triplicate culture flasks containing three different formulations of organically supplemented media (Table

4.2) and aged, sterile seawater as a presumed metabolite-free control. Chemical supplements were added to 15-ppt salinity f/2-Si prepared with aged natural seawater, and ultrafiltered (0.22 µm). Supplements for MLH1 were adapted from a formulation used to

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culture the free-living heterotrophic dinoflagellate Crypthecodinium cohnii (Tuttle and

Loeblich 1975). PKN ingredients were adapted from a medium used to propagate the parasitic apicomplexan Perkinsus marinus Levine (Coss et al. 2001). HDM was a modification of media used in culturing the parasitic dinoflagellates Amyloodinium ocellatum Brown & Hovasse (Oestmann and Lewis 1996) and Hematodinium sp. (Appleton and Vickerman 1998). Cultures were maintained in darkness at ca. 20 ˚C and microscopically examined twice daily for qualitative assessment of population vitality and evidence of cell proliferation. Subsamples for enumeration were removed after gentle mixing initially and at 2-day intervals thereafter.

To examine the potential for mixotrophic nutrition, cultures from three cryptoperidiniopsoid isolates with abundant flagellated cells feeding on cryptomonad prey were analyzed live using flow cytometry. Optical signals from dinoflagellates with autofluorescent inclusions (pigment-replete food vacuoles) were identified on bivariate plots of chlorophyll autofluorescence (640 nm) versus side angle light scatter, the same parameters used by Weisse and Kirchhoff (1997) to investigate feeding of Peridiniopsis berolinense (Lemmermann) Bourrelly on cryptophytes. Once identified, ca. 1.2 x 105 dinoflagellates with fluorescent inclusions from each culture were electrostatically sorted into f/2-Si media, thereby isolating them from cryptomonad prey cells. The sorted dinoflagellates were examined microscopically; all cells contained pigmented food vacuole(s) and no cryptomonad cells were observed. Each suspension was then split into six culture flasks containing 20 mL of f/2-Si media to effect final densities of ca. 1.0 x 103 cells·mL-1. Triplicate light and dark treatments were made by placing the cultures in an algal growth incubator under continuous cool white fluorescent lighting (150 µmol

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photons·m-2·s-1) for 48 h, with the dark treatment contained in a lightproof box next to the light treatment. Cultures were microscopically examined daily, and subsamples were removed for cell enumeration after gentle mixing. After 48 h, representative cultures from light and dark treatments were analyzed for autofluorescence by flow cytometry using the same instrument settings and parameters.

4.3.8 Data Analyses. Flow cytometric listmode data were plotted as univariate signal height distributions or bivariate frequency distributions using EXPO32 Analysis software. Potential doublets in DNA stained cells were discriminated by plotting DNA fluorescence signal peak height versus integral signal area, under the assumption that DNA doublets deviate from a linear ratio of the two signals (Wersto et al. 2001, Parrow et al.

2002). DNA-fluorescence intensity histogram deconvolution to yield peak numbers (area), means, coefficients of variation (CVs), and peak ratios were computed using MultiCycle

DNA Content and Cell Cycle Analysis Software (Phoenix Flow Systems, Inc., San Diego,

CA), substituting G1 and G2 phase descriptions with 1C and 2C, respectively. Tukey’s studentized range test was used to compare 1C DNA means (α = 0.01), and single factor

ANOVA (α = 0.05) was used to test for differences among sample and treatment means

(SAS Institute, Inc., 1999).

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Table 4.2. Media formulations tested for axenic growth / survival. The chemical supplements listed were added to 15-ppt salinity f/2-Sia prepared with diluted aged natural seawater, and ultra-filtered (0.22 µm). Percentages are final volumes unless otherwise specified.

MLH1 PKN HDM

d Na2Glycero-PO4 (0.8 mM) F12 (40%) FBS (10%)

e (NH4)2SO4 (1.5 mM) DME (20%) L-Glutamine (1%, w/v)

Na Acetate (15 mM) FBSf (5%) HEPES (10 mM)

Histidine·HCl (0.8 mM) HEPES (10 mM) Salinity 15, pH 7.0 b

D-Glucose (22 mM) Salinity 15, pH 7.5 b Antibioticsc

Betaine·HCl (9.7 mM) Antibioticsc

Tris (16 mM)

Salinity 15, pH 7.0 b

Antibioticsc

a f/2-Si contains inorganic nutrient salts, trace metals, biotin, thiamine, and vitamin B12. b Salinity adjusted with addition of synthetic sea salt (Instant Ocean, Aquarium Systems, Mentor, OH) or deionized water; pH adjusted using NaOH or HCl. c 25 IU penicillin·mL-1, 25 µg streptomycin·mL-1, 50 µg neomycin·mL-1 (dilution of Sigma P4083). d Nutrient mixture Ham’s F-12 (Sigma N4888). e Dulbecco’s Modified Eagle’s Medium (Sigma D1152). f Fetal bovine serum (Sigma F2442).

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4.4 Results

4.4.1 Life History. The cryptoperidiniopsoids used in this study were easy to culture and required only frequent feeding to maintain dense populations. For example, one culture was rejuvenated overnight by addition of dense cryptomonad prey (ca. 2.0 x 105 cells·mL-1) after > 6 months of storage in darkness at 4 ºC. An isolate cultured for 10 yr was subjected to repeated, prolonged starvation, but still exhibited similar cell production as newer isolates.

The seven cryptoperidiniopsoid isolates had the same morphology and behavior under LM. Two general phases were observed, motile (flagellates) and nonmotile (cysts), the proportions of which were dependent upon prey availability. Thecal plates were not visible using standard LM. Young (recently excysted, pre-feeding) biflagellate cells were typically colorless and irregularly ovate, with a sinistral descending girdle displacement of around 1.0-1.5 girdle widths (Fig. 4.2A). However, shape varied considerably in well-fed cultures as flagellated cells ranged in length from ca. 6-18 µm, apparently depending on the amount of ingested material. The relatively large nucleus was in the hyposome, food vacuoles occurred primarily in the episome, and small refractile lipid bodies were typically scattered throughout the cytoplasm.

Flagellated cells fed by attaching to cryptomonads with an extensible tube-like organelle, the peduncle, which emerged from the sulcus and through which prey cytoplasm was conveyed into episomal food vacuoles (Seaborn et al. 1999). In dense culture, cryptomonads that had been compromised by the peduncle of one dinoflagellate often appeared to attract others in the immediate vicinity, resulting in small aggregations that surrounded the compromised prey cell and consumed it as a group (as in Spero and Morée

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1981, Ucko et al. 1997). Up to four small cryptoperidiniopsoids were observed feeding on a single cryptomonad, wherein the pigmented cytoplasmic constituents of the prey cell passed into each feeding dinoflagellate. Larger cryptoperidiniopsoids often aspirated the cytoplasmic contents of each cryptomonad in a more singular action (video link, myzocytosis.mov). About four cryptomonad protoplasts were maximally ingested by a single cryptoperidiniopsoid. The prey-filled vacuoles of feeding cryptoperidiniopsoids typically aggregated into a single irregular-shaped episomal pigmented mass. Attempted cannibalism was observed, particularly on cells in the process of excystment when cryptomonad prey was scarce and flagellated cells were still abundant. When it occurred, a concerted attack by flagellated cells on a cyst was a good indicator that excystment was imminent. These attacks appeared to be directed toward some solute released during excystment, and were rarely successful.

In these isolates, cell division occurred in thin walled nonmotile cells (reproductive temporary cysts; Beam and Himes 1980, Popovský and Pfiester 1990, Anderson et al.

1995). Ecdysis was not observed during encystment. Encystment was rapid (seconds) and occurred by loss of the flagella, secretion of an imperceptible wall or other modification of the outer layer that increased resistance, and rounding of surface features. Encystment appeared to be an active process where flagellated cells swam along close to the culture vessel floor, made repetitive tight looping motions, and encysted suddenly. Patchy aggregations of cysts were common, suggesting active selection of encystment location as described for some other dinoflagellates (Lombard and Capon 1971, Ucko et al. 1997,

Parrow et al. 2002). As with flagellated cells, cryptoperidiniopsoid cysts varied considerably in size and amount of ingested material (Fig. 4.2B). When cell division

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occurred, the chromosomes expanded and underwent bipartition, and then cytoplasmic division followed (Fig. 4.2C). The outer cyst wall was not included in the division. When present, the pigmented food vacuole(s) were compressed into a single shrinking body situated equatorially in the division cyst (Fig. 4.2C). Two offspring cells resulted (Fig.

4.2D), each with a single nucleus (Fig. 4.2E). Release of two biflagellate offspring typically occurred 3-4 h after nuclear division. Offspring cells moved through a fissure in the translucent cyst wall and often remained attached to it by the distal ends of their longitudinal flagella for a few moments before swimming away. The food body, inherited by one offspring cell, was often quickly egested as reported for Peridiniopsis berolinense

(Calado and Moestrup 1997).

Reproductive cysts were rarely found in which the first two offspring protoplasts had distinctly rounded into oval cells, each apparently enclosed in its own membrane internal to the progenitor (primary) cyst outer wall (i.e. secondary cysts; Elbrächter and

Drebes 1978, Drebes and Schnepf 1988, Parrow et al. 2002) (Fig. 4.2F). Four flagellated cells (two from each secondary cyst) were released; thus, two consecutive cell divisions had occurred. Another general distinction was apparent between cysts that contained pigmented food bodies and those that did not (Fig. 4.2G). Colorless cysts were typically smaller than pigmented cysts in cultures with abundant prey (Fig. 4.2G). Initially they contained a single nucleus (Fig. 4.2G), but were also capable of protoplast division. Long term (weeks) prey-depleted cultures had few flagellated cells and many small colorless cysts (Fig. 4.2H).

Fusion of flagellated gametes was common in all isolates whether prey were plentiful or scarce. Paired gametes were equal or slightly different in size and usually colorless, although in well-fed cultures one of the pair often had a pigmented food body

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Figure 4.2

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Figure 4.2. Light micrographs (LMs) of cryptoperidiniopsoids. (A) A biflagellated cell without pigmented inclusions (ventral view). (B) Pre-division reproductive cysts with pigmented food vacuoles. (C) Reproductive cysts – the central cell is shown after nuclear division and partial cytoplasmic division, and the pigmented food body had migrated to the center early in division. The outer two cysts had not yet begun division. (D) Reproductive cyst in late development, with the offspring pair distinguished by a full line of division and an unequal inheritance of the pigmented food body. A slight contraction of the left cell revealed the surrounding thin cyst wall (arrow). (E) DNA-stained division cyst, revealing two distinct nuclei (green). (F) Reproductive cyst that had undergone two consecutive divisions, resulting in four offspring (two from each secondary cyst). All divisions were internal to the primary cyst outer wall. (G) Two common cyst types: with pigmented food vacuole (left, for comparison); and without prey pigments but full of refractile bodies

(right). Non-pigmented cysts were also capable of cell division. Inset: nuclear staining of a colorless cyst revealing a single nucleus. (H) A small, spherical quiescent cyst full of chromosomes, from a prey-starved culture. Scale bars = 10 µm.

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(Fig. 4.3A). Gametes oriented roughly perpendicular to one another and fusion began with the episome of one gamete fusing laterally into the episome (most common) or hyposome of the other (Fig. 4.3B,C). As fusion proceeded, the united pair continued swimming in characteristic clockwise pinwheel loops (video link, gametefusion.mov). Two trailing flagella were apparent as plasmogamy neared completion (Fig. 4.3D). Plasmogamy was typically complete in < 30 minutes. The resulting planozygote had a typical dinoflagellate shape (anterior episome, cingulum, and posterior hyposome) and two longitudinal flagella

(Fig. 4.3E).

Planozygotes were initially small (Fig. 4.3F) and highly motile, and thus escaped continuous observation, as described for some other small heterotrophic species (Beam and

Himes 1974, Drebes and Schnepf 1988). Planozygotes were found feeding on cryptomonads and those with prey-replete food vacuoles were larger (Fig. 4.3G).

Following ingestion and growth, planozygotes presumably became nonmotile by forming zygotic temporary cysts (von Stosch 1972, Beam and Himes 1980, Kita et al. 1993).

Zygotic cysts were morphologically indistinguishable under LM from other reproductive cysts, except by the occurrence of nuclear cyclosis (as in Spero and Morée 1981, Drebes and Schnepf 1988, Ucko et al. 1997, Parrow et al. 2002). During nuclear cyclosis the nucleus was rounded and the elongate string-like chromosomes swirled vigorously for > 3 h

(Fig 4.3H; video link, nuclearcyclosis.mov). The direction of rotation was typically uniform, although turbulent mixing of the chromosomes also occurred. One circulation minimally required ca.10-20 s, and near completion the speed and uniformity of the swirling gradually diminished. This striking phenomenon was easily followed by direct observation of living cysts, and occurred commonly in all seven isolates.

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A single division followed nuclear cyclosis (Fig. 4.3I,J), indistinguishable under LM from division in cysts that did not undergo nuclear cyclosis. Two biflagellate offspring were released from zygotic cysts 3-5 h after nuclear cyclosis ceased (Fig. 4.3K; video link, meiosis.mov). The entire process of nuclear cyclosis followed by division and release of two offspring dinoflagellates was confirmed in four or more examples from each isolate.

The two offspring appeared uninucleate, which was confirmed (n = 3 post-cyclosis excystments, randomly selected among the 7 isolates) by in situ fixation of the flagellated cells at the moment of excystment followed by DNA staining (Fig. 4.3L). The in vivo DNA staining technique successfully stained live cyst nuclei (Fig. 4.2E, G), and cell division occurred in cysts with fluorophore-bound nucleic acids (Fig. 4.2E). Nuclear cyclosis also occurred in stained cysts, although the process was rapidly terminated by fluorophore excitation.

Not all cryptoperidiniopsoid cysts were reproductive; excystment of single flagellated cells was also observed. Depletion of prey cells in culture led to increasing numbers of cysts and fewer flagellated cells. In cryptoperidiniopsoid cultures that were starved for long periods (weeks to months), cysts predominated and flagellated cells became rare. These cysts were small and colorless, and occurred in abundance adhered to the culture floor. It was unknown whether they were formed asexually or sexually.

However, a feeding population was quickly revived after addition of a dense inoculum of prey cells (ca. ≥ 105 cells·mL-1), whether following 1 month at room temperature or 6 months in cold storage. This rapid activity suggested that some if not all cysts formed under prey limitation were quiescent, suspended from growth by an unfavorable

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Figure 4.3

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Figure 4.3. LMs of cryptoperidiniopsoid sexual stages. (A) Paired gametes, one with a pigmented food vacuole. (B) Gamete pair in mid-fusion. C-E, series of observations. (C)

Non-pigmented gametes in mid-fusion. (D) Late gamete fusion, with two longitudinal flagella visible at the cell posterior. (E) Resultant planozygote, with two trailing flagella.

(F) Young planozygote prior to feeding. (G) Mature planozygote with a pigmented food vacuole and an enlarged episome. H-L, series of observations. (H) Cyst during nuclear cyclosis (nucleus, arrow). (I) Subsequent nuclear division, with equatorial placement of the food vacuole. (J) Protoplast division. (K) The two flagellated offspring and the remaining cyst wall (arrow). The upper dinoflagellate retained the residual food body. The lower dinoflagellate (having just emerged amorphously from the cyst) had not yet attained a typical shape. (L) In situ fixation and nuclear staining of the excysted pair, showing a single nucleus in each cell. Scale bars = 10 µm.

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environment (i.e. no food) and not truly ‘resting’ or dormant in the context of Anderson et al. (1995).

4.4.2 Population Synchronization. As for Pfiesteria spp., cryptoperidiniopsoid cysts were apparently undamaged by replacement of the saline culture media with deionized water, but remaining flagellated cells and prey cells lysed from osmotic stress.

Furthermore, reproductive cysts in deionized water were developmentally unaffected and underwent cell division as normal, yet the offspring cells apparently were blocked from excystment by the osmotically unfavorable medium. Thus, deionized water did not inhibit cytokinesis in cysts but arrested excystment. Submersion in deionized water also did not inhibit nuclear cyclosis.

Cryptoperidiniopsoid cells did not appear harmed by the 48 h arrest in deionized water with strong antibiotics. The length of arrest was considerably longer than the time interval required for normal cell division and excystment, presumably forcing all reproductive cysts to reach the same developmental state and halt (with excystment imminent). This premise was supported by the vigorous excystment (ca. 60–90% of cysts) that occurred in each treated isolate within 3 h of re-exposure to the previous culture salinity. The resulting flagellated cell densities depended on cyst abundance and excystment rate, and ranged from ca. 2.7 x 104 to 1.5 x 105 flagellated cells·mL-1, with a mean ± SD of ca. 7.0 x 104 ± 4.0 x 104 flagellated cells·mL-1. Excysted cells were colorless and rapidly swimming. Sexual fusions were common among the recently excysted dinoflagellates, indicating that some were gametes, and diploid planozygotes were rapidly formed within the populations. Attempted cannibalisms on excysting cells were observed

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during the 3 h excystment period prior to isolation of the flagellated cells, but were mostly unsuccessful as previously described.

4.4.3 Contaminant Tests. Contaminant-free cultures were necessary for comparison between pretreatment cultures with bacteria and those without, as well as investigation of axenic (in organic media) cryptoperidiniopsoid growth. Following the purification treatments, bacterial or fungal contaminants were not detected in four of the seven populations. For the other three isolates, both duplicate broth tubes clouded visibly within

5-7 days. Microscopic observation (phase contrast, 750x) of the contaminated broth showed single and chain-forming rod shaped bacteria (bacilli). Treatments that showed neither contaminant growth nor subsequent abundant bacteria-like particles were considered contaminant free (Guillard and Keller 1984). A purified cryptophyte culture passed the same tests and was considered axenic.

Cryptoperidiniopsoids showed little growth when the purified cells were inoculated into axenic cryptomonad culture, supporting the hypothesis of Alavi et al. (2001) that certain bacteria may interact directly with the dinoflagellates in some beneficial way. In this study, the purified dinoflagellates were normal in shape and swimming habit, but did not appear to feed on the cryptophytes as robustly as in pretreatment cultures. After one week, the cryptoperidiniopsoid populations were primarily small colorless cysts, as occurred in starving cultures with plentiful bacteria. However, sporadic dinoflagellates with pigmented food vacuoles were seen in all cultures, so limited feeding did occur. The three cryptoperidiniopsoid isolates with viable bacilli in the test medium also did poorly under these conditions. Cultures were placed in darkness to stop cryptomonad photosynthesis and after several days the occurrence of feeding dinoflagellates improved

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somewhat, but the overall feeding response was still considerably less than pre-treatment cultures. Subsamples were inoculated into flasks containing aged, untreated (+ bacteria) cryptophytes, and the dinoflagellates began vigorous feeding immediately. Within 1 h of inoculation, large reproductive cysts with pigmented food vacuoles were found, and the dinoflagellate populations developed as in pre-purification cultures. In separate bacteria- free preparations, feeding was also improved if the dinoflagellates were provided axenic cryptomonads that had been briefly microwaved long enough to turn them from red to green, stop motility, and partially burst some cells.

4.4.4 DNA Analyses. The 48 h deionized water treatment eliminated algal prey and limited the cryptoperidiniopsoid populations to cysts. Pigmented vacuoles, contained in many cells, were much smaller and pale in color by the end of the arrest period. Since the majority of the released cells were the direct products of cell division without the opportunity to feed, presumably they contained the minimal DNA content for the isolate

(1C, G1 cell cycle phase; but sexual fusions had been common prior to fixation). The synchronized excystment yielded clean flagellated cell suspensions of suitable density for repeated flow cytometric analyses. When analyzed, each of the cryptoperidiniopsoid isolates exhibited a similar DNA content (Table 4.1) and a bimodal flow cytometric DNA distribution, consisting of abundant 1C cells and less abundant 2C cells (Fig. 4.4). The 1C

DNA subpopulations appeared to be comprised of biflagellate vegetative cells and gametes, whereas the 2C DNA subpopulations likely were comprised mostly (some cells may have excysted with 2C DNA) or entirely of planozygotes that formed during the 3h post- excystment period prior to fixation. Among isolates, 1C peak CVs ranged from 7.3-8.4% and the calculated percentages of cells with 1C DNA ranged from ca. 80-92%. The

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percentages of cells with intermediate DNA content (S-phase, DNA synthesis) ranged from

0 (3 isolates) to 5%, and were considered artifacts or the result of vacuolar DNA from cannibalism after the 48 h arrest period. The ratio of 2C/1C peak means for each isolate was consistently 1.8, which is within the range of values reported in whole-cell ploidy investigations of protists (Edvardsen and Vaulot 1996, Green et al. 1996, Larsen and

Edvardsen 1998, Radke et al. 2001). We have consistently observed similar lowered (<

2.0) 2C/1C ratios in DNA analyses of other heterotrophic dinoflagellates. The effect may be due to a small change in the conformational state (increased chromosome compaction) of dinoflagellate 2C nuclei (Darzynkiewicz et al. 1977, Partensky et al. 1991), although data dispersion may also be an influence (Watson 1977).

Slight variability (but significantly different high and low values) in DNA content was measured among the cryptoperidiniopsoid isolates. Isolates with higher measured

DNA content had been in culture longer (correlation between DNA content culture age, r2 =

0.72; Fig. 4.5), but data were not available for isolates through time. The mean cryptoperidiniopsoid 1C DNA (averaging isolates) was also significantly different from means calculated using previously published values for closely related species P. piscicida and P. shumwayae (Parrow and Burkholder 2002; Fig. 4.6). The cryptoperidiniopsoid 1C

DNA values also had lower variance, attributed to reduced vacuolar DNA as a result of the arrest pretreatment.

4.4.5 Growth On Dissolved Organic Compounds, and Mixotrophy. Dinoflagellate populations inoculated into organically enriched culture media without bacteria initially were similar across treatments, and a diminishing frequency of sexual fusions was observed. Significant differences between treatments and isolates developed over the six-

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day incubation (Fig. 4.7). Short-term maintenance of flagellated cells was higher in HDM and PKN media than in MLH1 and seawater. Flagellated cell numbers in MLH1 and seawater treatments declined precipitously by day 4, and small colorless cysts increased.

Flagellated cells remained more abundant in HDM and PKN media, and many of the cysts that formed also appeared slightly larger and darker in color compared to those in MLH1 and seawater treatments. By day 6, although significantly higher flagellated cell abundance occurred in HDM and PKN media in comparison to MLH1 and seawater, the complete nutritional needs of the cryptoperidiniopsoids apparently were not met under these conditions, as none of these media supported significant population growth. Furthermore, subcultures placed in fresh media did not proliferate.

In the mixotrophy experiment, flagellated cells containing chlorophyll were identified flow cytometrically and isolated from cryptomonad prey by cell sorting (Fig.

4.8A). Initially all dinoflagellates isolated from cryptomonads had conspicuous red pigmented food inclusions. After 24 h, however, flagellated cells with pigmented inclusions were rare and many colorless cysts occurred in both light and dark treatments.

By 48 h no cells in either treatment were found with pigmented inclusions, and flow cytometric reanalysis verified that remaining cells did not contain detectable photopigments

(Fig. 4.8B). The light and dark treatments also did not differ over the short-term experiment in terms of flagellated cell numbers (Fig. 4.9), and no obvious differences in cell biomass or cyst morphology and abundance were apparent.

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Florida Bay-AC1 Charleston-AC1 Chesapeake-543A 1C 1C 1C 1C = 85 % 1C = 84 % 1C = 86 % CV = 7.5 CV = 7.4 CV = 7.7

2C 2C 2C beads beads beads

Virginia-1070 NOAA-AC1 1C 1C Varsity-AC1 1C 1C = 89 % 1C = 80 % 1C = 92 % CV = 8.1 CV = 8.4 CV = 7.3

Events beads 2C beads 2C beads 2C

1253-AC7 1C 1C = 91 % CV = 7.7

beads 2C

Relative DNA (AU)

Figure 4.4. Flow cytometric DNA profiles of cryptoperidiniopsoid flagellated cell populations, following a 48 h forced arrest and subsequent synchronized excystment.

Fluorescence intensity standards were 10-µm fluorescent beads. Each isolate had a similar

DNA content and distribution pattern, consisting of 1C and 2C relative DNA subpopulations. Percentages of cells with 1C DNA and 1C peak CV’s are listed for each isolate. Distribution analyses were performed on linear signals from the same datasets with

Multicycle DNA analysis software, using automatic software DNA peak detection and least squares curve fitting. AU = arbitrary units.

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5.3

5.2 5.1 5 4.9

4.8 r2 = 0.72 4.7 4.6

1C (CRBC units) DNA 1C 4.5

4.4 4.3 0246810 Years in Culture

Figure 4.5. Measured 1C DNA content versus years in culture for the cryptoperidiniopsoid isolates examined. The trend line coefficient of determination is 0.72 and data points indicate mean values ± standard deviations computed from linear 1C peak CVs, and normalized to CRBC units.

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8 n = 9

n = 7 5

n = 14 Mean 1C DNA (CRBC units) (CRBC DNA 1C Mean 3

2 P. piscicida cryptoperidiniopsoids P. shumwayae

Figure 4.6. Mean 1C DNA comparison between the cryptoperidiniopsoid isolates examined and previously published values for Pfiesteria piscicida and P. shumwayae isolates (Parrow and Burkholder 2002). Bars represent ± 2 standard errors (SEs) and (n) equals the number of different isolates available for each mean determination. Measured means were different (p < 0.01).

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12 Florida Bay-AC1 11 HDM 10 PKN MLH1 9 Seawater 8 7 6 * 5 4 3 2 1 • 0 8 Chesapeake-543A -1 HDM 7 PKN mL •

3 6 MLH1 Seawater 5 4

3 * 2

1 Flagellated Cells x 10 Flagellated Cells x • 0 7 1253T-AC7 HDM 6 PKN 5 MLH1 Seawater 4

3 2 * 1 0 • 0123456

Time (days)

Figure 4.7

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Figure 4.7. Time-course growth / survival responses of three different axenic cryptoperidiniopsoid isolates in various culture media. HDM, PKN, and MLH1 media were supplemented with different additions of organic nutrients, vitamins, amino acids, and serum (formulations given in Table 4.2). Sterile seawater (aged, salinity 15) was considered a metabolite-free negative control. Points represent replicate (n=3) means ± 2

SEs. Test conditions at time interval 6 with different symbols are significantly different (p

< 0.01).

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T = 0 h A T = 48 h B cryptoperidiniopsoids

cryptoperidiniopsoids

Side light scatter cryptomonads

Red autofluorescence (chl a, AU)

Figure 4.8. Representative flow cytometric logarithmic bivariate histograms of side angle light scatter versus red autofluorescence (chla). Each data point denotes 1 cell. (A) Time =

0, a mixed culture containing cryptomonad prey cells and feeding cryptoperidiniopsoids.

The feeding dinoflagellates exhibited higher side angle light scatter properties and could be distinguished from the cryptomonad cells. The boxed region demonstrates the sorting parameters used to select dinoflagellates with red autofluorescence of similar intensity to that of the algal prey. The dinoflagellates meeting these criteria were sorted into collection vessels containing f/2 media. (B) The sorted populations, reanalyzed under the same parameters 48 h later exhibited no detectable chla autofluorescence and therefore clustered at the y-axis. AU = arbitrary units.

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1000 1253T-AC7 900

800 700

600 500 400 Light 300 Dark

200 1200 NOAA-AC1 -1 1100

mL mL 1000 • 900 800

700 600 500 Light

Flagellated Cells 400 Dark 300 1200 Varsity-AC1 1100 1000 900

800 700 600 Light 500 Dark

400

Time (days) Figure 4.9

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Figure 4.9. Time-course phototrophic growth / survival responses of three different cryptoperidiniopsoid isolates in light and dark treatments. Flagellated cells with red autofluorescent inclusions (algal prey chlorophyll) were flow-cytometrically detected and sorted from cultures containing algal prey, and were maintained in f/2 media with light and dark treatments. No significant differences in population growth or survival were detected between treatment means. Points represent replicate means ± 2 SEs (n = 3).

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4.5 Discussion

4.5.1 Life History. Our observations on the life history of these cryptoperidiniopsoid isolates under the described culture conditions are summarized in Fig.

4.10. Biflagellated dinoflagellate cells encysted to divide, typically producing two offspring flagellated cells (Fig. 4.10-1,2,3). Some cysts did not divide and yielded a single flagellated cell, indicated by the double arrow between Fig. 4.10-1 and 10-2. In other cysts, two consecutive divisions occurred, producing four offspring (Fig. 4.10-4). Development of such cysts with evidence of meiosis (nuclear cyclosis) was not observed, but consecutive divisions while encysted probably occur both asexually and sexually (below). Gametes were produced in cysts, and fused to form planozygotes with two trailing flagella (Fig.

4.10-3,5,6,7). Planozygotes encysted to divide, and a pronounced nuclear cyclosis preceded a single division and release of two uninucleate, biflagellate offspring cells (Fig.

4.10-8, 9). It was unknown whether these offspring required a second postzygotic division

(sister chromatid segregation) before reentering mitotic interphase, as conventional meiosis would predict (Fig. 4.10-10).

The seven isolates examined here followed this pattern. Although the isolates were each initiated with a single cell, it is not yet possible to determine with certainty that sexual reproduction was homothallic. Persistent sexuality occurred, and the size range of planozygotes overlapped presumed haploid cells. Therefore, it remains a possibility that the single cells that were isolated could have been diploid, and carried with them heterozygous sex determinants as considered for Pfiesteria by Parrow et al. (2002). If allelic, these determinants would segregate at meiosis, yielding two opposite mating types in a culture initiated by a single cell. Such a culture would not technically be clonal

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4 6 5

7 1 3 10

N 2 9 8

Figure 4.10. Schematic from our observations of these strains for the life cycle of cryptoperidiniopsoids under the culture conditions used, based on flagellar and nuclear characters (gray ovals represent nuclei). The broken arrow indicates an inferred relationship. (1) Vegetative biflagellated cell. (2) Temporary cyst or quiescent cyst. (3)

Division cyst. (4) Division cyst with two consecutive divisions, producing four flagellated offspring. These probably occur in both the asexual and sexual cycle, following (3) or (9)

(see text). (5) Paired gametes. (6) Fusing gametes. (7) Planozygote. (8) Zygotic cyst with nuclear cyclosis. (9) Division after nuclear cyclosis. In each case (n > 30), two flagellated cells were released after nuclear cyclosis and subsequent cell division. (10) It was unknown whether a second postzygotic division was required prior to re-entry into mitotic interphase (see text).

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(Anderson et al. 1995). Thus, there remained the possibility for heterothallism, as well as physiological anisogamy in morphological isogamy (Chesnick and Cox 1989) in these cultures.

The life history of cryptoperidiniopsoids was strongly influenced by prey availability. When prey was abundant, feeding, mitosis, sexual fusion, and meiosis occurred. As prey were depleted, the flagellated cells became smaller on average and more commonly colorless. Dorsoventrally flattened flagellated cells were often found in starving populations. Prey exhaustion led to a significant reduction in flagellated cells and the appearance of abundant colorless, apparently quiescent cysts. Sexuality was common and persistent in all growing cultures. Both gametes and most recently excysted cells were smaller than mature feeding cells, so it was uncertain whether gametes were fully differentiated or had vegetative potential. All fusions were roughly isogamous.

Planozygotes were commonly phagotrophic, and then presumably encysted prior to division. Cysts with nuclear cyclosis were considered zygotic and subsequent division meiotic. Rarely, encysted cells underwent two consecutive divisions, releasing four flagellated cells. These probably resulted from flagellated cells that had acquired sufficient endogenous reserves for the extra division. Cysts that produced four offspring after nuclear cyclosis were not observed, but it is probable that cryptoperidiniopsoid zygotes with sufficient cytoplasmic reserves also undergo consecutive divisions.

Zygotic cysts that release two flagellated cells after nuclear cyclosis and a single division occur in several other dinoflagellates, such as the heterotrophic species

Crypthecodinium cohnii (Ucko et al. 1997), and Pfiesteria piscicida (Parrow et al. 2002), and the photosynthetic species Helgolandinium subglobosum von Stosch (von Stosch

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1972), Woloszynskia apiculata von Stosch (von Stosch 1973), and Alexandrium hiranoi

Kita & Fukuyo (Kita et al. 1993). In C. cohnii and W. apiculata release of more than 2 flagellated cells after nuclear cyclosis and subsequent divisions occurred less frequently, and similar variability is expected in cryptoperidiniopsoids and Pfiesteria spp. The first and second divisions of dinoflagellate zygotes are often separated by an interval similar to vegetative growth, and conventional meiosis might not occur in all dinoflagellates (Beam and Himes 1975,1980,1984, Raikov 1982, 1995, Pfiester 1984, 1989). Specific measurement of the DNA content of zygotic cyst nuclei and the offspring produced after division would help clarify meiosis in cryptoperidiniopsoids and similar species.

In other research (Seaborn et al. 1999, 2001, Marshall et al. 2000), cryptoperidiniopsoid cells have been described with amoeboid and chrysophyte-like morphologies similar to those described for Pfiesteria spp. (reviewed in Burkholder et al.

2001, Glasgow et al. 2001, Burkholder and Glasgow 2002). Thus, determination of the overall life history of cryptoperidiniopsoids requires further study and differences in behavior and morphology apparently occur depending on the isolate and environmental conditions. Sexual reproduction in these phagotrophic species differs from the classic pattern of dinoflagellate sexuality largely developed from photosynthetic species (Pfiester

1984, Walker 1984, Pfiester and Anderson 1987, Pfiester 1989), wherein sexual fusion is induced by nutrient depletion and results in a dormant hypnozygote. Other factors may contribute to expression of sexuality in heterotrophic species, such as prey type or abundance. Quiescence may be favored over true dormancy in non-over wintering marine heterotrophic species as an adaptational response to fluctuant prey availability.

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Cryptoperidiniopsoids feed on a variety of microalgae (Seaborn et al. 1999) as well as the tissues of fish (Burkholder et al. 2001). They also feed on human blood cells and injured microbial metazoa (Eriksen et al. 2002, our unpubl. data). Like K. fungiforme,

Pfiesteria spp., and Peridiniopsis berolinense, cryptoperidiniopsoids are generalist cytoplasm feeders that appear especially attracted through chemotaxis to physiologically stressed, damaged, or leaky organisms (Spero and Morée 1981, Calado and Moestrup 1997,

Burkholder et al. 1998, 2001a, Seaborn et al. 2001). Myzocytotic phagotrophy may be advantageous in prey-rich, shallow habitats where such dinoflagellates can feed on any organism of roughly equal size or larger, as long as the prey has a penetrable wall or an open wound (Spero 1979, Gaines and Elbrächter 1987). Cryptoperidiniopsoids would be expected to thrive in shallow estuarine waters near the sediment feeding on senescent plankton and soft-bodied benthic fauna, or in the water column with dense patches of favored prey such as cryptophytes (Seaborn et al. 2001).

4.5.2 DNA Analyses. The cell synchrony technique used here was simple, effective, and less disruptive or caustic than some techniques (Grdina et al. 1987), since pure water was the arresting agent. Synchronously excysted populations from each cryptoperidiniopsoid isolate consisted of 1C and 2C DNA subpopulations, similar to the P. piscicida strains examined by Parrow et al. (2002).

It is reasonable, based on persistent sexuality and abundant fusions that most if not all of the 2C events detected in these cryptoperidiniopsoid cultures were planozygotes.

Adherence to the pattern of conventional meiosis would predict that the two dinoflagellates released from zygotic cysts would also each have 2C DNA, but that remains hypothetical.

In many cultured auto- or auxotrophic dinoflagellates that divide while motile,

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contaminating vacuolar DNA is not an issue, sexual fusions are nonexistent or controlled by culture conditions, and mitotic cell cycle progression can be directly measured and related to growth rate, cell cycle control, and other physiology (Yentsch et al. 1983, Gerath and

Chisholm 1989, Cetta and Anderson 1990, Taroncher-Oldenburg et al. 1997, Pan and

Cembella 1998, Peperzak et al. 1998, Pan et al. 1999, Van Dolah and Leighfield 1999,

Leighfield and Van Dolah 2001). DNA histograms obtained from such dinoflagellates indicate that the populations consist of normal eukaryotic cycling cells, with a G1-S-G2+M distribution (Gray et al. 1987). In cryptoperidiniopsoids, however, feeding 1C cells can be misinterpreted as undergoing DNA synthesis due to vacuolar DNA contamination, and motile zygotes can be misinterpreted as G2 phase haploid cells, as described in P. piscicida

(Parrow et al. 2002). Furthermore, interpretation of cryptoperidiniopsoid DNA distributions is hampered because cultured populations are largely cell cycle asynchronous, and division in resistant nonmotile cells impedes straightforward measurement of cell cycle progression, additionally complicated by persistent sexual reproduction. This forced- quiescence technique minimized vacuolar DNA and controlled cell synchrony, resulting in more precise DNA measurements and more interpretable DNA distributions. Sexual fusions disturbed the nuclear synchrony of the excysted cells, but observation of those fusions during the excystment period allowed us to assign the measured 2C DNA cells to the result of those fusions, planozygotes.

Precision of DNA measurements in flow cytometry was interpreted considering the

CVs, and 1C CVs between 7.3-8.4% were consistently obtained for these cultures. In comparison to other dinoflagellates analyzed using the same stain (SYTOX Green), 1C

DNA CVs of 8.3–29.0% were reported for 8 different photosynthetic species (Veldhuis et

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al. 1997), < 10% for Prorocentrum spp. (Pan and Cembella 1998), 40.0% for heterotrophic

C. cohnii (Veldhuis et al. 1997), and < 15% for phagotrophic Pfiesteria spp. (Parrow and

Burkholder 2002). Peperzak et al. (1998) reported a 1C CV of 7.0% for Prorocentrum micans Ehrenberg stained with propidium iodide. Other reported 1C CVs for photosynthetic species from microfluorometric DNA studies include 6.4–9.0% for

Heterocapsa triquetra (Ehrenberg) Stein using DAPI (Chang and Carpenter 1988) and 8.7–

9.1% for Hoechst 33342 – stained Gonyaulax polyedra Stein (Cetta and Anderson 1990).

Considerably lower CVs have been obtained on extracted dinoflagellate nuclei, probably because cytoplasmic DNA variability (mitochondria, chloroplasts, etc.) was eliminated and more stringent nuclear staining occurred. Hayhome et al. (1987) reported CVs from 2.4–

5.3% for Peridinium volzii Lemmermann nuclei stained with mithramycin, and CVs from

2.0–6.0% were reported for isolated Oxyrrhis marina Dujardin nuclei stained with Hoechst

33258 (Whiteley et al. 1993). Thus, comparison of the whole-cell values obtained here with other reports indicates that vacuolar DNA was effectively minimized by the arrest treatment.

The cryptoperidiniopsoid isolates of this study differed slightly in DNA content, and differed significantly from published values for P. piscicida and P. shumwayae (Parrow et al. 2002; but note that the two studies used CRBC standards from different sources, and some variability has been reported between chicken lines; Johnston et al. 1999). There was a positive correlation between cryptoperidiniopsoid DNA content and isolate age, similar to trends in chromosome number reported for some P. piscicida isolates (Burkholder et al.

2001). Among photosynthetic dinoflagellates, a similar trend was found in chromosome number of six Peridinium spp., and was attributed to long-term culture-induced positive

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aneuploidy (Holt and Pfiester 1982). Similar chromosomal autodiploidy was reported in an old culture of Karenia brevis (Davis) G. Hansen & Moestrup (Loper et al. 1980, Walker

1982). Holt (1983) also reported a polyploid increase in chromosome number over time in asexually reproducing Peridinium willei Huitfeld-Kaas, and found that induced sexual fusion and subsequent meiotic division returned cells back to a much-reduced haploid number. Based on thecal morphology and chromosome number, Loeblich et al. (1981) suggested polyploidy as a possible speciation mechanism in dinoflagellates. It is uncertain whether changes in chromosome number consistently reflect changes in DNA content of dinoflagellates (Beam and Himes 1984) or other eukaryotes (Kraemer et al. 1972). The phenomenon of sibling species (genetic discrepancies within a stable phenotype) may also occur, but proof of reproductive isolation would be required for verification (Beam and

Himes 1977, 1982, 1984). Different isolates of dinoflagellate morphospecies often differ in measured DNA content (Beam and Himes 1984, Hayhome et al. 1987, Parrow and

Burkholder 2002), as do other protists (Vaulot et al. 1994, Edvardsen and Vaulot 1996,

Green et al. 1996). Hayhome et al. (1987) suggested that this variation is due to population isolation and a high degree of genetic redundancy in dinoflagellate nuclei, allowing them to tolerate a less than precise division of nuclear DNA during cell division. Sexual reproduction between isolates with different nuclear DNA contents has been shown as well, indicating that the same lability may also extend to cell fusion (Hayhome et al. 1987).

4.5.3 Cryptoperidiniopsoids and Bacteria. Isolation of algal zoospores after release from parental thalli is a venerable technique for obtaining contaminant-free isolates

(Pringsheim 1964). The application here was only successful in four of seven test cultures, likely because the goals of attaining high densities while simultaneously eliminating all

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contaminants were somewhat in conflict (Guillard 1973). The treatments can be improved for exacting use on fewer cells or with added washing steps. Once purified, the post- treatment cryptoperidiniopsoids proliferated poorly in bacteria-free cryptomonad cultures compared with pre-treatment cultures with abundant bacteria, in partial agreement with

Alavi et al. (2001). Cryptoperidiniopsoids were not observed to consume bacteria, although consumption of bacteria has been reported for other myzocytotic dinoflagellates via other feeding mechanisms (Burkholder and Glasgow 1997). Here, cryptoperidiniopsoids encysted in culture with abundant bacteria when cryptomonad prey were depleted. Alavi et al. (2001) speculated that certain epiphytic bacteria might directly influence cryptoperidiniopsoid survival, and in support of that premise, some photosynthetic dinoflagellates grow more slowly in axenic culture (Popovský and Pfiester 1990).

However, the role of specific extracellular bacteria as sole providers of a metabolite or other factor required for survival by an omnivorous, phagotrophic dinoflagellate in nature is unknown.

Barker (1935) described a cyst forming heterotrophic dinoflagellate (likely from the

C. cohnii species complex; Pringsheim 1956, Guillard and Keller 1984), cultured with rotting Fucus infusions, that apparently required mixed bacteria for development. However, members of this species complex can be cultured without bacteria (Provasoli and Gold

1962, Tuttle and Loeblich 1975, Beam and Himes 1980, 1982, Guillard and Keller 1984).

In contrast, survival of parasitic Amyloodinium ocellatum (considered a close relative of cryptoperidiniopsoids; Litaker et al. 1999) was severely reduced by the presence of bacteria in culture (Brown 1934, Noga and Bower 1987). Hematodinium spp., dinoflagellate parasites of crustaceans, do not require bacteria in culture (Appleton and Vickerman 1998).

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The free-living phagotrophic dinoflagellate Oxyrrhis marina Dujardin can be cultured without bacteria on various algae (Droop 1966), and axenically in defined chemical media

(Droop 1970, Droop and Pennock 1971). The omnivore Noctiluca scintillans Suriray can be grown in chemical media free of other organisms if glass particles are provided to elicit ingestion and vacuole formation (Lee 1990), but it grows more slowly than with live food

(McGinn 1969). Lee (1977) cultured heterotrophic Gyrodinium lebouriae Herdman axenically, and found that isolates containing intracellular bacteria fed by resorption, whereas those without were phagotrophic.

Despite these examples of heterotrophic dinoflagellates that do not require extracellular bacteria in culture, in natural assemblages bacteria and protozoa have many complex interactions (Hamilton 1973, Görtz and Brigge 1998, Lewis et al. 2001). Lee

(1990) described food size and quality as primary factors influencing the growth of protozoa, but clarified that it is sometimes difficult to separate the effects of feeding parameters from nutrition. Like some other myzocytotic dinoflagellates, cryptoperidiniopsoids appear to feed more selectively or effectively on physiologically stressed, damaged, or leaky prey. Bacteria can negatively impact the physiology of microalgae through competition for limiting nutrients, production of inhibitory compounds, cellular infection, and direct lysis (Cole 1982, Doucette 1995, Smayda 1996, Manage et al.

2000). Certain heterotrophic bacteria probably enhance release of labile exudates from aging cryptomonad cells (Fogg 1983), which would make the cells more readily detected by chemotactic dinoflagellates (Fenchel and Blackburn 1999). Limited feeding by cryptoperidiniopsoids on cryptomonads occurred in the absence of bacteria, but a higher chemotactic attraction was indicated by increased feeding activity on damaged

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(microwaved) cryptomonads. Moreover, when weakly feeding bacteria-free cryptoperidiniopsoids were added to an aged cryptomonad culture containing endemic bacteria, vigorous feeding commenced immediately. This further supports the premise that bacteria in the cryptophyte culture impacted the cryptoperidiniopsoid feeding response by modifying the cryptophytes in some way that made them more ‘leaky’ or otherwise increased their susceptibility to predation.

4.5.4 Growth On Dissolved Organic Compounds, and Mixotrophy. Proof of heterotrophic nutrition requires not only uptake, but also increased growth or survival of an organism by using organic substances (Gaines and Elbrächter 1987). In this study, cryptoperidiniopsoid flagellated cells placed in organically supplemented particle free media showed increased short-term persistence over those in plain seawater, suggesting that limited benefit occurred (Elbrächter 1972). Uptake of organic nutrients has been demonstrated in the closely related dinoflagellate, P. piscicida (Burkholder and Glasgow

1997, Glasgow et al. 1998, Lewitus et al. 1999b). Here, the two media that supported apparent increased maintenance of flagellated cells were both supplemented with animal serum. Serum is a complex mixture containing various proteins, polypeptides, hormones and lipids (Freshney 1986). Dissolved proteins and organic nitrogen may have been beneficial to the cryptoperidiniopsoids in these treatments, and the importance of lipids in the nutrition of other phagotrophic protozoa has also been suggested (Droop and Doyle

1966, Droop 1970, Droop and Pennock 1971). Although the organic media used here were not reliable for axenic cryptoperidiniopsoid propagation, other media formulations may be successful (e.g. as in Droop 1959, Droop and Pennock 1971), and the purification techniques used here could assist in such efforts (Droop 1970).

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Mixotrophy through temporary maintenance of cryptophyte chloroplasts

(kleptoplastidy) has been examined in Pfiesteria spp. (Lewitus et al. 1999a,b, Glasgow et al. 2001, Feinstein et al. 2002), and cryptoperidiniopsoids may also have kleptochloroplasts under some conditions (Parrow et al. 2001, Eriksen et al. 2002). The cryptoperidiniopsoid isolates of this study ingested the cytoplasm (including photopigments) of cryptomonad prey, and resulting food vacuoles were pigmented and autofluorescent. After feeding the dinoflagellates encysted to divide and during division the food vacuoles shrank and grew paler. The food body was unequally inherited, and the offspring cell that inherited it often egested the undigested remnants shortly after excystment. Depletion of cryptomonad prey quickly led to colorless cryptoperidiniopsoids. These observations did not indicate long- term maintenance of prey-derived chloroplasts for these isolates, but ephemeral (< 2 d) phototrophy may have occurred (Eriksen et al. 2002), as reported for other phagotrophic dinoflagellates (Skovgaard 1998, Feinstein et al. 2002).

Differentiation between kleptoplastidy and simple digestion is often problematic.

Based on flow cytometric particle sorting to isolate pigmented cryptoperidiniopsoids from cryptophytes, the isolated dinoflagellates lost (digested) the pigmented contents of their food vacuoles in < 2 days, and no difference in growth or survival was found between light and dark treatments. Thus, these cryptoperidiniopsoids did not measurably benefit from photosynthesis. However, the sorting treatment or some other unknown factor may have reduced the capacity for kleptoplastid photosynthetic activity or retention. Kleptoplastidy in obligately heterotrophic dinoflagellates has been interpreted as a short-term adaptation to prey limitation (Skovgaard 1998, Lewitus et al. 1999a,b). In these cryptoperidiniopsoid cultures, formation of quiescent cysts was the most obvious response to prey limitation.

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4.5.5 Future Directions. In the most recent major review of dinoflagellate sexuality

(Pfiester 1989), information on sexual reproduction was known for only 5 heterotrophic species out of a total of 31, and only partial accounts were available for several species.

Cryptoperidiniopsoids can be added to the relatively short list of heterotrophic dinoflagellates for which such information exists. However, the life cycle findings from this research are incomplete, and further elucidation of the life history will require examination of more strains under different conditions. More work is needed to better characterize meiosis in cryptoperidiniopsoids, and the importance of vegetative and sexual processes as survival strategies when prey are scarce or other environmental conditions are unfavorable. The nutritional findings (i.e. limited benefit from dissolved compounds and no measured benefit from kleptoplastidy), while true for these strains under the imposed culture conditions, are also indeterminate since different isolates within given dinoflagellate morphospecies commonly differ in behavior, physiology, and genetics (Lee 1977, Yentsch et al. 1978, Beam and Himes 1977, 1984, Hayhome et al. 1987, Wood and Leatham 1992,

Taylor 1993). External or endosymbiotic bacteria may be beneficial to cryptoperidiniopsoids in cryptophyte-fed culture, but the importance of that phenomenon in nature is unknown. Furthermore, cryptoperidiniopsoids and similar dinoflagellates may be species complexes of genetically divergent microorganisms (Beam and Himes 1982, Taylor

1987). Considering that cryptoperidiniopsoids are presently only a putative genera-level taxon, considerable variability beyond what was observed here is expected.

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Acknowledgments

We thank H. Marshall for some of the cryptoperidiniopsoid cultures; P. Rublee and D.

Oldach for assistance and counsel on molecular taxonomy; N. Allen and E. Johannes for

PM-DIC assistance; J. Alexander for assistance with cultures; and N. Deamer, H. Glasgow,

D. Stoecker, T. Tengs, and D. Kamykowski for counsel. Two anonymous reviewers improved the manuscript. Funding for this research was provided by the North Carolina

General Assembly, the National Science Foundation (#9912089), the U.S. Environmental

Protection Agency (#CR 827831-01-0), and the Department of Botany at North Carolina

State University.

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5. A CELL CYCLE SYNCHRONIZATION AND PURIFICATION TECHNIQUE FOR HETEROTROPHIC PFIESTERIA AND CRYPTOPERIDINIOPSOID DINOFLAGELLATES ANALYZED BY FLOW CYTOMETRY

Matthew W. Parrow, Nora J. Deamer, Jessica L. Alexander, and JoAnn M. Burkholder

Proceedings of the Tenth International Conference on Harmful Algae, 2002 (submitted)

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5.1 Abstract

A swarmer release method was described for obtaining dense, highly synchronous flagellated cell populations of phagotrophic Pfiesteria spp. and cryptoperidiniopsoid dinoflagellates. Cyst populations were held in deionized water for 2-5 days, and flagellated cells excysted upon resumption of previous culture salinity. The cells within the cysts apparently had not been harmed by the extended immersion in deionized water, and a simultaneous culture purification treatment using sodium hypochlorite, tested with

Pfiesteria shumwayae, was successful. Flow cytometric DNA analyses on the synchronized populations were improved in comparison to analyses on untreated populations, and initial synchrony of flagellated cell populations typically was greater than

90% 1C DNA. Vacuolar DNA contamination from previous phagotrophy was minimized.

The 2C DNA subpopulations that occurred consisted mostly or entirely of motile zygotes formed from sexual fusion of the released flagellated cells. The techniques described here may be useful in ecological, biochemical, and molecular investigations of these and similar heterotrophic dinoflagellates.

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5.2 Introduction

Progression through the cell cycle is defined by the physiological events of genome replication, mitosis, and cytokinesis. In protists, cell cycle events are closely linked to behavior, biochemistry, and life cycle differentiation (Wong, 1996). Study of synchronous populations can provide insights into the development, ultrastructure, and mean duration of specific stages in heteromorphic dinoflagellates (Kubai and Ris, 1969; Bhaud et al., 2000), and effective investigation of regulatory and cyclic biochemical and molecular events in proliferating cells often requires synchronized populations. Many photosynthetic dinoflagellates exhibit natural diel division phasing, which has allowed study of intrinsic and extrinsic factors influencing cell cycle-related physiology (e.g. Gerath and Chisholm,

1989; Van Dolah and Leighfield, 1999). In contrast, heterotrophic species rarely have been described with diel synchrony (e.g. Ucko et al., 1997), and very few have been manipulated into synchrony. Crypthecodinium species are unusual in that they can be grown on solid media and form vegetative cysts; synchronous excystment yields flagellated cells that can then be collected from the liquid media (Beam and Himes, 1980; Bhaud et al., 1991; Wong and Whiteley, 1996).

For some dinoflagellate species, reproductive cyst adherence has been used to isolate synchronous populations (Kubai and Ris, 1969; Franker et al., 1973), and resistance of some encysted species to lysis has been used to obtain synchronous populations by selective lysis of less resistant flagellated cells (Franker, 1971; Parrow et al., 2002).

Pfiesteria spp. and cryptoperidiniopsoids are estuarine phagotrophic dinoflagellates that commonly undergo cell division in adherent benthic cysts. Submersion of these cysts in deionized (DI) water does not inhibit cell division, but arrests excystment of flagellated

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offspring (Parrow et al. 2002; Parrow and Burkholder submitted). Here we described and tested a forced-quiescence swarmer release method for obtaining relatively dense, synchronized flagellated populations of these dinoflagellates with minimal ingested prey

DNA, and we tested a simple method for culture purification. The degree of synchrony and reduction of vacuolar DNA were analyzed by flow cytometry.

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5.3 Materials and Methods

Dinoflagellates in culture at the CAAE (n > 4 clones per taxon) were propagated in flasks with cryptomonads or in aquaria fed live fish (as in Burkholder et al., 2001). Cyst accumulation depended on culture growth and population density; frequent feedings were required to obtain dense cyst beds (covering the bottom of the culture vessel). Vessels with dense cyst accumulations were rinsed briefly with, and then immersed in, DI water. DI water rinsed away and/or lysed flagellated cells and many microbial contaminants, and reduced the dinoflagellate populations to cysts. Cysts were held in DI water for 24–120 h, and then were covered in sterile 15-psu artificial seawater. Within 3 h robust excystment typically occurred, and flagellates were decanted away from remaining cysts. Percent excystment was determined by monitoring fields of view of at least 50 cysts with light microscopy (LM; Olympus IX70, 200x), and marking excystment over time on a video monitor. Flagellate behavior (e.g. gamete fusion) was also tracked (LM; Olympus AX70, seawater immersion objectives, 200-400x). For flow cytometry, samples were fixed with

0.5% paraformaldehyde, treated with RNase A (1 g mL-1, Sigma), and stained with 5 M

SYTOX Green (Molecular Probes) (Parrow et al. 2002). Multicycle Software (Phoenix

Flow Systems) was used to compute peak numbers and CVs (relative standard deviation).

Vacuolar DNA was assessed on reduction of 1C peak CVs.

Effects of sodium hypochlorite treatment were tested on cysts from 2 P. shumwayae clones. P. shumwayae cysts were collected for purification in petri dishes placed in aquaria and incubated in DI water. Cysts were gently scraped free, re-suspended in 40 mL of DI water, shaken vigorously, pelleted by centrifugation (2000 rpm for 10 min), then re- suspended in DI water with 0%, 0.005%, 0.05% or 0.5% sodium hypochlorite (dilution of a

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6% sodium hypochlorite commercial bleach solution) and shaken. After 5 min cysts were re-pelleted and cleaned twice by shaking then pelleting in sterile DI water, then re- suspended in sterile 15-psu artificial seawater and observed for excystment and contaminants under LM. Sub-samples (1 mL) were inoculated into culture tubes with 20 mL 15-psu f/2 media enriched with 1 g L-1 bacto-peptone and 1 g L-1 glucose, and checked daily for turbidity as evidence of bacterial or fungal growth; un-inoculated tubes were clarity controls (Hoshaw and Rosowski 1973).

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5.4 Results and Discussion

Reproductive cyst development apparently was unaffected in DI water and divisions occurred, but flagellated offspring did not emerge in the osmotically unfavorable medium.

Respiration and digestion continued and minimized vacuolar DNA, supported by observations that the food vacuoles contained in many cells grew smaller and paler during the arrest. The duration of arrest was longer than the time typically required for completion of fissions and excystment, forcing the cysts into developmental synchrony. A robust excystment response typically occurred within 3 h of resumed salinity. Although the arrest conditions did not appear harmful over short durations, periods of arrest longer than optimal resulted in less vigorous excystment (Fig. 1A), probably due to cell death or induced dormancy. In this study, ca. 48 h of arrest optimized excystment for P. piscicida and the cryptoperidiniopsoid, and ca. 72 h was optimal for P. shumwayae. Arrest was also extended to 14 days for each taxon, and excystments still occurred after resumption of culture salinity. P. shumwayae cysts were also resistant to sodium hypochlorite (Fig. 1B).

Treatments of 0.05% and 0.5% hypochlorite typically resulted in cysts that appeared visually free of living contaminants (Fig. 1C). Inoculation into enriched broth indicated these treatments decreased contaminant abundance/diversity based on longer durations (ca.

4–7 days) of broth clarity compared to controls; all demonstrable contaminants were apparently eliminated in 40% of treatments using 0.5% hypochlorite.

Released flagellates were similar in size, colorless, and rapidly swimming. Cell pairs in apparent fusion occurred with variable frequency in all excysted populations, and on occasion were observed to complete fusion and form motile zygotes. Zygotes were highly mobile and escaped continuous observation. Some isolated flagellated cells

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(swarmers) produced small cysts that were morphologically consistent with non- reproductive temporary cysts (Parrow et al. 2002). Flow cytometry showed that the released swarmer populations of each taxon were composed primarily (≥ 90%) of 1C DNA cells (Fig. 2). There were 2C DNA subpopulations in each taxon, which likely were composed of sexual products based on observation of cell fusions among the released flagellated cells prior to fixation. In each species, fusions were only observed during the first several hours after swarmer release, and thereafter 2C DNA cell abundances were stable over time (days), indicating that fully differentiated gametes emerged from cysts.

Cells with 2C DNA were not detectably larger than 1C cells based on forward scatter signals. The 1C peak CVs were < 8.3, comparable to whole-cell values obtained for photosynthetic dinoflagellates using the same stain (e.g. Veldhuis et al. 1997), and lower than previously reported values for untreated Pfiesteria spp. (i.e. populations analyzed without use of this forced quiescence method; Parrow and Burkholder 2002). Thus vacuolar DNA apparently was minimized by digestion during the forced quiescence.

Most flagellated cells of Pfiesteria spp. and cryptoperidiniopsoids (except for gametes), whether haploid or diploid, apparently are trophic (feeding) and interphasic

(Parrow et al. 2002). Determination of the full cell cycle progression of these taxa will require inclusion of encysted stages, or at least the nuclei of those stages, in the analyses since a significant proportion of divisions occur in cysts. The relatively simple techniques described here yielded populations of abundant, highly synchronized cells, and could be beneficial in studies of the cell biology, physiology, and ecology of these heterotrophic dinoflagellates. These techniques also isolated cells with minimal vacuolar DNA

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contamination and external contaminants, which will be useful in biochemical and molecular investigations.

Acknowledgments

We thank Dr. H. Marshall for providing a cryptoperidiniopsoid culture, and Dr. P.

Rublee for assistance in molecular taxonomy. Funding was provided by the NSF

(#9912089) and the NC General Assembly.

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A 100 B 90 80

60 60

40 30 20 Relative Excystment ( % ) ) % ( Excystment Relative

) % within 3 h ( Excystment 0 0 0 0.005 0.05 0.5 0 24487296120 Duration of Arrest ( h ) Sodium Hypochlorite ( % )

Figure 5.1. (A) Percent excystment within 3 h of resumed culture salinity from representative P. piscicida cyst beds after increased duration of arrest in DI water (means +

2 standard errors [SEs]; n=4). (B) Percent excystment of P. shumwayae after hypochlorite treatment relative to that of untreated cysts (n=2). (C) Visually contaminant-free P. shumwayae cysts from an aquarium culture after treatment with 0.05% sodium hypochlorite

(bar = 10 µm).

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P. piscicida A B 1C = 93% CV = 7.6 1C

FS Number

2C 2C 1C

DNA DNA

P. shumwayae

1C = 96% C D

CV = 7.2 1C

FS Number 2C 1C 2C

DNA DNA

cryptoperidiniopsoid

1C = 90% E F CV = 7.1 1C

FS Number

2C 2C 1C

DNA DNA

Figure 5.2. Representative cellular DNA-fluorescence plots for released swarmers. Bottom axis is relative DNA-fluorescence in all plots; vertical axes in two-parameter scatter plots

(B,D,F) are linear forward scatter (a sizing parameter). A-B, P. piscicida; C-D, P. shumwayae; E-F, cryptoperidiniopsoid (beads = fluorescence standard).

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5.5 References

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Bhaud, Y., Guillebault, D., Lennon, J. F., Defacque, H., Soyer-Gobillard, M.O. and Moreau, H. 2000. Morphology and behavior of dinoflagellate chromosomes during the cell cycle and mitosis. J. Cell Sci., 113:1231--1239. Bhaud, Y., Salmon, J.M. and Soyer-Gobillard, M.O. 1991. The complex cell cycle of the dinoflagellate protoctist Crypthecodinium cohnii as studied in vivo and by cytofluorimetry. J. Cell Sci. 100:675-682. Burkholder, J. M., Glasgow, H. B. and Deamer-Melia, N. 2001. Overview and present status of the toxic Pfiesteria complex (Dinophyceae). Phycologia, 40(3):186-214. Franker, C.K., 1971. Division synchrony in primary cultures of an endozoic dinoflagellate. J. Phycol. 7, 165-169. C. K. Franker, L. F. Smith and L. M. Sakhrani. 1973. Morphotypic transition during synchronous growth of Crypthecodinium (Gyrodinium) cohnii. Arch. Mikrobiol. 90, 255-262. Gerath, M.W. and Chisholm, S.W. 1989. Change in photosynthetic capacity over the cell cycle in light/dark-synchronized Amphidinium carteri is due solely to the photocycle. Plant. Physiol. 91:999-1005. Hoshaw, R.W. and Rosowski, J.R. 1973. Methods for microscopic algae. In: Stein, J.R. [Ed.] Handbook of Phycological Methods: Culture Methods and Growth Measurements. Cambridge University Press, Cambridge, NY, pp. 53-67. Kubai, D.F., Ris, H., 1969. Division in the dinoflagellate Gyrodinium cohnii (Schiller). J. Cell Biol. 40, 508-528. Parrow, M.W. and Burkholder, J.M. Accepted. Estuarine heterotrophic cryptoperidiniopsoids (Dinophyceae): life cycle and culture studies. J. Phycol. Parrow, M.W. and Burkholder, J.M. 2002. Flow cytometric determination of zoospore DNA content and population DNA distribution in cultured Pfiesteria spp. (Pyrrhophyta). J. Exp. Mar. Biol. Ecol. 271:140-155. Parrow, M., Burkholder, J.M, Deamer, N.J. and Zhang, C. 2002. Vegetative and sexual reproduction of Pfiesteria spp. (Dinophycea) cultured with algal prey, and inferences for their classification. Harmful Algae 1:5-33.

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Ucko, M., Elbrächter, M. and Schnepf, E. 1997. A Crypthecodinium cohnii-like dinoflagellate feeding myzocytotically on the unicellular red alga Porphyridium sp. Eur. J. Phycol. 32:133-140. Van Dolah, F.M. and Leighfield, T.A. 1999. Diel phasing of the cell-cycle in the Florida red tide dinoflagellate, Gymnodinium breve. J. Phycol. 35:1404-1411. Veldhuis, M.J., Cucci, T.L. and Sieracki, M.E. 1997. Cellular DNA content of marine phytoplankton using two new fluorochromes: taxonomic and ecological applications. J. Phycol. 33:527-541. Wong, J.T.Y., 1996. Protozoan cell cycle control. Biol. Signals 5, 301-308. Wong, J.T.Y., Whiteley, A., 1996. An improved method of cell-cycle synchronization for the heterotrophic dinoflagellate Crypthecodinium cohnii Biecheler analyzed by flow cytometry. J. Exp. Mar. Biol. Ecol. 197, 91-99.

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6. REPRODUCTION AND SEXUALITY IN PFIESTERIA SHUMWAYAE

(DINOPHYCEAE)

Matthew W. Parrow and JoAnn M. Burkholder

Journal of Phycology (submitted)

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6.1 Abstract

Pfiesteria shumwayae is a heterotrophic dinoflagellate with a widespread distribution in temperate-subtropical estuarine waters. In this study, four clonal isolates from the eastern coast of North America, one from New Zealand, and a mixed composite of these clones were cultured in aquaria and fed live fish. Fission, sexuality, and phagotrophic feeding on fish were studied by light microscopy, scanning electron microscopy, and flow cytometry. The development of reproductive cysts isolated from aquaria was followed.

Synchronously excysted flagellate populations were examined for sexuality, then feeding behavior and reproduction when given larval fish. Reproductive cysts varied considerably in size and underwent protoplast division(s), most commonly producing 2-8 biflagellated offspring. Fusing gametes, resulting planozygotes, and nuclear cyclosis were documented as evidence of sexuality. Gametes emerged from cysts, and fusions were approximately isogamous. Resulting planozygotes had two longitudinal flagella and one transverse flagellum, and apparently fed before encysting. Distinct and lengthy chromosome movements (nuclear cyclosis) occurred in presumed zygotic cysts prior to nuclear division(s). These cysts did not exhibit dormancy in growing cultures, and produced 2 or 4 biflagellated offspring. Flagellated cells fed on surficial fish tissues and then encysted for reproduction. Stages indicating a completed sexual cycle (fusion, planozygotes, and nuclear cyclosis) were uncommon or absent in clonal cultures, but were relatively abundant in the mixed clone culture. Self-sterility factors apparently influenced sexuality. Starved populations formed quiescent cysts that released swimming cells when food was provided.

P. shumwayae reproduction and sexuality were similar to other and closely related species, with minor differences.

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6.2 Introduction

Dinoflagellates are an ancient group of aquatic protists with diversified ecologies.

Extant phototrophic and heterotrophic species are roughly equal in number, and mixotrophy is common (Schnepf and Elbrächter 1992, 1999, Stoecker 1999). They can be free-living, symbiotic in invertebrates, or parasitic on or in other protists, invertebrates, and vertebrates

(Cachon and Cachon 1987, Trench 1987, Gaines and Elbrächter 1987, Coats 1999).

Despite this adaptive divergence, certain basic life history traits appear to be largely conserved. Like other protists, dinoflagellate reproduction is dominated by asexual fission.

Von Stosch (1964, 1965, 1972, 1973) first authenticated dinoflagellate sexuality, and since then a comparatively rare but growing number of species have been shown to have a sexual cycle (reviewed in Beam and Himes 1980, Pfiester 1989). As presently understood, dinoflagellate sexuality begins with the mitotic production of functionally haploid gametes which fuse nuclei to produce a diploid zygote. Subsequent meiotic nuclear division(s) of the zygote returns the offspring to the haploid vegetative state. Thus, all dinoflagellates for which knowledge of sexuality exists are thought to demonstrate primarily haplontic life histories with zygotic meiosis (including the previously considered exception, Noctiluca scintillans (Macartney) Kofoid; Schnepf and Drebes 1993, Elbrächter and Qi 1998).

Predatory dinoflagellates are abundant and ecologically influential (Gaines and

Elbrächter 1987, Lessard 1991, Burkholder 1998, Jacobson 1999, Jeong 1999), but have been numerically underrepresented in studies of dinoflagellate reproduction and sexuality.

This is probably because relatively few phagotrophic species have been brought into sustained culture (Lessard 1993). Pfiesteria shumwayae Glasgow & Burkholder is a widespread, phagotrophic (via myzocytosis) dinoflagellate that can be continuously grown

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in culture (Burkholder et al. 2001a, Glasgow et al. 2001). Fission in this species generally occurs in nonmotile cells (cysts) that produce two to several flagellated offspring (Parrow and Burkholder 2002, Parrow et al. 2002), similarly to several other dinoflagellate species

(von Stosch 1973, Drebes and Schnepf 1988, Montresor 1995, Ucko et. al. 1997).

Although aspects of asexual proliferation and flagellate population DNA profiles in P. shumwayae have been described, the sexual cycle has not been well documented

(Burkholder et al. 2001a,b, Glasgow et al. 2001, Parrow and Burkholder 2002, Parrow et al.

2002). A recent investigation using clonal isolates fed microalgal prey was unable to substantiate sexuality in P. shumwayae, whereas it was found commonly in Pfiesteria piscicida Steidinger & Burkholder clonal isolates maintained the same way (Parrow et al.

2002). The purpose of this study was to document and evaluate reproduction and sexuality in P. shumwayae using clonal isolates and a mixed composite of those isolates given live fish as food.

Dinoflagellates were maintained in aquaria (as in Burkholder et al. 1995, 2001c), and cysts were collected for observation and treatment in petri dishes placed on the bottom of the aquaria. Cyst development and excystment were followed, and dinoflagellates from synchronized excystments were observed for sexual behavior as well as phagotrophic feeding and subsequent reproduction when given larval fish. Fusing flagellated cells

(gametes), resultant zygotes with two longitudinal flagella (planozygotes), and nuclear cyclosis were documented. Nuclear cyclosis is a prolonged (hours) and distinctive swirling of dinoflagellate zygotic chromosomes that is believed to coincide with the homologue pairing of meiotic prophase, and thus precedes meiotic division (von Stosch 1972, 1973,

Beam and Himes 1980, 1984, Pfiester 1984, 1989, Pfiester and Anderson 1987).

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Reproduction and sexuality in P. shumwayae were described and compared with similar and closely related species.

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6.3 Materials and Methods

6.3.1 Cultures. The five clonal P. shumwayae isolates examined in this study were in culture at the NCSU Center for Applied Aquatic Ecology. They had been obtained from water and sediment samples collected from estuaries in North Carolina and Maryland,

U.S.A. and New Zealand (Table 6.1). Clonal isolates were initiated with single flagellated cells obtained by flow cytometric cell sorting (Burkholder et al. 2001a). Species identification was confirmed by scanning electron microscopy (SEM) of suture-swollen biflagellated cells (Burkholder et al. 2001a, Glasgow et al. 2001, Rhodes et al. 2002), and by sequence-specific 18S rDNA based PCR molecular probes (Rublee et al. 1999, Bowers et al. 2000, Oldach et al. 2000). Isolates had been maintained in clonal culture on microalgal prey for 2 – 3 years prior to this study (Burkholder et al. 2001a). Although these isolates were each initiated with a single flagellated cell and were therefore considered clonal, diploid cells inadvertently could have been isolated (Parrow et al. 2002).

Subcultures of the five clonal, non-axenic isolates were also combined to make a mixed- clone P. shumwayae culture. All cultures were inoculated into separate 10 L aquaria filled with 15-ppt salinity synthetic seawater (Instant Ocean, Aquarium Systems, Mentor, OH).

They were fed juvenile tilapia (Oreochromis mossambicus Peters) and maintained for weeks to months as in Burkholder et al. (2001c).

P. shumwayae cysts were collected for observation and treatment in duplicate petri dishes placed on the bottom of the aquaria. After 3 – 5 days dishes were removed from aquaria and replaced to provide a regular supply of new cells. After removal from the aquaria, dishes were gently rinsed with deionized water and the outside surfaces were wiped clean and sterilized with ethanol. P. shumwayae cysts were adherent and apparently

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Table 6.1. The P. shumwayae isolates examined, estuary where collected, and time in culture.

Isolate Isolate age Source location designation (months)

102716 Neuse River Estuary, NC, USA 34

102539 Neuse River Estuary, NC 55

102540 Newport River Estuary, NC 24

102537 Chesapeake Bay, Maryland, USA 30

102558 Tasman Bay, New Zealand 32

102613 Mix of the above clonal isolates mixed (above)

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unharmed by temporary immersion in deionized water (Parrow et al. 2002), whereas flagellated cells and many potential contaminants were lysed and/or rinsed away.

For observation of cyst development and excystment, dishes were refilled with sterile 15 ppt-salinity synthetic seawater. For population excystment synchronization, cysts were held in deionized water for 72 h. As for related dinoflagellates (Parrow and

Burkholder accepted), P. shumwayae reproductive cyst development apparently was unaffected in deionized water, but release of flagellated offspring did not occur in the osmotically unfavorable medium. In other words, deionized water did not inhibit fission in cysts but arrested excystment. The duration of arrest was longer than the time typically required for completion of fissions and excystment. Thus, most reproductive cysts were synchronized to a similar state, pending excystment upon resumption of favorable culture salinity (Parrow and Burkholder submitted). Dishes were refilled with sterile synthetic seawater to end the arrest. After a robust excystment response had occurred (typically in 2–

4 h), flagellated cells were decanted away from the remaining adherent cysts into a new dish.

The rationale for subjecting the dinoflagellates to this forced quiescence synchronization procedure was multifold. First, cultures maintained in aquaria with fish invariably contain contaminating microorganisms (Burkholder et al. 2001c), and the treatment eliminated those that were unattached and/or less resistant to osmotic stress.

Second, since the encysted cells underwent a period of forced quiescence, cytoplasmic

DNA contamination from previous phagotrophy was minimized, increasing the resolution of nuclear DNA measurements on the excysted offspring. Also, since most released

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dinoflagellates were the direct products of cell division without having had the opportunity to feed, subpopulations with the lowest cellular DNA content could be equated to the basal nuclear state (functionally haploid, G1 cell cycle phase), and could be compared to other

DNA content subpopulations such as diploid planozygotes with greater certainty. Third, synchronized populations were simplified compared to asynchronous populations. The immediate history of the released dinoflagellates was known (i.e. they emerged from cysts), which offered diagnostic advantages in determining the sequence of life cycle transitions.

Food was provided or denied to the synchronous cells, and subsequent development compared. Fourth, life cycle events that occurred relatively infrequently and/or quickly

(such as gamete fusion) in asynchronous populations might be more readily observed or detected if they occurred more synchronously.

The excysted dinoflagellate populations were examined microscopically for sexuality (gamete fusions, planozygotes) and other behavior, and then were sampled for flow cytometric population DNA measurements. Dinoflagellate suspensions were inoculated into culture flasks or petri dishes with larval (length 10– 15 mm) Atlantic pupfish (Cyprinodon variegatus Lacepède) to follow feeding behavior and short-term reproductive development. Replicate dinoflagellate suspensions were left without food and observed for behavior. Duplicate petri dishes were removed from each aquarium culture weekly over a three-week period for observations of cyst development and excystments, synchronization treatments, sampling for microscopy and flow cytometry, and feeding treatments.

6.3.2 Microscopy. Live cells and life cycle events were observed, photographed, and video recorded in culture flask and petri dish subcultures. An Olympus IX70 inverted microscope (Olympus Corporation, Melville, NY; 40-400x) was used for routine

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examination. Critical observations and imaging were made using an Olympus AX-70 light microscope with Olympus 40x 0.8 NA and 60x 0.9 NA saltwater immersion objectives, a

0.8 NA condenser, and a DEI-750 cooled-chip CCD camera (Optronics Engineering,

Goleta, CA).

For verification of planozygotes using light microscopy, culture aliquots were mixed under a fume hood with a freshly made ice-cold fixation buffer to effect a final concentration of 1% glutaraldehyde, 0.1 M pH 7 sodium cacodylate buffer, and 0.005% osmium tetroxide. Drops of the fixed dinoflagellate suspensions were immediately placed on microscope slides and sealed under cover slips ridged with silicone grease. Flagella were well preserved using this method, provided that the slides were examined within a few hours. Cells with two longitudinal flagella were considered planozygotes, and were identified with a 20x 0.9 NA high-dry objective and imaged using a 60x 1.2 NA water objective. Preparations selected for lipid staining were fixed in 0.5% glutaraldehyde and stained with 1 µg·mL-1 (final concentration) Nile red (Molecular Probes, Inc., Eugene, OR) from a 250 µg·mL-1 stock solution prepared in acetone (Cooksey et al. 1987, Parrow et al.

2002) for 30 min in darkness prior to microscopic observation. Brightfield, differential interference contrast, and epifluorescence microscopy with mercury lamp excitation were used to image cells and fluorophore-bound complexes.

For SEM, culture aliquots were fixed with 1% osmium tetroxide, 2% reagent-grade glutaraldehyde, and 0.1 M sodium cacodylate, pH 7 (final concentrations) at 4 ºC in darkness for 20 min (Burkholder and Glasgow 1995). Cells were filtered onto 3 µm-pore size polycarbonate filters (Nucleopore Corp., Pleasanton, CA), rinsed in 0.1 M sodium cacodylate solution, and dehydrated through an ethanol series. Samples were critical-point- dried with carbon dioxide, sputter-coated with 30 nm Au/Pd, and viewed at an accelerating

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voltage of 15 kV on a Joel 5900LV scanning electron microscope (JOEL USA, Inc.,

Peabody, MA).

6.3.3 Flow Cytometry. Culture aliquots were filtered through 38 µm-porosity nylon screen (Wildlife Supply Co., Saginaw, MI) to remove potential debris, and were fixed with

0.5% paraformaldehyde (final concentration) overnight at 4ºC (Parrow et al. 2002). The dinoflagellates were then concentrated by centrifugation, treated with RNase A (1 µg·mL-1 final [w/v], Sigma, St. Louis, MO) for 1 h at room temperature, stained with 5 µM SYTOX

Green (Molecular Probes, Inc.), and returned to storage conditions for 12-14 h (Veldhuis et al. 1997, Eschbach et al. 2001). DNA measurements and cell sorting were performed with an

EPICS Altra flow cytometer (Coulter Corporation, Miami, FL) using a 15-mW argon laser (488 nm). Fluorescence emission of SYTOX-DNA complexes was detected at 525 nm.

Fluorescence and scatter signals were recorded on at least 104 cells per analysis, displayed using EXPO32 Analysis software (Applied Cytometry Systems, Sheffield, UK), and stored in listmode format. Cytometer amplification system linearity was checked using Immuno-Brite fluorospheres (Coulter Corporation) (Bagwell et al. 1989), and optical alignment and signal stability were monitored using Flow-Check fluorospheres (Coulter Corporation). Unstained samples were used as negative fluorescence controls. Cytoplasmic DNA contamination was assessed based on coefficients of variation (CVs, relative standard deviation) for the peak 1C

DNA fluorescence distributions. Electrostatically sorted dinoflagellates were collected on microscope slides for observation and sealed under cover slips ridged with silicone grease.

6.3.4 Data Analyses. Flow cytometric listmode data were plotted as univariate signal height distributions and bivariate frequency distributions. Artificial DNA doublets were identified from fluorescence signal peak height versus integral signal area plots,

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assuming that DNA doublets deviated from a linear ratio of the two signals (Wersto et al.

2001). MultiCycle DNA Content and Cell Cycle Analysis Software (Phoenix Flow

Systems, Inc., San Diego, CA) was used to compute peak numbers, means, coefficients of variation (CVs), and peak ratios from the population DNA-fluorescence distributions, substituting G1 and G2 phase descriptions with 1C and 2C.

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6.4 Results

6.4.1 Cyst Development and Excystment. Cysts typically were found in the petri dishes in a dense but somewhat patchy distribution after 3–5 days of incubation in the aquaria. P. shumwayae can be selective in encystment location (Parrow et al. 2002); it was assumed that the aquarium cyst populations were adequately sampled in the dishes, but preferential encystment on other aquarium surfaces may have created bias (Holt et al.

1994). Rinsing the dishes in deionized water limited the dinoflagellate populations to cysts that were observed for development and excystment. Fission in these P. shumwayae isolates occurred in nonmotile cysts that commonly produced two to eight swimming offspring (Parrow et al. 2002). Primary (pre-division) reproductive cysts were smooth- walled, roughly spherical, and varied in size (ca. 13–25 µm in diameter) (Fig. 6.1A). A distinct nucleus with condensed chromosomes was visible in many cysts, while in others it was partially or wholly obscured by a distended grayish food vacuole (Fig. 6.1A).

Apparently depending on initial size, the contents of primary reproductive cysts most commonly underwent 1-3 sequential nuclear and cytoplasmic division(s). The outer wall of the primary cyst was not included in division (Parrow et al. 2002) (Fig. 6.1B). Protoplast fission in larger cysts resulted in distinct internal secondary and sometimes tertiary cysts, in which flagellated offspring were produced by a further morphogenic division (as in

Elbrächter and Drebes 1978, Drebes and Schnepf 1982,1988, Parrow et al. 2002).

The protoplast of smaller primary cysts (ca. 13-17 µm) underwent a single nuclear and cytoplasmic division within the primary cyst outer wall, resulting in two offspring cells

(Fig. 6.1C). In medium-sized primary cysts (ca. 17-20 µm), the first nuclear division was

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followed by a pronounced equatorial protoplast cleavage forming two distinct secondary cysts, each internal to the outer primary cyst wall (Fig. 6.1D). A further division occurred

2–4 h later within each secondary cyst, resulting in two pairs of offspring cells. The protoplast of larger primary cysts (ca. 20–25 µm) divided to form two large secondary cysts

(Fig. 6.1E). Each secondary cyst protoplast cleaved again within 3–6 h, forming a group of four distinct tertiary cysts (Fig. 6.1F). On some occasions the outer primary cyst wall was no longer visible by this stage and may have dissolved. A few hours later a further division occurred within each tertiary cyst, resulting in a total of four pairs of offspring cells.

As reproductive cysts developed and division(s) occurred, the food vacuole grew smaller and was not divided, so that the residual body was inherited by only one offspring cell (Fig. 6.1F). In cysts with sequential divisions, each occurred in the same plane but perpendicular to the last. The final cell division (or only division in the case of smaller primary cysts) within primary, secondary, or tertiary cysts was zoosporogenic and produced two offspring that excysted as motile biflagellated cells. Thus, primary, secondary, or tertiary cysts functioned as zoosporangia (dinosporangia) (as in Drebes and Schnepf 1988), and the final number of flagellated offspring produced depended on the number of cysts formed before. Release of swimming cells typically occurred 3–5 h after the final division.

Excystments from paired secondary cysts and grouped tertiary cysts were asynchronous, but usually occurred within ca. 1 h of each other. Prior to excystment, flagella were often seen beating slowly inside the cyst. Upon excystment the tightly compressed offspring pair squeezed through a fissure in the cyst wall and assumed normal flagellated cell shape once they emerged. The dinoflagellate pair beat their flagella and separated, but remained attached to the remains of the cyst wall by the distal ends of their longitudinal flagella for

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several seconds before swimming away. Offspring that had inherited the residual food body occasionally egested it before departing (Calado and Moestrup 1997). Excysted dinoflagellates were typically 9–12 µm in length, biflagellated, and rapidly swimming (Fig.

6.1G).

Rarely it was observed that one secondary cyst of a pair released two flagellated offspring while the other divided again to form two tertiary cysts, each of which later released two swimming cells. In one observation a single tertiary cyst of a group formed two quaternary cysts, each of which later released two flagellated offspring. In these cases it was the cyst that retained the residual food body that underwent the asynchronous extra division (as in Drebes and Schnepf 1982). Thus, in our observations on these cultures, each encysted progenitor cell ultimately produced 2, 4, 6, 8, or 10 flagellated offspring. Non- reproductive temporary cysts from which a single flagellated cell emerged also occurred, and these were generally smaller than primary reproductive cysts (Fig. 6.1H). The sequences of reproductive cyst development most commonly observed in these isolates are summarized in Figure 6.2.

6.4.2 Population Synchronizations. Initial cyst populations isolated from the aquaria were in all stages of reproductive development, and were effectively synchronized without apparent harm to the dinoflagellates by 72 h immersion in deionized water. A robust and highly synchronous excystment response occurred within a few hours after resumption of previous culture salinity. The swimming dinoflagellates were decanted away from the remaining adherent cysts, which ended recruitment and resulted in isolated populations that consisted almost entirely of newly excysted flagellated cells. Some small cysts consistent with non-reproductive temporary cysts also occurred, but they were not

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A B C D

N FV

E F G

Figure 6.1

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Figure 6.1. Light micrographs of Pfiesteria shumwayae vegetative cells: (A) Primary reproductive cysts, showing the maximum size difference observed. The nucleus (N) occupies the majority of the small cyst on the left. The large primary cyst on the right is filled with a grayish food vacuole (FV). (B) Primary cyst after nuclear division, during cytoplasmic division. The outer membrane of the primary cyst was not included in division(s) (arrow). (C) A tightly compressed offspring pair prior to excystment (D) Two distinct secondary cysts, internal to the outer primary cyst wall (arrow). The right cell inherited the residual food body (darkened rounded structure). Four flagellated cells were produced, two from each secondary cyst. (E) Lower, two large secondary cysts before another fission produced tertiary cysts. Arrow = primary cyst outer wall. The upper cell was a temporary non-reproductive cyst. (F) Tertiary cysts, each of which had undergone a final internal zoosporogenic division. The upper cell inherited the residual food vacuole

(grayish spherical body). The outer wall of the original primary cyst was still apparent

(arrows). (G) An excysted biflagellated cell (zoospore, dinospore). (H) A small non- reproductive temporary cyst, with distinct chromosomes. Scale bars = 10 µm.

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excystment primary cysts A

secondary cysts excystment

tertiary cysts excystment

Figure 6.2. Diagram of the most commonly observed Pfiesteria shumwayae reproductive cyst development (gray circles, food vacuoles; black ovals, nuclei): (A) Smaller primary cysts (ca. 13-17 µm) underwent a single internal zoosporogenic division and produced two flagellated offspring. (B) Medium-sized primary cysts (ca. 17-20 µm) underwent distinct protoplast fission, producing two secondary cysts. Zoosporogenic division then occurred in each secondary cyst, and four flagellated offspring were produced. (C) The protoplast of the largest primary cysts (ca. 20-25 µm) underwent two sequential fissions to form secondary, then tertiary cysts. Zoosporogenic division occurred in each tertiary cyst, and eight flagellated offspring were produced. All fissions were internal to the original outer wall of the primary cysts, unless it dissolved. Food vacuoles shrank as division(s) occurred and were not divided among offspring. Asynchronous fissions and excystments also occurred (not shown; see text).

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abundant. Flagellated cell densities depended on initial cyst abundance and excystment rate, and typically were 104 – 105 cells mL-1.

6.4.3 Gamete Fusion and Planozygotes. P. shumwayae flagellated offspring pairs occasionally remained connected for a short time after excystment (Parrow et al. 2002) and misdivisions were a possible source of paired cells (Drebes and Schnepf 1988), so careful observations were needed to clearly identify fusing cells. Recognition behavior and initial contact between individual gametes were not observed. In all isolates, motile cell pairs in apparent fusion occurred among the newly excysted flagellate populations during the first several hours after isolation. Coupled cells were fast swimming and erratic, but generally swam in a spiraling pattern. Gamete fusion appeared to be much more abundant in the mixed clone culture compared to the clonal isolates, so the mixed clone was used to examine the process. Gametes were approximately isogamous and appeared to begin fusion by their episomes while in a perpendicular or oblique alignment (Fig 6.3A). As fusion proceeded, one gamete was progressively absorbed into the other. Resulting planozygotes with two trailing flagella (Fig. 6.3B) were produced in isolations from the mixed clone culture. Planozygotes were within the size range of other swimming cells and had a similar shape and rapid motility. SEM investigation of the mixed clone swimming cell isolations also showed apparent isogamous gamete pairs (Fig. 6.4A) and planozygotes with two longitudinal flagella and a single transverse flagellum (Fig. 6.4B, C). Cell pairs in apparent fusion were rare in the clonal isolates. Although resulting planozygotes were not found by microscopy in the clonal isolates, it is possible that they had occurred rarely and were undetected. Fusing cells were observed only during the first several hours after synchronized excystment, indicating that a subpopulation of fully differentiated gametes

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emerged from some cysts and the remaining flagellated cells lacked the potential or impetus for fusion under these conditions.

6.4.4 DNA Measurements. Flow cytometric analyses of the isolates fixed 6 h after release from cysts showed bimodal population DNA distributions composed of abundant

1C DNA cells and less abundant 2C DNA cells (Fig. 6.5). Clonal isolates differed slightly from one another in mean 1C DNA fluorescence. The 2C/1C peak means ratio for each isolate was 1.8, and 1C peak CVs were between 7.4-8.4%. In the clonal isolates, 2C DNA cells were less than 5% of the total flagellate populations, and most typically comprised less than 3%. In contrast, swimming dinoflagellates with 2C DNA comprised between 10-16% of the total populations from the mixed clone culture over three separate flagellated cell isolations. Cells with measured intermediate DNA content were less than 2% of the total flagellate populations for clonal isolates, and less than 5% of the total for the mixed clone isolations. Forward scatter signals (a sizing parameter; Shapiro 1995) from 2C DNA cells fell within the same range as cells with 1C DNA (Fig. 6.5). Thus, the 2C DNA dinoflagellates were not measurably larger or more optically dense.

Dinoflagellates with measured 1C DNA corresponded to the biflagellated cells that composed the great majority of each population. In preparations from the mixed clone culture, many if not all 2C dinoflagellates were likely the diploid products of fusion, based on observations of syngamy and the occurrence of planozygotes. Likewise, some or all of the more rare 2C dinoflagellates measured in clonal cultures may have been planozygotes that were not detected microscopically. In each culture, the percentage of 2C cells in the populations did not significantly increase between 6 and 24 h (Fig. 6.6). This supported observations that a finite number of fusions occurred soon after excystment, and also provided evidence that cannibalisms were not responsible for 2C DNA cells. Nearly all

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A B

Figure 6.3. Light micrographs of Pfiesteria shumwayae sexual cells from the mixed clone isolations: (A) Gametes, fusing by their epicones. The lower gamete progressively engulfed the upper gamete. (B) Planozygote with two trailing flagella. Scale bars = 10 µm.

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A C

Figure 6.4. SEMs of P. shumwayae sexual cells from the mixed clone isolations: (A)

Presumed gametes in early fusion. (B) Planozygote, antapical view, with a single transverse flagellum and two longitudinal flagella, and a deflated peduncle emerged from the distal pore. (C) Small planozygote, lateral view, with a single transverse flagellum and two longitudinal flagella. Scale bars = 5 µm.

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North Carolina AB

102716 1C – 97.6% 2C – 2.0% CV = 8.3 1C 2C beads 1C 2C

Maryland C D 102537

1C – 94.4% 2C – 4.3% CV = 7.8 1C 2C 1C 2C

New Zealand EF 102558 1C – 97.3%

2C – 2.1% CV = 8.2 1C 2C 1C 2C

Combined Clones G H 1C – 78.8%

2C – 16.4% CV = 8.4 1C Events 2C

Forward Scatter 2C 1C

Log DNA ( AU ) Linear DNA ( AU )

Figure 6.5

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Figure 6.5. Representative logarithmic flow cytometric DNA-fluorescence profiles (A, C,

E, G) and forward scatter signals versus linear DNA-fluorescence (B, D, F, H) scatter plots of P. shumwayae flagellated cell populations, fixed 6 h after excystment and isolation.

Measurements were taken on at least 104 dinoflagellates per analysis. Beads were fluorescence intensity standards (see text). Percentages of 1C and 2C relative DNA and 1C peak CVs are listed for each analysis shown. (A–B) North Carolina clonal isolate 102716.

(C–D) Maryland clonal isolate 102537. (E–F) New Zealand clonal isolate 102558. (G–H)

Combined clone culture. Dinoflagellates with 2C DNA occurred in each clone, but were significantly more abundant in the combined clone isolations. Forward scatter signals from

2C DNA dinoflagellates fell within the same range as signals from 1C DNA dinoflagellates.

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T = 6 h A T = 24 h B 1C – 95 % 1C – 96 % 2C – 4 % 2C – 3 % 1C

Events beads 2C

Flagellated Cell DNA Fluorescence (AU)

Figure 6.6. Representative logarithmic flow cytometric DNA-fluorescence profiles for

Pfiesteria shumwayae dinoflagellates from the same flagellated cell isolation, fixed 6 h (A) and 24 h (B) after synchronous excystment. Measurements were taken on at least 104 dinoflagellates per analysis. The 1C and 2C DNA subpopulations did not change significantly in relative abundance between the two sampling times.

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sorted dinoflagellates with 2C DNA from the mixed clone culture sampled 6 h after release from cysts had a single hyposomal brightly fluorescent nucleus and only comparatively faint cytoplasmic fluorescence, which suggested that karyogamy occurred early in planozygotes and the nucleus accounted for the measured 2C DNA. Dinoflagellates with

2C DNA from the clonal isolates were also primarily uninucleate. In all cultures, a small number of sorted 2C DNA cells were abnormally shaped and had two nuclei (Fig 6.7A), and may have been in the process of fusion at the time of fixation.

6.4.5 Nuclear Cyclosis. Many primary cysts collected from the mixed clone aquarium over the investigation period exhibited distinctive and lengthy chromosome movements prior to nuclear division (Fig. 6.7B, video link, cyclosis1.mov). This conspicuous phenomenon corresponded to nuclear cyclosis, a process known to occur specifically in the zygotic nuclei of some dinoflagellate species and thought to accompany the homologous chromosome pairing of meiotic prophase (von Stosch 1972, 1973, Barlow and Triemer 1988, Pfiester 1989, Kita et al. 1993). Cysts with nuclear cyclosis did not otherwise differ from primary cysts without nuclear cyclosis. During nuclear cyclosis the rounded nucleus was roughly 12 µm in diameter, and the chromosomes underwent turbulent swirling and mixing for more than 8 h in some observations. The chromosome movements ended gradually, and a few hours later a nuclear and cytoplasmic division occurred.

Cysts with nuclear cyclosis varied in size. In smaller cysts the food vacuole was minor and the moving chromosomes occupied most of the cell volume (Fig. 6.7B, C). The protoplast in these cysts divided once (Fig. 6.7D), and two flagellated offspring were released around 9–12 h after nuclear cyclosis ended. Nuclear cyclosis also occurred in larger cysts with greater cytoplasmic and vacuolar reserves (Fig. 6.7E, video link,

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cyclosis2.mov). In these cysts the nuclear division after cyclosis (Fig. 6.7F) was followed by fission, forming two secondary cysts (Fig. 6.7G). Each secondary cyst released two flagellated offspring about 12 h later (Fig. 6.7H, I). Nuclear cyclosis was found more commonly in smaller primary cysts that produced two swimming cells, although the distended food vacuoles in many larger cysts may have obscured the process.

It is common for dinoflagellate species that encyst for mitosis to also do so for zygotic division(s) (reviewed in von Stosch 1972, 1973, Beam and Himes 1980). Since unambiguous planozygote encystment was not observed, and no other characteristics were apparent for distinguishing sexual from asexual cysts, cysts with nuclear cyclosis were assumed to have been zygotic and the ensuing division(s) meiotic (as in Spero and Morée

1981, Drebes and Schnepf 1988, Ucko et al. 1997, Parrow et al. 2002, Parrow and

Burkholder accepted). Cysts with nuclear cyclosis rarely were found in some clonal cultures, and not at all in others. In contrast, nuclear cyclosis was common in cysts collected from the mixed clone culture over similar periods of observation.

6.4.6 Feeding Treatments and Encystment. Newly excysted dinoflagellates inoculated into flasks with larval fish swarmed about the fish with an obvious chemotactic attraction (Fenchel and Blackburn 1999, Cancellieri et al. 2001). The dinoflagellates attached to the fish surfaces with their peduncles and ingested fish cytoplasm as previously described (Burkholder and Glasgow 1995, 1997, Burkholder et al. 2001a,b, Vogelbein et al.

2001). The dinoflagellates also fed on free-floating cells that had apparently disassociated from the fish (Burkholder and Glasgow 1997), and a cytoplasmic disturbance was sometimes visible at the point of peduncle penetration (Fig. 6.7J). It is possible that the peduncle applies or injects some substance that aids penetration or ingestion, as suggested

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for Amyloodinium Brown & Hovasse (Lom 1981, Cachon and Cachon 1987) and tube- feeding Paulsenella Chatton and Crypthecodinium species (Drebes and Schnepf 1988,

Ucko et al. 1997). Fish cytoplasm could be seen streaming into the feeding dinoflagellates, which enlarged as ingestion occurred. Feeding by individual dinoflagellates on the fish lasted seconds to minutes, and many appeared to attach and feed more than once. After feeding the peduncle was stretched thin, detached from the fish, and retracted by the departing dinoflagellate. Swimming cells in various states of engorgement with grayish food vacuoles full of fish cytoplasm were observed within minutes of exposure to fish.

Some became greatly enlarged (up to 26 µm in length) with a distended food vacuole filling most of the cell (Fig. 6.7K).

DNA staining of the dinoflagellates after feeding demonstrated that the food vacuoles were rich in fish nucleic acids (Fig. 6.7L). Nile red staining showed that the vacuoles were also replete with fish lipids (Fig. 6.7M). Phagotrophic uptake of fish nucleic acids by the dinoflagellates was measured at the population level with flow cytometry (Fig.

6.8). Within 4 h of exposure to fish under these conditions, nearly half of the swimming population contained ingested nucleic acids greater than the basal amount attributable to the

1C nucleus (Fig. 6.8C). By 6 h and at 8 h the majority of the motile population (>70 %) contained ingested DNA (Fig 6.8D, E). After 8 h in this treatment, encystment of satiated cells reduced the proportion of swimming cells with increased cellular DNA from phagotrophy (Fig. 6.8F,G).

In synchronized dinoflagellate populations given fish, encysted cells with replete food vacuoles began to appear on the vessel floor within the first hour of exposure to fish.

Encystment was observed on occasion, wherein the engorged dinoflagellate swam along

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very close to the vessel floor, suddenly lost motility, rounded off in profile, and within seconds formed the typical rounded cyst shape. Ecdysis of thecal plates was not observed.

Cysts varied considerably in vacuolar content and cell size, as had been observed for cysts collected from the aquarium cultures. Within 6–8 h under these conditions, a large portion of the dinoflagellates had encysted. Cysts were found abundantly on the vessel bottom in a patchy distribution which suggested that active encystment location selection commonly occurred (as in Lombard and Capon 1971, Ucko et al. 1997, Parrow et al. 2002).

The culture volume was then decanted and the vessel was rinsed gently with deionized water and refilled with sterile 15 ppt-salinity artificial seawater. Since the time of encystment was known to within several hours, a general approximation of reproductive cyst development times was possible. In general, smaller primary cysts released two flagellated offspring within about 24 h. Medium-sized primary cysts formed secondary cysts and generally required an additional 3–6 h to produce flagellated offspring. Larger primary cysts formed secondary and then tertiary cysts, and required an additional 4–6 h to release swimming cells. A metabolic phase prior to division(s) occupied a major portion of the time spent encysted. During this time the food vacuole shrank as digestion apparently occurred, and cytoplasm increased. Once division(s) began, it progressed relatively quickly

(ca. 3–4 h per division) and equally. Release of swimming cells typically occurred within a few hours of the final (zoosporogenic) division.

As described, flagellated cell populations derived from the mixed clone culture produced abundant planozygotes. Nuclear cyclosis was also found in many of the primary cysts produced by these populations following exposure to fish. Cysts with nuclear cyclosis were found as early as 9 h after introduction of fish. These cysts presumably had been

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formed by the planozygotes and contained variable amounts of fish cytoplasm, indicating that planozygotes had been phagotrophic (Glasgow et al. 2001). As in previous observations, nuclear cyclosis lasted for over 8 h and subsequent division(s) produced two or, less commonly, 4 flagellated offspring.

Isolated dinoflagellate populations left to starve did not produce abundant reproductive cysts. After several days, most of the dinoflagellates were diminished in size and appeared to consist of little more than motile, membrane-bound nuclei (Parrow et al.

2002). Observations of cannibalism on (perhaps senescent) cells increased, and small temporary-like cysts became more abundant. Swimming cells were rare after four weeks.

Starved populations were placed in darkness at 22ºC for 3 months, after which no swimming cells were found, only a low abundance of small cysts similar to that shown in

Fig. 6.1H but often darker in color. It was unknown whether these cysts had been formed sexually or asexually, but it was apparent based on the low abundance that not all dinoflagellates had persisted as cysts. Larval fish were placed in half of the flasks, which stimulated the appearance within 24 h of a sparse population of swimming cells that fed on the fish and reproduced. No swimming cells were observed in the duplicate flasks without fish. The duplicates were returned to storage conditions for two more months, after which addition of fish into the flasks stimulated the appearance of swimming cells within 24 h.

This response suggested that some or all of the cysts were quiescent rather than resting, and were suspended from growth by lack of food rather than exhibiting a required dormancy

(Anderson et al. 1995).

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B C E A D N

N N

F G H I

J K

L M

Figure 6.7

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Figure 6.7. Light and fluorescence micrographs of Pfiesteria shumwayae sexual and feeding cells: (A) A sorted 2C DNA cell with two nuclei and an irregular shape, presumed to have been fusing gametes. (B) Primary cyst with the nucleus (N) in cyclosis. C-D, in series: (C) Small primary cyst with the nucleus (N) in cyclosis. (D) The same cyst, after division. Two flagellated offspring were produced 12 h after nuclear cyclosis ceased. E-I, in series: (E) Larger primary cyst with nucleus (N) in cyclosis, and a replete grayish food vacuole (below) (F) Nuclear division, 6 h after nuclear cyclosis had ceased. (G) Protoplast fission, producing two secondary cysts. The lower cyst received the residual food body.

(H) The lower secondary cyst remained after the upper had released two flagellated offspring. (I) Two offspring dinoflagellates were released from the lower cyst 1 h later.

The lower dinoflagellate inherited the residual food body. (J) Dinoflagellate (right) beginning to feed on a disassociated spherical fish cell (left) using its extended peduncle.

(K) A flagellated cell (dorsal view, transverse flagellum was displaced out of the cingulum

(C)) that had become greatly enlarged from ingestion of fish cytoplasm. A grayish food vacuole (FV) filled most of the cell. (L) Flagellated cell after feeding, with stained DNA.

The large food vacuole filled most of the cell and was rich in fish DNA. (M) Flagellated cell after feeding, with stained lipids. The large food vacuole was rich in fish lipids. Scale bars = 10 mm.

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Synchronized A T = 2 h B T = 4 h C T = 6 h D T = 0 h

beads 2C

Events T = 8 h E T = 10 h F T = 12 h G

Flagellated Cell DNA Fluorescence ( AU )

Figure 6.8. Flow cytometric DNA profile time series of a synchronized Pfiesteria shumwayae flagellated cell population fed larval fish. Measurements were taken on at least

104 dinoflagellates per time period; beads were fluorescence standards (see text). (A)

Synchronized population before larval fish were introduced. (B) 2 h, (C) 4 h, (D) 6 h, (E) 8 h, (F) 10 h, and (G) 12 h after introduction of fish.

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6.5 Discussion

6.5.1 Asexual Reproduction. The reproductive cycle observed in these isolates is diagrammatically summarized in Fig. 6.9. Biflagellated cells (zoospores, dinospores) were the predatory stage of the asexual cycle (Fig. 6.9-1). These cells could form non- reproductive temporary cysts (Fig. 6.9-2) perhaps in response to unfavorable stimuli. After feeding, they became nonmotile by forming primary cysts (Fig. 6.9-3). Primary cyst size appeared dependant on the amount of ingested material and cytoplasm. The protoplast of smaller primary cysts underwent a single zoosporogenic division and released two flagellated offspring (Fig. 6.9-4). The first protoplast fission of medium-sized primary cysts produced two distinct secondary cysts within the outer wall of the primary cyst.

Zoosporogenic division then occurred in each secondary cyst, producing four flagellated offspring (Fig. 6.9-5). Occasionally an extra asynchronous fission of secondary or tertiary cysts occurred prior to zoosporogenic division (Fig. 6.9-6). Larger primary cysts underwent two sequential protoplast fissions, forming four tertiary cysts in which zoosporogenic division occurred (Fig. 6.9-7). Thus, apparently depending on progenitor cell size, primary, secondary, or tertiary cysts functioned as zoosporangia, and each progenitor cell most commonly produced 2, 4, or 8 swimming offspring. P. shumwayae vegetative cells with amoeboid and chrysophyte-like morphologies have been described

(Glasgow et al. 2001), but are not typically considered forms of cell reproduction

(Burkholder and Glasgow 2002). The cycle of proliferation depicted here is therefore presented as an incomplete, but major reproductive pathway in P. shumwayae.

P. shumwayae cysts with sequential divisions demonstrated a form of reproductive sporogenesis similar to palintomy, defined as repeated binary fissions without an

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intermediate stage of nutrition and growth, leading to the formation of complete identical products of reproduction (Elbrächter and Drebes 1978). Palintomy is common in ectoparasitic dinoflagellates (Drebes 1984). For example, the dinoflagellate ectoparasite of copepod eggs Dissodinium Klebs (Pascher) also produces primary cysts after feeding, and then internal secondary cysts in which dinospores are formed (Drebes 1978, Elbrächter and

Drebes 1978). Tube-feeding (including pedunculate) heterotrophic dinoflagellate species show a gradual continuum of modifications in morphology and physiology from predation to parasitism (Cachon and Cachon 1987, Coats 1999). P. shumwayae falls along that continuum, but differs from species considered true ectoparasites in that the cells are omnivorous (Glasgow et al. 2001) and remain undifferentiated during feeding (flagella are retained) (Parrow et al. 2002).

P. shumwayae reproduction was similar to that of Paulsenella species, ectoparasites of marine diatoms. As in P. shumwayae, the satiated Paulsenella trophont also forms a primary cyst which often divides to form two secondary cysts, one or both of which may divide again forming tertiary cysts (Drebes and Schnepf 1982, 1988). Secondary and tertiary cysts are formed internal to the outer primary cyst wall, and inside each cyst a final morphogenic division produces a flagellated offspring pair. Like in P. shumwayae, smaller primary cysts of Paulsenella kornmannii Drebes and Schnepf release only two flagellated offspring.

SSU rDNA sequence analysis indicates a close phylogenetic relationship between

Pfiesteria and Amyloodinium, dinoflagellate ectoparasites of marine and estuarine fish

(Litaker et al. 1999). The two genera also have similar plate tabulations (Landsberg et al.

1994, Stiedinger et al. 1996). The pattern of reproduction in P. shumwayae was also

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similar to that of Amyloodinium ocellatum Brown & Hovasse; the two differ mainly in the size of the reproductive cyst and the number divisions that occur within it. The flagellated cells of A. ocellatum are roughly the same size as those of P. shumwayae, but upon feeding on fish surficial tissues for a period of days they develop into sac-like trophonts up to 150

µm in length (Lawler 1977, Lom 1981). Upon maturity, or death of the host, the trophont detaches and forms a cyst. The protoplast of this cyst undergoes sequential, equal, and perpendicular fissions into 2, 4, 8, 16, 32, 64, and then often 128 identical membrane-bound cysts, with all fissions occurring internal to the outer membrane of the progenitor cyst unless it dissolves (Brown 1934). A final morphogenic division then occurs within each cyst, producing two dinospores that excyst together as a tightly compressed pair (Brown

1934, Nigrelli 1936). Primary reproductive cysts of A. ocellatum also vary in size apparently as a result of incomplete feeding, with the smallest being around 30 µm in diameter. Smaller reproductive cysts undergo fewer fissions prior to formation of flagellated cells, with the smallest releasing flagellated cells after only the 8-cyst stage

(Brown 1934, Nigrelli 1936). P. shumwayae flagellated cells fed on fish more briefly than does A. ocellatum, and swelled during feeding to a smaller size (ca. < 26 µm in length).

Satiated cells then formed reproductive cysts that underwent fissions very similar to but fewer in number than those of A. ocellatum. Variability in the number of offspring produced occurs in each species, apparently related to the amount of food ingested. This phenomenon has also been described for Dissodinium (Drebes 1978, Elbrächter and Drebes

1978) and Paulsenella (Drebes and Schnepf 1982, 1988).

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e nt n e cy m s st tm y 1 e c n ex t 2

i excystment s

i 7 o

n f is s io n

4 6 5 fission

Figure 6.9. Diagram of asexual reproduction observed in these P. shumwayae cultures

(gray ovals, nuclei): (1) Biflagellated cell (zoospore or dinospore), the main feeding stage observed. (2) Non-reproductive temporary cyst. (3) Primary reproductive cysts. (4)

Smaller primary cyst with two offspring. (5) Secondary cysts with four offspring. (6)

Asynchronous divisions (see text). (7) Tertiary cysts with eight offspring.

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Reproduction in P. shumwayae was similar to that of P. piscicida (Parrow et al.

2002) and related cryptoperidiniopsoid species (Parrow and Burkholder accepted), which also divided in cysts. Reproductive cysts of P. piscicida and cryptoperidiniopsoids typically produce two flagellated offspring, but sequential divisions producing two secondary cyst then four offspring also occur. Reproductive progression to tertiary cysts appeared to be less common in P. piscicida and cryptoperidiniopsoids as compared to P. shumwayae, at least when cryptophyte microalgae were provided as food. These taxa are omnivorous, and food quality probably influences the number of offspring commonly produced. In P. shumwayae, primary cysts that progressed to secondary and tertiary cysts were relatively more common in this study with fish prey than with cryptophyte microalgae as prey (Parrow et al. 2002). Fish may have been more nutritive to P. shumwayae than cryptophytes, or a greater quantity of fish cytoplasm may have been ingested prior to primary cyst formation. Pfiesteria piscicida also has been reported to divide while motile

(Parrow and Burkholder 2002). Gamete misfusion and late separation of temporarily conjoined offspring (Parrow et al. 2002) resemble division and may have accounted for those apparent motile divisions. Apparent single offspring occasionally released from secondary cysts (Parrow et al. 2002) likely were tightly compressed pairs, based on the observations of this study. In each taxon, starvation in cultured populations leads to the formation of apparently quiescent cysts of unresolved ploidy (Parrow et al. 2002, Parrow and Burkholder accepted, this study).

6.5.2 Synchronizations and Flow Cytometry. The forced-quiescence synchronization technique used here resulted in highly synchronous excystments of flagellated cells. 1C peak CVs were lower than previously obtained values (Parrow and

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Burkholder 2002) and similar to published values for photosynthetic dinoflagellates

(reviewed in Parrow and Burkholder accepted). The data indicated that cytoplasmic DNA had been minimized in these heterotrophic dinoflagellates by the treatment, and that the measured DNA was primarily attributable to the dinoflagellate nuclei. Thus, more precise

DNA measurements and more interpretable DNA distributions were possible. The DNA distributions consisted of 1C and 2C relative nuclear DNA subpopulations. Dinoflagellates with 1C DNA were assumed to be functionally haploid and corresponded to the biflagellated cells that comprised the majority of each population. Dinoflagellates with 2C

DNA were much less abundant and were more difficult to interpret. Relatively abundant planozygotes were produced in flagellated cell isolations from the mixed clone culture, so many if not all measured 2C cells in those preparations were assumed to represent diploid planozygotes.

6.5.3 Sexuality. The sexual cycle observed more completely in the mixed clone culture is summarized in Fig. 6.10. Approximately isogamous gametes emerged from cysts and paired (Fig. 6.10-1). Fusion resulted in a planozygote with one transverse and two longitudinal flagella (Fig. 6.10-2). Planozygotes were the phagotrophic stage of the sexual cycle, and presumably encysted for division(s). Cysts with a distinct nuclear cyclosis occurred, and thus were assumed to be zygotic cysts formed by planozygotes (Fig. 6.10-3).

Under light microscopy, these cysts did not differ morphologically from primary cysts without nuclear cyclosis, and they exhibited no dormancy. After nuclear cyclosis, the protoplast of smaller zygotic cysts underwent a single division, assumed to be meiotic because it followed nuclear cyclosis (Fig. 6.10-4), and two uninucleate flagellated offspring emerged from the cyst (Fig. 6.10-5). It was unknown whether these two dinoflagellate

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offspring had nuclei with 2C DNA and required a further division (i.e. sister chromatid segregation) before reentering mitotic interphase, as adherence to the pattern of conventional meiosis would predict (Raikov 1982, 1995). If so, then the second postzygotic division presumably would have occurred in the next primary cyst, perhaps after feeding. The protoplast of larger zygotic cysts underwent two sequential divisions after nuclear cyclosis, and released four flagellated offspring (Fig. 6.10-6). Production of two flagellated offspring after nuclear cyclosis was much more commonly observed.

Progression to the tertiary cyst stage and release of eight flagellated offspring following nuclear cyclosis was not observed, but probably occurs if a planozygote acquires sufficient nutritional reserves for the extra division. P. shumwayae sexual cells (gametes, planozygotes) that acquire amoeboid morphology have been described (Glasgow et al.

2001). Therefore, this sexual cycle is presented as incomplete and only encompasses flagellated cells and cysts.

Sexual stages (fusing gametes, planozygotes, and cysts with nuclear cyclosis) were found more abundantly and completely in the mixed clone culture compared to individual clonal isolates. This observation was partly supported by population DNA measurements showing greater abundances of 2C DNA cells in swimming cell isolations from the mixed clone culture, many or all of which were newly formed diploid planozygotes based on microscopy. Swimming 2C DNA cells occurred at lower abundances (typically < 3%) in the clonal isolations, as previously reported for P. shumwayae (Parrow and Burkholder

2002, Parrow et al. 2002). Planozygotes were not observed in clonal isolations, and nuclear cyclosis was found only rarely in a few of the clonal cultures. Dinoflagellates with measured 2C DNA in those populations may have been be diploid planozygotes that were

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gametogenesis f us ion

1 Vegetative

cycle e x c y s t m 2 e

e n

n c

t 5 y s

6 N

fis 3 si on division 4

Figure 6.10. Illustration of the proposed sexual cycle of P. shumwayae (gray ovals, nuclei; broken arrow indicates an inferred relationship): (1) Coupled gametes. (2)

Planozygote. (3) Zygotic cyst with nuclear cyclosis. (4) Production of two flagellated offspring after nuclear cyclosis (more common). (5) It was unknown whether these offspring required a second postzygotic division prior to re-entry into mitotic interphase

(see text). (6) Production of two secondary cysts after nuclear cyclosis, with release of four flagellated offspring (less common).

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missed by microscopy, haploid G2 cell cycle phase cells that delayed mitosis and excysted, or persistent autodiploid subpopulations resulting from chromosome endoreplication (as in

Walker 1982). In the mixed clone culture, relatively abundant cysts with nuclear cyclosis occurred, and they commonly released two flagellated cells after a single division. The conventional pattern of meiosis would predict that two dinoflagellates released from a zygotic cyst would each have a 2C (but not technically diploid) nucleus containing previously replicated chromosomes. As with the other potential sources of non-diploid 2C

DNA flagellated cells, this remains a hypothesis for future examination.

Although it was not possible to conclude from this research that these P. shumwayae clones were heterothallic, it was clear that mixing clones enhanced the sexual cycle in these cultures. Mating types in heterothallic dinoflagellates have typically been regarded as binary (plus or minus; Pfiester and Anderson 1987). However, investigations of mating compatibility between geographically distinct clones have revealed more complex interactions. Apparent homothallic and heterothallic strains occur within some morphospecies (Hayhome et al. 1987, Steidinger et al. 1998, Burkholder et al. 2001a).

Apparent zygotes sometimes occur in clonal cultures of heterothallic species (von Stosch

1973, Destombe and Cembella 1990), and sexual compatibility factors may change over time in culture (Blackburn et al. 1989). In some dinoflagellates, the efficiency of sexual reproduction has been found to vary between clonal crosses and within clones, with sterility factors influencing compatibility not only at gamete recognition and fusion but also zygote and offspring viability (Blackburn et al. 1989, 2001, Destombe and Cembella 1990). The rare fusing cells observed by microscopy and potential zygotes detected by flow cytometry in clonal P. shumwayae cultures may have had reduced viability. The single clonal gamete

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fusion in P. shumwayae reported by Parrow et al. (2002) resulted directly in a nonmotile cell that did not develop, suggesting that self-incompatibility beyond gametic recognition may occur in P. shumwayae clonal cultures (as in Destombe and Cembella 1990). It was unknown in this study which clonal subpopulations were interbreeding in the mixed clone culture. Further determinations on mating compatibility in P. shumwayae will require examination of single crosses between clones. Reproductive compatibility and success in

P. shumwayae may prove more difficult to assess than in photosynthetic dinoflagellate species that produce morphologically distinctive, long-lived zygotes (Yoshimatsu 1981,

Anderson et al. 1988, 1995, Blackburn et al. 1989).

The sexual cycle of P. shumwayae is similar to that of related species P. piscicida and cryptoperidiniopsoids (Parrow et al. 2002, Parrow and Burkholder accepted). In each taxon, gametes emerge from cysts and are typically smaller than most vegetative cells.

However they are not easily distinguished prior to fusion because they fall within the size range of vegetative cells. Gametes typically fuse quickly (in < 1 h) and were primarily isogamous in these cultures. Since one gamete of a pair is progressively engulfed by the other, later stages of fusion can resemble anisogamy (as in Montresor 1995). Planozygotes have two trailing flagella, and Pfiesteria planozygotes have a single transverse flagellum

(Burkholder and Glasgow 1997, Burkholder et al. 2001b). The planozygotes of each taxon are within the size range of vegetative cells and are rapidly swimming, making individual planozygotes difficult to continuously observe. Planozygotes commonly engage in phagotrophy, as described for Katodinium (Gymnodinium) fungiforme Anissimova and

Paulsenella chaetoceratis (Paulsen) Chatton (Spero and Morée 1981, Drebes and Schnepf

1988). Zygotes presumably encyst for meiotic division(s).

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Identification of zygotic cysts was problematic in P. shumwayae because they exhibited no observed dormancy in growing cultures, and apparently did not differ under light microscopy from asexual cysts in external morphology or in the division process that produced offspring. However, nuclear cyclosis was observed upon direct observation of living cysts, and was used to identify encysted zygotes and subsequent meiotic division.

Structures diagnostic of meiotic prophase in other eukaryotes, such as the synaptonemal complex and chiasmata, have not been clearly demonstrated in dinoflagellates. Nuclear cyclosis has been shown to occur specifically before the first zygotic division in some dinoflagellate species, and accompanies a pairing of the chromosomes (von Stosch 1972,

1973, Coats et al. 1984, Barlow and Triemer 1988). It seems likely that dinoflagellate nuclear cyclosis is homologous to the unique chromosome movements that accompany homologue pairing during meiotic prophase in nearly all other studied eukaryotes (reviewed in Zickler and Kleckner 1998, Franklin and Cande 1999). The comparatively lengthy and vigorous nature of dinoflagellate nuclear cyclosis may be related to the unique structure of dinoflagellate chromosomes, and also to the typically large dinoflagellate genome (Spector

1984, Rizzo 1991). It has been proposed that meiotic homologue pairing events in organisms with long chromosomes in a crowded nucleus should create an especially prominent stirring of the nuclear contents (Zickler and Kleckner 1998).

Katodinium (Gymnodinium) fungiforme Anissimova, Paulsenella spp.,

Crypthecodinium cohnii, Pfiesteria spp., and cryptoperidiniopsoids all reproduce in temporary division cysts, and the occurrence of nuclear cyclosis was used to distinguish presumed zygotic cysts from otherwise identical vegetative cysts (Spero and Morée 1981,

Drebes and Schnepf 1988, Ucko et al. 1997, Parrow et al. 2002, Parrow and Burkholder

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accepted, this study). Production of two swimming cells from cysts after nuclear cyclosis occurred commonly in C. cohnii, Pfiesteria spp., and cryptoperidiniopsoids, and also occurred after nuclear cyclosis in the isolated, verified zygotic cysts of a few photosynthetic species (von Stosch 1972, 1973, Kita and Fukuyo 1993). Litaker et al. (2002) concluded that all P. piscicida cysts producing four flagellated offspring were zygotes, whereas all cysts producing two flagellated offspring were vegetative. This simplification is incorrect if nuclear cyclosis precedes meiosis in P. piscicida (Parrow et al. 2002). Unlike the coordinated divisions of conventional meiosis, the two postzygotic divisions of many dinoflagellate species are uncoordinated, and separated by an interval similar to the vegetative generation time (von Stosch 1973, Beam and Himes 1980, Coats et al. 1984,

Pfiester 1989).

Variations in sexuality occur among dinoflagellate species regarding cues for sexual induction, gamete morphology and fusion orientation, sterility factors, zygote morphology and development, and the timing and apparent segregational nature of meiosis (Beam and

Himes 1980, 1984, Pfiester 1984, 1989, Pfiester and Anderson 1987). These sexual phase variables are incompletely known for many species with described sexuality, and some may vary within species depending upon isolate or culture conditions (Uchida 2001). The complete sexual cycle of a dinoflagellate species can be difficult to document for a variety of reasons. Gametes usually look similar or identical to vegetative cells (Coats et al. 1984,

Blackburn et al. 1989, Gao et al. 1989, Probert et al. 1998), and fusion can be confused with division (Pfiester 1989). Sexual events can occur in low frequency (Ouchi et al. 1994, Seo

& Fritz 2001), and zygotes may fail to develop in culture (Walker 1984, Giacobbe &

Gangemi 1997, Steidinger et al. 1998). In some species certain zygotic stages are similar or

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identical in external morphology to other vegetative cells (Beam and Himes 1980, Drebes and Schnepf 1988, Giacobbe and Yang 1999).

Documentation of sexuality in P. shumwayae was difficult because zygotes were rare or absent in clonal cultures, and planozygotes were difficult to distinguish and follow in live cultures due to their small size and rapid speed. That nuclear cyclosis distinguishes zygotic cysts could be demonstrated in P. shumwayae by following the development of isolated fusing gametes in microcultures, as in von Stosch (1972, 1973) and Kita et al.

(1993). The reproductive cycle of P. shumwayae, like several other heterotrophic species, appears adapted to take advantage of fluctuant food availability and quality. Flagellated cells can persist for weeks without food, but eventually quiescent cysts are formed that apparently await more favorable conditions. When food is abundant or of high quality, the dinoflagellates can ingest sufficient material to produce multiple offspring. The sexual cycle may be influenced by self-sterility factors that favor outbreeding. Unlike many photosynthetic species, sexuality in P. shumwayae and several other heterotrophic dinoflagellates does not appear to be obligately linked to a dormant survival strategy, although that may occur under some environmental conditions. The possible influences of compatibility and sterility factors in P. shumwayae sexuality merits further exploration, as does the cytology of meiosis. Further studies are also needed on cell and population level nutrition and growth in P. shumwayae and related species, and the relative importance of asexual and sexual processes in population occurrence, survival, and distribution.

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Acknowledgments

We thank Dr. P. Rublee and Dr. D. Oldach for PCR assistance, E. Allen for SEM assistance, N. Deamer and K. Brewjo for culturing assistance, and H. Glasgow for general assistance. Funding for this research was provided by the North Carolina General

Assembly, the National Science Foundation (#9912089), the U.S. Environmental Protection

Agency (#CR 827831-01-0), and the Department of Botany at North Carolina State

University.

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